The JI
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     
 

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Egan, C. E.
Right arrow Articles by Carding, S. R.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Egan, C. E.
Right arrow Articles by Carding, S. R.
The Journal of Immunology, 2005, 175: 8191-8199.
Copyright © 2005 by The American Association of Immunologists

A Requirement for the V{gamma}1+ Subset of Peripheral {gamma}{delta} T Cells in the Control of the Systemic Growth of Toxoplasma gondii and Infection-Induced Pathology1

Charlotte E. Egan2,*, Jane E. Dalton*, Elizabeth M. Andrew*, Judith E. Smith{dagger}, Marc-Jan Gubbels3,{ddagger}, Boris Striepen{ddagger} and Simon R. Carding4,*

* Research Institute for Molecular and Cellular Biology and {dagger} Research Institute for Integrative and Comparative Biology, University of Leeds, Leeds, United Kingdom; and {ddagger} Center for Tropical and Emerging Global Diseases and Department of Cellular Biology, University of Georgia, Athens, GA 30602


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
{gamma}{delta} T cells are a diverse population of T cells that are widely distributed and are a common feature of pathogen-induced immune responses. It is not clear, however, whether different populations of {gamma}{delta} T cells have specific functions, and what factors determine the functional properties of individual populations. A murine model of peroral Toxoplasma gondii infection was used to determine the contribution V{gamma}1+ intestinal intraepithelial lymphocytes (IELs) vs systemic V{gamma}1+ T cells make to the acute and chronic stages of the host immune response, and whether the macrophage cytocidal activity of V{gamma}1+ T cells described in bacterial infections is seen in other, unrelated infectious disease models. In response to oral infection with virulent type 1 or avirulent type II strains of T. gondii, TCR-{delta}–/– mice rapidly developed severe ileitis. In contrast, in mice deficient in V{gamma}1+ T cells and IELs and wild-type mice, inflammation was delayed in onset and less severe. The protective effect of (V{gamma}1–) IELs to Toxoplasma infection was unrelated to their cytolytic and cytokine (Th1)-producing capabilities. Systemic V{gamma}1+ T cells were shown to play an essential role in limiting parasite growth and inflammation in peripheral tissues and, in particular, in the CNS, that was associated with their ability to efficiently kill parasite-elicited and infected macrophages. These findings suggest that macrophage cytocidal activity of V{gamma}1+ T cells may be a universal feature of pathogen-induced immune responses and that microenvironmental factors influence the involvement and function of {gamma}{delta} T cells in the host response to infection.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Although {gamma}{delta} T cells are relatively rare in lymphoid tissues, they are well represented in epithelia and in particular in the intestinal epithelium, where they can comprise the largest population of intraepithelial lymphocytes (IELs)5 (1, 2). The immunoregulatory properties of TCR-{gamma}{delta}+ IELs (3, 4) and their involvement in epithelial homeostasis (5, 6, 7) are consistent with their providing a first line of defense against infection (2, 8). Systemic populations of {gamma}{delta} T cells in contrast can contribute to both the early and the later stages of infection. The observation that pathogen infection of {gamma}{delta} T cell-deficient mice results in an exaggerated immune response, chronic inflammation, and tissue injury (9, 10, 11, 12, 13, 14, 15, 16, 17) suggests that systemic {gamma}{delta} T cells are of particular importance in controlling or limiting inflammatory responses and resolving pathogen-induced immune responses. The finding that {gamma}{delta} T cells play a role in macrophage homeostasis, especially in the elimination of activated pathogen-elicited macrophages (18, 19), and can influence {alpha}{beta} T cell responses (11, 20) is consistent with this interpretation. It is not clear, however, what the relationship is between {gamma}{delta} T cells that are active at different stages of pathogen-induced immune responses, whether different populations of {gamma}{delta} T cells have specific functions, and what factors determine the functional properties of individual populations. Two possibilities are that {gamma}{delta} T cell function is predetermined and that different populations have restricted or defined functions, or that {gamma}{delta} T cells are functionally heterogeneous with their response being shaped by environmental factors.

Based upon the functional analysis of individual subsets of {gamma}{delta} T cells in murine models of Coxsackievirus B3 infection and airway hypersensitivity, it has been proposed that {gamma}{delta} T cell function cosegregates with TCR variable (V{gamma}) gene usage (21, 22, 23, 24). In the Coxsackievirus infection model, the cytokines produced by different populations of {gamma}{delta} T cells have a significant influence on the CD4-Th cell phenotype with V{gamma}1+ (IL-4+) T cells being able to suppress and V{gamma}4+ (IFN-{gamma}+) T cells promote susceptibility to the development of virus-induced myocarditis (21, 22). In airway hypersensitivity, in contrast, V{gamma}1+ T cells can promote airway inflammation, whereas V{gamma}4+ (IFN-{gamma}+) pulmonary T cells can negatively regulate airway responsiveness (24). Our own studies of {gamma}{delta} T cell function in the murine model of listeriosis, however, have shown that certain populations of {gamma}{delta} T cell are functionally heterogeneous and that their response is not predetermined, but is instead influenced by environmental factors (25). Based upon cytokine profiling and TCR-CDR3 V{gamma}/V{delta} sequence analyses of Listeria-elicited V{gamma}1+ T cells, it was shown that the populations responding during the early and late stages of the immune response are distinct and nonoverlapping. The question of how {gamma}{delta} T cell responses are determined is, therefore, unresolved. In considering how to redress these contrasting findings from the different experimental model systems, it is noteworthy that the evidence for functional plasticity of {gamma}{delta} (V{gamma}1+) T cells is based upon the analysis of a single infectious disease model. To directly address this issue and determine the universality of the proposed function of V{gamma}1+ T cells, we have used a murine model of peroral Toxoplasma gondii infection. This model of infection is particularly well suited to assessing the impact TCR-V{gamma} gene usage vs tissue environmental influences have on V{gamma}1+ T cell responses because the contribution V{gamma}1+ T cells in the intestine (IELs) vs peripheral tissues make to protecting the host from infection can be assessed.

Toxoplasma infection is perhaps the most prevalent parasitic infection of mankind in the world (26) and can be divided into two temporally distinct phases, acute and chronic (27, 28, 29). The acute phase is characterized by widespread dissemination of rapidly dividing tachyzoites that can invade virtually any cell type. Chronic infection is associated with differentiation of tachyzoites into bradyzoites that form quiescent cysts within the brain and skeletal muscle. The generation of long-lived protection to Toxoplasma infection is dependent upon the development of a strongly polarized, IL-12-dependent, Th1-type response that is characterized by the production of IFN-{gamma} by NK and CD4+/CD8+ T cells (30, 31, 32, 33, 34). The failure to appropriately regulate the innate or adaptive response can lead to severe immunopathology, which is seen in susceptible inbred strains of mice that develop acute and lethal ileitis after oral infection (35, 36, 37). Circumstantial evidence suggests that {gamma}{delta}+ IELs may contribute to protection against Toxoplasma infection. The ability of adoptively transferred IELs to confer long-term protection against an otherwise lethal parasite challenge has been shown to be at least partially dependent upon the presence of TCR-{gamma}{delta}+ IELs in the host (38). In addition, {gamma}{delta} IELs may contribute to the local production of TGF-{beta} (3, 4) that is required for regulating T. gondii-induced ileitis (39). However, it is not clear which population(s) of {gamma}{delta} IELs might contribute to host protection. The evidence for the involvement of systemic {gamma}{delta} T cells in the host response to Toxoplasma infection once it has invaded the epithelium is more compelling. {gamma}{delta} T cells expand in response to T. gondii infection in genetically dissimilar strains of mice (40, 41, 42), rats (43), and humans (44, 45, 46, 47), the timing of which in rodents is dependent upon the strain and dose of Toxoplasma used and the route of administration. The increased susceptibility and mortality of {gamma}{delta} T cell-depleted mice to infection (40, 41), and that the adoptive transfer of cytolytic and IFN-{gamma}-producing {gamma}{delta} T cells prolongs survival against acute peroral parasite challenge (41) are consistent with a requirement for {gamma}{delta} T cells in immunity to Toxoplasma infection. Other studies, however, have found little or no evidence of a protective role for {gamma}{delta} T cells (38, 48). It is not clear, therefore, what role {gamma}{delta} T cells and the V{gamma}1 subset might play in the host response to Toxoplasma infection.

