Francisella tularensis-Infected Macrophages Release Prostaglandin E2 that Blocks T Cell Proliferation and Promotes a Th2-Like Response

Matthew D. Woolard, Justin E. Wilson, Lucinda L. Hensley, Leigh A. Jania, Thomas H. Kawula, James R. Drake and Jeffrey A. Frelinger
J Immunol February 15, 2007, 178 (4) 2065-2074; DOI: https://doi.org/10.4049/jimmunol.178.4.2065

Abstract

Francisella tularensis is a highly infectious bacterial pathogen, and is likely to have evolved strategies to evade and subvert the host immune response. In this study, we show that F. tularensis infection of macrophages alters T cell responses in vitro, by blocking T cell proliferation and promoting a Th2-like response. We demonstrate that a soluble mediator is responsible for this effect and identify it as PGE2. Supernatants from F. tularensis-infected macrophages inhibited IL-2 secretion from both MHC class I and MHC class II-restricted T cell hybridomas, as well as enhanced a Th2-like response by inducing increased production of IL-5. Furthermore, the soluble mediator blocked proliferation of naive MHC class I-restricted T cells when stimulated with cognate tetramer. Indomethacin treatment partially restored T cell proliferation and lowered IL-5 production to wild-type levels. Macrophages produced PGE2 when infected with F. tularensis, and treatment of infected macrophages with indomethacin, a cyclooxygenase-1/cyclooxygenase-2 inhibitor, blocked PGE2 production. To further demonstrate that PGE2 was responsible for skewing of T cell responses, we infected macrophages from membrane PGE synthase 1 knockout mice (mPGES1−/−) that cannot produce PGE2. Supernatants from F. tularensis-infected membrane PGE synthase 1−/− macrophages did not inhibit T cell proliferation. Furthermore, treatment of T cells with PGE2 recreated the effects seen with infected supernatant. From these data, we conclude that F. tularensis can alter host T cell responses by causing macrophages to produce PGE2. This study defines a previously unknown mechanism used by F. tularensis to modulate adaptive immunity.

Francisella tularensis is a facultative intracellular coccobacillus and the causative agent of tularemia. Because F. tularensis persists in the environment and has been produced as a bioweapon (1), interest in F. tularensis biology has risen. In mice, F. tularensis causes a fulminant infection that is similar to human tularemia (2). Cell-mediated responses are important in controlling and resolving F. tularensis infection in mice (3, 4, 5). Initial survival during infection is dependent on IFN-γ with NK cells being the likely source of this cytokine at early stages of the disease (6). IFN-γ is required in F. tularensis-infected mice for macrophage activation. Macrophages from IFN-γ-deficient mice fail to prevent intracellular growth of F. tularensis resulting in rapid death (7). Mice that lack adaptive immune responses are able to initially control bacterial infection; however, they are unable to resolve the infection and eventually die (8). T cell-mediated immune responses confer protection and immunity against F. tularensis, whereas B cell Ab responses seem to play a minimal role in protection, although a non-Ab B cell-mediated role has been reported (9). CD4+ T cells respond to exogenous proteins and CD8+ T cells respond against endogenous or cross-presented proteins. Both CD4+, MHC class II-restricted, and CD8+, MHC class I-restricted, T cells are independently capable of controlling F. tularensis, because mice depleted of either population are still able to clear infection (4). Interestingly, CD4−CD8−CD3+ T cells are also able to control intracellular F. tularensis growth in vitro (3), the MHC class restriction of these T cells is not known. Thus, a variety of T cells contribute to controlling F. tularensis infection. Any modulation or dampening of T cell responses would promote a selective advantage to F. tularensis in its ability to infect and replicate within the host.

PGs are a family of powerful short-lived lipid mediators of immune responses. PGs have been demonstrated to have both inhibitory and stimulatory effects on immune cell function (10, 11). PGE2 interacts with a variety of immune cells, leading to increased cAMP levels that alter activation (12, 13, 14). The production of PGE2 is a complex pathway (as reviewed in Ref. 15). The cyclooxygenase (COX)4-1/COX-2 enzymes convert arachidonic acid into PGH2, an unstable intermediate, PGES then converts PGH2 to PGE2. The COX-1 enzyme is constitutively expressed, whereas COX-2 is inducible. There are three PGE synthases membrane PGE synthase 1 (mPGES1), mPGES2, cPGES. Although all can convert PGH2 into PGE2, only mPGES1 has been shown to be responsible for induction of PGE2 from macrophages (16, 17). The ability of PGE2 to block T cell proliferation is mediated through the E prostanoid receptor 4 (18). Engagement of this receptor leads to decreased calcium flux, an early stage of T cell activation, and blocks the downstream production of IL-2 and IFN-γ, while augmenting production of IL-4 and IL-5 (18, 19, 20). IL-2 is an important mediator in T cell proliferation. Blocking of IFN-γ production directs the remaining T cells toward a Th2 phenotype. Although the effects of PG on the immune system are understood, their role in infectious disease is not. Recent studies have clearly demonstrated that the bacterial products LPS, peptidoglycan, and CpG DNA can initiate COX gene up-regulation and PG secretion (21, 22, 23, 24, 25). Thus, bacterial infections that can induce PGE2 are able to inhibit T cell function, and should gain an advantage in in vivo growth.

Intracellular bacteria use a myriad of mechanisms to evade immune responses. The mechanism behind this evasion is poorly understood and is likely specific to each bacterium. F. tularensis, Listeria monocytogenes, and Salmonella enterica all alter or escape the macrophage phagolysosomes to evade destruction (26, 27). Some intracellular bacteria down-regulate MHC class I or MHC class II surface expression (28, 29, 30, 31). Mycobacterium tuberculosis skews the cytokine response from a pro- to an anti-inflammatory response (32, 33). Legionella pneumophila causes IFN-γ-stimulated macrophages to produce PGE2, and blocks IFN-γ secretion from CD4+ T cells (34). S. enterica can also stimulate macrophages to secrete PGE2 (35). Thus, intracellular bacteria use multiple mechanisms to modulate and evade the adaptive immune response.

In this study, we identified one mechanism that F. tularensis uses to modulate T cell responses. Infection of macrophages by F. tularensis alters their ability to appropriately activate Ag-specific T cells. Furthermore, infected cells secrete a soluble product that can mediate this effect. We have shown the soluble factor to be PGE2, which leads to decreased T cell proliferation. Furthermore, PGE2 can skew cytokine production by T cells away from IL-2 and toward IL-5. This is the first such study to examine a direct effect of F. tularensis infection on subsequent T cell function.

Materials and Methods

Cell lines

The murine macrophage cell line ANA-1 (36) (provided by Dr. J. Frelinger, University of Rochester Medical Center, Rochester, NY) was cultured in RPMI 1640, supplemented with 5% FCS, 2 mM l-glutamine, 100 U/ml penicillin, 100 μg/ml streptomycin, and 50 μM 2-ME. Before infection, cells were washed extensively with antibiotic free-supplemented RPMI 1640.

The OVA-specific B3Z T cell hybridoma (37) (provided by N. Shastri, University of California, Berkeley, CA), which recognizes the OVA peptide SIINFEKL bound to Kb, was cultured in RPMI 1640, supplemented with 5% FCS, 2 mM l-glutamine, 100 U/ml penicillin, 100 μg/ml streptomycin, 50 μM 2-ME, and 50 μg/ml hygromycin.

RAW.Ak macrophages (provided by Dr. C. Harding, Case Western University, Cleveland, OH) were produced by transfection of the RAW246.7 murine macrophage cell line with mouse IAk MHC class II gene (38). RAW.Ak cells were cultured in RPMI 1640 supplemented with 10% heat-inactivated FBS, 1 mM sodium pyruvate, 10 mM HEPES buffer, 50 μM 2-ME, 50 U/ml penicillin, and 50 μg/ml streptomycin.

The IAk-HEL46–61-specific murine T cell hybridoma, h4Ly50.5 (provided by Dr. W. Wade, Dartmouth Medical School, Lebanon, NH), was cultured in DMEM supplemented with 10% heat-inactivated FBS, 1 mM sodium pyruvate, 2 mM l-glutamine, and 50 μM 2-ME.

Bacteria

The F. tularensis live vaccine strain (LVS) (29684; American Type Culture Collection (4) strain was used in these studies. Viable bacteria were quantified by serial dilution on chocolate agar.

