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The Journal of Immunology, 2004, 173: 2632-2640.
Copyright © 2004 by The American Association of Immunologists

Functional Inactivation of Immature Dendritic Cells by the Intracellular Parasite Toxoplasma gondii1

Amy S. McKee2,3,*, Florence Dzierszinski2,{dagger}, Marianne Boes{ddagger}, David S. Roos4,{dagger} and Edward J. Pearce4,*

Departments of * Pathobiology and {dagger} Biology, University of Pennsylvania, Philadelphia, PA 19104; and {ddagger} Department of Dermatology, Brigham and Womens Hospital, Harvard Institutes of Medicine, Boston, MA 02115


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Despite its noted ability to induce strong cellular immunity, and its known susceptibility to IFN-{gamma}-dependent immune effector mechanisms, the protozoan Toxoplasma gondii is a highly successful parasite, able to replicate, disseminate, and either kill the host or, more commonly, establish resistant encysted life forms before the emergence of protective immune responses. We sought to understand how the parasite gains the advantage. Using transgenic clonal parasite lines engineered to express fluorescent markers in combination with dendritic cells (DC) grown from the bone marrow of wild-type mice or transgenic mice expressing fluorescent protein-tagged MHC class II molecules, we used flow cytometry and fluorescence microscopy to analyze the responses of infected DC to both invasion by the parasite and subsequent DC maturation signals. We found that T. gondii preferentially invades immature dendritic cells but fails to activate them in the process, and renders them resistant to subsequent activation by TLR ligands or the immune-system-intrinsic maturation signal CD40L. The functional consequences of T. gondii-mediated suppression of DC activation are manifested in a relative inability of infected immature DC to activate naive CD4+ Th lymphocytes, or to secrete cytokines, such IL-12 and TNF-{alpha}, that play important roles in innate and/or adaptive immunity. The findings reveal that T. gondii suppresses the ability of immature DC to participate in innate immunity and to induce adaptive immune responses. The ability of T. gondii to temporarily evade recognition could provide a selective advantage that permits dissemination and establishment before adaptive immune response initiation.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Toxoplasma gondii is a highly prevalent intracellular protozoan pathogen that infects a broad range of vertebrates, including humans (1). The tachyzoite stage of this parasite actively invades a wide variety of nucleated cell types, establishing an intracellular parasitophorous vacuole (PV),5 within which the parasite replicates until lysis of the host cell (2). The most virulent (type I) isolates of T. gondii (e.g., strain RH) can cause death of the host, despite the development of a seemingly strong immune response. Type I parasite strains are not inherently resistant to immune effector mechanisms, as a secondary immune response is capable of providing protection against RH challenge in previously vaccinated animals (3). Infection with type II T. gondii isolates (e.g., strains ME49 and Prugniaud) are usually controlled by IFN-{gamma}-dependent mechanisms after the first week (4, 5), although a percentage of parasites differentiate into latent bradyzoite forms (6) that persist for the life of the host, and can reemerge as acute infection in immunocompromised individuals (7).

The virulence of type I strains appears to be associated with the more rapid intracellular replication and consequent higher tissue burdens observed for these parasites (8). Thus, the outcome of infection depends on a balance between the timely development of an effective cellular immune response, and the ability of parasites to replicate and disseminate before immune recognition. Previous studies have suggested that live T. gondii parasites may subvert detection upon initial interaction with cells of the innate immune system. In resting macrophages, infection inhibits nuclear localization of the transcription factors STAT1 and NF-{kappa}B (9, 10, 11, 12), suppressing the up-regulation of surface expression of MHC class I and class II (MHC II) peptide Ag-presenting molecules in response to IFN-{gamma} (9), and limiting the production of IL-12 and TNF-{alpha} in response to the Gram-negative bacterial cell wall component LPS (10, 11).

Dendritic cells (DC) are related to macrophages, but are markedly more capable of initiating new immune responses (13, 14), and therefore, are thought to play the critical role in the immunological recognition of never before encountered pathogens (15). Immature DC of the CD8{alpha}– subtype are concentrated at portals of entry into the body, such as the skin and mucosal surfaces, while CD8{alpha}+ DC predominantly reside in secondary lymphoid organs (16, 17). DC express pattern recognition receptors such as those of the TLR family, that allow them to recognize pathogens or microbial products containing foreign molecular motifs; ligation of TLRs initiates DC maturation (18, 19). Maturation is associated with the cessation of endocytosis, production of inflammatory cytokines (including IL-12 and TNF-{alpha}), migration to lymph nodes, and increased surface expression of MHC II, CD40, CD80 and CD86, enabling mature DC to activate naive CD4+ Th lymphocytes (Th cells) (20, 21, 22). The production of IL-12 is particularly important in toxoplasmosis, because this cytokine stimulates NK cells to make IFN-{gamma}, and promotes the development of IFN-{gamma}-producing Th1 cells (23, 24, 25).

