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The Journal of Immunology, 2007, 178: 3536-3543.
Copyright © 2007 by The American Association of Immunologists, Inc.

Increased Expression of Leukocyte Ig-Like Receptor-1 and Activating Role of UL18 in the Response to Cytomegalovirus Infection1

Claudia S. Wagner2,*, Gerdt C. Riise{dagger}, Tomas Bergström{ddagger}, Klas Kärre§, Ennio Carbone§,¶ and Louise Berg§

* Center for Infectious Medicine, Department of Medicine, Karolinska Institutet, Stockholm, Sweden; {dagger} Department of Respiratory Medicine and Allergology and {ddagger} Department of Clinical Virology, Göteborg University, Sahlgrenska University Hospital, Göteborg, Sweden; § Department of Microbiology, Tumor and Cell Biology and Strategic Research Center for Studies of Integrative Recognition in the Immune System, Karolinska Institutet, Stockholm, Sweden; and Department of Experimental and Clinical Medicine, University of Catanzaro "Magna Graecia," Catanzaro, Italy


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
NK and T cells are important for combating CMV infection. Some NK and T cells express leukocyte Ig-like receptor-1 (LIR-1), an inhibitory receptor recognizing MHC class I and the CMV-encoded homolog UL18. We previously demonstrated an early increase in LIR-1-expressing blood lymphocytes in lung-transplanted patients later developing CMV disease. We now show that NK and T cells account for the observed LIR-1 augmentation. Coincubation of PBMC from CMV-seropositive donors with virus-infected lung fibroblasts led to a T cell-dependent secretion of IFN-{gamma}, produced mainly by LIR-1+ T cells and by NK cells. Cytokine production during coculture with fibroblasts infected with virus containing the UL18 gene was augmented compared with the UL18 deletion virus, suggesting a stimulatory role for UL18. However, purified UL18Fc proteins inhibited IFN-{gamma} production of LIR-1+ T cells. We propose that cytokine production in the transplant induces NK and T cells to express LIR-1, which may predispose to CMV disease by MHC/LIR-1-mediated suppression. Although the UL18/LIR-1 interaction could inhibit T cell responses, this unlikely plays a role in response to infected cells. Instead, our data point to an activating role for viral UL18 during infection, where indirect intracellular effects cannot be excluded.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Like other members of the herpes virus family, human CMV (HCMV)3 causes a latent infection that can be reactivated under certain circumstances, such as during immune suppression after lung transplantation. HCMV infection is the single most common viral infection following lung transplantation, affecting more than half of the patients and representing one-third of all posttransplantation infections (1).

NK cells play a major role in combating herpes virus infections both in humans (2, 3) and in mice (4, 5). These cells constitute ~15% of human PBLs, are cytolytic, and have the capacity to produce a number of mainly proinflammatory cytokines. IFN-{gamma} produced by NK and T cells is important to limit murine CMV (MCMV) and HCMV infection (6, 7, 8). The activity of NK cells is dependent on signaling through activating and inhibiting receptors, many of which interact with MHC class I molecules. One family of immune regulating receptors, leukocyte Ig-like receptor (LIR)/immunoglobulin-like transcript (ILT)/monocyte/macrophage Ig-related receptor (MIR), are expressed by NK cells, T cells, B cells, monocytes, and dendritic cells (9, 10). LIR-1/ILT2/CD85j/LILRB1, which contains in its intracellular domain four ITIMs, has a broad recognition pattern for classical MHC class I (HLA-A, -B, -C), and nonclassical MHC class I (HLA-G).

In addition to NK cells, CD8+ T cells and to a lesser extent CD4+ T cells are crucial for control of both MCMV (11) and HCMV infection (12, 13). To evade immune responses, HCMV down-modulates HLA class I molecules on the surface of infected cells thereby reducing T cell recognition (14). Interestingly, the HCMV-encoded MHC class I-like protein UL18 (15) shows a >1000-fold higher affinity to LIR-1 compared with cellular MHC class I molecules (9, 16). The extremely high-binding affinity of UL18 to LIR-1 suggests that the interaction between these molecules may be a strategically important mechanism for the survival of HCMV in its host. Possible relevance for this interaction is shown by a high expression of LIR-1 especially on HCMV-specific T cells (17, 18) and on lymphocytes in lung-transplanted patients before development of CMV disease (19).

Data regarding the impact of UL18 on NK cell function are controversial: the MHC class I homolog seems to inhibit NK cell-mediated lysis in certain experimental settings (20, 21), but not in others (22, 23). Concerning T cells and UL18, there is one report (24) proposing that UL18 stimulates CD8+ T cell killing of infected cells via triggering of LIR-1, while many publications indicate the contrary, i.e., that LIR-1 engagement inhibits T cell function (25, 26, 27, 28).

In this study, we focused on the role of LIR-1 in the immune response to CMV, and potential functions of UL18. We addressed three questions: 1) which subsets of lymphocytes augment the LIR-1 expression previously observed early in lung transplant patients? 2) How is the LIR-1 expression induced–are cytokines involved, is it a consequence of interaction with virus infected cells? 3) Is there a functional consequence of LIR-1 expression in relation to the virally encoded ligand UL18?