Mice

B6.D2 TgN(tcr-Lcmv)327Sdz/Fre (P14) were bred at the University of North Carolina as described previously (39). C57BL/6 (B6) mice were purchased from either The Jackson Laboratory or Taconic Laboratories. The B6;129-mpges1tm1/Kol (mPGES1−/−), backcrossed to B6 for six generations (40), were provided by Dr. B. H. Koller (University of North Carolina at Chapel Hill, NC). All animals used in this study were maintained under specific pathogen-free conditions in the American Association of Laboratory Animal Care-accredited University of North Carolina Department of Laboratory Animal Medicine Facilities. B10.BR/SgSnJ (B10.BR) mice were purchased from The Jackson Laboratory and housed in the Animal Resource Facility at Albany Medical College under specific pathogen-free conditions.

Peptides

The gp33 peptide (KAVYNFATC), HY peptide (KCSRNRQYL), and OVA peptide (SIINFEKL) were synthesized by the University of North Carolina microchemical facility, purified by HPLC, and tested for purity by mass spectroscopy.

Tetramers

The P14 CD8+ T cell cognate tetramer Db-KAVYNFATC and irrelevant tetramer Db-KCSRNRQYL tetramer were produced as described previously (39, 41). All batches were routinely tested for LPS contamination.

Proliferation

Naive CD8+ T cells were purified from spleen of P14 transgenic mice by negative paramagnetic cell separation (Miltenyi Biotec) as described previously (41). We routinely achieved a 90% pure CD8+ T cell population as determined by flow cytometry. Standard [3H]thymidine incorporation was used to measure proliferation. In tetramer proliferation studies, purified P14 CD8+ T cells were cultured at a concentration of 2.5 × 104 cells/well of a 96-well plate and incubated with complete medium (RPMI 1640 with 5% FCS) and an equal volume of either mock-infected or infected supernatants for 2 h before the addition of various concentrations of tetramer. For PGE2 treatment, purified P14 CD8+ T cells were cultured at a concentration of 2.5 × 104 cells/well of a 96-well plate and incubated with complete medium (RPMI 1640 with 5% FCS) and an equal volume of either mock-infected or infected supernatants, or 15 ng/ml PGE2 (Cayman Chemical), or 15 ng/ml Iloprost (Sigma-Aldrich) along with various concentrations of tetramers. In peptide proliferation assays, P14 CD8+ T cells were purified. The remaining spleen cells (CD8−) were irradiated at 16 Gy emitted from a Gammacell 40 Cesium-137 source (Atomic Energy of Canada Limited, Ottawa, Canada) and used as APCs. CD8+ T cells and APCs were then plated in a 1:1 ratio at 2 × 105 cells/well in equal volumes of complete medium and either mock-infected or infected supernatants for 2 h before the addition of various concentrations of peptide.

Ab stimulation of T cell hybridoma

Flat-bottom 96-well plates were coated with 10 μg/ml hamster anti-mouse CD3ε (eBioscience; catalog no. 14-0031-86) for 15 h at 4°C. Following washing with PBS, 105 h4Ly50.5 T cells were added to the wells along with 10 μg/ml soluble anti-mouse CD28 (eBioscience) and cleared F. tularensis infection medium, mock-infected medium, or PGE2 (Cayman Chemical) for 24 h at 37°C. Following incubation, supernatants were screened for IL-2 and IL-5 using a mouse Th1/Th2 cytokine and cytometric bead array according to the manufacturer’s instructions (BD Biosciences; catalog no. 16-0281-86).

lacZ assays

After activation, cells were lysed and total lacZ activity in individual cultures was measured using chromogenic substrates as described previously (37). Briefly, individual cultures in 96-well plates were washed once with 100 μl of PBS, and lysed by addition of 100/μl Z buffer (100 mM 2-ME, 9 mM MgCl2, and 0.125% Nonidet P-40 in PBS) containing 0.15 mM CPRG (chlorophenol red/3-galactoside; Calbiochem). After 4-h incubation at 37°C, 50 μl of stop buffer (300 mM glycine and 15 mM EDTA in water) was added to each well, and absorption/fluorescence of each well was read using a 96-well plate reader. Absorption wavelength used for CPRG was 570 nm with 650 nm as the reference wavelength.

Bone marrow-derived macrophage generation

Bone marrow cells were flushed from B10.BR or B6 mouse femurs and incubated for 7 days on nontissue culture-treated 15-cm2 dishes with L cell-conditioned medium containing GM-CSF. Following differentiation, nonadherent cells were removed by multiple washes with PBS and bone marrow-derived macrophages were removed from plates by incubation with 1 mM EDTA in PBS.

Ag presentation assay

RAW.Ak or bone marrow-derived macrophages were pretreated where indicated for 24 h with 100 U/ml murine IFN-γ. After washing and suspension in cell culture medium, macrophages were placed into wells of a 96-well plate at a concentration of 2 × 105/well. Following 2-h incubation at 37°C to allow adherence to the plate, the macrophages were cocultured with the indicated concentration of filter-sterilized hen egg lysozyme (HEL) and 105 h4Ly50.5 T cells for 24 h at 37°C. Supernatants were removed and screened for IL-2 and IL-5 using a mouse Th1/Th2 cytokine kit and cytometric bead array according to the manufacturer’s instructions (BD Biosciences; catalog no. 551287).

Infection of cells and preparation of conditioned medium

Spleens were aseptically harvested from mice and a single-cell suspension was generated. RBC were removed by ACK lysis. Cells were plated at a concentration of 3.5 × 106 splenocytes/ml in antibiotic-free complete medium. Either splenocytes or macrophages were infected with F. tularensis at various multiplicity of infections (MOI) where indicated. Two hours after infection, extracellular F. tularensis was killed by the addition of 50 μg/ml gentamicin and the cultures were incubated for 45 min. Supernatant was removed and cells were washed with PBS, fresh antibiotic-free complete medium was added, and cells were incubated for a further 24 h. Supernatant was then collected and spun at 300 × g for 10 min to remove eukaryotic cells. Supernatant was then heat treated at 60°C for 10 min or UV inactivated to kill any extracellular F. tularensis. Supernatant was plated onto chocolate agar after heat treatment to ensure complete killing of F. tularensis. Supernatant was then stored at −80°C until needed. To inhibit COX-1/COX-2 activity, cells were pretreated with either ethanol (solvent control) or 10 μM indomethacin (Sigma-Aldrich) before infection. Indomethacin and ethanol were maintained in cultures throughout infection except in washes.

Flow cytometry

Cells were stained with either directly conjugated PE-labeled anti-Vα2 TCR or 2.5 nM of PE-labeled Dbgp33 tetramer and then analyzed by flow cytometry using FACSCalibur (BD Biosciences).

Enzyme immunoassay

PGE2 was measured using a commercial PGE2 enzyme immunoassay kit (Assay Design) according to the manufacturer’s instructions.

Statistical analysis

Data were analyzed using Student’s t test for differences in lacZ assays, proliferation assays, PGE2 levels, and CFUs. GraphPad Prism 4.03 (GraphPad) was used for analysis. Statistical significance is indicated as p ≤ 0.05, p ≤ 0.01, p ≤ 0.001.

Results

F. tularensis alters T cell activation through a soluble inhibitory factor

During generation of F. tularensis-specific T cell hybridomas for epitope discovery, we observed that F. tularensis-infected APCs were unable to stimulate F. tularensis-specific T cell hybridomas (data not shown). Furthermore, F. tularensis infection of APCs presenting soluble F. tularensis Ag resulted in a similar response by an Ag-specific T cell hybridoma. We hypothesized that F. tularensis is able to modulate T cell hybridoma activation. To better investigate this observation, we used epitope-defined MHC class I and MHC class II T cell hybridomas. We infected splenocytes with various MOI of F. tularensis. We then added the B3Z T cell hybridoma and cognate peptide. After 24 h we examined lacZ activity by the addition of CPRG. Addition of F. tularensis inhibited the production of lacZ of the B3Z T cell hybridoma at a MOI as low as one bacterium to four host cells (Fig. 1⇓A). These data demonstrates that F. tularensis infection of APCs can alter activation of MHC class I-restricted T cell hybridoma.

FIGURE 1.
FIGURE 1.

