Murine Innate Immune Response to Virulent Toxigenic and Nontoxigenic Bacillus anthracis Strains

Abstract

Effective treatment of anthrax is hampered by our limited understanding of the pathophysiology of Bacillus anthracis infection. We used a genetically complete (pXO1⁺ pXO2⁺) virulent B. anthracis strain and four isogenic toxin-null mutants to determine the effects of the anthrax edema toxin (ET; edema factor [EF] plus protective antigen [PA]) and lethal toxin (LT; lethal factor [LF] plus PA) on the host innate response during systemic infection. Using the spleen as an indicator for host response, we found that intravenous inoculation of LT-deficient mutants into C57BL/6 mice significantly increased production of several cytokines over that observed after infection with the parent strain or an EF-deficient mutant. Bacteria producing one or both of the toxins were capable of inducing significant apoptosis of cells present in spleens, whereas apoptosis was greatly reduced in mice infected with nontoxigenic mutants. Mice infected with toxin-producing strains also showed increased splenic neutrophil recruitment compared to mice infected with nontoxigenic strains and neutrophil depletion prior to infection with toxin-producing strains, leading to decreased levels of apoptosis. Together, these studies indicate that anthrax LT suppresses cytokine secretion during infection, but both EF and LF play roles in inducing neutrophil recruitment and enhancing apoptosis. Interestingly, in the absence of LF the effect of EF-induced cell recruitment is further enhanced, perhaps because LF so effectively suppresses the secretion of chemokines.

B. anthracis vaccine studies support an important role for the toxins in disease progression, but the pathophysiological roles of the Bacillus anthracis lethal toxin (LT) and edema toxin (ET) during systemic anthrax infection have not been defined in the context of host-pathogen interactions (7, 35, 39). Research to date has focused on the effects of purified toxin components on the murine host response or has been done using attenuated, noncapsulated B. anthracis strains lacking the virulence plasmid pXO2. This is the first study to use a genetically complete (pXO1⁺ pXO2⁺), virulent, capsulated B. anthracis strain and isogenic toxin gene mutants to specifically assess the effects of the individual toxin components on the murine host response during a systemic lethal anthrax infection.

B. anthracis secretes three toxin proteins—edema factor (EF), lethal factor (LF), and protective antigen (PA)—that act in binary combinations to form two AB-type toxins, the edema toxin (ET = PA+EF) and the lethal toxin (LT = PA+LF). PA, the “B” component, binds to at least two different receptors present on a variety of host cells (6, 38). Bound PA is proteolytically cleaved by furin, which results in release of a 20-kDa protein fragment and heptamerization of 63-kDa fragments to form a prepore (29). The oligomerized form of PA exposes three binding sites for LF and/or EF. The PA-EF-LF complex is taken up via clatherin-dependent endocytosis. Subsequently, a decrease in pH leads to a conformational change and the formation of a pore, thereby permitting the translocation of EF and LF into the host cell (4).

EF is a calmodulin-dependent adenylyl cyclase that alters water homeostasis by causing elevated levels of cyclic AMP (27). Tissue edema has been identified in some cases of anthrax-infected individuals (1), and intravenous injection of ET into rats leads to excessive accumulation of fluid (3), a finding similar to what is observed in cutaneous anthrax. In vitro, ET blocks the phagocytic activity of human polymorphonuclear neutrophils (43) and macrophages (10) and can modulate cytokine production by monocytes (22) and murine macrophage-like cell lines (10). Recent work has demonstrated that ET and LT may work synergistically to impair cytokine secretion from murine dendritic cells (40).

LF is a zinc-metalloprotease that cleaves the N terminus of mitogen-activated protein (MAP) kinase kinases 1, 2, 3, 4, 6, and 7 (MEK1, -2, -3, -4, -6, and -7) (12, 33, 41, 42) and is cytotoxic for certain cell types, including some types of macrophages and macrophage cell lines (17, 31). MAP kinase pathways are involved in a number of cellular functions, including cell growth and proliferation, differentiation, and apoptosis (26). By means of a series of phosphorylations, the MAP kinase pathway relays signals to the nucleus that results in changes in gene expression. In a variety of cell types, LT-induced disruption of the MAP kinase pathway resulted in increased propensity to undergo apoptosis (17, 23, 24, 31). However, the mechanism of LT cytotoxicity is not completely understood since there does not appear to be a direct link between MAP kinase cleavage and susceptibility to LT-induced cytolysis.

