TLR9 Contributes to Antiviral Immunity during Gammaherpesvirus Infection¹

Diseases caused by gammaherpesviruses continue to be a challenge for human health. The prototypic gamma 1-herpesvirus, EBV, is associated with lymphomas and nasopharyngeal carcinoma (1). Human herpesvirus-8 (also called Kaposi’s sarcoma-associated herpesvirus), a gamma 2-herpesvirus, is associated with lymphoproliferative disorders and Kaposi’s sarcoma (2). In vivo studies of gammaherpesvirus pathogenesis have been limited to clinical investigation of the infection because of the restricted host range of these viruses. Recently, the murine gammaherpesvirus 68 (MHV-68)³ has been established as a mouse model for the study of gammaherpesvirus pathogenesis (3, 4, 5, 6, 7, 8). MHV-68 is a natural pathogen of wild rodents (9) and is capable of infecting laboratory mice. The nucleotide sequence of MHV-68 is similar to EBV and even more closely related to Kaposi’s sarcoma-associated herpesvirus (10). In particular, MHV-68 is very useful to study the role of immunity in gammaherpesvirus infection (11, 12, 13, 14, 15).

Host immune responses play a pivotal role in the control of gammaherpesvirus infection and in pathogenesis. Whereas the adaptive immune response during gammaherpesvirus infection has been an area of intensive research, surprisingly little is known about the role of innate immunity in the control of gammaherpesvirus infection (16, 17). The TLR system is responsible for the primary recognition of infectious agents leading to the initiation of the innate and adaptive immune response (18, 19). Recently, a number of viruses, for example, HSV, CMV, respiratory syncytial virus, influenza A virus, and vesicular stomatitis virus, have been shown to activate cells via TLR family members (20, 21, 22). The activation of TLRs by viruses might lead to antiviral immune responses but viruses may also use these pathways to enhance their own replication (20). The important role of TLRs in antiviral immune responses is also mirrored by viral immune evasion strategies used against TLRs (22).

In a very recent study, it has been shown that EBV particles induce NF-κB activation in transfected human embryonic kidney cells and chemokine secretion by primary monocytes in a TLR2-dependent manner (23). The authors did not show whether intracellular TLRs like TLR9 also play a role after uptake of virus. TLR9 recognizes unmethylated CpG DNA motifs that are present in bacterial and viral DNA (19). Accordingly, it has been shown that TLR9 is required for IFN-α production in response to DNA viruses including murine CMV (MCMV) and HSV (19, 21, 22). There are some hints that gammaherpesviruses might also interact with TLR9. EBV-stimulated human plasmacytoid dendritic cells (DCs) promote the activation of NK cells and CD3⁺ T cells. This activation was dependent on cell-to-cell contact and was partially linked to TLR9 signaling (24). MHV-68 can induce IL-12 production in macrophages and DCs (25). HSV-1-induced IL-12 production during infection is mediated by TLR9 (26).

Thus, we considered TLR9 as a potential candidate to be activated by gammaherpesvirus infection and wanted to study its role in particular in vivo after MHV-68 infection. We demonstrate that TLR9 mediates the production of inflammatory cytokines by Flt3 ligand-cultured bone marrow cells (FL-DCs) in response to MHV-68 infection. By infection of TLR9^−/− mice, we show that TLR9 is involved in the antiviral immune response to MHV-68 infection. In the absence of TLR9 expression, lytic virus titers in the lung 6 days after intranasal (i.n.) infection were not affected but were increased in the spleen after i.p. infection. Similarly, in the absence of TLR9, the latent virus load in the spleen 17 days after infection was increased after i.p. infection but not after i.n. infection. Thus, we provide for the first time genetic evidence for an interaction of a gammaherpesvirus with TLR9. We demonstrate that TLR9 plays an important role in gammaherpesvirus immunity both during lytic infection and latency amplification and contributes to organ-specific immunity.

Baby hamster kidney cells (BHK-21) were maintained in Glasgow’s modified Eagle’s medium (Biochrom) supplemented with 5% FCS, 5% tryptose-phosphate broth, 100 U/ml penicillin, 100 μg/ml streptomycin, and 2 mM l-glutamine. NIH3T3 cells were grown in DMEM high glucose (Cell Concepts) supplemented with 10% FCS, 2 mM l-glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. Human embryonic kidney 293 cells were maintained in DMEM with 10% FCS, 2 mM l-glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. MHV-68 stocks were propagated and viral titers were determined by plaque assay on BHK-21 cells as described (27). Briefly, 10-fold dilutions were incubated on BHK-21 cells for 90 min at 37°C. After removing the inoculum, cells were incubated for 5 days at 37°C with fresh medium containing 1.5% methylcellulose. After 4–5 days, cells were stained with 0.1% crystal violet solution to determine the number of plaques.