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Subjects

Thirteen consecutive patients (median age, 56 (30–65) years) undergoing single lung (n = 11), bilateral lung (n = 1), or heart lung (n = 1) transplantation from March 2000 to February 2001 were studied. For comparison, blood samples from five healthy subjects (median age, 29 (29, 30, 31, 32, 33, 34, 35, 36, 37) years) were collected monthly. CMV disease was defined as symptoms of pneumonitis combined with presence of CMV inclusion bodies in transbronchial biopsy and/or bronchoalveolar lavage (BAL) samples following the criteria for CMV disease of the Fourth International Cytomegalovirus Workshop (29). The patients were divided into those who developed CMV disease during the study (CMVd+) and those who did not (CMVd–). Surveillance bronchoscopy with transbronchial biopsy and BAL was performed at 2, 4, 8, 12, 18, and 24 wk after transplantation. The clinical characterization and treatment of the patients has been described earlier (19).

For in vitro studies of lymphocyte function, buffy coats from healthy blood donors and laboratory personnel were included.

Collection of samples

Blood was drawn before bronchoscopy. PBMC were separated and frozen in FCS plus 10% DMSO at –70°C. BAL was performed by infusing PBS into a segmental middle lobe or lingula bronchus. Cellular components were frozen in FCS plus 10% DMSO at –70° C for later analysis. All cell samples from each patient were thawed and stained at the same time.

Quantitative HCMV PCR analysis

HCMV DNA in serum (copies per milliliter) was quantified using the Roche Cobas Amplicor HCMV Monitor (CMM) system according to the manufacturer’s instructions. One HCMV low-positive control, one HCMV high-positive control, and one HCMV-negative control were included in each experimental run. The limit of detection was 400 copies/ml (30) To avoid effects of between-assay variation, all samples from each patient were run in parallel in the same test session.

Analysis of cell surface molecule expression

Abs used were: PerCP- or allophycocyanin-conjugated anti-CD3 (BD Biosciences), allophycocyanin- or FITC-conjugated anti-CD56 (BD Biosciences), Abs to LIR-1 (M405) (31), UL16 binding protein (ULBP) (M-295), ULBP2 (M-311), ULBP3 (M-551), MHC class I polypeptide-related sequence (MIC)A (M-673), MICB (M-362) (provided by Amgen), ICAM-1 (BD Biosciences), and poliovirus receptor (PVR) (Abcam) in combination with PE-conjugated goat anti-mouse IgG (DakoCytomation). PE-conjugated anti-LIR-1 (HP-F1; Beckman Coulter), PE-conjugated anti-CD69 (DakoCytomation), PE-conjugated anti-Nectin-2 (BD Biosciences), allophycocyanin-conjugated anti-CD4, and anti-CD8 (BD Biosciences), FITC-conjugated (Fab')2 goat anti-human IgG1 (Jackson ImmunoResearch Laboratories Europe) for UL18Fc detection and allophycocyanin-labeled anti-mouse IgG1 (BD Biosciences) for detection of M405 and HP-F1 (anti-LIR-1 mAb, provided by Dr. M. Lopez-Botet, Universitat Pompeu Fabra, Barcelona, Spain). Mouse IgG1 was used as negative control for unconjugated Abs, UL16Fc as control for UL18Fc. For virally induced expression of FcRs, fibroblasts were stained with normal human serum and FITC-conjugated (Fab')2 goat anti-human IgG1. The lymphocyte gate was confirmed by gating on CD3+ cells.

Viruses

HCMV strains used were AD169, the UL18 deletion mutant of AD169 (a gift from H. Browne, University of Cambridge, Cambridge, U.K.) and the clinical strain 4636. Virus was propagated on human embryonic lung (HL) fibroblasts purchased from the Swedish Institute for Infectious Disease Control, batched, and stored at –70°C. Viral titers were determined by plaque assays as described (32). Viral stocks were tested negative for mycoplasma contamination using EZ-PCR Mycoplasma Test Kit (Biological Industries).

HCMV infection of human lung fibroblasts and coincubation with lymphocytes

HL fibroblasts (passages 5–20) were seeded in 24-well plates and HCMV virus (multiplicity of infection (MOI) 1) was added when cells were confluent. The level of infection was compared between viral strains at day 3 and 5 postinfection (p.i.) using an Ab for the viral protein UL44 (clone CCH2; DakoCytomation). The cell surface expression of MHC class I, MICA/B, ULBP1–3, PVR, Nectin-2, ICAM-1, and FcRs was analyzed day 4 p.i. by FACS analysis. NK cells and T cells were purified from healthy control PBMC by negative selection (StemSep and RosetteSep, respectively; StemCell Technologies). HCMV IgG serology was determined by Clinical Microbiology at Karolinska University Hospital using enzyme immunoassay. At day 3 p.i., 5 x 106 PBMC/3,5 x 106 purified T cells/0,5 x 106 NK cells were added with or without UL18Fc or UL16Fc protein 5 µg/ml and supernatant was removed at day 5 p.i. for cytokine analysis by ELISA (IFN-{gamma} analyzed with reagents from MabTech and IFN-{alpha} analyzed with reagents from Bender Medsystems). PBMC were stained for CD3, CD56, LIR-1/CD85j (Beckman Coulter), and CD69.

