Diverse roles for T-bet in the effector responses required for resistance to infection

Abstract

The transcription factor T-bet has been most prominently linked to natural killer (NK) and T cell production of interferon-γ (IFN-γ), a cytokine required for the control of a diverse array of intracellular pathogens. Indeed, in mice challenged with the parasite Toxoplasma gondii, NK and T cell responses are characterized by marked increases of T-bet expression. Unexpectedly, T-bet^−/− mice infected with T. gondii develop a strong NK cell IFN-γ response that controls parasite replication at the challenge site, but display high parasite burdens at secondary sites colonized by T. gondii and succumb to infection. The loss of T-bet had a modest effect on T cell production of IFN-γ but did not impact on the generation of parasite-specific T cells. However, the absence of T-bet resulted in lower T cell expression of CD11a, Ly6C, KLRG-1, and CXCR3 and fewer parasite-specific T cells at secondary sites of infection, associated with a defect in parasite control at these sites. Together, these data highlight T-bet independent pathways to IFN-γ production, and reveal a novel role for this transcription factor in coordinating the T cell responses necessary to control this infection in peripheral tissues.

Introduction

The T-box transcription factor T-bet is expressed in cells of the innate and adaptive immune system (1–4), but it is perhaps most prominently linked to the production of IFN-γ in T cells and NK cells (1, 2, 5). While many studies have focused on the role of T-bet in CD4⁺ T cell production of IFN-γ, there are studies in which it also impacts on the ability of NK and CD8⁺ T cells to produce IFN-γ (6–9). In addition, T-bet has been shown to bind to the promoter region of other genes, suggesting a broader function in the immune response. For example, T-bet controls key checkpoints in NK cell maturation (10) and also inhibits T cell production of IL-2, IL-4, and IL-17, thus repressing other CD4⁺ T cell differentiation fates that include T helper 2 (Th2) and Th17 cells (1, 2, 11). Furthermore, T-bet induces expression of the chemokine receptor CXCR3, and in vitro studies have identified a role for T-bet in the chemotaxis of CD4⁺ T cells (12). Consistent with a key role for T-bet in the development of Th1 cells, this transcription factor is required for IFN-γ mediated resistance to Leishmania major, Salmonella, Mycobacterium tuberculosis, and HSV-2 (2, 13–15). While production of IFN-γ is also required to limit replication of Listeria monocytogenes and LCMV, tahe IFN-γ produced in the T-bet^−/− mice is sufficient for controlling these pathogens (9, 16). One explanation for this phenomenon is that these T-bet independent pathways to IFN-γ production are mediated through a related T-box transcription factor, Eomesodermin (5, 17).

In current models, challenge of mice with the intracellular parasite T. gondii results in the production of interleukin 12 (IL-12) by dendritic cells and macrophages which promotes the activation and expansion of NK cell and T cell populations that express high levels of T-bet and are associated with the secretion of IFN-γ (18–22). The production of IFN-γ, which engages numerous antimicrobial mechanisms (23, 24), is essential for the local control of T. gondii in multiple tissues. The studies presented here demonstrate that when mice deficient in T-bet are challenged with T. gondii, they control parasite replication at the site of initial challenge, associated with strong NK cell responses, and can generate parasite-specific T cells that produce IFN-γ. However, these mice do not survive acute infection, and secondary sites of parasite colonization that include the heart, thymus, lung, and brain have high parasite burdens. The increased mortality is associated with alterations in the phenotype of parasite-specific T cell populations that include reduced expression of markers of activation (CD11a), effector status (Ly6C and KLRG-1), and trafficking (CXCR3) as well as a marked reduction in their numbers at these peripheral sites. The use of a mixed bone marrow chimeric approach revealed a cell-intrinsic requirement for T-bet for the development of appropriate parasite-specific effector T cells. Together, these data highlight a novel role of T-bet in coordinating the CD4⁺ and CD8⁺ T cell responses that are essential for the control of infection in peripheral tissues.

Materials and Methods

Mice, infection, and antibody treatment

T-bet deficient (T-bet^−/−), CBA/CaJ, Thy1.1⁺C57BL/6 and Swiss Webster mice were purchased from Jackson Laboratory. Wild-type C57BL/6 (WT) mice were purchased from Taconic. CD45.1⁺C57BL/6 mice were purchased from NCI. All mice were housed in a specific-pathogen free environment at the University of Pennsylvania School of Veterinary Medicine in accordance with federal guidelines and with approval of the Institutional Animal Care and Use Committee. The ME49 strain of T. gondii was maintained in Swiss Webster and CBA/CaJ mice and used as a source of tissue cysts for i.p. (10–20 cysts) infections. Soluble Toxoplasma Ag (STAg) was prepared from the RH strain of T. gondii as previously described (25). For IFN-γ neutralization experiments, mice were treated with 1mg αIFN-γ or isotype control on days 0, 3, and 6 of infection. For depletion of NK cells, mice were treated with 50ul αAsialo gm-1 or isotype control on days −1, 3, and 6 of infection. For bone marrow chimeras, CD45.1⁺ congenic mice were irradiated (1000rads) and i.v. injected with a 1:1 mixture of 6×10⁶ cells isolated from WT (CD45.2⁺Thy1.1⁺) and T-bet^−/− (CD45.2⁺Thy1.1⁻) bone marrow. Mice were given water containing sulfamethoxazol for the first 2 weeks post irradiation. Mice were allowed to reconstitute >8 weeks following irradiation, and were infected with T. gondii as described.

