Notice: Wiley Online Library will be unavailable on Saturday 27th February from 09:00-14:00 GMT / 04:00-09:00 EST / 17:00-22:00 SGT for essential maintenance. Apologies for the inconvenience.
Potential conflict of interest: Nothing to report.
Dendritic cells (DC) are crucially involved in the induction of immune responses; however, reports on DC functions in chronic hepatitis C are controversial. Function of DC includes proper cell trafficking between sites of infection and lympho-cellular compartments. Thus, we analyzed DC compartmentalization and changes in DC migration in hepatitis C virus (HCV)-infected patients. We found significantly lower numbers of circulating BDCA1+ and BDCA2+ DC in HCV(+) patients (n = 20) than in healthy controls (n = 12) (P < .05). Analyzing liver samples from HCV(+) patients (n = 15), HCV(−) controls (n = 15), and disease controls (n = 10), we demonstrated chronic hepatitis C to be associated with intrahepatic DC enrichment (P < .05). In vitro studies indicated that HCV E2-induced secretion of RANTES efficiently attracts CCR5(+) immature DC. Incubation of DC with sera derived from HCV(+) patients made DC unresponsive to CCL21, the chemokine recruiting DC to lymphoid tissues for T cell priming. Unlike attraction of CCR5+ DCs via RANTES, direct inhibition of DC migration in response to CCL21 was specific for patients with chronic hepatitis C and could be attributed to interaction of HCV E2 with CD81 on DC. In conclusion, migration of DC is markedly affected by interaction of HCV E2 with CD81. Failure of DC to recirculate to lymphoid tissue may be critically involved in impaired T cell priming during HCV infection. (HEPATOLOGY 2006;44:945–954.)
If you can't find a tool you're looking for, please click the link at the top of the page to "Go to old article view". Alternatively, view our Knowledge Base articles for additional help. Your feedback is important to us, so please let us know if you have comments or ideas for improvement.
Dendritic cells (DC) are professional antigen-presenting cells (APC) that play a key role in the induction of immune responses. In humans, DC are widely distributed throughout the organism and are crucially involved in the recruitment and activation of T cells, natural killer cells, macrophages, and B cells.1, 2 In general, DC can be divided in two different subsets. Myeloid DC, which represent approximately 0.5% to 1.0% of the peripheral blood mononuclear cells (PBMCs), can be identified on the basis of expression of blood dendritic cell antigen (BDCA) 1.3 Plasmacytoid DC make up 0.2% to 0.5% of PBMCs and can be identified using antibodies specific for BDCA-2.3, 4
Reports on DC in chronic hepatitis C are controversial. Initial studies suggested a functional impairment of DC from hepatitis C virus (HCV)-infected patients.5–7 However, clinically this should result in a generalized immune suppression, which is not observed in hepatitis C. More recent studies proposed a normal functional capacity of circulating dendritic cells in chronic hepatitis C,8–10 which raises the question why priming of a T cell response is ineffective in hepatitis C despite the presence of functional DC.
Until now DC function in hepatitis C was analyzed with respect to induction of allogeneic T cell proliferation and secretion of tumor necrosis cell alpha. However, effective antiviral function of DC also requires correct cell migration and homing to distinct lymphatic compartments. Importantly, inhibition of DC migration by human cytomegalovirus (HCMV) has been proposed as a potential viral strategy to paralyze the host's immune response.11
Dendritic cells display specific trafficking properties. Recruitment of immature DC (imDC) to sites of inflammation is regulated by chemotactic agonists such as the chemokines RANTES, MIP-1α, and MIP-1β, which are produced early in inflammatory tissues. These molecules represent ligands for chemokine receptors, such as CCR1, CCR2, and CCR5, which are expressed on circulating imDC.12 These chemokine receptors direct imDC from the peripheral blood to the sites of inflammation (e.g., the HCV-infected liver), where they can take up antigens. This step is followed by functional maturation of DC, which is accompanied by a dramatic change in chemokine receptor expression and migratory behavior. Mature DC are negative for receptors that bind inflammatory chemokines, such as CCR5, but express high levels of CCR7, the receptor for the chemokines CCL19 and CCL21, which are expressed in lymphoid organs. Thus, upregulation of CCR7 is crucial for homing of mature DC to T cell areas of lymphoid tissues, where priming of antigen-specific T cells takes place.13–16
CD81, a member of the tetraspanin family, is considered a putative cellular receptor for entry of the hepatitis C virus,17 and ligation of CD81 with HCV-E2 has been demonstrated to alter cellular functions of T cells and natural killer cells.18–22 In addition, Mantegazza and colleagues23 recently demonstrated that tetraspanins also may be importantly involved in the regulation of DC migration.23
Therefore, we put forward the hypothesis that HCV can interfere with the migratory capacity of dendritic cells by induction of chemokine secretion or directed binding of HCV E2 to CD81 on DC. To test this hypothesis, we analyzed compartmentalization of DC in chronic hepatitis C and studied HCV-induced alteration of DC migration.
