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


     
 

The Journal of Immunology, 2009, 182, 1069 -1078
Copyright © 2009 by The American Association of Immunologists, Inc.
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 HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Gigley, J. P.
Right arrow Articles by Bzik, D. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Gigley, J. P.
Right arrow Articles by Bzik, D. J.

Cell-Mediated Immunity to Toxoplasma gondii Develops Primarily by Local Th1 Host Immune Responses in the Absence of Parasite Replication1

Jason P. Gigley*,{dagger}, Barbara A. Fox* and David J. Bzik2,*

* Department of Microbiology and Immunology, Dartmouth Medical School, Lebanon, NH 03756; and {dagger} Department of Microbiology, Immunology and Tropical Medicine, George Washington University, Washington, DC 20037


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
A single inoculation of mice with the live, attenuated Toxoplasma gondii uracil auxotroph strain cps1-1 induces long-lasting immunity against lethal challenge with hypervirulent strain RH. The mechanism for this robust immunity in the absence of parasite replication has not been addressed. The mechanism of long-lasting immunity, the importance of route of immunization, cellular recruitment to the site of infection, and local and systemic inflammation were evaluated. Our results show that infection with cps1-1 elicits long-lasting CD8+ T cell- mediated immunity. We show that immunization with cps1-1-infected dendritic cells elicits long-lasting immunity. Intraperitoneal infection with cps1-1 induced a rapid influx of GR1+ neutrophils and two stages of GR1+CD68+ inflammatory monocyte infiltration into the site of inoculation. CD19+ B cells and CD3+ T cells steadily increase for 8 days after infection. CD8+ T cells were rapidly recruited to the site of infection and increased faster than CD4+ T cells. Surprisingly, cps1-1 infection induced high systemic levels of bioactive IL-12p70 and a very low level and transient systemic IFN-{gamma}. Furthermore, we show significant levels of these inflammatory cytokines were locally produced at the site of cps1-1 inoculation. These findings offer new insight into immunological mechanisms and local host responses to a non-replicating type I parasite infection associated with development of long-lasting immunity to Toxoplasma gondii.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
The uracil auxotroph strain cps1-1 developed from the virulent-type I strain RH does not replicate in the absence of uracil and is completely avirulent in both immune-competent and immune-compromised hosts such as IFN-{gamma}-deficient mice (1). A single inoculation of cps1-1 tachyzoites induces immunity against lethal type I parasite challenge; however, the mechanisms involved in cps1-1 vaccine-induced long-lasting immunity are unknown. Previous vaccine models of host response against infection have been complicated by parasite replication during the acute stage of T. gondii infection and a lack of immunity induced by dead parasites. These studies indicate that high systemic levels of IL-12p70 and IFN-{gamma} induced by live parasites are required to control acute T. gondii infections as well as for the development of immunity (1, 2, 3, 4, 5, 6, 7, 8, 9).

The current model of the host response leading to immunity suggests that acute T. gondii infection initiates the inflammatory response through both MyD88-independent and -dependent pathways (2, 3, 5, 10). Through these pathways several cell types produce high levels of bioactive heterodimer IL-12p70. IL-12p70 production appears to be dependent upon parasite genotype and during acute type II infections both IL-12p40 and IL-12p70 are produced while type I infections primarily induce only high levels of IL-12p40 (10, 11). Many cell types contribute to the production of IL-12p40 and IL-12p70 including GR1+ neutrophils, GR1+CD68+ monocytes, and dendritic cells (DCs)3 (3). Neutrophils are recruited early in high numbers to the site of infection, are required for control of acute infection, and produce early IL-12p70 and IL-12p40 (12, 13, 14, 15). High numbers of GR1+CD68+ monocytes are also recruited during acute T. gondii infection and contribute to the production of IL-12p40 and IL-12p70 and may be able to clear intracellular parasites by autophagy (16, 17, 18, 19). DCs also produce IL-12p40 and IL-12p70 and play a prominent role as an APC in the development of immunity against T. gondii infection (3, 6, 20, 21). Activation of T. gondii-specific CD4+ and CD8+ T cells by both IL-12p70 and recognition of T. gondii-specific Ags presented by APCs results in their expansion leading to high systemic levels of IFN-{gamma} required to control the acute infection. CD8+ T cells are the primary effector cells essential for long-lasting immunity and require CD4+ T cell help for priming (22, 23). The requirement for systemic IFN-{gamma} in the development of immunity and the kinetics of cellular recruitment to acute infection has not been previously investigated.

In this study, we examine the murine host immune response to the non-replicating type I vaccine strain cps1-1 and show that in the absence of replication innate and adaptive immune cells are recruited to the site of infection in a distinct kinetic pattern. Our data show that local cytokine responses are more potent than the systemic responses and a long-lasting CD8+ T cell immunity develops in the absence of significant systemic IFN-{gamma}. We show that cps1-1-infected DCs are highly effective at inducing long-lasting immunity. Our results distinctly highlight for the first time that after the initial response is initiated in draining lymphoid tissue, infection-localized low-level dynamic immunological mechanisms purely in response to the invading parasite are sufficient to lead to host control of and development of immunity against T. gondii infection. These findings advance the nonreplicating cps1-1 strain as a vaccine tool to more effectively guide vaccine design.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Experimental animals

Adult 6- to 8-wk-old C57BL/6, C57BL/6 B cell- deficient (µMT), IL-15–/– mice (B6), or age-matched BALB/c and BALB/c IFN-{gamma} –/– (IFN-{gamma} knockout) were obtained from The Jackson Laboratory. Mice were maintained in Tecniplast Seal Safe mouse cages on vent racks at the Dartmouth-Hitchcock Medical Center mouse facility (24). µMT mice were maintained under sterile food conditions. All mice were cared for and handled according to the Animal Care and Use Program of Dartmouth College using National Institutes of Health approved institutional animal care and use committee guidelines.

Parasites, parasite infections, immunizations, and preparations

Tachyzoites of the virulent type I strain RH or attenuated type I strain cps1-1 were obtained by serial passage in human foreskin fibroblast cell monolayers in Eagle’s MEM supplemented with 1% FBS and 250 nM uracil (cps1-1 only) (1). Viability of tachyzoites (30–50% infectious) was tested by plaque assay. Mice were immunized once i.v. or twice i.p. 2 wk apart with 1 x 106 cps1-1 tachyzoites. One month after final immunization, mice were challenged i.p. with either low 1 x 102 or high (1 x 103 or 1 x 105) doses of viable RH (25). Total tachyzoite lysate Ag was prepared from in vitro human foreskin fibroblast cultures of RH as previously described (26, 27). For cytokine assays, mice were infected i.p. with either 1 x 106 cps1-1 or 1 x 103 RH tachyzoites.

Ex vivo cell infections

Peritoneal exudate cells (PECs), peritoneal resident macrophages, and DCs were obtained from naive C57BL/6 mice. DCs were obtained from the spleens of wild-type (WT) mice and purified using EasySep CD11c-positive selection as per the manufacturer’s instructions (StemCell Technologies). PECs were obtained by peritoneal lavage with 5–7 ml of ice-cold sterile PBS and peritoneal resident macrophages were obtained by plating a portion of total PECs at 4 x 106 cells/ml in 24-well plates for 4 h at 37°C in DMEM supplemented with 10% FBS. Nonadherent cells were removed by gently washing and purity of macrophages was verified (>90% CD11b+) (28). The DCs, PECs, and PEC resident macrophages were plated at 2 x 106 cells/ml in infection medium consisting of Eagle’s minimal essential medium supplemented with 1% FBS and 250 nM uracil (Sigma-Aldrich). Purified cps1-1 tachyzoites were inoculated into individual wells at 5 x 105 parasites/ml for 12 h at 37°C. Infected cells were examined by light microscopy and contained one to four cps1-1 tachyzoites. Remaining extracellular tachyzoites were removed and mice were inoculated once with either 1 x 106 cps1-1 tachyzoites or 5 x 105 of infected DCs, PECs, or PEC resident macrophages.

Flow cytometric differential cell analysis

Total PECs were obtained at the indicated times after infection and were depleted of erythrocytes using buffered ammonium chloride and enumerated by trypan blue exclusion (29). PECs were fixed (1% paraformaldehyde) and stained using standard procedures in stain wash buffer containing 2% FBS in 1x PBS, 5% normal mouse serum (Sigma-Aldrich), and Mouse FC Block (BD Biosciences). Fixed PECs were singly or doubly stained with PE-conjugated anti-mouse CD3 (17A2), PE-conjugated anti-mouse GR1 (RB6-8C5), FITC-conjugated anti-mouse CD45R/B220 (RA3-6B2), FITC-conjugated anti-mouse CD11b (M1/70), FITC-conjugated anti-mouse CD19, anti-mouse F4/80, or the intracellular stain FITC-conjugated anti-mouse CD68. All Abs were from BD Biosciences except Abs to F4/80 and CD68, both from Serotec. For the intracellular stain (CD68), fixed PECs were permeabilized using stain wash buffer/0.5% saponin. Nonspecific Ig isotype-specific Abs were used as negative controls for all samples (BD Biosciences). Flow cytometric data was acquired and analyzed using a FACSCalibur flow cytometer and CellQuest software (BD Immunocytometry Systems). Absolute numbers are the product of the percentage of total events collected from 5 x 104 total events multiplied by total cell number per mouse.

