Liver-Resident CD103⁺ Dendritic Cells Prime Antiviral CD8⁺ T Cells In Situ

The liver maintains a tolerogenic environment as a result of continuous exposure to bacterial constituents and food-derived Ags (1). As a result, hepatic microbial pathogens, such as HCV and malaria, often establish persistent infection in the liver (2). Impaired T cell responses were reported to be associated with persistence of hepatotropic pathogens; however, little is known about the cellular and molecular basis for the immunoregulatory mechanisms linked to this persistence (3). Moreover, next-generation vaccine design must focus on inducing robust and durable T cell responses to clear pathogens from the liver. Adenovirus (Ad) has been considered for use as a vector for vaccines, because the recombinant Ad system is useful for delivery of Ag in vivo to generate vigorous Ag-specific immune responses (4). Thus, detailed analysis of the T cell responses to Ad infection in the liver may prove helpful for better vaccine design and efficacy.

To generate CD8⁺ T cells capable of clearing virus, naive CD8⁺ T cells must first come in contact with specialized APCs that have both processed viral Ag for MHC class I (MHC-I) presentation and upregulated expression of costimulatory molecules (5, 6). Activation of naive CD8⁺ T cells by APCs of hematopoietic origin (e.g., dendritic cells [DCs]) in response to local infections typically occurs in secondary lymphoid organs draining the sites of pathogen entry and its replication (7). However, in contrast to mucosal tissues (i.e., skin, lung, and gut), the contributing APCs and the location for CD8⁺ T cell activation during hepatic viral infections are poorly understood. Antiviral CD8⁺ T cells with competent effector activities (e.g., IFN-γ, granzyme) are generated in response to hepatic viral challenges (8, 9). Ag presentation by nonhematopoietic parenchymal cells (e.g., liver sinusoidal endothelial cell) was demonstrated to prime naive CD8⁺ T cells, although it resulted in activated T cells with poor effector functions (10–13). Thus, the cell type(s) responsible for initiating the functional hepatic CD8⁺ T cell response to viral infections remains unknown.

Conventional DCs are highly immunogenic APCs with the full capacity to capture, process, and present Ags to naive CD8⁺ T cells (14, 15). Immature DCs use broadly conserved pattern recognition receptors (such as TLRs) to become activated (16–18). Upon activation/maturation, they upregulate Ag-presenting molecules (i.e., MHC-I and MHC class II [MHC-II]) and costimulatory molecules (i.e., CD80 and CD86) (19). Lymph node (LN)-resident DCs, which are commonly divided into CD11b⁺ or CD8α⁺ subsets, express high levels of MHC-II and costimulatory molecules upon activation and serve as potent APCs (7, 20). Conversely, CD11b and CD103 expression demarcate nonlymphoid tissue DC subsets, with both populations primarily using CCR7 for migration out of the parenchyma via the lymphatic conduit system to enter the regional LNs (21, 22). Lymphoid tissue–resident CD8α⁺ and nonlymphoid tissue–resident CD103⁺ DCs play crucial roles in priming CD8⁺ T cells, because they are functionally specialized in MHC-I–restricted cross-presenting Ag (23, 24). These DC subsets also share a developmental pathway requiring the transcriptional factor Batf3 (25). Although the liver is believed to have DCs that patrol the tissue, the characterization of liver-resident DC subsets in a steady-state and their contribution to antiviral CD8⁺ T priming have yet to be investigated.

In this study, we sought to determine the primary APC regulating antiviral CD8⁺ T cell activation and differentiation in response to liver infections. Using the hepatotropic Ad engineered to express OVA (Ad-OVA) and the OT-I (OVA-specific) TCR-transgenic CD8⁺ T cell adoptive-transfer model, we demonstrate that liver-resident CD103⁺ DCs undergo maturation following Ad-OVA infection and are the major APCs capable of driving extensive CD8⁺ T cell proliferation and effector differentiation following infection. Thus, our study identifies the hepatic CD103⁺ DC subset as the primary cell type that establishes productive antiviral CD8⁺ T responses in situ to pathogens invading the liver. The implications of these findings for the design of vaccine strategies and therapeutic venues for chronic hepatic viral infections (e.g., hepatitis C virus [HCV]) are discussed (26–28).