For enumeration of circulating DC, 57 persons, all white individuals from the Bonn area in Germany, were enrolled into this study, including 26 patients with chronic hepatitis C, 21 subjects who cleared the virus under antiviral combination therapy with pegylated interferon 2α (180μg) and ribavirin (800-1200 mg) for 6 to12 months, depending on the HCV genotype, and 10 healthy HCV RNA-negative persons (Table 1).
The study conformed with the guidelines of the Declaration of Helsinki as approved by the ethics committee of Bonn University.
Cell Separation and Generation of Monocyte-Derived Dendritic Cells.
CD8+ T cells were separated from total PBMC using MACS cell separation kits following the manufacturer's recommendations (Miltenyi Biotec, Bergisch Gladbach, Germany). Monocyte-derived dendritic cells (MODC) were prepared as described before.24 In brief, PBMC were allowed to adhere in six-well plates at a density of 5 × 106 cells/mL for 1 hour at 37°C in RPMI-1640 with 10% autologous, heat-inactivated serum. Non-adherent cells were removed, and the adherent cells were cultured in RPMI-1640 with autologous, heat-inactivated serum, 750 U/mL human granulocyte macrophage–colony-stimulating factor and 500 U/mL human IL-4 per well for 7 days. Maturation of MODC was induced by culturing the cells for 24 hours in the presence of 100 ng/mL lipopolysaccharide (Sigma, St. Louis, MO).
FACS analysis was performed according to standard methods using the following antibodies: anti-CCR1, anti-CCR5, and anti-CCR7 (all from R&D Systems, Minneapolis, MN), anti-CD40, anti-CD80, anti-CD83, anti-CD86, anti-HLA-DR, and anti-MHC I (all from BD Biosciences, San Jose, CA). DC subsets were identified using antibodies specific for BDCA-2, which is strictly expressed on human plasmacytoid DC,3, 4 and BDCA-1, which is expressed on a major subpopulation of human mature DC.3 Both monoclonal antibodies (MAbs) were obtained from Miltenyi Biotech.
Surface expression of integrins was analyzed using anti-CD29 (β1 Integrin), -CD18 (Integrin β2), –CD11a (LFAα, α L Integrin chain), -CD11c (αX intergin), CD49c (α3 Integrin), CD49d (α4 Integrin), CD49f (α6 Integrin) (all from BD Biosciences). VCAM-1 expression was studied with anti-CD54.
Samples were analyzed on a FACScalibur flowcytometer, using the CellQuest software package (Becton Dickinson, Franklin Lakes, NJ).
Tissue Specimens and Immunohistochemistry.
Liver tissue specimens from chronic hepatitis C were obtained during routine diagnostic liver biopsies with a 1.4-mm Menghini needle after informed consent from the patients (n = 15). Normal liver specimens (n = 10) were taken from unaffected areas of liver resections for secondary hepatic malignancy and from cadaveric donors at the time of hepatectomy for liver transplantation.
Liver tissue was immediately embedded in Tissue Tek OCT compound (Miles Laboratories, Naperwille, IL) and snap-frozen in liquid nitrogen. Frozen tissue was kept at −80°C until examined. Sections of 5 to 7 μm were stained by an indirect immunoperoxidase technique using the following the DC-specific antibodies: anti-BDCA-1 and anti-BDCA2 (Miltenyi, Bergisch Gladbach, Germany).
In brief, endogenous peroxidase activity was blocked by 0.03% H2O2/NaNO3 (peroxidase blocking reagent, Dako, Carpinteria, CA). The sections were incubated with the respective specific primary antibody in phosphate-buffered saline (PBS)/1% fetal calf serum (FCS) in a moist chamber at room temperature for 90 minutes. After washing in PBS, peroxidase-coupled goat anti-mouse antibody (Dianova, Hamburg, Germany) was applied for 30 minutes. Bound antibody was detected with 3-amino-9-ethylcarbazole (Sigma Chemicals, Munich, Germany). All sections were then counterstained with Mayer's hemalum-eosin. Quantitative analysis was performed by manually counting BDCA-1(+) and BDCA-2(+) cells, respectively, in 20 visual fields at 200× magnification for each liver sample.
