Cross-Talk in the Innate Immune System: Neutrophils Instruct Recruitment and Activation of Dendritic Cells during Microbial Infection

Soumaya Bennouna, Susan K. Bliss, Tyler J. Curiel and Eric Y. Denkers
J Immunol December 1, 2003, 171 (11) 6052-6058; DOI: https://doi.org/10.4049/jimmunol.171.11.6052

Abstract

Type I inflammatory cytokines are essential for immunity to many microbial pathogens, including Toxoplasma gondii. Dendritic cells (DC) are key to initiating type 1 immunity, but neutrophils are also a source of chemokines and cytokines involved in Th1 response ignition. We found that T. gondii triggered neutrophil synthesis of CC chemokine ligand (CCL)3, CCL4, CCL5, and CCL20, chemokines that were strongly chemotactic for immature DC. Moreover, supernatants obtained from parasite-stimulated polymorphonuclear leukocytes induced DC IL-12(p40) and TNF-α production. Parasite-triggered neutrophils also released factors that induced DC CD40 and CD86 up-regulation, and this response was dependent upon parasite-triggered neutrophil TNF-α production. In vivo evidence that polymorphonuclear leukocytes exert an important influence on DC activation was obtained by examining splenic DC cytokine production following infection of neutrophil-depleted mice. These animals displayed severely curtailed splenic DC IL-12 and TNF-α production, as revealed by ex vivo flow cytometric analysis and in vitro culture assay. Our results reveal a previously unrecognized regulatory role for neutrophils in DC function during microbial infection, and suggest that cross-talk between these cell populations is an important component of the innate immune response to infection.

The intracellular protozoan parasite Toxoplasma gondii is a major opportunistic parasite in immunocompromised patients, and 30–50% of the population worldwide is asymptomatically infected with this microbial pathogen (1, 2). T. gondii normally elicits a robust type 1 cytokine response, in which CD4+ and CD8+ T lymphocytes produce IFN-γ that is essential in mediating resistance to infection (3, 4). However, in the genetic absence of IL-10 and during oral infection of certain inbred mouse strains, Toxoplasma triggers dysregulated type 1 cytokine production, leading to pathology and death (5, 6, 7).

Understanding how the type 1 cytokine response is initiated during infection with Toxoplasma and other microbial pathogens is an area of major interest. Dendritic cells (DC),3 through their ability to capture Ag, migrate to secondary lymphoid organs, simultaneously present antigenic peptide, and release IL-12, are important in driving Th1 differentation (8, 9). Indeed, injection of soluble tachyzoite (TZ) Ag, and infection with live parasites, results in rapid activation of IL-12-producing DC in the spleen (10, 11). Nevertheless, the cellular and molecular events leading to DC activation and Th1 differentiation during in vivo infection are ill-defined.

Recent studies have shown that T. gondii infection induces rapid influx of polymorphonuclear leukocytes (PMN) to the site of infection, a response that is dependent upon chemokine receptor CXCR2 (12, 13). Neutrophils produce several important proinflammatory cytokines and chemokines, including IL-12, TNF-α, CC chemokine ligand (CCL)3 (macrophage inflammatory protein (MIP)-1α) and CCL4 (MIP-1β) during in vitro stimulation with T. gondii and other microbial pathogens (14, 15). Most importantly, mice depleted of PMN using an Ab against Gr-1 (Ly6G) are unable to survive acute toxoplasmosis. Lack of resistance in neutrophil-depleted animals is associated with defective type 1 cytokine responses during infection with Toxoplasma and several other microbial pathogens (16, 17, 18, 19, 20). Collectively, these data suggest that PMN may play a role in orchestrating early immunity through production of cytokines and chemokines that promote development of Th1 T lymphocytes.

Although PMN are well-known as the first cell type to arrive at the site of infection, it is not clear how they could influence T cell differentiation, which is conventionally thought to be driven by DC in secondary lymphoid organs (21). In this study, we present an explanation for this conundrum. We found that parasite-triggered neutrophils release CCL3, CCL4, CCL5 (RANTES), and CCL20 (MIP-3α), chemokines that together display potent chemotactic activity for immature bone marrow-derived DC. Parasite-stimulated PMN also release soluble factors that trigger DC activation, as measured by IL-12(p40) and TNF-α production, as well as up-regulation of costimulatory molecules CD40 and CD86. We demonstrate that DC activation is driven at least in part by PMN-derived TNF-α. The physiological relevance of these data is suggested by the finding that in vivo PMN depletion results in defective splenic DC cytokine responses during infection. The data point to a model in which neutrophils instruct DC recruitment and activation, leading in turn to Th1 cell activation and ultimately immunity to microbial infection.