All experiments used 8–12-wk-old gender/aged-matched male and female mice. Thy1.2⁺ C57BL/6 (B6; H-2^b) mice and Thy1.1⁺ OT-I TCR-transgenic (OT-I; H-2^b) mice were purchased from Taconic Farms (Hudson, NY). CCR7^−/− (H-2^b) (29) mice were purchased from The Jackson Laboratory (Bar Harbor, ME). Batf3^−/− (H-2^b) (25) mice were kindly provided by Dr. Thomas Braciale (University of Virginia). All mice were housed in a pathogen-free facility and were tested routinely for mouse hepatitis virus and other pathogens. All mice were handled according to protocols approved by the University of Virginia Institutional Animal Care and Use Committee.

Replication-deficient recombinant Ad type 5 expressing OVA under the human CMV promoter and lacking E1 and E3 genes was purchased from the Iowa Gene Transfer Vector Core (Iowa City, IA). Recombinant murine CMV expressing OVA (MCMV-OVA) was gifted from Dr. Ann B. Hill (Oregon Health and Science University, Portland, OR). Mice were injected i.v. with 2.5 × 10⁷ IU Ad-OVA or 1 × 10⁴ PFU MCMV-OVA.

Livers were perfused with PBS via the portal vein until fully blanched and then put on ice in IMDM supplemented with 10% newborn calf serum. Whole livers were passed through a metal spleen screen and digested with 0.05% Collagenase IV (Sigma-Aldrich, St. Louis, MO) for 30 min at 37°C. Intrahepatic mononuclear cells were purified on a 21% Histodenz (Sigma-Aldrich) gradient after centrifugation at 1250 × g for 20 min without braking. Spleens and LNs were passed through a mesh spleen screen, followed by RBC lysis. All samples were washed and resuspended in IMDM plus serum, and leukocytes were counted via a hemocytometer.

Cells were labeled with Abs against CD45, B220, I-A/I-E, CD11c, CD11b, CD8α, CD103, CD80, CD86, PD-L1, H2-D^k, MHC-I^SIINFEKL, Thy1.1, CD25, CD69, IFN-γ, and granzyme B (all from eBioscience, San Diego, CA). For cell surface labeling, 1 × 10⁶ cells were blocked with anti-CD16/CD32 (2.4G2; University of Virginia) and incubated with the corresponding Abs for 30 min at 4°C in staining buffer (PBS with 2% FBS and 0.1% NaN₃). For cytokine staining, 1 × 10⁶ cells were incubated for 5 h in IMDM supplemented with 10% FBS, 10 U/ml penicillin G, 2 mM l-glutamine, 5 mM 2-ME, and 1 μl/ml GolgiPlug/GolgiStop (BD Biosciences). OT-I cells were restimulated with 2 μg/ml SIINFEKL peptide (AnaSpec). After incubation, the cells were surface labeled as described above and fixed using Cytofix/Cytoperm (BD Biosciences), according to the manufacturer’s instructions, prior to intracellular IFN-γ staining. All samples were run on a BD FACSCanto II (BD Immunocytometry Systems, San Jose, CA) and analyzed using FlowJo software 8.8.6 (TreeStar, Ashland, OR).