Flowcytometric Analysis of Cells in the Liver Specimens.
Liver biopsy specimens for FACS analysis were obtained from liver biopsies of five HCV RNA(+) patients using 1.5-mm diameter disposable biopsy needles. Fresh liver samples were washed twice in fresh medium and shaken gently to avoid blood contamination. Liver specimens were disrupted mechanically into small fragments in RPMI 1640 medium with 10% FCS using a forceps and scalpel. Then the fragments were homogenized on a cell strainer (BD Labware). The resulting cell suspension was washed and resuspended in RPMI 1640 medium. Intrahepatic dendritic cells were then analyzed by flowcytometry as described.
Cell Stimulation Assay.
PBMC of five healthy donors (three men, two woman; mean age, 30 ± 6 years) were isolated from peripheral blood by Ficoll–Paque density gradient centrifugation following standard protocols. Cell subpopulations were isolated as described. Anti-CD81 (clone JS-81) was obtained by PharMingen (San Diego, CA). The HCV E2-specific antibody 291A2, recombinant full-length HCV E2 protein, which was expressed in CHO cells and has a physiological glycosylation pattern, and recombinant full-length HCV core protein were a gift of Dr M. Houghton (Chiron Corp., Emeryville, CA). Immobilization of antibodies/protein on 96-well plates (no. 655161; Greiner, Frickenhausen, Germany) was performed as previously described.20 In brief, MAb was diluted in carbonate buffer (15 mmol/L Na2CO3, 35 mm NaHCO3, pH 9.6) and incubated overnight at 4°C. Next, plates were washed and saturated for 30 minutes at 37°C using FCS before the addition of cells. To immobilize recombinant HCV E2 protein, plates were first incubated with anti-HCV E2. After washing with PBS and saturation of the remaining free binding sites, HCV E2 was added and plates were incubated for 60 minutes at 37°C. After further washing, cells (PBMC or purified subpopulations) were added in complete medium.
Cell Migration Assay.
To analyze cell migration, 6 × 105 MODC were seeded into the upper compartment of a 5-μm nitrocellulose filter micro chamber system (Neuroprobes, Gaithersburg, MD) after incubation with or without immobilized HCV E2 for 6 hours. The lower compartment was filled with 300 μL RPMI 1640 medium containing either CCL21 (30 ng) or RANTES (25 ng) (PromoCell, Heidelberg, Germany) or no chemotactic supplement. Cells were then allowed to migrate through the filter at 37°C. After 5 hours, migrated cells were harvested from the lower compartment of the chemotaxis chamber and analyzed by flow cytometry. Migration indices were calculated as follows: MI = (number of cells migrating toward supernatants from stimulation experiments)/(number of cells migrating towards medium alone). Alternatively, MODC were cultured in the presence of 100 μL sera derived from HCV(+) or HBV(+) patients, subjects with primary sclerosing cholangitis (PSC), or healthy controls before performing migration assays.
Determination of Chemokine Levels.
Supernatants were collected in the stimulation experiments and levels of RANTES were determined by commercial ELISA according to the manufacturer's instructions (R&D Systems, Wiesbaden, Germany). Results are given as increase of chemokine release relative to control experiments without HCV E2.
Statistical analysis was performed with the Mann-Whitney U test. A P-value <.05 was considered to be statistically significant.
Quantitative Analysis of BDCA-1 and BDCA-2 Positive Cells in Peripheral Blood and Liver Tissue.