F. tularensis infection of APCs alters T cell activation. A, Unmanipulated B6 splenocytes were infected with F. tularensis at various MOI. B3Z T cell hybridomas and 50 ng/ml SIINFEKL were added, and 24 h after infection, T cell activation was determined by the addition of CPRG and OD of colored product was determined at 570 nM. ∗, Statistical difference (∗, p ≤ 0.05; ∗∗, p ≤ 0.01; ∗∗∗, p ≤ 0.001) from uninfected cells. B, RAW246.7 macrophage cell line was pretreated with IFN-γ to up-regulate MHC class II. Macrophages were infected at a MOI of 100:1 along with h4Ly50.5 HEL-specific T cell hybridoma, and various amounts of HEL was added. Twenty-four hours after infection, supernatants were analyzed for IL-2 and IL-5. ∗, Statistical difference (∗∗, p ≤ 0.01; ∗∗∗, p ≤ 0.001) from control levels. C and D, Supernatant from infected APCs were collected. Eukaryotic cells were removed by centrifugation, and extracellular F. tularensis was inactivated. Supernatant was diluted with fresh medium and added to naive splenocytes, B3Z T cell hybridoma, and 50 ng/ml SIINFEKL (C) or bone marrow-derived macrophages, h4Ly50.5, and HEL (D). Twenty-four hours after stimulation, lacZ activity (C) or IL-2 (D, left panel), or IL-5 (D, right panel) was determined. Representative experiments from three separate experiments are shown. Error bars represent intra-assay SEM of triplicates. ∗, Statistical difference (∗, p ≤ 0.05; ∗∗, p ≤ 0.01; ∗∗∗, p ≤ 0.001) from control.

We also tested the ability of F. tularensis infection to modulate activation of MHC class II-restricted T cell hybrids. Activation of h4Ly50.5 was determined by analyzing supernatants for IL-2 and IL-5. We infected RAW.Ak at a MOI of 100:1 and added the h4Ly50.5 and various concentrations of HEL. After 24 h IL-2 and IL-5 levels were determined in the supernatants. As seen in MHC class I-restricted T cells, there is a marked decrease in IL-2 secretion by the h4Ly50.5 T cell hybrid. We also observed increased production of IL-5 (Fig. 1⇑B). IL-5 is produced during Th2 responses and considered to be an anti-inflammatory cytokine (42, 43). These experiments demonstrate that F. tularensis alters T cell responses, both blocking IL-2 production and promoting a Th-2-like response.

We wanted to determine whether a soluble inhibitory factor was responsible for altering T cell hybridoma activation. To examine this hypothesis, we isolated supernatant from mock-infected or infected splenocytes or bone marrow-derived macrophages (MOI 100:1) as described previously. After 24 h, supernatant was collected and any F. tularensis LVS that had escaped from the APCs were inactivated with either UV treatment or heat treatment (100°C for 30 min). The serially diluted supernatant was added to uninfected splenocytes with the B3Z T cell hybridoma or bone marrow-derived macrophages and the h4Ly50.5 T cell hybridomas and appropriate cognate peptide. Twenty-four hours later, T cell hybridoma response was determined. Supernatant from infected splenocytes was able to block T cell hybridoma IL-2 secretion in both MHC class I- and MHC class II-restricted T cells (Fig. 1⇑, C and D). Infected supernatant also increased IL-5 production from the h4Ly50.5 T cell hybridoma (Fig. 1⇑D). Supernatants from infected RAW.AK cells had the same effect as infected bone marrow-derived macrophages (data not shown). The 100°C treatment should eliminate most proteins, especially cytokines, and would suggest that a very stable protein, lipid, or sugar structure is responsible for modulation of T cell hybridoma activation. The data demonstrate that F. tularensis infection of APCs leads to production of a soluble factor that can alter T cell activation.

Soluble inhibitory factor can directly inhibit T cell proliferation

We hypothesized that the soluble inhibitory factor either targets the APC and interferes with Ag presentation or directly affects the T cell. To examine this theory, we used an APC-free system of naive T cell activation. Soluble MHC class I tetramers can directly stimulate CD8+ T cells in vitro to proliferate (39). We used purified P14 transgenic CD8+ T cells that recognize the lymphocytic choriomeningitis virus gp33 epitope KAVYNFATC (44). Purified CD8+ T cells were stimulated with various concentrations of MHC class I tetramer-bearing cognate peptide (gp33 Db) in the presence of supernatant’s from mock-infected or infected splenocytes. [3H]Thymidine incorporation was used to determine proliferation. Incubation of T cells in infected supernatant led to a 10-fold shift in the dose response compared with mock-infected supernatant (Fig. 2⇓A). To determine whether the soluble inhibitory factor could make the T cells refractory to later stimulation, T cells were preincubated in mock-infected or infected supernatant for 2 h. The supernatant was removed and fresh medium along with various concentrations of cognate tetramer was added. Proliferation was again determined by [3H]thymidine incorporation. As described before, T cells incubated with infected supernatant were almost completely inhibited in their ability to proliferate compared with T cells incubated mock-infected supernatant (Fig. 2⇓B). Taken together, these experiments suggest that the soluble factor is interacting directly with the T cell to block proliferation and this treatment of the T cells makes them refractory to later stimulation.

FIGURE 2.
FIGURE 2.

Inhibitory factor exerts its action on T cells. Purified naive CD8+ T cells from P14 transgenic mice (A) were treated with infected or control supernatant along with various concentrations of tetramer or (B) pretreated for 2 h with supernatant that was washed away, and fresh medium was added along with various concentrations of cognate tetramer. Proliferation was determined by the addition of [3H]thymidine. ∗, Statistical difference (∗, p ≤ 0.05; ∗∗∗, p ≤ 0.001) from controls. C, Purified naive CD8+ T cells from P14 transgenic mice were treated or pretreated with infected or control supernatant; cells were then stained with either PE-Db gp33 tetramer or with anti-Vα2 TCR and anti-CD8. Cells were analyzed by flow cytometry and mean fluorescence intensity (MFI) was determined. There was no significant difference between control and infected supernatants in binding of cognate tetramer nor in TCR expression levels. D, Purified naive CD8+ T cells from P14 transgenic mice were treated with infected or control supernatant, and cells were then loaded with Indo-1AM (Invitrogen Life Technologies) and stimulated with 10 μg of tetramer. Break represents time for addition of tetramer. Calcium flux was then monitored by flow cytometry. Representative experiments from two separate experiments are shown. Error bars shown in A and B represent intra-assay SEM generated from triplicate samples.

Inhibition of T cell proliferation may be due to decreased expression of TCR. To determine whether the number of TCR molecules was down-regulated, we examined expression of Vα2. Binding of an anti-Vα2 Ab was unchanged (Fig. 2⇑C). It was also possible that the factor interfered with MHC-TCR interaction. To determine whether the soluble inhibitory factor was interfering with TCR binding to MHC peptide complex, we examined the ability of cognate MHC class I tetramer to bind to the P14 transgenic T cells. Purified CD8+ T cells were incubated or preincubated in mock-infected or infected supernatant and stained with PE-labeled MHC class I tetramer. The soluble inhibitory factor did not interfere with the ability of T cells to bind MHC tetramer (Fig. 2⇑C).

Because the binding of TCR and TCR-MHC are unaffected, it is likely that effects of the soluble factor are downstream of TCR engagement. Calcium mobilization is an early step in TCR signal transduction. We examined calcium mobilization of mock-infected and infected supernatant treated P14 transgenic T cells that were stimulated with cognate tetramer. Treatment of T cells with infected supernatant both decreased the amount of calcium mobilized after tetramer stimulation and slightly slowed the kinetics (Fig. 2⇑D), suggesting differential signaling. These data demonstrate that the soluble inhibitory factor is acting directly upon the T cell to block proliferation.

Indomethacin treatment partially reverses F. tularensis ability to alter T cell hybridoma cytokine secretion

The heat stability of the inhibitory factor suggests a small, stable protein, lipid or sugar. COX-2 is the inducible enzyme that is the important in the pathway leading to the production of many PGs. F. tularensis up-regulates COX-2 mRNA levels in human macrophages and dendritic cells (T. H. Kawula, unpublished data). Furthermore, PGs are well-known mediators of immune responses (18, 19, 20). Thus, we suggest that the mechanism that mediates F. tularensis modulation of T cells is the production of PGs by infected cells that then alters T cell activation. To test this theory, we infected bone marrow-derived macrophages with F. tularensis that were either untreated, treated with indomethacin in ethanol or the vehicle ethanol alone. Twenty-four hours after infection, supernatants were collected. Purified P14 CD8+ T cells were incubated in the supernatants along with cognate peptide and irradiated APCs. T cell proliferation was determined by [3H]thymidine incorporation. Treatment of APCs with indomethacin partially reversed T cell inhibition of proliferation (Fig. 3⇓A). To examine whether PGs were also affecting the MHC II-restricted T cells, we infected indomethacin-treated bone marrow-derived macrophages and added h4Ly50.5 T cell hybridomas and HEL protein. Twenty-four hours later, supernatants were tested for levels of IL-2 and IL-5. As seen in the P14 CD8+ T cells, indomethacin treatment partially restored IL-2 levels back to control levels (Fig. 3⇓B). However, indomethacin treatment reduced IL-5 to control levels (Fig. 3⇓C). These data taken together suggest that PGs are at least partially responsible for inhibiting T cell proliferation and completely responsible for increased production of IL-5 by the T cell hybridoma.