In this study, we show that whereas V{gamma}1+ IELs do not make a significant contribution to limiting parasite-induced inflammation and tissue injury in the intestine, systemic V{gamma}1+ T cells play a central role in limiting parasite growth and inflammation in peripheral tissues and, in particular, in the CNS, a property associated with the ability of V{gamma}1+ T cells to eliminate parasite-elicited and infected macrophages.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Mice and infection

Six- to 8-wk-old male C57BL/6 mice were purchased from Harlan Laboratories. C57BL/6 TCR-{delta}–/– and TCR-{beta}–/– mice were purchased from The Jackson Laboratory and bred in our facility. TCR-V{gamma}1–/– mice were generated in our laboratory (25) and backcrossed onto the C57BL/6 background for at least six generations. All mice were housed in specific pathogen-free conditions at Leeds University. Tachyzoites of the type I RH strain of T. gondii-expressing yellow fluorescent protein (YFP) (49) were maintained by serial passage through confluent monolayers of 3T3 fibroblasts in DMEM supplemented with 2 mM L-glutamine, 10% heat-inactivated FBS, 5 U/ml penicillin, and 10 g/ml streptomycin and incubated at 37°C, 5% CO2. All reagents were from Sigma-Aldrich. The type II ME-49 strain was maintained by passage in outbred, CD1 mice (Charles River Laboratories). Cysts were isolated from chronically infected brains by discontinuous Percoll (Amersham Biosciences) gradient centrifugation of brain homogenates. Bradyzoites were recovered from the cysts by digestion with an acid pepsin solution (50). Parasite numbers were determined by phase-contrast microscopy and diluted to a working concentration in PBS. Eight-week-old male mice were infected by oral gavage with 3000 ME-49 bradyzoites (equivalent of 6–10 cysts), or 106 YFP-RH tachyzoites. All animal experiments were conducted at the University of Leeds and in full accordance with the Animals Scientific Procedures Act 1986 under Home Office approval.

Cell isolations and flow cytometry

Spleen, liver, and mesenteric lymph node (MLN) were homogenized; contaminating erythrocytes were lysed with 0.84% (w/v) ammonium chloride solution; and the cell suspension was passed through 0.7-µm nylon filter and washed before Ab staining. Small intestinal IELs were isolated from Peyer’s patch-excised small intestines, as described previously (51). Abs used for surface staining included: F(ab')2 of mAbs specific for TCR-V{gamma}1 (clone 2.11) (52) and TCR-{delta} (GL3). The 2.11 hybridoma cell line was provided by P. Pereira (Institut Pasteur, Paris, France). Commercial anti-mouse mAbs used were anti-CD3 (145 2C11), anti-TCR-{gamma}{delta} (GL3), anti-TCR-{alpha}{beta} (H57-597), anti-B220 (RA3-6B2), anti-Gr-1, and anti-F4/80 conjugated to FITC, PE, or PE-Cy5.5 purchased from Caltag-Medsystems or BD Pharmingen. Streptavidin conjugates of PE, FITC (Caltag Laboratories), or Alexa Fluor 633 (Molecular Probes) were used as secondary reagents. To block nonspecific Ab binding, cells were preincubated with anti-FcR Ab mixture, anti-CD16/32 (20µg/ml) (Caltag Laboratories). Isotype-matched Abs of irrelevant specificity were used to determine the level of nonspecific staining. Stained cells were analyzed on a FACSCalibur flow cytometer using CellQuest software (BD Pharmingen).

Assessment of tissue injury

For histology, tissues were fixed overnight in phosphate-buffered 10% Formalin (Sigma-Aldrich) and embedded in paraffin. Five-micrometer sections were cut using a microtome, mounted on SuperFrost Plus microscope slides (BDH), and either stained directly with H&E or with biotinylated anti-active caspase 3 (BD Pharmingen), followed by avidin-biotin-HRP complexes (ARC amplification system; DakoCytomation) and diaminobenzidine substrate before counterstaining with hematoxylin. As an independent evaluation of liver injury serum, alanine aminotransferase (ALT) levels were measured on an ADVIA 2400 chemistry system analyzer (Bayer) following the manufacturer’s instructions.

Macrophage-T cell coculture

Plastic-adherent peritoneal exudate cells, spleen, or liver mononuclear cells (5 x 105) from day 10 ME-49-infected TCR-{delta}–/– mice were incubated with 1 x 106 nonadherent splenocytes from noninfected wild-type, TCR-{delta}–/–, TCR-{beta}–/–, or TCR-V{gamma}1–/– mice in 8-well chamber slides (Valeant Pharmaceuticals) and cultured at 37°C in RPMI 1640 (Sigma-Aldrich) with 10% FBS for 1 h, as previously described (53). For some experiments, {gamma}{delta} T cell-enriched preparations of splenocytes (>80% TCR-{gamma}{delta}+) prepared by immunomagnetic selection (25) from wild-type mice were used as effector cells. The extent of macrophage killing was determined by labeling target cells with the Live/Dead cell reagent (Molecular Probes) that contains fluorescent dyes that identify intracellular esterase activity of viable cells (3,3'-dioctadecyloxacarbocyanine) or that are incorporated in the nuclei of dead cells (propidium iodide). At least 100 live and/or dead cells were counted in four separate fields by UV microscopy.

Cytokine synthesis by {gamma}{delta} T cells

Intracellular cytokines were detected by cytoplasmic staining of IELs and splenocytes recovered from parasite-infected wild-type mice after culture in vitro for 5 h with 10 mg/ml brefeldin A (Sigma-Aldrich). Cells were then surface stained with anti-CD3, anti-TCR-{gamma}{delta}, and anti-TCR-V{gamma}1, or anti-TCR-V{gamma}7 mAbs, fixed in 1% paraformaldehyde, and permeabilized with 0.5% of saponin (Sigma-Aldrich) before cytoplasmic staining with PE-conjugated anti-mouse cytokine mAbs to IL-2, IL-4, IL-5, IL-6, IL-10, IFN-{gamma}, and TNF-{alpha} (Caltag-Medsystems or BD Pharmingen), or FITC-conjugated polyclonal Abs to MIP-1{beta} and MCP-1 (Sigma-Aldrich). Anti-TGF-{beta} and anti-latency-associated peptide (LAP) mAbs (R&D Systems) were conjugated to FITC and used for surface staining. PE- and FITC-conjugated isotype-matched mAbs of irrelevant specificity were used to determine levels of nonspecific staining of anti-cytokine mAbs. Serum cytokine levels were determined using flow cytometric cytokine bead arrays (BD Pharmingen), according to the manufacturer’s instructions.

Statistical analysis

All data were assessed for normal distribution using a Shapiro-Wilk test. For parametric data analysis was performed using Student t test, and for nonparametric data analysis was performed using Mann-Whitney U tests using the Statistical Package for the Social Sciences software, SPSS. Statistical significance was taken as p < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
The role of V{gamma}1+ T cells resident in the small intestine and peripheral lymphoid tissues in the host response to T. gondii infection was investigated by comparing the outcome of infecting wild-type (C57BL/6) mice and strains genetically deficient of all {gamma}{delta} T cells (TCR-{delta}–/–) or of the V{gamma}1+ T cell subset (V{gamma}1–/–) (25) with virulent type I tachyzoites or avirulent type II bradyzoites of T. gondii. For the analysis of the involvement of {gamma}{delta} IELs in the acute stage of Toxoplasma infection, a recombinant form of the RH strain of tachyzoites that express a highly fluorescent reporter gene (tandem YFP) (49) was used. Because YFP-reporter gene expression was lost during tachyzoite to bradyzoite stage conversion and RH is a noncyst-forming strain, it was not possible to use YFP-RH to study the chronic stage of Toxoplasma infection. Instead, the avirulent and cyst-forming type II ME-49 strain was used to study the chronic stage (2–6 wk postinfection) of T. gondii infection. In a series of preliminary experiments, an infectious dose of 106 tachyzoites and 3000 bradyzoites (equivalent to 5–10 cysts) was shown to produce a productive, but nonlethal acute and a chronic infection, respectively, in TCR-{delta}–/– and V{gamma}1–/– mice, and these doses were used in all subsequent experiments described in this study.

