Production of Outer Membrane Vesicles and Outer Membrane Tubes ...

Production of Outer Membrane Vesicles and Outer Membrane Tubes by Francisella novicida William D. McCaig,a Antonius Koller,b David G. Thanassia Center for Infectious Diseases and Department of Molecular Genetics and Microbiologya and Department of Pathology,b Stony Brook University, Stony Brook, New York, USA

Francisella spp. are highly infectious and virulent bacteria that cause the zoonotic disease tularemia. Knowledge is lacking for the virulence factors expressed by Francisella and how these factors are secreted and delivered to host cells. Gram-negative bacteria constitutively release outer membrane vesicles (OMV), which may function in the delivery of virulence factors to host cells. We identified growth conditions under which Francisella novicida produces abundant OMV. Purification of the vesicles revealed the presence of tube-shaped vesicles in addition to typical spherical OMV, and examination of whole bacteria revealed the presence of tubes extending out from the bacterial surface. Recently, both prokaryotic and eukaryotic cells have been shown to produce membrane-enclosed projections, termed nanotubes, which appear to function in cell-cell communication and the exchange of molecules. In contrast to these previously characterized structures, the F. novicida tubes are produced in liquid as well as on solid medium and are derived from the OM rather than the cytoplasmic membrane. The production of the OMV and tubes (OMV/T) by F. novicida was coordinately regulated and responsive to both growth medium and growth phase. Proteomic analysis of purified OMV/T identified known Francisella virulence factors among the constituent proteins, suggesting roles for the vesicles in pathogenesis. In support of this, production of OM tubes by F. novicida was stimulated during infection of macrophages and addition of purified OMV/T to macrophages elicited increased release of proinflammatory cytokines. Finally, vaccination with purified OMV/T protected mice from subsequent challenge with highly lethal doses of F. novicida.

F

rancisella tularensis is the causative agent of the zoonotic disease tularemia. F. tularensis is a Gram-negative, facultative intracellular pathogen that is capable of invading and replicating within a variety of host cell types. F. tularensis can be acquired by humans via several routes of infection (1). The most serious infections result from inhalation of aerosolized bacteria, a route that, if untreated, leads to a pneumonic form of tularemia with mortality rates as high as 60% (2). As a result of its high infectivity and low infectious dose (as few as 10 organisms), F. tularensis is classified as a category A agent of bioterrorism by the Centers for Disease Control and Prevention (http://www.bt.cdc.gov/agent/agentlist -category.asp). There are two clinically relevant subspecies of F. tularensis: subsp. tularensis, which is highly virulent, and subsp. holarctica, which causes a milder disease (1). An attenuated live vaccine strain (LVS) was derived from a subsp. holarctica strain, but the basis for its attenuation is not fully understood (2). An additional species of Francisella, F. novicida, has low virulence in humans but has proven highly useful as an experimental strain. F. novicida infection of host cells and pathogenesis in mice shares many similarities with F. tularensis, and the Francisella strains are ⬎98% similar at the genomic level (3). The molecular mechanisms underlying the extreme virulence of F. tularensis are just beginning to be understood. Although Francisella has a detectable extracellular phase in the host, the bacteria are thought to replicate primarily within the intracellular niche during infection (4, 5). Francisella are engulfed by macrophages within asymmetrical, spacious pseudopod loops (6), forming a Francisella-containing phagosome. Francisella prevent acidification and maturation of the phagosome, and within ca. 1 to 4 h escape into the host cell cytosol, where bacterial replication occurs (7, 8). The intracellular bacteria eventually activate host cell death pathways, leading to the release and spread of Francisella (9, 10).

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Critical to escape from the phagosome and intracellular replication is the activity of genes residing within the Francisella pathogenicity island (FPI), a conserved genomic region that is essential for virulence (11, 12). Another key feature of Francisella virulence is the ability of the organism to escape or subvert host immune responses. In contrast to typical lipopolysaccharide (LPS), the LPS expressed by Francisella has low proinflammatory activity and does not signal host cells through Toll-like receptor 4 (TLR4) (13, 14). In addition, the bacteria actively interfere with intracellular signaling pathways and the innate immune responses of host cells (7, 9, 15–18). Francisella spp. lack pathways typically used by intracellular Gram-negative pathogens to deliver virulence factors into host cells, such as the type III and type IV secretion systems (3). However, proteins encoded by the FPI share homology with components of the recently described type VI secretion system (T6SS), and experimental evidence supports a role for the FPI in the delivery of Francisella proteins to host cells (19). In addition to the putative T6SS, Francisella spp. encode a type I secretion system and a type IV pilus assembly pathway, both of which function in the secretion of proteins to the extracellular environment and contribute to pathogenesis (17, 20–22). Given its highly virulent and intracellular nature, Francisella spp. likely possess additional

Received 19 October 2012 Accepted 12 December 2012 Published ahead of print 21 December 2012 Address correspondence to David G. Thanassi, [email protected]. Supplemental material for this article may be found at http://dx.doi.org/10.1128 /JB.02007-12. Copyright © 2013, American Society for Microbiology. All Rights Reserved. doi:10.1128/JB.02007-12

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Francisella Outer Membrane Vesicles and Tubes

secretion systems, particularly for the delivery of virulence factors to the host cell cytosol. Outer membrane vesicles (OMV) are shed from the bacterial surface during normal growth of Gram-negative bacteria. Their production has been reported for nearly 40 years (23); however, interest in OMV has increased recently since it has become clear that they provide a mechanism for the secretion of virulence factors (24). Moreover, OMV are capable of delivering their cargo into the host cell cytoplasm, either through fusion with the plasma membrane or internalization via endocytic routes (25–27). There are several proposed models for the biogenesis of OMV, but the mechanisms by which they are created, whether they are produced in a regulated manner, and how specific proteins might be targeted to OMV have yet to be determined (28, 29). Previous ultrastructural studies of Francisella spp. during infection of macrophages noted the presence of vesicular material surrounding intracellular bacteria, suggesting that Francisella may release OMV during infection (30, 31). More recently, Pierson et al. demonstrated the production of OMV by F. novicida (32). The constituents of the F. novicida OMV included proteins known to contribute to the pathogenesis of tularemia and the OMV had activity against host cells (32). Thus, as found for other bacterial pathogens, Francisella may use OMV as a secretion system for the delivery of virulence factors to host cells. Pierson et al. examined OMV isolated from bacteria grown to late stationary phase (32). We show here that F. novicida also produces OMV during exponential and early stationary phases of growth. We identify specific conditions that favor production of OMV by F. novicida, showing that they are generated in a regulated manner. We demonstrate that in addition to producing OMV, F. novicida creates tubular extensions of its OM and that these tubes are released into the growth medium along with OMV. Our characterization of the F. novicida OMV and tubes (OMV/T) supports functions for these structures in pathogenesis and suggests that they may provide the basis for an effective componentbased vaccine. MATERIALS AND METHODS Bacterial strains and growth conditions. The media used for the growth of F. novicida strain U112 (ATCC 15482) were as follows: TS (tryptic soybean powder [BD Biosciences] at 30 g/liter, supplemented with 0.1% cysteine), BHI (brain heart infusion powder [BD Biosciences] at 37 g/liter, adjusted to pH 6.8), MHB (Mueller-Hinton II broth powder [BD Biosciences] at 22 g/liter, supplemented with 625 ␮M CaCl2, 530 ␮M MgCl2, 335 ␮M Ferric pyrophosphate, 5.6 mM D-glucose, and 2% IsoVitaleX), and CDM, which was made as described previously (33). For plates, Bacto agar (BD Biosciences) was added to 15 g/liter. Bacteria streaked on plates were incubated at 37°C in the presence of 5% CO2. Liquid media were incubated in the presence of 5% CO2 for 1 h prior to inoculation with bacteria, and cultures were grown at 37°C with shaking at 100 rpm. Starter liquid cultures were inoculated directly from frozen stocks, grown overnight, and diluted 1:100 to an optical density at 600 nm (OD600) of ⬃0.01. Day cultures were grown to exponential (OD600 of 0.5 to 0.8) or stationary (OD600 of 1.2 to 1.4) phase, as indicated. Purification of OMV/T. Bacterial cultures were grown in BHI in 400-ml batches in 2-liter flasks with baffles to exponential (OD600 of ⬃0.6) or early stationary (OD600 of ⬃1.4) phase. Bacteria were removed by successive low-speed centrifugation (5,000 ⫻ g, followed by 7,500 ⫻ g, 30 min for each), and the medium was passed through a 0.2-␮m-pore-size MF75 filter unit (Nalgene). Sodium azide was added to the cleared medium to 0.05%, and vesicles were harvested by ultracentrifugation (100,000 ⫻ g, 1 h, 4°C). For bacteria grown to exponential phase, prior to