Alternatively, PBMC, T cells, or NK cells were added in same numbers as above day 3 p.i. and cultures were incubated overnight before addition of GolgiStop (BD Biosciences) at a dilution of 1/1500. After additional 4-h incubation, surface staining for CD3, CD56, and LIR-1/CD85j (Beckman Coulter) was performed. Intracellular staining of IFN-{gamma} was done using Ab from clone 4S.B3 (BD Biosciences). For blockade of LIR-1–UL18 interactions, the purified UL18 mAb 10C7 at 10 µg/ml (hybridoma purchased from American Type Culture Collection) and 5% human serum was added to the fibroblasts 30 min before addition of PBMC.

Stimulation of T cells through CD3 in combination with LIR-1

PBMC or negatively isolated T cells prestimulated overnight with 50 U/ml IL-2 were stimulated with anti-CD3 mAb OKT3 (in-house production) on precoated PolySorp plates (Nunc; 1,5 x 105 cells/well). For costimulation/inhibition, wells were coated with 0.6 µg/ml OKT3 in combination with 10 µg/ml anti-CD28, UL16Fc protein (provided by Amgen) or UL18Fc fusion protein (plasmid provided by D. Cosman, Amgen). UL18Fc protein was produced in 293FT cells (Invitrogen Life Technologies) as described (33). GolgiStop was added after 40 min. Samples were incubated for an additional 4.5 h before surface staining and staining for intracellular IFN-{gamma} was performed as described above.

Statistics

Differences between patients groups were analyzed using Mann-Whitney and changes within patients over time using Wilcoxon signed rank Test. The Student paired t test or ANOVA analysis with subsequent Newman-Keuls multiple comparison test were used as indicated in figure legends. Graphs depict mean ± SEM or mean ± SD, as indicated in figure legends. Statistical significance was set to p < 0.05.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Increased LIR-1 expression on blood NK and T cells before HCMV diagnosis

We previously reported an early increase in the percentage of blood lymphocytes expressing LIR-1 in patients that later developed CMV disease (CMVd+ patients) (19). This augmented LIR-1 expression was evident before detection of CMV by PCR. The herein reported analysis of subsets showed that the increase in LIR-1 expression was most evident on NK cells (CD3–CD56+, Fig. 1a), reaching significant differences between CMVd+ and CMVd– patients early after transplantation (at weeks 4 and 8, p = 0.02 and 0.02, respectively). The increased expression of LIR-1 was also evident on CD3+CD56+ T cells (Fig. 1b, p = 0.02 at weeks 8 and 24) and on CD3+CD56– T cells (Fig. 1c, p = 0,05 at week 2, p = 0,004 at week 8, and p = 0,02 at week 24). We could not detect a change over time in the level of LIR-1 expression on monocytes as measured by median fluorescence intensity (MFI), nor a difference between the patient groups (data not shown).


Figure 1
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  		FIGURE 1. The percentage of blood lymphocytes expressing LIR-1 is increased in CMVd+ patients. a, CD3–CD56+ NK cells; b, CD3+CD56+ T cells; and c, CD3+CD56– T cells from CMVd+ patients (n = 8), CMVd– patients (n = 5), and healthy controls (n = 5) were analyzed for LIR-1 expression. The mean ± SEM percentage of cells expressing LIR-1 is plotted over time. Samples were collected before transplantation (pre) and at weeks 2, 4, 8, 12, 18, and 24 following transplantation in patients and monthly in healthy controls. The percentage of CD3–CD56+ NK cells expressing LIR-1 is shown for individual patients, in (d) CMVd+ patients, and in (e) CMVd– patients.

 
The proportion of T cells and NK cells within the lymphocyte gate did not differ between patient groups and decreased by approximately half and by a factor of seven, respectively, due to induction treatment. The proportion of T cells recovered to normal levels at week 8 (data not shown).

The early increase of the percentage of NK cells expressing LIR-1, from <8% before transplantation to >20% at week 2, was seen in all CMVd+ patients (Fig. 1d) and only in one CMVd– patient (Fig. 1e). This particular CMVd– patient (Fig. 1e, {circ}) was the only CMV-seronegative patient receiving an organ from a sero-positive donor. One patient who developed CMV disease never showed detectable virus levels in serum (Fig. 1d, {circ}). However, also this patient showed an increase in LIR-1 expression. Another patient, who did not develop CMV disease but who had HCMV DNA in serum (380 copies of virus/ml) at week 12 (Fig. 1e, triangle), showed an increase in LIR-1+ cells week 12. Data from before transplantation and weeks 2 and 4 following transplantation are missing for this particular patient because there were not enough lymphocytes for a reliable analysis.

Induction of LIR-1 on NK cells

As factors regulating the expression of LIR-1 on NK cells are unknown, possible mechanisms for the observed induction of LIR-1 in CMVd+ patients were investigated. First, we investigated whether cytokines were produced locally in the lung in CMVd+ patients analyzing BAL fluid by ELISA. We could detect IL-15 in six of eight CMVd+ patients (median: 85, range: 51–308 pg/ml) and in two of five CMVd– patients (51 and 101 pg/ml), but the difference between patient groups did not reach statistical significance. Low cell numbers impeded the investigation of LIR-1 expression on lymphocytes from BAL fluid.