To obtain FL-DCs, bone marrow cells from wild-type (wt) and knockout mice were cultured with 35 ng/ml human recombinant Flt3L (R&D Systems) for 8 days as described (28). FL-DCs contain a mixture of plasmacytoid DCs (pDCs) and myeloid DCs (28), and pDCs are known to recognize viruses via TLRs. FL-DCs were stimulated with the TLR4-ligand LPS (Sigma-Aldrich), the TLR7/8-ligand R848 (GLSynthesis), the TLR9-ligand CpG-oligodeoxynucleotide 2216 (gggggacgatcgtcgggggg; Ref. 29) (TIB MOLBIOL) and MHV-68 (multiplicity of infection (MOI) of 1 and 0.1, respectively) for 24 h. Afterward, supernatants were harvested and stored at −80°C for cytokine determinations.

TLR9^−/− mice, backcrossed to C57BL/6 mice for 10 generations, were bred under standard pathogen-free conditions in the animal facility of the Institute of Medical Microbiology, Immunology and Hygiene (Technical University Munich). Age-matched C57BL/6 mice (wt controls) were purchased from Charles River Laboratories. All mice were housed in individually ventilated cages during the MHV-68 infection period.

wt and knockout mice were anesthetized using ketamine/xylazine and infected i.n. with 5 × 10⁴ PFU of MHV-68 in 30 μl of sterile PBS. Alternatively, mice were infected i.p. with 5 × 10⁵ PFU in 250 μl of sterile PBS. To assess lytic viral replication, lungs were harvested on day 6 after i.n. infection and spleens on day 6 after i.p. infection. Viral titers in organ homogenates were determined as described previously. The detection limit of the assay is 50 PFU/organ as determined by spiking uninfected organs with known amounts of virus (30). On day 17 after either i.n. or i.p. infection, spleens were harvested; single splenocyte suspensions were prepared and analyzed in the ex vivo limiting dilution reactivation assay as described (30). Briefly, serial 3-fold dilutions of infected mouse splenocytes were plated on monolayers of 1 × 10⁴ low-passage NIH3T3 cells/well in 96-well tissue culture plates. Twenty-four wells were plated per dilution (starting with 1.5 × 10⁵ cells/well). NIH3T3 cells were screened microscopically for a viral cytopathic effect for up to 3 wk. To differentiate between latently infected cells and infectious virus in the samples, serial 3-fold dilutions of spleen cells were plated before or after mechanical disruption of viable cells (by two freeze-thaw cycles). No infectious virus was detected in samples of mechanically disrupted cells (data not shown). Frequencies of reactivating cells were calculated on the basis of the Poisson distribution by determining the cell number at which 63.2% of the wells scored positive for CPE. All animal experiments were in compliance with protocols approved by the local animal care and use committee.

Viral load in the spleens of infected mice was determined by quantitative real-time PCR using the ABI 7300 Real Time PCR System (Applied Biosystems). DNA was extracted from spleen cells using the QIAmp DNA Mini kit (Qiagen) and quantified by UV spectrophotometry. Amplification of 100 ng of DNA per reaction was performed with TaqMan universal PCR master mix and universal cycling conditions (Applied Biosystems). Using primers and probes as described (31), a 70-bp region of the MHV-68 glycoprotein B (gB) gene was amplified and viral DNA copy number was quantified. A standard curve was created using known amounts of a plasmid containing the HindIII-N fragment of MHV-68 encompassing the gB gene. The murine ribosomal protein L8 (rpl8) was amplified in parallel and used to normalize for input DNA between samples. The primer and probe sequences for L8 were as follows: forward: 5′-CATCCCTTTGGAGGTGGTA-3′; reverse: 5′-CATCTCTTCGGATGGTGGA-3′ and probe: 5′-ACCACCAGCACATTGGCAAACC-3′. A standard curve for rpl8 was generated by serial 10-fold dilution of a plasmid containing rpl8 (RZPD clone IRAVp968B01123D6). The data are presented as viral genome copy numbers relative to the copy number of L8. The quantification limit was set at 50 copies per sample, according to published recommendations (32).