Isolation and Analysis of Immune Populations

Single-cell suspensions from the spleens, lymph nodes (LNs), and peritoneal exudate cells (PECs) were prepared as previously described (26). Lungs were inflated with a solution of 1mg/ml Collagenase A (Roche) and 100ug/ml DNase (Roche) and then diced and digested in the same solution for 60 minutes at 37°C to obtain a single cell suspension. The resulting cells were then passed through a 70μm filter and used for FACS analysis. Cells were stained for surface markers in FACS buffer (0.5% BSA, 2mM EDTA in PBS), fixed with Foxp3 Fixation buffer (eBioscience), and stained for intracellular markers in Foxp3 Permeabilization buffer (eBioscience). To measure intracellular cytokine production, isolated cells were cultured in cRPMI (1% penicillin/streptomycin, 2 mM L-glutamine, 10% fetal bovine serum, 0.1% beta-mercaptoethanol, 1% nonessential amino acids, and 1 mM sodium pyruvate) in triplicate at 1 × 10⁶ cells/mL in a 96-well U-bottom plate, with PMA and ionomycin for 4 hr with Brefeldin A (Sigma) and monensin (BD). Cells were rinsed, stained for surface markers at 4°C, and fixed with 4% PFA in PBS for 11 min at room temperature. Intracellular cytokines were detected by staining in FACs buffer containing 0.5% saponin (Sigma). Splenocytes were also restimulated at 1 × 10⁵ cells/mL in cRPMI alone (media) or with either αCD3 (1ug/mL) or soluble T. gondii antigen (STAg, 12.5ug/mL) and incubated at 37°C for 72 hours. Sera were assayed for ALT, and supernatants and sera were assayed for IL-12p40 and IFN-γ by ELISA.

Flow Cytometry and Imaging

The following antibodies were purchased from BD: CD3 FITC, Ki67 FITC, CD49b (DX5) PE, Ki67 AF700, CD4 Pacific Blue, PSGL-1 BV421, CD8α PE-CF594, CD3 PE-CF594, CD122 Biotin, and Streptavidin PE-TexasRed. The following antibodies were purchased from eBioscience: CD25 FITC, Foxp3 AF488, CD11a FITC, NK1.1 FITC, CD19 FITC, CD4 FITC, KLRG-1 FITC, CD11a PE, Ly6C (clone HK1.4) PerCP-Cy5.5, CD69 PerCP-Cy5.5, NKp46 eFluor660, T-bet eFluor660, CD11c PE-Cy7, KLRG-1 PE-Cy7, IFN-γ PE-Cy7, CD25 APC-eFluor780, CD3 APC-eFluor780, Ly6C (clone HK1.4) APC-eFluor780, Thy1.1 APC-eFluor780, Foxp3 AF700, CD45.2 eFluor450, Ly6C (clone HK1.4) eFluor450, and CD4 eFluor650NC. The following antibodies were purchased from BioLegend: CD8 APC-Cy7, MHCII Pacific Blue, CD11a PerCP-Cy5.5, CXCR3 PE-Cy7, TCRβ AF700, CD3 Pacific Blue, and NK1.1 Pacific Blue. Invitrogen live/dead Aqua stain was used to determine viability. Biotinylated Tgd-057 MHC-I monomers were kindly provided by E. John Wherry (University of Pennsylvania) and tetramerized by incubation with streptavidin-conjugated PE. PE-conjugated AS-15 MHC-II tetramers were kindly provided by Marion Pepper (University of Washington). All samples were run on an LSRFortessa (BD) and analyzed using FlowJo software (Tree Star). Analysis and presentation of distributions was performed using SPICE version 5.35, downloaded from http://exon.niaid.nih.gov/spice/ (27). Images were obtained using the ImageStream (Amnis) and analyzed using IDEAS software (Amnis). To determine T-bet localization, nuclear and cytoplasmic masking functions were made using DAPI staining; these masks were then applied to T-bet expression.

Parasite Detection

For IHC detection of T. gondii, tissues were fixed in 10% formalin solution and then paraffin- embedded and sectioned. Sections were deparaffinized, rehydrated, antigen retrieved in 0.01M Sodium Citrate Buffer (PH 6.0) and endogenous peroxidase blocked by 0.3% H2O2 in PBS. After blocking with 2% normal goat serum, the sections were incubated either with anti-Toxoplasma antibody or isotype control. The sections were then incubated with Biotinylated goat anti-rabbit IgG (Vector, Burlington, CA), and ABC reagent was applied (Vectastain ABC Kit, Vector Labs). Then DAB substrate (Vector Labs) was used to visualize specific staining according to manufacturer’s instructions and slides were counterstained with hematoxylin. For fluorescence microscopy, brains were frozen in OCT, 6 μm sections prepared and stained with antibodies against CD4 and CD8 in Cy3 and against T. gondii in Alexa488 as well as DAPI for nuclear staining. To quantify parasite burden in the peritoneal exudate, 100,000 cells were used to prepare cytospins. Cells were methanol fixed and then stained with the Protocol Hema-3 Stain Set, and the ratio of infected cells to total cells in a field of vision was calculated. All images were obtained on a Nikon E600 microscope using a 20X (brain) or 40X (liver, lung, heart, thymus, PECs) objective and NIS Elements Imaging software. For quantitative PCR, DNA was isolated from tissues using the High Pure PCR Template Preparation Kit (Roche) followed by qPCR measuring the abundance of the T. gondii gene B1 using the primers 5′-TCTTTAAAGCGTTCGTGGTC-3′ (forward) and 5′-GGAACTGCATCCGTTCATGAG-3′ (reverse).