ultracentrifugation, 1 liter of cell-free medium was concentrated to ⬃50 ml using a tangential flow filtration unit (Pall) with a 100-kDa molecular mass cutoff membrane. The pelleted OMV/T were resuspended in buffer A (20 mM HEPES [pH 7.5], 0.05% sodium azide), subjected to an additional ultracentrifugation step (100,000 ⫻ g, 1 h, 4°C), and resuspended again in buffer A. The vesicles were then adjusted to 40% (vol/vol) OptiPrep (Axis-Shield) in buffer A in a total volume of 2 ml. The samples were loaded into a 13.2-ml ultracentrifuge tube, and lower-concentration OptiPrep solutions were layered on top (2 ml [35%], 2 ml [30%], 2 ml [25%], 2 ml [20%], 1 ml [15%], and 0.5 ml [0%]). The tubes were centrifuged (100,000 ⫻ g, 16 h, 4°C) in a swinging-bucket rotor, and then 1-ml fractions were collected from the top. Fractions were examined by SDSPAGE and Coomassie blue staining for protein content. Adjacent fractions with similar protein profiles were combined and diluted with 20 mM HEPES (pH 7.5) and recovered via ultracentrifugation (100,000 ⫻ g, 1 h, 4°C). Recovered pellets were resuspended in 20 mM HEPES (pH 7.5) containing 1% penicillin-streptomycin and 10 ␮g of gentamicin/ml, and aliquots were flash-frozen in liquid nitrogen and stored at ⫺80°C. The concentrations of OMV/T were determined using a bicinchoninic acid (BCA) protein assay (Pierce), according to the manufacturer’s instructions, with the addition of 2% SDS (34). Transmission electron microscopy (TEM). Samples were adsorbed to polyvinyl formal-carbon-coated grids (EMS) for 2 min, fixed with 1% glutaraldehyde for 1 min, washed twice with phosphate-buffered saline (PBS) and twice with water, and then negatively stained with 0.5% phosphotungstic acid (Ted Pella) for 30 s. For preparation of whole bacterial samples from liquid media, 1 ml of culture was centrifuged (8,000 ⫻ g, 5 min, 4°C) and then resuspended in 200 ␮l of PBS. For the preparation of whole bacterial samples from solid media, five colonies were directly suspended in 50 ␮l of PBS. Samples for thin sectioning were fixed with 2.5% EM-grade glutaraldehyde in 0.1 M PBS (pH 7.4) for at least 1 h. The samples were then placed in 1% osmium tetroxide in 0.1 M PBS, dehydrated in a graded series of ethyl alcohol, and embedded in Durcupan resin. Ultrathin sections of 80 nm were cut with a Reichert-Jung UltracutE ultramicrotome and placed on Formvar-coated slot copper grids. Sections were counterstained with uranyl acetate and lead citrate. All grids were viewed in a FEI Tecnai12 BioTwinG2 electron microscope at an 80-kV accelerating voltage, and images were obtained using an AMT XR-60 charge-coupled device digital camera system and compiled using Adobe Photoshop. Fractionation of F. novicida. Bacterial cultures were grown in BHI to early stationary phase (OD600 of ⬃1.4). For whole-cell lysates, 1 ml of culture was centrifuged (10,000 ⫻ g, 5 min, 4°C), the pellet was resuspended in 100 ␮l of SDS-PAGE sample buffer and heated at 95°C for 10 min. For bacterial fractionation, 100 ml of culture was centrifuged (10,000 ⫻ g, 5 min, 4°C), and the supernatant was removed and saved for analysis of secreted proteins as described below. The bacterial pellet was resuspended in 10 ml of 20 mM Tris-HCl (pH 8.0), moved to a fresh tube, and centrifuged again. The pelleted bacteria were resuspended in 1 ml of 20 mM Tris-HCl (pH 8.0) plus 20% sucrose, and 1 ml was moved to a clean tube. EDTA was added to 15 mM, lysozyme was added to 200 ␮g/ml, and the suspension was incubated on ice for 40 min. MgCl2 was then added to 26 mM, 4 ␮l of DNase I (10,000 U/ml; Thermo Scientific) was added, and the suspension was incubated for an additional 20 min on ice. The spheroplasted bacteria were pelleted by centrifugation (10,000 ⫻ g, 20 min, 4°C), and the supernatant was collected (equals periplasm sample). The bacterial pellet was resuspended in 1 ml of 20 mM Tris-HCl (pH 8.0) plus Complete protease inhibitor cocktail (Roche), and 1 ml was transferred to a clean tube. The sample was sonicated (Misonix Microson model XL-2000; power level 5) on an ice water bath, 15 s on and 15 s off, for 2 min. The sonicated bacteria then centrifuged (8,000 ⫻ g, 10 min, 4°C) to remove unbroken cells, and the supernatant was then ultracentrifuged (100,000 ⫻ g, 1 h, 4°C) to pellet membranes. The membrane pellet was resuspended in 1 ml of 20 mM Tris-HCl (pH 8.0) plus Complete protease inhibitor cocktail and Sarkosyl (sodium-n-lauroyl-sarcosinate; Fisher) was added to a final concentration

