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. Author manuscript; available in PMC: 2010 Jan 1.
Published in final edited form as: J Immunol. 2009 Jan 1;182(1):489–497. doi: 10.4049/jimmunol.182.1.489

Toxoplasma gondii Prevents Chromatin Remodeling Initiated by TLR-Triggered Macrophage Activation1

Jin Leng 1, Barbara A Butcher 1, Charlotte E Egan 1, Delbert S Abi Abdallah 1, Eric Y Denkers 1,2
PMCID: PMC2651083  NIHMSID: NIHMS86180  PMID: 19109180

Abstract

Macrophages infected with the opportunistic protozoan Toxoplasma gondii are unable to up-regulate many proinflammatory cytokine genes, including TNF (TNF-α), upon stimulation with LPS and other TLR ligands. In this study, we examined the influence of T. gondii on transcription factors associated with TNF-α transcription, as well as phosphorylation and acetylation of histone H3 at distal and proximal regions of the TNF-α promoter. During LPS stimulation, we found that Toxoplasma blocks nuclear accumulation of transcription factor c-Jun, but not that of cAMP response element-binding protein or NF-κB. However, chromatin immunoprecipitation studies revealed that binding of all of these transcription factors to the TNF promoter was decreased by T. gondii infection. Furthermore, the parasite blocked LPS-induced Ser10 phosphorylation and Lys9/Lys14 acetylation of histone H3 molecules associated with distal and proximal regions of the TNF-α promoter. Our results show that Toxoplasma inhibits TNF-α transcription by interfering with chromatin remodeling events required for transcriptional activation at the TNF promoter, revealing a new mechanism by which a eukaryotic pathogen incapacitates proinflammatory cytokine production during infection.

The opportunistic intracellular protozoan Toxoplasma gondii is a potent trigger of Th1 cytokines, a response that enables host survival and long-term parasite persistence (1, 2). Proinflammatory cytokine induction must be tightly regulated, because when overproduced these mediators cause immunopathology and host death. For T. gondii, this is exemplified during infection of IL-10 knockout mice that succumb from the inability to down-regulate parasite-induced proinflammatory cytokine production (3). From the perspective of the parasite, preventing overinduction of cytokines such as TNF-α, IL-12, and IFN-γ has the dual benefit of keeping the host alive to allow persistence while avoiding immune-mediated elimination.

In recent years, it has become clear that Toxoplasma takes an active role in interfering with intracellular signaling leading to proinflammatory mediators including IL-12, TNF-α, and NO (4–8). The exact molecular mechanisms by which this occurs remain largely unknown, although interference with MAPK (3) activation, NFκB translocation, and activation of STAT3 has been implicated (4, 9–13). In our studies, we have focused on T. gondii-infected macrophages, where a large panel of LPS-responsive genes, including those encoding IL-12p40 and TNF-α, is suppressed (5). For the case of IL-12, inhibition is relieved after ~12 h, but TNF-α remains potently suppressed throughout the time course of infection (14). Because of the dramatic effect of Toxoplasma on macrophage TNF-α production, we chose to focus in detail on the induction of this mediator to gain insight into how the parasite interferes with host cell signaling.

Regulation of TNF-α production is complex and is controlled in a tissue-specific and stimulus-specific manner (15–17). The primary control step of TNF-α gene expression resides in transcription initiation (18, 19). Studies have established that NFAT, ATF-2, Jun, Ets/Elk, and Sp-1 transcription factors and CBP/p300 coactivator proteins are involved in regulation of TNF-α transcription (16, 19). NF-κB binding to distal κB sites is also important for maximal induction of TNF-α (20). Downstream of these events, production of TNF-α protein is also dependent upon regulation of mRNA splicing, regulation of mRNA half-life, and regulation of mRNA translation (21, 22).

Transcriptional initiation of many genes, including TNF-α, requires changes in higher order chromatin structure surrounding the promoter (23–25). In general, the promoter region undergoes stimulus-induced chromatin remodeling to allow access of transcription factors and RNA polymerase II (pol II)3 machinery. Chromatin remodeling usually includes histone phosphorylation, acetylation, and methylation, accompanied by nucleosome disassembly (26). The particular pattern of modification, also known as the histone code, determines whether a gene is in an active or inactive conformation (27). Among the histone modifications, phosphorylation of histone H3 at serine 10, acetylation of histone H3 at lysines 9 and 14, and methylation of histone H3 at lysine 4 are associated with gene activation.

In this study, we show that macrophage infection by T. gondii inhibits recruitment of RNA pol II to the TNF-α promoter during LPS stimulation. Focusing on TNF-associated transcription factors, we found that nuclear accumulation of c-Jun, but not NF-κB p65 or cAMP-responsive element-binding protein (CREB), was blocked by the parasite. Despite LPS induction of nuclear NF-κB and CREB in infected cells, the parasite prevented recruitment of each to their respective sites on the TNF promoter. Furthermore, Toxoplasma interfered with LPS-induced histone H3 phosphorylation and acetylation surrounding the TNF-α promoter, providing an explanation for the inability to recruit transcription factors and RNA pol II. These results show that T. gondii targets the histone modification machinery to prevent TNF-α transcription, and they provide a likely explanation for the widespread suppressive effects of the parasite on proinflammatory genes induced by LPS and possibly other stimuli.