Statistics

Bar graphs and scatter plots were plotted as means with the standard error of the mean in Prism 5 software (GraphPad). All statistics were performed using an unpaired Student’s t test, except the bone marrow chimera experiments in which the statistics were performed using a paired Student’s t test in Prism 5 software.

Results

Expression of T-bet during the immune response to Toxoplasma gondii

As part of studies to understand the role of T-bet during the immune response against T. gondii, the populations involved in protective immunity were surveyed for infection-induced alterations in T-bet expression and localization. In naïve WT mice, there is heterogeneous expression of the integrin CD49b (DX5) within the NK cell population, with high expression indicative of NK cell maturity. In uninfected mice, T-bet is not highly expressed in NK cells, but following challenge with T. gondii, a population of mature DX5^hi NK cells that express high levels of T-bet emerged (Figure 1a). Similarly, based on the use of Ly6C and KLRG-1 as markers of CD4⁺ and CD8⁺ T cell effector populations, respectively (28–30), naïve mice had few effector cells, and this population expressed low levels of T-bet (Figures 1b and 1c). However, at day 9 post-infection, a subset of CD4⁺ T cells that expressed T-bet was evident and a portion of these cells also expressed high levels of Ly6C; the number of these T-bet⁺Ly6C^hiCD4⁺ T cells that emerged after infection was significantly increased over naïve mice (Figure 1b). Similarly, at this timepoint, there was an increase in the percentage and number of CD8⁺ T cells that expressed T-bet and KLRG-1 (Figure 1c).

Figure 1. T-bet expression increases in NK and T cells after infection with Toxoplasma gondii.

WT C57BL/6 mice were infected intraperitoneally (i.p.) with T. gondii. (A) After 5 or (B, C) 9 days, splenocytes from infected and uninfected mice were harvested and stained for analysis by flow cytometry. (A) NK cells were identified as NK1.1⁺CD3⁻ live cells. (B) CD4⁺ T cells were identified as CD3⁺CD4⁺CD8a⁻ live cells. (C) CD8⁺ T cells were identified as CD3⁺CD4⁻CD8a⁺ live cells. Data are representative of 3 experiments, n=3 per experiment.

* p<0.05, ** p<0.01

Further analysis revealed that in the CD4⁺ T cells from infected mice, T-bet expression was associated with expression of the activation markers CD11a, an integrin that, together with CD18, comprises lymphocyte function-associated antigen 1 (LFA-1), CXCR3, a Th1 associated chemokine receptor, and Ly6C (Figure 2a and Supplemental Figure 1). More than 95% of T-bet⁺ CD4⁺ T cells expressed at least one of these activation markers, and approximately 20% of these T-bet⁺ cells expressed all three activation markers (Figure S1). In contrast, only 25% of T-bet⁻ CD4⁺ T cells expressed any of these activation markers, and less than 5% of these cells expressed all three activation markers. Expression of CD11a has been used as a marker of activation for the polyclonal antigen-specific T cell population in a variety of infectious settings, including toxoplasmosis (31–34). Parasite-specific CD4⁺ T cells were identified using an MHC-II tetramer for AS15 (35) and, consistent with the polyclonal (CD11a^hi) population, these cells expressed T-bet, Ly6C, and CD11a (Figure 2b). Among the CD8⁺ T cells, T-bet was highly expressed in the infection-induced polyclonal CD11a^hi population (Figure 2c), and T-bet expression was closely associated with expression of CXCR3 and KLRG-1 (Figure S1). More than 95% of T-bet-expressing CD8⁺ T cells expressed CXCR3, high levels of CD11a, and/or KLRG-1, with nearly 50% expressing all three activation markers. Among the T-bet⁻ CD8⁺ T cell population, the majority (75%) did not express CXCR3, high levels of CD11a, or KLRG-1 and less than 1% of T-bet⁻ CD8⁺ T cells expressed all three of these activation markers (Figure S1). Moreover, consistent with the polyclonal CD11a^hi population, when parasite-specific CD8⁺ T cells were identified using an MHC-I tetramer for the parasite antigen Tgd057 (36) these cells highly expressed T-bet, KLRG-1, and CD11a (Figure 2d). Thus, following infection with T. gondii, T-bet is expressed in activated CD4⁺ and CD8⁺ T cells, and this correlates closely with the expression of CD11a, Ly6C and KLRG-1 by parasite-specific effector populations.

Figure 2. T-bet expressing T cells have a distinct phenotype after infection with T. gondii.

Splenocytes were harvested from T. gondii infected mice 9–11 days post-infection and stained for analysis by flow cytometry and Image Stream. (A) T-bet expression levels were assessed in the polyclonal CD11a^hi CD4⁺ T cell population compared to the CD11a^lo population. (B) T-bet, Ly6C, and CD11a expression levels were assessed on total CD4⁺ T cells (dashed lines) and tetramer⁺CD4⁺ T cells (solid lines). (C) T-bet expression levels were assessed in the polyclonal CD11a^hi CD8⁺ T cell population compared to the CD11a^lo population. (D) T-bet, KLRG-1, and CD11a expression levels were assessed on total CD8⁺ T cells (dashed lines) and tetramer⁺CD8⁺ T cells (solid lines). (E) T-bet localization was assessed within the Ly6C^loCD11a^hi and Ly6C^hiCD11a^hi CD4⁺ T cell population using the Image Stream. (F) T-bet localization was assessed within the KLRG-1^loCD11a^hi and KLRG-1^hiCD11a^hi CD8⁺ T cell population using the Image Stream. Data are representative of 2–4 experiments, n=3 per experiment.