As IL-15 was found in BAL fluid of CMVd+ patients and is important in NK cell proliferation in mouse CMV infection (34), we studied the effect of IL-15 on LIR-1 expression by healthy control PBMC in vitro. We also included IL-2, known to induce LIR-1 expression on activated CD8+ memory T cells (26), and IL-10, because HCMV encodes a IL-10 homolog (35). LIR-1 was found to be predominantly expressed by the CD56dim NK cell subset in peripheral blood and incubation of PBMC with IL-15 or with IL-2 increased the percentage of LIR-1-expressing NK cells by ~20% (data not shown). Also IL-10 slightly affected LIR-1 expression on NK cells, while TGF-beta had no effect (data not shown). In vitro BrdU incorporation indicated that the increased frequency of LIR-1-expressing NK cells upon cytokine stimulation was not due to specific proliferation of LIR-1-expressing cells, but rather induction of LIR-1 on previously LIR-1– NK cells. The percentage of LIR-1+ T cells did not change upon cytokine stimulation (data not shown).

To assess the possibility that virus infected cells can induce LIR-1 up-regulation on NK cells and/or T cells, PBMC from healthy donors were cocultured with uninfected or infected HL fibroblasts from days 3 to 5 p.i. The incubation time was chosen to allow even late viral transcripts like UL18 to be sufficiently expressed while infected fibroblasts were still viable. The clinical strain 4636 was included to detect a possible role of UL18 on LIR-1 regulation, because the 4636 strain encodes a UL18 protein with higher affinity for LIR-1 than AD169 (36). There was no consistent effect on the expression of LIR-1 on NK cells, neither in percentage of cells nor in level of expression (MFI) in this setting, using AD169-infected HL (Fig. 2a) or the clinical strain 4636 (Fig. 2b). Among the donors, four were seropositive for HCMV, and these showed no change or even a decreased percentage of LIR-1-expressing NK cells. NKG2C, NKp44, and NKp46 were other markers that were not significantly changed in the cocultures. Yet, all donors showed a dramatic increase in NK cells positive for the activation marker CD69 when cultured with infected fibroblasts (Fig. 2c). CD69 was also up-regulated on T cells (data not shown). LIR-1 expression on T cells was not affected by the 2-day coculture with virus infected HL, neither the percentage of positive cells nor MFI (data not shown).


Figure 2
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  		FIGURE 2. HCMV does not consistently up-regulate LIR-1 on NK cells, although it always induces high expression of CD69. PBMC (n = 11) were grown with uninfected or infected fibroblasts for 2 days (day 3–5 p.i.). Surface expression of LIR-1 was analyzed on NK cells after incubation with uninfected fibroblasts (HL) or HL infected with AD169 (a) or with the clinical isolate 4636 (b). In c, NK cells from the same cultures are analyzed for the expression of CD69. Each line represents one separate experiment.

 
Reduced cytokine production of PBMCs in response to dUL18 compared with AD169-infected fibroblasts

To investigate the role of LIR-1/UL18 interactions during activation of NK and T cells, we used AD169-infected fibroblasts or fibroblasts infected with a deletion virus for UL18 (dUL18) created on the AD169 background (37). We set up cocultures of PBMC from healthy donors and infected fibroblasts. Supernatants were analyzed by ELISA and were found to contain considerable levels of IFN-{gamma} and IFN-{alpha} (Fig. 3). It is likely that Ag-experienced effector cells participate in the IFN-{gamma} response to HCMV-infected fibroblasts, because seronegative donors, despite having substantial frequencies of LIR-1-expressing T cells, displayed only a minor IFN-{gamma} response (<2 ng/ml, data not shown). This indicates that the IFN-{gamma} response was not solely induced by UL18-LIR-1 interaction as reported for triggering of cytotoxicity in CD8+ T cells (24). Other contributing factors, either TCR-dependent or bystander effects (38) should be involved. Surprisingly, the deletion of UL18 reduced both IFN-{gamma} and IFN-{alpha} levels significantly (Fig. 3a). Virus-infected fibroblasts produced no detectable IFN-{alpha} and uninfected fibroblasts coincubated with PBMC induced IFN-{gamma} levels <1 ng/ml and no detectable IFN-{alpha}.