Materials and Methods

Mice and parasites

C57BL/6 female mice (6–8 wk of age) were purchased from The Jackson Laboratory. The mice were kept under specific pathogen-free conditions at the Transgenic Mouse Facility, Cornell University College of Veterinary Medicine. The facility is overseen by an Institutional Animal Care and Use Committee. T. gondii parasite strains RH, CC, ENT, and DEG were maintained by biweekly passage on human foreskin fibroblast monolayers in DMEM supplemented with 1% FCS, 100 U/ml penicillin, and 0.1 mg/ml streptomycin. In some experiments, we used transgenic RH strain tachyzoites expressing tandem copies of the gene encoding yellow fluorescent protein (provided by D. Roos, University of Pennsylvania, Philadelphia, PA, and B. Striepen, University of Georgia, Athens, GA). Parasite cultures were tested for Mycoplasma every 6–8 wk using a highly sensitive PCR-based ELISA (Roche Diagnostics).

Cell culture

Bone marrow cells were flushed from femur and tibia and cultured in complete DMEM consisting of DMEM supplemented with 10% FCS, 1 mM sodium pyruvate, 0.1 mM nonessential amino acids, 20% supernatant from L929 cells, 100 U/ml penicillin, and 0.1 mg/ml streptomycin. The cells were supplemented with fresh macrophage medium on day 3. After 5 days of culture, nonadherent cells were removed, adherent monolayers were washed in ice-cold PBS, and cells were harvested by gentle pipetting in DMEM supplemented with 1% FCS, 100 U/ml penicillin, and 0.1 mg/ml streptomycin. Infection of macrophages was accomplished by adding tachyzoites to cell cultures followed by brief centrifugation (200 × g for 3 min) to synchronize contact between cells and parasites. In most cases, LPS (100 ng/ml) was added 12 h after infection. Cells were recovered at varying times as indicated, depending upon the assay performed.

Semiquantitative real-time PCR

Real-time PCR were performed with a Power SYBR green kit according to the manufacturer’s instructions (catalog no. 4367659; Applied Biosystems). The primers used were as follows. TNF RNA pol II site 1 forward: GAAAAGCA AGCAGCCAACCA; TNF RNA pol II site 1 reverse: CGGATCATGCTTTC TGTGCTC; TNF RNA pol II site 2 forward: ACAGAAAGCATGATCCGC GA; TNF RNA pol II site 2 reverse: GCCACAAGCAGGAATGAGAAGA; TNF forward: CCTTGTTGCCTCCTCTTTTGC; TNF reverse: TCAGTGAT GTAGCGACAGCCTG; IL-10 forward: CCTGGCTCAGCACTGCTAT; IL-10 reverse: GCTCTTATTTTCACAGGGGAGAA; TNF promoter proximal forward: CCCCAACTTTCCAAACCCTCT; TNF promoter proximal reverse: CCCTCGGAAAACTTCCTTGGT; TNF promoter distal forward: GG CTTGTGAGGTCCGTGAATT; TNF promoter distal reverse: CCCTCGGA AAACTTCCTTGGT; IL-10 promoter forward: GCAGAAGTTCATTCCGA CCA; IL-10 promoter reverse: GGCTCCTCCTCCCTCTTCTA; GAPDH forward: CCTGAACAGAACAGCAATGGCT; and GAPDH reverse: GCTTGACGGTGTCTTTTGCCT.

Cytokine ELISA

IL-10 and TNF-α in cell cultures were measured using commercial kits according to the manufacturer’s recommendations (R&D Systems).

Immunoblotting

The following Abs were used in immunoblotting studies: anti-phospho-c-Jun (catalog no. 2361; Cell Signaling), anti-total-c-Jun (catalog no. 9165; Cell Signaling), anti-phospho-CREB (catalog no. 9191; Cell Signaling), anti-total CREB (catalog no. 9197; Cell Signaling), anti-poly(ADP-ribose) polymerase (catalog no. 9542; Cell Signaling), anti-NF-κB p65 (catalog no. SC-109; Santa Cruz Biotechnology), anti-phospho-histone H3 (Ser10; catalog no. 9701; Cell Signaling), anti-acetyl histone H3 (Lys9/14; catalog no. 9677; Cell Signaling), anti-total H3 (catalog no. 9715; Cell Signaling), and anti-phospho-MSK1 (catalog no. 9591; Cell Signaling). Cells (2 × 106/sample) were lysed in reducing SDS sample buffer, and DNA was sheared by forcing samples five times through a 27-gauge needle. In some experiments, nuclear and cytoplasmic portions of cell lysates were separated by using a nuclear extract kit (Active Motif). After 5 min at 100°C, samples were separated by 10% SDS-PAGE and proteins were subsequently electrotransferred onto nitrocellulose membranes. Membranes were then blocked in 5% nonfat dry milk containing 0.1% Tween 20 in TBS, pH 7.6 (TBST), for 1 h at room temperature, followed by incubation with Ab reconstituted in 5% BSA in TBST overnight at 4°C. After washing blots in TBST, primary Abs were detected with a HRP-conjugated secondary Ab in TBST containing 5% nonfat dry milk for 1 h at room temperature. After washing in TBST, protein bands were visualized using a chemiluminescence-based detection system (Cell Signaling).