Although splenic CD8{alpha}+ DC have been shown to respond to soluble T. gondii Ags (STAg) by producing IL-12 (26, 27, 28, 29), CD8{alpha}– DC were reported to be comparatively nonresponsive to STAg. Thus, it is not clear whether immature CD8{alpha}– DC, which would be expected to be the DC type to first encounter T. gondii following infection, are primed to become competent Ag-presenting cells following encounter with this parasite. The aim of this study was to address this issue. We show that T. gondii preferentially invades immature CD8{alpha}– DC, failing to activate them in the process, and rendering them insensitive to subsequent activation signals. Through suppression of dendritic cell maturation, T. gondii may gain a temporary advantage over the immune system, favoring its success as a parasite.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Hosts and parasites

C57BL/6 (B6) and BALB/c male and female mice and OTII male mice were purchased from The Jackson Laboratory (Bar Harbor, ME), and/or bred at the University of Pennsylvania (Philadelphia, PA). Transgenic mice in which the MHC II {beta}-chain gene was replaced with a version that codes for a class II molecule tagged with enhanced GFP (EGFP) (30), were maintained at Harvard Medical School (Boston, MA). Tachyzoites derived from the type II Prugniaud strain ({Delta}HXGPRT knockout parasites kindly provided by D. Soldati, Imperial College, London, U.K.), from the cps1-1 RH strain ({Delta}CPSII knockout parasites kindly provided by D. J. Bzik, Dartmouth Medical School, Lebanon, NH) (31), or from the virulent RH strain of T. gondii (transgenic lines engineered as described below) were maintained by serial passage in human foreskin fibroblast cell monolayers in DMEM containing 10% FBS (Invitrogen Life Technologies, Carlsbad, CA), as previously described (32).

Molecular methods

Transgenic parasite clonal lines were engineered to express the fluorescent markers yellow fluorescent protein (YFP), GFP, or red fluorescent protein (RFP) (DsRed; BD Clontech, Palo Alto, CA) in the cytosol, or to target reporters to dense granules for secretion into the PV. Vectors were based on Bluescript pKS+ (Stratagene, La Jolla, CA) and engineered as follows: 1) the T. gondii TUB1 promoter (33) (BglII site upstream of the initiation codon); 2) dense granule targeting sequences as described below in frame (AvrII site) with a fluorescent reporter or fluorescent reporter alone for cytosolic expression; 3) a 3' untranslated region from T. gondii dihydrofolate reductase-thymidylate synthase (DHFR-TS) gene terminating in a NotI site (34); and 4) a chloramphenicol acetyltransferase (CAT)-selectable marker expressed under the control of 5' and 3' untranslated regions from the T. gondii SAG1 gene (35). Plasmid ptubP30-GFP/sagCAT has been previously described (36, 37). Plasmid ptubGRA8-YFP/sagCAT was made using ptubFNR-YFP/sagCAT (kindly provided by F. Seeber, Philipps-Universität, Marburg, Germany), which contains the first 150 aa of the T. gondii ferredoxin-NADP+ reductase (38). Briefly, the FNR leader peptide was replaced by the GRA8 signal peptide (aa 1–24) obtained by PCR amplification from tachyzoite cDNA (primers 5'BglII-catagatctATGgctttaccattgcgtgtttcgg-3' and 5'NheI-ctagctagcgcgagctacaccaaagacagc-3') (M. J. Crawford, unpublished observations). Plasmids ptubRFP/sagCAT and ptubYFP/sagCAT were obtained by subcloning an RFP-3'dhfr or a YFP-3'dhfr fragment in place of the P30GFP-3'dhfr fragment in ptubP30-GFP/sagCAT via BglII/NotI restriction digests. RFP or YFP were originally subcloned via BglII/AflII restriction digests in ptub[BglII]GFP[AflII]3'dhfr[NotI]/sagCAT after PCR-amplification from pDsRed1-1 or pEYFP (BD Clontech). To improve expression levels of fluorescent reporters in T. gondii, second codons were modified to encode an alanine residue (39). Transfections were performed by electroporation as previously described (32), using 2 x 107 tachyzoites and 70 µg of NotI-linearized plasmid in a 2-mm gap cuvette (1.5 kEV pulse, 24 {Omega}; BTX, Holliston, MA). Stable transgenics were selected in the presence of 20 µM chloramphenicol, and parasite clones were then isolated by limiting dilution.

Abs and reagents

DC activation was analyzed by flow cytometry after staining with allophycocyanin- or PE-conjugated anti-CD11c mAb, and PE- or FITC-labeled anti-CD80, anti-CD40, and anti-CD86 mAb. When four-color analysis was required, biotinylated anti-MHC II or anti-CD40 mAb and PerCP-labeled streptavidin were used. Splenic DC were stained with allophycocyanin-labeled anti-CD11c, PerCP-labeled anti-CD8{alpha}, and FITC-labeled anti-CD80, anti-CD86, or anti-CD40 mAb (all mAb and PerCP-labeled streptavidin for flow cytometry; BD Pharmingen, San Diego, CA). Paired mAb in combination with recombinant cytokine standards were used to measure IL-12p40 (BD Pharmingen) and TNF-{alpha} (R&D Systems, Minneapolis, MN) in culture supernatants. Soluble CD40L (a gift from Immunex, Seattle, WA) and LPS (Sigma-Aldrich, St. Louis, MO) were each used at concentrations of 1 µg/ml. Soluble parasite extracts, used at a concentration of 50 µg/ml, were prepared from sonicated RH strain tachyzoites as previously described (40). Propionibacterium acnes ATCC:12930 (American Type Culture Collection, Manassas, VA) was heat-killed (HK) and used at a concentration of 10 µg/ml.

