Chemokines are critical for the recruitment of effector immune cells to sites of infection. Mice lacking the chemokine receptor CCR1 have defects in neutrophil trafficking and proliferation. In the present study, we tested the susceptibility of CCR1 knockout mice to infection with the obligate intracellular protozoan parasite Toxoplasma gondii. In comparison with parental wild-type mice, CCR1−/− mice exhibited dramatically increased mortality to T. gondii in association with an increased tissue parasite load. No differences were observed in Ag-specific T cell proliferation or in cytokine responses between mutant and wild-type mice. However, the influx of PMNs to the peripheral blood and to the liver were reduced in CCR1−/− mice during early infection. Our results suggest that CCR1-dependent migration of neutrophils to the blood and tissues may have a significant impact in controlling parasite replication.

T oxoplasma gondii induces a strong immune response in the infected host (1). Immune responses against the parasite can be divided into an innate acute response followed by an Ag specific cell-mediated immune response. During the acute phase, NK cells are an important element of host defense (2, 3), whereas development of a parasite specific T cell response is essential for long term protection (4). In a murine model, NK cells and T cells are critical for host defense against the parasite (4, 5). IFN-γ secreted by both of these cell types plays a pivotal role in acute infection and recrudescence in the chronically infected host (6, 7).

Recent studies have suggested that, in addition to T cells and NK cells, polymorphonuclear leukocytes (PMNs)3 may play a role in the generation of early resistance to T. gondii infection (8, 9) and an exacerbation of Toxoplasma infection has been reported in PMN-depleted mice (10, 11). Scharton-Kersten et al. (12) have demonstrated that mice deficient in inducible NO synthase succumbed to Toxoplasma infection after Ab depletion of PMNs. Antimicrobial effects of neutrophils are, in part, mediated by their ability to scavenge infected cells and secrete toxic products such as NO and reactive oxygen metabolites (13, 14, 15). These molecules have been shown to be involved in the intracellular killing of T. gondii (16, 17). Neutrophils also secrete chemokines, which are responsible for recruitment of other effector cell populations (9, 18, 19).

Chemokines are a large family of secreted proteins involved in regulating multiple steps in host defense mechanisms and during inflammatory responses (20). Chemokine classification is based on the arrangement of cysteine residues in their amino-terminal domains (21). CXC or α-chemokines mainly target neutrophils and T cells, whereas CC or β-chemokines target monocytes, eosinophils, and basophils with variable selectivity but usually do not affect neutrophils (22). Chemokines activate leukocytes by binding to G1-type G proteins (23). The 18 identified receptor subtypes can be divided into 2 major groups, CXCR and CCR, based on the 2 major classes of chemokines. One of the CCR receptors, CCR1, is expressed on neutrophils, monocytes, lymphocytes, and eosinophils and binds the leukocyte chemoattractant and hemopoiesis regulator macrophage-inflammatory protein (MIP-1α), as well as several other related chemokines (24, 25, 26). Previous studies have shown that mice deficient in the CCR1 gene had defects in the trafficking of neutrophils to the blood (27, 28). In the present study, we tested the ability of CCR1−/− mice to resolve acute T. gondii infection. CCR1−/− mice showed a higher susceptibility to Toxoplasma infection and had higher tissue parasite loads than did immunocompetent controls.

The generation of CCR1−/− mice has been described previously (27). The CCR1−/− mice used in this study were backcrossed for six generations onto the C57BL/6 background. Age- and sex-matched control C57BL/6 were obtained from The Jackson Laboratory (Bar Harbor, ME). Mice were challenged perorally (p.o.) with the 76K strain of T. gondii (kindly provided by Daniel Bout, Tours, France). This strain is maintained by continuous oral passage of cysts. Unless otherwise stated, each mouse was infected p.o. with 10–15 cysts.

Toxoplasma lysate Ag (TLA) was prepared from tachyzoites of the RH strain of T. gondii. Parasites were cultured in human fibroblasts and isolated from monolayers by forced extrusion through a 27-gauge needle. Host cell debris was removed using a Percoll gradient (1.04 g/ml). Purified parasites, essentially free of any fibroblast contamination, were washed twice and resuspended in PBS. The resulting preparation was pulse sonicated at 4°C (eight times for 10-s intervals at 18 kHz) before centrifugation at 10,000 × g for 15 min to remove insoluble material. The protein concentration of the Ag preparation was determined using a commercial assay kit (Bio-Rad Laboratories, Cambridge, MA) and stored aliquoted at −20°C.