FIGURE 3.
FIGURE 3.

Indomethacin treatment partially restores proliferation and completely reverses elevated IL-5 secretion. Bone marrow-derived macrophages were untreated, treated with indomethacin or vehicle. Macrophages were then infected or mock infected. A, Twenty-four hours after infection supernatant was collected. Supernatants were incubated with naive splenocytes and naive CD8+ T cells from a P14 transgenic mouse. Activation was then determined by [3H]thymidine incorporation. B and C, h4Ly50.5 HEL-specific T cell hybridoma cells and HEL was added to infected bone marrow-derived macrophages. Twenty-four hours later supernatants were tested for cytokine levels. Representative results from three separate experiments are shown. Error bars represent intra-assay SE generated from triplicate samples. ∗, Statistical difference (∗, p ≤ 0.05; ∗∗∗, p ≤ 0.001) from untreated infected bone marrow-derived macrophages.

F. tularensis infection causes PGE2 secretion

We have demonstrated that PGs are partially responsible for the skewing of T cell responses. Although several PGs have been demonstrated to influence immune responses, PGE2 has been the best characterized. Furthermore, PGE2 is a potent mediator of the immune response and has been demonstrated to inhibit IL-2 production by T cells and induce the secretion of IL-5, similar to the results seen with the T cell hybridomas during F. tularensis infection (18, 19, 20). We hypothesized that F. tularensis infection of bone marrow-derived macrophages will lead to the secretion of PGE2. To test this, we infected bone marrow-derived macrophages with F. tularensis at a MOI of 100:1. Twenty-four hours after infection, supernatants were tested for the presence of PGE2 by commercial ELISA. F. tularensis infection of bone marrow-derived macrophages leads to a significant increase in PGE2 secretion (Fig. 4⇓A). IFN-γ has been demonstrated to directly induce the production of PGE2 from peritoneal macrophages (45). As such, we wanted to determine whether IFN-γ was responsible for the release of PGE2. To examine this, we treated bone marrow-derived macrophages with 100 U/ml IFN-γ or were mock treated. Macrophages were infected or mock infected, and the levels of PGE2 were monitored after 24 h. PGE2 production by F. tularensis-infected bone-marrow-derived macrophages does not require IFN-γ treatment because untreated macrophages produced significant amounts of PGE2 (Fig. 4⇓B). To ensure that this increase represented newly synthesized PGE2, we treated the same cells with indomethacin before F. tularensis infection. Indomethacin treatment reduced PGE2 to control levels (Fig. 4⇓). These data demonstrate that F. tularensis induces PGE2 synthesis in macrophages.

FIGURE 4.
FIGURE 4.

F. tularensis infection induces PGE2 secretion from bone marrow-derived macrophages. A, Bone marrow-derived macrophages were either untreated, mock-treated (ethanol), or treated with indomethacin 2 h before infection. Macrophages were either mock infected or infected with F. tularensis at a MOI of 100:1. Twenty-four hours after infection supernatants were tested for PGE2. B, Bone marrow-derived macrophages were either untreated or treated with 100 U/ml IFN-γ 24 h before infection. Cells were then infected or mock infected. Twenty-four hours after infection supernatants were tested for PGE2 concentrations. Representative results from three separate experiments are shown. Error bars represents intra-assay SEM generated from triplicate samples. ∗, Statistical difference (∗∗, p ≤ 0.01; ∗∗∗, p ≤ 0.001) from control untreated.

PG enhances intracellular growth of F. tularensis LVS

Infectious intracellular organisms elicit PG production for multiple reasons. PGs can block T cell proliferation and skew the T cell response (14, 18, 46); it can also aid the infectious organism in intracellular replication (35, 47). We predicted that PGs might be beneficial to the ability of F. tularensis to invade and replicate inside infected cells. To address this theory, we examined intracellular growth of F. tularensis within indomethacin-treated macrophages as compared with controls. A B6-derived macrophage cell line, ANA-1, was treated with indomethacin or vehicle for 2 h before infection. Macrophages were than infected at a MOI of 100:1 and CFUs were determined at day 0, day 1, and day 2 postinfection. F. tularensis LVS readily replicated within untreated macrophages, because initial infection was 5 × 105 organisms and by day 2 there was almost a five log increase in the number of organisms (Fig. 5⇓A). In indomethacin-treated macrophages, the same number of organisms were able to invade as in untreated macrophages, suggesting that PGs are not needed for F. tularensis to invade macrophages. However, intracellular replication was severely restricted, because by day 2 there was only a two-log increase in the number of organisms. There was almost a 10,000-fold increase in the number of organisms from untreated macrophages as compared with indomethacin-treated macrophages. Indomethacin or PGE2 may be directly affecting F. tularensis replication. To examine this, we added indomethacin or PGE2 to broth cultures of F. tularensis and monitored growth. Neither indomethacin nor PGE2 had any effect on the organism’s ability to grow in broth culture (data not shown). The changes in replication between indomethacin-treated and untreated macrophages are due to the effects of PGs on the macrophage.

FIGURE 5.
FIGURE 5.

Blocking PGE2 synthesis by indomethacin treatment slows F. tularensis growth inside macrophages. A, ANA-1 macrophages were treated with indomethacin or vehicle 2 h before infection. Macrophages were then infected at a MOI of 100:1 with F. tularensis. At 0, 24, and 48 h postinfection CFUs were determined. ∗, Statistical difference (∗∗, p ≤ 0.01) from untreated control. B, Bone marrow-derived macrophages were either mock infected or infected at a MOI of 100:1 with untreated, ethanol fixed, UV heat-treated, or live F. tularensis. PGE2 levels were determined 24 h after infection. ∗, Statistical difference (p ≤ 0.05) from control. C, Bone marrow-derived macrophages were infected with either untreated or UV-treated F. tularensis. Twenty-four hours after infection, supernatants were tested for the ability to alter h4Ly50.5 T cell hybridoma cytokine secretion. D, Supernatants from mock-infected, UV-inactivated F. tularensis infected, or F. tularensis bone marrow-derived macrophages were incubated with purified naive CD8+ P14 transgenic T cell-irradiated APCs and cognate peptide. Proliferation was determined by [3H]thymidine incorporation. ∗, Statistical difference (∗∗, p ≤ 0.01) from control. Representative experiments from three separate experiments. Error bars represent intra-assay SEM generated from triplicate samples.

Indomethacin treatment partially restored T cell function and inhibited the ability of F. tularensis to replicate within host cells. Therefore, indomethacin’s ability to reverse F. tularensis-mediated T cell-altered activation may be due to limiting F. tularensis replication, rather than due to blocking the effects of PGs on the T cells. We hypothesized that F. tularensis did not need to replicate to induce PGE2 secretion. To test this, we treated F. tularensis to block replication. We inactivated F. tularensis with heat (60°C for 10 min), UV treatment, formaldehyde, or ethanol, and then added the organisms to bone marrow-derived macrophages. As expected, all treatments blocked replication as determined by growth on chocolate agar (data not shown). After 24 h PGE2 levels were determined. UV and formaldehyde-treated F. tularensis stimulated PGE2 production from the bone marrow-derived macrophages, whereas ethanol and heat treatment abolished F. tularensis’s ability to elicit PGE2 (Fig. 5⇑B). The fact that ethanol and heat-killing of bacteria ablated PGE2 induction from macrophages, whereas UV and formaldehyde did not, suggests that a protein is responsible for PGE2 production from infected macrophages. Ethanol and heat-killing will degrade or modify protein structure, whereas UV and formaldehyde treatment will likely conserve protein structure (48). However, future studies will be needed to better understand this process.

We next tested whether UV-treated F. tularensis could still block IL-2 production and increase IL-5 secretion. We isolated supernatant from bone marrow-derived macrophages that were incubated with UV-treated F. tularensis. This supernatant was then tested for its ability to inhibit IL-2 and augment IL-5 production from the h4Ly50.5 T cell hybridoma. As Fig. 5⇑C demonstrates, F. tularensis replication was not needed to alter T cell cytokine secretion. Similarly, supernatant from bone marrow-derived macrophages incubated with UV-treated bacteria also inhibits P14 C8+ T cell proliferation (Fig. 5⇑D). These data demonstrate that, although indomethacin inhibited the ability of F. tularensis to replicate in host cells, replication of F. tularensis was not necessary for either production of PGE2 or alteration of T cell responses induced by F. tularensis. Taken together, this would suggest that PGE2 secretion by F. tularensis-infected cells can suppress T cells, and the effect can be reversed by the addition of indomethacin.