Accelerated onset of parasite-induced ileitis in the absence of {gamma}{delta} IELs

Both the onset and severity of T. gondii-induced ileitis were accelerated and more severe in TCR-{delta}–/– mice infected with RH tachyzoites compared with wild-type mice (Table I and Fig. 1A). The first signs of inflammation were evident in TCR-{delta}–/– mice 4 days postinfection (Table I), some 3 days before the involvement of {alpha}{beta} T cells (35). Pathology was characterized by the presence of small distinct foci of infiltrating polymorphonuclear cells (PMNC) and villus edema and blunting. This was in contrast to the virtually normal histology of the intestine of wild-type and TCR-V{gamma}1–/– mice (Table I). In contrast, TCR-{beta}–/– were more susceptible to RH infection, with the majority (8 of 10) developing a lethal ileitis within 7 days postinfection (Table I), consistent with the requirement for {alpha}{beta} T cells (Th1) for protection from Toxoplasma infection (30, 31, 32, 33, 34).


View this table:
[in this window]
[in a new window]
  		Table I. Histopathology of ileum and liver of wild-type and transgenic mice after peroral infection with T. gondii

 


View larger version (61K):
[in this window]
[in a new window]
  		FIGURE 1. Accelerated onset of parasite-induced ileitis in {gamma}{delta} T cell-deficient mice. A, Groups of C57BL/6 wild-type (WT) and {gamma}{delta} T cell (TCR-{delta}–/–)- and V{gamma}1 T cell (TCR-V{gamma}1–/–)-deficient mice were infected orally with YFP-RH parasites, and 7 days later the ileum was removed and processed for histology. Representative sections from >10 mice of each strain are shown. The arrows identify inflammatory infiltrate in mucosa of TCR-{delta}–/– mice. Magnification, x60. B, The frequency of apoptotic cells in the intestinal mucosa of WT, TCR-{beta}–/–, TCR-{delta}–/–, and TCR-V{gamma}1–/– before (0) and 4 days after oral infection with YFP-RH was determined by immunohistochemical analysis of ileal section using anti-active caspase 3 Abs. The graph shows the average (±SEM) number of apoptotic epithelial cells per villi, as determined by counting 10–20 villi on each section and 4–5 sections per sample of at least 5 independently acquired samples for each mouse strain. C, The percentage of cytokine-synthesizing V{gamma}1+ and V{gamma}7+ IELs was determined by cell surface and intracellular staining of IELs directly ex vivo from day 7 RH-infected WT mice. The data shown represent the mean (±SEM) frequency of cells compiled from six mice.

 
By 7 days postinfection, inflammation was more extensive and severe in TCR-{delta}–/– mice (Fig. 1A and Table I), with the almost complete loss of normal epithelial architecture; hemorrhaging in the lamina propria; massive accumulation of inflammatory cells comprising monocytes, macrophages, and lymphocytes throughout the ileum; and extensive necrosis. Pathology was also evident at this time in TCR-V{gamma}1–/– mice, although it was less severe than that seen in TCR-{delta}–/– mice and was more comparable to that of wild-type mice, with fewer foci of inflammation and necrosis throughout the ileum and less extensive changes in villous architecture (Fig. 1A and Table I). The increased susceptibility of TCR-{delta}–/– mice to Toxoplasma infection was not parasite strain or stage specific because a similar outcome was seen after infection with ME-49 bradyzoites (Table I) and with the less virulent type II PRU strain of tachyzoites (J. Dalton and S. Carding, unpublished observations). Although PRU- and ME-49-mediated inflammation in TCR-{delta}–/– mice was as extensive as that of RH infection, it was less severe. Also, the onset of PRU- and ME-49-induced inflammation was delayed compared with RH infection and was first evident after 7–10 days in both wild-type and TCR-{delta}–/– (Table I, and data not shown), which most likely reflects the decreased tissue transmigratory and long distance migratory capabilities of type II strains of Toxoplasma (54). Compared with TCR-{delta}–/–-infected mice, TCR-V{gamma}1–/– mice resembled the wild type in being more resistant to ME-49 infection, with the first signs of ME-49-induced ileitis apparent only after 2 wk postinfection (Table I). Collectively, these findings demonstrate that compared with TCR-{delta}–/– mice, V{gamma}1–/– mice are more resistant to parasite-induced inflammation and tissue injury.

IEL responses have been associated with epithelial cell-directed cytotoxicity, the release of immunoregulatory cytokines/chemokines, and up-regulation of Th1 cytokines (reviewed in Ref.2). We, therefore, investigated whether these responses involved or required V{gamma}1+ IELs during the initial response to T. gondii infection in vivo. Staining of sections of ileum with Abs specific for active caspase 3, an early indicator and mediator of apoptotic cell death (55), identified very few, but equivalent numbers of apoptotic cells in the epithelial mucosa of wild-type, {gamma}{delta}-, or V{gamma}1+-deficient mice at 4 days postinfection, a time point coincident with the onset of ileitis in TCR-{delta}–/– mice (Fig. 1B). Analysis of cytokine production by {gamma}{delta} IELs during the development of ileitis (Fig. 1C) showed that the majority (>90%) did not synthesize any of the cytokines analyzed, including examples of those known to be constitutively expressed (e.g., TGF-{beta}) or up-regulated by IELs upon activation (e.g., IFN-{gamma}, TNF-{alpha}, and TGF-{beta}) (2). Among the small number of cytokine-producing V{gamma}1+ IELs, synthesis of IL-4, IL-6, and IL-10 was detected. Among V{gamma}7+ IELs, cytokine (primarily IL-6) production was restricted to a small subset (~10%) of cells (Fig. 1C).

Systemic V{gamma}1+ T cells contribute to restricting parasite growth and chronic inflammation

By 1 wk postinfection, the parasite burden in the majority of infected tissues, including the liver, peritoneum, and lungs of TCR-{delta}–/– and TCR-V{gamma}1–/– mice, was increased compared with wild-type mice (Fig. 2A, and data not shown). Within the spleen, the majority of parasites were intracellular within F4/80+ macrophages (Fig. 2A). This increase in parasite burden in TCR-{delta}–/– and TCR-V{gamma}1–/– mice occurred in parallel with the development of the peripheral {gamma}{delta} and V{gamma}1+ T cell response to Toxoplasma infection in wild-type mice (Fig. 2B). The {gamma}{delta} T cell response was dominated by V{gamma}1+ T cells that comprised 60–90% of all {gamma}{delta} T cells in the MLN, spleen, and liver at the time points analyzed. This pattern of the {gamma}{delta} and V{gamma}1+ T cell response was seen in both tachyzoite- and bradyzoite-induced infections (data not shown). {gamma}{delta} T cell expansion was first evident 7 days postinfection in both the lymphoid tissues draining the initial site of infection and in primary lymphoid organs, reaching maximum numbers at ~10 days postinfection, after which they declined (Fig. 2B).



View larger version (29K):
[in this window]
[in a new window]
  		FIGURE 2. V{gamma}1+ T cells contribute to restricting parasite growth and chronic inflammation. A, Seven days after oral infection of wild-type (WT), TCR-{delta}–/–, and TCR-V{gamma}1–/– mice with YFP-RH tachyzoites, spleens were removed, homogenized, and analyzed for the presence of fluorescent parasites by flow cytometry. The percentage values shown in each plot represent the proportion of electronically gated events that are parasites (circled region). The inset figure shows that parasites (YFP-RH, x-axis) reside primarily within macrophages (F4/80, y-axis) in the spleen at 7 days postinfection with YFP-RH. The percentage values shown represent the proportion of gated events within each quadrant; the upper right quadrant identifies the majority (>97%) of parasites within macrophages. The data shown are representative of that obtained from seven mice. B, Kinetics of the {gamma}{delta} and V{gamma}1+ T cell response in MLN, spleen, and liver of wild-type mice to ME-49 infection, as determined by harvesting tissue mononuclear cells before (0) and at regular intervals postinfection (PI) and staining with anti-CD3, anti-TCR-{gamma}{delta}, and anti-TCR-V{gamma}1 Abs and flow cytometric analysis. The data shown represent the mean values (±SEM) compiled from more than six mice per time point.