Figure 3
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  		FIGURE 3. Cytokine production in PBMC is induced by virus-infected fibroblasts and is reduced in the absence of UL18. a, Cytokine detection in supernatants by ELISA. PBMC (n = 17 for IFN-{gamma}, n = 9 for IFN-{alpha}) were cultured with dUL18- or AD169-infected HL day 3–5 p.i. (p = 0,001 for IFN-{gamma}, p = 0,003 for IFN-{alpha}, paired t test). For IFN-{gamma}, only CMV-seropositive individuals were included. b, The role of different cell types in IFN-{gamma} production. ELISA was performed on supernatants of PBMC or separated cells cultured with virus infected HL as indicated, n = 8 for PBMC and T cells, n = 5 for CD14-depleted PBMC, n = 6 for isolated T cells plus NK cells, n = 5 for AD169 and n = 3 for dUL18-isolated NK cells. *, p < 0,05, paired t test. c, The difference in cytokine detection between AD169 and dUL18 is not due to different infection efficiency or MOI. The gray line in the histogram plot represents IgG1 control staining of AD169-infected HL, the black dotted line represents UL44 staining of dUL18-infected HL, and the black solid line UL44 staining of AD169-infected HL. The bar chart summarizes (mean + SD) UL44 staining of AD169- and dUL18-infected HL for three independent experiments.

 
To dissect the role of different cell types for the IFN-{gamma} response to infected fibroblasts, PBMC or separated cultures of either T cells, NK cells, combined NK plus T cells or CD14-depleted PBMC were cultured as above with AD169- or dUL18-infected HL (Fig. 3b). Isolated T cells produced low levels of IFN-{gamma} during culture with AD169-infected HL and significantly less in absence of UL18. A combination of purified NK and T cells augmented levels of IFN-{gamma}, while the trend for less production with dUL18 was maintained. NK cells alone did not produce detectable levels of IFN-{gamma} (Fig. 3b). Monocyte depletion decreased IFN-{gamma} levels compared with PBMC, while the dUL18 trend was still discernable. We conclude that T cells alone can produce IFN-{gamma} in response to virus-infected cells, while monocytes and NK cells augment cytokine production. The reduced ability of dUL18-infected fibroblasts to induce IFN-{gamma} is seen also using isolated T cells, indicating that UL18, or other nearby viral genes, may have an activating effect on T cells.

To ensure that the difference in cytokine response to dUL18 compared with AD169 was not due to different efficiency of infection, several control experiments were performed. Viral titers were determined by plaque assays, infectivity was monitored by microscope, different batches of viruses were used, and virus was titrated to determine the range of MOI where cytokine response was constant (data not shown). Furthermore, infected fibroblasts were stained intracellularly for the virally encoded protein UL44 to measure infection level (Fig. 3c).

Both LIR-1+ and LIR-1– NK cells contribute substantially to IFN-{gamma} production in cocultures

Because the only known receptor for UL18 is LIR-1, we wanted to relate cytokine production to cell type and LIR-1 status. Intracellular staining for IFN-{gamma} was performed on PBMC cocultured with infected fibroblasts (AD169 or dUL18, Fig. 4).


Figure 4
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  		FIGURE 4. NK cells are a major source of IFN-{gamma} and their IFN-{gamma} production is not dependent on the presence of LIR-1. PBMC (5 million/well) were coincubated with HL fibroblasts day 3 p.i. GolgiStop was added after overnight incubation, and intracellular staining for IFN-{gamma} was performed after 4 h. Cells were identified as T cells (CD3+) or NK cells (CD3–CD56+) by surface staining. Data depict mean + SD of six CMV-seropositive blood donors. *, p < 0,05 paired t test.

 
In coculture of bulk PBMC with infected fibroblasts, T cells and NK cells contributed equally in absolute numbers (data not shown) to the IFN-{gamma}+ cell fraction. Among T cells, the majority of IFN-{gamma}-producing cells were LIR-1+ (Fig. 4), in line with previously published observations that couple the expression of LIR-1 to virus-specific T cells with an effector memory phenotype (18, 39). No significant decrease in the percentage of IFN-{gamma}+ T cells in PBMC upon contact with dUL18-infected HL compared with AD169 was detectable in this setup, reflecting a decreased production of IFN-{gamma} per T cell rather than a decreased proportion of T cells responding dUL18 (Fig. 3). However, NK cells of both LIR-1+ and LIR-1– subsets showed a reduced response to dUL18-infected fibroblasts compared with AD169, and LIR-1 status had no impact on IFN-{gamma} production (Fig. 4). Only a very small fraction of isolated T cells stained positive for IFN-{gamma} after contact with infected fibroblasts, and isolated NK cells did not respond at all (data not shown). A combination of NK and T cells resulted surprisingly in IFN-{gamma} production by NK cells only (data not shown), indicating that the IFN-{gamma} produced by NK cells in response to infected cells is T cell dependent and that the IFN-{gamma} detected by ELISA in supernatants of NK+T cells (Fig. 3b) is mostly produced by NK cells.

UL18Fc fusion protein reduces T cell IFN-{gamma} production

We wanted to study the direct effect of UL18 on T cells in an isolated setup using UL18Fc fusion proteins, because PBMC cultured with infected fibroblasts produced less IFN-{gamma} in the absence of UL18 compared with wild-type AD169-infected HL (Fig. 3, a and b), and because IFN-{gamma} production by T but not NK cells was LIR-1 dependent (Fig. 4) and T cells were involved in triggering NK cell responses to infected cells (data not shown).