For fluorescence microscopy, mouse liver tissues were perfused with 1× PBS and periodate-lysine-paraformaldehyde fixative, excised, incubated in periodate-lysine-paraformaldehyde for 3 h at 4°C, and passed over a sucrose gradient. Tissues were frozen in OCT medium, sectioned at 5 μm thickness, blocked with 2.4G2 solution (2.4G2 supernatant containing anti-CD16/32, 10% each of chicken, donkey and horse serum, and 0.1% NaN₃), and stained with Abs from BioLegend, eBioscience, Santa Cruz Biotechnology, and Sino Biological (Beijing, China). Confocal microscopy was performed on a Zeiss LSM-700, and data were analyzed using Zen 2009 Light Edition software (Carl Zeiss MicroImaging, Jena, Germany). For histology, mouse liver tissues were perfused with 1× PBS, excised, incubated in 10% buffered formalin acetate (Fisher Scientific) overnight at 4°C, washed with 70% ethanol, and embedded in paraffin wax. Tissues were sectioned at 5 μm thickness and stained with H&E. Light microscopy was performed on an Olympus BX51 microscope.

CD8⁺ T cells were isolated from the spleens and mesenteric LNs of Thy1.1⁺OT-I⁺ mice using anti-CD8α Ab–conjugated magnetic bead separation kits (Miltenyi Biotec). Cells were labeled with 1.8 μM CFSE for 8 min at room temperature and transferred by i.v. injection into naive Thy1.2⁺ recipients. In the experiments that used splenectomized mice, recipients were treated i.v. with either 150 μg IgG or MEL-14 (anti-CD62L) 1 d prior to i.v. transfer of CFSE-labeled OT-I T cells. One day post–OT-I T cell transfer, the mice were infected i.v. with 2.5 × 10⁷ IU Ad-OVA.

Liver DCs were isolated from B6 mice (n = 7–10) that were either left uninfected or infected with 2.5 × 10⁷ IU Ad-OVA for 12 h prior to analyses. CD11c⁺ cells were first enriched by a positive selection using magnetic bead–based purification kits (Miltenyi Biotec). DCs were stained using Abs for CD45, MHC-II, CD11c, B220, CD11b, and CD103 and sorted by flow cytometry using a BD FACSVantage SE sorter at the Flow Cytometry Core Facility (University of Virginia). DCs were identified by staining with a mixture of Abs against CD45, MHC-II, CD11c, CD11b, and CD103; plasmacytoid DCs (pDCs) were determined by staining with Abs against CD45, MHC-II, CD11c, and B220.

FACS-sorted liver DCs were pulsed with OVA (SIINFEKL) or β-gal (ICPMYARV) peptides at 0, 25, or 500 ng/ml for 20 min at 37°C and washed. DCs (4 × 10³) and CFSE-labeled OT-I T cells (4 × 10⁴) were cocultured for 4 d at 37°C in vitro. FACS-sorted liver DCs isolated 12 h postinfection (hpi) from livers of infected B6 mice were cultured with CFSE-labeled OT-I T cells at a ratio of 1:10 for 4 d at 37°C in vitro. All DC/T cell coculture media included IMDM supplemented with 10% heat-inactivated FBS (HyClone), 10 U/ml penicillin G, 2 mM l-glutamine, 5 mM 2-ME, 20 mM HEPES, and 100 μg/ml gentamicin (all from Invitrogen).

Student t tests (two-tailed) were used to evaluate the significance of the differences. A p value < 0.05 was regarded as statistically significant.