 
The effects of the absence of {gamma}{delta} and V{gamma}1+ T cells during the acute stages of Toxoplasma infection were also evident in the primary sites of parasite invasion, including the liver, lungs, and CNS after infection with either RH tachyzoites (Table I) or ME-49 bradyzoites (Fig. 3 and Table I). The onset and extent of inflammation were more rapid and severe after infection with virulent RH tachyzoites compared with avirulent ME-49 bradyzoites. By 4 days postinfection, liver pathology was first apparent in RH-infected TCR-{delta}–/– mice and was characterized by portal tract inflammation and the presence of PMNC and lymphocytes, and by 7 days postinfection had developed into severe inflammation with extensive necrosis (Table I). Liver pathology in infected TCR-{beta}–/– mice resembled that of TCR-{delta}–/– with evidence of inflammatory foci and lesions, although they were fewer in number and smaller compared with those seen in TCR-{delta}–/– mice (data not shown). Severe liver pathology and injury in TCR-{delta}–/– mice were also reflected in changes in serum levels of the liver enzyme, ALT, which at 7 days postinfection was significantly (p < 0.05) higher in TCR-{delta}–/– (333 ± 31 IU/L) compared with wild-type (174 ± 23 IU/L) and TCR-{beta}–/– (199 ± 14 IU/L) mice. The extent of liver pathology in T. gondii-infected TCR-V{gamma}1–/– mice was similar to that seen in TCR-{delta}–/– mice, although the onset of inflammation and necrosis was delayed. Up to 10 days postinfection, the liver histology in TCR-V{gamma}1–/– mice resembled that of wild-type mice, whereas by 14 days it was comparable to that of TCR-{delta}–/– mice (Fig. 3). Changes in serum ALT levels in TCR-V{gamma}1–/– mice (241 ± 34 IU/L at day 7, 284 ± 28 at day 10, and 379 ± 32 at day 14) also reflected the delay in the development of liver pathology. A similar pattern of pathology was seen in the lungs of T. gondii-infected mice, with the almost complete ablation of normal tissue architecture due to a massive infiltrate of inflammatory cells (primarily PMNC and lymphocytes) apparent in TCR-{delta}–/– mice by 7 days postinfection and by 10–14 days postinfection in TCR-V{gamma}1–/– mice (data not shown).



View larger version (109K):
[in this window]
[in a new window]
  		FIGURE 3. Exaggerated and chronic inflammation in Toxoplasma-infected {gamma}{delta}- and V{gamma}1+ T cell-deficient mice. Histological analysis (H&E stained) of the liver and CNS of mice orally infected with ME-49 bradyzoites showing extensive inflammation and necrosis in the liver of TCR-{delta}–/– and TCR-V{gamma}1–/– mice and inflammation in the brain of {gamma}{delta} and V{gamma}1+ T cell-deficient mice. The histology shown is representative of >10 mice of each strain. The arrows identify inflammatory cell foci and stars identifying cysts in the CNS at 6 wk postinfection; m = meningitis. Magnification, x100.

 
Toxoplasmic encephalitis was more severe in TCR-V{gamma}1–/– mice infected with ME-49 bradyzoites compared with wild-type mice (Fig. 3). By 3 wk postinfection, both TCR-{delta}–/– and TCR-V{gamma}1–/– mice developed severe encephalitis characterized by vascular cuffing and an extensive infiltrate of inflammatory cells comprising of macrophages, granulocytes, and lymphocytes in the meninges and in brain tissue itself, where it encircled cysts. Inflammation persisted in the brain of TCR-V{gamma}1–/– mice for >6 wk after the initial infection (Fig. 3, and data not shown), consistent with the requirement for V{gamma}1+ T cells in controlling inflammation.

Macrophage homeostasis is disrupted in Toxoplasma-infected V{gamma}1–/– mice

In view of the chronic inflammation seen in Toxoplasma-infected TCR-V{gamma}1–/– mice and the known involvement of V{gamma}1+ T cells in macrophage homeostasis (25, 53), we sought evidence for the disruption of macrophage homeostasis in T. gondii-infected TCR-V{gamma}1–/– mice and the involvement of macrophage-reactive V{gamma}1+ T cells in controlling toxoplasmosis. The chronic inflammatory reaction in tissues of Toxoplasma-infected TCR-{delta}–/– and TCR-V{gamma}1–/– mice was characterized by the accumulation of activated macrophages and granulocytes (Fig. 4), which persisted for up to 4 wk after initial infection. This chronic macrophage and granulocyte response was seen in all major sites of Toxoplasma infection in TCR-V{gamma}1–/– mice, including the liver, spleen, lungs, peritoneal cavity (Fig. 4, and data not shown), and the CNS, where it was still evident 6 wk after initial infection (Fig. 3). In wild-type mice, the kinetics of the macrophage response was mirrored by the V{gamma}1+ T cell response to Toxoplasma infection (Fig. 2B). Further evidence of an uncontrolled and ineffective inflammatory anti-parasite response in TCR-{delta}–/– and TCR-V{gamma}1–/– mice was provided by the analysis of serum levels of the proinflammatory cytokines TNF-{alpha} and IFN-{gamma} (Fig. 4) and CNS parasite burden (Fig. 5A). Significantly higher levels of both cytokines were present in the serum of TCR-{delta}–/– and TCR-V{gamma}1–/– mice throughout the first 2 wk of infection, with the highest levels seen between 7 and 10 days postinfection. Significantly higher numbers of parasite cysts were recovered from the brain of TCR-{delta}–/– and TCR-V{gamma}1–/– mice (Fig. 5A) with occasional histological evidence of cyst rupture in the brain of TCR-{delta}–/– and TCR-V{gamma}1–/– mice, suggestive of reactivation that was not seen in wild-type mice (data not shown) and of a role for V{gamma}1+ T cells in containing parasite growth in the CNS during the chronic phase of T. gondii infection.



View larger version (26K):
[in this window]
[in a new window]
  		FIGURE 4. Exaggerated systemic inflammatory response in Toxoplasma-infected {gamma}{delta}- and V{gamma}1+ T cell-deficient mice. The kinetics of the liver macrophage and granulocyte response in ME-49-infected wild-type mice (WT), {gamma}{delta} T cell-deficient (TCR-{delta}–/–), and V{gamma}1+ T cell-deficient (TCR-V{gamma}1–/–) mice was determined by Ab staining and flow cytometry. The data shown represent the mean (±SEM) values compiled from six mice per time point for each strain of mouse. *, = p < 0.01; **, p < 0.05. Serum TNF-{alpha} and IFN-{gamma} levels were determined by cytokine bead array assays and flow cytometry, with the data shown representing mean (±SEM) values obtained from five to seven animals of each strain. *, p < 0.05; **, p < 0.001; ***, p < 0.0003.

 


View larger version (11K):
[in this window]
[in a new window]
  		FIGURE 5. V{gamma}1+ T cells are required to contain parasite growth during the chronic stage of Toxoplasma infection. A, CNS cyst burden determined by counting of cysts recovered from homogenates of infected brains at 21 days postinfection. The values shown represent the mean (±SEM) of values obtained from >10 mice of each strain. *, p < 0.05 comparing wild type (WT) with TCR-V{gamma}1–/–, and **, p < 0.01 comparing WT with TCR-{delta}–/–. B, The level of macrophage killing by V{gamma}1+ T cells was determined by culturing peritoneal macrophages from day 10 infected TCR-{delta}–/– mice with {gamma}{delta} T cell-enriched splenocytes from noninfected WT, TCR-{beta}–/–, and TCR-V{gamma}1–/– mice and with splenocytes from TCR-{delta}–/– mice, as described in Materials and Methods. Levels of spontaneous macrophage death were determined by culturing macrophages alone (Media). The results represent the mean (±SEM) level of killing in three independent experiments. *, p < 0.001 comparing WT and TCR-{beta}–/– with TCR-{delta}–/– and TCR-V{gamma}1–/–, and **, p < 0.05 comparing WT with TCR-{delta}–/– and TCR-V{gamma}1–/– mice. C, The frequency of V{gamma}1+ T cells synthesizing representative pro- and anti-inflammatory cytokines was determined by surface and intracellular staining of splenocytes directly ex vivo from day 10 infected wild-type mice, as described in Materials and Methods. The data shown represent the mean (±SEM) frequency of cytokine-positive cells in three independent experiments, four to six mice in each experiment.