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of 0.5% to solubilize the cytoplasmic membrane. The tube was rocked at room temperature for 5 min, and then ultracentrifuged (100,000 ⫻ g, 1 h, 4°C) to pellet the OM. The final pellet was resuspended in 20 mM TrisHCl (pH 8.0)– 0.3 M NaCl (equals OM sample). Protein concentrations were determined by using the BCA assay, and aliquots were mixed with SDS-PAGE sample buffer and heated at 95°C for 10 min. For analysis of the secreted proteins, 3-ml aliquots of the saved culture supernatants were filtered through 0.22-␮m-pore-size syringe filters (Sarstedt), and sodium azide was added to 0.05%. Trichloroacetic acid was then added to 1 ml of the filtered supernatants to a 9% final concentration. Samples were placed on ice for 30 min and centrifuged (16,000 ⫻ g, 4°C, 5 min) to pellet precipitated proteins. The pellets were washed twice with ice-cold acetone, followed each time by centrifugation (16,000 ⫻ g, 4°C, 5 min). Final pellets were resuspended in 20 ␮l of SDS-PAGE sample buffer and heated at 95°C for 10 min. All samples were subjected to SDS-PAGE and stained with Coomassie blue. Protease accessibility assay. Purified OMV/T were left untreated or treated with proteinase K (10 ␮g/ml), SDS (0.02%), or proteinase K plus SDS for 1 h at room temperature. Phenylmethylsulfonyl fluoride (0.1 mM) was added to inhibit the protease, and samples were processed for TEM as described above or heated at 95°C for 10 min in SDS sample buffer for subsequent SDS-PAGE analysis. For immunoblotting, the proteins separated by SDS-PAGE were transferred to a polyvinylidene difluoride (Osmonics) membrane and probed with 1:10,000 anti-FopA (35) or antiFipB (36) antibodies. Immunoblots were developed with alkaline phosphatase-conjugated secondary antibodies and BCIP (5-bromo-4-chloro3-indolylphosphate)/NBT (nitroblue tetrazolium) substrate (KPL). Heat treatment of OMV/T. Purified OMV/T were left at room temperature, heated at 60°C for 5, 15, or 30 min, or 80°C for 1 h, and then left to cool at room temperature for 1 h. Samples were processed for TEM as described above. For quantification of OMV/T at different time points during the 60°C heat treatment, 10 random TEM fields at each time point were chosen, and the numbers of OMV or tubes were determined by visual inspection. The values for the 10 fields were then averaged. Lysozyme treatment of OMV/T and whole bacteria. Treatment of bacteria to generate spheroplasts was performed as described above for bacterial fractionation, except that after the 40-min incubation in the presence of lysozyme-EDTA, a 20-␮l aliquot was removed and processed for TEM as described above. For the treatment of purified OMV/T, the samples were pelleted and resuspended in 20 mM Tris-HCl (pH 8.0), EDTA was added to 15 mM, lysozyme was added to 200 ␮g/ml, and the suspension was incubated on ice for 40 min before processing for TEM. MudPIT. Detailed mass spectrometry and data analysis methods are provided in the supplemental material. Purified OMV/T were analyzed using the multidimensional protein identification technology (MudPIT) method (37). Spectral count normalization (38) was applied to spectra identified for each independent OMV/T sample isolated to generate a normalized spectral abundance factor (NSAF) value. Three independent experiments were performed at each time point, and only proteins identified in all experiments at their respective time points were considered vesicle associated. Predicted localization of F. tularensis subsp. novicida U112 proteins was determined using the PSORTb v3.0 (39) precomputed proteome. Comparison of individual F. novicida proteins to F. tularensis Schu S4, F. tularensis LVS, or to the OMV-associated content of other bacteria was accomplished using BLAST (Basic Local Alignment Search Tool) (40). Preparation of macrophages. Murine bone marrow-derived macrophages (muBMDM) were obtained as previously described (41) from C3H/HeN mice (Charles River) and cultured for 5 days in bone marrow medium (BMMHI) composed of Dulbecco modified Eagle medium (DMEM; Invitrogen) containing 2 mM L-glutamine, 1 mM sodium pyruvate, 20% heat-inactivated fetal bovine serum (FBS; HyClone), and 30% medium previously conditioned by L929 cells. The muBMDM were seeded into 24-well plates at a concentration of 1.5 ⫻ 105 cells per well in bone marrow assay medium (BMAM) composed of DMEM (Invitrogen)

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containing 2 mM L-glutamine, 1 mM sodium pyruvate, 1% heat-inactivated FBS (HyClone), 1% penicillin-streptomycin, and 4 ␮g of gentamicin/ml. The muBMDM were then incubated at 37°C and 5% CO2 and used for experiments the next day. The L-cell conditioned medium was obtained by plating 2 ⫻ 105 L929 cells in 75-cm2 culture flasks in minimum essential medium (Invitrogen) containing 2 mM L-glutamine, 1 mM sodium pyruvate, 1 mM nonessential amino acids (Invitrogen), and 10% FBS and then collecting the medium after 10 days. All protocols involving animals were approved by the Institutional Animal Care and Use Committee of Stony Brook University. Cytotoxicity assays. Purified OMV/T were resuspended in room temperature BMAM at concentrations of 0.1, 1, 10, and 20 ␮g/ml. The supernatant was removed from muBMDM previously seeded into 24-well plates, the cells were washed twice with room temperature PBS, and 1 ml of vesicle-containing medium was added to the wells. The plates were incubated at 37°C and 5% CO2 for 24 or 48 h, the supernatants were collected, and a lactate dehydrogenase (LDH) assay (CytoTox 96 nonradioactive cytotoxicity assay; Promega) was performed according to the manufacturer’s instructions. Background LDH release was measured in medium lacking vesicles, while total LDH release (⫽100%) was measured from uninfected cells that were lysed by freezing and thawing. The percentage of LDH release was calculated by subtracting the background LDH release value from the LDH release value of the samples, and this number was then divided by the total LDH release value and multiplied by 100. The values for each experiment were determined from the average of triplicate wells; three independent experiments were performed. Detection of cytokine secretion. OMV/T were added to muBMDM as described above. After 24 h of incubation, conditioned media from the wells were clarified by centrifugation (200 ⫻ g, 5 min) and stored at ⫺20°C until assayed. Quantikine ELISA kits (R&D Systems) were used to detect tumor necrosis factor alpha (TNF-␣), CCL2, or CXCL2 release from the macrophages according to the manufacturer’s instructions. Heat and protease treatment of the OMV/T was performed as described above. A sham OMV/T preparation was created by incubation of 400 ml of BHI media (without bacteria) in a 2-liter flask, followed by all steps, as done for purification of OMV/T. The final ultracentrifugation tubes containing the sham “pellet” were washed with 20 mM HEPES buffer containing 1% penicillin-streptomycin and 10 ␮g of gentamicin/ml, and aliquots were flash frozen and stored at ⫺80°C. The values for each experiment were determined from the averages of triplicate wells; three independent experiments were performed for the dose-response experiments, and two independent experiments were performed for the experiments involving heat and proteinase K treatment. Macrophage coincubation with F. novicida for TEM. Coincubation experiments were performed as previously described for phagocytic uptake analysis with minor modifications (42). MuBMDM were resuspended at a concentration of 6 ⫻ 106 in BMMHI. Cells were pelleted (1,000 ⫻ g, 10 min, 4°C), and the supernatant was removed. A 1-ml portion of F. novicida in BMMHI was added to the pelleted macrophages at an approximate multiplicity of infection of 2,000. The tube was centrifuged twice (200 ⫻ g, followed by 800 ⫻ g, for 10 min each time, 4°C), and the supernatant was removed. The tube containing pelleted bacteria and cells was placed in a 37°C water bath for 5 min. Cells and bacteria were then fixed with 1 ml of 2.5% glutaraldehyde at 37°C for 2 min, followed by incubation on ice for 30 min. The tube was centrifuged (10,000 ⫻ g, 10 min, 4°C), and the pellet was resuspended in 1 ml of ice-cold PBS. The sample was then processed for thin sectioning and TEM as described above. Vaccination and challenge experiments. Groups of 3 or 4 BALB/c mice (6 to 8 weeks old; Charles River) were intranasally inoculated with 20 ␮g of purified OMV/T in 20 ␮l of PBS or with PBS alone as a control. At 6 weeks after vaccination, mice were challenged intranasally with 620 (n ⫽ 3) or 960 (n ⫽ 4) CFU of F. novicida grown in BHI broth to exponential phase. The infectious doses were determined by retrospective CFU counts. Mice were monitored for 21 days after bacterial challenge.