Materials and Methods

Mice

C57BL/6 and Swiss-Webster female mice, 6–8 wk of age, were obtained from Taconic Farms (Germantown, NY). TNF knockout and wild-type counterparts (B6129SF2/J) were purchased from The Jackson laboratory (Bar Harbor, ME). Animals were housed in filter-covered isolator cages in the animal facility of the College of Veterinary Medicine at Cornell University (Ithaca, NY) which is accredited by the American Association for Accreditation of Laboratory Care.

Parasites and infections

TZ of the virulent T. gondii strain RH were maintained by biweekly passage on human foreskin fibroblasts in complete medium (cDMEM) composed of DMEM (Life Technologies, Gaithersburg, MD) supplemented with 1% FCS (HyClone Laboratories, Logan, UT), 100 U/ml penicillin, and 100 μg/ml streptomycin (PenStrep; Life Technologies). T. gondii cysts were harvested from brain homogenates of Swiss-Webster mice that were infected 1 mo earlier with the ME49 parasite strain. Infections were accomplished by i.p. injection of 20 ME49 cysts. In vivo neutrophil depletion was accomplished by i. p. injection of 200 μg of RB6-8C5 or NIMP-R14 mAb every 48 h.

Reagents and Ab

LPS (Escherichia coli strain 0111:B4) and fMLP were purchased from Sigma-Aldrich (St. Louis, MO). FITC-conjugated Abs specific for CD11c and Ly6G, PE-conjugated Abs specific for IL-12 (p40/p70) (C15.6), TNF-α (MP6-XT22), class II MHC I-Ab (AF6-120.1), CD40 (3/23), CD80 (16-10A1), CD86 (GL1), Gr-1(Ly-6G), and purified rat-anti MIP-1β (A 65-2), and anti-TNF-α blocking Abs (MP6-XT3 and G281-2626) were obtained from BD PharMingen (San Diego, CA). Purified antisera specific for MIP-1α (M20), MIP-3α (A-20), and RANTES (C-19) were obtained from Santa Cruz Biotechnology (Santa Cruz, CA). Normal rat and goat Ig were purchased from Jackson ImmunoResearch Laboratories (West Grove, PA). Anti-CD11c and anti-MHC class II magnetic microbeads were purchased from Miltenyi Biotec (Auburn, CA). RB6-8C5 mAb (IgG2b) was kindly provided by Dr. R. L. Coffman (DNAX Research Institute, Palo Alto, CA). NIMP-R14, a rat IgG2b Ab that selectively binds to mouse neutrophils (22), was provided by Dr. F. Tacchini-Cottier (World Health Organization Immunology Research and Training Center, University of Lausanne, Epalinges, Switzerland).

Bone marrow PMN purification

Neutrophils were isolated from mouse bone marrow following a previously published protocol (23). Briefly, single cell suspensions of bone marrow cells were collected from femur and tibia and resuspended in DMEM supplemented with 5% FCS and 1% PenStrep. Cells were then centrifuged at 500 × g for 7 min at 4°C and resuspended in HBSS (Ca2+-free) supplemented with 0.38% sodium citrate. The cell suspension was layered on top of a step gradient consisting of 52, 65, and 75% Percoll diluted in Ca2+-free HBSS, and centrifuged at 1500 × g for 30 min at 4°C. Neutrophils were recovered at the interface of the 65 and 75% Percoll layers. The proportion of neutrophils, determined by Diff-Quik staining of cytospin preparations, was routinely >90%.

Isolation of peritoneal neutrophils

Mice were i.p. injected with 1 ml of 10% thioglycollate (Difco Laboratories, Detroit, MI). Eighteen hours later, peritoneal exudate cells (PEC) were obtained by lavage with ice-cold PBS. PEC were washed in PBS, passed through a 70-μm nylon cell strainer, and erythrocytes in the suspension were lysed using Red Cell Lysis Buffer (Sigma-Aldrich). PEC were then washed, resuspended to 2 × 107 cells/ml in MACS buffer composed of Dulbecco’s PBS (Life Technologies) containing 0.5% BSA (Sigma-Aldrich), 1 mM EDTA (Fisher Scientific, Pittsburgh, PA), and incubated for 15 min at 4°C with anti-MHC class II magnetic microbeads. After washing, the mixture was transferred to columns installed within a magnetic apparatus to remove MHC class II-expressing cells, according to the manufacturer’s instructions (Miltenyi Biotec).