Chromatin immunoprecipitation (ChIP)

The following ChIP grade Abs were used for immunoprecipitation: anti-RNA pol II (catalog no. 39097; Active Motif), anti-NF-κB p65 (catalog no. SC-109; Santa Cruz Biotechnology), anti-phospho-CREB (catalog no. 9197; Cell Signaling), anti-phospho-c-Jun (catalog no. 2361; Cell Signaling), anti-phosphohistone H3 (Ser10; catalog no. 9701; Cell Signaling), and anti-acetyl histone H3 (Lys9/Lys14; catalog no. 9677; Cell Signaling). Assays were performed using the ChIP-IT enzymatic express kit (Active Motif) according to the manufacturer’s instructions. Briefly, cells (1.5 × 107/sample) were fixed using 1% formaldehyde at room temperature for 10 min. Fixation was stopped by adding glycine to the mixture. The cells were then collected by scraping in buffer containing PMSF (100 mM). After brief centrifugation, the cells were resuspended in lysis buffer (Active Motif) with a protease inhibitor mixture (Active Motif) and incubated for 30 min on ice. The cells were then resuspended in cell digestion buffer (Active Motif) and subjected to enzymatic digestion for 10 min at 37°C. The reaction was stopped with addition of 0.5 M EDTA. Abs were added into the sheared chromatin preparations and the mixture was incubated with Protein G Magnetic Beads (Active Motif) overnight at 4°C. The precipitated DNA-protein-Ab complexes were then washed and the cross-linking was reversed by incubation at 65°C for 4 h. Proteinase K was added to digest protein and DNA subsequently was purified using ethanol extraction, air dried, and redissolved in 100 µl of H2O. The retrieved DNA was then subjected to real-time RT-PCR amplification using promoter-specific primers.

Transcription factor DNA-binding ELISA

The presence of NF-κB p65 in nuclear extracts was determined by binding to plate-bound target oligonucleotides exactly according to the manufacturer’s instructions (Active Motif).

Immunofluorescence microscopy

Macrophages were plated onto coverslips and infected with RH strain tachyzoites at a parasite:cell ratio of 4:1. After overnight incubation, cells were treated with LPS (100 ng/ml) for 30–60 min. Coverslips bearing macrophages were fixed with 3% paraformaldehyde (20 min, room temperature), then permeabilized with methanol for 10 min at −20°C. Coverslips were blocked with 5% normal goat serum in PBST for 60 min, then rabbit anti-phospho-H3 (Ser10) Ab (catalog no. 9701; Cell Signaling) or rabbit anti-acetyl histone H3 (Lys9/14) (catalog no. 9677; Cell Signaling) was added in PBST to the coverslips, and cells were incubated overnight at 4°C. Coverslips were washed with PBS, then goat anti-rabbit Ab conjugated to Alexa Fluor 594 and anti-Toxoplasma p30 conjugated to FITC was added and cells were incubated for 2 h at 4°C. After washing in PBS, coverslips were mounted with ProLong Antifade containing 4′,5-dia-midino-2-phenylindoleI (Molecular Probes). Images were collected with a BX51 fluorescence microscope (Olympus) equipped with a DP 70 camera using DP controller software (version 1.1.65; Olympus) and DP manager software (version 1.1.1.71; Olympus).

Flow cytometry

Peritoneal cells from infected mice were washed in complete DMEM, resuspended at 1 × 107/ml, and plated in a 24-well plate. Brefeldin A (Golgi Plug; BD Biosciences) was added into the culture to block the protein transport. The cells were then stimulated with LPS. After 6 h of incubation, the cells were washed in FACS wash buffer (1% BSA in PBS) and plated 2 × 106/well in a 96-well plate. F4/80 APC Ab (catalog no. MF48005; Caltag Laboratories) was added and incubated for 20 min at 4°C. The cells were then washed twice with FACS wash buffer and incubated with 200 µl of Fix/Permeabilization buffer (BD Biosciences) for 15 min. The cells were washed again and resuspended in Ab mixture containing anti-TNF PE (catalog no. 554419; BD Biosciences) and anti-p30 FITC (catalog no. 12–132; Argene). After 1 h of incubation, cells were washed and resuspended in FACS wash buffer. Samples were analyzed on a FACSCalibur flow cytometer (BD Biosciences) collecting at least 50,000 events. The data were subsequently analyzed using FlowJo software (Tree Star).

Statistics

The statistical significance of the data was determined by an unpaired Student’s t test. Values of p < 0.05 were considered significant. All experiments were performed at least three times.