Cytokine assays

Serum was obtained from cps1-1- or RH-infected mice at indicated times from whole blood incubated at room temperature for 2 h and then centrifuged for 10 min at 14,000 rpm at 4°C. Total PECs and splenocytes from infected mice were harvested and then depleted of erythrocytes, counted, and seeded in 24-well plates with DMEM supplemented with 10% FBS. PECs were plated at 1 x 106 cells/well and splenocytes at 5 x 106 cells/well and cultured for 24 h without Ag restimulation (16). Cell cultures were lysed by freeze-thawing three times. Debris were removed from the supernatants by centrifugation at 14,000 rpm at 4°C for 10 min. Concentrations of the mouse cytokines IFN-{gamma}, IL-12p40, and IL-12p70 in serum (1/4 dilution) and cell supernatants were then determined using OptEIA ELISA kits (BD Biosciences) (11). ELISA was developed with tetramethylbenzidine substrate (Pierce) and stopped using 0.2 N H2SO4 (30).

T. gondii lysate preparation and serum IgG assay

Anti-toxoplasma-specific serum IgG responses were measured in cps1-1-immunized mice by coating each well of a 96-well plate (Nunc) with 100 µl of Toxoplasma lysate Ag at a concentration of 1 µg/ml in 0.1 M Na2HPO4 (pH 9.0) overnight at 4°C. Plates were washed three times with 1x PBS with 0.05% Tween 20 (PT), then blocked with PT/1% BSA. Sera were serially diluted beginning with 1/100 in PT/0.1% BSA and applied to each well in triplicate and incubated overnight at 4°C. Plates were washed with PT and then mouse anti-Toxoplasma Igs were detected with either anti-mouse IgG H + L, anti-mouse IgG1, or anti-mouse IgG2a (Jackson ImmunoResearch Laboratories). All secondary Abs were HRP-conjugated and developed as described for cytokine ELISA.

Adoptive transfer and T lymphocyte Ab depletion

All procedures were performed 1 mo after final cps1-1 immunization. Spleens were harvested and after disruption and RBC lysis, CD19+ B cells or CD8+ T cells were purified using EasySep Mouse positive selection kits (StemCell Technologies) (29). Naive WT mice received either 1 x 107 CD8+ T cells, 5.0 x 106 CD19+ B cells, or 4 x 107 immune or naive splenocytes (4 x 107 splenocytes contained ~2 x 106 CD8+ T cells and 4 x 106 CD4+ T cells) via tail vein injection. Twenty-four hours after cell transfer, mice were i.p. challenged with 1 x 103 RH tachyzoites (31). T cell subpopulations were depleted by i.p. injection of 500 µg of either control rat IgG, anti-CD4 Ab (GK1.5), anti-CD8 Ab (TIB210), or both anti-CD4/CD8 Ab on days –3, –2, –1, and 0, then every other day until day 21 after challenge (32). On day 0 of Ab treatment, T cell depletion was confirmed and mice were i.p. challenged with 1 x 103 RH.

Quantitative real-time PCR for parasite equivalents

Spleens and brains from infected animals were harvested and DNA was extracted from an entire organ using a Qiagen DNeasy Tissue Kit (Qiagen Sciences) and pooled. Amplification of parasite DNA from 400 ng of purified tissue DNA was performed using primers specific for the T. gondii B1 gene (forward primer, GGAACTGCATCCGTTCATG and reverse primer, TCTTTAAAGCGTTCGTGGTC) at 10 pmol of each per reaction (Integrated DNA Technologies) (12, 33) and amplified by real-time fluorogenic PCR using SMartMix HM (Cepheid) on a Cepheid Smart Cycler. Each reaction contained one lyophilized SMartMix HM bead and 1/20,000 SYBR Green I (Cambrex). Parasite equivalents were determined by extrapolation from a standard curve.

Statistical analysis

The Kaplan-Meier product limit test was used to measure significant differences between survival curves of i.v. and ex vivo-loaded cell type route experiments (GraphPad PRISM software). All other means comparisons were subjected to a Student t test and are represented as the mean ± SEM.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
cps1-1 tachyzoites induce protective immunity capable of clearing a high-dose lethal-type I RH challenge

A single inoculation of live-attenuated cps1-1 tachyzoites protect type II T. gondii-resistant BALB/c mice against a low-dose lethal-type I challenge (1). Other studies report that a single inoculation of irradiation inactivated cps1-1-induce immunity in Tyk2–/– and C57BL/6 mice in a lethal challenge model (34). We sought to determine the mechanism of immune protection and efficacy of cps1-1-induced immunity to clear a challenge infection in the highly sensitive C57BL/6 mouse background. cps1-1-immunized C57BL/6 mice were completely protected (Fig. 1A) and completely clear lethal-type I challenge (Fig. 1B), whereas all naive mice uniformly succumbed to infection by day 10 after challenge and had high parasite burdens. All cps1-1-immunized mice regardless of challenge dose were continuously monitored for 18 mo and uniformly survived challenge infection to old age, showing that cps1-1 induces long-lasting immunity (data not shown).


Figure 1
View larger version (10K):
[in this window]
[in a new window]

  		FIGURE 1. Percent survival of cps1-1-immunized C57BL/6 (B6) mice. B6 mice were immunized i.p. with 106 cps1-1 parasites twice 14 days apart or not immunized. At 21 days after the final immunization, all mice were challenged i.p. with either 103 RH parasites for survival (A) or 105 RH parasites also i.p. for measure of parasite clearance (B). A, Percent survival of cps1-1-immunized ({diamondsuit}) or unimmunized ({square}) mice after challenge. B, Brains and spleens were harvested from ({diamondsuit}) cps1-1-immunized or ({square}) unimmunized naive at given time points after challenge and quantitative real-time PCR for parasite equivalents was performed. Survival data represented are the results from one experiment repeated three times with a n = 4 for each experiment. Quantitative PCR data are representative of two experiments with a n = 4 for each and are expressed as the mean ± SEM.

 
Intravenous immunization with cps1-1 tachyzoites alone or ex vivo cps1-1-infected DCs or PECs induce long-lasting protective immunity

In determining vaccine effectiveness, numerous studies have investigated different routes and cell-specific requirements for successful immunization (30, 35, 36). To explore the importance of route and cell type-specific effects on the ability of cps1-1 vaccination to elicit long-lasting immunity, mice were immunized i.v. once with cps1-1 alone, ex vivo cps1-1-infected DCs, PECs, or macrophages derived from resident PECs and groups of mice were challenged at 2 (Fig. 2A) or 6 mo (Fig. 2B) after immunization with a lethal dose of RH tachyzoites (LD100) and monitored for survival. Immunization with ex vivo cps1-1-infected DCs, PECs, or PEC-derived macrophages resulted in nearly complete survival of RH-challenged mice at 2 mo after immunization and the afforded protection was similar to that of mice immunized with tachyzoites of cps1-1 (Fig. 2A). In contrast, when lethal challenge was administered at 6 mo (Fig. 2B), significant differences in percent survival of mice immunized with ex vivo cps1-1-infected DCs (83% survival), PECs (50% survival), and PEC-derived macrophages (33% survival) were observed. Percent survival of mice immunized with ex vivo cps1-1-infected DCs or PECs was not statistically different from mice immunized i.v. with cps1-1 tachyzoites alone. Although peritoneal macrophages loaded with cps1-1 were effective at inducing short-term protection at 2 mo, they were significantly less effective at inducing long- lasting immunity (6 mo, p = 0.02; Fig. 2, A and B). Short-term protection to lethal RH challenge (LD1000) was also effectively induced following cps1-1 immunization of IL-15–/– mice (Fig. 2C).