Bone marrow-derived DC cultures

Generation of bone marrow-derived DC was accomplished following a previously published protocol (24). Briefly, single cell bone marrow preparations were obtained as described above, cells were washed in RPMI 1640 (Fisher Scientific) and resuspended at 2 × 105 cells/ml in bone marrow-derived DC medium composed of RPMI 1640 supplemented with 1% PenStrep, 10% FCS, 50 μM 2-ME, and 20 ng/ml GM-CSF (Peprotech, Rocky Hill, NJ). Cells were plated on 100 × 15 mm standard sterile polystyrene Petri dishes (Fisher Scientific) and cultured for 9 days at 37°C in 5% CO2. Fresh DC medium, containing GM-CSF, was added on days 3, 6, and 8 after culture initiation. On day 9, cells were resuspended in cDMEM alone or in the presence of different stimuli at 37°C in 5% CO2. Eighteen hours later, supernatants from the cultures were recovered and either used immediately or stored at −80°C.

Splenic CD11c+ DC isolation

DC in the spleen were isolated as described elsewhere (11). Briefly, splenocyte suspensions were prepared in DMEM supplemented with PenStrep, 10 mM HEPES, and 1 mM EDTA and erythrocytes were lysed in Red Cell Lysis Buffer (Sigma-Aldrich). The remaining cells were washed, resuspended at 2.5 × 108 cells/ml in MACS buffer, then incubated with CD11c-conjugated magnetic microbeads for 15 min at 4°C. After washing, the cell suspension was passed through a column (25 MS; Miltenyi Biotec) in the presence of a magnet, and unbound cells were removed by several washes in MACS buffer. Columns were then removed from the magnetic field, and CD11c+-enriched cells were flushed through, washed, and resuspended in cDMEM for culture. Cell populations isolated in this manner were routinely >70% CD11c+.

PMN cultures

Bone marrow and peritoneal PMN (2 × 106/ml) were cultured in the presence of TZ (0.5:1 ratio of parasites to cells) at 37°C in 5% CO2 in 96-well tissue culture plate (Corning Costar, Cambridge, MA). Eighteen hours later, supernatants from the cultures were recovered, filtered through a 0.2-μm membrane (Corning, Corning, NY), and either used immediately or stored at −80°C. To remove specific chemokines, peritoneal PMN supernatants were incubated with anti-chemokine Ab (each at 15 μg/ml) at 4°C on a shaker for 2 h, then protein G-agarose beads (Santa Cruz Biotechnology) were added to supernatants for an additional 4 h under the same conditions. The supernatants were then spun at 1000 rpm for 4 min and the bead-free supernatants were collected and either used immediately or stored at −80°C. Anti-TNF-α treatments were performed with a combination of two neutralizing Ab at saturating concentrations (BD PharMingen) or control Ab in PMN supernatants during DC treatment.

RT-PCR

RNA was isolated, reverse-transcribed, and subjected to PCR amplification as described (25). The primer sequences used were: β-actin, TGACGGGGGTCACCCACACTGTGCCCATCTA (sense), CTAGAAGCATTGCGGTGGACGATGGAGGG (antisense); CCL3, CGGAAGATTCCACGCCAATTC (sense), GGTTGAGGAACGTGTCCTGAAG (antisense); CCL4, CCCACTTCCTGCTGTTTCTCTTAC (sense), AGCAGAGAAACAGCAATGGTGG (antisense); CCL5, CCACGTCAAGGAGTATTTCTACACC (sense), CTGATTTCTTGGGTTTGCTGTG (antisense); CCL20, TACTCCACCTCTGCGGCGAATCAGAA (sense), GTGAAACCTCCAACCCCAGCAAGGTT (antisense). The cDNA was amplified 27 cycles (CCL3), 29 cycles (CCL4, CCL5 and β-actin), and 35 cycles (CCL20) (20).