After general anesthesia, spleens were removed and homogenized in a petri dish, and contaminating erythrocytes were lysed in RBC lysis buffer (Sigma, St. Louis, MO). After two to three washes in HBSS (Life Technologies, Gaithersburg, MD) supplemented with 3% FBS (HyClone Laboratories, Logan, UT). Cells were suspended in RPMI 1640 (Life Technologies) containing 10% FBS and cultured in 96-well flat-bottom plates in a 200-μl volume at a starting concentration of 2 × 105 cells/well. Cells were stimulated with either 5 μg/ml Con A or 15 μg/ml TLA. After culture for 72 h, [3H]thymidine (0.5 μCi/well; Amersham, Arlington Heights, IL) was added to the wells. Pulsed splenocytes were harvested onto glass filters using an automated multiple sample harvester and dried. Incorporation of radioactive thymidine was determined by liquid scintillation counting according to a standard protocol.

Spleen cells from infected mice were isolated as described above, washed, and then suspended in 3% BSA-PBS. Phenotypic analysis of splenocytes was conducted using direct immunofluorescent staining and FACS analysis (Becton Dickinson, San Jose, CA). Cells (1 × 106/ml) were incubated with FITC- and/or PE-labeled Ab (all from PharMingen, San Diego, CA) on ice for 45 min. After incubation, cells were washed several times in buffer, fixed in 1% methanol-free formaldehyde, and stored at 4°C in dark before FACS analysis.

Tissues (intestine, spleen, liver, lung, and brain) from T. gondii-infected animals were collected at day 7 postinfection (p.i.). DNA was extracted from tissues using the Qiamp tissue kit (Qiagen, Chatsworth, CA), and 400 ng of each sample were analyzed by quantitative PCR. Amplification of parasite DNA was performed using primers specific for a 35-fold repetitive sequence of the Toxoplasma B1 gene (5′-GGAACTGCATCCGTTCATGAG-3′ and 5′-TCTTTAAAGCTTCGTGGT C-3′), which is found in all known parasite strains (29). A 134-bp competitive internal standard containing the same primer template sequences as the 194-bp B1 PCR fragment was also synthesized (30). Amplification of this 194-bp segment of the B1 gene and the 134-bp segment of the internal standard was performed using in a 50-μl reaction mixture containing 1.25 U Amplitaq DNA polymerase; 1× buffer (Perkin-Elmer, Norwalk, CT); 0.2 mM concentrations each of dGTP, dATP, dTTP, and dCTP; and 0.4 μM concentrations of each B1 primer. For each reaction, a known amount of DNA from the tissues was amplified with varying amounts of the internal standard. Levels of parasite load were estimated by comparison to the internal controls. To determine the parasite load in infected tissues, PCR was performed under the same conditions using a known number of parasites. The level of internal control was calculated per parasite (30).

Tissues from infected CCR1−/− animals and parental control animals were fixed in 10% buffered formalin, paraffin processed, and used to prepare 5-μm histological sections. Sections were stained with hematoxylin and eosin and photographed on an Olympus Van Ox microscope with Kodak Elite 100 film. The resulting images were digitized with a Polaroid Sprint scanner and processed using Adobe Photoshop software.

Tissues from T. gondii-infected animals were collected at day 7 p.i. RNA from the tissue samples was collected using Trizol (Life Technologies) according to the manufacturer’s instructions. Reverse transcription was performed using Moloney murine leukemia virus reverse transcriptase (Life Technologies) and random hexamer primers. (Promega, Madison, WI). Levels of mRNA for IFN-γ and IL-10 were measured by quantitative PCR using the PQRS quantitative method (31). The tissues from uninfected mice were used to establish a baseline value of 1.0 against which the level of message for cytokine in the test mice was quantitated.

The splenocytes from infected wild-type and CCR1−/− mice were cultured in a 24-well plate in presence of 15 μg/ml TLA. After incubation for 72 h, culture supernatants were collected and stored at −70°C. These supernatants were analyzed for IFN-γ production by ELISA (Endogen, Woburn, MA).

Relative PMNS numbers in uninfected and infected, wild-type and homozygous CCR1 knockout mice (three or four animals per group) were determined by dual-color immunofluorescent staining. Whole mouse blood (100 μl) was incubated for 1 h at 4°C with FITC-labeled anti-CD45.2 (clone 104; PharMingen), a pan leukocyte marker, and PE-labeled anti-mouse GR-1 (RB6-8C5; PharMingen), a PMN-specific marker. Samples were then washed three times by centrifugation, and the erythrocytes were lysed using FACS lysing solution (Becton Dickinson) before fixation with 1% paraformaldehyde. Duplicate cytospin slides were then prepared for each sample and analyzed using a Bio-Rad MR1000 Confocal Scanning Laser Microscope system as described previously (32). At least 300 cells on each slide were counted. Relative PMN numbers were then calculated as a percentage of the total blood leukocytes.