Figure 2
View larger version (18K):
[in this window]
[in a new window]

  		FIGURE 2. Short-term and long-term immune protection induced by ex vivo cps1-1-loaded DCs, PECs, and peritoneal macrophages. A and B, B6 mice were unimmunized (*) or immunized i.v. once with either cps1–1 ({blacktriangleup}) alone, or DCs ({square}), PECs (x), or peritoneal macrophages ({circ}) loaded ex vivo with cps1-1. C, Il-15–/– mice (B6 background) were immunized i.p. with 106 cps1-1 parasites ({diamondsuit}) twice 14 days apart or not immunized ({square}). Mice were then challenged with a lethal dose of 102 RH i.p. for either 2 mo (A) or 6 mo (B) or a lethal dose of 103 RH i.p. (C) 21 days after immunization and percent survival was monitored for 30 days. A and B, 6 Month cps1-1-loaded macrophage survival was significantly lower than all others with p = 0.02. Data represent one experiment performed with six mice per immunization group and significant differences were calculated via the Kaplan-Meier product limit test. Survival data in C were repeated twice with n = 4/group.

 
Long-lasting immunity elicited by cps1-1 immunization is primarily Th1 biased, CD8+ T cell-mediated, and can be adoptively transferred

The Th1-biased inflammation produced by infection with T. gondii leads to primarily CD8+ T cell-mediated immunity and control of chronic infection (23, 26, 37, 38). To examine whether cps1-1-induced immunity is also CD8+ T cell mediated, we used Ab depletion of CD4+ and CD8+ T cell subsets from immunized mice and measured T cell-depleted mouse survival following lethal-type I RH challenge. In addition, we measured whether B lymphocytes were also involved as effectors in cps1-1-induced immunity by immunizing B cell-deficient mice (µMT) and then measuring their survival against lethal challenge. We observed that all of the anti-CD4+ and control Ig-treated mice survived, whereas only 25% of either CD8+ or CD8+ and CD4+ depleted mice survived the challenge infection (Fig. 3A). Although cps1-1-immunized µMT mice survived longer than nonimmunized naive µMT mice, all immunized µMT mice died by day 27 after challenge (Fig. 3B). These data are consistent with previous results indicating that B cells may be required as effector cells or are necessary for the development or recall of an effective memory CD8+ T cell population (39, 40, 41).


Figure 3
View larger version (17K):
[in this window]
[in a new window]

  		FIGURE 3. The effect of Ab depletion of T cells, lack of B cells, and adoptive transfer of immune cells on survival against lethal T. gondii challenge. B6 and µMT (B6) mice were immunized following our standard protocol. One month after the final immunization, (A) immunized B6 mice were treated with either control Ig ({square}) or Ab specific for CD4 ({diamondsuit}), CD8 ({circ}), or both CD4 and CD8 (x) while naive B6 ({triangleup}) were left untreated. B, Both immunized ({square}) and unimmunized ({diamondsuit}) µMT mice were left untreated. A and B, All mice were challenged with 103 RH parasites i.p. and percent survival was measured. C, B6 mice were immunized as described above and then 3 wk after the final immunization whole splenocytes, CD19+, and CD8+ splenocytes were harvested and either 4 x 107 total immune splenocytes (*), 1 x 107 CD8+T cells ({square}), 5 x 106 CD19+ B cells ({blacktriangleup}), or 4 x 107 total naive splenocytes ({circ}) were transferred to naive recipient mice. Twenty-four hours after transfer, mice were challenged with 103 RH parasites i.p. and monitored for survival. All experiments were performed with n = 4/group and repeated twice with similar results.

 
To confirm that cps1-1-induced CD8+ T cells were the primary effector mechanism of immune protection, we adoptively transferred naive total splenocytes or purified CD8+ T cells, CD19+ B cells, or total splenocytes from cps1-1- immunized mice into naive recipients and measured their survival following lethal-type I RH challenge. Mice receiving cps1-1 immune splenocytes, purified CD19+ B cells, or naive splenocytes succumbed by day 10 after RH challenge. By contrast, all naive mice receiving purified CD8+ T cells from cps1-1-immunized mice survived challenge infection (Fig. 3C). To determine whether cps1-1 immunization induced a Th1-biased response, we measured the anti-toxoplasma serum titer of IgG2a as an indicator of Th1-biased isotype switching (42, 43). Consistent with reports from other T. gondii vaccine models, high levels of IgG2a were measured (Table I) (30, 44, 45). Our results demonstrate that a Th1-biased CD8+ T cell-mediated immune response is the major mechanism required for the protective immunity induced by immunization with cps1-1.


View this table:
[in this window]
[in a new window]

  		Table I. Anti-toxoplasma-specific Ab serum titer after i.p. immunization with cps1-1

 
Inflammatory cells are recruited early into the peritoneum with a dynamic pattern in the absence of replicating and disseminating T. gondii infection

During T. gondii infection, specific inflammatory cells infiltrate into the site of infection and can indicate the type of immune response occurring and provide mechanisms of direct or indirect control of active infection (4, 16, 17, 46, 47). We examined the total PEC number and kinetics of specific inflammatory cell infiltration by flow cytometry in response to nonreplicating cps1-1 infection compared with RH infection (Fig. 4). After i.p. infection, a significant (p = 0.005) increase in total PEC numbers occurs by day 2 after cps1-1 inoculation followed by a steady increase (p < 0.001) through day 8 as compared with day 0 naive controls (Fig. 4B). By contrast, in RH infection an unexpected early decrease was followed by a greater after number of inflammatory cells infiltrating into the peritoneum (1.5-fold over cps1-1 (p = 0.002) by day 6 after infection (Fig. 4B).


Figure 4
View larger version (26K):
[in this window]
[in a new window]

  		FIGURE 4. Peritoneal exudate inflammatory cell recruitment in response to infection with cps1-1 as compared with highly virulent strain RH. B6 mice were infected i.p. with 106 cps1-1 ({diamondsuit}) or 103 RH ({square}) parasites and total PECs were harvested at days 0, 2, 4, 6, and 8. A, Representative typical dot plot of total PECs indicating three gated regions: R1, SSClowSClow lymphocytes; R2, SSCmidFSChigh monocytes; and R3, SSChighFSClow granulocytes. B, Total PECs numbers were counted, then the composition of PECs was analyzed by flow cytometry and absolute numbers were calculated. C, Granulocytes by GR1+CD68– in R3; D, macrophages by CD68+; E, inflammatory macrophages by GR1+CD68+; F, B lymphocytes by CD19; G, total T lymphocytes by CD3+; H, CD3+CD4+ T lymphocytes; and I, CD3+CD8+ T lymphocytes. Flow cytometric data represent the mean ± SEM from a n = 4 and was repeated twice. Significant differences for cps1-1 ({blacksquare}) and RH (•) are p < 0.05.

 
Inflammatory cells such as granulocytes (polymorphonuclear neutrophils), macrophages, and B and T lymphocytes are important for the development of protective immunity and for direct control of T. gondii infection (1, 13, 16, 47). To determine what cell types may be contributing to the development of immunity, we used flow cytometry-based differential cell analysis of PECs from mice infected with either cps1-1 or RH. We characterized three general cell populations composing total PECs (Fig. 4A). In cps1-1 infection, we observed a significant increase in the absolute number of GR1+CD68– granulocytes (p = 0.03) by day 2 after inoculation with a 9-fold increase, after which GR1+CD68– granulocytes return to uninfected control levels. By contrast, during RH infection, the absolute numbers of GR1+CD68– granulocytes did not rise until day 4 (day 0 vs day 4, 6.8-fold, p = 0.001) and reached maximal numbers at day 6 with a 6.0-fold increase from day 4 (p = 0.001; Fig. 4C). Measurement of the percentage of total PECs and direct examination of Diff-Quik- stained cytospins confirmed these results for both cps1-1 (day 2) and RH (days 4–6) infections and the latter reveal by nuclear morphology that these GR1+CD68– cells are neutrophils (data not shown).

We measured the kinetics of CD68+ cells (resident peritoneal macrophages and monocytes) and GR1+CD68+ inflammatory monocytes infiltrating after cps1-1 or RH infection. The absolute numbers of CD68+ cells infiltrating into the peritoneum with cps1-1 infection increases 1.5-fold (p < 0.001) by day 4 and a 2.1-fold (p = 0.001) from day 0 controls by day 6 after infection (Fig. 4D). Even though early after RH infection the absolute numbers of CD68+ cells decreased, their numbers significantly increased through day 6 (day 4, p < 0.001; day 6, p = 0.03) after infection (Fig. 4D). However, the rate of influx measured as CD68+ cells as a percentage of total PECs for either cps1-1 or RH infection did not significantly change over time (data not shown). Analysis of the CD68+ population revealed an interesting pattern of cellular infiltration. Two days after cps1-1 infection, we observed a 168-fold increase (p < 0.001) in the absolute numbers of GR1+CD68+ inflammatory monocytes (Fig. 4E). This rapid increase was followed by a 2.5-fold decrease (p < 0.001) by day 4, then a second stage of GR1+CD68+ monocytes infiltrated into the site of cps1-1 inoculation at day 6 (2.5-fold increase, p = 0.002; Fig. 4E). In RH infection, GR1+CD68+ inflammatory monocyte infiltration is delayed until day 4 (p < 0.001) and increases continuously through day 8 after infection (Fig. 4E). The >99% replacement of total CD68+ macrophages (Fig. 4, D and E) suggests some cells may express GR1+ in response to infection; however, in vitro infection of peritoneal-derived macrophages with cps1-1 under replicating or nonreplicating conditions did not elicit the expression of GR1 (data not shown).