Results

T. gondii potently and specifically inhibits TLR-induced TNF-α production in macrophages

Previously, we established that T. gondii infection inhibits LPS induced production of TNF and other proinflammatory mediators in mouse macrophages (5, 14). To confirm and extend these results, we used the type I T. gondii RH strain to infect bone marrow-derived macrophages, cells that are readily invaded by the parasite (Fig. 1, A and B). When infected cells were subjected to LPS/TLR4 triggering, TNF-α cytokine release was potently suppressed (Fig. 1C). Parasite-induced suppression of TNF-α was maintained for up to 36 h after LPS stimulation (data not shown). This was not a nonspecific cytotoxic effect, because LPS-induced IL-10 production was unaffected by parasite infection (Fig. 1D). We also examined cytokine mRNA levels during LPS stimulation of infected cells. In accord with the cytokine protein data, LPS-induced TNF mRNA induction was inhibited by preinfection with T. gondii (Fig. 1E). In contrast, there was no consistent effect of the parasite on IL-10 mRNA during LPS stimulation (Fig. 1F).

FIGURE 1. T. gondii inhibits LPS-induced TNF-α, but not IL-10 production.

FIGURE 1

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A and B, Bone marrow-derived macrophages were infected (4:1 ratio of tachyzoites:cells) with a T. gondii RH strain genetically engineered to express tandem copies of yellow fluorescence protein (YFP). After 12 h, cells were prepared for fluorescence microscopy (A) and flow cytometry (B). C and D, Twelve-hour-infected macrophages were stimulated with LPS (■) or cultured in medium alone (□), and then supernatants were collected 6 h later for TNF-α (C) and IL-10 (D) ELISA. E and F, Macrophages were infected (4:1 ratio of parasites: cells) and then 12 h later subjected to LPS stimulation. At the indicated time points, RNA was extracted, reverse transcribed into cDNA, and real-time PCR was performed for TNF-α (E) and IL-10 (F) transcripts. The data are normalized to GAPDH expression levels. E and F, ■ and □ indicate LPS-induced responses of infected and noninfected cells, respectively.

T. gondii inhibits TNF-α induction by multiple TLR ligands and multiple parasite strains interfere with TNF-α induction

To examine whether T. gondii blocks TNF release triggered through other TLR pathways in addition to that initiated by LPS, we used Pam3Cys (TLR2 ligand) and CpG oligodinucleotides (TLR9 ligand) to stimulate infected macrophages. Fig. 2A shows that both TLR2-and TLR9-induced TNF responses were inhibited by RH strain infection. We also examined the activity of agonists directed at TLR3, 5, 7, and 8, and although none of these induced significant amounts of TNF-α, IL-12 production was suppressed during triggering through these TLR (data not shown).

FIGURE 2. T. gondii inhibits TNF-α induction by multiple TLR ligands and multiple parasite strains inhibit TNF-α induction.

FIGURE 2

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A, Bone marrow-derived macrophages were infected with RH strain tachyzoites for 12 h, followed by a 6-h stimulation with LPS, Pam3Cys, or CpG. Supernatants were collected for subsequent TNF ELISA. B, Macrophages were infected with RH, CC, ENT, or DEG strain tachyzoites for 12 h, followed by a 6-h stimulation with LPS. Supernatants were collected for subsequent TNF ELISA.

Next, we compared highly virulent type I Toxoplasma strains with low virulence type II strains to ask whether inhibition was specific for RH or whether other strains could suppress TNF-α production. As shown in Fig. 2B, although RH was the most potent suppressor of TNF-α, another type I strain (ENT) also blocked TNF production. In addition, the type II strains CC and DEG also possessed suppressive activity.

T. gondii blocks RNA pol II recruitment to the TNF gene

Regulation of TNF-α is complex, with control being exerted at transcriptional and posttranscriptional levels. To determine whether Toxoplasma acts at the transcriptional level to interfere with TNF-α production, we examined recruitment of RNA pol II to the TNF-α gene start codon using ChIP analysis with three independent sets of primers. As shown in Fig. 3A, LPS stimulated RNA pol II recruitment to the TNF-α promoter. In contrast, parasite infection alone failed to trigger recruitment. Importantly, when cells were preinfected with T. gondii, LPS stimulation failed to induce upregulation of RNA pol II activity at the TNF-α promoter. To substantiate this finding, we switched to a real-time PCR-based approach to examine ChIP products using two additional primer sets for amplification. In both cases, although LPS induced increased RNA pol II recruitment to the promoter, parasite infection blocked this response (Fig. 3, B and C). We conclude that T. gondii prevents TNF-α protein release, at least in part, through interference with recruitment of RNA pol II to the TNF gene transcription start site.

FIGURE 3. T. gondii blocks RNA pol II recruitment to the TNF gene.

FIGURE 3

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Macrophages were infected with RH strain tachyzoites for 12 h, followed by a 30-min stimulation with LPS. Nuclear extracts were prepared and subjected to a ChIP assay using Ab against RNA pol II. A, ChIP DNA was amplified by standard PCR methodology using primers spanning the 5′ region of the TNF gene, and input DNA was amplified as a control. B and C, ChIP DNA was amplified by real-time PCR using two different primer sets spanning the 5′ end of the TNF gene. The data was normalized to input DNA and amplification was expressed as relative enrichment compared with cells in medium (defined as 1). Med, Cells incubated in medium alone; Tg, cells infected with Toxoplasma. *, p < 0.01 comparing LPS and Tg + LPS.