It was first determined whether the UL18Fc proteins bound exclusively to LIR-1 expressing T cells (Fig. 5a). T cells were stained with UL18Fc and conjugated secondary Ab (Fig. 5a, left panel), with anti-LIR-1/CD85j-PE (Fig. 5a, middle panel) or with the combination of the two (Fig. 5a, right panel). To account for possible Fc binding, UL16Fc was used as control and found not to stain (data not shown). Fig. 5a illustrates that UL18Fc recognized only LIR-1+ T cells, and because staining with both reagents create a diagonal staining profile, it is probable that UL18Fc and the anti-LIR-1 Ab stain the same protein. Furthermore, the data show that the binding of UL18Fc could not block the epitope for this Ab (clone HP-F1).


Figure 5
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  		FIGURE 5. UL18 binds to LIR-1 and inhibits T cell IFN-{gamma} production induced by stimulation via CD3. a, UL18Fc and anti-LIR-1/CD85j-PE clone HP-F1 stain the same T cells. Purified T cells were stained with UL18Fc, detected by FITC-conjugated anti-human IgG1 Ab (Fc{gamma} specific; left panel), anti-LIR-1/CD85j-PE (clone HP-F1; middle), and a combination of both (right panel). Purified T cells (b and c) or PBMC (d) were stimulated overnight with 50 U IL-2/ml and added to wells coated with anti-CD3 (OKT3, 0.6 µg/ml) alone or in combination with anti-CD28 (10 µg/ml), together with either no protein, UL16Fc control protein, or UL18Fc protein (both 10 µg/ml). After 40 min, GolgiStop was added and cells were stained for the surface markers CD3, CD8, LIR-1, and intracellular IFN-{gamma}. b, Representative example of IFN-{gamma} production by CD3+ T cells upon stimulation with OKT3 in combination with UL16Fc or UL18Fc. c, Mean + SD of n = 6 for OKT3-stimulated CD3+ T cells and n = 8 for OKT3 plus anti-CD28-stimulated CD3+ T cells in combination with indicated proteins. Percentage of IFN-{gamma}-positive cells is shown separately for LIR-1-positive and LIR-1-negative cells. d, Mean + SD for PBMC (n = 5) stimulated with OKT3 ± anti-CD28 in combination with indicated proteins. *, p < 0.05; **, p < 0.01; ***, p < 0.001, ANOVA.

 
Intracellular IFN-{gamma} production was measured to assess the impact of UL18Fc binding to LIR-1 during T cell stimulation by anti-CD3 (OKT3) with and without costimulation through CD28 (Fig. 5, b–d). Suboptimal stimulation with OKT3 induced IFN-{gamma} production mainly by LIR-1+ T cells. This response was markedly reduced by addition of UL18Fc, but not UL16Fc (Fig. 5, b and c). Engagement of the costimulatory molecule CD28 resulted in increased IFN-{gamma} production in both LIR-1+ and LIR-1– T cells (Fig. 5c). UL18Fc almost completely blocked also the costimulated induction of IFN-{gamma} production by LIR-1+ T cells in all donors, while there was no such effect by UL16Fc. Similar results were obtained using bulk PBMC from seropositive donors (Fig. 5d). A certain degree of blocking was also seen for UL16Fc and UL18Fc proteins in LIR-1– T cells in some donors. This effect could be a contribution by the Fc portion or unspecific protein competition during coating. It was not possible to block the UL18Fc effect by including the LIR-1 Ab HP-F1 in the assay, because this Ab was found to bind to a different epitope on LIR-1 (Fig. 5a and data not shown).

In our setup, the frequency of LIR-1+ T cells of healthy blood donors was higher on CD8+ T cells (12.7 (4.2–63.5%)) compared with CD4+ T cells (3.2 (0.5–18.5%), p = 0,02), corroborating previous publications (17, 39). Nevertheless, LIR-1+CD4+ T cell constituted a substantial subset in most donors and results obtained when gating on CD8+ or CD8– T cells separately did not differ from results described above (data not shown).

UL18Fc protein decreases IFN-{gamma} production in PBMC-HL cocultures but cannot abrogate the difference between AD169 and dUL18

In line with previous results (40), we were not able to stain by flow cytometry for surface-expressed UL18 on AD169-infected HL using the 10C7 mAb (41), while fibroblasts infected with recombinant vaccinia virus expressing UL18 were clearly stained with this Ab (data not shown). Neither did we observe a difference between AD169- or dUL18-infected HL when staining with LIR-1Fc proteins (data not shown). In contrast, UL18 expression was clearly detected by Western blot using the anti-UL18 mAb M71 (Amgen), either on whole cell lysates of AD169-infected cells (and dUL18 as control) or after immunoprecipitation of cell lysates with 10C7 (data not shown). The lack of detectable surface UL18 on AD169-infected HL argues against a direct interaction between LIR-1 on leukocytes and viral UL18. However, UL18 could be expressed in amounts below detection limit of our method. To further investigate a possible interaction of LIR-1 and viral UL18, we added UL18Fc, or UL16Fc as control, to cocultures of PBMC and HL infected with AD169 or the UL18 deletion virus. The amount of IFN-{gamma} detected in the supernatant of such cocultures was decreased substantially in presence of UL18Fc, but not in presence of UL16Fc, for both viral strains (Fig. 6, a and b). The inhibitory effect of UL18Fc in both viral cultures recapitulates the strong reduction of cytokine production by UL18Fc observed in experiments with OKT3 stimulated T cells (Fig. 5, b and d). It is likely that FcRs expressed on leukocytes and on infected fibroblasts account for cross-linking of UL18Fc bound to LIR-1 in the cocultures. Thus, addition of UL18Fc proteins did not merely block a possible interaction of low-level surface expressed viral UL18 with LIR-1, but mainly led to inhibition by triggering LIR-1. It is noteworthy that despite the general inhibitory effect of UL18Fc proteins in PBMC-HL cocultures, the significant difference in IFN-{gamma} production upon contact with AD169- or dUL18-infected HL persisted (Fig. 6c).