Analysis of CD19+ B lymphocytes reveals a similar pattern of flux for either cps1-1 or RH inoculation only through day 4 after infection, after which the pattern is distinctly different. The absolute number of CD19+ B cells increases 2-fold (p = 0.001) by day 6 and is maintained after cps1-1 infection, whereas by day 6 with RH infection, a steady reduction in absolute numbers occurs (day 4, p = 0.01). This reduction continues to near undetectable levels by day 8 after RH infection (Fig. 4F).

When measuring total T cells by CD3+ as well as CD3+CD4+ and CD3+CD8+ T cell flux, our analyses revealed that the absolute number of total CD3+ T cells after cps1-1 infection significantly increases early in the peritoneum 1.9-fold (p = 0.02) by day 2 after infection and continues to increase through day 8 after infection (Fig. 4G). By contrast, total CD3+ T cells did not increase significantly until day 4 after RH infection (Fig. 4G). We observed that despite lower total PEC numbers after cps1-1 infection, the absolute numbers of CD3+ T cells were significantly greater (day 2, p = 0.004; day 4, p < 0.001; day 6, p < 0.001; day 8, p < 0.001) than during RH infection at each time point (Fig. 4G). The absolute numbers of CD3+CD4+ T cells markedly increase in numbers at day 6 after cps1-1 infection (Fig. 4H). Interestingly, CD3+CD8+ T cells increase in absolute numbers earlier than CD3+CD4+ T cells at day 2 (2.8-fold, p = 0.007) and day 4 (2.2-fold, p = 0.01) after cps1-1 infection, after which absolute numbers do not significantly change (Fig. 4I). By contrast, during RH infection, CD3+CD8+ T cells have the greatest increase in absolute numbers at the site of infection while CD3+CD4+ T cells remain nearly unchanged (Fig. 4, H and I).

Systemic production of IFN-{gamma} is low and transient after cps1-1 inoculation

Previous studies of immunity using replicating and disseminating T. gondii strains revealed high-level systemic production of IFN-{gamma}, IL-12p40, and variable levels of IL-12p70 production that is dependent on parasite genotype (types I, II, and III) (5, 11, 47). Replication and dissemination during acute infection is likely to enhance the lethal overproduction of inflammatory cytokines (48, 49). Therefore, the level of proinflammatory cytokines necessary for the development of immunity is unclear. We measured systemic (serum) levels of proinflammatory Th1 cytokines IL-12p40, IL-12p70, and IFN-{gamma} by ELISA in mice infected with cps1-1 or RH at days 0, 2, 4, 6, and 8 after infection. Infection with replicating and disseminating T. gondii is known to result in enhanced inflammation arising from host cell lysis and tissue damage during acute infection (11, 48, 50). As shown in Fig. 5, A and C, systemic IFN-{gamma} and IL-12p40 in serum of mice infected with RH did not significantly increase until day 4 after infection and quickly rose to extremely high levels by days 6 and 8. Confirming previous reports, systemic IL-12p70 was not significantly produced in response to RH over the course of infection (Fig. 5B) (11). By contrast, systemic IFN-{gamma} is detected at a very low level only early (day 2, p = 0.03; Fig. 5A) and is not detected systemically after day 2 after cps1-1 infection and through day 8 after cps1-1 infection. IL-12p70 production was induced early (day 2) and continued to increase through day 6 (p = 0.04) after cps1-1 infection (Fig. 5B) and was significantly higher than in response to RH (p < 0.001) for the time points of days 4, 6, and 8 after infection. Interestingly, essentially all systemic IL-12 produced in response to cps1-1 was in the form of bioactive IL-12p70 (Fig. 5, B and C). By days 6 and 8 after RH infection, little bioactive IL-12p70 was present even though a large excess of IL-12p40 was detected systemically (Fig. 5, B and C). Additionally, although systemic IFN-{gamma} was only detectable at a low level day 2 after infection, IFN-{gamma} is essential for immunity in this vaccine model (Fig. 5D).


Figure 5
View larger version (14K):
[in this window]
[in a new window]

  		FIGURE 5. Systemic Th1 cytokine production in response to infection with cps1-1 or RH and requirement of IFN-{gamma} in development of adaptive immunity. B6 mice were infected i.p. with 106 cps1-1 ({diamondsuit}) or 103 RH ({square}) parasites and sera were taken at days 0, 2, 4, 6, and 8. Serum levels of IFN-{gamma} (A) (note log scale in A is different than in B and C), IL-12p70 (B), and IL-12p40 (C) were measured by ELISA. D, Groups of six WT or IFN-{gamma}–/– mice were infected i.p. with a single dose of 106 cps1-1 parasites or were unimmunized. Thirty days later, immunized WT ({diamondsuit}), IFN-{gamma}–/– ({square}), or unimmunized IFN-{gamma}–/– ({circ}) were challenged with LD100 (200) of RH tachyzoites. Experiments shown in A–C were performed with n = 4 mice/group. The data presented here are representative of two experiments with similar results and indicate the mean ± SEM with p < 0.05 as calculated by Student’s t test and denoted by cps1-1 ({blacksquare}) and RH (•).

 
IFN-{gamma} and IL-12p70 production is primarily localized to the site of cps1-1 inoculation

Because systemic production of IFN-{gamma} was low and transient after cps1-1 infection, we measured the kinetics of local (PECs) and peripheral (spleen) IFN-{gamma} and IL-12p70 production induced by cps1-1 or RH infection. PECs from mice infected with RH produced moderate IFN-{gamma} detectable (p < 0.001) at day 4, which was maintained through day 8 after infection (Fig. 6A). Moderate levels of IL-12p70 were produced by PECs from RH-infected mice beginning on day 4, then decreasing through day 8 (Fig. 6C). Splenocytes from RH-infected mice produced IFN-{gamma} at manifold greater levels than PECs (Fig. 6, A and B) as previously observed (16, 48). IFN-{gamma} production by splenocytes significantly increased by day 4 after infection (day 4 vs day 2, p = 0.001) and continued to increase through day 8 (Fig. 6B). Minimal splenocyte production of IL-12p70 was observed on days 4 and 6 after RH infection (Fig. 6D).


Figure 6
View larger version (16K):
[in this window]
[in a new window]

  		FIGURE 6. Local and peripheral Th1 cytokine production in response to infection with cps1-1 or RH. B6 mice were infected i.p. with 106 cps1-1 ({diamondsuit}) or 103 RH ({square}) parasites and total PECs or splenocytes were taken at days 0, 2, 4, 6, and 8. PECs (A and C) were plated at 1 x 106 cells/ml, splenocytes (B and D) at 5 x 106 cells/ml, and cultured for 24 h. Supernatants were then assayed for IFN-{gamma} (A and B) and IL-12p70 (C and D) by ELISA. All experiments were performed with n = 4 mice/group. The data presented here are representative of two experiments with similar results and indicate the mean ± SEM with p < 0.05 as calculated by Student’s t test and denoted by cps1-1 ({blacksquare}) and RH (•).

 
After cps1-1 infection, a novel and primarily local pattern of cytokine production is observed (Fig. 6). PEC IFN-{gamma} production was detectable as early as day 2 and was maximal by day 6 (day 4 vs day 6, p = 0.04) followed by a rapid decrease by day 8 after cps1-1 infection. Interestingly, PECs from cps1-1-infected mice produced significantly greater IFN-{gamma} (p = 0.005) than PECs from RH-infected mice by day 6 after infection (Fig. 6A). In Fig. 6C, we reveal that PECs from cps1-1-infected mice rapidly produce significantly greater levels of IL-12p70 by day 2 after infection as compared with RH (p = 0.007). These IL-12p70 levels remained consistently high (days 2–8) after cps1-1 inoculation, whereas IL-12p70 production from PECs after RH infection was delayed to day 4 and significantly lower by days 6 and 8 with p = 0.012 and p = 0.001, respectively (Fig. 6C). PEC cytokine production, on a per cell basis, of IFN-{gamma} (Fig. 6, A and B) and IL-12p70 (Fig. 6, C and D) was higher than that of splenocytes from cps1-1-infected mice. In contrast to hypervirulent RH infection, Th1 cytokines including IL-12p70 were produced more rapidly and at greater levels locally in response to cps1-1 infection.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
T. gondii infection elicits a Th1-biased inflammatory response leading to the control of both acute and chronic infection and establishing lifelong CD8+ T cell-mediated immunity (2, 3, 23, 37, 38, 51). Understanding development of immunity is essential for vaccine and therapeutic development to combat infection and to prevent congenital transmission or toxoplasmic encephalitis in the immune-compromised individual. Although the IL-12 pathways triggered by MyD88 in the innate response are prominent, recent data with the cps1-1 infection model suggests that MyD88 is not required for the development of adaptive immunity (52). CD4+ and CD8+ T cells are activated by DC-, macrophage-, and neutrophil- produced IL-12 and recognition of Ag. Once activated, they produce IFN-{gamma} and bias the host response toward a Th1 inflammatory environment which leads to the development of immune protection mediated primarily by memory CD8+ T cells (2, 3, 5, 7, 51, 53). The only definitive mediators of T. gondii infection that have been proven to be absolutely essential for immunity are CD8+ T cells and IFN-{gamma}.