We found a significantly lower proportion of circulating BDCA-1 expressing DC in HCV-infected patients as compared with healthy individuals (0.6 ± 0.08 vs. 1.2 ± 0.16%, P < .01). Similarly, HCV-infected patients had a significantly lower proportion of BDCA-2 expressing dendritic cells as compared with healthy subjects (0.33% ± 0.03% vs. 0.53% ± 0.11%, P < .05). Interestingly, patients who became HCV RNA(−) under combined antiviral therapy with interferon-alpha and ribavirin displayed BDCA-1 (1.09% ± 0.12%) and BDCA-2 (0.52% ± 0.9%) expression, which was identical to that found in healthy individuals (Fig. 1A). Figure 1B exemplifies that, unlike peripheral blood, numbers of BDCA-1+ and BDCA-2+ DC were markedly increased in the livers of patients with chronic hepatitis C as compared with only a few positive cells in our normal control specimens. When the differences in intrahepatic DC between patients with chronic hepatitis C and normal control specimens were analyzed in a quantitative manner, both the increases in BDCA-1+ (20.9 ± 6.8 vs. 6.9 ± 2.3 cells per visual field, mean ± SEM; P = .036) and BDCA-2+ (10.2 ± 2.7 vs. 0.6 ± 0.3 cells per visual field; P = .03) DC were found to be highly significant. Intrahepatic accumulation of BDCA1(+) and BDCA2(+) dendritic cells was also seen in non–HCV-related hepatic diseases such as chronic hepatitis B (BDCA-1+ DC: 46.9 ± 3.0; BDCA-2+ DC: 12.4 ± 2.9) and PSC (BDCA-1+ DC: 6.8 ± 3.0; BDCA-2+ DC: 6.8 ± 3.0) (Fig. 1C). No statistically significant differences were found with respect to expression of maturation markers (CD40, CD86, and HLA-DR).
Unfortunately, intrahepatic dendritic cells could not be analyzed in patients who became HCV RNA(−) under therapy because routine biopsies at the end of antiviral therapy are not performed in our department.
In Vitro Interaction of HCV E2 With CD8+ T Cells Recruits Immature DC Attributable to Release of RANTES.
Recently, we showed that incubation of circulating PBMC with HCV E2 results in enhanced secretion of RANTES, the natural ligand of CCR5.25 Importantly, we found that HCV E2 stimulation of intrahepatic PBMC induced a dose-dependent secretion of RANTES, comparable to that of circulating PBMC (Fig. 2A). As dendritic cells migrate toward increasing amounts of CC chemokines, we speculated that interactions of HCV E2 with CD8(+) T lymphocytes should attract DC.
Figure 2B illustrates migration of monocyte-derived dendritic cells toward supernatant of CD8(+) T cells incubated with or without immobilized recombinant HCV E2. We found that markedly more DC were attracted by the supernatant obtained from HCV E2–stimulated cells as compared with the supernatants of cells cultured in the absence of HCV E2.
Incubation of DC with the supernatant obtained from E2-stimulated CD8+ T cells resulted in downregulation of CCR5 (Fig. 3A), due to internalization of the receptor after ligand binding. In line with this in vitro observation, we found reduced in vivo expression of CCR5 on intrahepatic BDCA1+ and BDCA2+ cells as compared with circulating DC (Fig. 3B).
Stimulation of DC With HCV E2 Results in Reduced Responsiveness to CCL21 In Vitro.
Recent reports demonstrated that tetraspanins delay migration of tumor cells and DC. Because MODC bear high levels of CD81 (Fig. 4A), we analyzed whether binding of HCV E2 to CD81 on DC interferes with DC migration. Thus, we performed a chemotactic assay using the lymphoid chemokine CCL21 as chemoattractant. We observed that overnight incubation of maturated DC with immobilized anti-CD81 resulted in a dramatically reduced responsiveness of the cells to CCL21 (Fig. 4B). Likewise, migration of mature dendritic cells was markedly reduced when DC were stimulated with immobilized HCV-E2 protein (Fig. 4C). This effect could be blocked by pre-incubation of DC with a soluble CD81-specific MAb before incubation with plate-bound HCV E2 (Fig. 4D). Specificity of altered DC migration with respect to the HCV E2/CD81 interaction was further confirmed by the fact that recombinant full-length HCV core protein had no effect on the migratory responses of DC toward CCL21 (data not shown). To analyze the in vivo relevance of HCV-mediated inhibition of DC migration, we incubated MODC with sera derived from patients with chronic hepatitis B, chronic hepatitis C, primary sclerosing cholangitis, or healthy controls in vitro. As shown in Fig. 4E, we found specifically that DC cultured in the presence of HCV-derived sera showed a markedly reduced migration toward CCL21 as compared with DC incubated with sera from healthy subjects. Specificity of this observation was confirmed by the fact that neither sera from HBV(+) patients nor sera from PSC patients reduced DC trafficking.