 
V{gamma}1+ T cells kill parasite-elicited macrophages, a mechanism by which V{gamma}1+ T cells restrict parasite growth and chronic inflammation in toxoplasmosis

Because macrophages are a major target of Toxoplasma infection (Fig. 2A), we determined whether the killing of parasite-elicited and activated macrophages might be a mechanism by which macrophage cytocidal V{gamma}1+ T cells can help restrict parasite dissemination and growth. The killing of T. gondii-elicited, activated macrophages was assessed using a fluorescence-based cytotoxicity assay that we have previously used to demonstrate the killing by V{gamma}1+ T cells of Listeria-elicited macrophages (53), which is Fas-Fas ligand mediated (56). Peritoneal macrophages from day 10 ME-49-infected TCR-{delta}–/– mice were used as target cells, and bulk or {gamma}{delta} T cell-enriched splenocytes (>80% {gamma}{delta} T cells) from noninfected wild-type, TCR-{delta}–/–, TCR-V{gamma}1–/–, or TCR-{beta}–/– mice were used as effector cells (25). As shown in Fig. 5B, T. gondii-elicited macrophages were efficiently killed (>80%) by {gamma}{delta} T cells from wild-type mice and TCR-{beta}–/– mice, but not from TCR-V{gamma}1–/– mice or by splenocytes from TCR-{delta}–/– mice. Due to the difficulty of identifying parasites within the harvested macrophages, it was not possible to determine the extent of killing of infected vs noninfected, but activated macrophages by V{gamma}1+ T cells. Assuming that at least some of the target macrophages were infected, it was not possible to detect the release of any viable parasites from lysed macrophages either microscopically or in in vitro fibroblast reinfection assays (data not shown). Comparable levels of killing were also seen using the spleen or liver as a source of target macrophages (data not shown). Evidence that this killing was directly attributable to V{gamma}1+ T cells was obtained from experiments showing that in using {gamma}{delta} T cells from V{gamma}1–/– mice (Fig. 5B) the level of macrophage killing was not significantly different from the level of spontaneous macrophage death in cultures of macrophages in the absence of T cells. As observed previously (53), the susceptibility of macrophages to lysis by {gamma}{delta} T cells was dependent upon their prior activation, as {gamma}{delta} T cells did not kill macrophages from noninfected wild-type mice (data not shown). Activation was not, however, a requirement for {gamma}{delta} T cell macrophage cytocidal activity (Fig. 5B).

Analysis of cytokine production by peripheral, liver and splenic, V{gamma}1+ T cells responding to parasite infection showed that <6% of T cells produced any of the cytokines tested with IL-4, IL-10, and LAP being synthesized by a small subset of liver- and/or spleen-derived V{gamma}1+ T cells. The poor cytokine response of peripheral V{gamma}1+ T cells resembled that of V{gamma}1+ IELs (Fig. 1C).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Although {gamma}{delta} T cells are found in a number of different anatomical sites, with localization often associated with the expression of distinct TCR-V{gamma} chains, it is not clear what the relative contribution of the different populations is to protecting the host from pathogens, and whether their functional phenotype is predetermined and cosegregates with TCR-V{gamma} chain usage or is influenced by environmental factors. Using the protozoan parasite T. gondii as a model enteric pathogen to investigate the contribution that V{gamma}1+ IELs vs V{gamma}1+ peripheral T cells make to infection, we have shown that whereas V{gamma}1+ IELs do not appear to make a significant contribution to the acute phase of infection or to preventing or limiting parasite-induced ileitis, systemic V{gamma}1+ T cells play an essential role in limiting parasite growth and inflammation in peripheral tissues and in the CNS. These findings also demonstrate that the macrophage cytocidal activity of V{gamma}1+ T cells contributes to antiparasitic as well as antibacterial responses and may therefore be a universal feature of pathogen-induced immune responses.

More extensive accumulation and rapid dissemination of parasites and accelerated onset of ileitis in mice deficient of {gamma}{delta} IELs are consistent with these cells playing an important role in resisting pathogen invasion in the small intestine. Although we have been unable to establish the identity of the protective {gamma}{delta} IEL populations, our data suggest that whereas V{gamma}1+ IELs do not contribute to epithelial defense, the increased resistance of V{gamma}1–/– mice to parasite invasion in which ~90% of {gamma}{delta} IELs are V{gamma}7+ (25) suggests that V{gamma}7+ IELs have a direct role in limiting pathogen invasion and the development of intestinal inflammation. The nature of this protective function of V{gamma}1– {gamma}{delta} IELs is not known, although it appears to be independent of the cytolytic Th1-skewed immunoregulatory properties ascribed to IELs (2). These findings do not, however, discount the importance of {gamma}{delta} IEL cytotoxicity and cytokine production that may be of more relevance to the host response later in the course of infection once parasites have migrated from the intestinal mucosa. Indeed, as ileitis develops ~1 wk after infection, there is evidence of increased IEL activity and cytokine production (42), which precedes lamina propria or Peyer’s patch T cell involvement (35, 57). IEL-mediated killing of parasite-infected cells is only apparent after 1 wk (~9 days) postinfection (58). In addition, the ability of adoptively transferred bulk IELs, which include {alpha}{beta} and {gamma}{delta} T cells to confer protection against severe and lethal ileitis in otherwise susceptible mice (38, 51, 59), may be attributable to IELs killing parasite-infected cells (58) and secreting anti-inflammatory cytokines such as TGF-{beta} and IL-10 (39, 60), or by influencing the local activity or homeostatic regulation of other immune cells (53, 61, 62), including pathogenic {alpha}{beta} (CD4+) T cells (35, 36, 57).

Although there is no obvious requirement for TCR-V{gamma}1+ IELs in limiting parasite invasion and the development of intestinal inflammation, systemic anti-parasite responses are severely compromised in the absence of V{gamma}1+ T cells. Macrophage cytocidal V{gamma}1+ T cells dominate the systemic {gamma}{delta} T cell response to T. gondii as well as a number of other pathogens (63). In comparison with other experimental infection models, however, the Toxoplasma-elicited V{gamma}1+ T cell response is more potent, resulting in greater numbers of V{gamma}1+ T cells and elimination of the vast majority (>80%) of parasite-elicited, activated macrophages (Fig. 5B). Assuming that some of these target macrophages are infected with parasites, V{gamma}1+ cells are likely to play a central and important role in Toxoplasma containment because macrophages are major cellular targets for Toxoplasma infection and dissemination in peripheral tissues (this study) (64) and their elimination would be an effective means of controlling parasite infection. In particular, the unresolved inflammation and increased parasite cyst burden in the brain of infected TCR-{delta}–/– and V{gamma}1–/– mice suggest that V{gamma}1+ T cells can contribute to resolving inflammation in the brain of T. gondii-infected mice by killing activated macrophages that make up a large part of the CNS infiltrate (this study) (65) and can contribute to the development of inflammatory lesions (66). In particular, they may help prevent or limit the extent of CNS inflammation by eliminating macrophages that during the acute phase are infected with cyst-forming bradyzoites (65) and can serve as a kind of "Trojan horse" and transport mechanism for entry of parasite into the brain (64).

The significance of the production of immunoregulatory cytokines, including IL-10, by a small subset of systemic V{gamma}1+ T cells is difficult to interpret due to their small numbers and the fact that these cytokines can be produced by more than one cell type in both the liver and spleen. The production of IL-10 and LAP by V{gamma}1+ T cells may, however, be significant in view of their potent immunosuppressive properties and role in down-modulating pathogenic CD4 T cells and limiting infection-induced pathology (36, 37). Similarly, the significance of the production of IL-6 by a small subset of {gamma}{delta} (V{gamma}7+) IELs is also uncertain, although it might play a role in limiting parasite invasion and the development of intestinal inflammation (67). It is also worth noting that because our analysis of cytokine production by {gamma}{delta} IELs was not exhaustive and did not include all of the cytokines and chemokines known to be produced by these cells (3, 4), it is possible that V{gamma}1+ as well as other {gamma}{delta} IELs populations may contribute to epithelial defense by the production of cytokines other than those analyzed in this study.