Statistical analysis. Cytotoxicity and cytokine release results were analyzed for significance using data obtained from three independent experiments with three replicates each, unless otherwise noted. Probability (P) values were calculated by one-way analysis of variance and Bonferroni’s multiple-comparison post test against the negative control value. The log-rank test was used to calculate the P value for the mouse challenge experiments. Statistical calculations were performed using Prism 4.0 (GraphPad Software). P values of ⬍0.05 were considered significant.

RESULTS

F. novicida produces OMV and OM tubes in response to growth phase and medium. The F. novicida OMV characterized by Pierson et al. were isolated from bacteria after 44 h of growth (32), a time in late stationary phase when many bacteria are likely dying and releasing contents due to cell lysis. The protein composition and other properties of OMV may change with growth phase (43), and vesicles produced by dying bacteria are expected to be different from vesicles produced during bacterial growth. Therefore, we examined F. novicida strain U112 for the production of OMV at earlier stages of growth. After dilution of an overnight culture to an OD600 of 0.01, strain U112 grows exponentially in brain heart infusion (BHI) broth for ⬃8 h, until reaching an OD600 of ⬃1.0 (see Fig. S1 in the supplemental material). The bacteria then enter stationary phase and remain at an OD600 of ⬃1.4 for an additional 16 h before beginning to decrease in OD, indicating cell death (see Fig. S1 in the supplemental material). We chose mid-exponential-phase (OD600 ⫽ 0.6, ⬃4 h of growth) and early-stationary-phase (OD600 ⫽ 1.4, ⬃9 h of growth) time points to examine strain U112 for the production of OMV. Cell-free culture supernatant fractions from bacteria grown in BHI to the two time points were ultracentrifuged to harvest OMV. The OMV pellets were then purified by flotation through an OptiPrep density gradient. The majority of proteins in the pellets floated to the top, lower-density region of the gradient, as expected for vesicle-associated proteins (see Fig. S2 in the supplemental material). We recovered vesicles from this same lowerdensity region of the gradient, but not from other fractions, confirming flotation of the vesicles up the gradient. OMV pellets were obtained for both the exponential- and stationary-phase BHI cultures. However, the exponential-phase bacteria produced fewer vesicles compared to the stationary-phase cultures; 2 liters of exponential-phase supernatant yielded ⬃0.25 mg of purified vesicles, whereas 2 liters of stationary-phase supernatant yielded ca. 1 to 2 mg of purified vesicles. Thus, there was a 4- to 8-fold increase in vesicle yield despite an only ⬃2 fold increase in the bacterial OD. In addition, we were unable to isolate OMV from strain U112 grown to either exponential or early stationary phase in different rich media—tryptic soy (TS) or Mueller-Hinton (MH) broth— demonstrating that vesicle production by F. novicida is responsive to growth media as well as growth phase. Examination of the gradient-purified vesicles by TEM revealed the presence of typical, spherical OMV for both the exponentialand stationary-phase BHI cultures (Fig. 1). Surprisingly, elongated, tube-shaped vesicles were also present in the samples from both growth phases (Fig. 1). Although there were differences in the staining of the EM grids (Fig. 1), the overall appearance and relative numbers of spherical and tube-shaped vesicles was similar for the two time points. The spherical OMV ranged from ⬃50 to 300 nm in diameter; the tubular vesicles were ⬃40 nm in diameter and ranged from ⬃0.3 to 1.5 ␮m in length. Previous studies noted the presence of large protrusions on the surface of Francisella spp.,


FIG 1 TEM images of purified OMV/T. OMV/T were purified from U112 cultures grown in BHI broth to exponential phase (A) or early stationary phase (B). The purified samples from both time points contain mixtures of spherical (white arrows) and tubular (black arrows) vesicles. Scale bars, 500 nm.

similar in appearance to the tubular vesicles (44, 45). In addition, recent publications have described morphologically similar structures, termed nanotubes, extending from the surfaces of bacteria grown on solid medium (46, 47). However, production of nanotubes by liquid-grown bacteria or the release of nanotubes into the culture medium has not been reported. To determine whether F. novicida produced tubes on its cell surface, we examined bacteria grown to early stationary phase in BHI broth. TEM imaging of whole bacteria revealed the presence of tubes projecting out from the bacterial surface, similar in diameter and appearance to the tubular structures present in the purified vesicles (Fig. 2A). As expected, spherical OMV were also frequently observed associated with the bacterial surface or released into the surrounding medium (data not shown). Nanotubes produced by Bacillus subtilis were shown to be extensions of the cytoplasmic membrane and to connect neighboring bacteria to allow the exchange of cytoplasmic constituents (47). In contrast, as revealed by thin section TEM, the F. novicida tubes are formed by extensions of the OM (Fig. 2B). In addition, most of the tubes were not in contact with neighboring bacteria (Fig. 2 and 3), arguing against a role in direct bacterial-bacterial bridging. As found for the vesicles, production of tubes on the surface of strain U112 was greater for bacteria grown to early stationary compared to exponential phase, and for bacteria grown in BHI compared to TS broth (data not shown). Plate-grown bacteria also produced OM tubes (Fig. 3). Tubes produced by plate-grown bacteria were more numerous and longer than those seen in broth-grown cultures. This suggests that the tubes may be sensitive to shear forces generated during growth in liquid cultures. However, many of the tubes produced by the plate-grown bacteria were also detached from the bacterial surface, suggesting that release of the OM tubes is not solely driven by shear forces (Fig. 3). Similar to broth-grown F. novicida, we observed a dramatic decrease in the amount of tubes produced by bacteria grown on TS compared to BHI agar (Fig. 3). Bacteria grown on MH or Chamberlain’s defined medium (CDM) plates also had markedly fewer tubes compared to BHI-grown bacteria (data not shown). Taken together, these results show that production of tubes on the bacterial surface and release of OMV/T into the culture medium are similarly regulated processes and responsive to both growth medium and growth phase. Initial characterization of the F. novicida OMV/T. We were unable to separate the OMV from the tubes using either density

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FIG 2 F. novicida produce tubes on their cell surfaces that are extensions of the OM. (A) TEM image of U112 whole bacteria grown in BHI broth to early stationary phase. The asterisks mark whole bacteria and the arrows point to tubes. Scale bar, 500 nm. (B and C) Thin-section TEM images of bacteria grown as in panel A. The bacterial cytoplasm is marked with an asterisk, the locations of the periplasm (Peri) and OM are indicated, and the arrowhead points to a tube. Scale bars, 100 nm.

gradient flotation or velocity sedimentation, suggesting that the OMV/T are similar in composition (W. D. McCaig and D. G. Thanassi, unpublished data). Analysis of the purified OMV/T by silver staining or immunoblotting with an anti-LPS antibody confirmed the presence of LPS in the vesicles, consistent with their derivation from the OM (McCaig and Thanassi, unpublished). The purified OMV/T had a distinct protein profile compared to U112 total cell lysates, periplasm, OM, and total secreted proteins (Fig. 4). This differential protein profile is in keeping with OMV from other bacteria (48–51). Notably, there were substantial differences in the protein profiles for OMV/T isolated from exponential- versus stationary-phase cultures (Fig. 4). Thus, the content of the F. novicida vesicles changes with growth phase. A similar dynamic protein content has been reported for OMV isolated from P. aeruginosa (43). To obtain a qualitative measure of luminal versus surface-exposed proteins present in the OMV/T, we incubated purified vesicles with proteinase K in the absence or presence of 0.02% SDS. Addition of 0.02% SDS disrupted the integrity of both the OMV and OM tubes (data not shown), allowing access of the protease to the interior content of the vesicles. Incubation of purified OMV/T with proteinase K alone resulted in loss of a number of presumably

FIG 3 TEM images of whole bacteria isolated from solid medium. U112 bacteria were isolated from TS (A) or BHI (B) agar plates. The white arrows indicate whole bacteria, and the black arrows indicate tubes. There is increased tube production for BHI-grown bacteria. Scale bars, 500 nm.