T. gondii blocks LPS-induced nuclear accumulation of c-Jun, but not CREB or NF-αB p65

To elucidate the molecular basis by which T. gondii inhibits TNF-α gene induction, we first focused on transcription factors that target the TNF promoter (Fig. 4A). Previous studies have established that both proximal and distal regions of the promoter are important for maximal TNF induction (16, 19, 20). The distal region (−500 to −900) contains four NF-κB binding sites, and of these there is evidence that the κB2 and κB3 sites are the most active (20). The proximal region (0 to −200) contains binding sites for multiple transcription factors including CREB, ATF-2, c-Jun, Ets/Elk, Egr-1, and Sp1 (Fig. 4A). CREB, ATF-2, and c-Jun each recognize a conserved cAMP-responsive element palindrome. CBP/p300 serves as a platform to bring all of the transcription factors together to form an enhanceosome and also functions as a histone acetylase (28, 29).

FIGURE 4. T. gondii blocks LPS-induced nuclear translocation of c-Jun, but not that of CREB or NF-κB p65.

FIGURE 4

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A, Schematic map of the mouse TNF promoter showing distal and proximal sites involved in transcriptional initiation. B, Macrophages were either infected or left uninfected for 12 h and then subjected to LPS stimulation for the indicated time periods (min). Western blotting for total and activated forms of c-Jun and CREB was performed on nuclear lysates. Blotting for the nuclear enzyme poly(ADP-ribose) polymerase was performed to confirm equal protein loading. C, Cytoplasmic and nuclear extracts were prepared from infected and noninfected macrophages stimulated with LPS for the indicated time periods (min). The extracts were subsequently probed with Ab to NF-κB p65. In D, the in vivo-binding capability of nuclear NF-κB p65 was determined using an ELISA-based method to measure transcription factor binding to solid-phase target oligonucleotides. In this panel, addition of soluble target oligonucleotide (Oligo Comp) blocked p65 binding, confirming the specificity of the assay. Med, Cells cultured in medium alone; Tg, T. gondii-infected cells; n.s., not significant comparing LPS and Tg plus LPS.

We chose to examine the activation and binding properties of CREB, c-Jun, and NF-κB p65 upon TLR4 triggering of infected and noninfected cells. In Fig. 4B, we examined total and phosphorylated levels of c-Jun and CREB in macrophage nuclear extracts from noninfected and infected cells stimulated with LPS. Stimulation through TLR4 led to strong phosphorylation of c-Jun that was apparent at 30 min and sustained for at least 2 h. Interestingly, in cells preinfected with Toxoplasma, levels of nuclear phospho-c-Jun activation were lower. Most strikingly, we found a major decrease in total nuclear c-Jun following LPS stimulation of infected macrophages (Fig. 4B). LPS stimulation also resulted in modestly increased amounts of phospho-CREB in the nuclei of noninfected macrophages, but, with the possible exception of the 120-min time point, T. gondii preinfection failed to have a major effect on phosphorylated or total nuclear CREB levels (Fig. 4B).

Next, we examined the influence of T. gondii on LPS-induced NF-κB nuclear translocation. We and others previously reported that the parasite interferes with nuclear accumulation of this transcription factor, but this block was not sustained beyond 6 h of infection (9). In Fig. 4C, we show that LPS induces nuclear accumulation of NF-κB p65 within 30 min of stimulation that was maintained for at least 60 min. In accord with previous data, 12-h preinfected cells responded with similar rapid NF-κB p65 translocation in response to TLR4 triggering (Fig. 4C). We then assessed the binding activity of nuclear NF-κB p65 after LPS stimulation in the presence and absence of infection. As shown in Fig. 4D, LPS induced a 5-fold increase in NF-κB p65-binding activity, as measured in a binding assay using plate-bound κB consensus target oligonucleotides. As expected, parasite infection alone failed to increase nuclear NF-κB-binding activity and the presence of tachyzoites in macrophages did not significantly diminish the LPS-induced increase in nuclear p65 binding (Fig. 4D). We attempted a similar assay to assess CREB-binding activity, but were unable to obtain consistent results (data not shown).

T. gondii blocks binding of NF-κB p65 and phosphorylated c-Jun to the TNF promoter

We next determined the influence of Toxoplasma on recruitment of NF-κB p65, phospho-CREB, and phospho-c-Jun to the TNF-α promoter region in live cells using ChIP assays. As shown in Fig. 5A, LPS stimulated strong binding of p65 to the TNF distal promoter region. In sharp contrast, T. gondii infection potently blocked recruitment of p65 to κB sites on the TNF-α promoter. Thus, even though NF-κB translocated to the nucleus normally in infected cells (Fig. 4C) and even though this transcription factor was functional based upon in vitro binding assays (Fig. 4D), it was prevented from binding to its target promoter sequence (Fig. 5A). We observed a similar pattern when we examined the effect of Toxoplasma on phospho-CREB recruitment to the TNF promoter, although in this case parasite-mediated inhibition did not reach statistical significance. When we examined phospho-c-Jun, we found that LPS triggered strong binding to the TNF promoter and that this response was strongly down-modulated by prior T. gondii infection (Fig. 5C). Although the latter result was not entirely unexpected based upon decreased levels of total c-Jun in the nucleus during infection, it is notable that substantial amounts of phosphorylated nuclear c-Jun remained in infected cells (Fig. 4B).