DC cultures and isolation of splenic DC

DC were grown from mouse bone marrow using GM-CSF as described in detail previously (41). Using this protocol, we obtained cell populations that were >90% CD11c+. As expected, no contaminating B cells, macrophages, CD4 or CD8 T cells, or CD8+ DC were generated under these conditions, as determined by FACS using mAbs specific for B220, F4/80, CD4, and CD8 (Ref. 42 , and data not shown); non-DC were primarily neutrophils, as assessed by staining with anti-GR1 (Ref. 42 , and data not shown). DC were harvested on day 10, counted by trypan blue exclusion, resuspended to a concentration of 2 x 106/ml in DC medium (5 x 105/ml; RPMI containing 100 U/ml penicillin, 100 µg/ml streptomycin (Mediatech, Washington, DC), 0.5 µM 2-ME (Sigma-Aldrich), 10% FCS (HyClone, Logan, UT), and 2 mM L-glutamine (Mediatech)) with 5 ng/ml GM-CSF (PeproTech, Rocky Hill, NJ). CD11c+ splenic DC were isolated from single-cell suspensions of splenocytes using CD11c microbeads and magnetic sorting (Miltenyi Biotec, Auburn, CA), and resuspended to a concentration of 5 x 105/ml in DC medium. DC were plated in 24- or 48-well plates before incubation with parasites or extracts.

In vitro DC activation experiments and flow cytometry

DC were exposed for 18 h to live or HK (56°C for 20 min) tachyzoites, or to parasite extract. In some experiments, infected DC were stimulated with CD40L or LPS for 3 or 6 h, or copulsed with LPS for 18 h. At this time, supernatants were collected and stored at –20°C for subsequent cytokine measurements, and DC were collected and stained for flow cytometric analysis; DC were washed in flow wash (Dulbecco’s PBS containing 1% FCS and 0.05% sodium azide) and incubated with Fc Block (BD Pharmingen) for 15 min on ice, followed by incubation with allophycocyanin-conjugated anti-CD11c mAb and PE- or FITC-conjugated anti-MHC II IAb, -MHC II IAd, -CD80, -CD86, or -CD40 mAbs for 30 min on ice. For IL-12 intracellular staining, DC were incubated for 15 min in 3% paraformaldehyde (Sigma-Aldrich) in PBS, permeabilized in saponin buffer (Dulbecco’s PBS containing 0.075% saponin (Sigma-Aldrich), 5% normal mouse serum (Cedarlane Laboratories, Hornby, Ontario, Canada)) for 15 min on ice, and stained with either a PE-labeled anti-IL-12p40 or PE-labeled IgG isotype control mAb for 15 min at room temperature. Cells were then washed once in saponin buffer and twice in flow wash. Data were acquired on a FACSCalibur Flow Cytometer using CellQuest software (BD Biosciences, San Jose, CA) and analyzed using FlowJo software (Tree Star, Ashland, OR). In some experiments, infected DC isolated by FACS using a FACSVantage SE flow cytometer (BD Biosciences).

Microscopy and imaging

DC (2 x 106/ml) were cultured on sterile glass coverslips in 24-well plates with or without live or HK tachyzoites, or soluble extracts, for 18 h. Cells were fixed in 4% paraformaldehyde, permeabilized in saponin buffer, and stained with biotin-conjugated anti-MHC II IAb mAb and AlexaFluor594-conjugated streptavidin (Molecular Probes, Eugene, OR) for 30 min. Reagents were diluted in 10% FBS, 0.1% Triton X-100 in PBS. For time-lapse video microscopy, 4 x 106 DC-expressing EGFP-tagged MHC II molecules (EGFP-MHC II) (30) were transferred onto poly-L-lysine-treated, gridded, coverslip-bottom, 35-mm dish (MatTek, Ashland, MA) before parasite infection (106 tachyzoites), and imaging was performed at 37°C using a heated microscope stage and lens objective heater (Bioptechs, Butler, PA). YFP, GFP, RFP, and secondary reagent fluorescence was detected using a Zeiss Axiovert 35 microscope equipped with a 100 W Hg-vapor lamp (Thornwood, NY), appropriate barrier/emission filters, and an interline transfer chip CCD camera (Hamamatsu, Hamamatsu City, Japan). Images were captured, and color and contrast adjusted using Openlab software (Improvision, Lexington, MA).

Ag presentation assay

DC (2 x 106/ml) were cultured in 24-well plates in the presence or absence of cps1-1 RH tachyzoites (multiplicity of infection, 0.5) for 18 h, after which infected cells were FACS-sorted, stimulated with 1 µg/ml LPS for 3 h, counted, and resuspended to a concentration of 5 x 104/ml. A total of 100 µl of the suspension was added per well to 96-well round-bottom plates in the presence of 100 ng/ml OTII peptide (ISZAVHAAHAEINEAGR; Invitrogen Life Technologies). CD4+ T cells were isolated from a single-cell suspension of OTII splenocytes (CD4 T cell isolation kit; Miltenyi Biotec), and resuspended at 5 x 105/ml; 100 µl aliquots of this suspension were added to peptide- or medium-pulsed control, or infected DC as described above. After 24 h, CD4+ T cells were stained with a PE-labeled anti-CD25 mAb and a PerCP-conjugated anti-CD4 mAb, and analyzed using FACSCalibur as described above.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Immature DC are activated by exposure to dead parasites or parasite extracts, but not as a result of invasion by living parasites