Despite much effort, the mechanisms by which lifelong CD8+ T cell-mediated immunity develops is still not well defined. Because type I strains are universally lethal, only the replicating yet slowed ts-4 and type II strain infections result in the development of protective immunity (54, 55). Replicating parasites lyse host cells, disseminate, and lead to extensive host tissue destruction and enhanced inflammation (48, 49). Enhanced inflammation and lethality of replication competent parasites in immune-compromised hosts have greatly complicated the study of host immune responses specifically required for the development of immunity (5, 7, 16, 47, 48, 49, 56). Current evidence supports that only live parasites in contrast to dead parasites are capable of inducing immunity, suggesting that active invasion of the parasite is required for development of immunity (8, 9). In contrast, the critically attenuated and completely avirulent-type I parasite cps1-1 elicits a potent and long-lasting immunity. Consequently, the cps1-1 vaccine model provides an excellent nonreplicating model to investigate immunity to T. gondii in the absence of parasite replication (1). Despite several highly important studies focused on intracellular host control of T. gondii infection using the cps1-1 model, how the host responds and the type of immunity elicited remains unknown (1, 18, 34, 57). In this study, we investigated the mechanism of protective immunity elicited by the vaccine strain cps1-1. Our investigation indicates that the Th1-biased inflammatory response induced by cps1-1 infection is primarily local. We also reveal that an attenuated type I parasite strain can induce and sustain significant IL-12p70 production. Our analysis of cell infiltration into the peritoneum indicates that there is early innate inflammatory cell infiltration marked by GR1+CD68– neutrophil and GR1+CD68+ inflammatory monocytes. GR1+CD68+ inflammatory monocytes are recruited in two separate waves and completely replace resident peritoneal macrophages in the first wave. CD8+ T cells are recruited earlier than CD4+ T cells and recruitment of both T cell subsets corresponds with production of local IFN-{gamma}. We find that DCs are more capable of eliciting long-lasting immunity than resident peritoneal macrophages when loaded ex vivo with cps1-1. We also established that the development of cps1-1-induced immunity requires B cells. Although cps1-1 only induced a limited Th1-biased inflammatory response, infected mice still developed a highly effective and long-lasting CD8+ T cell-mediated immunity protection capable of clearing a high lethal dose virulent challenge. IL-15–/– mice were also completely protected from lethal RH challenge after short-term immunization with cps1-1. It will be interesting to determine whether cps1-1 has any ability to elicit long-lasting immunity in IL-15-deficient mice.

Control of acute infection and long-lasting immunity against chronic and recurrent T. gondii infection is dependent on Th1-biased inflammation and memory CD8+ T cells (5, 23, 26, 51, 55). Since we observed that cps1-1 immunization protects against virulent challenge for at least 18 mo, this suggests that a Th1-biased immunity and T cell memory was elicited in response to cps1-1 infection. We demonstrate this to be the case because depletion of CD8+ T cells from cps1-1-immunized mice abrogates immune protection against virulent challenge. In addition, we demonstrated that adoptive transfer of purified CD8+ T cells from cps1-1-immunized mice to naive recipients conveys immune protection against virulent challenge. cps1-1 immunization also elicits Th1-biased serum titers of IgG2a.

DCs are required for development of immunity to T. gondii (21, 50). A single i.v. immunization with ex vivo cps1-1-infected DCs, but not peritoneal macrophages, provides immune protection against virulent challenge at 6 mo. These results support the hypothesis that DCs are required for the development of long-lasting immunity. One reason the longevity of immunity is enhanced by DCs could be through better stimulation of immune responses. Evidence from other studies support this role for DCs since mice immunized with parasite extract alone do not develop robust immunity while parasite extract-exposed DCs enhance the immunity against type II challenge infection (36). Ex vivo infection of DCs with cps1-1 could result in high levels of IL-12p70 production and more efficient presentation of T. gondii Ags enhancing the activation of T cells. Alternatively, the i.v. route may deliver parasite-containing DCs directly to the immunogenic environment of the spleen, promoting DC-T cell interactions and development of T. gondii-specific immunity. Since we observed that cps1-1 administered alone i.v. induces protective immunity in a single inoculation, Ag processing and presentation is likely to be more efficient when infection is delivered directly into an immunogenic site.

Recently, GR1+CD68+ monocytes were shown to infiltrate into the site of infection and appear to be required for control of infection (16, 17). In addition to GR1+CD68+ monocytes, neutrophils are also important for early control of T. gondii infection (13, 58). These studies suggest that the inflammation induced by high parasite numbers stimulates the recruitment of these cells. Infection with a high number of cps1-1 tachyzoites recruits GR1+CD68+ monocytes and GR1+CD68– neutrophils early and GR1+CD68+ monocytes infiltrate in two distinct waves, completely replacing the resident peritoneal macrophages early after infection. The recruited neutrophils are most likely secreting early IL-12; however, the role GR1+CD68+ monocytes could play is unknown. They could act as innate effector cells by providing IL-12p40 and IL-12p70 or may have the ability to destroy intracellular parasites by autophagy (13, 17, 18). Interestingly, we observed during virulent RH infection that both GR1+CD68– neutrophils and GR1+CD68+ macrophages appear later and that the GR1+CD68+ macrophages compose >90% of the total macrophages present. These data suggest that during RH infection innate responses fail to establish adaptive immunity. The low levels of systemic IL-12p70 in RH infection may be due to the lack of early granulocyte and GR1+CD68+ macrophage infiltration.

Beside innate responses playing a key role in controlling the acute T. gondii infection, adaptive immunity mediated by B cells, CD4+ cells, and CD8+ T cells is important for long-term control of recurrent and chronic infection (23, 26, 39, 45, 55). Influx of lymphocytes into the site of inoculation has not been well defined with the i.p. model of inoculation. B cells may be required for optimal Th1 responses enhancing CD8+ T cell responses in mice and by providing IL-12p70 in humans (40, 41, 59). Interestingly, we found that B cells are required for the development of cps1-1-induced immune protection as cps1-1-immunized µMT mice survived longer, but universally succumbed to infection. B cells have been implicated for T. gondii and other disease models to act as effectors by producing Ab, to enhance CD8+ T cell memory by production of IL-12p70 or interaction with the T cells, or to be essential for CD8+ T memory cell recall (39, 40, 41, 45, 60, 61, 62). We found that CD19+ B cells infiltrate into the site of infection with cps1-1 where they remain in contrast to replicating RH infection where they are cleared, suggesting that B cell loss during a virulent infection could decrease the host’s ability to respond to and control infection and acute infection and to diminish the development of protective immunity.

Our data also indicate that CD8+ T cells respond more rapidly to cps1-1 at the site of infection than CD4+ T cells and could therefore in an Ag-dependent manner establish some control of the parasite through an IL-12p70-dependent production of IFN-{gamma} (5, 8). Recent studies support our hypothesis in that in the absence of IL-12p70 (p35–/– mice) cps1-1 infection results in a lack of the dominant effector CD8 T cell population (63). However, we have no direct evidence that these early CD8+ T cells are activated or capable of killing infected cells. At later times during cps1-1 infection, we show that CD4+ T cells dominate the T lymphocyte population and could provide CD8+ T cell help. A subpopulation of CD4+ T cells could also represent T regulatory cells helping to control the inflammatory response. Further studies are needed to determine how rapidly CD8+ T cells can respond to cps1-1 infection in an Ag-specific manner and also to more clearly determine the phenotypes and role(s) of CD4+ T cells in the development of CD8+ T cell immunity in the cps1-1 vaccine model.