DCs are represented widely in both lymphoid (i.e., spleen, LNs) and nonlymphoid (i.e., lung, skin) tissues. Lymphoid DCs, such as CD11c^hiMHC-II^hiB220⁻ splenic DCs, are conventionally defined by the surface expression of CD8α or CD11b molecules. In contrast, a distinct DC subset in nonlymphoid tissues, such as the lung, expresses CD103 (αE integrin, a ligand for E-cadherin) rather than CD8α, whereas another subset retains the CD11b⁺ expression. To determine whether the liver displays characteristics of DC subsets from nonlymphoid tissue, we probed the diversity of DC populations in the liver by analyzing the expression of CD11b, CD103, and CD8α. Gating strategies for analyzing liver-resident DCs are shown in Supplemental Fig. 1. CD11c^hiMHC-II^hiB220⁻ DCs resident in the livers of naive mice could be discriminated into CD11b⁺CD103⁻, CD11b⁻CD103⁺, and CD11b⁻CD103⁻ subsets (Fig. 1A, upper panels), which were further distinguished based on CD8α expression (Fig. 1A, lower panels). This analysis revealed that liver CD11b⁻ DCs consist of four distinct subsets: CD103⁻CD8α⁻, CD103⁺CD8α⁻, CD103⁺CD8α⁺, and CD103⁻CD8α⁺ cells, suggesting that liver-resident DCs segregate into subsets displaying unique cell surface markers with phenotypic characteristics of both lymphoid and nonlymphoid DCs. These findings suggest that the liver has the physiological properties of both lymphoid and nonlymphoid tissues, which is consistent with the view that it serves as a tertiary lymphoid organ.

Because most of the current emphasis in DC biology focuses on the unique differences between CD11b⁺CD103⁻ and CD11b⁻CD103⁺ subsets (henceforth designated as CD11b⁺ and CD103⁺ DCs, respectively), we limited our analysis to these two dominant liver DC populations. Accordingly, CD103⁺ DCs are the primary subset of naive liver-resident DCs in both percentage and total cell number (Fig. 1B).

Early reports describing liver DCs often note their localization to the portal tracts of the liver parenchyma (30, 31). Consistent with these earlier observations, we found that CD103⁺ DCs were localized most frequently around the portal tracts (Fig. 1C). The liver anatomy is unique in that the highest volume of blood flow comes directly from the gastrointestinal tract, entering from the portal veins. Because this blood is highly concentrated in microbial products and other food Ags, it is likely that liver-resident DCs congregate closely to these gateways to sample newly arriving potential Ags, including pathogenic microorganisms.

Although murine liver DCs were reported to exhibit a less mature phenotype than splenic DCs, the expression of MHC-II and costimulatory molecules (i.e., CD80 and CD86) are readily detectable on DCs in the liver (Fig. 1D) (32). Interestingly, PD-L1 expression is lower on CD103⁺ liver-resident DCs compared with the CD11b⁺ subset, suggesting that CD103⁺ cells may be less inhibitory as APCs and, thereby, display an increased capacity to support naive lymphocyte activation (33).

In view of the display of costimulatory ligands by hepatic DCs, it was of interest to assess their capacity to support the activation of Ag-specific CD8⁺ T cells. To this end, we sorted hepatic DCs from the livers of uninfected donors into CD103⁺ and CD11b⁺ subsets and analyzed the capacity of these DCs to support proliferative expansion of naive TCR-transgenic OT-I CD8⁺ T cells in vitro following pulsing of the DC with varying concentrations of the OT-I T cell ligand SIINFEKL peptide. As assessed by CFSE dilution, CD103⁺ DCs stimulated more potent peptide dose-dependent proliferative responses than did their peptide-pulsed CD11b⁺ DC counterparts (Fig. 2A). Compared with DC subsets, pDCs were least efficient in triggering CD8⁺ T proliferation. This enhanced stimulatory capacity of CD103⁺ DCs also was evident from the elevated numbers of total OT-I CD8⁺ T cells recovered at the end of the in vitro culture (Fig. 2B). Taken together, these findings suggest that the hepatic CD103⁺ DC subset in a steady-state displays an intrinsically enhanced capacity to present the processed antigenic peptides to naive CD8⁺ T cells compared with CD11b⁺ DCs.