Flow cytometry

To analyze DC surface markers, Fc receptors were blocked in FACS buffer (PBS, 1% BSA, and 0.1% sodium azide) containing 10% normal mouse serum for 15 min at 0°C, then cells were stained with optimal concentrations of FITC-conjugated anti-CD11c in combination with PE-conjugated antisera specific for class II MHC, CD40, CD80, and CD86 for 30 min at 0°C. For intracellular cytokine detection, splenic CD11c+ were blocked, stained with FITC-conjugated anti-CD11c, then cells were fixed in 3% paraformaldehyde (Sigma-Aldrich), 0.1 mM CaCl2, and 0.1 mM MgCl2 for 30 min at 0°C. CD11c+ cells were subsequently washed in permeabilization buffer (PBS with 0.075% saponin) and incubated for 15 min at 0°C in permeabilization buffer containing 10% normal mouse serum. After two washes in permeabilization buffer, PE-conjugated anti-IL-12 or anti-TNF-α or control Ab was added, cells were incubated for 30 min at 0°C and subsequently washed for flow cytometric analysis. Data was acquired on a FACSCalibur system (10,000 events per sample) and analyzed with CellQuest software (BD Immunocytometry Systems, San Jose, CA).

Chemotaxis

Cell migration was assessed using a disposable 96-well chemotaxis chamber (ChemoTx no. 101-5; Neuroprobe, Gaithersburg, MD). The wells contained either medium, fMLP (10−6 M), or test sample supernatants. The framed polycarbonate filter (5-μm pore size) was installed over wells and 30 μl of cells, resuspended in cDMEM (1.5 × 106/ml), were added to the filter surface. In control wells, cells were directly resuspended in supernatants from PMN-TZ cocultures. After incubation (90 min at 37°C in 5% CO2), the number of cells that migrated through the filter into the wells was counted in five high power fields under a phase-contrast microscope.

Cytokine ELISA

IL-12(p40) was measured as previously described (26), and TNF-α was measured using a commercially obtained kit (BD PharMingen).

Statistical analysis

The statistical significance of the data was analyzed using an unpaired two-tailed Student’s t test.

Results

Toxoplasma stimulated PMN release factors chemotactic for DC

To determine whether PMN exposed to T. gondii released soluble factors capable of influencing DC activity, we analyzed cell-free supernatants from PMN-TZ cocultures for their ability to elicit bone marrow DC chemotaxis. Under the conditions used, ∼60% of PMN were infected at the termination of the cocultures. Supernatants from PMN-TZ cocultures contained factors strongly chemotactic for immature DC (Fig. 1⇓A). T. gondii was required to elicit this response, since PMN cultured in medium alone (PMN-M) failed to release DC chemotactic factors. To distinguish increased chemotaxis vs chemokinesis, PMN-TZ supernatant was placed in the same chamber as the DC in the cell migration assay (Fig. 1⇓A, Contr). In this case, movement into the counting chamber did not occur. Supernatants collected from parasites in the absence of PMN failed to induce DC chemotaxis, implicating PMN factors released in response to the parasite (data not shown).

FIGURE 1.
FIGURE 1.

Toxoplasma stimulated PMN release DC chemotactic factors. A, Bone marrow-derived DC chemotaxis assay in presence of medium alone (M), cell-free supernatants derived from PMN cultured in medium alone (PMN-M), or from PMN infected with TZ (PMN-TZ). Contr, control for chemotaxis, in which PMN-TZ supernatant was placed in the same chamber as the DC. B, RT-PCR analysis of chemokines expressed by PMN incubated with medium alone or cocultured with TZ for 2 h. C, DC chemotaxis assay with cell-free supernatants incubated with control Ab (□), a combination of anti-CCL3 (MIP-1α), -CCL4 (MIP-1β), and -CCL5 (RANTES) Ab (▨), anti-CCL20 (MIP-3α) mAb (▦), and a combination of anti-CCL3, -CCL4, -CCL5, and -CCL20 Ab (▪). Data are representative of four different experiments. ∗, p < 0.01 (PMN-TZ supernatants + control Ab vs PMN-TZ supernatants + anti-CCR5 ligands); ∗∗, p < 0.01 (PMN-TZ supernatants + anti-CCR5 ligands vs PMN-TZ supernatants in presence of a combination of all four Ab).