Supernatants from mPGES1−/− bone marrow-derived macrophages infected with F. tularensis cannot inhibit T cell proliferation

We demonstrated that inhibition of PG production partially reversed F. tularensis’s ability to alter T cell responses. Of the known PGs, only PGE2 has been demonstrated to have the immunomodulatory properties that we have observed (11, 18, 19, 20, 49). We decided to use genetic ablation to further examine the possible role of PGE2 in T cell modulation during F. tularensis infection of bone marrow-derived macrophages. We tested mice that had the mPGES1 gene inactivated by insertional mutagenesis (40). As mentioned earlier, the mPGES1 and COX-2 are responsible for inducible PGE2 production (40, 50). We hypothesized that F. tularensis infection of bone marrow-derived macrophages from the mPGES knockout mice would not block T cell proliferation. Bone marrow-derived macrophages from the mPGES1−/− and B6 (wild-type control) were infected with F. tularensis LVS at a MOI of 100:1. After 24 h, supernatants were transferred to uninfected irradiated APCs and CD8+ naive P14 transgenic T cells and cognate peptide, and proliferation was examined. Supernatant from mPGES1−/−-infected bone marrow-derived macrophages was unable to block T cell proliferation, unlike wild-type (Fig. 6⇓). This demonstrates that PGE2 is responsible for the skewing of T cell activation during F. tularensis infection of APCs.

FIGURE 6.
FIGURE 6.

PGE2 is responsible for blocking of T cell proliferation and altering the response from IL-2 to IL-5. A, Bone marrow-derived macrophages from wild-type (B6) and mPGES1−/− mice were infected with F. tularensis at a MOI of 100:1. Twenty-four hours after infection, supernatants were collected and incubated with purified, naive, CD8+ P14 T cells and irradiated APCs along with cognate peptide. Proliferation was determined by [3H]thymidine incorporation. B, Purified naive, P14 CD8+ T cells were either mock treated or treated with 15 ng/ml PGE2 or Iloprost (prostacyclin analog). T cells were then stimulated with 5 nM cognate tetramer. Proliferation was determined by [3H]thymidine incorporation. C, h4Ly50.5 HEL T cell hybridomas were either mock treated or treated with 10 ng/ml PGE2 and then stimulated with anti-CD3/CD28 Abs. Twenty-four hours after stimulation supernatants were analyzed for cytokine concentrations. Representative experiments from three separate experiments. Error bars represent intra-assay SEM generated from triplicate samples. ∗, Statistical difference (∗, p ≤ 0.05; ∗∗, p ≤ 0.01; ∗∗∗, p ≤ 0.001) from control.

It is possible that the infected macrophages are making PGE2, which in an autocrine fashion, acts on the macrophage to make another factor that is skewing T cell responses. To determine whether this was true or whether the PGE2 was directly acting on the T cells, we examined the effect of PGE2 on T cells stimulated in the absence of APCs. Purified naive CD8+ P14 transgenic T cells were mock treated or treated with 15 ng/ml commercial PGE2, this concentration is similar to what is seen from F. tularensis-infected bone marrow-derived macrophages, or 15 ng/ml Iloprsot (a prostacylcin analog). Iloprost was added because it has been suggested to also block T cell activation, and mPGES1−/− mice have been demonstrated to have lower production of PGI2 (51). The T cells were then stimulated with 5 nM of their cognate tetramer. T cell proliferation was determined by [3H]thymidine incorporation. The addition of PGE2 inhibited the proliferation of the P14 CD8+ T cells (Fig. 6⇑B). Furthermore, the level of inhibition was similar to infected supernatant T cell inhibition. To further demonstrate that PGE2 is altering T cell activation and skewing the response toward a Th2-like environment, the h4Ly50.5 T cell hybridoma was treated with various concentrations of commercial PGE2, F. tularensis infection supernatant, or mock supernatant in the presence of immobilized anti-CD3ε and soluble anti-CD28. After 24 h of stimulation the levels of IL-2 and IL-5 were determined. Once again, treatment of T cells with concentrations of PGE2 that are similar to F. tularensis-treated bone marrow-derived macrophages block IL-2 secretion and increased IL-5 secretion (Fig. 5⇑C). Interestingly, a lower concentration of PGE2 still blocked IL-2 secretion, whereas a higher concentration of PGE2 was needed to augment IL-5. This may explain why indomethacin treatment partially recovered IL-2 inhibition but did not completely reverse IL-5 levels, as seen in Fig. 3⇑. The fact that cells lacking the mPGES1 gene are unable to inhibit T cell proliferation like wild type and that PGE2, at concentrations similar to what is seen in infected supernatant, is able to block T cell proliferation stimulated in the absence of APCs demonstrates that PGE2 is the inhibitory factor.

Discussion

Infection by F. tularensis has several distinct clinical patterns. Each form is dependent upon the mode of transmission. F. tularensis can gain entry through the skin, the lung, or though the gut (52). Each of these portals of entry is a unique immunological niche so F. tularensis needs multiple ways to evade and/or modulate the immunological response. To date, the best-characterized F. tularensis immune evasion is its ability to invade host macrophages and escape the phagolysosomal degradation pathway (53, 54, 55). In this study, we identified a mechanism used by F. tularensis to alter both innate and adaptive immunity. F. tularensis infection of bone marrow-derived macrophages leads to the secretion of PGE2. This F. tularensis-elicited PGE2 inhibits IL-2 production and shifts the T cell response from a Th1-type-mediated response to a Th2-type response. Furthermore, F. tularensis-induced PGs also modulate macrophages, allowing F. tularensis to better replicate intracellularly. This is the first such study on F. tularensis to demonstrate a potential shift in the adaptive immune response.

Appropriate T cell responses are critical for clearing F. tularensis and establishing long-term immunity. Both CD4+ and CD8+-mediated responses can clear F. tularensis from the host (3, 4, 5, 56), and the critical factor is the secretion of IFN-γ (8, 57, 58, 59). Therefore, a Th1-like response consisting of IL-2 and IFN-γ production would be beneficial, whereas a Th2-like mediated response, dominated by IL-4 and IL-5, would be ineffective in controlling F. tularensis. NK cells are also capable of producing and secreting IFN-γ in response to F. tularensis (6). It is likely that NK-derived IFN-γ is important in initially controlling F. tularensis but is not sufficient to clear the organism. Consistent with this, SCID mice, which produce NK cells, can survive up to 40 days before succumbing to infection (8). Thus, the response of T cells and their IFN-γ is needed to clear infection. F. tularensis mechanisms that promote the production of small molecules that block IFN-γ and IL-2 and promote a Th2 response would have beneficial effects on bacterial survival. We showed that F. tularensis infection inhibits an effective T cell response, because we see decreased IL-2 production from both MHC class I- and MHC class II-restricted T cell hybrids, and decreased proliferation from naive transgenic T cells. We also see the promotion of a Th2-mediated response from the MHC class II T cell hybrids. Thus, F. tularensis has used a mechanism that can target both CD4+ and CD8+ T cells, which is important because both can control F. tularensis (3, 4). This alteration of T cell activation is mediated by the soluble compound PGE2.

PGs are known to be powerful mediators of immune responses. The immunoregulatory properties of PGE2 are the best characterized of the known PGs. Due to the multiple receptors that PGE2 can bind, it has both inflammatory and anti-inflammatory effects on immune cells (11). However, it is well established that PGE2 leads to increased cAMP levels within T cells that blocks IFN-γ and IL-2 production and subsequent T cell proliferation, and promotes the production of the Th2 cytokine IL-5 (18, 19, 20). Furthermore, PGE2 modulates APCs, by blocking maturation and suppressing the secretion of the Th1-promoting cytokine IL-12 (11, 49). PGE2 can also down-regulate inducible NO synthase (iNOS) transcription and interfere with the proper phagolysome function (60, 61, 62). Although these effects of PGE2 on the immune system have been known for several years, the role of PGE2 during infections is just starting to be examined. Recent studies show that bacterial components LPS, CpG DNA, and peptidoglycan wall can all induce COX-2 expression and PGE2 production (21, 22, 25). Interestingly, L. pneumophila and Escherichia coli both use PGE2 to modulate T cell function (31, 63). Inhibition of PGE2 production by either indomethacin treatment or deletion of the mPGES1 gene reverses the inhibition of Th1-like T cell responses by restoring IL-2 production and T cell proliferation and lowering IL-5 production down to wild-type levels. Furthermore, treatment of T cells with PGE2 recreates the effects seen with infected supernatant. It is possible that PGE2 is not the final molecule in modulating the T cell response, but rather causes the T cell to produce another factor that works in an autocrine fashion to alter T cell activation. Regardless of the final mechanism, this study clearly demonstrates that F. tularensis can elicit PGE2 upon infection of macrophages and this PGE2 directly alters T cell response.