To test whether HCV E2 interferes with maturation of DC, we analyzed markers of DC maturation (i.e., CD40, CD80, CD83, CD86, MHC I, and HLA-DR) on MODC that had been stimulated with lipopolysaccharide at day 5 of culture in either the absence or presence of immobilized HCV E2 overnight. As illustrated in Fig. 5A, exposure to immobilized HCV E2 did not alter expression of these markers, excluding any obvious effects of HCV E2 on maturation of DC. Likewise, we did not observe any HCV E2–mediated effects on the expression of chemokine receptors CCR5 and CCR7. Most of the functions associated with tetraspanins relate to their ability to facilitate interactions with other proteins generating functional complexes. Integrins are frequently associated in such complexes and in particular have been demonstrated to be involved in tetraspanin-mediated regulation of DC migration.23 Thus, we analyzed a panel of integrins with respect to altered surface expression after CD81 cross-linking. CD29 was the only component, of which the expression was slightly increased on HCV E2-stimulated MODC as compared with unstimulated MODC, whereas surface expression of neither the integrins CD11a, CD11c, CD18, CD49c, CD49d, and CD49f nor of CD54 was not affected by HCV E2 (Fig. 6A-B).
Infection with HCV often results in chronic disease, indicating an inefficient immune response. In this context, several authors proposed that defective function of DC in HCV-infected patients might result in insufficient priming of T cell responses as a potential mechanism contributing to HCV-specific immune dysfunction.5–7 In contrast to this hypothesis, more recent studies failed in demonstrating functional differences in DC from patients with chronic hepatitis C as compared with DC from healthy controls in terms of lipopolysaccharide-induced secretion of interferon gamma and tumor necrosis factor alpha, as well as their allostimulatory capacity.8–10
However, to efficiently fulfill their function as antigen-presenting cells, DCs must migrate from the peripheral blood to the infected liver to take up antigens followed by the subsequent migration of antigen-loaded DC to the lymphoid organs, where they present the processed antigens to T cells.
Because DC migration has not yet been studied in chronic hepatitis C, we analyzed migratory behavior and compartmentalization of DC in chronic hepatitis C.
In line with previous reports, we observed significantly reduced numbers of circulating DC in chronic hepatitis C as compared with HCV-negative controls.26, 27 This might reflect intrahepatic accumulation of dendritic cells during chronic hepatitis C, as we found significantly increased numbers of BDCA-1 and BDCA2-positive DCs in HCV-infected livers.
In line with this hypothesis, we found that HCV E2–mediated RANTES secretion of CD8+ T cells was capable of recruiting immature CCR5+ dendritic cells in in vitro migration assays. Because chronic hepatitis C is associated with increased intrahepatic levels of T cells and T cell–secreted chemokines such as RANTES, this mechanism might also be operative in vivo.28–30 Thus, accumulation of DC in livers of chronically HCV-infected patients can in part be explained by hepatic recruitment of circulating CCR5-expressing DC because of increased intrahepatic levels of inflammatory chemokines, including RANTES. However, other inflammatory hepatic diseases such as chronic HBV infection and PSC also display enhanced intrahepatic levels of chemoattractive cytokines/chemokines, which result in intrahepatic recruitment and accumulation of DC. In line with this suggestion, we found enhanced numbers of BDCA1+ and BDA2+ DC also in non–HCV-related hepatic diseases.
After recruitment to areas of inflammation and uptake of antigens, DC have to migrate to the secondary lymph organs to prime antigen-specific T cells. HCV is a hepatotropic virus. Thus, DC encounter large amounts of viral antigens within the liver, the site of infection. Confirming previous results by other groups,23 we found that DC express CD81, a tetraspanin considered to act as a cellular binding receptor for hepatitis C virions. Tetraspanins have been proposed to be importantly involved in the regulation of cell morphology and cell motility, possibly also comprising DC trafficking. Thus, we analyzed the effects of HCV binding to CD81 on the migratory behavior of DC. In vitro studies demonstrated that cross-linking of CD81 with HCV E2 resulted in impaired migration of DC towards the chemokine CCL21, which regulates trafficking of DC from the sites of infection to the lymphoid organs. Importantly, the specificity of this inhibitory effect was confirmed by the fact that inhibition was blocked by pre-incubation of DC with anti-CD81 and the observation that inhibition was not induced by recombinant HCV core protein, which does not bind to CD81. Importantly, inhibition of DC migrations toward CCL21 was a specific finding in hepatitis C because incubation of DC with sera derived from HCV RNA(+) patients resulted in impaired DC trafficking, whereas incubation of DCs with sera from patients with chronic hepatitis B or primary sclerosing cholangitis did not. Thus, our in vitro results suggest that recruitment of circulating CCR5+ DCs into the liver owing to enhanced intrahepatic levels of cytokines/chemokines is a general feature of inflammatory hepatic diseases, whereas inhibition of DC re-migration toward lymphoid tissues seems to be a specific feature of HCV infection.