When examined by flow cytometry, bone marrow-derived DC consisted of CD11c+CD8{alpha}– DC, definable as immature or mature based on their relative surface expression (lower on immature cells, higher on mature cells) of MHC II, CD80, CD86, and CD40 (Fig. 1A, and data not shown). To assess how DC respond to T. gondii infection, we inoculated DC cultures with live tachyzoites, a parasite soluble extract (STAg), or HK parasites, and used flow cytometry to compare surface expression levels of MHC II, CD80, CD86, and CD40 with those on DC cultured in the absence of any stimuli (medium controls), or with HK P. acnes, which we have reported previously to be a strong maturation stimulus for DC (42). Although parasite extract and, especially, HK parasites induced notable DC maturation (albeit not as marked as that stimulated by HK P. acnes), live tachyzoites had remarkably little effect on inducing activation as measured by up-regulated surface expression of MHC II, CD80, or CD40 (Fig. 1A). We did note a significant increase in the population of DC expressing high levels of CD86 following infection (Fig. 1A). Consistent with the absence of changes in surface expression of MHC II, CD80, and CD40, IL-12p40 secretion was not enhanced by T. gondii infection (Fig. 1B). Consistent with previous findings (26), T. gondii extract also failed to induce IL-12 production by CD8{alpha}– DC, but HK parasites provided a strong stimulus (Fig. 1B). HK P. acnes induced the production of ~10-fold more IL-12 than was provoked by HK tachyzoites (data not shown). We were unable to detect production of IL-6, IL-10, or TNF-{alpha} following exposure of DC to live or HK tachyzoites, or to T. gondii extracts (data not shown).



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  		FIGURE 1. Immature DC are not activated by T. gondii infection. DC from B6 mice were pulsed for 18 h with culture medium (negative control), live RH strain T. gondii tachyzoites, a soluble tachyzoite extract (T. gondii extract, also known as STAg), HK RH strain parasites (HK T. gondii), or HK strain ATCC:12930 P. acnes bacteria (positive control). A, DC activation was assessed by examining surface expression of MHC II, CD80, CD86, and CD40. Gates were drawn to delineate mature DC based on expression in P. acnes-pulsed positive controls, and numbers indicate percentage of DC within the gates. Scales are log. B, IL-12 levels in culture supernatants were assayed by ELISA (triplicate assays; error bars = SD). Similar results were observed for DC from BALB/c (data not shown). Data shown are from one experiment, and are representative of results from three independent experiments.

 
Tachyzoites establish productive infection in DC and preferentially infect immature cells

To specifically assess the phenotype of infected vs uninfected DC, we engineered both transgenic RH strain (virulent) and Prugniaud strain (cyst-forming) parasites expressing fluorescent reporter proteins in dense granules, allowing straightforward detection of infected cells by fluorescence microscopy and flow cytometry (Fig. 2). These reporters (YPF and GFP) are secreted into the PV space upon active host cell invasion, but not when parasites are phagocytosed (36, 37). A small number of phagocytosed HK parasites remained detectable by microscopy due to residual reporter protein, but this low level of fluorescence was not discernable by flow cytometry (Fig. 2C, bottom panel).



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  		FIGURE 2. Tachyzoites establish productive DC infection. B6 DC were incubated 18 h with either live or HK T. gondii tachyzoites expressing GFP or YFP targeted to dense granules and secreted into the PV. Cultures were then examined by fluorescence microscopy (A and B, Prugniaud strain, Pru GRA8-YFP) or flow cytometry (C, RH strain, RH P30-GFP; log scales). Numbers indicate percentage of GFP-positive DC. Similar results were obtained when RH P30-GFP parasites were observed by microscopy, and Pru GRA8-YFP were analyzed by flow cytometry (data not shown). Live parasites entered CD11c-positive DC and established active infection, as indicated by the large number of intracellular parasites and secretion of the YFP marker. Microscopic images show phase, fluorescence, and overlay for three representative cells. Data shown are from one experiment, and are representative of results from three independent experiments.

 
Use of these transgenic parasites allowed us to specifically examine changes in maturation markers at the level of individual infected cells. Analysis of infected vs uninfected cells after addition of live T. gondii to DC cultures revealed that a disproportionately high percentage of infected DC exhibited an immature phenotype with low levels of surface MHC II, CD80, CD86, and CD40 expression (Fig. 3), and that there was an ~2- to 3-fold higher ratio of immature to mature DC in the infected compared with uninfected subpopulation (Fig. 3B). Despite inoculation with ~1 infectious tachyzoites per DC, most mature DC remained uninfected, while immature cells contained multiple PVs, reflecting multiple invasion events (Fig. 3A). Interestingly, the total numbers of immature DC, characterized by relatively low expression of surface MHC II, in cultures infected with living tachyzoites were higher than in cultures that did not contain parasites (Fig. 3B). For example, 67% of medium controls were MHC IIhigh, vs 34% of infected cultures. Moreover, a lower percentage of MHC IIhigh DC were infected (in the example shown, 41% of MHC IIhigh cells were infected, while 68% of MHC IIlow cells were infected). Similar results were found when expression of CD40, CD80, and CD86 were analyzed in infected vs uninfected DC (Fig. 3B). These data suggest either deactivation of infected mature cells, or preferential invasion of immature DC, followed by suppression of spontaneous maturation during the 18-h culture period.