FIGURE 5. T. gondii blocks LPS-induced binding of phosphorylated c-Jun and NF-κB p65 to the TNF promoter.

FIGURE 5

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Bone marrow-derived macrophages were infected with RH strain tachyzoites and then 12 h later subjected to a 30-min LPS stimulation. ChIP assays were performed using Abs specific for NF-κB p65 (A), phospho-CREB (B), and phospho-c-Jun (C). The immunoprecipitated DNA was amplified by real-time PCR using primers spanning either proximal (for c-Jun and CREB) or distal (for NF-κB p65) sites of the TNF promoter. The results were normalized to the input DNA and expressed as relative quantitation (RQ) where medium is defined as 1. Med, Medium; Tg, cells infected with T. gondii; n.s., not significant. *, p < 0.01 comparing LPS and Tg plus LPS.

Toxoplasma interferes with histone H3 modification associated with gene transcriptional activation

The finding that T. gondii potently blocked recruitment of transcription factors to the TNF-α promoter without interfering with activation or nuclear translocation suggested that the parasite might target chromatin structure rather than transcription factors per se. Therefore, we sought to determine the influence of Toxoplasma on Lys9/14 acetylation and Ser10 phosphorylation of histone H3, modifications that are associated with chromatin decondensation and gene activation (30). We found that even nonstimulated macrophages expressed high amounts of acetylated histone H3 and levels were not changed by LPS stimulation in the presence or absence of infection (Fig. 6A). These results were confirmed by immunofluorescence microscopy (Fig. 6B). However, we found that LPS induced phosphorylation of histone H3 within 60 min of infection, and this response was strongly inhibited in parasite-infected macrophages (Fig. 6A). Likewise, immunofluorescence microscopy confirmed inhibition of histone H3 phosphorylation in TLR4-triggered infected cells, in contrast to noninfected macrophages that were stimulated with LPS (Fig. 6, B and C). The kinase MSK1 elicits histone H3 phosphorylation on the Ser10 residue (31, 32). Therefore, we asked whether LPS triggered activation of MSK1, and, in turn, whether T. gondii blocked the response. Although LPS stimulated rapid activation of MSK1, the response was unaffected by prior infection with Toxoplasma (Fig. 6D). We also examined histone H3 Thr3 and Ser28 modification, but found little or no effect of Toxoplasma on the phosphorylation status of these residues (data not shown).

FIGURE 6. T. gondii globally blocks LPS-induced phosphorylation of histone H3 at Ser10 but has no effect on Lys9/14 histone H3 acetylation.

FIGURE 6

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A, Macrophages were infected and then 12 h later subjected to LPS stimulation for the indicated time periods (min). Total cell lysates were extracted and subjected to Western blotting using Abs specific to phospho-Ser10 histone H3, acetyl Lys9/14, and total H3. In B, fluorescence microscopy was used to examine Lys9/14 acetylation (top panels) and Ser10 phosphorylation (bottom panels) of histone H3 in infected and noninfected cells stimulated for 30 min with LPS. Macrophages were stained with Ab specific for acetylated or phosphorylated histone H3 as indicated (red) and Toxoplasma p30 (green). Nuclei are stained with 4′,6-diamidino-2-phenylindole (blue). In C, cells that were positive for phospho-Ser10 at histone H3 staining were counted in each group of cells (~200 cell/condition). *, p < 0.01 comparing infected and noninfected cells. D, Twelve-hour-infected and control macrophages were stimulated with LPS (100 ng/ml) and cell lysates were prepared at the indicated time points (min). Immunoblotting for phospho-MSK1 was subsequently performed. As a control for protein loading, levels of histone H3 were assessed in the same samples. Med, Medium; Tg, T. gondii.

We next wanted to specifically examine the status of histone H3 modification surrounding distal and proximal regions of the TNF-α promoter during macrophage activation in the presence and absence of infection, since global levels of H3 phosphorylation and acetylation might not be reflective of histone modification at the TNF promoter. Therefore, we examined histone modifications at this site by ChIP assay. We found that LPS induced elevated levels of phosphorylation of histone H3 at Ser10 at both distal and proximal TNF promoter regions (Fig. 7, A and C). Notably, this response was blocked by Toxoplasma infection. Furthermore, we also examined acetylation of histone H3 at Lys9 and Lys14 at proximal and distal locations of the TNF promoter (Fig. 7, B and D). Although LPS induced H3 acetylation at both sites, this response was blocked in infected macrophages. Finally, we examined the status of histone H3 surrounding the IL-10 promoter, an LPS-induced gene that is not affected by Toxoplasma (Fig. 1). In this case, there was no evidence for histone H3 modification following TLR4 stimulation as measured by phosphorylation (Fig. 7E) or acetylation (Fig. 7F), and infection with T. gondii had no influence in the presence or absence of LPS. Therefore, although global levels of acetylated histone H3 are not affected by Toxoplasma, the parasite down-regulates acetylated H3 at the TNF promoter. In contrast, the parasite blocks LPS-induced Ser10 H3 phosphorylation at the global level and this extends to the activity at the TNF promoter.