The effect of LT on the host inflammatory cytokine response during B. anthracis infection is currently under debate. Hanna et al. (17) found that sublytic concentrations of LT induced interleukin-1β (IL-1β) and tumor necrosis factor alpha (TNF-α) expression in macrophages. These authors subsequently proposed the idea that macrophages are the primary effectors of cell death during a systemic anthrax infection via the release of cytokines and reactive oxygen intermediates, leading to a massive inflammatory response (18). In support of this hypothesis, Pickering and Merkel found that A/J mice infected with Sterne strain (pXO1⁺ pXO2⁻) spores had elevated levels of IL-6 and gamma interferon (IFN-γ) in the sera of some of the mice. Furthermore, the levels of TNF-α, IL-6, IL-12, IL-10, IFN-γ, and granulocyte-macrophage colony-stimulating factor (GM-CSF) in A/J mice infected with Sterne and then treated with lipopolysaccharide (LPS) did not differ significantly from uninfected LPS-treated control mice, suggesting that LT does not function to suppress the inflammatory response in vivo. Pickering and Merkel (34) also demonstrated that the addition of B. anthracis Sterne strain spores did not prevent LPS-induced cytokine production from a macrophage cell line J774.A1, further buttressing this hypothesis. In sharp contrast to these findings, other researchers have found that LT functions to inhibit cytokine production during infection (14, 33). These groups demonstrated that LT exposure inhibits LPS-induced cytokine and NO production from macrophages. LT was also found to impair murine dendritic cells in their ability to prime T cells and mount an effective immune response (2). Most recently, it was demonstrated that both LT and ET are able to suppress human T-cell activation and proliferation in vitro (30) and murine T-cell activation in vivo (9). These data support the idea that the toxins function to inhibit an effective host immune response by disabling multiple branches of the immune system, allowing the bacteria to replicate to high numbers and remain undetected.

To examine and clarify the roles of the toxins during a lethal B. anthracis infection, we analyzed the host response to infection with our genetically complete, virulent strain UT500 and isogenic toxin mutants with specific toxin genes deleted. We mimicked the systemic phase of the disease by administering vegetative bacilli intravenously, taking advantage of the fact that wild-type and toxin-deficient B. anthracis replicate to similar numbers in the murine host. The synchronicity of infection and similar mean times of death allowed us to directly compare the host response between mice infected with parent or toxin mutant B. anthracis strains. Cytokine profiling experiments were implemented to assess the effect of the toxins on activating or suppressing the immune response in vivo. We also performed experiments to measure the levels of splenic cell apoptosis in parent and toxin mutant B. anthracis-infected mice. Finally, we compared the numbers and types of cells present in spleens of parent and nontoxigenic B. anthracis-infected mice using cell surface staining and differential cell staining techniques to further elucidate the roles of the toxins during infection. We demonstrate that mice infected with the parent B. anthracis strain have vastly different host responses than mice infected with nontoxigenic mutants and clarify previous findings concerning the role of B. anthracis toxins during infection.

MATERIALS AND METHODS

Bacterial strains.

UT500 (pXO1⁺ pXO2⁺) was constructed as described previously (5). Briefly, pXO2 from Pasteur strain 6602 (pXO1⁻ pXO2⁺) was transduced into Sterne strain 7702 (pXO1⁺ pXO2⁻) using CP51 (15) to create UT500. UT500 is toxigenic and capsulated (5). UT539 (Δlef [Kan^r]), UT540 (Δcya [Kan^r]), UT541 (Δlef Δcya [Kan^r Spc^r]), and NM1 [ΔpagA(R) (Kan^r)] are all isogenic derivatives of UT500. The construction of UT539, UT540, UT541, and NM1 was described previously (21).

Preparation of B. anthracis vegetative cells for intravenous (i.v.) infections.