IL-12 p40/p70 (BD Pharmingen), IL-6 (BD Pharmingen), and IFN-α (PBL Biomedical Laboratories) in supernatants of FL-DCs were determined by ELISA as recommended by the manufacturers.

For FACS analysis, 10⁶ cells were suspended in 100 μl of FACS buffer (PBS, 2% FCS). Nonspecific binding of Abs to FCR was blocked by incubating cells with 1 μg of the anti-CD16/CD32 mAb 2.4G2 (BD Pharmingen). After 5 min at 4°C, the relevant mAbs were added at a concentration of 0.5 μg/10⁶ cells and cells were incubated for 30 min at 4°C. The following mAbs from BD Pharmingen were used: FITC-conjugated anti-mouse CD45R/B220 (RA3-6B2), FITC-conjugated anti-mouse CD3 molecular complex (17A2), FITC-conjugated anti-mouse CD4 (L3T4) (GK1.5), FITC-conjugated anti-mouse CD8a (Ly-2) (53-6.7), FITC-conjugated anti-mouse CD14 (rmC5-3), PE-conjugated anti-mouse Ly-6A/E (Sca-1) (E13-161.7), allophycocyanin-conjugated anti-mouse NK1.1 (PK136), FITC-conjugated rat IgG2a, κ isotype standard (R35-95), and FITC-conjugated rat IgG2b, κ isotype standard (A95-1). After staining, cells were washed twice, resuspended in 500 μl of 0.5% paraformaldehyde in PBS and analyzed on a FACSCalibur using CellQuest software (BD Biosciences).

Lungs of mice were harvested on day 6 after intranasal infection and fixed in 10% formalin in PBS. For histopathological analysis, organs were embedded in paraffin, sections were cut, stained with H&E and analyzed by light microscopy.

If not otherwise indicated, data were analyzed by Student’s t test.

To investigate a potential role of TLR9 in gammaherpesvirus-host cell interaction, we stimulated FL-DCs, generated from either wt or TLR9^−/− mice, with MHV-68 (MOI = 0.1). After 24 h, the supernatants were analyzed for IL-12 p40/p70 and IFN-α by ELISA. As positive control, a type A CpG-oligodeoxynucleotide (2216) was used for stimulation. Although FL-DCs generated from wt mice produced IL-12 in response to both MHV-68 and CpG-DNA, the production of IL-12 was abolished in the absence of TLR9 in either case (Fig. 1,A). Similarly, the production of IFN-α in response to CpG-DNA was abolished in the absence of TLR9. In contrast, the production of IFN-α in response to MHV-68 was reduced but not completely abolished in the absence of TLR9 (Fig. 1,B). To further define the role of TLR9 in the production of inflammatory cytokines by FL-DCs in response to MHV-68, we examined the production of IL-6, a cytokine produced at high levels during MHV-68 infection (33). FL-DCs generated from wt mice produced IL-6 in response to both MHV-68 and CpG-DNA. In the absence of TLR9, the production of IL-6 was abolished in response to CpG-DNA and significantly reduced in response to MHV-68 (Fig. 1 C). The induction of IL-6 by MHV-68 in FL-DCs generated from wt mice was dose-dependent (MOI of 1 and 0.1, respectively). In control cultures, FL-DCs generated from both wt or TLR9^−/− mice produced IL-6 in response to LPS (TLR4 ligand) and R848 (TLR7/8 ligand). In addition, heat-inactivated MHV-68 (1 h, 65°C) was used as control. Heat denaturation, which leads to the disruption of the viral envelope and thereby prevents infection, prevented the induction of IL-6. Taken together, these results strongly suggested a role of TLR9 in gammaherpesvirus-host cell interaction.