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surface-exposed proteins, but the overall protein profile was mostly unchanged (Fig. 5A). Incubation of the samples with SDS alone had no effect on the protein profile; however, incubation with proteinase K in the presence of 0.02% SDS resulted in a dramatic loss of protein bands (Fig. 5A), showing that a large number of proteins are protected by the intact vesicles. These results suggest that, as shown for other OMV, these protected proteins could be delivered to the cytoplasm of host cells via fusion of the vesicles with the plasma membrane or endocytosis (25–27). Of note, incubation of the OMV/T with proteinase K alone did not cause changes in the appearance or number of tubes (data not shown), indicating that the structural component for the formation of the tubes is not a surface-accessible protein. The tubes are formed by extension of the bacterial OM (Fig. 2B) and thus could be structured by an internal peptidoglycan backbone. To test this, we treated whole bacteria with EDTA

FIG 4 Protein profiles of F. novicida cellular fractions and purified OMV/T. (A) Whole bacterial lysates (WB), OM, periplasm (Peri), total secreted proteins, and OMV/T were isolated from U112 bacteria grown in BHI broth to early stationary phase and subjected to SDS-PAGE and Coomassie blue staining. The abundant protein noted by the black arrowhead is lysozyme. (B) SDS-PAGE profile of OMV/T isolated from exponential-phase U112 grown in BHI medium. The abundant protein noted by the white arrowhead is Fsp53.



FIG 5 Protease accessibility and verification of specific OMV/T-associated proteins. OMV/T purified from U112 grown to early stationary phase were treated with proteinase K in the presence or absence of 0.02% SDS to disrupt vesicle integrity. The vesicles were subjected to SDS-PAGE and stained with Coomassie blue (A), blotted with anti-FopA (B), or blotted with anti-FipB (C). The white arrowheads in panel A point out proteins susceptible to proteinase K digestion in intact vesicles (no SDS added). The black arrowhead in panel C notes a fragment of FipB generated by proteinase K digestion of intact vesicles.

and lysozyme to digest the peptidoglycan and generate spheroplasts. TEM examination showed that the tubes remained intact on the spheroplasted bacteria (data not shown). In addition, incubation of purified vesicles with EDTA and lysozyme had no

effect on tube structure (data not shown). We next examined the sensitivity of the vesicles to heat treatment. Purified OMV/T were held at room temperature, incubated at 80°C for 1 h, or incubated at 60°C for 5, 15, or 30 min and then left to cool at room temperature. OMV/T were stable at room temperature, but incubation at 80°C caused a nearly complete disruption of the vesicles (data not shown). The tubes were sensitive to heat treatment, since no tubular vesicles remained after heating to 60°C for as little as 5 min (Fig. 6). In contrast, the number of spherical vesicles increased (Fig. 6B), suggesting denaturation of a factor responsible for structuring the tubes and conversion to spherical shape. Consistent with this, some vesicles at the 5-min time point appeared to be transitioning from a tubular to a spherical shape (Fig. 6A, arrowhead). Similar tubes in transition were observed in 50% of the images (n ⫽ 20) taken at the 5-min time point but were absent in images taken at other incubation times. The total numbers of remaining spherical vesicles decreased with longer incubation (Fig. 6), demonstrating a general sensitivity to lysis by heat. Taken together, these results indicate that a heat-sensitive factor, presumably a protein(s), is responsible for structuring the OM tubes. Identification of F. novicida OMV/T-associated proteins. To identify OMV/T-associated proteins, purified vesicles were analyzed by mass spectrometry using the MudPIT method (37). Three independent analyses were performed for each time point (exponential and early stationary phases), and only proteins appearing in all three analyses were considered as vesicle associated for that time point. A normalized spectral abundance factor (NSAF) was used to quantify the relative amounts of individual proteins in each sample. The MudPIT analysis identified 99 proteins from the exponential-phase vesicles and 286 proteins from the stationary-phase vesicles (see Tables S1 and S2 in the supple-

FIG 6 Sensitivity of OMV/T to heat. OMV/T purified from early-stationary-phase U112 were held at room temperature (RT) or incubated at 60°C for 5, 15, or 30 min. (A) TEM images of OMV/T heated as indicated. The arrows point to tubular vesicles present in the RT-treated sample. The black arrowhead notes a vesicle in the sample heated for 5 min that appears to be transitioning from tubular to spherical shape. Scale bars, 500 nm. (B) Quantitation of tubular and spherical vesicles per TEM field at each time point. Bars represent the means ⫾ the standard errors of the means (SEM) from 10 fields. **, P ⬍ 0.01 for comparison of heated with room temperature OMV.


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TABLE 1 The fifteen most abundant OMV/T-associated proteins Gene

Locus

Molecular mass (Da)

Description

NSAF

Exponential phase fsp53 fopB lpnA Unknown Unknown Unknown Unknown fopA atpF Unknown Unknown ompH pal fipB Unknown

FTN_1261 FTN_0119 FTN_0427 FTN_1451 FTN_1734 FTN_0714 FTN_0340 FTN_0756 FTN_1650 FTN_0429 FTN_0428 FTN_1481 FTN_0357 FTN_0771 FTN_1448

54,755 19,437 15,802 19,786 13,688 196,823 12,166 41,176 17,369 19,364 17,692 18,839 23,261 39,446 51,986

Hypothetical protein OM protein of unknown function Lipoprotein of unknown function Hypothetical protein Hypothetical protein Hypothetical protein Hypothetical protein OmpA family protein F0F1 ATP synthase subunit B Hypothetical protein Hypothetical protein OM protein OmpH Peptidoglycan-associated lipoprotein Protein-disulfide isomerase Hypothetical protein

0.0975 0.0472 0.0387 0.0377 0.0286 0.0278 0.0237 0.0221 0.0200 0.0177 0.0172 0.0145 0.0127 0.0127 0.0110

Early stationary phase fopB Unknown lpnA fipA ompH fipB tufA Unknown ahp1 Unknown pal ugpQ Unknown atpF Unknown

FTN_0119 FTN_1451 FTN_0427 FTN_0772 FTN_1481 FTN_0771 FTN_1576 FTN_1448 FTN_0973 FTN_1734 FTN_0357 FTN_0637 FTN_0120 FTN_1650 FTN_0643

19,437 19,786 15,802 10,466 18,839 39,446 43,390 51,986 21,746 13,688 23,261 38,840 15,823 17,369 18,148

OM protein of unknown function Hypothetical protein Lipoprotein of unknown function Hypothetical protein OM protein OmpH Protein-disulfide isomerase Elongation factor Tu Hypothetical protein AhpC/TSA family peroxiredoxin Hypothetical protein Peptidoglycan-associated lipoprotein Glycerophosphoryl diester phosphodiesterase Rhodanese-related sulfurtransferase F0F1 ATP synthase subunit B Hypothetical protein