B. anthracis spores were streaked onto nutrient broth yeast agar containing 0.8% bicarbonate (NBY-CO₃) and antibiotics when appropriate (kanamycin [50 μg/ml] and/or spectinomycin [100 μg/ml]). The plates were incubated in an atmosphere of 5% CO₂ at 37°C for approximately 24 h to ensure the maintenance of pXO2. A few colonies from each plate were used to inoculate 15 ml of Luria broth containing 0.5% glycerol (LB-goh) and antibiotics (when appropriate). Cultures were shaken at 200 rpm for approximately 12 h at 30°C in air. Cells were subcultured in LB-goh (no antibiotics, initial optical density at 600 nm [OD₆₀₀] of 0.1) at 37°C to an OD₆₀₀ of approximately 0.4. At this OD₆₀₀, all cultures contained ∼10⁷ CFU/ml and were found to have less than 5% heat-resistant CFU (spores). Cultures were then centrifuged for 3 min at 16,000 × g, the medium was removed, and the cells were washed with Dulbecco phosphate-buffered saline (DPBS; pH 7.2; catalog no. 20012-050; Invitrogen, Carlsbad, CA) using a volume equivalent to the initial culture volume. The cells were washed two more times before they were resuspended in a volume equivalent to the original culture volume. Prior to infection, an aliquot of washed cells was diluted and plated on blood agar plates to obtain the final CFU/ml used in the infection (inoculating dose).

Mouse infection.

All mouse protocols were approved by the University of New Mexico Institutional Animal Care and Use Committee. Female 7- to 8-week-old C57BL/6 mice were purchased from Harlan Sprague-Dawley (Madison, WI) and maintained as described previously (28). Mice were infected intravenously using a 30-gauge needle via the tail vein with approximately 100 μl of 10⁷ CFU/ml or ∼10⁶ B. anthracis CFU. For assessing CFU over time, spleens were harvested at 4, 8, and 11 h postinfection and subjected to bead beating for 90 s prior to dilution in DPBS and plating on sheep blood agar plates.

Cytokine profiling.

Approximately 11 h postinfection, mice were euthanized with 1 ml of Avertin (1.24% 2,2,2-tribromoethanol, 0.78% tert-amyl alcohol) administered intraperitoneally. At this time, spleens were harvested in 1 ml of DPBS containing protease inhibitor cocktail (DPBS/MPI; catalog no. P8340; Sigma-Aldrich, St. Louis, MO), homogenized, and then subjected to three snap-freeze cycles. The samples were then centrifuged at 7,000 × g, and the supernatant was filtered through a Whatman Puradisc 25-mm 0.45 μM syringe column (catalog no. 6751-2504; Fisher Scientific, Hampton, NH). Cytokine levels present in infected and control mice were measured by using Bio-Plex mouse cytokine 18-Plex (catalog no. 171-F11181) according to the protocol of the manufacturer (Bio-Rad, Hercules, CA). Undiluted and samples diluted 1/10 in DPBS/MPI were assayed in duplicate. Similar cytokine profiling results were observed when mice were i.v-infected with the 1,000 CFU that reproducibly gives 100% death, and the tissues were harvested 20 h postinfection. The IL-12p70 cytokine levels shown were measured by using the Beadlyte mouse multicytokine detection system (catalog no. 48-004; Upstate, Charlottesville, VI) and the protocol suggested by the manufacturer.

Preparation of spleen cells for flow cytometry experiments.

Spleens were removed from infected mice approximately 11 h postinfection and were ground between the ends of two frosted slides in 10% heat-inactivated fetal bovine serum (FBS)-RPMI. The cells were then transferred to a 15-ml conical tube, and Hanks buffered saline solution was added to a final volume of 15 ml. Cells were centrifuged at 300 × g for 10 min at 4°C. Supernatant was removed, and 2 ml of red blood cell lysis buffer (0.826% NH₄Cl, 0.1% KHCO₃, and 0.0037% disodium EDTA in double-distilled H₂O [pH 7.3]) was then added to the cells for 1 to 2 min. Hanks balanced salt solution was then added to 15-ml final volume, and the cells were centrifuged again at 300 × g for 10 min at 4°C. Supernatant was poured off, and the cells were resuspended in 2 ml of staining buffer (DPBS containing 2% heat-inactivated FBS and 40 μg of EDTA/ml). The cells were counted on a hemacytometer for total cell enumeration, and cell viability was assessed by using trypan blue exclusion.