The CCR5 ligands CCL3, CCL4, and CCL5, as well as the CCR6 ligand CCL20, function as factors chemotactic for immature DC (27, 28). Therefore, we asked whether T. gondii could induce PMN synthesis of these chemokines. As shown in Fig. 1⇑B, mRNA for CCL3, CCL4, CCL5, and CCL20 were rapidly up-regulated in PMN cocultured with T. gondii. To determine whether these chemokines were involved in the DC chemotactic activity of PMN-T. gondii coculture supernatants, we depleted chemokines using anti-chemokine Ab conjugated to protein G-agarose beads. Depletion of CCL3, CCL4, and CCL5 decreased the chemotactic effect of supernatants derived from PMN-TZ cultures. Anti-CCL20 Ab induced a greater decrease in the chemotactic index and a combination of both anti-CCR5 ligand and anti-CCL20 Ab reduced the level of migration to that obtained in medium alone (Fig. 1⇑C). Importantly, these Ab had no effect on DC chemotaxis mediated by fMLP, a chemotactic peptide that exerts its effects through binding to its own unique receptors (29) (Fig. 1⇑C).

TZ-stimulated PMN activate immature DC

The ability of PMN factors, as well as T. gondii itself, to induce immature DC maturation was determined by measuring up-regulation of the costimulatory molecule CD40. As shown in Fig. 2⇓, relative to medium (Fig. 2⇓A), LPS (Fig. 2⇓B) induced strong CD40 up-regulation. Direct DC infection (Fig. 2⇓C) and DC incubation with secreted parasite products (Fig. 2⇓D) induced low levels of activation. Supernatants from unstimulated PMN also induced a low amount of CD40 up-regulation (Fig. 2⇓E), but culture fluid from PMN-T. gondii cocultures displayed strong DC activating capability (Fig. 2⇓G). Importantly, the ability of PMN supernatants to induce CD40 up-regulation was abrogated with neutralizing anti-TNF-α mAb (Fig. 2⇓H). Similar results were obtained when MHC class II and CD86 expression was examined (data not shown).

FIGURE 2.
FIGURE 2.

TZ-stimulated neutrophils release factors inducing DC CD40 up-regulation. Bone marrow-derived DC were cultured with (A) medium, (B) LPS (1 μg/ml), (C) live TZ (0.5:1 ratio of parasites to cells), (D) TZ supernatants, (E) cell-free supernatants from PMN in medium (PMN-M) in presence of control Ab, (F) PMN-M with anti-TNF-α blocking Ab, (G) cell-free supernatants from PMN cocultured with live TZ (PMN-TZ) in presence of control Ab, (H) PMN-TZ with anti-TNF-α blocking Ab. After 18 h, DC were subjected to FACS analysis using anti-CD11c and anti-CD40 Ab. The results show the gated CD11c+ population stained with anti-CD40 (bold line) and an isotype control (thin line). The numbers indicate percent of cells positive for CD40. This experiment was repeated twice with similar results.

These results suggested involvement of TNF-α in DC activation by PMN-TZ, but it was possible that PMN-TZ supernatants contained factors eliciting DC TNF-α, which then activated the cells in an autocrine manner. To determine the source of TNF-α, neutrophils were isolated from TNF-α knockout mice and subjected to T. gondii coculture. Supernatants from TNF-α-deficient PMN-TZ cultures failed to induce DC up-regulation of either CD40 or CD86 (Fig. 3⇓). In contrast, wild-type PMN-TZ supernatants induced DC costimulatory molecule up-regulation that achieved levels similar to that induced by LPS. The results demonstrate that neutrophil-derived TNF-α plays an important role in DC activation, as measured by CD40 and CD86 up-regulation.

FIGURE 3.
FIGURE 3.

PMN-derived TNF-α is required to induce DC costimulatory molecule up-regulation. DC were cultured in the presence of medium (M), LPS, or PMN supernatants (SN) prepared from either wild-type (WT) or TNF-α knockout (KO) neutrophils. PMN SN were prepared by incubation with either medium or TZ. After 18 h, expression of CD40 and CD86 was assessed on CD11c+ cells. This experiment was repeated twice with essentially identical results.

Next, we determined whether TNF-α alone could activate bone marrow-derived DC. Accordingly, DC were incubated for 18 h with increasing amounts of exogenous TNF-α, then expression of CD40 and CD86 was evaluated by FACS analysis. As shown in Fig. 4⇓, high levels of costimulatory molecule up-regulation were seen only in the presence of 1 ng/ml or higher of TNF-α.