Recently, human effector CD8⁺ T cells have been shown to have heterogeneous levels of T-bet protein that are associated with differential localization of T-bet in the nucleus and cytoplasm (37). Since T-bet expression closely correlated with expression of Ly6C and KLRG-1, proteins that aid in defining effector populations in murine T cells, ImageStream analysis was combined with these surface molecules to visualize T-bet localization. Because there were relatively few numbers of cells that bound to the MHC tetramers, the polyclonal CD11a^hi CD4⁺ and CD8⁺ T cell populations were analyzed. Among the Ly6C^loCD11a^hiCD4⁺ T cell (memory precursor) population, T-bet was present primarily in the nucleus, as quantified by colocalization with DAPI (Figure 2e). In contrast, within the Ly6C^hiCD11a^hiCD4⁺ T cell (effector) population, the expression of T-bet in approximately 75% of cells did not colocalize with DAPI, indicating that T-bet was cytoplasmic in these cells (Figure 2e). Analysis of the CD8⁺ T cells revealed that within the KLRG-1^loCD11a^hiCD8⁺ T cell (memory precursor) population, a portion of cells expressed low levels of T-bet that was difficult to localize, but in those cells that highly expressed this protein it was present primarily in the nucleus (Figure 2f). However, the majority of KLRG-1^hiCD11a^hiCD8⁺ T cells (effectors) had T-bet present in the cytoplasm (Figure 2f). Together, these data demonstrate that among the antigen-specific (CD11a^hi) T cells, T-bet is differentially localized among the effector and memory precursor T cell populations.

T-bet knockout mice are susceptible to infection with T. gondii

To directly assess the significance of the infection-induced increases of T-bet in activated NK and T cells, WT and T-bet^−/− mice were challenged with ME49, an avirulent strain of T. gondii. WT mice survived this challenge, yet mice lacking T-bet succumbed to infection between days 9 and 14 post-infection (Figure 3a). In many instances, susceptibility to T. gondii is either a consequence of an inability to control parasite replication (21, 38, 39) or the development of T cell-mediated immune pathology (40–43). In order to determine the cause of the increased mortality seen in T-bet^−/− mice, parasite burden and levels of immune-mediated damage were assessed. Analysis of the peritoneal exudate cells (PECs) revealed that there was a small but reproducible increase in parasite burden in the T-bet^−/− mice at day 5 (Figure 3b) and day 9 post-infection (data not shown). A more comprehensive analysis of the tissues from infected WT and T-bet^−/− mice by quantitative PCR (Figure 3c) and immunohistochemistry (Figures 3d–g) revealed that in WT mice few parasites were detected at peripheral sites of infection such as the thymus, heart, lung, and brain. However, in T-bet^−/− mice, these tissues all contained areas of extensive parasite replication. Liver lesions (areas of coagulative necrosis in the liver that are typically associated with CD4⁺ T cell-mediated immunopathology) were present in the tissues from WT mice but were absent or reduced in those from T-bet^−/− mice (Figure 3h). While there was no difference in the frequency of NK cells in the livers from infected WT and T-bet^−/− mice, there was a marked reduction in the number of T cells in the livers from the T-bet^−/− mice (data not shown). Furthermore, WT mice had higher levels of infection-induced alanine aminotransferase (ALT), which is indicative of liver damage, in the sera than T-bet^−/− mice at day 9 post-infection (Figure 3i), consistent with the elevated pathology. Thus, the major cause of susceptibility of the T-bet^−/− mice is a failure to control parasite replication at secondary sites of infection.

Figure 3. T-bet is necessary for survival and regional parasite control during T. gondii infection.

(A) WT and T-bet deficient (T-bet^−/−) mice were infected i.p. with T. gondii and were monitored for survival. (B) At day 9 post-infection, peritoneal lavage was performed and peritoneal exudate cells (PECs) were collected for cytospin preparation and analysis of parasite burden. (C) DNA was extracted from spleens (●), lungs (■), hearts (◆), and thymi (▲) from infected WT and T-bet^−/− mice and levels of parasite DNA were assessed using qPCR. (D–G) Tissues from WT and T-bet^−/− mice were isolated 9–11 days post infection and fixed in formalin solution, sectioned, and stained for T. gondii antigen. (H) Liver samples were harvested from mice 9 days post-infection and stained by H&E. Area of necrosis is outlined with a dashed white line. (I) Serum was harvested from uninfected and T. gondii infected WT and T-bet^−/− mice and analyzed for ALT levels. Data are representative of 2–3 experiments, n=3–5 per experiment.