Cell surface staining and cell profiling.

Spleen cells were first incubated with purified anti-CD16/CD32 (2.4G2, rat immunoglobulin G2b [IgG2b]) to block both FcR-mediated binding of staining antibody and cell aggregation caused by FcγR binding of antibody-stained cells by FcγR⁺ cells in the populations. Subsequently, cells were stained with combinations of the following monoclonal antibodies (MAb) in three separate reaction tubes: (i) to identify T-cell subsets, anti-CD3e-PE (145-2C11, Ar Hamster IgG1), anti-CD8-PerCP (53-6.7, rat IgG2a), and anti-CD4-APC (RM4-5, rat IgG2a) were used; (ii) to identify macrophages and neutrophils, anti-Ly-6G-fluorescein isothiocyanate (1A8, rat IgG2a), anti-Ly-6G and Ly-6C (Gr-1)-biotin (RB6-8C5, rat IgG2b), and anti-F4/80-APC (BM8, rat IgG2a; Caltag Laboratories) were used; and (iii) to identify B cells, anti-CD19-APC (1D3, rat IgG2a) was used. The following isotype controls were used in the appropriate combinations: rat IgG2a (R35-95), rat IgG2b (A95-1), or Ar hamster IgG1 (A19-3). All antibodies were purchased from BD Biosciences unless otherwise noted. Cells were stained at 4°C, followed by two washes with DPBS containing 2% FBS and 40 μg of EDTA per ml. Cells that used the biotinylated antibody Gr-1 were subsequently incubated with SA-PerCP for 30 min at 4°C, followed by two washes with DPBS containing 2% FBS and 40 μg of EDTA/ml. Cells were fixed with 1% paraformaldehyde in DPBS (wt/vol). The data acquisition was performed on a BD Biosciences FACSCalibur (San Jose, CA), and analysis was performed with WinList software (Verity Software House, Topsham, ME). After acquiring the percentages of each cell type in the total cell population, we used these percentages along with the total cell numbers as obtained from the hemacytometer to determine the numbers of each cell type present.

Apoptosis TUNEL assay.

Spleens were prepared as described above for flow cytometry experiments. The APO-BRDU TUNEL (terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling) apoptosis kit (Phoenix Flow Systems, Inc., San Diego, CA) was used for TUNEL analysis by flow cytometry; the procedures suggested by the manufacturer were followed.

Neutrophil depletion in mice.

Mice were injected intraperitoneally with 100 μg of RB6-8C5 antibody (8) at 24 h prior to infection with B. anthracis. Control mice were injected with 100 μg of IgG2b isotype control SFR3-DR5 antibody (36) 24 h prior to infection. Prior to infection, neutrophil depletion was assessed by randomly selecting two mice from each antibody treatment group for differential staining of splenic cell preparations. The average neutrophil depletion was ca. 95%.

Statistical analysis.

The data shown for cytokines, trypan blue cell viability analysis, cell surface profiling, and apoptosis experiments represent an average of three independent experiments (± the standard error of the mean) with three to five mice per B. anthracis infection group. Error was determined by using one-way analysis of variance with Dunnett's post-hoc test to determine significance relative to PBS mock-infected control mice (see Fig. 2 and 3) or UT500-infected mice (see Fig. 4). The data shown for neutrophil depletion experiments are an average of two independent experiments (± the standard error of the mean) with three to five mice per group.

FIG. 2.

Cytokine or chemokine levels (in pg/ml) present in splenic lysates of B. anthracis-infected C57BL/6 mice at 11 h postinfection. *, P < 0.05; **, P < 0.01 (relative to PBS-mock-infected control mice).

FIG. 3.

Numbers of viable spleen cells as assessed by trypan blue staining. *, P < 0.01 (relative to PBS-mock-infected control mice).

FIG. 4.

Numbers of cells as assessed by immunofluorescence labeling and flow cytometry of cells isolated from spleens of B. anthracis-infected C57BL/6 mice at 11 h postinfection. The dashed line indicates the numbers of cells in PBS-mock-infected control mice. *, P < 0.05; **, P < 0.01 (relative to UT500-infected mice).