0.0578 0.0454 0.0320 0.0262 0.0257 0.0217 0.0206 0.0164 0.0163 0.0160 0.0155 0.0103 0.0098 0.0096 0.0094

mental material), with a combined identification of 292 unique OMV/T-associated proteins. The 15 most-abundant proteins for each time point are listed in Table 1. Consistent with their different gel electrophoresis profiles (Fig. 4), there are a number of changes in proteins identified by mass spectrometry for exponential-phase versus stationary-phase vesicles. Most notably, although 93 of the 99 proteins present in exponential-phase OMV/T are also found in the stationary phase, the stationary-phase vesicles contain almost 200 additional proteins. The relative abundance of most of the 93 shared proteins remained consistent at both time points; however, 31 proteins exhibited ⬎2-fold changes in NSAF values between samples, with 28 being found in lower abundance and 3 in greater abundance in stationary-phase compared to exponential-phase OMV/T (see Table S3 in the supplemental material). The 292 unique OMV/T-associated proteins comprise ⬃17% of the F. novicida genome and are distributed among multiple functional categories and cellular locations (see Fig. S3 in the supplemental material). Approximately 16% of the vesicle-associated proteins were previously shown to be OM associated in Francisella spp. (see Table S4 in the supplemental material) (52, 53), and 20% have homologs that are OMV-associated in other bacteria (see Table S5 in the supplemental material) (48, 54, 55). Consistent with the derivation of the vesicles from the OM, OM-associated proteins are prominent among the most abundant vesicle-associ-

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ated proteins (comprising ⬃15% of NSAF values) and include the major Francisella antigens and T cell epitopes FopA, FopB, and LpnA (Table 1) (53, 56). FopA is an integral OM protein that is highly immunogenic and serves as a protective antigen for experimental tularemia caused by the LVS (35, 57). Immunoblotting with anti-FopA antibodies confirmed its presence in the purified OMV/T (Fig. 5B). Integral OM proteins are often resistant to protease digestion in intact bacteria and maintain some resistance to digestion when the OM is disrupted. In agreement with this, FopA was largely resistant to digestion by proteinase K in intact vesicles and only partly sensitive to digestion in the presence of 0.02% SDS (Fig. 5B), indicating the maintenance of proper protein structure and membrane integrity in the purified vesicles. Approximately 22% of the vesicle-associated proteins are secreted or associated with virulence in Francisella spp. (see Table S6 in the supplemental material) (11, 20, 36, 58–63). Four of these proteins—PepO, BglX, ChiA, and Fsp53—are secreted by strain U112 in a type IV pilus-dependent manner (20). Notably, Fsp53 is the most abundant protein identified in the exponential-phase OMV/T (Table 1) and appears as the most abundant band in the gel electrophoresis profile (Fig. 4). The identity of this band as Fsp53 was confirmed by mass spectrometry (data not shown). These results suggest that the secreted proteins might associate with the OMV/T for delivery to host cells, similar to the secretion of heat-labile enterotoxin in enterotoxigenic Escherichia coli (24,



49). Lee et al. identified 12 major extracellular proteins in cell-free culture supernatants of F. tularensis (60). Five of these proteins are present in the purified OMV/T, including the peroxidase/catalase KatG, the succinyl-CoA synthetase subunits SucC and SucD, the peroxiredoxin Ahp1, and the chaperonin GroEL (see Table S6 in the supplemental material). A number of FPI-associated proteins were also detected in the purified vesicles: IglB, IglC, IglI, PdpB, and PdpD. Each of these FPI proteins is essential for intramacrophage growth and Francisella virulence (12, 59, 64–68). Additional abundant OMV/T-associated proteins known to be virulence factors of Francisella spp. include FipB and the hypothetical proteins FTN_0714, FTN_0340, FTN_0429, and FTN_0643 (Table 1) (36, 59, 61, 62). Each of the hypothetical proteins was identified in transposon mutant screens of strain U112 as defective for colonization of mice after either pulmonary or intraperitoneal infection (59, 61, 62). All have predicted signal sequences and therefore are likely to be exported outside the cytoplasm where they could associate with the OMV/T. FipB was identified as an essential virulence factor in the fully virulent F. tularensis Schu S4 strain (36). FipB is a predicted lipoprotein with a DsbA periplasmic disulfide isomerase domain and a domain homologous to the surface-exposed Mip host cell invasion protein of Legionella pneumophila (69). F. novicida FipB is 98.9% identical to the Schu S4 protein. In addition to FipB, stationary-phase OMV/T contained high levels of the FipA protein (Table 1). FipA is encoded immediately upstream of fipB and also has homology with Mip proteins. FipA is not required for the virulence of F. tularensis in mice but contributes to intracellular survival and may influence the activity of FipB (36). We confirmed the presence of FipB in the purified vesicles by immunoblotting with anti-FipB antibody (Fig. 5C). FipB was partly protected from proteinase K digestion when vesicles were incubated with proteinase K alone, with a small amount of an ⬃30-kDa cleavage product appearing (Fig. 5C). In contrast, FipB was completely degraded by proteinase K in the presence of 0.02% SDS (Fig. 5C). This indicates a primarily luminal location for FipB, a finding consistent with a periplasmic location in the bacteria, but also suggests that at least some FipB may be surface exposed, similar to Mip proteins (69). The presence of multiple virulence factors and secreted proteins supports a role for the OMV/T in the pathogenesis of tularemia. Effects of the F. novicida OMV/T on host cells. OMV are enriched in immunostimulatory molecules, such as LPS and lipoproteins, and may contain toxins and other factors that are active against host cells (24, 25, 70–72). Although Francisella LPS is not proinflammatory (13, 14), the F. novicida OMV/T contain numerous lipoproteins, as well as known Francisella virulence factors and antigenic proteins (Table 1 and see Table S6 in the supplemental material). To determine effects of the F. novicida OMV/T on host cells, we first examined cytotoxicity using a lactate dehydrogenase (LDH) release assay. Increasing amounts of purified vesicles isolated from stationary-phase cultures were added to muBMDM, and LDH release was measured after 24 and 48 h of incubation. We observed no significant cell death after 24 h of incubation (data not shown). After 48 h of incubation, an apparent dose-dependent cytotoxic response was triggered by the OMV/T, but only the value obtained with the highest dose (20 ␮g of OMV/T) was significantly different from untreated cells (Fig. 7A). Given the extended incubation time required and the general lack of significant effect, we conclude that the F. novicida OMV/T have minimal cytotoxicity to host cells.