Innate responses in the absence of T-bet

To better understand the underlying cause of the increased susceptibility of the T-bet^−/− mice, the innate response to T. gondii was characterized. These studies revealed no significant differences in serum levels of infection-induced IL-12 or IFN-γ in the WT and T-bet^−/− mice at days 5 (data not shown) and 9 post-infection (Figures 4a and 4b). Moreover, at day 5 post-infection, a period of enhanced NK cell activity (44, 45), there was equivalent expansion of DX5⁺ NK cells from T-bet^−/− and WT mice (Figure 4c) and these cells expressed similar levels of the activation marker CD69 (Figure 4d). Analysis of NK cells from the spleens of infected and uninfected mice showed that there were basal differences in IFN-γ production from WT and T-bet^−/− NK cells (data not shown), but after challenge these cells produced similar levels of IFN-γ (Figures 4e and 4f). Additionally, when WT and T-bet^−/− mice were treated with an IFN-γ blocking antibody, there was a marked increase in the percentage of infected PECs over the isotype control treated mice (Figures 4g and 4h). Furthermore, depletion of NK cells in T-bet^−/− during infection with T.gondii also resulted in a similar increase in the percentage of infected PECs (Figure 4i). Together, these data establish that T-bet is dispensable for NK cell activation and the early IFN-γ produced in the absence of T-bet contributes to the control of parasite replication at the local site of infection.

Figure 4. NK and cytokine responses are intact in infected T-bet^−/− mice.

(A–F) WT and T-bet^−/− mice were infected i.p. with T. gondii. (A, B) Serum was harvested from day 9 infected and uninfected WT and T-bet^−/− mice and assayed by ELISA for (A) IL-12 and (B) IFN-γ. (C) Splenocytes from uninfected and day 5 infected WT and T-bet^−/− mice were analyzed by flow cytometry for mature (DX5⁺) NK cells (NK1.1⁺CD3⁻ live cells). (D) NK cells (NK1.1⁺CD3⁻ live cells) from day 5 infected WT and T-bet^−/− mice were analyzed by flow cytometry for expression of CD69 (WT CD69 MFI=1246±89.41, T-bet^−/− CD69 MFI=1086±24.83). (E, F) Whole splenocytes from uninfected and day 5 infected WT and T-bet^−/− mice were restimulated in vitro with PMA and ionomycin in the presence of Brefeldin A and monensin for 4 hours. Cells were then fixed and stained for cytokine production by NK cells (E) and quantified (F). (G, H) T-bet^−/− mice were infected i.p. with T. gondii. On days 0, 3, and 6, mice were given 0.5mg αIFN-γ or isotype control (αIgG) i.p. On day 7 post-infection peritoneal lavage was performed and PECs were collected for cytospin preparation and analysis of percentage of cells infected. (I) T-bet^−/− mice were treated with 50ul αAsialo gm-1 or isotype control on days −1, 3, and 6 of T. gondii infection. At day 9 post-infection, peritoneal lavage was performed and peritoneal exudate cells (PECs) were collected for cytospin preparation and analysis of parasite burden. Data are representative of 2–3 experiments, n=3 per experiment. * p<0.05

Role of T-bet in the T cell response to T. gondii

In order to assess the role of T-bet in the generation of the adaptive T cell response to T. gondii, WT and T-bet^−/− mice were infected with T. gondii and responses were analyzed 9–11 days later. Splenocytes from infected and uninfected mice were harvested and stimulated in vitro with either anti-CD3 or with soluble T. gondii antigen (STAg) and assessed for IFN-γ production (Figures 5a and 5b). Intracellular staining revealed that among WT cells, the majority of CD4⁺ T cells producing IFN-γ were T-bet⁺ (Figure 5c). While infection induced an increase in IFN-γ production by both WT and T-bet^−/− CD4⁺ T cells (13-fold increase in the WT and 10-fold increase in the T-bet^−/−), the percentage of IFN-γ-producing CD4⁺ T cells was reduced in the T-bet^−/− mice compared to the WT mice (Figure 5c), but this was not associated with increases in IL-4 (data not shown). Similarly, the majority of WT CD8⁺ T cells producing IFN-γ also expressed T-bet; however, WT and T-bet^−/− CD8⁺ T cells produced comparable levels of IFN-γ (Figure 5d). Thus, there is a significant proportion of CD4⁺ and CD8⁺ T cells that produce IFN-γ independently of T-bet.

Figure 5. T-bet-deficient T cells produce IFN-γ during infection with T. gondii.

WT and T-bet^−/− mice were infected i.p. with T. gondii. Splenocytes were restimulated in vitro with (A) αCD3 or (B) STAg and supernatants were assayed for IFN-γ by ELISA. (C, D) Splenocytes from uninfected and infected WT and T-bet^−/− mice were restimulated in vitro stained for IFN-γ production by (C) CD4⁺ T cells (CD3⁺CD4⁺CD8a⁻ live cells) and (D) CD8⁺ T cells (CD3⁺CD4⁻CD8a⁺ live cells). Data are representative of 3 experiments, n=3–4 per experiment. ** p<0.01