FIGURE 4.
FIGURE 4.

Direct DC activation by in vitro stimulation with exogenous TNF-α. Bone marrow-derived DC were stimulated for 18 h with the indicated concentrations of rTNF-α, then cells were stained with mAb to CD40 and CD86 (bold lines) and subjected to FACS analysis, gating on CD11chigh cells. Isotype controls are represented by thin lines. The numbers indicate percent of cells falling within the indicated gate.

T. gondii-stimulated neutrophils secrete factors that induce IL-12(p40) and TNF-α production by DC

We next asked whether supernatants from PMN-T. gondii cocultures triggered DC cytokine production. As shown in Fig. 5⇓A, supernatants from PMN-T. gondii cocultures induced DC IL-12 release to levels similar or greater than that seen with directly infected DC. Interestingly, while T. gondii alone elicited minimal DC TNF-α production, supernatants from PMN-T. gondii cocultures triggered robust production of this proinflammatory cytokine (Fig. 5⇓B). As shown in the figure, and as previously reported (26), PMN themselves produced IL-12 and TNF-α when stimulated with T. gondii. Nevertheless, levels produced by neutrophils are low relative to that induced by DC. This is of interest because high levels of exogenous TNF-α are required to fully activate DC (Fig. 4⇑). The results suggest that PMN-derived TNF-α acts in concert with other presently undefined factors to achieve full DC costimulatory molecule up-regulation.

FIGURE 5.
FIGURE 5.

T. gondii-stimulated neutrophils secrete factors that stimulate DC IL-12 and TNF-α production. Bone marrow-derived DC were cultured with TZ (0.5:1 ratio of parasites to cells), PMN-M, and PMN-TZ. Supernatants were collected for IL-12(p40) (A) and TNF-α (B) cytokine ELISA 18 h after culture initiation. M, medium; PMN-M, cell-free supernatants from PMN stimulated with medium; PMN-TZ, cell-free supernatants from PMN stimulated with T. gondii. nd, none detected. Data are representative of greater than five different experiments.

DC from neutrophil-depleted, infected mice are impaired in IL-12 and TNF-α production during in vivo T. gondii infection

The above results leave unresolved the question of whether PMN exert effects on DC during in vivo infection. To address this critical issue, PMN-negative mice, generated by anti-neutrophil mAb administration, were infected with T. gondii and splenic DC cytokine production was examined at the peak of acute infection. We have previously shown that PMN-depleted animals cannot survive acute infection (16). As shown in Fig. 6⇓A, FACS intracellular cytokine staining of an enriched CD11c+ DC population shows 50 and 54% of cells in the infected group expressing IL-12 and TNF-α, respectively. In striking contrast, neutropenic infected animals displayed a 2-fold decrease in both TNF-α and IL-12 expression (Fig. 6⇓A). Nevertheless, there was also the possibility that the anti-Gr-1 mAb used to deplete neutrophils was also removing Gr-1+ IL-12+ DC. Accordingly, we examined the Gr-1 phenotype of IL-12-positive cells in spleens from infected mice. As shown in Fig. 6⇓B, among total splenocytes virtually all IL-12-positive cells were Gr-1 negative. In addition, we enriched for CD11c+ DC and found that the small population that expressed Gr-1 was IL-12-negative (Fig. 6⇓B). To confirm these results, an enriched population of CD11c+ DC was isolated from 7-day T. gondii-infected mice and cultured in medium or with TZ. Correlating with the intracellular staining results, DC-enriched cells from infected animals produced both IL-12(p40) and TNF-α during ex vivo stimulation (Fig. 6⇓C). In contrast, DC isolated from neutropenic infected animals were severely impaired in ability to produce IL-12(p40) and most strikingly, TNF-α. As with bone marrow-derived DC responses to TZ (Fig. 5⇑), splenic DC from noninfected mice produced IL-12 but not TNF-α when cultured with the parasite. In control experiments, depletion of Gr-1-expressing cells in spleen populations from noninfected mice had no significant impact on parasite-stimulated IL-12 production (control, 6.0 ± 0.2 ng/ml vs 6.8 ± 0.3 ng/ml). These results confirm that splenic Gr-1+ CD11c+ DC do not contribute to IL-12 production during either in vivo or in vitro infection.