Conflicting evidence for both detrimental (61) and beneficial (16) roles of {gamma}{delta} T cells in influencing CNS inflammation has been reported. However, because the identity of the {gamma}{delta} T cells involved was not established, the contribution made by V{gamma}1+ T cells to pathogenic or protective {gamma}{delta} T cell responses in these studies needs to be determined. If V{gamma}1+ T cells are involved, then the timing and nature of their response will be an important factor in determining what the nature of their involvement is. The V{gamma}1+ T cell response to infection is functionally heterogeneous and biphasic with early responding cells of a proinflammatory phenotype and late responding cells with anti-inflammatory properties (25), the timing and magnitude of both being strongly influenced by the nature of the pathogen (virulence), the route, dose, and site of infection, as well as host factors (68). The apparently conflicting roles for {gamma}{delta}, and possibly V{gamma}1+, T cells in the pathogenesis of CNS disease may therefore be explained by differences in the nature of the pathogen and conditions under which {gamma}{delta} T cells are recruited and whether the prevailing microenvironmental conditions favor the involvement of pro- or anti-inflammatory populations.

In demonstrating differences in the involvement of populations of mucosal and systemic {gamma}{delta} T cells that use the same TCR-V{gamma} chain in regulating host responses to infection, this study increases our understanding of the factors that influence {gamma}{delta} T cell function during both acute and chronic stages of infections. Although the clonal relationship of peripheral vs intestinal V{gamma}1+ T cells has not been established, their contrasting level of involvement and contribution to the host response to Toxoplasma infection suggests that they are functionally distinct, reflecting microenvironmental influences unique to the different anatomical sites in which they reside and operate.


   Acknowledgments
 
We thank the personnel of the pathology laboratory of the Leeds Teaching Hospital Trust for their help in developing immunohistochemical protocols, and Dr. Antonio Barragan (Karolinska Institut, Stockholm, Sweden) and Professor Jim Alexander and Dr. Craig Roberts (Strathclyde University, Glasgow, Scotland) for helpful comments and discussions during the preparation of the manuscript.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
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 by grants from The Wellcome Trust to S.R.C. and by a Biotechnology and Biological Sciences Research Council studentship to C.E.E. Back

2 Current address: Department of Microbiology and Immunology, College Veterinary Medicine, Cornell University, Ithaca, NY 14853. Back

3 Current address: Department of Biology, Boston College, Chestnut Hill, MA 02467. Back

4 Address correspondence and reprint requests to Dr. Simon R. Carding, University of Leeds, Research Institute for Molecular and Cellular Biology, Irene Manton Research Building, Room 8.91h, Leeds LS2 9JT, W. Yorkshire, U.K. E-mail address: S.R.Carding{at}Leeds.ac.uk Back

5 Abbreviations used in this paper: IEL, intestinal intraepithelial lymphocyte; ALT, alanine aminotransferase; LAP, latency-associated peptide; MLN, mesenteric lymph node; PMNC, polymorphonuclear cell; YFP, yellow fluorescent protein. Back