To characterize the impact of T-bet on T cell activation, a panel of markers that included CD25, CD11a, Ly6C, KLRG-1 and CXCR3 was used to compare the T cell populations in WT and T-bet^−/− mice. This analysis revealed that following challenge WT CD4⁺ T cells had high levels of CD11a and this was decreased in the T-bet^−/− CD4⁺ T cells (Figure 6a). Interestingly, although there was no difference in CD25 expression between naïve WT and T-bet^−/− mice (data not shown), T-bet^−/− CD4⁺ T cells from infected mice displayed higher levels of CD25 than CD4⁺ T cells from WT mice (Figure 6a). However, there was no difference in the frequency or numbers of Foxp3 expressing cells between the WT and T-bet^−/− CD4⁺ T cell populations (data not shown). In the CD8⁺ T cell population, there were CD11a^hi and CD11a^lo populations of WT CD8⁺ T cells, and this heterogeneity was less apparent in the T-bet^−/− CD8⁺ T cells (Figure 6b). Moreover, similar to the CD4⁺ T cells in the T-bet^−/− mice, CD8⁺ T cells expressed higher levels of CD25 than WT CD8⁺ T cells (Figure 6b). Initially, these data on CD11a expression suggested that in infected T-bet^−/− mice there are fewer parasite-specific CD4⁺ T cells. However, the use of a T. gondii-specific AS15 MHC-II tetramer (35) revealed that in the spleens of WT and T-bet^−/− mice there was an equivalent expansion of the these CD4⁺ T cells (Figure 6c). Additionally, there were no differences in PSGL-1, CD44 or CD62L expression between the WT and T-bet^−/− tetramer⁺ CD4⁺ T cells (data not shown). Nonetheless, there were phenotypic differences between the WT and T-bet^−/− cells that mirrored the polyclonal population. Thus, the WT AS15-specific CD4⁺ T cells expressed high levels of the activation markers CD11a, Ly6C, and CXCR3, but these were reduced in parasite-specific CD4⁺ T cells from the T-bet^−/− mice (Figure 6d). In addition, while these T cells did produce IFN-γ, the MFI was reduced compared to WT T cells (Figure 6d).

Figure 6. Altered phenotype of parasite-specific T cells in infected T-bet^−/− mice.

WT and T-bet^−/− mice were infected i.p. with T. gondii. (A, B) Splenocytes from infected mice were harvested and stained for analysis by flow cytometry for CD11a and CD25 expression levels on (A) CD4⁺ and (B) CD8⁺ T cells. (C, D) Splenocytes from infected or uninfected mice were stained for parasite-specific CD4⁺ T cells using an MHC II tetramer and analyzed for (C) quantification, and (D) activation and effector status markers CD11a, Ly6C, CXCR3, and IFN-γ by flow cytometry. In the IFN-γ flow plots, the IFN-γ FMO is light grey (E, F) Splenocytes from infected or uninfected mice were stained for parasite-specific CD8⁺ T cells using an MHC I tetramer and this population was (E) quantified and analyzed for (F) activation and effector status markers CD11a, KLRG-1, CXCR3, and IFN-γ by flow cytometry. In the IFN-γ flow plots, the IFN-γ FMO is light grey. Data are representative of 3–5 experiments, n=3–4 per experiment.

In naïve WT and T-bet^−/− mice, the frequency of splenic Tgd057-specific CD8⁺ T cells is low and infection with T. gondii results in an equivalent expansion of this population (Figure 6e), which expressed similar levels of CD11a (Figure 6f). Additionally, there were no differences in CD44 or CD62L expression between the WT and T-bet^−/− Tetramer⁺ CD8⁺ T cells (data not shown). However, while the majority of Tgd057-specific WT CD8⁺ T cells expressed KLRG-1 and CXCR3, the parasite-specific CD8⁺ T cells from T-bet^−/− mice expressed significantly lower levels of these proteins (Figure 6f), but the WT and T-bet^−/− parasite-specific CD8⁺ T cells expressed comparable levels of IFN-γ (Figure 6f). These data indicate that T-bet is not required for the activation and expansion of parasite-specific CD4⁺ and CD8⁺ T cells but is required for these cells to acquire an effector phenotype associated with expression of CD11a, CXCR3, Ly6C and KLRG-1.

Role of T-bet in T cell responses in peripheral sites

Given that the T-bet^−/− mice had an increased parasite burden in peripheral tissues, the decreased expression of CXCR3 and CD11a by parasite-specific T cells from the spleens of these mice suggested a defect in their ability to either access distal sites of infection or function within these sites to control parasite replication. Therefore, WT and T-bet^−/− mice were infected and the T cell responses were assessed at peripheral sites. In the lungs of WT mice, there was a large population of parasite-specific CD4⁺ T cells characterized by high levels of Ly6C. In the T-bet^−/− mice, the numbers of these cells were markedly reduced and those that were present expressed reduced levels of Ly6C (Figure 7a, b). Similarly, there were significantly more parasite-specific CD8⁺ T cells in the lungs of WT mice than in the T-bet^−/− mice (Figure 7c), and the WT cells expressed higher levels of KLRG-1 (Figure 7d). Immunohistochemical analysis of brain sections taken at day 9 post-infection showed T cells associated with areas of parasite replication in WT mice (Figure 7e), but in T-bet^−/− mice there was a marked absence of T cells despite the increased parasite burden (Figure 7e). Thus, the susceptibility of the T-bet^−/− mice is not due to a defect in the numbers of parasite-specific T cells generated, but rather is associated with an altered phenotype and reduced numbers of effector T cells at local sites of parasite replication.

Figure 7. T-bet is required for T cell localization at secondary sites of parasite colonization.

WT and T-bet^−/− mice were infected i.p. with T. gondii. (A, B) Lungs were harvested and cells were isolated and stained for parasite-specific CD4⁺ T cells using an MHC II tetramer and Ly6C expression was assessed on these cells. (C, D) Lung cells were also stained for parasite-specific CD8⁺ T cells using an MHC I tetramer and KLRG-1 expression was assessed on these cells. (E) Brains were harvested from WT and T-bet^−/− mice and stained for T cells (Cy7, red), T. gondii (FITC, green), and DAPI (blue). Data are representative of 2–3 experiments, n=3 per experiment.