FIGURE 7. T. gondii blocks LPS-induced histone H3 modification at the TNF promoter.

FIGURE 7

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Macrophages were infected with T. gondii and then 12 h later subjected to LPS stimulation for 30 min. The cells were fixed with formaldehyde and a ChIP assay was performed using Abs specific for Ser10 phosphorylated (A and C) or Lys9/14 acetylated (B and D) histone H3. The resulting DNA was amplified by real-time PCR using primers spanning either proximal (A and B) or distal (C and D) sites of the TNF promoter region. As a control, the same ChIP DNA was amplified using primers spanning the IL-10 promoter region (E and F). The results were normalized to the input DNA and expressed as relative quantitation (RQ), with medium defined as 1. Med, Cells incubated in medium alone; Tg, cells infected with Toxoplasma. *, p < 0.05 comparing LPS and Tg plus LPS.

Toxoplasma suppresses TNF-α production during in vivo infection

We wanted to determine whether parasite suppression of bone marrow-derived macrophage TNF-α production extended to in vivo infection. Therefore, we assessed production of this cytokine by inflammatory macrophages recruited to the peritoneal cavity during infection. Mice were inoculated by i.p. injection with Toxoplasma and peritoneal wash-out cells were collected 12 h later. The cells were cultured in vitro in medium or LPS, and TNF-α production was assessed in infected and noninfected populations of F4/80-positive macrophages (Fig. 8A). As shown in Fig. 8B, in cells cultured without further stimulation, 13% of noninfected macrophages produced TNF-α. In contrast, only 3% of infected macrophages produced this cytokine in the same overall population. When F4/80-positive macrophages were stimulated ex vivo with LPS (Fig. 8C), 84% of noninfected macrophages produced TNF-α. Strikingly, in the infected cell population, only 25% stained positive for TNF-α in response to ex vivo TLR4 triggering. We note that in the F4/80-positive population, a distinct subset of cells harbors low parasite levels and can respond to LPS (Fig. 8C), a distribution pattern that we do not see in vitro (Fig. 1B). We do not know the origin of these macrophages at present. One possibility is that these are cells that were newly infected during the in vitro response to LPS. Another possibility is that this population represents a macrophage subset that is less permissive to T. gondii replication and is also less sensitive to parasite-induced inhibition of TNF-α. Regardless, our data for the first time show that Toxoplasma shuts down proinflammatory signaling in an in vivo situation.

FIGURE 8. T. gondii inhibits TNF-α production during in vivo infection of F4/80-positive macrophages.

FIGURE 8

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Mice were infected by i.p. injection of 106 RH strain tachyzoites. After 12 h, cells in the peritoneal cavity were collected and cultured for 6 h in either medium alone or LPS in the presence of brefeldin A. Cells were then subjected to surface staining for F4/80 and intracellular staining for TNF-α and Toxoplasma p30 (SAG-1) to detect tachyzoites. A, F4/80-positive cells in the peritoneal cavity. The rectangle delineates cells selected for analysis. B, TNF-α and infection levels in F4/80-positive cells cultured in medium. C, TNF-α and infection levels in F4/80-positive cells stimulated in vitro with LPS.

Discussion

Microbial pathogens such as Toxoplasma must evade the potent antimicrobial effects of proinflammatory mediators produced by the innate immune system if they are to establish persistent infection in the host. For intracellular pathogens, direct interference with gene transcription is a highly effective strategy to achieve this end (33, 34). Inflammatory gene expression is controlled by multiple regulatory signal transduction cascades (35). First, gene-specific transcription factors must be activated and translocated to the nucleus. Second, it is becoming increasingly realized that in many cases chromatin structure surrounding target genes must be modified to allow access of transcription factors (36–38). There are many examples of pathogens that target pathways leading to transcription factor activation. A more limited number of recent studies have found that some bacterial pathogens epigenetically affect gene function (39–42). This study is the first example of a eukaryotic pathogen that targets regulatory cascades controlling chromatin structure to subvert host cell function.

The four core histones (H2A, H2B, H3, and H4) that make up the nucleosome are arranged as an octamer, around which DNA is wound. The N termini of the histones are subject to covalent modification that can be associated with repression or induction of genes (36–38). Transcriptional activation is strongly associated with histone H3 modification. The pattern of chemical changes is complex but among the modifications, phosphorylation of Ser10 and acetylation of Lys9 and Lys14 are linked to changes in chromatin structure that permit access of transcription factors (30).