Figure 6
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  		FIGURE 6. UL18Fc addition to PBMC-HL cocultures inhibits cytokine production. PBMC (5 million/well) were cocultured with HL fibroblasts infected with (a) AD169 or (b) dUL18 in presence of UL18Fc or UL16Fc proteins (5 µg/ml) or without added protein (control) day 3–5 p.i. Supernatants were analyzed for IFN-{gamma} by ELISA. Each line represents results from one CMV-seropositive healthy blood donor (n = 6) identified by the same marker in a and b. c, Summary of results for IFN-{gamma} production in response to AD169- and dUL18-infected HL upon addition of UL18Fc protein, mean + SD, n = 6. *, p < 0,05, paired t test.

 
Instead of using UL18Fc, the addition of anti-UL18 Ab 10C7 to the same coculture setup should only disrupt binding of potential surface expressed UL18 and LIR-1. However, the use of 10C7 could not abrogate the difference in cytokine response induced by dUL18 compared with AD169-infected HL (data not shown).

Changes in cell surface molecules on infected HL cannot explain the different cytokine response to AD169- or dUL18-infected cells

Our interpretation of the presented data is that a potential interaction between LIR-1 and UL18 expressed by virally infected cells has at best a minor effect on leukocyte function, while the increased cytokine production seen in response to AD169 infected fibroblasts may be due to some as yet unidentified, possibly intracellular, function of UL18. To test whether UL18 interferes with expression of ligands for NK and T cells on infected fibroblasts, cells were stained for MHC class I, MICA, MICB, ULBP1–3, PVR, Nectin-2, and ICAM-1 (Fig. 7). HCMV infection induced expression of FcRs, ULBP1, 2, 3 (42), MICA, B (43), ICAM-1 (44), PVR and Nectin-2, while class I levels were decreased (45). Expression of ULBP1/2 and Nectin-2 were slightly more elevated on dUL18 compared with AD169-infected fibroblasts, whereas expression levels of the other molecules did not differ between viral strains (Fig. 7). The observed slight increase in expression of activating NKR ligands on dUL18-infected fibroblasts cannot explain the decrease in cytokine induction.


Figure 7
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  		FIGURE 7. Expression of cell surface ligands for NK and T cells does not differ substantially between AD169- and dUL18-infected HL. Representative histograms of at least three independent experiments, displaying surface expression of the indicated markers on fibroblasts infected with AD169 (black solid line), dUL18 (gray dashed line), or uninfected (gray solid line). Cell count is indicated on the y-axis.

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
We have in this study extended our previously published findings of an increased expression of LIR-1 on lymphocytes following lung transplantation in patients that later develop CMV disease (19). We herein report that the increased LIR-1 expression is most clearly seen on NK cells and is also evident on T cells. Coculture with virus infected fibroblasts induced IFN-{gamma} secretion by LIR-1+ T cells and by NK cells irrespective of LIR-1 phenotype. A deletion mutant of viral strain AD169 lacking the LIR-1 ligand UL18 unexpectedly induced less cytokine response in NK and T cells during such coculture. On the contrary, isolated UL18Fc fusion protein could inhibit IFN-{gamma} production of anti-CD3-activated T cells.

Possible explanations for the described LIR-1 induction in lung-transplanted patients include HCMV-dependent and -independent events. Two-day cocultures of PBMC with infected fibroblasts did not up-regulate frequencies (Fig. 3) or expression levels (data not shown) of LIR-1+ NK or T cells, while anti-CD3 stimulation has been shown to enhance numbers of LIR-1+ T cells after 2 days (25). Another study (46) has reported an increase in the proportion of LIR-1-expressing T cells after 10 days coculture with HCMV-infected fibroblasts. This up-regulation was seen also after coculture with fibroblasts infected with UV-irradiated virus or addition of virus without fibroblasts to PBMCs, pointing toward specific T cell activation rather than molecular interactions between LIR-1 and UL18 as reason for induction of LIR-1. In our hands, LIR-1 was only slightly up-regulated on T cells cocultured with infected fibroblasts day 3–11 p.i. compared with uninfected fibroblasts, and LIR-1 expression by NK cells was not consistently affected under these circumstances (data not shown).