The elicitation of PGs by F. tularensis also modulates macrophage function. Other pathogens have been shown to use PG production to allow for better intracellular replication (35, 47). Treatment of a macrophage cell line with indomethacin slows the intracellular growth of F. tularensis. PGs down-regulates iNOS mRNA levels within macrophages (60, 61), and iNOS responses are necessary for macrophages to degrade F. tularensis (52). Further studies will be needed to further characterize the response of PGs on macrophages that allow F. tularensis to grow. However, these data show that induction of PGs by F. tularensis has many beneficial properties for the infection and survival of the bacterium. PGE2 production happens within 24 h so that both innate and adaptive immune responses are impacted. This early production of PGE2 likely gives F. tularensis an advantage in replication, and likely initial infection. Furthermore, the continued production of PGE2 will then alter subsequent T cell responses, allowing the organism to persist. This would suggest that blocking PGE2 would allow for quicker clearance of the organism from the host. However, PGE2 inhibits inflammatory-induced tissue damage. For example, in E. coli infection PGE2 protects against liver damage but hampers clearance of the organism (63). Thus, blocking PGE2 may lead to increased tissue damage and quicken time to death. However, in vivo studies using indomethacin during infection with the intracellular parasite Leishmania have demonstrated reduced symptoms of disease compared with infected controls (64). Further studies are being done to analyze the in vivo role of PGE2 during F. tularensis infection.

In summary, we demonstrate that F. tularensis elicits PGE2 production from infected bone marrow-derived macrophages. This F. tularensis-induced PGE2 skews the generation of subsequent T cell responses. PGE2 blocks the production of IL-2, and subsequently the T cell’s ability to proliferate, and augments the production of IL-5. The inhibition of PGE2, either by indomethacin or deletion of the mPGES1 gene, reversed F. tularensis’s ability to modulate and skew the T cell response. Furthermore, the inhibition of PGs by indomethacin hampers the ability of F. tularensis to replicate intracellularly. Clearly, the induction of PGE2 secretion by F. tularensis appears to be advantageous to the organism. This research sheds new light onto F. tularensis’s ability to modulate host immune responses, and will need to be taken into consideration in the future development of vaccine therapy for this organism.

Acknowledgments

We thank Michael Johnson and Debbie MacArthur for technical assistance; Drs. Karen Elkins, Michelle Lennartz, Dan Loegering, Toufic Nashar, Paul Hess, and Robert Maile for helpful conversations and critical comments on this manuscript; and the Albany Medical College Microbiology Core Facility and Albany Medical College Immunology Core Facility.

Disclosures

The authors have no financial conflict of interest.

Footnotes

  • The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

  • ↵1 This work was supported in part by National Institutes of Health Grants PO1 AI056321, Subproject 1, National Institutes of Health/National Institute of Allergy and Infectious Diseases Southeast Regional Center of Excellence for Emerging Infections and Biodefense (Grant no. U54 AI 057157). M.D.W. was supported by a T32 Training Grant (AR07369-25). J.E.W. was supported by a T32 Training Grant (AI-49822) to the Center for Immunology and Microbial Disease.

  • ↵2 M.D.W. and J.E.W. contributed equally to this manuscript.

  • ↵3 Address correspondence and reprint requests to Dr. Jeffrey A. Frelinger, Department of Microbiology and Immunology, University of North Carolina, CB 7290, 804 Mary Ellen Jones Building, Chapel Hill, NC 27599. E-mail address: jfrelin{at}med.unc.edu

  • ↵4 Abbreviations used in this paper: COX, cyclooxygenase; LVS, live vaccine strain; CPRG, chlorophenol red/3-galactoside; HEL, hen egg lysozyme; MOI, multiplicity of infection; iNOS, inducible NO synthase; mPGES, membrane PGE synthase.

  • Received July 26, 2006.
  • Accepted November 16, 2006.