RESULTS

Parent and toxin-deficient strains of B. anthracis grow at similar rates in the spleens and lungs of C57BL/6 mice.

We showed previously that parent B. anthracis strain UT500 and isogenic toxin mutants caused similar mean times to death (MTD) with the same 50% lethal doses (LD₅₀) in BALB/c mice by using intratracheal and i.v. methods of infection (21). In the present study, we compared the murine host systemic response after i.v. inoculation with parent or toxin mutant B. anthracis strains in C57BL/6 mice. In C57BL/6 mice, the i.v. LD₅₀ for infection with UT500, UT540 (EF⁻), UT539 (LF⁻), and NM1 (PA⁻) were all approximately 50 to 100 CFU per mouse. Intravenous inoculation with ∼10³ CFU/mouse of all five strains resulted in an MTD of approximately 18 to 24 h when the bacterial burden was approximately 10⁷ to 10⁸ bacteria in the lungs and spleens just prior to death (data not shown).

For cytokine profiling, apoptosis, and cell profiling experiments, mice were inoculated i.v. with a dose of 10⁶ CFU of vegetative bacilli/mouse. At this dose, all mice had similar numbers of CFU in their lungs and spleens at 4, 8, and 11 h, and all mice died approximately 11 h to 12 h postinfection with bacterial burdens of 10⁷ to 10⁸ (Fig. 1). This burden was similar to that observed at 18 to 24 h postinfection in mice infected with a lower dose (10³ CFU/mouse) (data not shown). The narrow MTD observed in mice infected with the 10⁶ vegetative bacillus dose provided the optimal condition for comparing the host response to infection with either the UT500 or the toxin-deficient B. anthracis strains that is not possible when mice are infected intratracheally where the MTD is between 2 and 3 days.

FIG. 1.

Growth of UT500 (▪) or NM1 (PA⁻) (□) in the lungs and spleens of C57BL/6 mice after i.v. inoculation of 10⁶ vegetative bacteria per mouse. Each bar represents the average CFU of five mice (± the standard deviation). The data shown represent one of three independent experiments. Strains UT539 (LF⁻), UT540 (EF⁻), and UT541 (EF⁻ LF⁻) display similar growth patterns (data not shown).

The splenic cytokine response in B. anthracis lethal toxin mutant-infected C57BL/6 mice is amplified.

We sought to determine the effects of B. anthracis lethal and edema toxins on the host cytokine response during a systemic anthrax infection. A broad panel of cytokines (IL-2, IL-3, IL-4, IL-5, IL-6, IL-10, IL-12p70, IL-17, IL-1α, IL-1β, IFN-γ, TNF-α, macrophage inflammatory protein 1α [MIP-1α], G-CSF, GM-CSF, KC, and RANTES) produced by lymphocytes and phagocytes were measured by using multicytokine Luminex detection analysis. Cytokine profiles of splenic lysates in mice infected with parent or isogenic toxin mutants were determined 11 h postinfection when the mice were clinically ill.

Three major patterns of cytokine profiles in B. anthracis-infected mice were noted. The most discerning pattern was observed in mice infected with any LT-deficient strain (UT539, UT541, or NM1). These mice had similar elevated levels of several inflammatory cytokines or neutrophil chemokines (IL-6, IL-12p70, IL-1α, IL-1β, G-CSF, and KC) compared to mice infected with the parent or EF⁻ (UT540) strain that had significantly lower levels of these cytokines (Fig. 2A to F). For example, IL-6 was present at concentrations 223- to 361-fold higher in LT⁻ strain-infected mice than in UT500- or UT540 (EF⁻)-infected mice (Fig. 2A). These cytokines are mainly produced by phagocytic cell types such as macrophages and monocytes, suggesting that LT suppresses the secretion of inflammatory mediators from these cells during a systemic anthrax infection.