Type I and type II infections induce high systemic levels of IFN-{gamma} and IL-12 necessary for control of the acute T. gondii infection and development of immunity (7, 48, 49, 64, 65, 66). We show that development of immunity after cps1-1 infection is dependent on IFN-{gamma}. Paradoxically, production of systemic IFN-{gamma} is very low and transient after cps1-1 infection. Interestingly, we find that cps1-1 elicited significant levels of locally produced IFN-{gamma} and IL-12p70 and that, of these cytokines, only IL-12p70 is detected at higher levels systemically. Our results suggest that systemic production of inflammatory cytokines in response to infection with replicating parasites may be enhanced by widespread host tissue destruction during acute infection. Our results using the cps1-1 vaccine model show that only minor systemic levels and mostly local production of these cytokines is sufficient for the development of long-lasting CD8+ T cell-dependent immunity.

Recent data suggest that the IL-12p70 heterodimer is only produced in response to type II infections and is not significantly produced by type I infections (11). Our data confirm that virulent-type I RH parasites induce limited amounts of IL-12p70. In contrast to RH, our data demonstrate that the attenuated type I parasite cps1-1 induces high systemic as well as significant local production of IL-12p70. One reason IL-12p70 could be induced by cps1-1 and not during RH infection is that nonreplicating parasites may fail to elicit a stress or other replication-dependent response in the infected host cell that disrupts IL-12p70 production. Alternatively, during pyrimidine starvation cps1-1 may express genes and gene products not expressed by replicating RH that stimulates production of IL-12p70.

Proper IL-12p70 secretion requires intracellular assembly of the IL-12p35 and IL-12p40 subunits (67). IL-12p40 is the regulated subunit and is produced in excess after cellular stimulation. RH infection results in high levels of excess IL-12p40 production systemically, while paradoxically excess IL-12p40 is not produced systemically during cps1-1 infection. Secreted IL-12p40 is reported to be agonistic as well as antagonistic by extracellularly forming an IL-12p80 homodimer that can either recruit cells or inhibit IL-12 receptor binding of the bioactive IL-12p70 heterodimer and in both cases enhance the inflammatory response induced by virulent parasites (68, 69). In the absence of cps1-1 parasite replication, less secreted IL-12p40 monomer may be produced because of limited host cell stimulation compared with replicating RH infection where parasite numbers continue to drastically increase. Clearly, our results demonstrate that the inability of replicating type I parasites to stimulate IL-12p70 production is not due to the absence of a parasite molecule that type II strains possess (10, 70).

Recent studies suggest that IL-12 controls the contraction phase of the CD8 T cell response and therefore controls the level of memory T cell development (71). Additionally, recent studies reveal an essential role of IL-12p70 in immunity induced by cps1-1. One study shows that in IL-12p35–/– mice, CD62LlowKLRG1+ CD8 effector T cells do not develop in response to cps1-1 infection (63). Another study shows that IL-12p40 is required for protective immunity in the cps1-1 infection model (52). Therefore, cps1-1 may induce sufficient IL-12p70 in the absence of potentially inhibitory excess IL-12p40 leading to effective long-lasting CD8 T cell immunity.

In this study, we reveal that the long-term protective immunity induced by immunization with cps1-1 clears virulent RH challenge parasites, is induced most efficiently by the i.v. route and by DCs, is dependent on CD8+ T cells, and is biased toward a Th1 response based on local production of IL-12p70 and IFN-{gamma}. Using cps1-1, we also provide the first kinetic model of the host response specific to only parasite infection in the absence of significant parasite replication. These results reveal that local and relatively limited systemic inflammation is sufficient for the development of potent CD8+ T cell-mediated immunity. This study more clearly establishes the cps1-1 vaccine model as an effective tool to further elucidate the host immune response specific to only parasite infection without inflammatory complications caused by parasite replication-associated host tissue destruction.


   Acknowledgments
 
We thank Dr. Edward J. Usherwood, Dr. William F. Wade, and Dr. Imtiaz A. Khan for critical review of this 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 National Institutes of Health Grants AI-41930 and J.P.G. was supported by National Institutes of Health Grants T32 AI07363-13 and T32 AI007519. Back

2 Address correspondence and reprint requests to Dr. David J. Bzik, Department of Microbiology and Immunology, Dartmouth Medical School, Room 654, East Borwell Building, 1 Medical Center Drive, Lebanon, NH 03756. E-mail address: David.J.Bzik{at}Dartmouth.edu Back