To estimate the relative index of PMNs in the tissues of the infected animals, the livers and intestines from wild-type and CCR1−/− mice were collected at days 1, 2, and 3 p.i. Histological sections were stained with hematoxylin and eosin and examined by Olympus Van Ox microscope.

Statistical analysis of lymphoproliferation studies and relative PMN numbers were conducted by unpaired Student t tests (33).

To determine whether the deletion of the CCR1 gene altered susceptibility to T. gondii infection, CCR-1−/− and parental C57BL/6 mice were challenged p.o. with 15 cysts of T. gondii. As shown in Fig. 1, all of the knockout mice succumbed to infection by day 14 postchallenge. In contrast, almost 80% of the animals from the control group survived Toxoplasma infection. None of the survivors exhibited any signs of clinical sickness, and they continued to live until the termination of the experiment.

FIGURE 1.

Survival of CCR1−/− mice against T. gondii infection. Female CCR1−/− (n = 18) and parental wild type (WT, n = 18), 5–6 wk old, were infected with 15 cysts of the 76K strain of T. gondii p.o. The survival was monitored on a daily basis. There were six animals per group, and study was performed three times. Data are represented as a cumulative percent of all three experiments.

FIGURE 1.

Survival of CCR1−/− mice against T. gondii infection. Female CCR1−/− (n = 18) and parental wild type (WT, n = 18), 5–6 wk old, were infected with 15 cysts of the 76K strain of T. gondii p.o. The survival was monitored on a daily basis. There were six animals per group, and study was performed three times. Data are represented as a cumulative percent of all three experiments.

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Tissues from both CCR1−/− and parental mice were analyzed by quantitative PCR to determine whether levels of parasite multiplication are increased in knockout animals. The relative abundance of the B1 gene, a genetic marker for T. gondii, was determined at 12 days p.i. The majority of organs from knockout mice (spleen, liver, lungs, and brain) exhibited a 10- to 20-fold increase in parasite burden as compared with parental C57BL/6 mice (Fig. 2). A nearly 6-fold increase in the parasite load was noted in the intestines of CCR1−/− mice (Fig. 2).

FIGURE 2.

Number of parasites per μg tissue DNA in the organs of CCR1−/− and parental mice infected with T. gondii. Mice (n = 3/group) were infected p.o. with 15 cysts of T. gondii. At day 12 p.i., the organs from both wild-type and knockout mice were collected, and levels of parasite load in the tissues was determined by competitive DNA PCR. This experiment was performed twice with similar results.

FIGURE 2.

Number of parasites per μg tissue DNA in the organs of CCR1−/− and parental mice infected with T. gondii. Mice (n = 3/group) were infected p.o. with 15 cysts of T. gondii. At day 12 p.i., the organs from both wild-type and knockout mice were collected, and levels of parasite load in the tissues was determined by competitive DNA PCR. This experiment was performed twice with similar results.

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Sections of the liver of day 8 p.i. CCR1−/− mice (Fig. 3,A) showed preservation of overall tissue architecture, with multiple scattered foci of individual hepatocyte necrosis accompanied by a mixed lymphocyte infiltrate and the occasional granulocyte. Many small foci of intracellular T. gondii replication were seen throughout the liver (Fig. 3,A,inset, arrows). Parental control mice (Fig. 3,B) showed the expected small lymphocytic foci scattered throughout the parenchyma and the focal fatty metamorphosis typical of T. gondii-infected C57BL/6 mice (34). The small intestine of both the CCR1−/− mice and the parental controls (Fig. 3, C and D) showed scattered focal necrosis. This necrosis was more pronounced and more likely to be full-thickness in the parental controls as compared with the CCR1−/− mice. Thus, it appears that the inflammatory intestinal pathology reported in wild-type C57BL/6 mice (34, 35) is comparatively less severe in the CCR1−/−animals. The spleens of the CCR1−/− mice (Fig. 3,E) showed lymphocyte depletion of the primary follicles, with evidence of intracellular replication of T. gondii in macrophages and stromal cells (Fig. 3,E, inset, arrows). The parental controls (Fig. 3 F) showed preservation of the basic splenic architecture with scattered perifollicular large clear cells typical of the pattern seen in infected C57BL/6 mice.

FIGURE 3.

Photomicrographs of liver, ileum, and spleen from CCR-1−/− mice infected with T. gondii and similarly infected parental C57BL/6 controls at day 8 p.i. A, Infected liver of a CCR-1−/− mouse showing foci of hepatocyte necrosis and mixed inflammatory infiltrate. Inset, Individual hepatocytes harbor replicating T. gondii (arrows). B, Parental control mice show fatty metamorphosis of hepatocyte cytoplasm and small islands of lymphocytic infiltration. Replication of T. gondii is not evident. C, Ileum of infected CCR1−/− mouse with a focus of superficial epithelial necrosis with mixed inflammatory infiltrate. D, Ileum of parental control showing full-thickness necrosis and hemorrhage. CCR1−/− spleen showing lymphocyte depletion in primary follicles and evidence of intracellular replication of T. gondii within macrophages and stromal cells (inset, arrows). F, Spleen of infected parental mouse showing preservation of follicular architecture and scattered large clear reticuloendothelial cells. Size bars = 100 μm.

FIGURE 3.

Photomicrographs of liver, ileum, and spleen from CCR-1−/− mice infected with T. gondii and similarly infected parental C57BL/6 controls at day 8 p.i. A, Infected liver of a CCR-1−/− mouse showing foci of hepatocyte necrosis and mixed inflammatory infiltrate. Inset, Individual hepatocytes harbor replicating T. gondii (arrows). B, Parental control mice show fatty metamorphosis of hepatocyte cytoplasm and small islands of lymphocytic infiltration. Replication of T. gondii is not evident. C, Ileum of infected CCR1−/− mouse with a focus of superficial epithelial necrosis with mixed inflammatory infiltrate. D, Ileum of parental control showing full-thickness necrosis and hemorrhage. CCR1−/− spleen showing lymphocyte depletion in primary follicles and evidence of intracellular replication of T. gondii within macrophages and stromal cells (inset, arrows). F, Spleen of infected parental mouse showing preservation of follicular architecture and scattered large clear reticuloendothelial cells. Size bars = 100 μm.

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Cytokine analysis of the splenocytes from infected animals was performed at day 7 p.i. using quantitative PCR. Message for IFN-γ was increased in both the wild-type and the knockout mice in response to T. gondii infection. Almost equal levels of IFN-γ message were observed in the spleens of infected CCR1−/− and control wild-type animals at day 7 p.i. (Fig. 4,A). IL-10 levels were elevated in both knockout and wild-type infected mice as compared with uninfected mice. No noticeable differences in the production of IL-10 message were observed in the spleens of infected knockout vs the spleens of wild-type mice (Fig. 4 B). Both of these cytokines play an important immunoregulatory role in infectious diseases (36, 37, 38, 39).

FIGURE 4.

Cytokine mRNA expression after T. gondii infection in CCR1−/− and parental wild-type mice. Mice were infected p.o. with 15 cysts of T. gondii as mentioned earlier. At day 7 p.i., splenocytes from each infected and uninfected controls (three mice/group) were harvested and pooled. mRNA expression for IFN-γ and IL-10 was assayed by RT-PCR. The differences in the transcriptional level for both the genes are expressed relative to uninfected mice (assigned as 1). The cDNA concentration examined at each time point was standardized to hypoxanthine phosphoribosyltransferase mRNA level (not shown).

FIGURE 4.

Cytokine mRNA expression after T. gondii infection in CCR1−/− and parental wild-type mice. Mice were infected p.o. with 15 cysts of T. gondii as mentioned earlier. At day 7 p.i., splenocytes from each infected and uninfected controls (three mice/group) were harvested and pooled. mRNA expression for IFN-γ and IL-10 was assayed by RT-PCR. The differences in the transcriptional level for both the genes are expressed relative to uninfected mice (assigned as 1). The cDNA concentration examined at each time point was standardized to hypoxanthine phosphoribosyltransferase mRNA level (not shown).

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In addition to IFN-γ message, levels of IFN-γ protein produced by the splenocytes of infected animals in response to antigenic stimulation were determined by ELISA. As shown in Table I, minimal differences in the IFN-γ release were noted between wild-type and CCR1−/− mice.

Table I.

IFN-γ levels (picograms per milliliter) from T. gondii-infected micea

CCR1−/−Wild-Type
UnstimulatedStimulatedUnstimulatedStimulated
Uninfected ND 9.5 ± 3.4 ND 6.8 ± 2.3 
Infected 456 ± 73.5 1436 ± 251 512 ± 1208 1548 ± 246.4 
CCR1−/−Wild-Type
UnstimulatedStimulatedUnstimulatedStimulated
Uninfected ND 9.5 ± 3.4 ND 6.8 ± 2.3 
Infected 456 ± 73.5 1436 ± 251 512 ± 1208 1548 ± 246.4 
a

CCR1−/− and parental C57BL/6 mice were infected with the 76K strain of T. gondii p.o. At day 8 p.i., animals were sacrificed, and splenocytes isolated and pooled (3 mice/group). Spleen cells (1 × 106) were cultured in 24-well plates in the presence of 15 μg TLA per ml. After 72 h incubation, the cultures were collected, centrifuged, and stored at −70°C. The supernatants were assayed for IFN-γ by ELISA. ND, Not detected.

To determine whether the mortality observed in the CCR1−/− mice was due to their inability to elicit a normal T cell response, phenotypic analysis of splenocytes from infected animals was performed. As shown in Table II, similar increases in both the percentage and the absolute numbers of T cells were observed in the infected CCR1−/− and wild-type mice. No significant differences were seen within either the CD4+ or the CD8+ T cell populations of infected animals. As NK cells play an important role in acute T. gondii infection, the splenocytes were analyzed for the presence of NK cells. A nominal difference in the NK cell population was observed between the two strains of mice.

Table II.

Phenotypic analysis of CCR1−/− mice infected with T. gondiia

MouseCD3CD4CD8NK
% positiveAbs. No.% positiveAbs. No.% positiveAbs. No.% positiveAbs. No.
WT UI 33.5 ± 2.12 0.61 ± 0.03 19.50 ± 2.80 0.36 ± 0.05 11.25 ± 1.41 0.28 ± 0.02 4.40 ± 0.56 0.08 ± 0.01 
WT Inf 47.00 ± 4.24 1.57 ± 0.14 26.46 ± 1.41 0.88 ± 0.04 20.35 ± 2.56 0.68 ± 0.08 8.10 ± 1.47 0.27 ± 0.05 
CCR1−/− UI 29.5 ± 2.28 0.51 ± 0.04 18.55 ± 3.53 0.32 ± 0.06 9.55 ± 0.75 0.22 ± 0.01 5.57 ± 0.72 0.10 ± 0.01 
CCR1−/− Inf 45.50 ± 4.94 1.42 ± 0.16 27.55 ± 2.15 0.83 ± 0.11 18.45 ± 1.33 0.57 ± 0.04 7.36 ± 2.52 0.23 ± 0.08 
MouseCD3CD4CD8NK
% positiveAbs. No.% positiveAbs. No.% positiveAbs. No.% positiveAbs. No.
WT UI 33.5 ± 2.12 0.61 ± 0.03 19.50 ± 2.80 0.36 ± 0.05 11.25 ± 1.41 0.28 ± 0.02 4.40 ± 0.56 0.08 ± 0.01 
WT Inf 47.00 ± 4.24 1.57 ± 0.14 26.46 ± 1.41 0.88 ± 0.04 20.35 ± 2.56 0.68 ± 0.08 8.10 ± 1.47 0.27 ± 0.05 
CCR1−/− UI 29.5 ± 2.28 0.51 ± 0.04 18.55 ± 3.53 0.32 ± 0.06 9.55 ± 0.75 0.22 ± 0.01 5.57 ± 0.72 0.10 ± 0.01 
CCR1−/− Inf 45.50 ± 4.94 1.42 ± 0.16 27.55 ± 2.15 0.83 ± 0.11 18.45 ± 1.33 0.57 ± 0.04 7.36 ± 2.52 0.23 ± 0.08 
a

CCR1−/− and age-matched parental wild-type (WT) mice were infected with T. gondii as mentioned above. Splenocytes were isolated and pooled (three animals/group) at day 7 p.i. the cells were phenotyped for the expression of CD3, CD4, CD8, and NK receptors by direct immunofluorescence using FACS. Data are represented as mean ± SD of two similar experiments. Abs No., Absolute number of phenotypic cells × 108; UI, uninfected; Inf, infected.

To determine whether there were differences in lymphocyte activation between the two strains of mice, splenocytes were dual-stained for CD44 or CD62L. Increased expression of surface marker CD44 (CD44hi) is characteristic of activated T cells (40), whereas CD62L is expressed at low levels (CD62Llow) on the activated T cells (40). On CD4+ T cells, minor differences in the percentage of cells expressing these activation markers were observed: wild-type (82 ± 4 CD44high, 44 ± 8 CD62Llow) and CCR1−/− mice (78 ± 8 CD44high, 47 ± 10 CD62Llow). Similar (nominal) differences in the expression of activation markers on CD8+ T cells between wild-type (62 ± 12 CD44high, 55 ± 9 CD62Llow) and CCR1−/− mice (67 ± 14 CD44high, 49 + 8 CD44low) were noted.

We and others have demonstrated that acutely infected mice exhibit an immune down-regulation at 7 days p.i., which manifests as a decreased proliferative response to Con A stimulation (41, 42). It has been reported that this altered response is mediated by NO released by IFN-γ-activated macrophages (41, 42, 43). To further evaluate the immune response generated against T. gondii in CCR1−/− mice, lymphocyte proliferation assays were performed. Splenocytes were harvested from CCR1−/− mice at day 7 p.i., and proliferative responses to Con A and TLA were compared with those of age-matched wild-type controls. As shown in Fig. 5,A, splenocytes from both CCR1−/− and wild-type infected mice failed to respond to Con A stimulation. In contrast, spleen cells from uninfected control groups showed significant mitogenic response (p < 0.001). However, when the NO synthase antagonist, N-monomethyl-l-arginine (l-NMMA), was added to the splenocyte cultures, mitogenic responses of splenocytes from both CCR1−/− and wild-type mice were partially restored. Similarly, spleen cells from infected CCR1−/− mice failed to proliferate in response to TLA stimulation in the absence of l-NMMA (Fig. 5 B). Treatment with the NO antagonist significantly neutralized the suppression in the CCR1−/− splenocyte culture (p = 0.03). Similar observations were made with the spleen cell cultures from infected wild-type controls (p = 0.02).

FIGURE 5.

Ag-specific proliferation of spleen cells from T. gondii-infected animals. Pooled splenocytes (n = 3) from infected CCR1−/− and wild-type (WT) controls were collected at day 7 p.i. Spleen cells (2 × 105/well) were cultured in the presence of Con A or TLA. Some of the Con A or Ag stimulated well were treated with 0.5 mM l-NMMA. There were four replicate wells for each condition. After 72 h incubation, proliferation was measured by [3H]thymidine incorporation. Data are representative of two separate experiments.

FIGURE 5.

Ag-specific proliferation of spleen cells from T. gondii-infected animals. Pooled splenocytes (n = 3) from infected CCR1−/− and wild-type (WT) controls were collected at day 7 p.i. Spleen cells (2 × 105/well) were cultured in the presence of Con A or TLA. Some of the Con A or Ag stimulated well were treated with 0.5 mM l-NMMA. There were four replicate wells for each condition. After 72 h incubation, proliferation was measured by [3H]thymidine incorporation. Data are representative of two separate experiments.

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Relative numbers of peripheral blood PMNs were estimated in both uninfected and orally infected wild-type and homozygous CCR1 knockout mice. When expressed as a percentage of total leukocytes, PMNs were not significantly different in the peripheral blood of uninfected CCR1−/− mice vs uninfected wild-type mice (Fig. 6). At day 1 p.i., PMNs as a percentage of peripheral blood leukocytes dropped significantly in the infected CCR1−/− mice vs uninfected CCR1−/− mice (p < 0.05, unpaired t test), compared with the uninfected wild-type mice (p = 0.1, NS) and the day 1 p.i. wild-type mice (p < 0.05, unpaired t test). A similar drop in PMN numbers at this time point was not observed in the control wild-type mice. However, numbers in the wild-type mice were reduced at day 2 p.i. as compared with the day 1 p.i. levels (p < 0.05, unpaired t test). In contrast, the percentage of PMNs in the knockout mice had increased from the day 1 p.i. levels (not significant). At 3 day p.i., numbers in the wild-type mice remained at day 2 levels and were significantly lower than levels at day 1 p.i. (p < 0.05, unpaired t test). PMN numbers in the CCR1−/− mice were now significantly higher than the day 1 p.i. levels (p < 0.05, unpaired t test), having recovered to levels equivalent to those seen in uninfected mice.

FIGURE 6.

Comparison of PMN frequency in wild type (□) and homozygous CCR1 knockout mice (▪) in the presence and absence of Toxoplasma infection. Wild-type and homozygous knockout mice, three animals/group, were either mock-infected or infected p.o. with 15 cysts of T. gondii. Animals were sacrificed at days 1, 2, and 3 p.i., and peripheral blood PMNs were estimated as a percent of total leukocytes. Bars in each group represent the mean percentage (±SEM) of the total leukocytes that were PMNs. Significant differences (p < 0.05, unpaired t test) between treatment pairs are indicated by brackets. The experiment was performed three times with similar results.

FIGURE 6.

Comparison of PMN frequency in wild type (□) and homozygous CCR1 knockout mice (▪) in the presence and absence of Toxoplasma infection. Wild-type and homozygous knockout mice, three animals/group, were either mock-infected or infected p.o. with 15 cysts of T. gondii. Animals were sacrificed at days 1, 2, and 3 p.i., and peripheral blood PMNs were estimated as a percent of total leukocytes. Bars in each group represent the mean percentage (±SEM) of the total leukocytes that were PMNs. Significant differences (p < 0.05, unpaired t test) between treatment pairs are indicated by brackets. The experiment was performed three times with similar results.

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To evaluate the trafficking of PMNs to the tissues of the infected animals, hematoxylin and eosin-stained sections of liver and intestine were examined, and relative PMNs indices determined. A slightly lower PMN index was observed in the liver and intestine of CCR1−/− mice at day 1 p.i. This drop was more pronounced in both tissues at day 2 p.i. (Table III) and returned to the levels of wild-type mice by day 3 p.i. The reduced tissue load is probably due to a significant reduction in the levels of circulating PMNs observed in the blood at day 1 p.i. in the CCR1−/− mice. However, by day 3 p.i. both relative peripheral blood PMN numbers and tissue PMN load had recovered, indicating that redundant mechanisms of PMN maturation and mobilization eventually lead to an increase in PMN production and mobilization in the CCR1−/− mice.

Table III.

Relative PMN index in the tissues of T. gondii-infected micea

Days p.i.MouseWild-Type LiverCCR1−/− LiverWild-Type IntestineCCR1−/− Intestine
1+ 1+ 1+ 1+ 
 1+ 1+ 1+ 1+ 
 1+ 1+ 1+ 1+ 
      
3+ 2+ 4+ 3+ 
 3+ 3+ 4+ 3+ 
 4+ 2+ 3+ 4+ 
      
4+ 1+ 4+ 1+ 
 3+ 2+ 4+ 2+ 
 3+ 1+ 4+ 2+ 
      
2+ 3+ 3+ 3+ 
 2+ 3+ 2+ 3+ 
 3+ 2+ 3+ 3+ 
Days p.i.MouseWild-Type LiverCCR1−/− LiverWild-Type IntestineCCR1−/− Intestine
1+ 1+ 1+ 1+ 
 1+ 1+ 1+ 1+ 
 1+ 1+ 1+ 1+ 
      
3+ 2+ 4+ 3+ 
 3+ 3+ 4+ 3+ 
 4+ 2+ 3+ 4+ 
      
4+ 1+ 4+ 1+ 
 3+ 2+ 4+ 2+ 
 3+ 1+ 4+ 2+ 
      
2+ 3+ 3+ 3+ 
 2+ 3+ 2+ 3+ 
 3+ 2+ 3+ 3+ 

a Female CCR1−/− mice, 5 to 6 wk old, and age-matched parental C57BL/6 mice were infected p.o. with 15 cysts of the 76K strain of T. gondii. At days 1, 2, and 3, p.i., the animals were sacrificed; the livers and intestines were removed, and fixed in 10% buffered formalin. The histological sections were cut and examined as described in Materials and Methods. For each tissue, multiple sections were examined and table illustrates the findings typical for the organ. The experiment was performed three times with similar results. The PMN index values were defined as follows: 1+, no, rare single PMN within inflammatory nodules; 2+, few, or less than one-third of the cells in the inflammatory nodules are PMNs; 3+, moderate, or one-time to two-thirds of the cells in the nodule are PMNs; 4+, two-thirds or more of the cells are PMNs.

The results presented in these studies demonstrate a nonredundant role for CCR1 during acute T. gondii infection. Mice deficient in CCR1 succumbed to low dose Toxoplasma challenge. Mortality in knockout mice was related to a marked increase in the replication of parasites in the tissues of these animals. The increased susceptibility of CCR1−/− mice could not be attributed to an inadequate cellular immune response generated against the parasite. Cell-mediated immunity plays a critical role in protection against T. gondii infection (44). Both NK cells and T cells have an important function during murine toxoplasmosis (4, 45). Based on phenotypic analysis, the raised NK and T cell numbers in infected CCR1−/− mice were comparable with those in the parental wild-type animals. No major differences in the cytokine patterns between the wild-type and parental knockout mice in response to T. gondii infection was observed. Apparently, an absence of the CCR1 gene does not have an effect on the induction of cellular immune responses against the parasite. However, despite evoking an apparently normal adaptive immune response against T. gondii, CCR1−/− mice had decreased intestinal necrosis as compared with parental wild-type animals. It has been reported that this parasite-driven intestine pathology is primarily mediated by IFN-γ-producing CD4+ T cells (35). Because CCR1−/− mice showed a normal CD4+ T cell immune response, it is possible that other cell types such as neutrophils may play a secondary role in the hyperimmune response in the intestine during T. gondii infection. In the present study, a reduced PMN index in the intestines of CCR1−/− mice in comparison with parental wild-type mice was observed at day 2 p.i. The role of neutrophils in the gut inflammation, although well described during other infectious diseases (46, 47), needs to be evaluated in the T. gondii model.

Although CCR1 is expressed on wide variety of lymphoid and myeloid cells, the overt phenotypic difference in the knockout mice is one of impaired myeloid progenitor cell function. Normal T cell responses in infected CCR1 −/− mice suggest that CCR1 has a redundant function in these cells. However, recent studies with a cardiac allograft model have reported that mice lacking CCR1 receptor showed significant prolongation of allograft survival (48). The lack of allograft rejection by CCR1−/− mice was attributed to a defect in the influx of IFN-γ-producing CD4+ T cells in these animals. Contrary to these findings, lung lymph nodes from CCR1−/− mice injected with Schistosoma eggs showed increased IFN-γ production (27). These reports raise the possibility that the role of chemokines may vary according to the disease model. Earlier studies have shown that neutrophils from CCR1−/− mice are nonresponsive to MIP-1α suggesting that CCR1 may be the dominant MIP-1α receptor subtype in these cells (27). CCR1 may have evolved to play a more important role in mouse neutrophils to compensate for the lack of other chemokine receptors, such as CXCR1 which is present on human neutrophils. Based on our current observations with T. gondii and earlier findings with MIP1-α-treated mice it can be postulated that CCR1 is important for the mobilization of bone marrow PMN pool.

Recent reports have demonstrated that neutrophils have the ability to produce both IL-12 and IFN-γ after stimulation with microbial Ags (11). IL-12 is an important cytokine that leads to type 1 cytokine (IFN-γ) synthesis in the infected host (49, 50). IFN-γ is known to be critical for survival against Toxoplasma infection (5, 6). Our results showed no differences in the levels of IFN-γ message in the spleens of the T. gondii-infected parental and CCR1−/− mice. These findings are in agreement with those of Sayles et al. (10), who demonstrated that depletion of neutrophils had no effect on IFN-γ production of T. gondii-infected animals. However, Sayles et al. reported reduced numbers of CD4+ and CD8+ T cells in response to T. gondii infection in neutrophil-depleted mice. The differences between our findings and those of Sayles et al. can be attributed to the fact that unlike neutrophil depletion, lack of CCR1 does not result in the complete blockade of neutrophil migration in these animals. This is also further proved by the fact that at later stages of infection (day 3 p.i.), no apparent differences in the neutrophil count between the tissues of knockout and wild-type mice were observed. Moreover, it is likely that there are other, redundant, mechanisms by which T cells can be activated in these mice.

Studies over the last few years have emphasized an important role for neutrophils as mediators of effective immune responses against microbial infections (11, 51). Neutrophils are often the first cell type recruited to an area of infection or inflammation. Neutrophil-depleted mice succumb to a mildly virulent strain of Candida albicans (51). Neutrophils have also been reported to contribute to early resistance against T. gondii infection (52). Depletion of PMNs with antigranulocyte Ab impairs the ability of mice to survive acute infection with a low virulence strain of T. gondii (12). In the present study, CCR1−/− mice, known to have disordered trafficking and mobilization of PMNs (28), exhibited increased susceptibility to T. gondii infection. However, based on our findings, it appears that CCR1−/− mice do not have any major problem in the trafficking of PMNs from the blood to the tissues. The relative PMN indices in the tissues of knockout and parental wild-type mice are more or less similar at day 1 p.i. Nevertheless, there is a very obvious decrease in the percentage of PMNs in the blood at this time point. This can be explained by an inability of the knockout mice to replenish the peripheral blood PMNs from the bone marrow pool, as circulating PMNs are recruited into the infected tissues. The decreased PMN levels in the blood of CCR1−/− mice results in a reduced PMN index in the tissues at day 2 p.i. However, the levels of PMNs in the peripheral blood and tissues are restored, respectively, at days 2 and 3 p.i. Thus, it seems that in the absence of CCR1, mobilization of PMNs to the peripheral blood during T. gondii infection can occur by alternative mechanisms. Recent studies have demonstrated that G-CSF plays an important role in granulopoiesis during infections (53). The role of G-CSF in the mobilization of PMNs during T. gondii infection is currently being investigated in our laboratory. A decrease in the percentage of PMNs was also observed in the peripheral blood of the wild-type C57BL/6 mice. However, the fall in PMN levels took place at day 2 p.i. as compared with day 1 p.i. in the knockout mice. Moreover, it appears that circulating PMNs levels in the wild-type mice are rapidly restored because the PMN index in the tissues remains unchanged. An interesting feature of these findings is that a very short delay in the mobilization of PMNs during T. gondii infection can have a pronounced effect on the outcome of infection. Consistent with our data, earlier studies have demonstrated that CCR1−/− mice had accelerated mortality when challenged with Aspergillus fumigatus; a fungal infection that is principally controlled by neutrophils (27).

Based on reports from other laboratories and our current observations, the hypothesis we propose is as follows. Natural immunity against T. gondii is dependent on the induction of strong parasite-specific immunity in the host (1). However, before adaptive immunity is established, the rapid multiplication of T. gondii tachyzoites in the infected host needs to be contained. Because neutrophils migrate very early during infection, they may be necessary to restrict the replication of the parasites. Earlier findings by Gao et al. (27) suggest that CCR1 may be essential for influx of neutrophils to the infected tissues. The absence of CCR1 probably results in defective neutrophil migration from bone marrow to the blood. This could lead to uncontrolled parasitic replication in T. gondii-infected mice, which ultimately overwhelms the capacity of the adaptive immune system to contain the infection.

1

This work was supported by National Institutes of Health Grant AI33325.

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