FIGURE 6.
FIGURE 6.

CD11c+ DC from neutrophil-negative infected mice are defective in production of IL-12(p40) and TNF-α. A, C57BL/6 mice were neutrophil-depleted by i.p. injection of RB6C6.8C5 mAb. After Ab administration, mice were i.p. infected with 100 ME49 cysts, and 7 days later, spleens were harvested and CD11c+ DC enriched using immunomagnetic beads conjugated to CD11c mAb. Flow cytometric analysis after gating on CD11c+ cells shows cells stained with PE-labeled anti-IL-12 or anti-TNF-α mAb. Numbers represent the percent of cells falling within the indicated gates. In B, splenocytes from day 7-infected mice were stained with PE-labeled anti-IL-12 and FITC-labeled anti Gr-1 mAb. Also shown in B, CD11c+ DC were enriched from the same population and assessed for IL-12 and Gr-1 expression. Percent of cells falling within the indicated gates is shown. In C, C57BL/6 mice were neutrophil-depleted by i.p. injection of NIMP-R14 and splenocytes prepared 7 days postinfection. ELISA was performed on 18 h supernatants from CD11c+-enriched cells cultured with medium or TZ (0.5:1 ratio of parasites to cells). CD11c+ DC were obtained from mice that were: NI, not infected; Inf, infected; PMN neg Inf, neutrophil-depleted and infected. Data are representative of three different experiments.

Discussion

Production of IL-12 by the innate immune system is essential in triggering a Th1 response and in surviving T. gondii infection (30, 31, 32, 33). Dendritic cells, macrophages, and neutrophils have each been suggested as a source of this cytokine during Toxoplasma infection, and the ability of multiple cell types to serve as an IL-12 source may, in part, account for the strong Th1 response elicited by infection (10, 12, 25, 26, 30, 34, 35). Here, we reveal that cytokine and chemokine cross-talk in the innate immune system, in particular between PMN and DC, further promotes the Th1-inducing properties of the latter cell type.

Neutrophils cocultured with live Toxoplasma produce soluble factors displaying several important effects on DC. In chemotaxis assays, supernatants from parasite-stimulated PMN cultures possessed strong chemotactic activity toward immature bone marrow-derived DC. CCL3 (MIP-1α), CCL4 (MIP-1β), CCL5 (RANTES), and, in particular CCL20 (MIP-3α) are chemotactic for immature DC (27, 28). Our results and those of others show that PMN produce chemokines including CCR5 ligands and CCL20 (25, 27, 36). Ab blocking studies revealed that DC chemotaxis induced by PMN factors was due to the combined activities of these chemokines.

DC also released high levels of IL-12 in response to stimulation with supernatants from T. gondii-PMN cocultures. In addition, direct infection of bone marrow and spleen-derived DC led to high amounts of IL-12 release, as did incubation with products released by extracellular tachyzoites (data not shown). Recent studies reveal a role for CCR5 and MyD88 in Toxoplasma-triggered DC IL-12 production (34, 37). This is due, at least in part, to Toxoplasma cyclophilin-18, a protein that mediates its effects by binding to CCR5 on DC (38). Here, we found that Ab blocking of CCL3, CCL4, and CCL5 failed to inhibit the IL-12 response, making it unlikely that CCR5-binding chemokines present in the PMN supernatants trigger DC IL-12 (data not shown). We cannot presently distinguish the extent to which IL-12-inducing activity in the parasite-stimulate PMN supernatants is attributable to Toxoplasma Ag itself, vs neutrophil factors released in response to the parasite.

Given the above results with IL-12, it is striking that TNF-α release could not be attributed to the activity of parasite products on DC. This is because direct DC infection, as well as incubation with secreted TZ factors (data not shown), elicited minimal TNF-α. We similarly found that splenic DC fail to produce TNF-α after parasite stimulation. Indeed, lack of TNF-α in parasite-infected cells reflects active suppression by the parasite in bone marrow-derived DC (our unpublished observations) and macrophages (39, 40). Nevertheless, we found that splenic DC released TNF-α when cells were isolated at day 7 of infection, and in vitro incubation with parasite Ag increased the amount of TNF-α released. The results suggest that CD11c+ DC are capable of responding directly to the parasite by releasing TNF-α when isolated from a proinflammatory environment.