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  		FIGURE 3. T. gondii-infected DC lack maturation markers. DC from B6 mice were incubated with live Prugniaud strain (Pru) parasites expressing GRA8-YFP (A) or RH strain parasites expressing P30-GFP (A and B), and stained for MHC II in permeabilized cells (A), or MHC II, CD40, CD80, and CD86 on intact cells (B). Contrast was intentionally boosted in the anti-MHC II ({alpha}MHC II) panels to visualize low levels of expression in immature DC (A). Small numbers in B indicate percentages in the corresponding quadrants, drawn to distinguish infected (right), uninfected (left), immature (lower), and mature (upper) DC. Bold numbers below each plot indicate the percentage of uninfected DC (left) or infected DC (right) that are immature, and were calculated by dividing percentage of immature uninfected or immature infected DC by total uninfected or infected DC and multiplying by 100. Italicized numbers indicate the percentage of mature (upper) and immature (lower) DC infected by T. gondii and were calculated by dividing the percentage of infected mature or infected immature DC by total mature or immature DC and multiplying by 100. Similar results were observed using DC from BALB/c mice (data not shown). Data shown are from one experiment, and are representative of results from three independent experiments.

 
To examine these possibilities, we infected live bone marrow-derived DC from mice expressing EGFP-MHC II (30) with transgenic RH strain parasites expressing cytosolic RFP, and followed these cultures for 27 h by time-lapse video microscopy. These experiments clearly show that infected immature DC remained in an immature state, indicated by low surface expression of MHC II over the course of 27 h (Fig. 4, white arrows). Infected DC continued to express low levels of EGFP-MHC II in intracellular compartments (e.g., Fig. 4A, and data not shown). Although some uninfected DC were seen to undergo spontaneous maturation (data not shown), this was never observed for infected immature DC.



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  		FIGURE 4. Time-lapse video microscopy of EGFP-MHC II transgenic DC infected with fluorescently labeled parasites. B6 DC from EGFP-MHC II transgenic mice (green) were inoculated with ptub-RFP transgenic RH tachyzoites (red) and tracked over the course of infection. MHC IIlow cells infected with tachyzoites (white arrows) failed to mature during the course of infection (note parasite replication). MHC IIhigh cells were only infrequently infected, but continued to mature (yellow arrowheads in B; note high surface expression at 27.5 h (A); note parasite replication). Data shown are from one experiment, and are representative of results from two independent experiments.

 
Mature DC (expressing high levels of MHC II) were occasionally infected (Fig. 4B, yellow arrowhead), and these cells progressively developed a more highly dendritic morphology and slightly higher levels of MHC II surface expression, indicating that MHC II expression was not down-regulated over time as a result ofinfection. We observed that parasites remained viable and able to replicate within either immature or mature DC.

Mature splenic and bone marrow-derived DC remain activated following invasion

During the time-lapse analysis, we became aware that mature DC would occasionally become infected, and that these cells retained their mature status. To examine this issue more carefully, we focused on a comparison of the effects of infection on bone marrow-derived DC that had been matured by exposure to LPS before infection, or on isolated splenic DC, which mature spontaneously once placed in culture (43). As expected, activation of bone marrow-derived DC with LPS for 6 h before infection led to increased surface expression of MHC II (Fig. 5A). When pulsed with live tachyzoites, these mature DC became infected, and continued to express high levels of MHC II (Fig. 5A), indicating that their maturation status was not reversed by infection. Infected splenic DC (including both CD8{alpha}– and CD8{alpha}+ subsets) also retained their mature phenotype upon infection (Fig. 5B, and microscopy studies, data not shown). Thus, in situations where the numbers of immature DC are limiting, T. gondii will infect mature DC, which subsequently continue to exhibit a mature phenotype. An additional finding to emerge from these experiments is that parasite replication is less extensive in DC that were activated before infection. This is evident as decreased intensity of parasite-associated fluorescence in infected LPS-activated DC compared with infected unactivated-DC (Fig. 5A). Possibly this reflects the production of NO by LPS-activated DC (42).



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  		FIGURE 5. Mature DC remain mature following infection with T. gondii. A, Bone marrow-derived DC were activated for 6 h with LPS, infected with RH ptub-YFP for 18 h, and stained for surface MHC II. Numbers indicate percentages of total DC within each quadrant. B, Splenic DC isolated from naive B6 mice were infected with RH P30-YFP for 18 h, washed, and stained for surface MHC II, CD11c, and CD8{alpha}. Quadrants show MHC II expression and parasite fluorescence for gated CD11c+CD8{alpha}– DC (upper panels) or gated CD11c+CD8{alpha}+ DC (lower panels). Numbers indicate percentage of gated DC in each quadrant. This experiment was performed twice with similar results; data from one experiment are shown.