To analyze the role of TLR9 in gammaherpesvirus pathogenesis, we infected C57BL/6 and TLR9^−/− mice with MHV-68. In a first series of experiments, mice were infected i.n. with 5 × 10⁴ PFU of MHV-68. Lytic viral replication in the lungs was determined by plaque assay on day 6 postinfection (p.i.), the time point at which viral titers usually reach a peak. Comparable viral titers were observed in both groups of mice (Fig. 2). In addition, histopathological analysis of lung tissue revealed no obvious differences either (data not shown). The establishment of latency in the spleen is associated with a marked splenomegaly and an increase in the number of latently infected B cells which peaks around 2–3 wk p.i. Thus, spleens of infected mice were harvested 17 days p.i. to assess the role of TLR9 during latent infection. The number of latently infected cells reactivating virus, as determined by an ex vivo reactivation assay, was comparable between wt and TLR9^−/− mice. In wt mice, the frequency of reactivating splenocytes was 1 in 6,145 cells. The corresponding number in TLR9^−/− mice was 1 in 5,837 (Fig. 3,A). Preformed infectious virus was not detected in spleens harvested 17 days p.i. (data not shown). Consistent with the reactivation data, spleens of infected wt and TLR9^−/− mice harbored similar amounts of viral genomes as determined by quantitative real-time PCR (Fig. 3,B). Thus, both lytic and latent MHV-68 infection were not altered in TLR9^−/− mice after i.n. infection. In a second series of experiments, mice were infected i.p. with 5 × 10⁵ PFU of MHV-68. On day 6 p.i., spleens were harvested and analyzed. As shown in Fig. 4, i.p. infection resulted in significantly higher lytic viral titers in the spleens of TLR9^−/− mice compared with wt mice, as determined both by plaque assay (Fig. 4,A) and quantitative real-time PCR (Fig. 4,B). To assess the role of TLR9 during latency amplification after i.p. infection, spleens of MHV-68-infected mice were harvested 17 days p.i. and analyzed both by ex vivo reactivation assay and quantitative real-time PCR. The number of spleen cells reactivating virus was significantly higher in TLR9^−/− mice than in wt controls (frequency of reactivation: 1 in 40,124 in TLR9^−/− mice and 1 in 88,878 in wt mice) (Fig. 5,A). In accordance with these findings, the viral genomic load in spleens of knockout mice was significantly higher than in wt mice (Fig. 5 B). Preformed infectious virus was not detected in spleens harvested 17 days after i.p. infection (data not shown).

In this study, using MHV-68 as a model, we demonstrated that TLR9 plays an important role in gammaherpesvirus immunity both during lytic infection and latency amplification. We could show that TLR9 mediates the production of inflammatory cytokines by FL-DCs in response to MHV-68 infection in vitro. Although the production of IL-12 was abolished in the absence of TLR9, some residual production of IFN-α and IL-6 was observed. These results suggest that MHV-68 may, as shown for other viruses, also engage mechanisms other than TLR9 to induce cytokine secretion. For example, HSV-1 can activate both TLR2 and TLR9 (26, 34). For MCMV, both TLR9/MyD88-dependent and -independent processes for IFN-α release have been described (35). MHV-68 infection induces a number of cytokines, for example, IFN-α/IFN-β, IFN-γ, IL-6, IL-10, and IL-12 (25, 33, 36). The type I IFNs have been shown to play a key role in the control of early (37) as well as latent MHV-68 infection (38). MHV-68-induced IL-12 functions to limit the viral burden but also contributes to virus-mediated splenomegaly (25). Although the cellular sources of the MHV-68-induced IFN-α/IFN-β have not yet been analyzed in detail, DCs have been shown to be a source for MHV-68-induced IL-12 (25). MHV-68-induced IL-10 increases the viral load but limits the virus-induced splenomegaly (39). IL-6 appears to be not essential for the development of an effective immune response to MHV-68 (40). IL-10 and IL-6 have been shown to be produced both by T cells and non-T cells (33).