The second cytokine pattern was observed in mice infected with nontoxigenic B. anthracis strains (UT541 and NM1). These mice had high cytokine levels relative to mice infected with any toxin-producing strain (Fig. 2G to H). TNF-α and MIP-1α levels were 20- to 27-fold higher in UT541 and NM1-infected mice relative to that observed in toxigenic B. anthracis-infected mice. Notably, ET and LT may act synergistically in limiting MIP-1α and also IL-1α production in the murine host. EF⁻ (UT540) and LF⁻ strain (UT539)-infected mice had modest elevated levels of both IL-1α and MIP-1α, while mice infected with nontoxigenic strains (UT541 or NM1) had much higher levels of both IL-1α and MIP-1α compared to all B. anthracis-infected mice.

Finally, for IL-10 we observed a unique cytokine pattern. IL-10 levels were elevated in UT539 (LF-)-infected mice (11-fold) relative to parent strain-infected mice (Fig. 2I). A small increase in IL-10 was detected in some nontoxigenic (UT541 and NM1) B. anthracis-infected mice, but overall the IL-10 levels were not significantly different from UT500-infected mice (P > 0.05). No significant increases in the cytokines or chemokines IL-2, IL-3, IL-4, IL-5, IL-17, IFN-γ, GM-CSF, or RANTES were observed in any of the B. anthracis-infected mice (data not shown). Many of these cytokines are produced by lymphocytes, suggesting that LT may not have a significant effect on the secretion of cytokines from lymphocytes during this stage of infection.

C57BL/6 mice infected with toxin-producing B. anthracis strains demonstrate altered splenic cell numbers and neutrophilic infiltrates.

The dissimilar cytokine profiles of mice infected with parent versus toxin mutant B. anthracis strains suggested that different populations of immune cells might be recruited and/or survive within the spleens of these mice during a fatal systemic anthrax infection. Therefore, we evaluated the spleens of infected mice for total cell numbers and performed cell phenotyping to identify the types and numbers of immune cells present. Host cell viability of B. anthracis parent-infected mice relative to PBS mock-infected control mice revealed that B. anthracis-infected mice had significantly fewer viable splenic cells, ca. 25% less (Fig. 3). In contrast, mice infected with the toxin-deficient B. anthracis strains did not show significantly different numbers of total viable spleen cells from PBS-inoculated control mice (Fig. 3).

We used flow cytometry for enumeration of CD19⁺ B cells, F4/80⁺ macrophages, Gr1⁺ Ly-6G⁺ granulocytes, CD3⁺ CD4⁺ T cells, and CD3⁺ CD8⁺ T cells present in the spleens of PBS mock-infected control mice compared to B. anthracis parent- and toxin-deficient-infected mice (Fig. 4; PBS mock-infected control mice are represented by the horizontal dashed line). Relative to the PBS mock-infected control mice, UT500-infected mice had fewer CD19⁺ B cells and CD3⁺ CD4⁺ and CD3⁺ CD8⁺ T cells (Fig. 4). Although more Gr1⁺ Ly-6G⁺ granulocytes were detected in UT500-infected mice compared to the PBS control mice, similar numbers of F4/80⁺ macrophages were detected in UT500-infected mice and the healthy PBS mock-infected control mice (Fig. 4).

A comparison of the types of cells present in mice infected with toxin-deficient strans versus parent B. anthracis strains revealed that UT539 (LF⁻)-infected mice had the highest number of granulocytes present in their spleens, more than twice that observed in UT500-infected mice (Fig. 4A) and more than four times that observed in PBS mock-infected control mice. We confirmed the presence of neutrophils in spleens of B. anthracis-infected mice by using differential staining. The pattern of neutrophil influx in B. anthracis-infected mice closely corresponded with the pattern of granulocytes observed with flow cytometry (data not shown). UT541 (EF⁻ LF⁻)- and NM1 (PA⁻)-infected mice had numbers of CD19⁺ B cells similar to those for PBS control mice, which was greater than the number of CD19⁺ B cells found in the spleens of UT500-infected mice (Fig. 4B). Quantitation of macrophages (F4/80⁺ cells) (Fig. 4C) showed that only mice infected with UT540 (EF⁻) had significantly higher numbers of macrophages relative to UT500-infected mice (P < 0.05). Similar to the pattern observed for macrophages, CD3⁺ CD4⁺ and CD3⁺ CD8⁺ T cells were found to be elevated in UT540 (EF⁻)-infected mice compared to UT500-infected mice (Fig. 4D and E) (P < 0.01). In addition, the number of CD3⁺ CD8⁺ T cells was also greater in UT539 (LF⁻)-infected mice than in wild-type-infected mice (P < 0.05).