References

  1. ↵
    Dennis, D. T., T. V. Inglesby, D. A. Henderson, J. G. Bartlett, M. S. Ascher, E. Eitzen, A. D. Fine, A. M. Friedlander, J. Hauer, M. Layton, et al 2001. Tularemia as a biological weapon: medical and public health management. J. Am. Med. Assoc. 285: 2763-2773.
    OpenUrlCrossRefPubMed
  2. ↵
    Fortier, A. H., M. V. Slayter, R. Ziemba, M. S. Meltzer, C. A. Nacy. 1991. Live vaccine strain of Francisella tularensis: infection and immunity in mice. Infect. Immun. 59: 2922-2928.
    OpenUrlAbstract/FREE Full Text
  3. ↵
    Cowley, S. C., E. Hamilton, J. A. Frelinger, J. Su, J. Forman, K. L. Elkins. 2005. CD4−CD8− T cells control intracellular bacterial infections both in vitro and in vivo. J. Exp. Med. 202: 309-319.
    OpenUrlAbstract/FREE Full Text
  4. ↵
    Cowley, S. C., K. L. Elkins. 2003. Multiple T cell subsets control Francisella tularensis LVS intracellular growth without stimulation through macrophage interferon γ receptors. J. Exp. Med. 198: 379-389.
    OpenUrlAbstract/FREE Full Text
  5. ↵
    Ericsson, M., M. Kroca, T. Johansson, A. Sjostedt, A. Tarnvik. 2001. Long-lasting recall response of CD4+ and CD8+ αβ T cells, but not γδ T cells, to heat shock proteins of Francisella tularensis. Scand. J. Infect. Dis. 33: 145-152.
    OpenUrlCrossRefPubMed
  6. ↵
    Lopez, M. C., N. S. Duckett, S. D. Baron, D. W. Metzger. 2004. Early activation of NK cells after lung infection with the intracellular bacterium, Francisella tularensis LVS. Cell. Immunol. 232: 75-85.
    OpenUrlCrossRefPubMed
  7. ↵
    Fortier, A. H., T. Polsinelli, S. J. Green, C. A. Nacy. 1992. Activation of macrophages for destruction of Francisella tularensis: identification of cytokines, effector cells, and effector molecules. Infect. Immun. 60: 817-825.
    OpenUrlAbstract/FREE Full Text
  8. ↵
    Elkins, K. L., T. R. Rhinehart-Jones, S. J. Culkin, D. Yee, R. K. Winegar. 1996. Minimal requirements for murine resistance to infection with Francisella tularensis LVS. Infect. Immun. 64: 3288-3293.
    OpenUrlAbstract/FREE Full Text
  9. ↵
    Culkin, S. J., T. Rhinehart-Jones, K. L. Elkins. 1997. A novel role for B cells in early protective immunity to an intracellular pathogen, Francisella tularensis strain LVS. J. Immunol. 158: 3277-3284.
    OpenUrlAbstract
  10. ↵
    Narumiya, S.. 2003. Prostanoids in immunity: roles revealed by mice deficient in their receptors. Life Sci. 74: 391-395.
    OpenUrlCrossRefPubMed
  11. ↵
    Nataraj, C., D. W. Thomas, S. L. Tilley, M. T. Nguyen, R. Mannon, B. H. Koller, T. M. Coffman. 2001. Receptors for prostaglandin E2 that regulate cellular immune responses in the mouse. J. Clin. Invest. 108: 1229-1235.
    OpenUrlCrossRefPubMed
  12. ↵
    Aronoff, D. M., C. Canetti, M. Peters-Golden. 2004. Prostaglandin E2 inhibits alveolar macrophage phagocytosis through an E-prostanoid 2 receptor-mediated increase in intracellular cyclic AMP. J. Immunol. 173: 559-565.
    OpenUrlAbstract/FREE Full Text
  13. ↵
    Roper, R. L., B. Graf, R. P. Phipps. 2002. Prostaglandin E2 and cAMP promote B lymphocyte class switching to IgG1. Immunol. Lett. 84: 191-198.
    OpenUrlCrossRefPubMed
  14. ↵
    Sottile, A., I. Venza, M. Venza, A. Valenti, D. Teti. 1996. PGE2-induced immunoregulation mediated by cytokine production from cultures of human peripheral T lymphocytes. Immunopharmacol. Immunotoxicol. 18: 27-36.
    OpenUrlCrossRefPubMed
  15. ↵
    Park, J. Y., M. H. Pillinger, S. B. Abramson. 2006. Prostaglandin E2 synthesis and secretion: the role of PGE2 synthases. Clin. Immunol. 119: 229-240.
    OpenUrlCrossRefPubMed
  16. ↵
    Bezugla, Y., A. Kolada, S. Kamionka, B. Bernard, R. Scheibe, P. Dieter. 2006. COX-1 and COX-2 contribute differentially to the LPS-induced release of PGE2 and TxA2 in liver macrophages. Prostaglandins Other Lipid Mediat. 79: 93-100.
    OpenUrlCrossRefPubMed
  17. ↵
    Ikeda-Matsuo, Y., Y. Ikegaya, N. Matsuki, S. Uematsu, S. Akira, Y. Sasaki. 2005. Microglia-specific expression of microsomal prostaglandin E2 synthase-1 contributes to lipopolysaccharide-induced prostaglandin E2 production. J. Neurochem. 94: 1546-1558.
    OpenUrlCrossRefPubMed
  18. ↵
    Kvirkvelia, N., I. Vojnovic, T. D. Warner, V. Athie-Morales, P. Free, N. Rayment, B. M. Chain, T. W. Rademacher, T. Lund, I. M. Roitt, P. J. Delves. 2002. Placentally derived prostaglandin E2 acts via the EP4 receptor to inhibit IL-2-dependent proliferation of CTLL-2 T cells. Clin. Exp. Immunol. 127: 263-269.
    OpenUrlCrossRefPubMed
  19. ↵
    Betz, M., B. S. Fox. 1991. Prostaglandin E2 inhibits production of Th1 lymphokines but not of Th2 lymphokines. J. Immunol. 146: 108-113.
    OpenUrlAbstract
  20. ↵
    Harris, S. G., J. Padilla, L. Koumas, D. Ray, R. P. Phipps. 2002. Prostaglandins as modulators of immunity. Trends Immunol. 23: 144-150.
    OpenUrlCrossRefPubMed
  21. ↵
    Chen, B. C., Y. S. Chang, J. C. Kang, M. J. Hsu, J. R. Sheu, T. L. Chen, C. M. Teng, C. H. Lin. 2004. Peptidoglycan induces nuclear factor-κB activation and cyclooxygenase-2 expression via Ras, Raf-1, and ERK in RAW 264.7 macrophages. J. Biol. Chem. 279: 20889-20897.
    OpenUrlAbstract/FREE Full Text
  22. ↵
    Chen, Y., J. Zhang, S. A. Moore, Z. K. Ballas, J. P. Portanova, A. M. Krieg, D. J. Berg. 2001. CpG DNA induces cyclooxygenase-2 expression and prostaglandin production. Int. Immunol. 13: 1013-1020.
    OpenUrlAbstract/FREE Full Text
  23. ↵
    Smith, R. S., R. Kelly, B. H. Iglewski, R. P. Phipps. 2002. The pseudomonas autoinducer N-(3-oxododecanoyl) homoserine lactone induces cyclooxygenase-2 and prostaglandin E2 production in human lung fibroblasts: implications for inflammation. J. Immunol. 169: 2636-2642.
    OpenUrlAbstract/FREE Full Text
  24. ↵
    Treffkorn, L., R. Scheibe, T. Maruyama, P. Dieter. 2004. PGE2 exerts its effect on the LPS-induced release of TNF-α, ET-1, IL-1α, IL-6 and IL-10 via the EP2 and EP4 receptor in rat liver macrophages. Prostaglandins Other Lipid Mediat. 74: 113-123.
    OpenUrlCrossRefPubMed
  25. ↵
    Uematsu, S., M. Matsumoto, K. Takeda, S. Akira. 2002. Lipopolysaccharide-dependent prostaglandin E2 production is regulated by the glutathione-dependent prostaglandin E2 synthase gene induced by the Toll-like receptor 4/MyD88/NF-IL6 pathway. J. Immunol. 168: 5811-5816.
    OpenUrlAbstract/FREE Full Text
  26. ↵
    Sjostedt, A.. 2006. Intracellular survival mechanisms of Francisella tularensis, a stealth pathogen. Microbes Infect. 8: 561-567.
    OpenUrlCrossRefPubMed
  27. ↵
    Cossart, P., P. J. Sansonetti. 2004. Bacterial invasion: the paradigms of enteroinvasive pathogens. Science 304: 242-248.
    OpenUrlAbstract/FREE Full Text
  28. ↵
    Kirveskari, J., Q. He, M. Leirisalo-Repo, O. Maki-Ikola, M. Wuorela, A. Putto-Laurila, K. Granfors. 1999. Enterobacterial infection modulates major histocompatibility complex class I expression on mononuclear cells. Immunology 97: 420-428.
    OpenUrlCrossRefPubMed
  29. ↵
    Mitchell, E. K., P. Mastroeni, A. P. Kelly, J. Trowsdale. 2004. Inhibition of cell surface MHC class II expression by Salmonella. Eur. J. Immunol. 34: 2559-2567.
    OpenUrlCrossRefPubMed
  30. ↵
    Weiss, D. J., O. A. Evanson, D. J. McClenahan, M. S. Abrahamsen, B. K. Walcheck. 2001. Regulation of expression of major histocompatibility antigens by bovine macrophages infected with Mycobacterium avium subsp. paratuberculosis or Mycobacterium avium subsp. avium. Infect. Immun. 69: 1002-1008.
    OpenUrlAbstract/FREE Full Text
  31. ↵
    Neumeister, B., M. Faigle, D. Spitznagel, A. Mainka, A. Ograbek, H. Wieland, N. Mannowetz, H. G. Rammensee. 2005. Legionella pneumophila down-regulates MHC class I expression of human monocytic host cells and thereby inhibits T cell activation. Cell Mol. Life Sci. 62: 578-588.
    OpenUrlCrossRefPubMed
  32. ↵
    Bonecini-Almeida, M. G., J. L. Ho, N. Boechat, R. C. Huard, S. Chitale, H. Doo, J. Geng, L. Rego, L. C. Lazzarini, A. L. Kritski, et al 2004. Down-modulation of lung immune responses by interleukin-10 and transforming growth factor β (TGF-β) and analysis of TGF-β receptors I and II in active tuberculosis. Infect. Immun. 72: 2628-2634.
    OpenUrlAbstract/FREE Full Text