Inhibition of DC migration by HCV resembles findings in human cytomegalovirus (HCMV) infection. Moutaftsi and colleagues31 recently demonstrated impaired lymphoid chemokine-mediated migration of HCVM-infected DC, and Varani et al.11 observed reduced migration of HCMV-infected immature DC in response to inflammatory chemokines. Thus, inhibition of DC migration by HCMV has been proposed as a potential viral strategy to paralyze the host's immune response. The observed changes in migratory behavior after HCMV infection could be attributed to an altered surface expression of CCR7 and CCR5, respectively. However, this mechanism is unlikely to operate in hepatitis C; we did not observe any differences in CCR5 or CCR7 expression in DC when they were cultured with or without HCV E2 proteins. Furthermore, impaired migration of DC in vitro could not be attributed to defects in maturation because cross-linking of CD81 did not affect surface expression of CD40, CD80, CD83, CD86, MHC I, or HLA-DR, which are common markers associated with DC maturation.
Mantegazza and colleagues23 recently showed that tetraspanins such as CD63 regulate DC migration in response to inflammatory chemokines, which was associated with an altered surface expression of the integrins CD29, CD11b, and CD18. They proposed that tetraspanins may modify motility of DC by altering the function/expression of integrins associated with the multimolecular tetraspanin complexes. In this context, CD81 can form complexes with α3β1, α4β1, α6β1, and other integrins,32, 33 and changes in motility induced by α6β1 have been proposed to be mediated via a mechanism involving the integrin-associated protein CD81.34 Thus far, however, we only observed a minute increase in the expression of CD29 (β1-integrin) but none of the other studied integrins (α3 (CD49c)-, α4(CD49d)-, and α6 (CD49f)), when MODC were incubated with HCV E2. Furthermore, Domanico et al.34 have demonstrated that anti-CD81 strongly inhibited cell migration, although cell adhesion was not affected. Thus, it remains unclear at present which signaling molecules are involved in CD81-mediated inhibition of DC migration.
Taking into account that DC and Langerhans cells use ICAM-1 (CD54) for migration and entry into lymph nodes,35 we also studied expression of ICAM-1 but failed to detect any effects on CD54 expression after stimulation of CD81 with HCV E2.
In our studies, we used recombinant HCV E2 protein, which represent a frequently used substitute to study binding of viral particles to cellular receptors.18, 20, 21
As discussed elsewhere, likely intact HCV particles can cross-link CD81 by binding to cells in a similar manner as was demonstrated for recombinant HCV E2 in our experimental system.21 In particular, HCV envelope proteins E1 and E2 form heterodimers that are thought to be present on the virus surface in an ordered array of hundreds of molecules.21 Thus, our in vitro observation may reflect a mechanism, which is likely also to operate in natural HCV infection. This suggestion is supported by the fact that incubation of MODC with containing hepatitis C virions also resulted in impaired DC migration toward CCL21. Of note, this effect was HCV-specific because culturing DC in the presence of sera derived from HBV-infected patients or subjects with PSC did not alter trafficking.
Taken together, our findings are consistent with the following model of HCV E2–mediated alterations of DC migration in chronic hepatitis C: within the HCV-infected liver, HCV E2 binds to CD8+ T cells via CD81, resulting in release of RANTES. After the gradient of attracting CC-chemokines such as RANTES, CCR5+ immature DC migrate into the liver. There, intrahepatic binding of RANTES to CCR5 results in internalization of the CCR5 receptor. Direct interaction of CD81 with the HCV E2 protein makes DC unresponsive to the chemokine CCL21 expressed in lymphoid tissues, involving a hitherto unknown mechanism (Fig. 7).
Dysregulation of DC migration in hepatitis C may have important clinical implications, because DC become trapped within the infected liver. This may prevent their migration to lymphoid organs to fulfill their function as antigen-presenting cells priming antigen-specific cellular immune responses. Conversely, this mechanism might represent an interesting target for therapeutic interventions because blocking of HCV E2-CD81 interactions might restore migratory capacity of DC.