3 Abbreviations used in this paper: DC, dendritic cell; PEC, peritoneal exudate cell; WT, wild type. Back

Received for publication October 24, 2007. Accepted for publication November 9, 2008.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Fox, B. A., D. J. Bzik. 2002. De novo pyrimidine biosynthesis is required for virulence of Toxoplasma gondii. Nature 415: 926-929. [Medline]
  2. Mun, H. S., F. Aosai, K. Norose, M. Chen, L. X. Piao, O. Takeuchi, S. Akira, H. Ishikura, A. Yano. 2003. TLR2 as an essential molecule for protective immunity against Toxoplasma gondii infection. Int. Immunol. 15: 1081-1087. [Abstract/Free Full Text]
  3. Scanga, C. A., J. Aliberti, D. Jankovic, F. Tilloy, S. Bennouna, E. Y. Denkers, R. Medzhitov, A. Sher. 2002. Cutting edge: MyD88 is required for resistance to Toxoplasma gondii infection and regulates parasite-induced IL-12 production by dendritic cells. J. Immunol. 168: 5997-6001. [Abstract/Free Full Text]
  4. Bennouna, S., S. K. Bliss, T. J. Curiel, E. Y. Denkers. 2003. Cross-talk in the innate immune system: neutrophils instruct recruitment and activation of dendritic cells during microbial infection. J. Immunol. 171: 6052-6058. [Abstract/Free Full Text]
  5. Scharton-Kersten, T., P. Caspar, A. Sher, E. Y. Denkers. 1996. Toxoplasma gondii: evidence for interleukin-12-dependent and-independent pathways of interferon-{gamma} production induced by an attenuated parasite strain. Exp. Parasitol. 84: 102-114. [Medline]
  6. Aliberti, J., C. Reis e Sousa, M. Schito, S. Hieny, T. Wells, G. B. Huffnagle, A. Sher. 2000. CCR5 provides a signal for microbial induced production of IL-12 by CD8{alpha}+ dendritic cells. Nat. Immunol. 1: 83-87. [Medline]
  7. 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]
  8. Subauste, C. S., M. Wessendarp. 2000. Human dendritic cells discriminate between viable and killed Toxoplasma gondii tachyzoites: dendritic cell activation after infection with viable parasites results in CD28 and CD40 ligand signaling that controls IL-12-dependent and -independent T cell production of IFN-{gamma}. J. Immunol. 165: 1498-1505. [Abstract/Free Full Text]
  9. Butcher, B. A., E. Y. Denkers. 2002. Mechanism of entry determines the ability of Toxoplasma gondii to inhibit macrophage proinflammatory cytokine production. Infect. Immun. 70: 5216-5224. [Abstract/Free Full Text]
  10. Kim, L., B. A. Butcher, C. W. Lee, S. Uematsu, S. Akira, E. Y. Denkers. 2006. Toxoplasma gondii genotype determines MyD88-dependent signaling in infected macrophages. J. Immunol. 177: 2584-2591. [Abstract/Free Full Text]
  11. Robben, P. M., D. G. Mordue, S. M. Truscott, K. Takeda, S. Akira, L. D. Sibley. 2004. Production of IL-12 by macrophages infected with Toxoplasma gondii depends on the parasite genotype. J. Immunol. 172: 3686-3694. [Abstract/Free Full Text]
  12. Khan, I. A., P. M. Murphy, L. Casciotti, J. D. Schwartzman, J. Collins, J. L. Gao, G. R. Yeaman. 2001. Mice lacking the chemokine receptor CCR1 show increased susceptibility to Toxoplasma gondii infection. J. Immunol. 166: 1930-1937. [Abstract/Free Full Text]
  13. Bliss, S. K., B. A. Butcher, E. Y. Denkers. 2000. Rapid recruitment of neutrophils containing prestored IL-12 during microbial infection. J. Immunol. 165: 4515-4521. [Abstract/Free Full Text]
  14. Bliss, S. K., L. C. Gavrilescu, A. Alcaraz, E. Y. Denkers. 2001. Neutrophil depletion during Toxoplasma gondii infection leads to impaired immunity and lethal systemic pathology. Infect. Immun. 69: 4898-4905. [Abstract/Free Full Text]
  15. Del Rio, L., S. Bennouna, J. Salinas, E. Y. Denkers. 2001. CXCR2 deficiency confers impaired neutrophil recruitment and increased susceptibility during Toxoplasma gondii infection. J. Immunol. 167: 6503-6509. [Abstract/Free Full Text]
  16. Mordue, D. G., L. D. Sibley. 2003. A novel population of Gr-1+-activated macrophages induced during acute toxoplasmosis. J. Leukocyte Biol. 74: 1015-1025. [Abstract/Free Full Text]
  17. Robben, P. M., M. LaRegina, W. A. Kuziel, L. D. Sibley. 2005. Recruitment of Gr-1+ monocytes is essential for control of acute toxoplasmosis. J. Exp. Med. 201: 1761-1769. [Abstract/Free Full Text]
  18. Ling, Y. M., M. H. Shaw, C. Ayala, I. Coppens, G. A. Taylor, D. J. Ferguson, G. S. Yap. 2006. Vacuolar and plasma membrane stripping and autophagic elimination of Toxoplasma gondii in primed effector macrophages. J. Exp. Med. 203: 2063-2071. [Abstract/Free Full Text]
  19. Zhao, Y., D. Wilson, S. Matthews, G. S. Yap. 2007. Rapid elimination of Toxoplasma gondii by {gamma} interferon-primed mouse macrophages is independent of CD40 signaling. Infect. Immun. 75: 4799-4803. [Abstract/Free Full Text]
  20. Schulz, O., A. D. Edwards, M. Schito, J. Aliberti, S. Manickasingham, A. Sher, C. Reis e Sousa. 2000. CD40 triggering of heterodimeric IL-12 p70 production by dendritic cells in vivo requires a microbial priming signal. Immunity 13: 453-462. [Medline]
  21. Liu, C. H., Y. T. Fan, A. Dias, L. Esper, R. A. Corn, A. Bafica, F. S. Machado, J. Aliberti. 2006. Cutting edge: dendritic cells are essential for in vivo IL-12 production and development of resistance against Toxoplasma gondii infection in mice. J. Immunol. 177: 31-35. [Abstract/Free Full Text]
  22. Casciotti, L., K. H. Ely, M. E. Williams, I. A. Khan. 2002. CD8+-T-cell immunity against Toxoplasma gondii can be induced but not maintained in mice lacking conventional CD4+ T cells. Infect. Immun. 70: 434-443. [Abstract/Free Full Text]
  23. Suzuki, Y., J. S. Remington. 1988. Dual regulation of resistance against Toxoplasma gondii infection by Lyt-2+ and Lyt-1+, L3T4+ T cells in mice. J. Immunol. 140: 3943-3946. [Abstract]
  24. Kitamura, D., J. Roes, R. Kuhn, K. Rajewsky. 1991. A B cell-deficient mouse by targeted disruption of the membrane exon of the immunoglobulin µ chain gene. Nature 350: 423-426. [Medline]
  25. Villegas, E. N., M. M. Elloso, G. Reichmann, R. Peach, C. A. Hunter. 1999. Role of CD28 in the generation of effector and memory responses required for resistance to Toxoplasma gondii. J. Immunol. 163: 3344-3353. [Abstract/Free Full Text]
  26. Gazzinelli, R. T., F. T. Hakim, S. Hieny, G. M. Shearer, A. Sher. 1991. Synergistic role of CD4+ and CD8+ T lymphocytes in IFN-{gamma} production and protective immunity induced by an attenuated Toxoplasma gondii vaccine. J. Immunol. 146: 286-292. [Abstract]
  27. El-Malky, M., L. Shaohong, T. Kumagai, Y. Yabu, M. S. Noureldin, N. Saudy, H. Maruyama, N. Ohta. 2005. Protective effect of vaccination with Toxoplasma lysate antigen and CpG as an adjuvant against Toxoplasma gondii in susceptible C57BL/6 mice. Microbiol. Immunol. 49: 639-646. [Medline]
  28. Da Gama, L. M., F. L. Ribeiro-Gomes, U. Guimaraes, Jr, A. C. 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]
  29. Obar, J. J., S. G. Crist, E. K. Leung, E. J. Usherwood. 2004. IL-15-independent proliferative renewal of memory CD8+ T cells in latent gammaherpesvirus infection. J. Immunol. 173: 2705-2714. [Abstract/Free Full Text]
  30. Bourguin, I., M. Moser, D. Buzoni-Gatel, F. Tielemans, D. Bout, J. Urbain, O. Leo. 1998. Murine dendritic cells pulsed in vitro with Toxoplasma gondii antigens induce protective immunity in vivo. Infect. Immun. 66: 4867-4874. [Abstract/Free Full Text]
  31. Ely, K. H., L. H. Kasper, I. A. Khan. 1999. Augmentation of the CD8+ T cell response by IFN-{gamma} in IL-12-deficient mice during Toxoplasma gondii infection. J. Immunol. 162: 5449-5454. [Abstract/Free Full Text]
  32. Obar, J. J., D. C. Donovan, S. G. Crist, O. Silvia, J. P. Stewart, E. J. Usherwood. 2004. T-cell responses to the M3 immune evasion protein of murid gammaherpesvirus 68 are partially protective and induced with lytic antigen kinetics. J. Virol. 78: 10829-10832. [Abstract/Free Full Text]
  33. Kirisits, M. J., E. Mui, R. McLeod. 2000. Measurement of the efficacy of vaccines and antimicrobial therapy against infection with Toxoplasma gondii. Int. J. Parasitol. 30: 149-155. [Medline]
  34. Shaw, M. H., G. J. Freeman, M. F. Scott, B. A. Fox, D. J. Bzik, Y. Belkaid, G. S. Yap. 2006. Tyk2 negatively regulates adaptive Th1 immunity by mediating IL-10 signaling and promoting IFN-{gamma}-dependent IL-10 reactivation. J. Immunol. 176: 7263-7271. [Abstract/Free Full Text]
  35. McLeod, R., J. K. Frenkel, R. G. Estes, D. G. Mack, P. B. Eisenhauer, G. Gibori. 1988. Subcutaneous and intestinal vaccination with tachyzoites of Toxoplasma gondii and acquisition of immunity to peroral and congenital toxoplasma challenge. J. Immunol. 140: 1632-1637. [Abstract]
  36. Aline, F., D. Bout, S. Amigorena, P. Roingeard, I. Dimier-Poisson. 2004. Toxoplasma gondii antigen-pulsed-dendritic cell-derived exosomes induce a protective immune response against T. gondii infection. Infect. Immun. 72: 4127-4137. [Abstract/Free Full Text]
  37. Subauste, C. S., A. H. Koniaris, J. S. Remington. 1991. Murine CD8+ cytotoxic T lymphocytes lyse Toxoplasma gondii-infected cells. J. Immunol. 147: 3955-3959. [Abstract]