Cell intrinsic versus extrinsic requirement for T-bet

Altered CD4⁺ T cell activity during toxoplasmosis leads to changes in the CD8⁺ T cell response (46, 47) and the studies described above do not distinguish the cell intrinsic versus cell extrinsic effects of T-bet deficiency on the development and function of parasite-specific T cells. In order to start to address this issue, a competitive mixed-bone marrow chimera approach was used in which irradiated recipient mice received a 1:1 ratio of WT:T-bet^−/− bone marrow (Figure 8a). After >8 weeks post-reconstitution, the mice were sacrificed and tissues were examined to determine levels of chimerism. Similar to previous reports (7), the majority of the reconstituted cells were derived from the T-bet^−/− donor. At steady state in all four experiments performed, cells from the T-bet^−/− donor comprised 65–75% of the total CD4⁺ T cells in the spleen as well as the lung (Figure 8b), and this ratio of WT:T-bet^−/− CD4⁺ T cells was maintained for more than 6 months post-reconstitution (data not shown). These observations indicate that T-bet^−/− CD4⁺ T cells have a competitive advantage over WT CD4⁺ T cells in this experimental setting. In contrast, the CD8⁺ T cell compartment had equal proportions of WT and T-bet^−/− cells in the spleen, and an increased frequency of WT CD8⁺ T cells in the lung (Figure 8c).

Figure 8. T-bet is intrinsically required for T cell localization at secondary sites of parasite colonization.

(A–H) Bone marrow was harvested from WT and T-bet^−/− mice, and then transferred via i.v. injection into mice irradiated (1000rads). (B, C) >8 weeks post-transfer, cells from the spleen and lung were analyzed for reconstitution in the (B) CD4⁺ T cell compartment, and the (C) CD8⁺ T cell compartment. (E–J) >8 weeks post-transfer, the mice were infected with T. gondii i.p. and 9–11 days post-infection cells from the spleen and lung were (E) analyzed for numbers of parasite-specific (Tet⁺) CD4⁺ T cell and replicates were concatenated and (F) assessed for expression of Ly6C on WT and T-bet^−/− Tet⁺CD4⁺ T cells. (G) IFN-γ production was assessed in WT and T-bet^−/− CD4⁺ T cells. (H, I) The parasite-specific (Tet⁺) CD8⁺ T cell compartment was (H) quantified, and concatenated samples were (I) assessed for KLRG-1 expression on WT and T-bet^−/− Tet⁺CD8⁺ T cells. (J) IFN-γ production was assessed in WT and T-bet^−/− CD8⁺ T cells. Data are representative of 4 experiments, n=4–5 per experiment.

When the chimeras were infected with T. gondii and analyzed 9–11 days later (Figure 8d), there was a 70-fold expansion of the WT parasite-specific CD4⁺ T cells but only a 6-fold expansion of the T-bet^−/− parasite-specific CD4⁺ T cells (data not shown), resulting in no difference in the number of WT and T-bet^−/− parasite-specific CD4⁺ T cells in the spleen or lung after infection (Figure 8e). However, the WT parasite-specific CD4⁺ T cells from both the spleen and lung expressed higher levels of Ly6C (Figure 8f) and CD11a (data not shown) than T-bet^−/− parasite-specific CD4⁺ T cells after infection. Additionally, there was reduced IFN-γ from the T-bet^−/− CD4⁺ T cell compartment compared to the WT CD4⁺ T cells (Figure 8g). In contrast, there were significantly more WT parasite-specific CD8⁺ T cells in both the spleen and lung after challenge (Figure 8h), and these WT parasite-specific CD8⁺ T cells from the spleen and lung expressed higher levels of KLRG-1 (Figure 8i) and CD11a (data not shown) than the T-bet^−/− parasite-specific CD8⁺ T cells. Surprisingly, there was more IFN-γ production from the WT CD8⁺ T cells than from the T-bet^−/− CD8⁺ T cells (Figure 8j). These data indicate that following challenge with T. gondii, there is a cell-intrinsic requirement for T-bet for the activation and acquisition of effector status of the CD4⁺ and CD8⁺ T cell populations, and these effector populations are required for control of the parasite in peripheral tissues.

Discussion

The experiments presented here reveal that during experimental toxoplasmosis T-bet is not required for the innate NK cell dependent mechanism of resistance nor for the development and expansion of parasite-specific CD4⁺ and CD8⁺ T cells. Indeed, the ability of NK and CD8⁺ T cells to produce IFN-γ appears intact. However, T-bet is required for optimal IFN-γ production by CD4⁺ T cells. It has been reported that in response to restimulation with αCD3 and αCD28, a significant proportion of T-bet^−/− CD4⁺ T cells from mice infected with T. gondii produced IFN-γ, and a smaller subset produced IL-4 (22). In the studies presented here, the parasite-specific T-bet^−/− CD4⁺ T cell restimulated with STAg also produce IFN-γ, but not IL-4, and thus appear to be Th1 cells (48). Because the loss of CD4⁺ T cells alone does not lead to acute susceptibility to T. gondii (49, 50), this partial defect in CD4⁺ T cell IFN-γ production is not sufficient to explain the susceptibility of the T-bet^−/− mice challenged with T. gondii. Rather, the increased parasite numbers at many secondary sites of parasite dissemination was associated with reduced numbers of parasite-specific effector T cells. However, in the spleen, despite the presence of parasite specific effector T cells, there was still a marked increase in parasite burden. These latter results have to be interpreted with care as the parasite burden detected in the spleen also reflects the numbers of T. gondii in the blood, but suggest that the reduced expression of CD11a, CXCR3, Ly6C and KLRG1 may also compromise the function of these parasite-specific T cells. Thus, these data highlight that additional functions of T-bet are required for the ability of effector T cells to access and operate within local sites of infection. To the best of our knowledge, this is the first report that, in the context of infectious disease, highlights this key role for T-bet in coordinating multiple facets of the effector response and localization to peripheral sites and may help to explain the basis for the susceptibility of T-bet^−/− mice in other infectious settings (2, 13–15, 51).