Received for publication August 19, 2005. Accepted for publication October 7, 2005.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Hayday, A. C.. 2000. {gamma}{delta} T cells: A right time and a right place for a conserved third way of protection. Annu. Rev. Immunol. 18: 975-1026. [Medline]
  2. Hayday, A., E. Theodoridis, E. Ramsburg, J. Shires. 2001. Intraepithelial lymphocytes: exploring the Third Way in immunology. Nat. Rev. Immunol. 2: 997-1003.
  3. Shires, J., E. Theodoridis, A. C. Hayday. 2001. Biological insights into TCR{gamma}{delta}+ and TCR{alpha}{beta}+ intraepithelial lymphocytes provided by serial analysis of gene expression (SAGE). Immunity 15: 435-444. [Medline]
  4. Fahrer, A. M., Y. Konigshofer, A. M. Kerr, G. Ghandour, D. H. Mack, M. M. Davis, Y. H. Chien. 2001. Attributes of {gamma}-{delta} intraepithelial lymphocytes suggested by their transcriptional profile. Proc. Natl. Acad. Sci. USA 98: 10261-10266. [Abstract/Free Full Text]
  5. Havran, W. L., Y. Chien, J. Allison. 1991. Recognition of self antigens by skin derived T cells with invariant {gamma}{delta} antigen receptors. Science 252: 1430-1432. [Abstract/Free Full Text]
  6. Boismenu, R., W. L. Havran. 1994. Modulation of epithelial cell growth by intraepithelial {gamma}{delta} T cells. Science 266: 1253-1255. [Abstract/Free Full Text]
  7. Komano, H., Y. Fujiura, M. Kawaguchi, S. Matsumoto, Y. Hashimoto, S. Obana, P. Mombaerts, S. Tonegawa, H. Yamamoto, S. Itohara, et al 1995. Homeostatic regulation of intestinal epithelia by intraepithelial {gamma}{delta} T cells. Proc. Natl. Acad. Sci. USA 92: 6147-6151. [Abstract/Free Full Text]
  8. Janeway, C. A., Jr, B. Jones, A. C. Hayday. 1988. Specificity and function of T cells bearing {gamma}{delta} receptors. Immunol. Today 9: 73-76. [Medline]
  9. Mombaerts, P., J. Arnoldi, F. Russ, S. Tonegawa, S. H. E. Kauffman. 1993. Different roles of {alpha}{beta} and {gamma}{delta} T cells in immunity against an intracellular bacterial pathogen. Nature 365: 53-56. [Medline]
  10. Fu, Y.-X., C. E. Roark, K. Kelly, D. Drevets, P. Campbell, R. O’Brien, W. K. Born. 1994. Immune protection and control of inflammatory tissue necrosis by {gamma}{delta} T cells. J. Immunol. 153: 3101-3115. [Abstract]
  11. Roberts, S. J., A. L. Smith, A. B. West, L. Wen, R. C. Findly, M. J. Owen, A. C. Hayday. 1996. T-cell {alpha}{beta}+ and {gamma}{delta}+ deficient mice display abnormal but distinct phenotypes towards a natural, widespread infection of the intestinal epithelium. Proc. Natl. Acad. Sci. USA 93: 11774-11779. [Abstract/Free Full Text]
  12. D’Souza, C. D., A. M. Cooper, A. A. Frank, R. J. Mazzaccaro, B. R. Bloom, I. M. Orme. 1997. An anti-inflammatory role for {gamma}{delta} T lymphocytes in acquired immunity to Mycobacterium tuberculosis. J. Immunol. 158: 1217-1221. [Abstract]
  13. Moore, T. A., B. B. Moore, M. W. Newstead, T. J. Standiford. 2000. {gamma}{delta} T cells are critical for survival and early proinflammatory gene expression during murine Klebsiella pneumonia. J. Immunol. 165: 2643-2650. [Abstract/Free Full Text]
  14. Seixas, E. M. G., J. Langhorne. 1999. {gamma}{delta} T cells contribute to control of chronic parasitemia in Plasmodium chabaudi infection in mice. J. Immunol. 162: 1837-1841.
  15. Moretto, M., B. Durell, J. D. Schwartzman, I. A. Khan. 2001. {gamma}{delta} T cell-deficient mice have a down-regulated CD8+ T cell immune response against Encephalitozoan cuniculi infection. J. Immunol. 166: 7389-7397. [Abstract/Free Full Text]
  16. Wang, T., E. Scully, Z. N. Yin, J. H. Kim, S. Wang, J. Yan, E. Fikrig. 2003. IFN{gamma}-producing {gamma}{delta} T cells help control murine West Nile virus infection. J. Immunol. 171: 2524-2531. [Abstract/Free Full Text]
  17. Selin, L. K., P. A. Santolcuito, A. K. Pinto, E. Szomolanyi-Tsuda, R. M. Welsh. 2001. Innate immunity to viruses: control of vaccinia virus infection by {gamma}{delta} T cells. J. Immunol. 166: 6784-6794. [Abstract/Free Full Text]
  18. Egan, P. E., S. R. Carding. 2000. Down-modulation of the inflammatory response to bacterial infection by {gamma}{delta} T cells cytotoxic for activated macrophages. J. Exp. Med. 191: 2145-2158. [Abstract/Free Full Text]
  19. Skeen, M., M. Freeman, H. Ziegler. 2004. Changes in myeloid populations and their proinflammatory cytokine expression during infection with Listeria monocytogenes in the absence of {gamma}{delta} T cells. J. Leukocyte Biol. 76: 104-115. [Abstract/Free Full Text]
  20. Kaufmann, S. H. E., C. Blum, S. Yamamoto. 1993. Crosstalk between {alpha}{beta} T cells and {gamma}{delta} T cells in vivo: activation of {alpha}{beta} T cell responses after {gamma}{delta} T cell modulation with the monoclonal antibody GL3. Proc. Natl. Acad. Sci. USA 90: 9620-9624. [Abstract/Free Full Text]
  21. Huber, S. A., D. Graveline, M. K. Newell, W. K. Born, R. L. O’Brien. 2000. V{gamma}1+ T cells suppress and V{gamma}4+ T cells promote susceptibility to Cocksackievirus B3-induced myocarditis in mice. J. Immunol. 165: 4174-4181. [Abstract/Free Full Text]
  22. Huber, S. A., D. Graveline, W. K. Born, R. L. O’Brien. 2001. Cytokine production by V{gamma}+ T cell subsets is an important factor determining CD4+ Th-phenotype and susceptibility of BALB/c mice to Coxsackievirus B3-induced myocarditis. J. Virol. 75: 5860-5869. [Abstract/Free Full Text]
  23. Hahn, Y.-S., C. Taube, N. Jin, L. Sharp, J. M. Wands, M. K. Aydintug, M. Lahn, S. A. Huber, R. L. O’Brien, E. W. Gelfand, W. K. Born. 2004. Different potentials of {gamma}{delta} T cell subsets in regulating airway responsiveness: V{gamma}1+ cells, but not V{gamma}4+ cells, promote airway hyperreactivity, Th2 cytokines, and iarway inflammation. J. Immunol. 172: 2894-2902. [Abstract/Free Full Text]
  24. Lahn, M., A. Kanehiro, K. Takeda, J. Terry, Y.-S. Hanh, M. K. Aydintug, A. Knowal, K. Ikuta, R. O’Brien, E. W. Gelfand, W. K. Born. 2002. MHC class I-dependent V{gamma}4+ pulmonary T cells regulate {alpha}{beta} T cell-independent airway responsiveness. Proc. Natl. Acad. Sci. USA 99: 8850-8855. [Abstract/Free Full Text]
  25. Andrew, E. M., D. J. Newton, J. E. Dalton, C. E. Egan, S. J. Goodwin, D. Tramonti, P. Scott, S. R. Carding. 2005. Delineation of the function of a major {gamma}{delta} T cell subset during infection. J. Immunol. 175: 1741-1750. [Abstract/Free Full Text]
  26. Hill, D. B., J. P. Dubey. 2002. Toxoplasma gondii: transmission, diagnosis and prevention. Clin. Microbiol. Infect. 8: 634-640. [Medline]
  27. Suzuki, Y., M. A. Orellana, S. Y. Wong, F. K. Conley, J. S. Remington. 1993. Susceptibility to chronic infection with Toxoplasma gondii does not correlate with susceptibility to acute infection in mice. Infect. Immun. 61: 2284-2288. [Abstract/Free Full Text]
  28. Brown, C., R. McLeod. 1994. Mechanisms of survival of mice during acute and chronic Toxoplasma gondii infection. Parasitol. Today 10: 290-292. [Medline]
  29. Darcy, F., F. Santoro. 1994. Toxoplasmosis. F. Kierszenbaum, Jr, ed. Parasitic Infections and the Immune System 163-201. Academic, San Diego.
  30. Suzuki, Y., C. Orellana, R. D. Schreiber, J. S. Remington. 1988. Interferon-{gamma}: the major mediator of resistance against Toxoplasma gondii. Science 240: 516-518. [Abstract/Free Full Text]
  31. Gazzinelli, R. T., F. T. Hakim, S. Hieny, G. M. Shearer, A. Sher. 1991. Synergistic role for CD4+ and CD8+ T lymphocytes in IFN-{gamma} production and protective effect immunity induced by attenuated T. gondii vaccine. J. Immunol. 146: 6155-6162.
  32. Gazzinelli, R. T., Y. J. Xu, S. Hieny, A. Cheever, A. Sher. 1992. Simultaneous depletion of CD4+ and CD8+ T lymphocytes is required to reactivate chronic infection with Toxoplasma gondii. J. Immunol. 149: 175-180. [Abstract]
  33. Gazzinelli, R. T., M. Wysocka, S. Hayashi, E. Y. Denkers, S. Hieny, P. Caspar, G. Trinchieri, A. Sher. 1994. Parasite-induced IL-12 stimulates early IFN-{gamma} synthesis and resistance during acute infection with Toxoplasma gondii. J. Immunol. 153: 2533-2543. [Abstract]
  34. Scharton-Kersten, T. M., T. A. Wynn, E. Y. Denkers, S. Bala, E. Grunvald, S. Heiny, R. T. Gazzinelli, A. Sher. 1996. In the absence of endogenous IFN-{gamma}, mice develop unimpaired IL-12 responses to Toxoplasma gondii while failing to control acute infection. J. Immunol. 157: 4045-4054. [Abstract]
  35. Liesenfeld, O., J. Kosek, J. S. Remington, Y. Suzuki. 1996. Association of CD4+ T cell-dependent, interferon-{gamma}-mediated necrosis of the small intestine and early mortality in genetically susceptible mice to peroral infection with Toxoplasma gondii. J. Exp. Med. 184: 597-607. [Abstract/Free Full Text]