  38. Hakim, F. T., R. T. Gazzinelli, E. Denkers, S. Hieny, G. M. Shearer, A. Sher. 1991. CD8+ T cells from mice vaccinated against Toxoplasma gondii are cytotoxic for parasite-infected or antigen-pulsed host cells. J. Immunol. 147: 2310-2316. [Abstract]
  39. Sayles, P. C., G. W. Gibson, L. L. Johnson. 2000. B cells are essential for vaccination-induced resistance to virulent Toxoplasma gondii. Infect. Immun. 68: 1026-1033. [Abstract/Free Full Text]
  40. Langhorne, J., C. Cross, E. Seixas, C. Li, T. von der Weid. 1998. A role for B cells in the development of T cell helper function in a malaria infection in mice. Proc. Natl. Acad. Sci. USA 95: 1730-1734. [Abstract/Free Full Text]
  41. Matter, M., S. Mumprecht, D. D. Pinschewer, V. Pavelic, H. Yagita, S. Krautwald, J. Borst, A. F. Ochsenbein. 2005. Virus-induced polyclonal B cell activation improves protective CTL memory via retained CD27 expression on memory CTL. Eur. J. Immunol. 35: 3229-3239. [Medline]
  42. Snapper, C. M., W. E. Paul. 1987. Interferon-{gamma} and B cell stimulatory factor-1 reciprocally regulate Ig isotype production. Science 236: 944-947. [Abstract/Free Full Text]
  43. Sornasse, T., V. Flamand, G. De Becker, H. Bazin, F. Tielemans, K. Thielemans, J. Urbain, O. Leo, M. Moser. 1992. Antigen-pulsed dendritic cells can efficiently induce an antibody response in vivo. J. Exp. Med. 175: 15-21. [Abstract/Free Full Text]
  44. Waldeland, H., E. R. Pfefferkorn, J. K. Frenkel. 1983. Temperature-sensitive mutants of Toxoplasma gondii: pathogenicity and persistence in mice. J. Parasitol. 69: 171-175. [Medline]
  45. Johnson, L. L., P. C. Sayles. 2002. Deficient humoral responses underlie susceptibility to Toxoplasma gondii in CD4-deficient mice. Infect. Immun. 70: 185-191. [Abstract/Free Full Text]
  46. Kelly, M. N., J. K. Kolls, K. Happel, J. D. Schwartzman, P. Schwarzenberger, C. Combe, M. Moretto, I. A. Khan. 2005. Interleukin-17/interleukin-17 receptor-mediated signaling is important for generation of an optimal polymorphonuclear response against Toxoplasma gondii infection. Infect. Immun. 73: 617-621. [Abstract/Free Full Text]
  47. Scharton-Kersten, T. M., T. A. Wynn, E. Y. Denkers, S. Bala, E. Grunvald, S. Hieny, 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]
  48. Gavrilescu, L. C., E. Y. Denkers. 2001. IFN-{gamma} overproduction and high level apoptosis are associated with high but not low virulence Toxoplasma gondii infection. J. Immunol. 167: 902-909. [Abstract/Free Full Text]
  49. Mordue, D. G., F. Monroy, M. La Regina, C. A. Dinarello, L. D. Sibley. 2001. Acute toxoplasmosis leads to lethal overproduction of Th1 cytokines. J. Immunol. 167: 4574-4584. [Abstract/Free Full Text]
  50. Aliberti, J., J. G. Valenzuela, V. B. Carruthers, S. Hieny, J. Andersen, H. Charest, C. Reis e Sousa, A. Fairlamb, J. M. Ribeiro, A. Sher. 2003. Molecular mimicry of a CCR5 binding-domain in the microbial activation of dendritic cells. Nat. Immunol. 4: 485-490. [Medline]
  51. Suzuki, Y., M. A. 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]
  52. Sukhumavasi, W., C. E. Egan, A. L. Warren, G. A. Taylor, B. A. Fox, D. J. Bzik, E. Y. Denkers. 2008. TLR adaptor MyD88 is essential for pathogen control during oral Toxoplasma gondii infection but not adaptive immunity induced by a vaccine strain of the parasite. J. Immunol. 181: 3464-3473. [Abstract/Free Full Text]
  53. Gately, M. K., L. M. Renzetti, J. Magram, A. S. Stern, L. Adorini, U. Gubler, D. H. Presky. 1998. The interleukin-12/interleukin-12-receptor system: role in normal and pathologic immune responses. Annu. Rev. Immunol. 16: 495-521. [Medline]
  54. Waldeland, H., J. K. Frenkel. 1983. Live and killed vaccines against toxoplasmosis in mice. J. Parasitol. 69: 60-65. [Medline]
  55. Gazzinelli, R., Y. 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]
  56. Nguyen, T. D., G. Bigaignon, D. Markine-Goriaynoff, H. Heremans, T. N. Nguyen, G. Warnier, M. Delmee, M. Warny, S. F. Wolf, C. Uyttenhove, et al 2003. Virulent Toxoplasma gondii strain RH promotes T-cell-independent overproduction of proinflammatory cytokines IL-12 and {gamma}-interferon. J. Med. Microbiol. 52: 869-876. [Abstract/Free Full Text]
  57. Andrade, R. M., M. Wessendarp, M. J. Gubbels, B. Striepen, C. S. Subauste. 2006. CD40 induces macrophage anti-Toxoplasma gondii activity by triggering autophagy-dependent fusion of pathogen-containing vacuoles and lysosomes. J. Clin. Invest. 116: 2366-2377. [Medline]
  58. Bliss, S. K., Y. Zhang, E. Y. Denkers. 1999. Murine neutrophil stimulation by Toxoplasma gondii antigen drives high level production of IFN-{gamma}-independent IL-12. J. Immunol. 163: 2081-2088. [Abstract/Free Full Text]
  59. Wagner, M., H. Poeck, B. Jahrsdoerfer, S. Rothenfusser, D. Prell, B. Bohle, E. Tuma, T. Giese, J. W. Ellwart, S. Endres, G. Hartmann. 2004. IL-12p70-dependent Th1 induction by human B cells requires combined activation with CD40 ligand and CpG DNA. J. Immunol. 172: 954-963. [Abstract/Free Full Text]
  60. Parent, M. A., K. N. Berggren, L. W. Kummer, L. B. Wilhelm, F. M. Szaba, I. K. Mullarky, S. T. Smiley. 2005. Cell-mediated protection against pulmonary Yersinia pestis infection. Infect. Immun. 73: 7304-7310. [Abstract/Free Full Text]
  61. Woelbing, F., S. L. Kostka, K. Moelle, Y. Belkaid, C. Sunderkoetter, S. Verbeek, A. Waisman, A. P. Nigg, J. Knop, M. C. Udey, E. von Stebut. 2006. Uptake of Leishmania major by dendritic cells is mediated by Fc{gamma} receptors and facilitates acquisition of protective immunity. J. Exp. Med. 203: 177-188. [Abstract/Free Full Text]
  62. Frenkel, J. K., D. W. Taylor. 1982. Toxoplasmosis in immunoglobulin M-suppressed mice. Infect. Immun. 38: 360-367. [Abstract/Free Full Text]
  63. Wilson, D. C., S. Matthews, G. S. Yap. 2008. IL-12 signaling drives CD8+ T cell IFN-{gamma} production and differentiation of KLRG1+ effector subpopulations during Toxoplasma gondii Infection. J. Immunol. 180: 5935-5945. [Abstract/Free Full Text]
  64. McLeod, R., P. Eisenhauer, D. Mack, C. Brown, G. Filice, G. Spitalny. 1989. Immune responses associated with early survival after peroral infection with Toxoplasma gondii. J. Immunol. 142: 3247-3255. [Abstract]
  65. Gazzinelli, R. T., S. Hieny, T. A. Wynn, S. Wolf, A. Sher. 1993. Interleukin 12 is required for the T-lymphocyte-independent induction of interferon {gamma} by an intracellular parasite and induces resistance in T-cell-deficient hosts. Proc. Natl. Acad. Sci. USA 90: 6115-6119. [Abstract/Free Full Text]
  66. Pepper, M., F. Dzierszinski, A. Crawford, C. A. Hunter, D. Roos. 2004. Development of a system to study CD4+-T-cell responses to transgenic ovalbumin-expressing Toxoplasma gondii during toxoplasmosis. Infect. Immun. 72: 7240-7246. [Abstract/Free Full Text]
  67. Trinchieri, G.. 2003. Interleukin-12 and the regulation of innate resistance and adaptive immunity. Nat. Rev. Immunol. 3: 133-146. [Medline]
  68. Cooper, A. M., S. A. Khader. 2007. IL-12p40: an inherently agonistic cytokine. Trends Immunol. 28: 33-38. [Medline]
  69. Gately, M. K., D. M. Carvajal, S. E. Connaughton, S. Gillessen, R. R. Warrier, K. D. Kolinsky, V. L. Wilkinson, C. M. Dwyer, G. F. Higgins, Jr, F. J. Podlaski, et al 1996. Interleukin-12 antagonist activity of mouse interleukin-12 p40 homodimer in vitro and in vivo. Ann. NY Acad. Sci. 795: 1-12. [Medline]
  70. Del Rio, L., B. A. Butcher, S. Bennouna, S. Hieny, A. Sher, E. Y. Denkers. 2004. Toxoplasma gondii triggers myeloid differentiation factor 88-dependent IL-12 and chemokine ligand 2 (monocyte chemoattractant protein 1) responses using distinct parasite molecules and host receptors. J. Immunol. 172: 6954-6960. [Abstract/Free Full Text]
  71. Pearce, E. L., H. Shen. 2007. Generation of CD8 T cell memory is regulated by IL-12. J. Immunol. 179: 2074-2081. [Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 Infect. Immun.Home page
J. P. Gigley, B. A. Fox, and D. J. Bzik
Long-Term Immunity to Lethal Acute or Chronic Type II Toxoplasma gondii Infection Is Effectively Induced in Genetically Susceptible C57BL/6 Mice by Immunization with an Attenuated Type I Vaccine Strain
Infect. Immun., December 1, 2009; 77(12): 5380 - 5388.
[Abstract] [Full Text] [PDF]

Home page
 Infect. Immun.Home page
K. A. Jordan, E. H. Wilson, E. D. Tait, B. A. Fox, D. S. Roos, D. J. Bzik, F. Dzierszinski, and C. A. Hunter
Kinetics and Phenotype of Vaccine-Induced CD8+ T-Cell Responses to Toxoplasma gondii
Infect. Immun., September 1, 2009; 77(9): 3894 - 3901.
[Abstract] [Full Text] [PDF]

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 HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Gigley, J. P.
Right arrow Articles by Bzik, D. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Gigley, J. P.
Right arrow Articles by Bzik, D. J.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS