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Journal of Virology, August 2005, p. 9831-9841, Vol. 79, No. 15
0022-538X/05/$08.00+0 doi:10.1128/JVI.79.15.9831-9841.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.
Institute of Medical Microbiology and Immunology, University of Aarhus, Aarhus, Denmark
Received 19 November 2004/ Accepted 5 April 2005
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ABSTRACT |
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INTRODUCTION |
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/ß) (23). At later time points during infection, the adaptive immune system takes over, and CD8+ T cells specifically kill cells displaying the appropriate viral peptides in the context of MHC I (35). Maturation of antigen-specific T cells is initiated by antigen-loaded dendritic cells (DCs) and other antigen-presenting cells that migrate to secondary lymphoid organs and present antigens to T cells in the T-cell zones. Cells that receive signals through the T-cell receptor, as well as through costimulatory pathways, rapidly start to expand. T-cell proliferation is controlled both through cell-cell interactions and cytokine secretion. Once the T cells become activated, receptors are expressed on the cell surface, which allows the cells to migrate to infected areas of the body to kill virus-infected cells.
Regulation of leukocyte migration to lymphoid organs and inflamed areas is a complex process in which chemokines play pivotal roles (18). Chemokines constitute a large family of secreted peptides, which are produced by tissue cells or leukocytes in both constitutive and inducible manners (19). Chemokines are categorized into the subfamilies CXC (), CC (ß), C (
) and CX3C (
), according to the positions of conserved amino-terminal cysteines (16, 19). Chemokine receptors are seven-transmembrane-spanning, G-protein-coupled receptors. They are classified based on the class of chemokines they bind, e.g., CCRs bind CC chemokines (CCLs), and CXCRs bind CXC chemokines (CXCLs). Among chemokines, there is a great degree of redundancy; i.e., many receptors bind several chemokines, and many chemokines use more than one receptor. Chemokine receptors are expressed primarily on leukocytes but also on other cells, such as endothelial cells (16).
With regard to the role of chemokines in defense and disease during viral infections, some information is available, mainly from studies in murine models (20). For instance, defense against intracerebral infection by mouse hepatitis virus (MHV) is impaired if either of the two CXCR3 ligands CXCL9 or CXCL10 is inactivated, which was suggested to be due to important roles of these chemokines in T-cell recruitment to the central nervous system (8, 15). Mice lacking CCR1 or the ligand CCL3 display impaired ability to clear infection by a paramyxovirus in the lower respiratory tract (7), which was associated with higher mortality and reduced inflammatory response (7). Also, clearance of systemic infection with the herpesvirus cytomegalovirus (CMV) was delayed in CCL3–/– mice, which correlated with decreased infiltration of NK cells to the liver (27). For some other virus infections, CCR1 or CCL3 do not contribute to host defense but are involved in the pathogenesis of the infections (3, 6, 32).
CCR5 is coreceptor for human immunodeficiency virus (HIV), and this molecule is thus a potential target for anti-HIV drugs (33). There is therefore a need for evaluation of the role of CCR5 in the immune response against infections. Although most work done in animal models has concentrated on adult mice (1, 5, 10, 14, 22, 36), little information is available on the function of CCR5 during infection in young mice, which have a less developed cellular immune response and hence are more prone to infections in general (21, 25). Host defense against the viruses lymphocytic choriomeningitis virus, influenza A virus or MHV, or the intracellular bacterium Listeria monocytogenes has been found not to be hampered in adult CCR5-deficient mice (5, 10, 22, 36), whereas lack of this receptor did impair defense against the encapsulated yeast Cryptococcus neoformans and the parasite Toxoplasma gondii (14, 24). In humans, natural CCR5 mutations are well described (13). By far the one most studied is the CCR5 
32 frameshift mutation, which is relatively abundant (17). Individuals homozygous for the CCR5 32 mutation are fully immunocompetent with respect to defense against many infections but have been shown to display enhanced susceptibility to acute and chronic hepatitis C virus infection (34), thus suggesting a role for this receptor in both innate and adaptive immunity against some virus infections.
We have examined here the role of CCR1 and CCR5 in defense against a systemic herpes simplex virus type 2 (HSV-2) infection in 4- versus 8-week-old mice and found that CCR5 modulates both the innate and adaptive immune system in young animals and hence is essential for optimal antiviral defense early in life.
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MATERIALS AND METHODS |
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Mice and virus. The mice used in the present study were 4- and 8-week-old female C57BL/6, CCR1–/– and CCR5–/– mice. The animals were bred at Taconic M&B (Ry, Denmark). The CCR1–/– and CCR5–/– mice were generated as described elsewhere (9, 14). The virus used was the MS strain of HSV-2. The mice were infected intraperitoneally (i.p.) on day 0 with 106 PFU of virus suspended in 100 µl of phosphate-buffered saline (PBS). The animals were sacrificed on the indicated days postinfection (p.i.), and peritoneal cells (PCs) were harvested by lavage of the peritoneal cavity with cold PBS (pH 7.4) supplemented with 5% FCS and 0.4% heparin. Samples of spleen, liver, and brain were kept in RPMI 1640 or MEM on ice for further treatment or frozen at the time of harvest.
Virus plaque assay. Samples of liver, spleen, and brain were weighed, thawed, and homogenized three times for 5 s each time in MEM supplemented with 5% FCS just before use in the plaque assay. After homogenization the organ suspensions were pelleted by centrifugation at 1,620 x g for 30 min, and the supernatants were used for plaque assay as well as ELISAs (see below). Plaque assays were done on Vero cells seeded in MEM supplemented with 5% FCS at a density of 1.2 x 106 in 5-cm diameter plates and left overnight to settle. The cells were infected by incubation for 1 h at 37°C with 100 µl of serial dilutions of the organ suspension and 400 µl of medium, during which the tissue culture plates were rocked every 15 min to ensure even distribution of the virus. Subsequently, the organ suspensions were removed, and 8 ml of MEM was added to the plates. The medium was supplemented with 2.5 to 5% FCS, depending on how confluent the cells were at the time of incubation and with 0.2% human immunoglobulin. The cells were incubated at 37°C for 2 days and stained with 0.03% methylene blue, and the plaques were counted.
ELISA.
Murine CCL3, CCL5, interleukin-12 (IL-12) p40, and IFN- were detected by ELISA. Maxisorp plates were coated overnight at room temperature with 100 µl of antibody (anti-CCL3, 4 µg/ml [R&D Systems]; anti-CCL5, 2 µg/ml [R&D Systems]; or anti-IL-12, 8 µg/ml [BD Pharmingen]) in coating buffer (Na2CO3, 15 mM; NaHCO3, 35 mM; NaN3, 0.02% [pH 9.6]). After blocking for at least 1 h at room temperature with 300 µl of 1% BSA in blocking buffer (PBS with 5% sucrose 0.05% NaN3 [pH 7.4]), successive culture supernatants or recombinant murine CCL3, CCL5 (both R&D Systems), or IL-12 p40 (BD Pharmingen) were added to the wells (100 µl each) and incubated overnight at 4°C. Subsequently, wells were incubated at room temperature for 2 h with 100 µl biotinylated, anti-murine detection antibody (CCL3, 50 ng/ml [R&D Systems]; CCL5, 100 ng/ml [R&D Systems]; IL-12, 1 µg/ml [BD Pharmingen]) in a 0.1% suspension of BSA in TBS (Trizma, 20 mM; NaCl, 150 mM; 0.05% Tween 20 [pH 7.3]). Streptavidin-horseradish peroxidase (R&D Systems) diluted in 1:200 in Tris-buffered saline with 0.1% BSA was added, followed by incubation for 20 min. As a substrate 100 µl of the TMB substrate was added, and plates were incubated in the dark for an appropriate amount of time. The color reaction was stopped with 50 µl of 5% H2SO4, and the absorbance was measured at 450 nm with 570 nm as reference. Between each step the plates were washed three times with PBS-0.05% Tween 20 (pH 7.4). For detection of IFN- we used the monoclonal antibody Duoset (R&D Systems), and the protocol recommended by the manufacturer was followed. The detection limits of the ELISA were as follows: 31.25 pg/ml (CCL3) and 15.63 pg/ml (CCL5, IL-12 p40, and IFN-).
IFN-/ß bioassay.
IFN-/ß bioactivity was measured by using a L929-cell-based bioassay. L929 cells (2 x 104 cells/well in 100 µl) in MEM with 5% FCS were incubated overnight at 37°C in successive twofold dilutions of samples or murine IFN-/ß as a standard. Subsequently, vesicular stomatitis virus (VSV/V10) was added to the wells, and the cells were incubated for 2 to 3 days. The dilution mediating 50% protection was defined as 1 U of IFN-/ß/ml.
RT-PCR. Total RNA was extracted from PCs and organ homogenates with TRIzol according to the recommendations of the manufacturer. Briefly, cells were lysed in TRIzol, and chloroform was added, followed by phase separation by centrifugation. RNA was precipitated with isopropanol and pelleted by centrifugation. Pellets were washed with 70% ethanol and redissolved in RNase-free water. Then, 2 to 4 µg of RNA was subjected to reverse transcription (RT) with oligo(dT) as primer and Expand Reverse Transcriptase (both from Roche). The cDNA was amplified by PCR with the following primers: CCL3, 5'-GAA GAG TCC CTC GAT GTG GCT A-3' (sense) and 5'-CCC TTT TCT GTT CTG CTG ACA AG-3' (antisense); CCL5, 5'-ATA TGG CTC GGA CAC CAC TC-3' (sense) and 5'-GAT GCC GAT TTT CCC AGG AC-3' (antisense); and ß-actin, 5'-CCC ACT CCT AAG AGG AGG ATG-3' (sense) and 5'-AGG GAG ACC AAA GCC TTC AT-3' (antisense). The products spanned 561 bp (CCL3), 330 bp (CCL5), and 215 bp (ß-actin).
Isolation of splenocytes (SCs). Spleens were harvested and kept in 1 ml of RPMI 1640 supplemented with 10% FCS. Spleens were homogenized manually five times with a glass homogenizer in 1 ml of RPMI supplemented with FCS. The suspensions were filtered through gauze, which was washed with 10 ml of medium. The suspensions were pelleted by centrifugation at 180 x g for 10 min. The erythrocytes were lysed in NaCl 0.2% for 30 s, followed by the addition of an equal volume of NaCl 1.6% supplemented with 10% FCS. The leukocytes were pelleted by centrifugation at 115 x g for 12 min and resuspended in RPMI 1640 supplemented with 10% FCS.
Isolation of IHLs. Livers were isolated and washed repeatedly in RPMI 1640 supplemented with 5% FCS, followed by manual homogenization with a glass homogenizer. The homogenate was incubated in RPMI 1640 with 0.02% (wt/vol) collagenase IV and 0.002% (wt/vol) DNase I at 37°C for 40 min. Cells were precipitated by centrifugation at 720 x g for 10 min, resuspended in RPMI 1640, underlaid with 26% (wt/vol) metrazamide in PBS, and centrifuged at 1,500 x g for 20 min. Intrahepatic leukocytes (IHLs) were isolated in the interface, and erythrocytes were lysed in 0.2% NaCl for 30 s, followed by the addition of an equal volume of 1.6% NaCl. The leukocytes were pelleted by centrifugation for 10 min at 180 x g.
Purification of T cells. To isolate T cells from the spleen, we used the Pan T Cell Isolation kit (Miltenyi), following the instructions of the manufacturer, and separated the cells with an AutoMACS cell sorter. Cell purification with this kit yields untouched T cells depleted of B cells, NK cells, DCs, macrophages, granulocytes, and erythroid cells.
NK cell cytotoxicity assay. YAC-1 cells were loaded with BATDA [bis(acetoxymethyl)2,2':6',2"-terpyridine-6,6"-dicarboxylate] by incubating the cells for 20 min at 37°C in RPMI supplemented with 10 µM BATDA. The cells were washed four times and seeded in 96-well plates with 5 x 103 cells per well. PCs were added to the target cells in the indicated ratios to reach a final volume of 200 µl per well, and effector and target cells were brought into contact by centrifugation for 5 min at 50 x g. The cells were incubated for 2 h at 37°C, after which the cells were resuspended and centrifuged to allow even distribution of the released BATDA. From each well 25 µl was mixed with 200 µl of a 50 µM Eu3+ solution. Fluorescence was measured, and the percent cell lysis was calculated as follows: 100 x (experimental release – spontaneous release)/(maximum release – spontaneous release).
Measurement of ex vivo production of IFN- by splenic T cells.
To assay production of IFN- by T cells, spleens were harvested on day 6 p.i and kept in 1 ml of cold RPMI 1640 supplemented with 5% FCS. Splenic T cells were isolated as described above and set in culture in 24-well culture plates (2 x 106 cells/well in 1 ml) with irradiated syngeneic SCs as antigen-presenting cells. The cells were either left untreated or received 3 x 107 PFU of heat-inactivated HSV-2. Supernatants were collected after 72 h of culturing and IFN- was measured by ELISA as described above.
Measurement of SC proliferation. To assay proliferation of SCs, spleens were harvested on day 5 p.i., and SCs were isolated as described above. The cells were set in culture in 96-well plates in a CO2 humidified atmosphere. Proliferation was measured by using the EZ4U kit (Biomedica) according to the instructions of the manufacturer. Briefly, the substrate was dissolved and 20 µl was added per well to reach a final volume of 220 µl/well. After 4 h, absorbance was measured at 450 nm, with 620 nm as a reference.
Flow cytometry. PCs were harvested, counted, and resuspended in RPMI 1640 supplemented with 10% FCS to a concentration of 107 cells/ml, The cells were incubated in a 96-well plate on ice in the dark for 40 min with FITC-conjugated anti-mouse monoclonal anti-NK1.1 antibody at a concentration of 16.22 µg/ml. SCs and IHLs were isolated as described above. The leukocytes were incubated in a 96-well plate on ice in the dark for 40 min with FITC-conjugated anti-mouse monoclonal anti-CD3a antibody (16.22 µg/ml), PE-conjugated rat anti-mouse monoclonal anti-CD8a and CD4 antibodies (6.5 µg/ml), or combinations of the FITC- and PE-conjugated antibodies at the concentrations mentioned above. We used FITC-conjugated monoclonal mouse immunoglobulin G1 (16.22 µg/ml) and PE-conjugated monoclonal mouse immunoglobulin G2a (6.5 µg/ml) as isotype control antibodies.
The labeled cells were fixed by using 1% paraformaldehyde diluted in PBS and kept at 4°C until they were analyzed. Acquisition and analysis were performed with a flow cytometer (Coulter FS500). The data were stored in list mode files. A total of 20,000 cells were analyzed in each experiment by using a single laser system with a wavelength of 488 nm. Compensation was determined before the acquisition of data.
Adoptive transfer. Mouse donors (C57BL/6 and CCR5–/– mice) were infected i.p. with 106 PFU of HSV-2 prior to isolation of donor cell populations. For isolation of NK cell activity, PCs were harvested 24 h p.i. and cultured for 2 h in RPMI supplemented with 5% FCS before isolation of nonconfluent cells, which contained >95% of the cytotoxic activity. The cells (4 x 106 per mice) were injected i.p. into CCR5–/– mice infected with 106 PFU of HSV-2 2 h previously. The nonadherent cells contained between 10 and 400 virus particles per 4 x 106 cells as determined by plaque assay, with no differences between wild-type (WT) and CCR5–/– mice. For isolation of SCs, mice were sacrificed 6 days p.i., and cells were isolated as described above. The SCs were resuspended in Hanks balanced salt solution, and the desired number of viable cells were inoculated slowly intravenously into 4-week-old female C57BL/6 recipient mouse, which had been infected 6 h previously i.p. with 106 PFU of HSV-2 suspended in 100 µl of PBS. The SCs contained between 0 and 10 virus particles per 5 x 107 cells as determined by plaque assay, with no differences between WT and CCR5–/– mice. At 2 days after infection, livers were harvested and the viral load was determined by plaque assay.
Statistical analysis. The data are presented as means ± the standard error of the mean (SEM). The statistical significance was estimated with the Student t test for unpaired observations or Wilcoxon rank sum test. P values of <0.05 were considered significant.
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RESULTS |
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In the 4-week-old mice a generalized infection developed (Fig. 1A to C and G to I). In the spleen, virus was detected from day 1 p.i., and virus titers remained high through day 4, after which the amount of virus declined (Fig. 1B and H). No differences between viral load in the spleen of C57BL/6 and any of the knockout mouse strains were found. In the brain, virus was detected on day 6 in all mice strains (Fig. 1C and I). However, significantly higher virus titers were observed in CCR5-deficient animals on day 6 compared to WT C57BL/6 mice (Fig. 1I) (P = 0.029). In the livers of 4-week-old C57BL/6 mice, the virus was detectable from day 1 and peak virus titers were reached on day 4, followed by a pronounced decline in virus titer between days 4 and 6 (Fig. 1A and G). No significant differences between WT and CCR1–/– mice were observed (Fig. 1A). For CCR5-deficient mice, however, significantly higher virus titers were found on day 4 (P = 0.014) and day 6 (P = 0.007) compared to WT littermates (Fig. 1G). The strong decline between day 4 and day 6 seen in the WT and CCR1 mice was not found in the CCR5 mice, in which the virus titer remained high throughout the course of infection.
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Expression of CCL3 and CCL5 during HSV-2 infection. To investigate the expression of ligands for CCR5 in vivo, 4-week-old C57BL/6 were infected i.p. with 106 PFU of HSV-2, and livers, spleens, and PCs were harvested on days 1, 4, and 6. RNA and soluble fractions of organ homogenates were prepared and used for RT-PCR and ELISA, respectively. PCs were either lysed for RNA purification immediately after the mice were sacrificed or cultured for 24 h for measurement of ex vivo chemokine production. As shown in Fig. 2A to C, HSV-2 infection triggered expression of CCL3 and CCL5 in the peritoneal cavity, with clear induction observed on day 1 p.i. The levels remained elevated until day 6 p.i. for CCL5 but not for CCL3, whose mRNA levels decreased significantly between days 1 and 4 p.i. At the protein level, a clear induction of both CCL3 and CCL5 was observed on day 1 p.i., after which a slight decrease was observed on days 4 and 6. In the liver, we observed elevated levels of both CCL3 and CCL5 proteins on day 1, which subsequently decreased modestly (Fig. 2D to E). The mRNA for CCL3 was found to be induced on day 1 and to remain high through the 6 days of the experiment (Fig. 2F), whereas for CCL5 the infection did not seem to affect mRNA levels. In the spleen, a clear induction of CCL3 protein and mRNA was observed on day 1 p.i., which decreased to basal levels on day 4 (Fig. 2G to I). CCL5 mRNA and protein was constitutively present and appeared not to be affected by the HSV-2 infection. Thus, CCL3 and CCL5, two ligands for CCR1 and CCR5, are induced during a generalized HSV-2 infection with distinct expression patterns in different organs.
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IL-12 and IFN-/ß are two well-described activators of NK cell activity (2, 31). Therefore, reduced production of any of these cytokines in response to infection could potentially explain the impaired cytotoxic activity. To test this, PCs were harvested from C57BL/6 and CCR5–/– mice and infected with HSV-2 for 24 h, and supernatants were examined for IL-12 and IFN-/ß. As seen in Fig. 4, HSV-2 infection triggered expression of both IL-12 and IFN-/ß, and no differences were observed between cells from C57BL/6 and CCR5–/– mice.
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Immune responses in the spleen.
Since CCR5-deficient mice displayed a strikingly impaired ability to control infection between days 4 and 6 p.i. and also did not develop splenomegaly in response to infection, we wanted to explore the nature of the adaptive immune response to the infection in these mice. First, we looked at IFN- production by splenic T cells in response to antigen restimulation. Spleens were harvested from mice infected for 6 days or from control mice, and T cells were isolated and cultured in vitro with irradiated syngeneic SCs as antigen-presenting cells in the presence or absence of heat-inactivated virus. Supernatants were harvested after 72 h of culture, and IFN- levels were measured by ELISA. As seen in Fig. 7, T cells from uninfected C57BL/6 mice produced no IFN- after treatment in vitro with virus antigen. When T cells from infected C57BL/6 mice were examined, restimulation in vitro led to a significant elevation of IFN- production compared to cells from uninfected mice. T cells from infected CCR5-deficient mice produced IFN- in response to antigen restimulation to an even larger extent than T cells from infected C57BL/6 mice. This indicates that although splenic proliferation of lymphocytes in response to HSV-2 infection is impaired in CCR5–/– mice, antigen-specific T cells do develop in these mice after HSV-2 infection.
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produced, since T cells from CCR5–/– mice produce IFN- to levels at least comparable to what is seen in WT mice after virus rechallenge. Effector cell migration to the liver. Since the generalized HSV-2 infection in mice develops most strikingly in the liver, and since antiviral activity in this organ was impaired in CCR5-deficient mice, we examined accumulation of T-cell populations in the liver during infection. Livers were harvested from C57BL/6 mice and CCR5-deficient mice on day 5 p.i. and examined for the presence of cells positive for CD3/CD4 or CD3/CD8 (Fig. 9). With respect to CD3/CD4-positive cells, no infection-induced changes were observed, nor were any significant differences between C57BL/6 and CCR5–/– mice apparent (Fig. 9A to E). In C57BL/6 mice, we observed an increase in the number of CD3/CD8-positive cells on day 5 p.i. compared to uninfected mice. This increase was not seen in the livers from infected CCR5–/– mice (Fig. 9F to J). Thus, in 4-week-old mice lacking CCR5, fewer CD8 T cells are recruited to the liver during an acute generalized infection.
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DISCUSSION |
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Initially, we compared 4- and 8-week-old C57BL/6 mice with CCR1- and CCR5-deficient mice and found that young, but not adult, CCR5–/– mice exhibited impaired clearing of the infection in liver and brain. No significant differences between C57BL/6 and CCR1–/– mice were found with respect to clearance of HSV-2 infections. It has previously been demonstrated that CCR5 is important for clearance of infections by Cryptococcus neoformans and Toxoplasma gondii (14, 24) but not lymphocytic choriomeningitis virus, MHV, influenza virus, and Listeria monocytogenes (5, 22, 26, 36). Our work has thus disclosed an age-dependent role of CCR5 in defense against a virus infection. At least two potential explanations may underlay this finding. First, it is well-known that an age-dependent resistance against infections develops in mice when the animals reach sexual maturity (21, 25), and it is possible that the involvement of multiple mechanisms allows a sufficient antiviral response in adult CCR5–/– mice despite the lack of an antiviral mechanism dependent on this receptor. Second, young mice might harbor a CCR5-dependent antiviral defense mechanism not used in mature animals. For instance, T cells, which do express CCR5 (11), are more abundant early in life than during adulthood (12) and have recently been shown to be required for a protective immune response against an intestinal parasite in 4-week-old mice but not to contribute to defense in 7-week-old mice (25).
NK cells have long been recognized to play an important role in early defense against many viruses, including herpesviruses (2, 23, 28), and Salazar-Mather et al. (27) demonstrated that CCL3-deficient mice had higher levels of NK cells in the blood and spleen, but these cells could not migrate to the liver during a murine CMV infection, hence leading to impaired antiviral defense. When we examined the innate antiviral response at the site of infection, we found a rapid increase in the number of leukocytes and NK1.1-positive cells in the peritoneal cavity with no apparent differences between C57BL/6 mice and any of the knockout mice tested. It should be noted that NK1.1 is not expressed exclusively on NK cells but also on activated CD4- and CD8-positive T cells (30). However, since we examined the cells for NK1.1 expression on day 1 p.i., i.e., before activation of T cells, we believe that the vast majority of NK1.1-positive cells are indeed NK cells. When the cytotoxic activity of the recruited NK cells was examined we found that NK cells harvested from infected CCR5-deficient mice, but not from CCR1-deficient mice, displayed significantly lower activity than NK cells harvested from infected C57BL/6 mice. The underlying mechanism appears to be independent of IL-12 and IFN-/ß production, two well-known activators of NK cell cytotoxic activity (2, 31). The observed reduction in NK cell activity in CCR5–/– mice was associated with lower antiviral capacity in vivo, as assessed by adoptive transfer of NK activity to recipient CCR5–/– mice. In our protocol we transferred nonadherent cells and hence cannot exclude that other cell types, including plasmacytoid DCs with the potential to produce large amounts of IFN-/ß, could also have been delivered to the recipient mice and so contribute to the observed differences between the WT and CCR5–/– mice. However, since C57BL/6 and CCR5–/– mice did not differ with respect to HSV-induced IFN-/ß production we do not think contaminating cell types have affected the result significantly and thus conclude that the differences in viral load after adoptive transfer of NK activity can indeed be ascribed to differences in NK cell activity between C57BL/6 and CCR5–/– mice.
When the animals were examined macroscopically, we observed that, whereas the spleens of WT and CCR1–/– mice increased in size as the infection developed, spleens from CCR5–/– mice retained the same size. No difference between WT and CCR5–/– animals was found when we examined 8-week-old mice (data not shown). In agreement with the differences in spleen size between C57BL/6 and CCR5-deficient mice, we found that SCs harvested from 4-week-old infected CCR5-deficient mice proliferated significantly less ex vivo than cells harvested from infected C57BL/6 mice. When we looked at the T-cell subsets in the spleen, we noted that both CD4 and CD8 T cells were impaired with respect to expansion in response to infection in young CCR5-deficient animals. As for the underlying mechanisms behind this finding, several possibilities exist. First, it is possible that recruitment of immature DCs to the site of infection is impaired in the absence of CCR5. This would lead to reduced antigen presentation to T cells in lymphoid organs and hence the generation of a lower T-cell response. Second, it is possible that antigens are presented to the naive T cells, but proliferative signals are not sufficiently strong in the CCR5-deficient mice. For instance, low production of IL-2 would hamper proliferation. Alternatively, costimulatory molecules on T cells also play essential roles in the regulation of T-cell proliferation (4).
We observed reduced migration of CD8-positive T cells to the liver in the CCR5–/– mice in response to HSV-2, which suggests that the lack of CCR5 cannot be compensated for by other chemokine receptors. A similar defect in CD8+-T-cell trafficking has been observed in CCR5-deficient mice after disseminated Cryptococcus neoformans infection (14) or intracranial infection with MHV (10). Adoptive transfer of equal numbers of SCs from immune C57BL/6 and CCR5–/– mice to HSV-2-infected C57BL/6 mice did not recover the antiviral activity of CCR5 SCs, thus suggesting that in addition to the reduced T-cell proliferation in CCR5–/– mice, these cells also display reduced antiviral activity, either due to a defect capacity to migrate to the infected organs or due to reduced function of infiltrating T cells. CCR5 and its ligands have previously been demonstrated to contribute to recruitment of antiviral effector cells to the liver. The CCR5 ligand CCL3 is required for optimal recruitment of NK cells to the liver during CMV infection in mice (27). In humans, ca. 90% of hepatitis B virus (HBV)-specific CD8 T cells that infiltrate the liver of HBV patients are CCR5 positive compared to less than 30% among HBV-specific CD8+ T cells in the blood (29). Moreover, individuals that lack CCR5 exhibit increased susceptibility to hepatitis C virus infection (34).
In summary, we have demonstrated that CCR5, which serves as a coreceptor for HIV and is a potential target for antiviral drugs (33), is required for antiviral defense during a generalized HSV infection in young but not in adult mice. Lack of CCR5 led to impairment of both the innate and adaptive immune response. The present study therefore suggests that treatment of patients with CCR5 antagonists may have negative effects on the immune system in children and immunosuppressed individuals and also highlights the point that modulation of the immune system, as a means of treatment, requires fundamental knowledge of the immune system in different patient groups.
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ACKNOWLEDGMENTS |
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This study was supported by grants from The Danish Health Science Research Council (grants 22-02-0144, 22-03-0011, 22-03-0117, and 22-03-0183), the LEO Pharma Research Foundation, and the Kathrine and Vigo Skovgaards Fond.
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FOOTNOTES |
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