Our results for the first time show that T. gondii prevents chromatin remodeling surrounding the TNF-α promoter by interfering with phosphorylation and acetylation of histone H3 that would otherwise be induced by TLR signaling. We found that Toxoplasma blocks Ser10 phosphorylation of total histone H3 in the nucleus, as well as at distal and proximal TNF-α promoter regions. Our studies in addition show that T. gondii interferes with LPS-induced acetylation of histone H3 at Lys9 and Lys14 surrounding the TNF promoter. Interestingly, this was a gene-specific effect because global levels of acetylated H3 were constitutively high in macrophages, and neither LPS nor the combination of Toxoplasma and LPS altered these levels. Ser10 phosphorylation of histone H3 can recruit several histone acetyl transferases to catalyze acetylation of Lys9 and Lys14 (43, 44). Therefore, it is possible that the effects of Toxoplasma on histone H3 acetylation at the TNF promoter are indirectly mediated by inhibition of H3 Ser10 phosphorylation. Insofar as chromatin modification is a prerequisite to allow transcription factor access at many proinflammatory genes, our results may provide a unifying model for the parasite’s ability to simultaneously down-regulate production of several proinflammatory mediators in addition to TNF-α (5, 14).

Although Toxoplasma down-modulates many TLR4-inducible genes, some escape inhibition. Most prominently, we found that LPS-triggered IL-10 is not blocked by the parasite. We also found no evidence that LPS altered chromatin modification surrounding the IL-10 promoter. Thus, the relatively low amount of LPS-triggered IL-10 compared with TNF-α appears to be produced independently of histone H3 Ser10 phosphorylation or Lys9/14 acetylation and therefore escapes inhibition by the parasite. In contrast, there is evidence that high level IL-10 transcription induced by other stimuli such as combined treatment with LPS and immune complexes involves chromatin remodeling at the IL-10 promoter (45). We are currently examining whether these responses are likewise affected by Toxoplasma.

Although interference with chromatin remodeling is likely to be a potent parasite mechanism to manipulate the host cell transcriptome, we also found evidence for interference with c-Jun activity. Thus, total levels of nuclear c-Jun rapidly decreased following LPS stimulation of infected macrophages. One interpretation of this result is that TLR4 triggering causes phosphorylation of nuclear c-Jun, and Toxoplasma increases nuclear export of the activated form of this transcription factor. Unlike the results of the 12-h infections reported here, the parasite also prevents nuclear accumulation of NF-κB during short-term infection. Whether this reflects decreased nuclear import or increased export is unclear (4, 10, 46).

Toxoplasma is sequestered within a parasitophorous vacuole, but recent findings show that the parasite delivers molecules to the host cell to modify signal transduction (47). During host cell invasion, apical organelles called rhoptries discharge and at least some rhoptry proteins enter the host cell. The ROP16 molecule, a rhoptry protein with predicted kinase activity, is released during invasion and targets STAT3/6 for activation (12). STAT3 activation by T. gondii has previously been implicated in suppression of IL-12 by the parasite (11). In addition a protein phosphatase 2C released by the parasite is targeted to the host cell nucleus, although in this case host target molecules have not yet been identified (48). Nevertheless, the emerging opinion is that T. gondii takes an active part in manipulation of the host cell (13, 47, 49), and the results of the present study provide strong mechanistic evidence for this view.

Toxoplasma inhibits signaling through TLR, yet the parasite itself expresses its own TLR ligands. Tachyzoite profilin activates TLR11 and glycosylphosphoinositol anchors on the parasite surface are capable of triggering TLR2 and TLR4 (50, 51). The parasite cannot survive without profilin, which is used in invasion (52). Likewise, the parasite is not viable without the ability to synthesize glycosylphosphoinositol-anchored proteins (53). Possibly for these reasons T. gondii has adopted potent strategies to down-modulate proinflammatory signaling that would otherwise result from recognition of its own essential molecules by host pattern recognition receptors.

Several bacterial pathogens target proinflammatory signaling pathways in macrophages. The Yersinia virulence proteins YopP/J block phosphorylation of IκB and MAPK kinase, effectively terminating both NF-κB and MAPK signal transduction (54–57). Along similar lines, Bacillis anthracis lethal factor enzymatically cleaves MAPK kinases (58). Some bacterial pathogens can also affect chromatin structure. Mycobacterium tuberculosis down-regulates histone acetylation around the MHC2TA locus, and Listeria monocytogenes dephosphorylates histone H3 and deacetylates histone H4 during early infection (39–41). It has recently been reported that Shigella flexneri escapes inflammatory chemokine induction through effects of the OspF protein on histone H3 phosphorylation (42). Intracellular protozoans are now also emerging as master evaders of immunity (34). Toxoplasma has negative effects on MAPK, NF-κB, and STAT1 signaling pathways, although the mechanisms involved are less well defined (9, 59, 60). Our present studies reveal gene-specific targeting of chromatin structure as a new and profound strategy used by this intracellular protozoan to manipulate host cell responses.

Acknowledgments

We thank S. Coonrod for critical review of this manuscript.

Footnotes

1

This work was supported by National Institutes of Health Grant AI50617 (to E.Y.D.) and a Genomics Scholars Award from the Cornell University Center for Vertebrate Genomics (to J.L.).

3

Abbreviations used in this paper: pol II, polymerase II; CREB, cAMP-responsive element-binding protein; ChIP, chromatin immunoprecipitation.

Disclosures

The authors have no financial conflict of interest.

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