 
Invaded immature DC are nonresponsive to maturation signals

The data presented above suggested that the spontaneous maturation observed in some bone marrow DC during culture is prevented when these cells are infected by T. gondii. To test whether infected immature DC are indeed refractory to maturation signals, we assayed the ability of infected cells to make IL-12p40 and TNF-{alpha}, and to up-regulate MHC II expression upon treatment with LPS. As shown in Fig. 6A, increasing the parasite inoculum reduced the production of IL-12 and TNF-{alpha} by DC concomitantly stimulated with LPS. Up-regulation of MHC II surface expression by LPS was also inhibited in infected immature DC. As shown in Fig. 6B, the majority (89%) of uninfected MHC IIlow DC were induced to express surface MHC II by LPS treatment. In contrast, very few (6%) infected DC up-regulated MHC II expression following treatment with LPS (Fig. 6B). These results suggest the active inhibition of DC maturation by the parasites.



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  		FIGURE 6. Infected DC do not respond to LPS. DC from B6 mice were inoculated with P30-GFP transgenic RH tachyzoites at increasing multiplicity of infection (A) or at a ratio of 1 Tz:DC (B), and simultaneously stimulated plus (A, {blacksquare}) or minus (A, {diamond}) 1 µg/ml LPS. Levels of IL-12p40, TNF-{alpha}, and surface MHC II were assayed 18 h postinfection. Infection with live parasites markedly inhibited LPS-induced cytokine and surface MHC II expression (shown on a log scale). Data points in A indicate levels detected in triplicate wells ± SD. Although uninfected MHC IIlow DC were induced to differentiate in response to LPS (compare upper and lower left quadrants, ± LPS), infected DC failed to respond to LPS treatment (compare upper and lower right quadrants, ± LPS). Data shown are from one experiment, and are representative of results from three independent experiments.

 
Previous reports indicated that DC exposed to T. gondii extracts and tachyzoites are primed to subsequently make IL-12 in response to the immune-system-intrinsic maturation signal provided by CD40 ligation (44, 45). In light of this finding, we compared the ability of DC exposed to live or HK T. gondii, parasite extract, or HK P. acnes to produce IL-12 and elevate MHC II surface expression in response to CD40L. Ligation of CD40 with soluble CD40L led to a >3-fold increase in the percentage of DC (all MHC IIhigh) making IL-12, while having no detectable effect on MHC II expression levels per se (Fig. 7). Infection of DC with live T. gondii profoundly inhibited the ability of DC to produce IL-12 in response to CD40L (Fig. 7). In contrast to the suppression of DC responsiveness to CD40L in T. gondii-infected cells, DC maintained responsiveness to this stimulus following treatment with parasite extract, and were activated directly by HK parasites or HK P. acnes; in the latter two cases, CD40 ligation caused small but measurable additional increases in the percentages of DC making IL-12 (Fig. 7). In summary, infection of DC by T. gondii leads to a significant defect in the ability of these cells to respond to microbial and to immune-system-intrinsic maturation signals.



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  		FIGURE 7. Infected DC do not respond to CD40L. B6 DC were inoculated with control medium, live or HK P30-GFP transgenic RH tachyzoites, soluble parasite Ag, or HK P. acnes, and incubated 18 h, followed by 6 h exposure to CD40L (1 µg/ml), before assaying MHC II and IL-12 expression. Numbers indicate percentages of DC that are MHC IIhigh and MHC IIlow, and positive and negative for IL-12 production. In contrast to other treatments with T. gondii or P. acnes Ags (large nonbold numbers), CD40L failed to up-regulate IL-12 expression in MHC IIhigh T. gondii-infected DC (circled large number); see text for further discussion. Scales are log. Data shown are from one experiment, and are representative of results from two independent experiments.

 
Inhibition of DC maturation in infected cells does not require parasite replication

To address whether the apparent effects of infection on DC maturation might be attributable to infection-induced DC morbidity, we used the uracil auxotroph mutant RH strain cps1-1 (31). Parasites possessing this null mutation in the regulatory enzyme required for de novo pyrimidine biosynthesis (carbamoyl phosphate synthase II) invade nucleated mammalian cells normally, but require uracil supplementation to replicate within the host cell (31). cps1-1 knockout mutants (31) engineered to express cytosolic YFP were inoculated into DC in uracil-free medium, and the DC were examined for activation following exposure to LPS. cps1-1-infected DC, like DC infected with wild-type parasites, were incapable of responding normally to LPS-stimulation, as a lower percentage of cells up-regulated MHC II, CD80, and CD40 (compare expression in uninfected vs infected cells in Fig. 8). Thus, infection with a single parasite can have a rapid, profound effect on the ability of immature DC to undergo the changes considered essential for them to be able to activate T cells.



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  		FIGURE 8. Parasite replication is not required for repression of DC activation. DC from B6 mice were infected for 18 h with ptub-YFP transgenic cps1-1 parasites in the absence of uracil (to inhibit intracellular replication of these mutants), followed by 6 h treatment ± 1 µg/ml LPS. Quadrants delineate infected and uninfected DC expressing high or low levels of MHC II, CD80, CD86, or CD40, and small numbers indicate percentage of DC within each quadrant; scales are log. Bold numbers below each pair of plots indicate the percentage of immature uninfected (left) or infected (right) DC that have up-regulated expression of each activation marker in response to LPS. LPS stimulated expression of all four markers in uninfected DC, but this up-regulation was inhibited by T. gondii infection. Data shown are from one experiment, and are representative of results from three independent experiments.

 
Infection impairs the ability of DC to activate Th cells

Maturation prepares DC for interactions with Th cells, increasing surface expression of a panel of molecules, including MHC II CD80, and CD86, that allow DC to deliver primary and secondary activation signals to Th cells. In addition, increased expression of CD40 on mature DC primes these cells to receive reciprocal activation signals from CD40L expressed on Th cells. The fact that T. gondii preferentially infects immature DC, and blocks their ability to mature, suggests that infected DC are unlikely to function as competent activators of naive T cells. To test this hypothesis, uninfected DC or DC infected with cps1-1 mutant T. gondii were cultivated in the presence or absence of LPS for 6 h, followed by incubation with or without OVA peptide and purified OTII Th cells. As shown in Fig. 9, in the absence of OVA peptide, DC failed to induce Th cell activation, as assessed by the expression of the early response marker CD25. Pulsing uninfected DC with OVA peptide allowed them to activate 16% of cocultured CD4 cells, a process that was considerably enhanced (from 16 to 36%) by prior treatment with LPS (Fig. 9); this result is consistent with the pronounced DC maturation induced by LPS (Fig. 6). In contrast, infected DC were impaired in the ability to activate Th cells. Only 7% of OTII cells cocultured with OVA peptide-pulsed infected DC up-regulated expression of CD25 (Fig. 9). Moreover, infected DC stimulated with LPS and pulsed with peptide, though more capable of activating Th cells than infected peptide-pulsed cells that had not been exposed to LPS, were nevertheless measurably less capable of activating OTII cells than were immature uninfected DC (Fig. 9). Thus, when immature DC are infected with T. gondii, they are unable to mature into functional Th-activating cells, even following stimulation with LPS.



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  		FIGURE 9. Infected DC are deficient in their ability to activate naive T cells. B6 bone marrow-derived DC were infected with cps1-1 ptub-YFP mutant and incubated 18 h without uracil (preventing parasite replication and DC lysis). Cultures were incubated 6 h ± 1 µg/ml LPS, sorted into infected and uninfected populations, resuspended for 1 h in fresh medium ± 100 ng/ml OTII peptide, and mixed with OTII CD4+ cells at a ratio of 1 DC:10 CD4+ T cells. Activation of CD4+ cells was assessed by staining for CD25 after 24 h; numbers indicate percentage of CD25+, based on comparison with isotype control staining. Data shown are from one experiment, and are representative of results from two independent experiments.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
T. gondii is a ubiquitous human and animal pathogen, infecting a wide range of tissues in virtually any vertebrate. During the course of a natural infection, the parasite is likely to encounter, and infect, immature DC at peripheral mucosal sites. Previous studies have shown that toxoplasmosis leads to the activation of splenic DC (46), as can the injection of T. gondii extracts (26). However, the effects on DC of their direct exposure to living Toxoplasma have remained unclear. In the studies described here, we found that T. gondii was highly effective at invading immature DC, preferring immature rather than mature cells when given the choice (Figs. 2–4). In contrast to the stimulation provoked by HK parasites or parasite extracts, live parasites failed to up-regulate surface expression of MHC II, CD80, or CD40, or to produce detectable levels of IL-12 or other inflammatory cytokines (Figs. 1 and 3), even in response to stimulation with TLR or CD40 ligands (Figs. 6–8). These cells were also inefficient at activating CD4+ T cells (Fig. 9), indicating that DC remained functionally as well as phenotypically immature. These findings reveal that T. gondii invasion of immature DC suppresses the ability of these cells to participate in innate immunity and to induce adaptive immune responses.

The basis for preferential invasion of immature DC is currently unclear. DC morphology and behavior change dramatically during differentiation, resulting in the extension of dendrites and increased motility (20), but both of these factors might have been expected to enhance, rather than inhibit, parasite binding. T. gondii uses multiple adhesins and surface Ags to promote attachment and entry (47, 48, 49), and it has been proposed that tachyzoites attach to common carbohydrate modifications on host cell membrane molecules, allowing them to invade virtually any cell type (2). It is possible that maturation-associated changes in glycosylation inhibit the ability of tachyzoites to invade DC. Alternatively, differences in invasion may be attributable to maturation-associated changes in the expression of DC surface proteins that are important for the recognition of host cells by the parasite (47, 48, 49).

Active invasion by the parasite (but not intracellular replication; Fig. 8) is required for suppression of DC maturation. The failure of infection to stimulate DC maturation might be explained, in part, by the exclusion of transmembrane signal transducing receptors from the PV membrane upon parasite entry (50). In contrast, incubation with HK parasites or soluble parasite extracts resulted in the up-regulation of MHC II and other surface markers, presumably via signaling through TLR and/or CCR5 (28), which are known to be expressed in immature CD8{alpha}– DC (51).