  36. Gazzinelli, R. T., M. Wysocka, S. Heiny, T. Scharton-Kersten, A. Cheever, R. Kuhn, W. E. G. Muller, G. Trichieri, A. Sher. 1996. In the absence of endogenous IL-10, mice acutely infected with Toxoplasma gondii succumb to lethal immune response dependent on CD4+ T cells and accompanied by overproduction of IL-12, IFN-{gamma} and TNF-{alpha}. J. Immunol. 157: 798-805. [Abstract]
  37. Suzuki, Y., A. Sher, G. Yap, D. Park, L. E. Neyer, O. Liesenfeld, M. Fort, H. Kang, E. Gufwoli. 2000. IL-10 is required for prevention of necrosis in the small intestine and mortality in both genetically resistant BALB/c and susceptible C57BL/6 mice following peroral infection with Toxoplasma gondii. J. Immunol. 164: 5375-5382. [Abstract/Free Full Text]
  38. Lepage, A. C., D. Buzoni-Gatel, D. T. Bout, L. H. Kasper. 1998. Gut-derived intraepithelial lymphocytes induce long term immunity against Toxoplasma gondii. J. Immunol. 161: 4902-4908. [Abstract/Free Full Text]
  39. Buzoni-Gatel, D., H. Debbabi, M. Moretto, I. H. Dimier-Poisson, A. C. Lepage, D. T. Bout, L. H. Kasper. 2001. Murine ileitis after intracellular parasite infection is controlled by TGF-{beta}-producing intraepithelial lymphocytes. Gastroenterology 120: 914-924. [Medline]
  40. Hisaeda, H., H. Nagasawwa, K.-I. Maeda, Y. Maekawa, H. Ishikawa, Y. Ito, R. A. Good, K. Himeno. 1995. {gamma}{delta} T cells play an important role in hsp65 expression and in acquiring protective immune responses against infection with Toxoplasma gondii. J. Immunol. 154: 244-251.
  41. Kasper, L. H., T. Matsuura, S. Foneska, J. Arruda, J. Y. Channon, I. A. Khan. 1996. Induction of {gamma}{delta} T cells during acute murine infection with Toxoplasma gondii. J. Immunol. 157: 5521-5527. [Abstract]
  42. Lee, Y.-H., D.-W. Shin. 2002. T cell phenotype and intracellular IFN-{gamma} production in peritoneal exudate cells and gut intraepithelial lympocytes during acute Toxoplasma gondii infection in mice. Kor. J. Parasitol. 40: 119-129.
  43. Kempf, M. C., M. F. Cesbron-Delauw, D. Deslee, U. Gross, T. Herrmann, P. Sutton. 1999. Different manifestations of Toxoplasma gondii infection in F344 and LEW rats. Med. Microbiol. Immunol. 187: 137-142. [Medline]
  44. De Paoli, P., G. Basaglia, D. Geannari, M. Crovatto, M. L. Modolo, G. Santini. 1992. Phenotypic profile and functional characteristics of human {gamma} and {delta} T cells during acute toxoplasmosis. J. Clin. Microbiol. 30: 729-731. [Abstract/Free Full Text]
  45. Scalise, F., R. Gerli, G. Castellucci, F. Spinozzi, G. M. Fabietti, S. Crupi, L. Sensi, R. Britta, R. Vaccaro, A. Bertotto. 1992. Lymphocytes bearing the {gamma}-{delta} T cell receptor in acute toxoplasmosis. Immunology 76: 668-670. [Medline]
  46. Subauste, C. S., J. Y. Chung, D. D. Koniaris, C. A. Hunter, J. G. Montoya, S. Porcelli, J. S. Remington. 1995. Preferential activation and expansion of human peripheral blood {gamma}{delta} T cells in response to Toxoplasma gondii in vitro and their cytokine production and cytotoxicity against T. gondii-infected cells. J. Clin. Invest. 96: 610-619. [Medline]
  47. Hara, T., S. Ohashi, Y. Yamashita, T. Abe, H. Hisaeda, K. Himeno, R. A. Good, K. Takeshita. 1996. Human V{delta}2+ {gamma}{delta} T cell-tolerance to foreign antigens of Toxoplasma gondii. Proc. Natl. Acad. Sci. USA 93: 5136-5140. [Abstract/Free Full Text]
  48. Sayles, P. C., A. L. Rakhmilevich, L. L. Johnson. 1995. {gamma}{delta} T cells and acute primary Toxoplasma infection in mice. J. Infect. Dis. 171: 249-257. [Medline]
  49. Gubbels, M.-J., C. Li, B. Striepen. 2003. High throughput growth assay for Toxoplasma gondii using yellow fluorescent protein. Antimicrob. Agents Chemother. 47: 309-316. [Abstract/Free Full Text]
  50. Dubey, J. P.. 1998. Re-examination of resistance of Toxoplasma gondii tachyzoites and bradyzoites to pepsin and trypsin digestion. Parasitology 116: 43-50.
  51. Buzoni-Gatel, D., H. Debbabi, M. Moretto, I. H. Dimier-Poisson, A. C. Lepage, D. Bout, L. H. Kasper. 1999. Intraepithelial lymphocytes traffick to the intestine and enhance resistance to Toxoplasma gondii oral infection. J. Immunol. 162: 5846-5852. [Abstract/Free Full Text]
  52. Pereira, P., D. Gerber, S. Y. Huang, S. Tonegawa. 1995. Ontogenetic development and tissue distribution of V{gamma}1-expressing {gamma}/{delta} T lymphocytes in normal mice. J. Exp. Med. 182: 1921-1929. [Abstract/Free Full Text]
  53. Dalton, J. E., J. Pearson, P. Scott, S. R. Carding. 2003. The interaction of {gamma}{delta} T cells with activated macrophages is a property of the V{gamma}1 subset. J. Immunol. 171: 6488-6494. [Abstract/Free Full Text]
  54. Barragan, A., D. L. Sibley. 2002. Transepithelial migration of Toxoplasma gondii is linked to parasite motility and virulence. J. Exp. Med. 195: 1625-1633. [Abstract/Free Full Text]
  55. Martinon, F., J. Tschopp. 2004. Inflammatory caspases: linking an intracellular innate immune system to autoinflammatory diseases. Cell 117: 561-574. [Medline]
  56. Dalton, J. E., G. Howell, J. Pearson, P. Scott, S. R. Carding. 2004. Fas/FasL interactions are essential for binding to and killing of activated macrophages by {gamma}{delta} T cells. J. Immunol. 204: 3660-3667.
  57. Mennechet, F. J. D., L. H. Kasper, N. Rachinel, W. Li, A. Vandewalle, D. Buzoni-Gatel. 2002. Lamina propria CD4+ T lymphocytes synergize with murine intestinal epithelial cells to enhance proinflammatory response against an intracellular pathogen. J. Immunol. 168: 2988-2996. [Abstract/Free Full Text]
  58. Chardes, T., D. Buzoni-Gatel, A. C. Lepage, F. Bernard, D. Bout. 1994. Toxoplasma gondii oral infection induces specific cytotoxic CD8{alpha}/{beta}+ Thy-1+ gut intraepithelial lymphocytes, lytic for parasite-infected enterocytes. J. Immunol. 153: 4596-4603. [Abstract]
  59. Buzoni-Gatel, D., A. C. Lepage, I. H. Dimier-Poisson, D. Bout, L. H. Kasper. 1997. Adoptive transfer of gut intraepithelial lymphocytes protects against murine infection with Toxoplasma gondii. J. Immunol. 158: 5883-5889. [Abstract]
  60. Kasper, L. H., N. Courret, S. Darche, S. Luangsay, F. J. D. Mennechet, L. Minns, N. Rachinel, C. Ronet, D. Buzoni-Gatel. 2004. Toxoplasma gondii and mucosal immunity. Int. J. Parasitol. 34: 234-242.
  61. Cardona, A. E., P. A. Gonzalez, J. M. Teale. 2003. CC chemokines mediate leuykocyte trafficking into the central nervous system during murine neurocysticercosis: role of T cells in amplification of the host response. Infect. Immun. 71: 2634-2642. [Abstract/Free Full Text]
  62. Tagawa, T., H. Nishimura, T. Yajima, H. Hara, K. Kishihara, G. Matsuzaki, I. Yoshino, Y. Maehara, Y. Yoshikai. 2004. V{delta}1+ {gamma}{delta} T cells producing CC chemokines may bridge a gap between neutrophils and macrophages in innate immunity during Escherichia coli infection in mice. J. Immunol. 173: 5156-5164. [Abstract/Free Full Text]
  63. Andrew, E. M., S. R. Carding. 2005. Murine {gamma}{delta} T cells in infection: beneficial or deleterious?. Microbes Infect. 7: 529-536. [Medline]
  64. Da Gamma, L. M., F. L. Ribeiro-Gomes, U. Guimaraes, Jr, A. C. V. Arnholdt. 2004. Reduction in adhesiveness to extracellular matrix components, modulation of adhesion molecules and in vivo migration of murine macrophages infected with Toxoplasma gondii. Microbes Infect. 6: 1287-1296. [Medline]
  65. Schluter, D., J. Lohler, M. Deckert, H. Hof, G. Schwendenmann. 1991. Toxoplasma encephalitis of immunocompetent and nude mice: immunohistochemical characterization of Toxoplasma antigen, infiltrates and major histocompatibility complex gene products. J. Neuroimmunol. 31: 185-198. [Medline]
  66. Tackey, R. N., O. O. Kassim. 1999. Macrophage responses to Toxoplasma antigens in vitro: a possible role in inflammatory lesions in toxoplasmosis. Am. J. Trop. Med. Hyg. 61: 272-279. [Abstract]
  67. Lieberman, L. A., C. A. Hunter. 2002. The role of cytokines and their signaling pathways in the regulation of immunity to Toxoplasma gondii. Int. Rev. Immunol. 21: 373-403. [Medline]
  68. Belles, C., A. L. Kuhl, A. Donoghue, Y. Sano, R. L. O’Brien, W. Born, K. Bottomly, S. R. Carding. 1996. Bias in the {gamma}{delta} T cell response to Listeria monocytogenes. J. Immunol. 156: 4280-4289. [Abstract]




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrow Request Permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Egan, C. E.
Right arrow Articles by Carding, S. R.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Egan, C. E.
Right arrow Articles by Carding, S. R.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS