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Journal of Virology, September 2007, p. 10101-10112, Vol. 81, No. 18
0022-538X/07/$08.00+0     doi:10.1128/JVI.01242-07
Copyright © 2007, American Society for Microbiology. All Rights Reserved.

CCR5 and CXCR3 Are Dispensable for Liver Infiltration, but CCR5 Protects against Virus-Induced T-Cell-Mediated Hepatic Steatosis

P. J. Holst,¹ C. Orskov,² K. Qvortrup,² J. P. Christensen,¹ and A. R. Thomsen^1*

Institute of Medical Microbiology and Immunology,¹ Institute of Medical Anatomy, University of Copenhagen, the Panum Institute, 3C Blegdamsvej, DK-2200 N, Copenhagen, Denmark²

Received 7 June 2007/ Accepted 29 June 2007

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
CCR5 and CXCR3 are important molecules in regulating the migration of activated lymphocytes. Thus, the majority of tissue-infiltrating T cells found in the context of autoimmune conditions and viral infections express CCR5 and CXCR3, and the principal chemokine ligands are expressed within inflamed tissues. Accordingly, intervention studies have pointed to nonredundant roles of these receptors in models of allograft rejection, viral infection, and autoimmunity. In spite of this, considerable controversy exists, with many studies failing to support a role for CCR5 or CXCR3 in disease pathogenesis. One possible explanation is that different chemokine receptors may take over in the absence of any individual receptor, thus rendering individual receptors redundant. We have attempted to address this issue by analyzing CCR5^–/–, CXCR3^–/–, and CCR5/CXCR3^–/– mice with regard to virus-induced liver inflammation, generation and recruitment of effector cells, virus control, and immunopathology. Our results indicate that CCR5 and CXCR3 are largely dispensable for tissue infiltration and virus control. In contrast, the T-cell response is accelerated in CCR5^–/– and CCR5/CXCR3^–/– mice and the absence of CCR5 is associated with the induction of CD8⁺ T-cell-mediated immunopathology consisting of marked hepatic microvesicular steatosis.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Chemokines play a key role in virus-induced inflammation and virus clearance, acting as regulators of leukocyte trafficking (18, 49). Among the chemokine receptors, CCR5 has drawn considerable attention as the main coreceptor for human immunodeficiency virus and CXCR3 is a promising target for antiinflammatory therapeutics (41, 52). Notably, both receptors have been associated with type 1 cytokine T-cell responses and the majority of tissue-infiltrating T cells in several human inflammatory diseases are found to coexpress these receptors (7, 42, 48). During viral infection, CCR5 and CXCR3 are expressed on NK cells and activated CD4⁺ and CD8⁺ T cells and the ligands are generally expressed in virus-infected tissues (30, 38, 50). Further support for the important role of CCR5 and CXCR3 comes from the existence of virally encoded chemokine receptor antagonists targeting these receptors (23, 27). In this context, it is not surprising that CXCR3 or its ligands have been experimentally demonstrated to be important for immune-mediated virus control in the central nervous system (CNS), the lungs, the ovaries, and the islet of Langerhans in the pancreas (9, 15, 20, 26, 51). In contrast, despite the suggestive expression patterns, several studies have indicated that CCR5 or CCR5 ligands are less important in antiviral immunity (30, 37). The positive findings suggest a role for CCR5 in macrophage and CD4⁺ T-cell recruitment, but the receptor appears not to be required for CD8⁺ T-cell migration except in young mice, in which the immune system has not yet reached full maturity (3, 17). The reason for this difference between the two receptors could lie in a difference in biological redundancy. Thus, several CCR5 ligands can be expressed simultaneously and these can also signal through CCR1, which can be expressed on CD8⁺ T cells. On the other hand, CXCR3 is the only lymphocyte receptor for CXCL9 through CXCL11 and this may explain why the requirement for this receptor is less easily circumvented (31, 36).

Another factor clouding the interpretation of chemokine function in vivo remains organ-specific effects (8). Several reports have presented convincing, yet different conclusions regarding the requirement for these receptors in liver inflammation. For example, during chronic infections of hepatitis C, a disease entity affecting more than 200 million people worldwide (43), CCR5, CXCR3, and CXCR6 are expressed on liver-infiltrating lymphocytes (LILs) and the expression correlates with disease severity (53). Yet, contrary to expectations, the CCR532 human chemokine defect is associated with spontaneous hepatitis C clearance and lower inflammatory scores (19). In contrast, the CCR532 polymorphism or genetically engineered deficiency of CCR5 results in accelerated graft-versus-host disease in mice and reduced graft-versus-host disease in humans. Notably, the data are not unequivocal and the effect seems to depend on the regimen of pretransplant conditioning (35, 54, 55). The strongest evidence for a role of CCR5 in hepatic immunity in humans comes from the discovery that homozygosity for the CCR532 chemokine defect is associated with primary sclerosing cholangitis and an increased risk of ischemic-type billiary lesion following liver transplantation (13, 33). Additionally, the CCR5 ligand CCL3 has been suggested to play an essential role in virus control through the induction of gamma interferon (IFN-) and the production of CXCR3 ligand during acute murine cytomegalovirus (MCMV) infection (45, 46).

We have recently taken the rational approach and attempted to unravel redundancy within the chemokine system by generating CCR5/CXCR3 double-deficient (CCR5/CXCR3^–/–) mice. In a prior study, we infected these mice intracerebrally with the model virus, lymphocytic choriomeningitis virus (LCMV), and in that case, we found opposing actions of CCR5 and CXCR3 in relation to T-cell-mediated inflammation in the CNS (10). We have now taken the system further to evaluate chemokine receptor requirements for liver inflammation following intraperitoneal (i.p.) infection with a rapidly invasive strain of LCMV. This model system is characterized by dramatic expansion of virus-specific cytotoxic T lymphocytes that mediate nearly all virus-associated pathology as well as virus control during the acute phase of infection. Surprisingly, we find that both chemokine receptors are redundant for the elimination of virus infection in the liver but not for the regulation of virus-induced immunopathology.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Mice. The generation of CXCR3-deficient (CXCR3^–/–) mice has been described before (21). The animals used in these experiments were the progeny of breeder pairs kept at the Panum Institute, University of Copenhagen. CCR5-deficient (CCR5^–/–) mice (B6; 129P-CmKbr5<tm/Kn2>) were bred locally from breeder pairs obtained from the Jackson Laboratory (Bar Harbor, ME). CCR5/CXCR3^–/– mice were produced as recently described (10). Wild-type (WT) C57BL/6 mice were purchased from Taconic M & B (Ry, Denmark). Mice from outside sources were always allowed to rest for at least a week before being entered into experiments; by that time, the animals were about 7 to 9 weeks old. Animals were housed under controlled (specific-pathogen-free) conditions, and experiments were conducted according to national guidelines.

Virus infection. Mice were infected i.p. with a virus dose of 10⁵ PFU of LCMV clone 13 in a volume of 0.3 ml.

Organ virus titers. To determine virus titers in organs, the organs were first homogenized in phosphate-buffered saline (PBS) to yield 10% (vol/wt) organ suspensions and then serial 10-fold dilutions were prepared. Each dilution was then plated in duplicate onto MC57G cells. Forty-eight hours after infection, infected cell clusters were detected using monoclonal rat anti-LCMV (VL-4) antibody, peroxidase-labeled goat anti-rat antibody, and o-phenylenediamine (substrate) (6). The numbers of PFU were counted, and results were expressed as PFU per gram of tissue.

Splenocyte preparation. Single cell suspensions of splenocytes were obtained by pressing the organs through a fine steel mesh, and cells were counted in a hemocytometer.

Preparation of LILs. The protocol for liver leukocyte isolation was modified from that of Liu et al. (28). Mice were sacrificed by careful cervical dislocation. The abdomen was opened, and the portal vein was cut to create an outlet. The caval vein was then cannulated and perfused with 5 to 10 ml of PBS. Perfused livers were homogenized in 20 ml of Hank's balanced salt solution (HBSS) by passing the tissue through a fine steel mesh, and cells were pelleted by centrifugation at 500 x g. Pellets were resuspended by vortexing in gradient buffer (92.5 ml Percoll, 3.6 ml 20x PBS, 750 ml of 50 ku/µl of heparin, 2 ml of 4.16% NaHCO₃, and 152 ml of HBSS for each 250 ml). Leukocytes were then pelleted by 20 min of centrifugation at 800 x g, and the pellets were resuspended in 0.83% NH₄Cl. Cell suspensions were incubated for 10 min to allow erythrocytes to lyse, and cells were washed twice in HBSS, counted in a hemocytometer, and subjected to fluorescence-activated cell sorter analysis.

Antibodies and dextramers for flow cytometry. The following monoclonal antibodies were purchased from BD Pharmingen (San Diego, CA) as rat anti-mouse antibody: phycoerythrin (PE)-conjugated and Cy-chrome-conjugated anti-CD8, allophycocyanin-conjugated anti-CD4, fluorescein isothiocyanate (FITC)-conjugated anti-CD44, FITC-conjugated anti-Mac-1 (CD11b), FITC-conjugated CD11c, PE-conjugated anti-B220 (CD45R), PE-conjugated anti-NK1.1, Alexa 488-conjugated anti-T-cell receptor alpha/beta (TCR-/ß), PE-conjugated anti-IFN-, and PE-conjugated immunoglobulin G1 isotype standard. The detection of LCMV-specific CD8⁺ T cells was performed with PE-conjugated H-2D^b/gp_33-41 or H-2D^b/np_396-404 dextramers kindly provided by Dako (Glostrup, Denmark).

Flow cytometric analysis. Surface staining of cells for flow cytometry was performed according to standard laboratory procedure (2). For the evaluation of cytokine production by LCMV-specific CD8⁺ T cells, splenocytes were incubated in vitro at 37°C in 5% CO₂ with or without gp_33-41 peptide (0.1 µg/ml) in the presence of monensin (3 µM; Sigma Chemicals Co., St. Louis, MO), and murine recombinant interleukin-2 (10 U/well; R&D Systems Europe Ltd., Abingdon, United Kingdom). After incubation, cells were surface stained, washed, and permeabilized using 0.5% saponin. Cells were then stained with anti-IFN- or immunoglobulin G1 isotype control for 20 min at 4°C. Samples were analyzed using a Becton Dickinson FACSCalibur, and at least 10⁴ mononuclear cells were gated using a combination of low angle and side scatter to exclude dead cells and debris. Data analysis was conducted using Cell Quest Pro (B&D Biosciences).

In vivo CD8⁺ T-cell depletion. The depletion of CD8⁺ T cells was obtained by i.p. injection on day –1 and +1 relative to virus infection with the ascites from mice carrying the 2.43 hybridoma. The efficiency of the depletion was verified by flow cytometric analysis of splenocytes from the treated animals.

Detection of mRNA in the liver. Livers from mice deeply anesthetized and exsanguinated were immediately removed, snap frozen on dry ice, and stored at –80°C until processed. Total RNA was extracted by homogenizing livers in 4 M guanidine isothiocyanate (GITC), followed by centrifugation to pellet debris with double extraction of RNA from the supernatant by phenol and chloroform isoamylalcohol. The aqueous supernatant was then precipitated in isopropanol, and the pellet was resuspended in 4 M GITC, precipitated again using isopropanol, pelleted, and subsequently washed twice in ethanol. Pellets were resuspended in diethyl pyrocarbonate-treated water. Transcription levels were then studied using the RiboQuant multiprobe RNase protection assay (RPA) system (Pharmingen, San Diego, CA). The following templates sets (from Pharmingen) were used: cytokine marker mRNA (tumor necrosis factor beta [TNF-ß], LTß, TNF-, interleukin-6, IFN-, IFN-ß, transforming growth factor ß1 [TGF-ß1] to TGF-ß3, migration inhibition factor [MIF]), and chemokine marker mRNA (XCL1 [lymphotactin], CCL5 [RANTES], CCL4 [MIP-1ß], CCL3 [MIP-1], CXCL1 and CXCL2 [MIP-2], CXCL10 [IP-10], and CCL2 [MCP-1]). All sets of probes included templates for the housekeeping genes L-32 and GAPDH (glyceraldehyde-3-phosphate dehydrogenase) to serve as loading controls. The RPA was performed according to the manufacturer's instructions. Briefly, [-³²P]UTP-labeled antisense RNA transcript was generated from the template sets using T7 RNA polymerase. RNA from each sample was allowed to hybridize to the labeled probe for 16 to 20 h at 56°C. Single-stranded RNA was digested with an RNase/T1 mixture, and the hybrids were analyzed on a denaturing urea-polyacrylamide gel. For qualitative and quantitative results, gels were subjected to PhosphorImager analysis (Amersham Pharmacia Biotech) and the data were subsequently analyzed using ImageMaster TotalLab software with L-32 as the normalization control (Amersham Pharmacia Biotech).

Quantification of cytokine and chemokine levels in the liver. The amounts of cytokine or chemokine protein in the liver were evaluated using tissue homogenates containing protein inhibitor. Levels of IFN- were measured using a standard sandwich enzyme-linked immunosorbent assay, and chemokine levels were assessed using a multiplex fluorescent bead assay from Biosource and a Luminex analyzer. All analyses were carried out according to the manufacturers' instructions.

CD8 staining. Frozen liver sections were fixed in acetone and stained for CD8 immunoreactivity by using the rat anti-mouse CD8 antibody (catalog no. 550281; BD Biosciences, San Jose, CA) diluted 1:1,000. Immunoreactive cells were visualized using biotin-labeled anti-rat antibody, the tyramide signal amplification kit as instructed by the manufacturer (PerkinElmer, Boston, MA), and diaminobenzidine. No staining was seen in the absence of primary antibody. The tissue sections were lightly counterstained with hematoxylin.

Oil Red staining. Frozen liver sections were fixed in paraformaldehyde and stained with Oil Red O solution for 10 min at room temperature, followed by a light counterstain with hematoxylin. The percentage of Oil Red O staining was measured in four random visual fields from each slide using Image Pro, and the mean percent ± standard error of the mean (SEM) was then calculated for each slide.

Transmission electron microscopy. WT and CCR5/CXCR3^–/– mice were fixed by vascular perfusion through the left ventricle of the heart with 2% glutaraldehyde in 0.05 M sodium phosphate buffer (pH 7.2) for 2 min. Following fixation, the abdomen was opened and the liver was removed and stored in the same fixative. Following isolation of suitable specimen blocks, the samples were rinsed three times in 0.15 M sodium cacodylate buffer (pH 7.2) and subsequently postfixed in 1% OsO4 in 0.12 M sodium cacodylate buffer (pH 7.2) for 2 h. The specimens were dehydrated in graded series of ethanol, transferred to propylene oxide, and embedded in Epon according to standard procedures. Ultrathin sections were cut with a Reichert-Jung Ultracut E microtome and collected on 200 mesh copper grids with Formvar supporting membranes. The sections were stained with uranyl acetate and lead citrate and examined with a Philips CM 100 transmission electron microscope operated at an accelerating voltage of 80 kV and equipped with a SIS MegaView II camera. Digital images were recorded with the analySIS software package.

Statistical analysis. Quantitative results were compared using the Mann-Whitney U test. A P value of <0.05 was considered evidence of statistical significance.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Characterization of LCMV-induced hepatitis following i.p. infection with clone 13. To define the basic parameters of our hepatitis model, we analyzed the kinetics of leukocyte accumulation in the livers of mice infected i.p. with 10⁵ PFU of LCMV clone 13. Mice were sacrificed either before infection or 2, 5, and 7 days after infection, and the numbers of CD4⁺ and CD8⁺ T cells, NK cells, NKT cells, B cells, and Mac-1⁺ cells isolated from the perfused liver were determined (Fig. 1A). Two days after the infection, we did not find any increase in the number of LILs. Substantial cellular infiltration was noted 5 days after infection, with the most marked increases in numbers of NK and CD8⁺ T cells. Interesting also is that the number of B220⁺ cells increased significantly, raising the possibility that some of these cells were plasmacytoid dendritic cells. However, additional phenotypic characterization revealed that none of these cells were CD11c⁺ cells, demonstrating that few if any were dendritic cells. At day 7 after infection, the number of infiltrating CD8⁺ T cells had increased even further, whereas numbers of other cell types were either unchanged or had begun to decline.

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FIG. 1. Kinetics of the intrahepatic immune response in WT mice infected with 105 PFU of LCMV clone 13 i.p. (A) Leukocytes were purified from uninfected and infected animals at the indicated times after infection and were stained with monoclonal antibodies to reveal the total numbers of each of the main hepatic leukocyte subsets defined by CD4+/NK1.1– (CD4+ T cells), CD8+/CD8ß+ (CD8+ T cells), TCR-/ß+/NK1.1+ (NKT), TCR-/ß–/NK1.1+ (NK), B220+ (B cells), and Mac-1+ (macrophages). (B) Following virus infection, animals were sacrificed at the indicated times and viral titers in the liver were determined by plaque assay on homogenates. (C) Chemokine expression within infected and uninfected livers as determined by RPA assay and normalized to L32 expression. All data shown are averages for four to eight mice. Bars represent standard deviations.

To determine how the T-cell influx correlated with virus control and clearance, we quantified the viral load within the liver at days 2, 3, 4, 5, and 7 after infection (Fig. 1B). Viral titers were found to peak between days 3 and 5 after infection, and virus was present, but only at a very low level on day 7 postinfection.

To map the chemokines that might be involved in effector cell recruitment, we next screened broadly for likely candidates by using an RPA to look for relevant gene transcripts in the livers of uninfected mice and of mice infected 2, 5, and 7 days previously (Fig. 1C). We found that the expression of all of the chemokines studied here was increased in the livers of infected mice. Notably, the CXCR3 ligand CXCL10 was found to be maximally expressed on day 5 after infection, coinciding with peak viral titers, while the expression levels of the CCR5 ligands CCL3, CCL4, and CCL5 were stable between days 5 and 7 after infection. These correlations seem to match previous reports suggesting CXCL10 to be produced predominantly from virally infected cells and the CCR5 ligands to be produced from activated NK and T cells (4, 30). Based on these results, we selected days 5 and 7 postinfection as the most optimal time points for studying NK cell recruitment, the early phase of the CD8⁺ T-cell infiltration, and the resulting virus control.

Unimpaired leukocyte infiltration in CCR5^–/–, CXCR3^–/–, and CCR5/CXCR3^–/– mice. To define the actual chemokine/receptor interactions involved in effector cell recruitment, we infected chemokine receptor-deficient mice and WT mice in parallel. First, we investigated whether antiviral effector T cells were efficiently generated and capable of homing to the liver in the CCR5^–/–, CXCR3^–/–, and CCR5/CXCR3^–/– mice. To this end, we quantitated leukocyte subsets in the spleen and liver on days 5 and 7 after i.p. LCMV infection and compared the response of the chemokine receptor-deficient mice to that of WT mice.

All in all, there were few substantial differences in the cellular response patterns of the four different genotypes; however, a few trends could be discerned. On day 7 after infection, both CXCR3^–/– strains had significantly more CD8⁺ T cells in the spleen (Fig. 2A) and this was true also for cells with known specificity for LCMV (Fig. 2B). In contrast, both CCR5^–/– strains had higher numbers of liver-infiltrating CD8⁺ T cells, including LCMV-specific cells, on day 7 after infection than did CCR5-replete mice (Fig. 2A and B).

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FIG. 2. Liver and splenic lymphocyte subsets following infection of WT, CXCR3–/–, CCR5–/–, and CCR5/CXCR3–/– mice with 105 PFU of LCMV clone 13 i.p. Animals infected either 5 or 7 days previously were sacrificed, and spleen and liver leukocytes were collected. (A) Total numbers of NK cells (NK1.1+/TCR-/ß–), NKT cells (NK1.1+/TCR-/ß+), CD4+ T cells (CD4+/NK1.1–), and CD8+ T cells (CD8+/CD8ß+) (n = 8 mice/group). (B) Total numbers of LCMV-specific CD8+ T cells defined by CD8 staining and binding of either gp33-41 or np396-404 dextramers. All results shown are averages ± standard deviations (error bars) of four animals. *, P was less than 0.05, Mann-Whitney rank sum test versus WT mice.

Immunohistochemical visualization of CD8⁺ cells failed to reveal any obvious differences between the different mouse strains in the intrahepatic localization of the infiltrating cells (Fig. 3). In summary, these results clearly revealed that all of the tested mouse strains efficiently support the generation of antigen-specific CD8⁺ T cells following LCMV infection and that the generated cells are capable of infiltrating the liver.

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FIG. 3. No major difference in intrahepatic CD8+ T-cell localization in infected WT, CXCR3–/–, CCR5–/–, and CCR5/CXCR3–/– mice. Frozen liver sections from uninfected WT mice and sections taken 5 or 7 days after infection with 105 LCMV clone 13 were stained with rat CD8 antibody, followed by peroxidase-labeled -rat antibody. Shown are representative micrographs from each group (n = 4). dpi, days postinfection.

The number of NK and NKT cells also increased in the spleen, but no differences were observed between the genotypes, neither here nor in the liver, indicating that both of these cell types could infiltrate the LCMV-infected liver in the absence of CCR5 and CXCR3. Notably, no differences were found if uninfected WT and CCR5/CXCR3^–/– mice were compared (data not shown).

Slightly impaired virus control in CCR5 knockout mice, similar parenchymal liver damage, and similar inflammatory responses in WT, CXCR3, CCR5, and CCR5/CXCR3 double-knockout mice. In addition to studying the hepatic cell infiltrates, we also evaluated the ability of chemokine receptor-deficient mice to control and eliminate infectious virus from the liver, a process which requires direct cellular contact between virus-specific CD8⁺ T cells and virus-infected target cells. Remarkably, on day 5 postinfection, only CCR5^–/– mice had intrahepatic viral titers that were significantly higher than those of WT, CXCR3, and CCR5/CXCR3^–/– mice. Indeed, WT, CXCR3^–/–, and CCR5/CXCR3^–/– mice had almost identical viral titers (Fig. 4A). In WT mice and in both single-knockout strains, the virus load in the liver decreased significantly between days 5 and 7 postinfection, as would be expected from the marked influx of virus-specific CD8⁺ T cells. In contrast, hepatic infection was not efficiently controlled in CCR5/CXCR3^–/– mice. The differences in viral titers between the groups on day 7 did not, however, reach statistical significance.

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FIG. 4. Hepatic virus titers, parenchymal cell damage, and IFN- levels in WT, CXCR3–/–, CCR5–/–, and CCR5/CXCR3–/–, mice following i.p. infection with 105 PFU of LCMV clone 13. (A) Following virus infection, animals were sacrificed at the indicated times and virus titers in the liver were determined by plaque assays on the homogenates. Results represent averages ± SEMs of 4 to 13 mice; *, P was less than 0.05, Mann-Whitney rank sum test versus WT mice. (B) Five and 7 days after infection, mice were bled and serum was collected. Samples were analyzed for alanine aminotransferase (ALAT) and aspartate aminotransferase (ASAT) activity, and results were expressed in international units/milliliter. Results represent averages ± standard deviations (error bars) of four to eight mice; stippled lines denote average levels in uninfected mice. (C) Five and 7 days after infection, the concentration (conc.) of IFN- in liver homogenates was analyzed. *, significant difference at a P value of <0.05. Results represent averages ± SEMs; n = 4/group.

To supplement the evaluation of virus elimination, we also investigated the extent of parenchymal liver damage by measuring serum transaminase levels. We found these levels to be elevated roughly similarly in all infected mice regardless of genotype, with a trend of higher peak values for double-knockout mice (Fig. 4B).

As all the mouse strains investigated had a substantial influx of leukocytes and roughly similar viral loads and serum transaminase levels, we decided to see if any compensatory changes were occurring in any of the knockout strains regarding the virus-induced cytokine or chemokine response. To this end, we performed RPA analysis on cytokine and chemokine panels and found no differences between the genotypes in transcripts for the cytokines IFN-, TNF-, and TGF-ß (Fig. 5, top panels). With regard to IFN-, we also measured protein levels in the liver and, although statistically significant differences were observed (Fig. 4C), the absolute differences were small and probably not biologically important. The chemokine panels reproduced the patterns seen in WT mice (Fig. 1C), with the CXCR3 ligand CXCL10 peaking on day 5 and the CCR5 ligands CCL3 through CCL5 peaking on day 7 after infection (Fig. 5, bottom panels). Thus, the lack of CCR5 and CXCR3 did not induce any demonstrable compensatory changes in the chemokine profile, which supports the conclusion that CCR5 and CXCR3 are not essential for the accumulation of LILs following LCMV infection.

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FIG. 5. Lack of compensatory changes in cytokine or chemokine expression in CXCR3–/–, CCR5–/–, or CCR5/CXCR3–/– mice infected with 105 PFU of LCMV clone 13 i.p. RNA was purified from livers taken 5 or 7 days after infection in each of the above strains and from livers from WT mice, and relevant transcripts were quantified using RNase protection assays. Shown are the signals from the most highly expressed cytokines and chemokines normalized to the expression of the housekeeping gene L32; for levels in uninfected mice, see Fig. 1C. Shown are averages ± standard deviations (error bars); n = 3 in each group.

Profound microvesicular steatosis in infected CCR5^–/– strains. Though our initial readouts (leukocyte infiltration, viral titers, and parenchymal cell damage) failed to reveal any important role for CCR5 and CXCR3 in hepatic immunity, we were able to observe an antiinflammatory role of CCR5. Thus, during the dissection of i.p. infected CCR5^–/– and CCR5/CXCR3^–/– mice, we noted a macroscopic whitening of the liver surface and, upon the homogenization of liver tissue that we performed for leukocyte purification, a whitish fatty supernatant appeared. For this reason, tissue sections from infected WT, CXCR3^–/–, CCR5^–/–, and CCR5/CXCR3^–/– mice were stained with Oil Red O to reveal fat accumulation, and the relative hepatic fat accumulation was determined from consecutive randomized sections. This analysis revealed a dramatic accumulation of fat droplets within hepatocytes that contained central nuclei. This pattern was clearly dependent on the absence of CCR5 and protracted in CCR5/CXCR3^–/– mice (Fig. 6A). Steatosis was markedly reduced in all groups on day 9 after infection, and little if any fat accumulation was found at 15 and 30 days after infection (percentages of liver sections staining with Oil Red O were below 2% for both CCR5 and CCR5/CXCR3^–/– mice).

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FIG. 6. Hepatic steatosis and mitochondrial abnormalities in infected CCR5–/– strains. (A) Shown are representative images of frozen liver sections taken from mice infected either 5 or 7 days previously with 105 PFU of LCMV clone 13 i.p. The sections have been stained with Oil Red solution to reveal intracellular fat content. Numbers in the lower part of each picture show average area stained with Oil Red in percent of total liver section area ± SEM of four mice in each group. dpi, days postinfection. (B) Representative transmission electron micrographs from WT and CCR5/CXCR3–/– mice infected 5 days previously. Disrupted mitochondrial architecture with loss of clearly definable christae is evident in CCR5/CXCR3–/– mice. Bar = 1 µm.

To further characterize the nature of the apparent microvesicular steatosis, tissue sections from perfusion-fixed livers of WT and CCR5/CXCR3^–/– mice were investigated by transmission electron microscopy. This analysis supported the diagnosis of microvesicular steatosis and documented disorganized christae in the mitochondria of CCR5/CXCR3^–/– mice (Fig. 6B). This sign of mitochondrial dysfunction is often found under conditions of microvesicular steatosis associated with inflammation (16, 24, 25).

LCMV-induced hepatic steatosis is dependent on CD8⁺ T cells. The virus-induced hepatic steatosis in CCR5-deficient mice either could reflect direct virus-induced cell damage confined to mice with this genotype or could be the indirect result of a modified inflammatory response. As virus-specific CD8⁺ T cells cause most known LCMV-induced pathology, we performed a depletion of CD8⁺ T cells before and during LCMV infection in CCR5/CXCR3^–/– mice. The effect of this depletion was analyzed in CCR5/CXCR3^–/– mice on day 7 after infection, as this was the setting of the most consistent and dramatic steatosis. As can be seen in Fig. 7, virus-induced steatosis was almost completely abolished in CD8⁺ T-cell-depleted CCR5/CXCR3^–/– mice, despite increased hepatic infection in these mice. Notably, supplementary analysis of chemokine levels in the liver revealed that except for a significant reduction in the concentration of CXCL10, levels of all relevant chemokines, including the CCR5 ligands CCL3 through CCL5, were unaffected by the CD8⁺ T-cell depletion (Fig. 8).

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FIG. 7. Elimination of CD8+ T cells prevents LCMV-induced hepatic steatosis. (A) Representative micrographs of frozen liver sections from CCR5/CXCR3–/– mice infected 7 days previously with 105 PFU of LCMV clone 13 i.p. The top micrograph shows steatosis in infected mice, whereas the bottom micrograph depicts almost completely absent steatosis in mice treated with the CD8+ cell-depleting antibody 53.6.7. (B) Quantitation of hepatic steatosis and liver virus titers in infected CD8-depleted (53.6.7) mice and infected, untreated CCR5/CXCR3–/– mice (control). Bars depict the average area stained with Oil Red in percent of total liver section area ± SEM of four mice in each group. Numbers associated with the bars represent liver virus titers as PFU per gram of organ ± standard deviations of four animals. In both cases, the differences between CD8-depleted and undepleted mice were statistically significant (P < 0.05; Mann-Whitney rank sum test).

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FIG. 8. Elimination of CD8+ T cells only marginally affects chemokine levels in the liver. CCR5/CXCR3–/– mice were infected with 105 PFU of LCMV clone 13 i.p. Some of the mice were depleted of CD8+ cells using the antibody 53.6.7, and on day 7 after infection, chemokine levels in liver homogenates were assayed. Results represent averages ± standard deviations (error bars) for four mice per group.

Absence of CCR5 leads to a T-cell-intrinsic defect in the regulation of IFN- synthesis. Although the analysis of whole-organ homogenates for content of IFN- did not point to this cytokine as the crucial factor in mediating liver steatosis, the extent and distribution of the morphological changes did point to a T-cell-derived soluble factor as the underlying mechanism. Therefore, to study the regulation of cytokine synthesis in CD8⁺ T cells from CCR5^–/– mice more directly, we compared ex vivo production of IFN- in CD8⁺ T cells from these mice and matched wild types in the presence or absence of overt stimulation (plus/minus relevant peptide). The cells were incubated in vitro for 1, 3, or 5 h, and monensin was added for the last hour before analysis.

Expanding on our own published results, the most remarkable difference was an increased and sustained synthesis of IFN- in CD8⁺ T cells from CCR5^–/– mice in the absence of added peptide (Fig. 9, top panel). Given that previous analysis failed to reveal substantial differences between CCR5^–/– mice and matched wild types in the total number of gp₃₃-specific splenic CD8⁺ T cells (Fig. 2B), this pattern could reflect either an increased frequency of cells expressing virus naturally in the splenocyte cultures from CCR5^–/– mice or a less-stringent regulation of cytokine synthesis intrinsic to CD8⁺ T cells from these mice. In order to discriminate between these possibilities, we separated splenocytes from both types of mice into CD8⁺ and non-T cells and subsequently cultured the cells of either type with syngeneic cells or cells of the opposite genotype for 5 h. As can be seen clearly in Fig. 9, bottom panel, the capacity of the CD8⁺ T cells to "spontaneously" synthesize IFN- followed the origin of the T cells and not the genotype of the cells with which they were cocultured. This finding is most consistent with a less-stringent regulation of cytokine synthesis intrinsic to the CD8⁺ T cells from CCR5^–/– mice and suggests that once activated, the CD8⁺ T cells from CCR5^–/– mice would go on producing cytokine for a longer time period than would similarly stimulated WT CD8⁺ T cells.

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FIG. 9. Absence of CCR5 leads to impaired regulation of IFN- synthesis. (A) Splenocytes from WT and CCR5–/– mice infected with 105 PFU of LCMV clone 13 i.p. 7 days earlier were incubated in vitro with or without LCMV-derived peptide (gp33-41). At the indicated time points, monensin was added to the cultures, and CD8+ cells were analyzed for cytokine production by intracellular staining 1 h later; cells from uninfected mice were included as a control. Means of duplicate samples are presented; results are representative of two experiments. (B) Splenocytes from infected WT and CCR5–/– mice were separated into CD8+ cells and non-T cells, and each population was incubated with syngeneic or allogeneic cells of the opposite phenotype. After 5 h of incubation, CD8+ cells were analyzed for intracellular cytokine. Representative plots are depicted (n = 2/group).

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 
Our study is one of several attempting to address the requirement for CCR5 and/or CXCR3 in hepatic immunity, and it is the first to address liver immunology in mice expressing neither receptor (1, 3, 11, 34, 35, 56). It is also the first time liver inflammation has been investigated in mice lacking chemokine receptors in the LCMV system. Previous studies of liver immunology using LCMV have mostly been using the WE strain, and this is one of a few using the Armstrong clone 13 strain. Using this model, we achieved a markedly lower hepatic virus load and lower increases in serum transaminase levels than what was reported using the WE strain (5), yet an appreciable inflammatory infiltrate is evident (Fig. 1A to C). These observations suggested that this system would be useful for elucidating intrahepatic virus-induced immune responses without excessive induction of unspecific inflammation.

Our results in the chemokine knockout strains clearly demonstrate that neither NK nor CD4⁺ or CD8⁺ T cells absolutely require the expression of CCR5 and/or CXCR3 for liver infiltration (Fig. 2). This seemingly contrasts with previous reports for cytomegalovirus-infected mice lacking CXCR3 or the CCR5 ligand MIP-1, a system in which both MIP-1 and the CXCR3 ligand CXCL9 are needed for efficient virus control (22, 45); yet there are some possible explanations. First, LCMV is a much simpler virus than MCMV and induces many different chemokines with few viral countermeasures (Fig. 1C) (49). Second, MCMV directly infects hepatocytes, whereas LCMV (the Armstrong strain in particular) primarily causes infection of Kuppfer cells and the periportal areas (12, 39). Therefore, different chemokine requirements may exist for cell infiltration and virus control in these two viral systems. In contrast to the intact or even increased hepatic inflammatory infiltrate in mice lacking CCR5, virus control was modestly impaired compared to that in WT, CXCR3^–/–, and intriguingly, CCR5/CXCR3^–/– mice (Fig. 2 and 4). Thus, our attempt to unravel redundancy by using double-chemokine-receptor-knockout mice eliminated the advantage it was supposed to confer. This result is, however, similar to what we recently reported for LCMV-induced inflammation in the CNS (10). It must be stressed that the mechanism by which CXCR3 deficiency improves initial virus control in CCR5^–/– mice remains undetermined, yet the literature offers a possible explanation. Shields et al. (47) studied chemokine expression during hepatitis C virus infection in the liver and found CCR5 ligands to be expressed in the periportal areas, whereas the CXCR3 ligand was expressed from the liver sinusoids and hepatocytes. Accordingly, the balance between different stimuli would determine the localization of CCR5/CXCR3 double-positive cells (a similar mechanism is in fact determining splenic leukocyte positioning) (44). When applied to our system, a lack of CCR5 might drive the CD8⁺ T cells away from the periportal areas, where most of the virus is located, deeper into the liver parenchyma, where CXCR3 ligands would be present. In this situation, concomitant loss of CXCR3 stimulation might restore periportal localization through other signals. Unfortunately, the majority of our LILs are not antigen specific and we have not been able to costain sections with both antigen-specific major histocompatibility complex dextramers and portal tissue-specific stains. Thus, a direct demonstration of an altered intrahepatic localization remains elusive. In any case, the effect of selective CCR5 deficiency on viral titers is transient and modest and any differences in intrahepatic localization could likewise be subtle.

Remarkably and quite unexpected, despite almost normal viral control, morphological analysis demonstrated a marked microvesicular steatosis in both strains of CCR5-deficient mice (Fig. 6). In our system, the presence of any appreciable fatty change within the livers of infected mice clearly depended on the absence of CCR5, yet similar steatotic changes were observed previously in WT mice infected with high doses of the rapidly invasive WE strain of LCMV and indeed in the context of viral infections other than LCMV (29, 32). We are, however, the first to demonstrate that such pathological changes can be CD8⁺ T-cell dependent in a system where other features of virus-associated pathology are rather limited (Fig. 7). Indeed, our observation of specific inflammatory changes in the absence of CCR5 may add to the understanding of our own recent report of opposing effects of CCR5 and CXCR3 on LCMV-induced meningitis (10). LCMV-induced hepatic steatosis was found to be pronounced and short lived in CCR5^–/– mice, yet it was maintained at increasing levels until day 7 in CCR5/CXCR3^–/– mice. We suggest that this pattern may reflect the fact that only double-knockout mice failed to reduce the viral load between days 5 and 7 postinfection.

We noticed that the clinical history of LCMV-induced steatosis has several features in common with virus-induced microvesicular steatosis in humans, i.e., classical Reye's syndrome, an apparently virus-triggered, acute liver failure and encephalopathy in children hallmarked by similar light and electron microscopical findings in the liver (16, 24). Indeed, the requirement for CCR5 also seems to vary according to age in a herpes simplex model of liver inflammation (3). However, aspirin administration, the use of which is associated with Reye's syndrome (14) in humans, failed to increase the pathology in our mice (data not shown). If the human CCR5 polymorphism plays a similar role in classical Reye's syndrome, other predisposing factors are likely to vary considerably. Thus, although identical mechanisms at the genetic level between our model and Reye's syndrome remains elusive, we have demonstrated that microvesicular steatosis can be triggered by CD8⁺ T-cell activities even when viral replication in the liver is quite limited, and this could very well be a mimic to the situation in influenza-infected humans.

Although the observation of CCR5-dependent protection from virus-induced hepatic steatosis is unique, recent reports have demonstrated that CCR5 protects mice against the hepatotoxic effects of systemic concanavalin A administration (1, 34). Taken together with our study, it seems that evidence for an important role of CCR5 in controlling hepatic immunopathology is accumulating.

Precisely how the CD8⁺ T cells induce liver steatosis is not clear. Given the extent of the observed steatosis, a soluble factor seems most likely to be the underlying mechanism. Yet, variation in neither whole-organ levels of mRNA for several proinflammatory mediators nor protein levels of IFN- or CCR5 ligands readily explains the association between steatosis and the lack of CCR5 expression. On the other hand, our in vitro analysis of CD8⁺ T cells from CCR5^–/– mice points to a prolonged synthesis of IFN- by the activated T cells in these mice, which could support the assumption that this cytokine and/or other associated secreted molecules may play an important role in the observed pathology. As to the reason for the discrepancy between our in vivo and in vitro results, it could be argued that the in vitro assay much more precisely detects differences in cytokine synthesis by activated CD8⁺ T cells, whereas analysis of the whole organ simply gives a snapshot of the accumulated potential of these cells, many of which may not realize this normally.

In summary, we have shown that neither CCR5 nor CXCR3 is necessary for virus-induced NK cell or CD4⁺ and CD8⁺ T-cell infiltration into the liver. Lack of CCR5 resulted in a very modest impairment of virus control that was initially counteracted by a concomitant lack of CXCR3. Lack of CCR5 or both CCR5 and CXCR3 resulted in marked virus-induced CD8⁺ T-cell-mediated steatosis. Our findings may shed light on studies showing adverse effects of CCR5 deficiency in studies of human immunopathology (13, 33, 40).

 
This work was supported in part by the Novo Nordisk Foundation, the Danish Cancer Society, the Lundbeck Foundation, the Sophus C. E. Friis Foundation, the Foundation for the Advancement of Medical Sciences, the Aase and Ejnar Danielsen Foundation, the Hede Nielsen Family Foundation, the Leo Nielsen Foundation, Merchant Foght's Foundation, the Christian and Ellen Larsen Foundation, and the Leo Pharma Research Foundation. P.J.H. is the recipient of a research fellowship from the Faculty of Health Science, University of Copenhagen, Copenhagen, Denmark.

We thank Grethe Thorner Andersen and Lone Malte for expert laboratory assistance and Grazyna Hahn for technical assistance. Major histocompatibility complex dextramers were kindly donated by Jørgen Schøller, Dako, Glostrup, Denmark.

 
* Corresponding author. Mailing address: Institute of Medical Microbiology and Immunology, University of Copenhagen, The Panum Institute, building 22.5.16, 3C Blegdamsvej, DK-2200 Copenhagen N, Denmark. Phone: 45-35-32-78-71. Fax: 45-35-32-78-74. E-mail: a.r.thomsen{at}immi.ku.dk

Published ahead of print on 11 July 2007.

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Journal of Virology, September 2007, p. 10101-10112, Vol. 81, No. 18
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