How does infection suppress the ability of immature DC to respond to strong microbial and immune-system-intrinsic activation signals? In macrophages, T. gondii infection is known to interfere with STAT1 and NF-{kappa}B signaling pathways, although the mechanistic basis underlying these effects has not been defined (9, 10, 11). It seems unlikely that inhibition of STAT1 signaling could fully explain the effect of T. gondii infection on DC, because factors not known to use the STAT1 pathway (LPS, CD40L, TNF-{alpha}, etc.) are still able to induce transcription of MHC II, CD80, CD86, and CD40 (52, 53, 54, 55, 56), and MHC II surface expression (21, 22, 57). Inhibition of NF-{kappa}B signaling provides a more attractive mechanism through which DC maturation and the production of IL-12 and TNF-{alpha} could be prevented in immature DC, as these processes do not occur when nuclear translocation of NF-{kappa}B is inhibited by I-{kappa}B overexpression (58). Interestingly, while those relatively few mature DC that became infected with T. gondii remained activated by the criteria of MHC II surface expression, they were unable to produce IL-12 in response to CD40L. Thus, T. gondii appears to be able to suppress at least two distinct steps in DC maturation, preventing increased MHC II expression, and IL-12 production in MHC IIhigh DC in response to additional stimuli (55, 59).

IL-10 is a well-recognized inhibitor of DC maturation (60, 61), but this cytokine is unlikely to underlie the suppressive action of tachyzoite invasion on DC maturation, as IL-10 was not detected in culture supernatants (data not shown), and uninfected DC in the same cultures were able to mature, suggesting that a soluble factor is not responsible for the inhibitory activity. Similarly, IL-10 was not implicated in the macrophage studies noted above (10, 12, 62).

During an active T. gondii infection, parasites are likely to encounter, and infect, immature DC in peripheral tissues. Infection-induced inhibition of the ability of these cells to make cytokines, such as IL-12, would be expected to have a negative impact on the production of IFN-{gamma} by NK cells, and therefore, on the expression of IFN-{gamma}-dependent innate microbicidal mechanisms. Moreover, by infecting immature DC, parasites may be able to disseminate from the site of infection within cells undergoing steady-state migration to draining lymphoid organs (63, 64). Our data, and the findings of a previous study (65), indicate that infected DC are compromised in their ability to activate T cells, suggesting that the arrival of infected DC in lymphoid tissues will not result in efficient priming of a T cell response. Together, these processes would be expected to enhance the likelihood that parasites will reach sites such as the brain and skeletal muscles, where tachyzoites undergo differentiation into the encysted bradyzoite forms that establish chronic infection, ensuring parasite transmission. In light of these findings, it is possible that the initiation of innate and adaptive immune responses during toxoplasmosis is accomplished by DC activated by dead parasites or parasite products (26). Additionally, neutrophils have been reported to be recruited to the site of infection and produce cytokines, such TNF-{alpha}, which can initiate DC maturation (66), and thus, may play an important role in immune response development (67).

Inhibition of DC maturation has previously been reported during infection by the malaria parasites Plasmodium falciparum and Plasmodium yoelii, where the ability of DC to activate T cells is greatly inhibited by their interaction with adhesive P. falciparum-infected RBC (68), and P. yoelii blood-stage infection induces the secretion of soluble factors by DC, leading to the suppression of CD8+ T cell responses against the liver stage of the parasite (69). Although the mechanisms by which the related pathogens T. gondii and Plasmodium sp. suppress DC maturation undoubtedly differ in detail, the fact that two highly successful genera of the phylum Apicomplexa are able to decommission DC from participating in immune responses indicates that this may be an important strategy for successful parasitism by this group of organisms. The experimental accessibility of T. gondii raises the possibility of addressing this issue by using powerful genetic approaches to identify parasite molecules that are responsible for suppressing DC maturation.


   Acknowledgments
 
We thank Hidde Ploegh for support and helpful discussions, and Amy Crawford, EuiHye Jung, and Tamika Seals for expert technical assistance.


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

1 This work was supported in part by National Institutes of Health Grants RO1-AI53825 (to E.J.P.) and R37-AI28724 (to D.S.R.). D.S.R. and E.J.P. are Burroughs Wellcome Scholars in Molecular Parasitology. D.S.R. is an Ellison Foundation Scholar in Global Infections Diseases. Back

2 A.S.M. and F.D. contributed equally to this work. Back

3 Current address: Department of Immunology, National Jewish Medical and Research Center, 1400 Jackson Street, Denver, CO 80296. Back

4 Address correspondence and reprint requests to Dr. Edward J. Pearce, University of Pennsylvania, Room 203D Johnson Pavilion, Hamilton Walk, Philadelphia, PA 19104-6076. E-mail address: ejpearce{at}mail.med.upenn.edu; or Dr. David S. Roos, University of Pennsylvania, Room 305 Goddard Laboratories, Hamilton Walk, Philadelphia, PA 19104-6018. E-mail address: droos{at}sas.upenn.edu Back

5 Abbreviations used in this paper: PV, parasitophorous vacuole; B6, C57BL/6; CAT, chloramphenicol acetyltransferase; DC, dendritic cell; EGFP, enhanced GFP; HK, heat killed; MHC II, MHC class II; RFP, red fluorescent protein; STAg, soluble T. gondii Ag; YFP, yellow fluorescent protein; EGFP-MHC II, EGFP-tagged MHC II molecule. Back

Received for publication April 7, 2004. Accepted for publication June 4, 2004.


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