In contrast, our results are consistent with the possibility that NK cells are induced to express LIR-1 by exposure to cytokines produced locally in the lung. Although three of the eight CMVd+ patients showed >150 pg IL-15/ml BAL fluid on at least one occasion (and none of the five CMVd– patients), the difference between the groups was not significant. The results therefore do not allow us to conclude whether local IL-15 levels were responsible for the differences in LIR-1 expression and additional factors in the CMVd+ patients (e.g., early undetectable reactivation of HCMV) can play a role. The question thus remains: is the increase in LIR-1 expression the consequence of early HCMV reactivation, or does it precede and predispose to it. In the latter case, one may speculate that the LIR-1 induction can be more prominent in some patients due to genetic- or disease-associated factors, and this early strong LIR-1 up-regulation may subsequently down-modulate immune reactions through cognate MHC ligands and to a limited extend contingently also through interactions with viral UL18. This would then contribute to immunosuppression leading to HCMV reactivation and disease.

To study the direct effect of UL18 on T cells, we used UL18Fc proteins and observed reduced IFN-{gamma} production upon stimulation of T cells via the TCR in the presence of UL18. One previous report shows that UL18 can induce CD8+ T cell lysis of HCMV-infected cells or cells infected with vaccinia virus expressing UL18 in a MHC class I/TCR independent manner (24). Yet the mechanism by which the inhibitory receptor LIR-1 can deliver activating signals remains to be elucidated. In contrast, the same group and others have demonstrated that engagement of LIR-1 can inhibit CD8+ T cell cytokine production (25, 26, 27) and in some (26, 28), but not all (25), settings also cytotoxicity.

However, our data obtained with the UL18 deletion virus argue against the simple notion that UL18 acts to inhibit the immune response. IFN-{gamma} and IFN-{alpha} responses were diminished when PBMC or purified T cells were cocultured with fibroblasts infected with UL18 deletion virus compared with the parental strain AD169, a rather surprising finding considering the inhibitory functions of the UL18R LIR-1 (25, 27, 31, 47, 48). Other studies using dUL18 virus to detect a potential inhibitory effect of UL18 on NK cells also observe rather an enhancement of killing instead of inhibition (22, 23). One possible explanation is that UL18 binds to another unknown receptor on NK or T cells. Costaining of T cells with anti-LIR-1 Ab and UL18Fc protein resulting in an exact overlapping T cell subset (Fig. 5) as well as immunoprecipitations with UL18 and LIR-1 on CD8+ T cells (24) argue against this hypothesis. Yet, UL18 may have other ligands on monocytes or dendritic cells, or alternatively, ligation of LIR-1 on those cells may result in enhanced effector functions of T and NK cells via cell cross-talk. In contrast, as isolated T cells responded similarly compared with T cells in PBMC bulk cultures (Figs. 3b and 5, c and d), the involvement of other LIR-1 expressing cell types in the culture is not a prerequisite. A third possibility is that UL18 interacts intracellularly with either cellular or other viral proteins, leading to an altered expression of surface molecules on infected cells that in turn influence immune responses to HCMV. This scenario could explain why studies on the effect of UL18 on NK cells using UL18 expressed in isolation (20, 21) reach other conclusions compared with studies with HCMV-infected cells (22, 23). The lack of detection of UL18 surface expression on infected fibroblasts by flow cytometry (data not shown and Ref. 40) as well as the failure to abrogate the difference in cytokine response toward AD169 and dUL18 by addition of either UL18Fc proteins (Fig. 6c) or the anti-UL18 Ab 10C7 (data not shown) point toward an intracellular role of UL18.

In summary, our data indicate that the net effect of UL18 expressed in infected cells does not inhibit, but rather activate NK and T cell function. We speculate that UL18 may have as yet unidentified effects regulating immune responses, possibly by inhibition of negative regulatory pathways, leading to a more potent activation of lymphocytes. As to LIR-1, one must also consider the possibility that increased expression can lead to a general inhibition via interaction with MHC class I molecules, thus explaining the observed connection between elevated LIR-1 expression and CMV disease.


   Acknowledgments
 
We thank members of the Klas Kärre and Hans Gustaf Ljunggren groups for fruitful discussions, in particular Alexander Rölle, Adnane Achour, and Hans-Gustaf Ljunggren. Miguel Lopez-Botet is acknowledged for sharing reagents.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by grants from the Swedish Heart-Lung Foundation, Swedish Cancer Society, King Gustaf V 80th Jubilee Fund, Västra Götaland’s Region, and the Swedish Society of Medicine. E.C. was supported by European Union Grant ERBFM-BICT 972110 and by Associazione Italiana per la Ricerca Cancro. Back

2 Address correspondence and reprint requests to Dr. Claudia S. Wagner, Center for Infectious Medicine, F59, Karolinska University Hospital Huddinge, S-141 86 Stockholm, Sweden. E-mail address: claudia.wagner{at}ki.se Back

3 Abbreviations used in this paper: HCMV, human CMV; MCMV, mouse CMV; BAL, bronchoalveolar lavage; HL, human embryonic lung; LIR, leucocyte Ig-like receptor; MOI, multiplicity of infection; p.i., postinfection; MFI, mean fluorescence intensity; MIC, MHC class I polypeptide-related sequence; PVR, poliovirus receptor; ULBP, UL16 binding protein; CMVd+/CMVd–, CMV disease positive/negative. Back

Received for publication August 8, 2006. Accepted for publication December 21, 2006.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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