The absence of TLR9 expression resulted in increased lytic virus titers in the spleen after i.p. infection but not in the lung after i.n. infection. Similarly, in TLR9-deficient mice, the latent virus load in the spleen 17 days after infection was increased after i.p. but not after i.n. infection. Thus, the role of TLR9 in gammaherpesvirus immunity seems to depend on the route of infection and to be organ specific. The natural route of MHV-68 infection is unknown but intranasal infection is believed to reproduce mucosal infection which is characteristic of natural herpesvirus transmission (15). After i.n. infection, primary lytic replication takes place in lung epithelial cells. Virus is then transported to lymphoid tissue, most likely by infected DCs. Infected B cells from the mediastinal lymph nodes then traffic to the spleen and other lymphoid organs and establishment of lifelong latency takes place (5, 15). The establishment of latency in the spleen is associated with a strong increase in the number of latently infected B cells (41). In addition to B lymphocytes which are the major reservoir harboring latent MHV-68 (42), macrophages (43), DCs (44), and lung epithelial cells (45) have also been shown to harbor latent virus. In contrast to i.n. infection, i.p. infection seeds lytic virus directly to the spleen and thus allows splenocytes to be infected by direct lytic spread (46). As a consequence, TLR9-expressing cells may come in direct contact with lytic virus in the spleen after i.p. infection but not after i.n. infection, providing a possible explanation as to why the effect of TLR9 on latency amplification is apparent only after i.p. but not after i.n., infection. With regard to lytic replication, we again observed an effect of TLR9 only in the spleen but not in the lung. Clearly, in this case, lytic virus is present in both organs and thus could interact with TLR9-expressing cells. However, it has been shown by Northern blot analysis that TLR9 is much stronger expressed in the spleen than in the lung and a variety of other tissues (47). In addition, it has recently been demonstrated that both myeloid DCs and pDCs in the lung show no detectable expression of TLR9 while both subsets in the spleen express TLR9 (48). Consequently, CpG oligonucleotides exerted differential effects on lung and spleen DCs when administered to mice (48). Similarly, in our study, the absence of TLR9 resulted in differential effects in the lung and spleen. In TLR9-deficient mice, lytic virus replication in the lung was undistinguishable from wt mice, which would be consistent with the above-mentioned fact that TLR9 expression was undetectable in lung DCs of wt mice. In contrast, absence of TLR9 in the spleen, an organ where TLR9 is regularly expressed in DCs of wt mice, resulted in significantly higher lytic virus titers in TLR9-deficient mice. Although differences in TLR9 expression between lung and spleen might explain our results, there are also other possibilities which can be envisaged: dependent on the route of infection (i.n. vs i.p.), different cell types in the spleen may be infected. However, it has been demonstrated that establishment and maintenance of gammaherpesvirus latency are independent of the infective dose and route of infection (49). In contrast, analyses of MHV-68 mutants have shown that viral genes such as M2 play a role specifically after i.n. but not after i.p. infection. This suggests that the requirements for the establishment of latency are affected by the route of infection (50). Thus, it might be possible that subtle differences in the cell types infected in the spleen by different routes of infection may account for our results. This deserves further studies.

It has been hypothesized that the biological significance of the absence of TLR9 in lung DCs may be a protective measure that has evolved to protect the lung from the development of diseases that are associated with high cytokine (IL-6) production such as pulmonary fibrosis (48). Thus, it is tempting to speculate that the same applies to MHV-68 lung infection, namely that the “physiological” absence of TLR9 in lung DCs may prevent an overwhelming innate immune response, for example, inflammatory cytokine production, and thereby limits pulmonary disease while having only little effect on virus replication. Histopathological analysis of lung tissue revealed indeed no obvious differences between infected wt and TLR9-deficient mice. It has very recently been suggested that innate immunity functions in an organ-specific fashion designed to sustain organ physiology, for example, by different expression profiles of pattern-recognition receptors between organs (51). Supporting observations in this direction have been made with HSV-1 and West Nile virus. In the case of HSV-1, TLR2-mediated induction of inflammatory cytokines in the brain of infected mice was not protective but associated with lethal encephalitis. As a result, TLR2-deficient mice showed reduced mortality when compared with wt mice (34). In the case of West Nile virus, TLR3-induced inflammatory responses contribute to pathogenesis rather than to protection by triggering a breakdown of the blood-brain barrier. As a consequence, TLR3-deficient mice survive an otherwise lethal infection because of reduced virus entry into the brain (52). In contrast, activation of TLRs is often associated with protective antiviral innate immune responses (20, 22). For example, both TLR3- and TLR9-deficient mice are more susceptible to MCMV infection (35, 53). MyD88, a key intermediate of multiple TLR-signaling pathways, is essential for the induction of type I IFN, the production of neutralizing Abs and protection of mice from lethal infection after i.n. but not after i.v. infection with vesicular stomatitis virus (54). Thus, TLR activation may either reduce or exacerbate disease, depending on the pathogen and the location of the infection (34).

In summary, we provide for the first time genetic evidence for an interaction of a gammaherpesvirus with TLR9. We demonstrate that TLR9 plays an important role in gammaherpesvirus immunity both during lytic infection and latency amplification, and that TLR9 contributes to organ-specific immunity.

We are grateful to Dr. B. Adler for critical reading of the manuscript and to Dr. R. Kammerer for providing reagents.

The authors have no financial conflict of interest.