Apoptosis of splenocytes is increased in parental and single toxin mutant B. anthracis-infected C57BL/6 mice.

LT causes activation-induced apoptosis of certain cell lines in vitro. Here, we used a TUNEL flow cytometry assay to assess the apoptotic cells in the spleens of infected mice. Mice infected with UT500 had elevated levels of apoptosis (32.3%) compared to PBS control mock-infected mice (3.1%) (P < 0.01). Mice infected with UT539 (LT⁻) demonstrated even higher levels of apoptosis (46.5%). UT540 (EF⁻)-infected mice had significantly increased levels of apoptosis (16.2%) compared to PBS mock-infected control mice (P < 0.05) but less than parent or UT539 (LT⁻) mutant-infected mice. The double toxin mutants, UT541 and NM1, had the lowest levels of apoptosis (6.9 and 6.8%, respectively; P > 0.05 relative to PBS mock-infected control mice).

We hypothesized that the increased apoptosis found in spleens of UT500-, UT539 (LF⁻)-, and UT540 (EF⁻)-infected mice may be due to the apoptosis of recruited neutrophils because increased neutrophil influx was found in mice infected with these B. anthracis strains. Thus, we depleted neutrophils by the administration of a neutrophil-depleting MAb (RB6-8C5) (8) or an equivalent amount of an isotype control MAb (SFR3-DR5) (36) 24 h prior to infection with UT500, UT539 (LF⁻), or UT540 (EF⁻) bacilli. We harvested the spleens from infected mice at 9 h postinfection and measured the number of apoptotic cells. At this time point, the neutrophil-depleted mice appeared moribund and, therefore, they were harvested 2 h earlier than in previous apoptosis experiments. Although the total levels of apoptotic cells were lower than previously found at 11 h postinfection, the results indicate that neutrophil-depleted mice, infected with UT500 or UT539 (LF⁻) had significantly lower percentages of apoptotic cells relative to the non-neutrophil-depleted isotype control mice (Fig. 5).

FIG. 5.

Percentage of apoptotic cells in spleens of neutrophil-depleted B. anthracis-infected mice as detected by using a TUNEL assay. ND, neutrophil depleted; IC, isotype control. *, P < 0.01.

DISCUSSION

We utilized a virulent pXO1⁺ pXO2⁺ B. anthracis parent strain and isogenic toxin gene mutants to specifically assess the effects of the secreted toxins on the host immune system using splenic cells from infected mice as an indicator. We demonstrated three major findings. (i) LT suppresses the host innate immune response during systemic anthrax infection, as evidenced by the high levels of proinflammatory cytokines and chemokines in mice infected with UT539 (LF⁻), UT541 (EF⁻ LF⁻), or NM1 (PA⁻) compared to mice infected with strains that express LT (UT500 and UT540 [EF⁻]). (ii) The presence of LT and/or ET induces neutrophilic infiltrates into the spleen. (iii) Both ET and LT induce apoptosis, and the presence of EF alone induces the greatest amount of apoptosis.

Previously conflicting conclusions regarding the effects of LT and ET on the cytokine response of the host may be due to several factors. First, a large majority of the research was performed using purified toxin components. The actual levels of toxins present in tissues during an infection are unknown, making it difficult to accurately mimic the amount of toxin present during infection. Second, attenuated pXO1⁺ pXO2⁻ B. anthracis strains were commonly used in these studies. The capsule is essential in the murine model for virulence (11) and might modulate how the host cells interact with and respond to the toxins. Finally, many investigations were performed in vitro using a single cell type, e.g., macrophages or dendritic cells. In vitro experiments cannot accurately reflect the in vivo microenvironments that are present when different types of cells are interacting in whole tissues during infection, and therefore the same cells may respond differently to B. anthracis in vivo.