We next examined the ability of the F. novicida OMV/T to stimulate proinflammatory responses of host cells. We incubated purified OMV/T with muBMDM for 24 h and measured the release of the cytokines TNF-␣, CXCL2 (MIP-2), and CCL2 (MCP-1) by enzyme-linked immunosorbent assay (ELISA). A dose-dependent increase for each of the cytokines was observed, with significantly increased release for the 1- and 10-␮g vesicle doses compared to buffer only or sham vesicle preparation controls (Fig. 7B). To examine the mechanism by which the F. novicida OMV/T trigger proinflammatory responses, we incubated the vesicles with proteinase K to digest surface-accessible proteins (as in Fig. 5A) prior to addition to muBMDM. Proteinase K treatment of intact OMV/T had no effect on release of the cytokines compared to untreated OMV/T (Fig. 7C), indicating that surfaceaccessible proteins are not responsible for triggering the host cell responses. A previous study demonstrated that OMV must be intact to cause a cytotoxic effect on host cells (25). Therefore, we next treated the F. novicida OMV/T at 80°C for 1 h to disrupt the vesicles, prior to the addition of 1 ␮g to the muBMDM. The disrupted OMV/T triggered 2- to 3-fold-lower levels of cytokine release from the host cells compared to untreated vesicles (Fig. 7C). Treatment of the heat-disrupted vesicles with proteinase K did not cause further changes in cytokine release compared to heattreated OMV/T alone (Fig. 7C). Thus, the OMV/T must be intact to trigger maximal proinflammatory responses from host cells. F. novicida produces tubes during infection of host cells. The OMV/T released into the surrounding medium by F. novicida would need to diffuse away from the bacteria to interact with host cells. In contrast, the tubes extending from the bacterial surface could mediate direct contact with host cells. To determine whether OM tubes are produced during infection of host cells, we examined F. novicida U112 during early stages of infection of muBMDM. The bacteria were grown in BHI medium to early log phase (OD600 ⫽ 0.4, ⬃2 h of growth), a time when few to no bacteria express tubes (ca. ⱕ2%; similar to Fig. 3A), and placed in suspension with muBMDM on ice. Samples were heated to 37°C to initiate phagocytosis, fixed after 5 min, and then processed for TEM imaging by thin sectioning (42). Tubes were seen extending from bacteria in close proximity to macrophages, as well as from those that had been taken up by phagocytosis (Fig. 8). In some cases, the tubes appeared to be initiating contact with the host cell plasma membrane (Fig. 8). Of the 142 bacteria observed in the EM images (n ⫽ 83), 50% expressed tubes. Moreover, 76% of the bacteria expressing tubes were associated with macrophages, with the remainder free in the surrounding medium. Given that only ⬃2% of the input bacteria expressed tubes, these data show that production of the OM tubes by F. novicida is stimulated by interaction with host cells, and suggest a role for the tubes in host cell contact and bacterial uptake. Of note is that addition of infection medium to the bacteria in the absence of host cells did not induce the production of tubes (data not shown), suggesting that a host cell-derived signal triggers tube production. Vaccination with F. novicida OMV/T generates a protective immune response. OMV have proved effective as vaccines for a number of bacterial pathogens (72–76), and Pierson et al. showed that vaccination of mice with OMV isolated from late-stage F. novicida cultures provided limited protection against subsequent bacterial challenge (32). We vaccinated mice by intranasal administration of 20 ␮g of purified OMV/T isolated from early-stationary-phase U112 cultures, or PBS as a control. We then challenged

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FIG 7 Effects of OMV/T on host cells. (A) The indicated amounts of OMV/T purified from early-stationary-phase U112 were incubated with muBMDM for 48 h, and cytotoxicity was quantified by measuring LDH release. (B) The indicated amounts of OMV/T or a sham vesicle preparation were incubated with muBMDM for 24 h, and the release of TNF-␣, CXCL2, and CCL2 was quantified by ELISA of conditioned media. (C) MuBMDM were incubated for 24 h with buffer only, 1 ␮g of OMV/T, proteinase K (PK) only, 1 ␮g of OMV/T treated with proteinase K, 1 ␮g of OMV/T heated to 80°C for 1 h, or 1 ␮g of OMV/T heated and then treated with proteinase K. Cytokine release was quantified as in panel B. Bars indicate the means ⫾ the SEM for three independent experiments (A and B) or a representative of two experiments (C). **, P ⬍ 0.01 for comparison with the buffer-only (0 ␮g of OMV/T) control.

the mice 6 weeks later with highly lethal doses of F. novicida by the intranasal route (620 or 960 CFU; the 50% lethal dose of U112 is ⬍10 CFU [32]). As shown in Fig. 9, the OMV/T-vaccinated mice had significantly increased survival compared to the control mice. All OMV/T-vaccinated mice infected at the lower challenge dose of 620 CFU (n ⫽ 3) survived the entire course of infection, whereas OMV/T-vaccinated mice challenged at the higher dose of 960 CFU (n ⫽ 4) exhibited significantly delayed time to death, with two mice surviving until day 17. This demonstrates that the F. novicida OMV/T are capable of eliciting a protective immune response in vivo. DISCUSSION

We report here the isolation and characterization of native OMV from F. novicida and show that F. novicida produces novel tubelike extensions of its OM and releases tube-shaped vesicles into the surrounding medium. Several pieces of evidence indicate roles for the OMV/T in the virulence of F. novicida: (i) the vesicles contain known antigens, secreted proteins, and virulence factors; (ii) the production of the tubes is stimulated during infection of macro-

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phages; and (iii) the OMV/T elicit proinflammatory responses from host cells. In support of a general role for OMV/T in the pathogenesis of tularemia, we have also observed tubes on the surface F. tularensis LVS (subsp. holarctica) (44), and the production and release of both tubes and OMV by the fully virulent Schu S4 strain (subsp. tularensis) (McCaig and Thanassi, unpublished). OMV appear to be produced by all Gram-negative bacteria as part of their normal growth. However, mechanisms by which OMV are produced, and whether and how they might be regulated, remain poorly defined. Our results provide strong evidence in support of regulated production of OMV and the specific packaging of vesicle-associated proteins. F. novicida produced low levels of OMV/T during exponential-phase growth, but production increased upon entry into stationary phase. Moreover, the content of the vesicles was dynamic, with different protein profiles for the exponential-phase and early-stationary-phase OMV/T, and nearly three times more protein was present in the latter. Similar amounts of OMV/T from both time points were analyzed by mass spectrometry, so the difference in numbers of proteins detected was not due to vesicle yield. The dynamic content of the F. novi-



FIG 8 Thin-section TEM images of F. novicida during infection of muBMDM. U112 bacteria grown in BHI broth to early log phase were used to infect muBMDM. (A) Bacteria inside a phagosome. (B and C) Bacteria interacting with the macrophage plasma membrane. The bacteria are marked by asterisks. (D to F) Higher magnification images of bacteria expressing OM tubes in panels A to C. The arrows point to tubes. Scale bars: A to C, 500 nm; D to F, 100 nm.

cida vesicles suggests that their function may change with growth state. The production of OMV/T by F. novicida was also highly responsive to growth medium. Production was stimulated in BHI, but repressed in other media. Hazlett et al. demonstrated that the protein expression profile of F. tularensis grown in BHI closely resembles that of organisms isolated from infected macrophages (77). Together with previous studies that observed vesicles associated with phagocytized Francisella (30, 31), this supports a functional role for OMV/T during infection. Importantly, we found that the production of OM tubes by F. novicida was induced by interaction with host cells. The images obtained from these experiments suggest that the tubes may initiate contact with macrophages to trigger bacterial uptake. If true, the tubes could be considered as a specialized form of OMV production, allowing directed contact with or delivery of virulence factors to host cells, rather than nonspecific release into the extracellular medium. The production of tube-shaped structures, termed nanotubes, by bacteria has recently been described (46, 47). These structures were produced on solid media and appeared to allow direct bridging between bacteria and the exchange of cytoplasmic molecules,

FIG 9 Vaccination of mice with OMV/T. Mice were inoculated intranasally with PBS as a control or OMV/T purified-form early-stationary-phase U112. The mice were challenged after 6 weeks with 620 (Lo; n ⫽ 3) or 960 (Hi; n ⫽ 4) CFU of F. novicida wild-type bacteria and monitored for survival for 21 days. *, P ⬍ 0.05.