A common feature of the acute phase of many intracellular pathogens is the presence of an IFN-γ dependent, NK cell mediated mechanism of resistance (52–54). Because T-bet controls key checkpoints in NK maturation (10) and has been linked to the ability of these cells to produce IFN-γ (2, 15) it seemed likely that the NK response during toxoplasmosis would be T-bet dependent. However, others have reported that T-bet is not required for the production of IFN-γ from these cells (8, 10) and following challenge of the T-bet^−/− mice with T. gondii, NK cell IFN-γ production appeared normal. Similarly, when T-bet^−/− mice are infected with L. monocytogenes, the early NK cell response is intact and is associated with acute resistance to the bacteria (16) but the long-term consequences of T-bet deficiency on the T cell responses to this bacterium remain unclear. Similarly, T-bet^−/− mice survive for more than 2 weeks after challenge with Salmonella (13) indicating that early IFN-γ production in this model is intact. These results from multiple experimental systems are consistent with a model in which high levels of inflammation associated with many parasitic and bacterial infections provide sufficient signals to overcome the requirement for T-bet in NK cell maturation.

Previous in vitro studies have identified a role for T-bet in T cell chemotaxis through its induction of CXCR3 (12), while more recent work has implicated T-bet in trafficking of Tregs in vivo (55). The findings presented here that T-bet is required in vivo for the ability of parasite-specific T cells to express CXCR3 and the integrin CD11a (a component of LFA-1) reinforce this notion. CXCR3 and/or its ligands have a key role in the trafficking of effector T cells necessary to control of number of pathogens including T. gondii, Respiratory Syncytial Virus, and influenza (56–59). Thus, the reduced number of effector T cells at peripheral sites of infection in the T-bet^−/− mice could be a result of defective T cell trafficking to these sites. However, this alteration in trafficking is not always detrimental; during toxoplasmosis, immune-mediated pathology in the liver is caused by CD4⁺ effector T cells (50, 60, 61), and in our studies T-bet^−/− mice have reduced levels of liver pathology. Likewise, the loss of T-bet or CXCR3 provides protection against experimental cerebral malaria, associated with a reduced number of T cells in the brains of infected mice (62, 63), and T-bet-deficiency confers protection in a murine model of Type 1 diabetes that is associated with reduced islet infiltration by T cells (7). These findings from diverse infectious and autoimmune models highlight that the ability to target T-bet (64) may also influence T cell trafficking and limit inflammation. Interestingly, T-bet was not restricted to the nucleus in highly activated cells. Although the function of T-bet as a cytoplasmic protein remains unknown, differential T-bet localization has been linked to effector status (37), activation (65), cell cycle (66), and protein stability (67). The biological significance of the cytoplasmic localization of T-bet, the mechanisms that underlie this partitioning, and whether localization influences T cell phenotype and/or migration remain unclear but may provide opportunities to target different functions of this transcription factor.

Although the studies presented here highlight the impact of T-bet on expression of CD11a and CXCR3 and their links to T cell trafficking, these molecules are involved in many facets of T cell activation, which may contribute to the defect in effector populations. For example, CD11a is upregulated on T cells after TCR signaling and its expression has been used as a marker of activation for infection-induced polyclonal T cell populations (31–33). Consequently, the reduced levels of CD11a expression on T cells from infected T-bet^−/− mice initially suggested that T-bet was required for their ability to generate parasite-specific populations. However, the use of tetramers demonstrated that the generation of parasite-specific CD4⁺ and CD8⁺ T cells was intact in the T-bet^−/− mice and implied that upregulation of this integrin was, in part, dependent on T-bet. Thus, because CD11a is important during priming for the generation of antigen-specific effector CD8⁺ T cells (68, 69), upregulation of CD11a is a potential mechanism by which T-bet influences the phenotype of the effector population. Recent studies have also shown that CXCR3 is important for T cell differentiation and behavior (70, 71), therefore, the combination of decreased CD11a and CXCR3 expression on T-bet^−/− T cells could impact the differentiation and function of these populations. Indeed, the T-bet^−/− T cells from infected mice display altered effector phenotypes, with a decrease in the Ly6C^hi CD4⁺ and KLRG-1^hi CD8⁺ effector T cells. While there was a decrease in the numbers of KLRG-1^hi effector CD8⁺ T cells in the T-bet^−/− mice, it should be noted that the KLRG-1^lo CD8⁺ memory precursor T cell population was intact in the absence of T-bet. Similarly, during LCMV infection, memory-precursor CD8⁺ T cells display lower T-bet expression (28, 72), and T-bet deficiency results in enhanced generation of memory CD8⁺ T cells (72), but the ability of this T-bet^−/− population to protect against secondary challenge is unclear (6, 72). Thus, while the studies described here help to provide a better understanding of the role of T-bet in resistance to T. gondii, they also highlight the need to investigate the role of T-bet in T cell priming and the generation protective memory responses.