Parasite-stimulated neutrophil supernatants were capable of strong CD40 up-regulation on DC, and this contrasted with the relatively weak increase in expression occurring during direct DC infection. Furthermore, DC activation, as measured by up-regulated CD40 expression, was abrogated in the presence of a neutralizing TNF-α Ab. In this regard, we have also found that T. gondii-stimulated human peripheral blood PMN induce TNF-α-dependent increases in costimulatory molecule expression in human monocyte-derived DC (our unpublished observations). Supernatants prepared from TNF-α knockout neutrophils fail to induce DC activation, implicating PMN as the source of this cytokine. Nevertheless, the amount of recombinant TNF-α required to induce similar levels of DC activation exceeded by 4-fold the amount released by parasite-stimulated PMN. This suggests that another factor, derived from either PMN or DC, may synergize with neutrophil-derived TNF-α to promote DC activation. The identity of such factor(s) is currently under investigation in our laboratory. Regardless, the ability of parasite-triggered PMN to induce DC CD40 up-regulation is likely to be important, because CD40L (CD154) is required for splenic DC activation during toxoplasmosis (11).

To address the physiological relevance of the data, mice were administered neutrophil-depleting mAb before infection. This treatment has previously been shown to result in defective Th1 responses during infection with Toxoplasma and other microbial pathogens (16, 18, 41). Our present studies show that CD11c+ splenic DC from infected neutrophil-depleted mice display severe defects in IL-12 and TNF-α production, suggesting that pathogen-triggered PMN influence in vivo DC activation. Although one of the depleting Ab used in this study (RB6C6.8C5) also recognizes a subset of plasmacytoid-like Gr-1 (Ly6G)+ DC in the spleen, it is, nevertheless, highly unlikely that removal of this subset accounts for the defective DC cytokine response. This is because it has been shown that IL-12 produced by T. gondii Ag-triggered splenic DC derives from CD11c+Gr-1 (Ly6G)− populations (42), and we also do not detect IL-12 expression among Gr-1+ splenic DC from normal mice in the infection model used here (Fig. 6⇑B). However, we cannot yet exclude the possibility that other non-PMN Gr-1+ cells play an in vivo role in instructing DC IL-12 production during Toxoplasma infection.

It is not yet known how neutrophils recognize TZ. Inasmuch as MyD88-negative PMN fail to produce IL-12 in response to the parasite, it is likely that Toll-like receptors are involved in molecular recognition (37). The recent identification of Toxoplasma cyclophilin-18 as a protein that induces DC IL-12 release implicates this molecule in neutrophil cytokine production (38). Indeed, we find that PMN express high levels of CCR5 (L. Del Rio and E.Y. Denkers, unpublished observations), the ligand through which T. gondii cyclophilin exerts its effects in DC. We are currently examining molecular recognition of parasite Ag by PMN.

Regardless of how neutrophils sense Toxoplasma, our data suggest a model in which PMN recruited to a site of infection are triggered by the parasite to release DC chemotactic molecules. The increase in the local DC population would favor direct interactions with the parasite, promoting Ag uptake and infection, and initiating IL-12 synthesis. Parasite-triggered soluble PMN factors would also induce DC TNF-α production. Additionally, DC recruited to the developing inflammatory focus would be activated by PMN products, in a process dependent upon TNF-α. The DC, primed by neutrophils and armed with Ag, could then traffic to tissues of the draining lymph node, where they would initiate T cell activation and Th1 subset selection.

Acknowledgments

We thank Dr. M. Hesse for critical comments on this manuscript and Dr. L. Del Rio for insightful discussion.

Footnotes

  • ↵1 This work was supported by National Institutes of Health Grants AI47888 (to E.Y.D.) and AI44322 (to T.J.C.).

  • ↵2 Address correspondence and reprint requests to Dr. Eric Y. Denkers, Department of Microbiology and Immunology, College of Veterinary Medicine, Cornell University, Ithaca, NY 14853-6401. E-mail address: eyd1{at}cornell.edu

  • 3 Abbreviations used in this paper: DC, dendritic cell; PMN, polymorphonuclear leukocyte; TZ, tachyzoite; CCL, CC chemokine ligand; MIP, macrophage inflammatory protein; PEC, peritoneal exudate cell.

  • Received June 6, 2003.
  • Accepted September 30, 2003.

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