as well as bridging between bacteria and eukaryotic cells (46, 47). In contrast, the F. novicida tubes are produced in liquid as well as on solid media and are formed by extensions of the OM and thus are continuous with the periplasm rather than the cytoplasm. The F. novicida tubes do not appear to bridge bacteria together and readily detach from the bacteria into the extracellular milieu. Therefore, the F. novicida tubes are distinct from the previously reported structures, and their presence suggests a novel mechanism by which Francisella can interact with its environment. A shared mechanism may be responsible for generating the OMV/T and some additional feature, potentially a tube-specific cargo molecule, likely imparts shape to the tubes. The heat-sensitive nature of the tubes suggests that this structuring molecule is a protein. Proteomics analysis of the purified OMV/T identified 292 vesicle-associated proteins, including prominent Francisella antigens, secreted proteins, and virulence factors. Seven proteins are secreted by the type IV pilus system in F. novicida (20), four of which we found to be associated with the OMV/T. These proteins might first be secreted outside the bacteria by the type IV pilus pathway and then associate with the surface of the vesicles. This would be similar to the secretion of heat-labile enterotoxin in enterotoxigenic E. coli (24, 49). Alternatively, the proteins might enter the lumen of the vesicles from the periplasm. Fsp53 was the most abundant protein in exponential-phase OMV/T but was ⬎16-fold decreased in the stationary-phase vesicles. This suggests a role for Fsp53 during the exponential growth of F. novicida. The function of Fsp53 is unknown; however, an F. novicida strain containing a deletion mutation of both Fsp53 and the homologous upstream gene FTN_1260 (which is also present in the exponential phase OMV/T; see Table S1 in the supplemental material) is attenuated for replication in macrophages and virulence in mice (N. P. Mohapatra and J. S. Gunn, personal communication). Proteins associated with the putative FPI T6SS were also detected in the OMV/T. A role for OMV in T6S has not been established, but the vesicles could mediate delivery of FPI proteins to host cells. The large number of secreted proteins and virulence factors present indicate that the OMV/T represent a viable secre-

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tion pathway in F. novicida and that the vesicles are likely to impact interactions with the host during infection. Pierson et al. previously isolated OMV from U112 cultures grown to late stationary phase in TS broth and identified 416 vesicle-associated proteins (32). Of these, 185 were also identified in the present study and include Francisella antigens (FopA, FopB, and LpnA), secreted proteins (KatG, SucC, SucD, Ahp1, GroEL, PepO, BglX, ChiA, and Fsp53), FPI proteins (IglB, IglC, and PdpD), and additional known and hypothetical virulence factors (FipB, FTN_0714, FTN_0340, FTN_0429, and FTN_0643). However, the relative abundance of the proteins differs markedly between the two studies, and more than half of the proteins identified by Pierson et al. were not present in our OMV/T preparations. Notably, GroEL was the most abundant protein in the OMV from the prior study (32), representing ⬎20% of the sample, whereas this protein was absent from our exponential-phase vesicles and present at only 0.07% of spectra in our early-stationary-phase vesicles. These differences likely reflect the different time points and growth media used for vesicle isolation, as well as the fact that the OMV in previous study were not purified using density gradient flotation (32). In addition, Pierson et al. did not report the presence of tube-shaped vesicles in their OMV preparations. We did not attempt to replicate the conditions of the previous study to explore the basis for this difference; however, since production of the tubes is responsive to both growth phase and growth medium, we hypothesize that the absence of tubes was due to differences in these parameters between the two studies. A range of activities has been associated with OMV, including delivery of virulence factors to host cells and modulation of innate and adaptive immunity (24). We found that the F. novicida OMV/T had only minor cytotoxicity toward muBMDM. However, the OMV/T elicited a robust, dose-dependent release of proinflammatory cytokines. These results suggest that the proinflammatory, but not the cytotoxic, effects of the OMV/T may be physiologically relevant during infection. Although Pierson et al. did not examine the ability of their OMV to stimulate proinflammatory responses of host cells, these researchers did detect hemolytic and acid phosphatase activities (32). In contrast to our results, Pierson et al. reported significant cytotoxicity of their OMV to the J774A.1 murine macrophage-like cell line (32). This disparity in results may be due to several factors, including differences in the composition of the vesicles, the use of primary cells in our study versus a cell line by Pierson et al., and the use of a much higher vesicle dose (4.3 mg) in the previous study (32). Innate immune responses to Francisella are primarily mediated by TLR2, with recognition of lipoproteins being a major part of this response (78). The F. novicida OMV/T are enriched in lipoproteins, including the known TLR2 agonists LpnA and FipB (78), and delivery of the lipoproteins to host cells may underlie the inflammatory activity of the vesicles. Although the immunostimulatory component of lipoproteins is heat resistant (79), disruption of the OMV/T by heat treatment resulted in a significant decrease in proinflammatory activity, indicating that the vesicles must be intact for greatest potency and arguing against nonspecific activation of the BMDM due to components released from the vesicles or from contaminating molecules. An alternative explanation is that a heat-sensitive factor such as a protein may be responsible for the proinflammatory activity of the OMV/T. A role for the immunomodulatory activity of the F. novicida OMV/T during infection remains to be determined; however, a recent study demonstrated

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that a TLR2-dependent inflammatory response conferred by membrane vesicles contributes to the virulence of mycobacteria (71). A number of studies have shown that Francisella spp. actively interfere with intracellular signaling pathways and innate immune responses of host cells (7, 9, 15–17). In addition, fully virulent strains of F. tularensis are capable of inducing a broad immunosuppression during early stages of infection (18). In the context of these studies, it will be important to determine whether vesicles released by fully virulent strains behave similarly toward host cells as the F. novicida OMV/T or whether fully virulent bacteria modulate their vesicles to avoid activating host responses. The potential for OMV to serve as protective vaccines has been demonstrated for diverse bacteria (72–76). In addition, Pierson et al. showed that vaccination with their OMV provided limited protection against subsequent F. novicida infection (32). We show here that mice vaccinated intranasally with a single dose of purified OMV/T were afforded significant protection against subsequent challenge with highly lethal doses of F. novicida. The stronger protective immune response observed in our study compared to that obtained by Pierson et al. may be due to differences in experimental procedures, differences in the composition of the vesicles as noted above, and the presence of OM tubes in addition to OMV in our samples. There currently exists no licensed vaccine for tularemia and a vaccine based on OMV is attractive since it would be safe (no live bacteria), the LPS of Francisella has low toxicity, and known antigens are present in the OMV in their native membrane context. Thus, the method for OMV/T isolation detailed here may provide the foundation for the generation of a safe and effective subunit vaccine for tularemia. ACKNOWLEDGMENTS We thank Susan Van Horn of the Central Microscopy Imaging Center at Stony Brook University for assistance with electron microscopy. We thank Barbara Mann (University of Virginia) for providing the anti-FipB antibody and for critical reading of the manuscript. We thank Jorge Benach and Anne Savitt (Stony Brook University) for providing the antiFopA antibody. We thank Nrusingh Mohapatra and John Gunn (The Ohio State University) for sharing unpublished data. We thank Martha Furie (Stony Brook University) for helpful suggestions and critical reading of the manuscript. This study was supported by Public Health Service grant AI055621 from the National Institute of Allergy and Infectious Diseases and by Targeted Research Opportunity and Seed Grant awards from Stony Brook University.

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