The above findings suggested that liver-resident CD103⁺ DCs were potent APCs and potentially could serve as the primary APC to support CD8⁺ T cell differentiation following exposure to a hepatotropic virus infection (e.g., high dose of i.v. Ad). To examine the impact of CD103⁺ liver-resident DCs in response to infection we used mice deficient in the transcription factor Batf3, which is reported to be necessary for the development of the CD103⁺ (and corresponding CD8α⁺) DC lineage (25). As expected, Batf3^−/− mice lacked both CD103⁺ and CD8α⁺ liver-resident DCs (Supplemental Fig. 2A). Moreover, the expression of CD103⁺ liver DCs was not increased following Ad infection (Supplemental Fig. 2B), suggesting that this subset was not induced in response to Ad-mediated liver inflammation.

To explore the impact of CD103⁺ DC deficiency on naive CD8⁺ T cells responding in the liver to virus infection, we adoptively transferred naive CFSE-labeled OT-I T cells into wild-type and Batf3^−/− mice 1 d prior to i.v. infection with the replication-deficient recombinant Ad expressing the OVA gene. This administration route primarily results in virus infection of the liver, including hepatocyte and nonhepatocyte cell populations. Livers, spleens, and liver-draining and nonliver-draining LNs (DLNs) were analyzed at various time points postinfection for accumulation of OT-I T cells and for CFSE dilution. OT-I T cells were not detectable in any site at 2 d postinfection (dpi; data not shown). OT-I T cells responding to Ad were readily detectable at 3 dpi, with the highest accumulation of T cells evident in the liver (Fig. 3A). In contrast, infected Batf3^−/− T cell recipients displayed decreased frequency and total numbers of responding OT-I T cells in all tissues examined (Fig. 3B, 3C). Because of the defect in Ad replication, virus clearance and titers cannot be assessed directly. Using quantitative real-time PCR to measure the message of Ad hexon protein showed no differences between the wild-type and Batf3^−/− livers at 2 or 3 dpi (data not shown). Furthermore, we performed histological studies to assess liver damage. Notably, there was increased mononuclear cell infiltrates detectable in wild-type livers compared with Batf3^−/− livers at 3 dpi (Fig. 3D).

When the CFSE-dilution profile of OT-I T cells responding in the wild-type and Batf3^−/− T cell recipients was evaluated, we noted that the tempo of OT-I cell division was more rapid in the livers of wild-type mice. However, this accelerated CFSE-dilution profile and OT-I T cell accumulation were not evident in the liver-DLNs of the wild-type and Batf3^−/− recipients (Fig. 3E) (34, 35). Activated (responding) OT-I T cells isolated from the livers of wild-type and Batf3^−/− mice at 3 dpi display comparable levels and frequencies of the activation markers CD69 and CD25 (Supplemental Fig. 3). Similarly, responding OT-I T cells isolated from the two liver sources demonstrated a comparable capacity to secrete IFN-γ in terms of both cell frequency and mean fluorescence intensity (MFI) upon in vitro stimulation with cognate synthetic peptide epitope (Supplemental Fig. 3). Lastly, i.v. infection with MCMV-OVA, another hepatotropic virus, results in similar loss of OT-I T cell accumulation in Batf3^−/− livers, with no significant differences in IFN-γ expression (Supplemental Fig. 4).

The pattern of responsiveness of transferred OT-I T cells in the livers of wild-type and Batf3^−/− mice following Ad infection was not influenced by the cell inoculum dose of OT-I T cells transferred. Adoptive transfer of 2 × 10⁶ naive OT-I T cells into wild-type and Batf3^−/− mice (i.e., 10-fold higher cell inoculum dose than used above) allows us to identify the transferred T cells as early as 1 dpi. At this early time point of infection, the frequency of transferred OT-I T cells in wild-type and Batf3^−/− livers were comparable (Fig. 4). Thus, the enhanced expansion of the adoptively transferred T cells observed in infected wild-type recipients was not due to a difference in homing efficiency to the livers of Batf3^−/− mice.