Earlier findings of in vitro studies implicated ET in modifying the mammalian cell cytokine response (22, 30, 40). In the present study, we found that UT540 (EF⁻)-infected mice did not have significantly different levels of inflammatory cytokines from UT500-infected mice. The presence of the capsule or other virulence factors acting together with ET may mask the host cytokine response to ET since we used virulent pXO1⁺ pXO2⁺ strains in contrast to previous studies that used Sterne strain derivatives (pXO1⁺ pXO2⁻) (40) or purified toxin components (22, 30). ET may cooperate with LT to modify the host's cytokine response, as has been previously suggested (40). Very high levels of TNF-α and MIP-1α were only observed in mice infected with mutants lacking both toxins (UT541 or NM1). It is possible that the reduced levels of TNF-α and MIP-1α present in UT539 (LF⁻)-infected mice were due to the high levels of IL-10 present in these mice. IL-10 is a potent suppressor of macrophage functions, including cytokine secretion (16). In fact, IL-10 negatively regulates LPS-induced TNF-α production but not IL-6 production via activation of Bcl-3 in macrophages (25). This is in accordance with our findings because UT539 (LF⁻)-infected mice displayed high levels of IL-10 and IL-6 and reduced levels of TNF-α.

LT has been shown to cleave MAP kinases that are essential for cytokine gene expression (12, 33, 41, 42). Accordingly, mice infected with LT⁻ strains showed high levels of IL-1α, IL-1β, IL-6, KC, and G-CSF compared to mice infected with B. anthracis strains capable of producing LT during infection. The increased levels of these splenic cytokines observed in LT⁻ B. anthracis-infected mice were not evident until the mice were moribund. Sera collected from infected mice at earlier time points did not demonstrate an increase (data not shown), suggesting that the toxins play a key role in suppressing the phagocyte cytokine response late during infection once the toxins have accumulated in tissues to a significant concentration.

Cell surface profiling studies by flow cytometry and differential staining of splenic cell cytospin preparations revealed decreased numbers of splenic B cells in mice infected with bacilli expressing ET, LT, or both. There was also a statistically significant decrease (∼25%) in both CD3⁺ CD4⁺ and CD3⁺ CD8⁺ T cells. Although the biological relevance of a 25% reduction is difficult to predict, together with the reduction in the number of B cells, all major arms of acquired immune response were negatively affected, making the significance more likely.

We also examined the effect of LT and ET on the apoptosis of splenic cells during systemic anthrax infection. In mice infected with nontoxigenic B. anthracis strains (UT541 or NM1), only low levels of apoptosis were detected relative to parent strain-infected mice even though the mice were moribund and appeared to be as sick as UT500-infected mice. The high percentage of apoptotic cells detected in UT539 (LF⁻)-infected mice (46.5%) was initially surprising because previous in vitro studies demonstrated a role for LT in sensitizing cells to LPS/TLR4-induced apoptosis (24, 31, 32). Significantly, however, cell surface phenotyping and differential staining of cells present in spleens of these mice revealed elevated numbers of neutrophils, a finding consistent with previous findings that demonstrate that LT hinders neutrophil migration (13). In the present study we demonstrated that a significant number of apoptotic splenic cells of UT500- and UT539 (LF⁻)-infected mice were neutrophils, in agreement with previous studies demonstrating that activated neutrophils recruited to tissues during infection undergo programmed cell death in order to limit and resolve the inflammatory response (19, 20, 37). Ultimately, it is still uncertain whether the toxins actively induced apoptosis of cells in vivo or whether the elevated levels of apoptotic cells detected during B. anthracis infection was simply due to the increased recruitment of neutrophils during terminal systemic anthrax infection when one or both toxins were present. Future work will focus on distinguishing between these two processes.

In summary, this study sheds important information on the B. anthracis secreted toxins and their individual impact during the infectious process. Although wild-type and toxin-deficient B. anthracis strains caused lethality in mice with similar MTD and LD₅₀ values, the mice clearly responded differently to infection with wild-type B. anthracis compared to infection with nontoxigenic B. anthracis strains. We acknowledge that some of the alterations in the host response in toxin mutant-infected mice may have been due to interactions of the toxins with other virulence factors (32). Overall, the murine model is an important animal model to study the effects of the B. anthracis toxins in vivo due to the tools available for cell and cytokine analysis. It is now necessary to investigate whether other common B. anthracis experimental models, e.g., rabbit and nonhuman primate models, will yield similar or disparate results.