  33. ↵
    Briken, V., S. A. Porcelli, G. S. Besra, L. Kremer. 2004. Mycobacterial lipoarabinomannan and related lipoglycans: from biogenesis to modulation of the immune response. Mol. Microbiol. 53: 391-403.
    OpenUrlCrossRefPubMed
  34. ↵
    Neild, A. L., S. Shin, C. R. Roy. 2005. Activated macrophages infected with Legionella inhibit T cells by means of MyD88-dependent production of prostaglandins. J. Immunol. 175: 8181-8190.
    OpenUrlAbstract/FREE Full Text
  35. ↵
    Uchiya, K., T. Nikai. 2004. Salmonella enterica serovar Typhimurium infection induces cyclooxygenase 2 expression in macrophages: involvement of Salmonella pathogenicity island 2. Infect. Immun. 72: 6860-6869.
    OpenUrlAbstract/FREE Full Text
  36. ↵
    Cox, G. W., B. J. Mathieson, L. Gandino, E. Blasi, D. Radzioch, L. Varesio. 1989. Heterogeneity of hematopoietic cells immortalized by v-myc/v-raf recombinant retrovirus infection of bone marrow or fetal liver. J. Natl. Cancer Inst. 81: 1492-1496.
    OpenUrlAbstract/FREE Full Text
  37. ↵
    Shastri, N., F. Gonzalez. 1993. Endogenous generation and presentation of the ovalbumin peptide/Kb complex to T cells. J. Immunol. 150: 2724-2736.
    OpenUrlAbstract
  38. ↵
    Hockett, R. D., J. R. Cook, K. Findlay, C. V. Harding. 1996. Interferon-γ differentially regulates antigen-processing functions in distinct endocytic compartments of macrophages with constitutive expression of class II major histocompatibility complex molecules. Immunology 88: 68-75.
    OpenUrlCrossRefPubMed
  39. ↵
    Wang, B., R. Maile, R. Greenwood, E. J. Collins, J. A. Frelinger. 2000. Naive CD8+ T cells do not require costimulation for proliferation and differentiation into cytotoxic effector cells. J. Immunol. 164: 1216-1222.
    OpenUrlAbstract/FREE Full Text
  40. ↵
    Trebino, C. E., J. L. Stock, C. P. Gibbons, B. M. Naiman, T. S. Wachtmann, J. P. Umland, K. Pandher, J. M. Lapointe, S. Saha, M. L. Roach, et al 2003. Impaired inflammatory and pain responses in mice lacking an inducible prostaglandin E synthase. Proc. Natl. Acad. Sci. USA 100: 9044-9049.
    OpenUrlAbstract/FREE Full Text
  41. ↵
    Kerry, S. E., J. Buslepp, L. A. Cramer, R. Maile, L. L. Hensley, A. I. Nielsen, P. Kavathas, B. J. Vilen, E. J. Collins, J. A. Frelinger. 2003. Interplay between TCR affinity and necessity of coreceptor ligation: high-affinity peptide-MHC/TCR interaction overcomes lack of CD8 engagement. J. Immunol. 171: 4493-4503.
    OpenUrlAbstract/FREE Full Text
  42. ↵
    Del Prete, G., E. Maggi, S. Romagnani. 1994. Human Th1 and Th2 cells: functional properties, mechanisms of regulation, and role in disease. Lab. Invest. 70: 299-306.
    OpenUrlPubMed
  43. ↵
    Seder, R. A.. 1994. Acquisition of lymphokine-producing phenotype by CD4+ T cells. J. Allergy Clin. Immunol. 94: 1195-1202.
    OpenUrlCrossRefPubMed
  44. ↵
    Pircher, H., K. Burki, R. Lang, H. Hengartner, R. M. Zinkernagel. 1989. Tolerance induction in double specific T-cell receptor transgenic mice varies with antigen. Nature 342: 559-561.
    OpenUrlCrossRefPubMed
  45. ↵
    Blanco, J. C., C. Contursi, C. A. Salkowski, D. L. DeWitt, K. Ozato, S. N. Vogel. 2000. Interferon regulatory factor (IRF)-1 and IRF-2 regulate interferon γ-dependent cyclooxygenase 2 expression. J. Exp. Med. 191: 2131-2144.
    OpenUrlAbstract/FREE Full Text
  46. ↵
    Makoul, G. T., D. R. Robinson, A. K. Bhalla, L. H. Glimcher. 1985. Prostaglandin E2 inhibits the activation of cloned T cell hybridomas. J. Immunol. 134: 2645-2650.
    OpenUrlAbstract
  47. ↵
    Liu, W., S. Dubinett, S. L. Patterson, K. A. Kelly. 2006. COX-2 inhibition affects growth rate of Chlamydia muridarum within epithelial cells. Microbes Infect. 8: 478-486.
    OpenUrlCrossRefPubMed
  48. ↵
    Dombek, K. M., L. O. Ingram. 1984. Effects of ethanol on the Escherichia coli plasma membrane. J. Bacteriol. 157: 233-239.
    OpenUrlAbstract/FREE Full Text
  49. ↵
    Yang, L., N. Yamagata, R. Yadav, S. Brandon, R. L. Courtney, J. D. Morrow, Y. Shyr, M. Boothby, S. Joyce, D. P. Carbone, R. M. Breyer. 2003. Cancer-associated immunodeficiency and dendritic cell abnormalities mediated by the prostaglandin EP2 receptor. J. Clin. Invest. 111: 727-735.
    OpenUrlCrossRefPubMed
  50. ↵
    Kudo, I., M. Murakami. 2005. Prostaglandin e synthase, a terminal enzyme for prostaglandin E2 biosynthesis. J. Biochem. Mol. Biol. 38: 633-638.
    OpenUrlCrossRefPubMed
  51. ↵
    Kamei, D., K. Yamakawa, Y. Takegoshi, M. Mikami-Nakanishi, Y. Nakatani, S. Oh-Ishi, H. Yasui, Y. Azuma, N. Hirasawa, K. Ohuchi, et al 2004. Reduced pain hypersensitivity and inflammation in mice lacking microsomal prostaglandin E synthase-1. J. Biol. Chem. 279: 33684-33695.
    OpenUrlAbstract/FREE Full Text
  52. ↵
    Lindgren, H., L. Stenman, A. Tarnvik, A. Sjostedt. 2005. The contribution of reactive nitrogen and oxygen species to the killing of Francisella tularensis LVS by murine macrophages. Microbes Infect. 7: 467-475.
    OpenUrlCrossRefPubMed
  53. ↵
    Santic, M., M. Molmeret, K. E. Klose, Y. Abu Kwaik. 2006. Francisella tularensis travels a novel, twisted road within macrophages. Trends Microbiol. 14: 37-44.
    OpenUrlCrossRefPubMed
  54. ↵
    Santic, M., M. Molmeret, K. E. Klose, S. Jones, Y. A. Kwaik. 2005. The Francisella tularensis pathogenicity island protein IglC and its regulator MglA are essential for modulating phagosome biogenesis and subsequent bacterial escape into the cytoplasm. Cell Microbiol. 7: 969-979.
    OpenUrlCrossRefPubMed
  55. ↵
    Clemens, D. L., B. Y. Lee, M. A. Horwitz. 2004. Virulent and avirulent strains of Francisella tularensis prevent acidification and maturation of their phagosomes and escape into the cytoplasm in human macrophages. Infect. Immun. 72: 3204-3217.
    OpenUrlAbstract/FREE Full Text
  56. ↵
    Yee, D., T. R. Rhinehart-Jones, K. L. Elkins. 1996. Loss of either CD4+ or CD8+ T cells does not affect the magnitude of protective immunity to an intracellular pathogen, Francisella tularensis strain LVS. J. Immunol. 157: 5042-5048.
    OpenUrlAbstract
  57. ↵
    Rhinehart-Jones, T. R., A. H. Fortier, K. L. Elkins. 1994. Transfer of immunity against lethal murine Francisella infection by specific antibody depends on host γ interferon and T cells. Infect. Immun. 62: 3129-3137.
    OpenUrlAbstract/FREE Full Text
  58. ↵
    Polsinelli, T., M. S. Meltzer, A. H. Fortier. 1994. Nitric oxide-independent killing of Francisella tularensis by IFN-γ-stimulated murine alveolar macrophages. J. Immunol. 153: 1238-1245.
    OpenUrlAbstract
  59. ↵
    Anthony, L. S., E. Ghadirian, F. P. Nestel, P. A. Kongshavn. 1989. The requirement for γ interferon in resistance of mice to experimental tularemia. Microb. Pathog. 7: 421-428.
    OpenUrlCrossRefPubMed
  60. ↵
    Harbrecht, B. G., Y. M. Kim, E. A. Wirant, R. L. Simmons, T. R. Billiar. 1997. Timing of prostaglandin exposure is critical for the inhibition of LPS- or IFN-γ-induced macrophage NO synthesis by PGE2. J. Leukocyte Biol. 61: 712-720.
    OpenUrlAbstract
  61. ↵
    D’Acquisto, F., L. Sautebin, T. Iuvone, M. Di Rosa, R. Carnuccio. 1998. Prostaglandins prevent inducible nitric oxide synthase protein expression by inhibiting nuclear factor-κB activation in J774 macrophages. FEBS Lett. 440: 76-80.
    OpenUrlCrossRefPubMed
  62. ↵
    Rangel Moreno, J., I. Estrada Garcia, M. De La Luz Garcia Hernandez, D. Aguilar Leon, R. Marquez, R. Hernandez Pando. 2002. The role of prostaglandin E2 in the immunopathogenesis of experimental pulmonary tuberculosis. Immunology 106: 257-266.
    OpenUrlCrossRefPubMed
  63. ↵
    Takano, M., H. Nishimura, Y. Kimura, J. Washizu, Y. Mokuno, Y. Nimura, Y. Yoshikai. 1998. Prostaglandin E2 protects against liver injury after Escherichia coli infection but hampers the resolution of the infection in mice. J. Immunol. 161: 3019-3025.
    OpenUrlAbstract/FREE Full Text
  64. ↵
    Farrell, J. P., C. E. Kirkpatrick. 1987. Experimental cutaneous leishmaniasis II: a possible role for prostaglandins in exacerbation of disease in Leishmania major-infected BALB/c mice. J. Immunol. 138: 902-907.
    OpenUrlAbstract
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