Upon activation, tissue-resident DCs upregulate CCR7 and follow the CCL19/21 chemokine gradient into the lymphatics to ultimately reside in secondary lymphoid organs, particularly LNs draining the site of Ag deposition in the target organ. Previous work from our laboratory implicated liver parenchymal cells (e.g., hepatocytes, liver sinusoidal endothelial cells) as APCs for antiviral CD8⁺ T cells responding to virus infection of the liver (36). It was of interest to determine the contribution of liver-resident DCs to the induction of CD8⁺ T cell responses following Ad infection. In particular, we wanted to ascertain whether liver DCs, upon encountering viral Ag, must migrate to secondary lymphoid organs to stimulate responses from naive CD8⁺ T cells trafficking through the sites. To explore this possibility, we carried out adoptive transfer of naive CCR7⁺ OT-I T cells into CCR7^−/− mice, which limit DC migration, and examined the proliferative expansion and tissue localization of the transferred T cells following i.v. Ad infection (29).

The frequency and absolute number of responding OT-I T cells were dramatically reduced in secondary lymphoid organs (e.g., spleen, liver-DLNs) of CCR7^−/− mice at 3 dpi, exhibiting ∼10-fold decrease (Fig. 5A). The frequency and absolute number of OT-I T cells present in the liver were unaffected by CCR7 deficiency in the liver DCs (Fig. 5B).

To more directly assess in situ priming of CD8⁺ T cells by hepatic CD103⁺ DCs, we adoptively transferred naive OT-I T cells into B6 and Batf3^−/− mice that were first splenectomized and treated with anti-CD62L (MEL-14) Ab to prevent naive OT-I T cells from entering secondary lymphoid tissues. Splenectomized and MEL-14–treated wild-type mice exhibit increased hepatic OT-I accumulation at 3 dpi compared with either sham and isotype-treated controls or splenectomized and isotype-treated controls. The level of OT-I accumulation in the livers of splenectomized and MEL-14–treated Batf3^−/− mice increases to the level of sham and isotype-treated B6 livers and is consistent with the differences seen in Fig. 3C, where there are fewer numbers compared with the equally treated splenectomized and MEL-14 B6 livers. Taken together, these data suggest that CD103⁺ DCs contribute significantly to the accumulation of liver OT-I T cells in situ.

Because hepatic-resident DCs are capable of priming naive OT-Is in situ, we next characterized the maturation status of DCs in the liver directly following infection. Although there is a difference in the frequency and numbers of CD11b⁺ and CD103⁺ DCs by 12 hpi (Fig. 6A, 6B), hepatic CD103⁺ DCs exhibit the highest expression of the costimulation markers CD80 and CD86, common indicators of DC activation (Fig. 6C). Additionally, hepatic CD103⁺ DCs stained highest for MHC-I^SIINFEKL by 12 hpi (Fig. 6D), suggesting that they are the primary DCs presenting Ag during the early phase of viral infection. As a positive control for MHC-I^SIINFEKL staining, total splenocytes were pulsed with SIINFEKL peptide for 20 min at 37°C. LN DCs were below the limit of detection for MHC-I^SIINFEKL at 12 hpi (data not shown).

To test whether hepatic CD103⁺ DCs present endogenous viral Ag to naive CD8⁺ T cells in vitro, we isolated liver DCs 12 hpi and cocultured them with naive OT-I T cells. After a 4-d incubation, there were more CFSE-diluted OT-I CD8⁺ T cells in the CD103⁺ DC coculture compared with CD11b⁺ DC stimulation (Fig. 6E). These results reinforce a critical role for CD103 DCs in Ag presentation and contribution to CD8⁺ T cell priming at the early viral infection.

In the current study, we examined the role of liver-resident CD103⁺ DCs in the activation of naive CD8⁺ T cells and the site of their accumulation after activation in response to hepatic viral infection using a hepatotropic Ad-infection model. The liver is believed to differ from other organs, such as lung and skin, in that naive CD8⁺ T cells can be activated within the hepatic parenchymal site (37). Studies demonstrated that nonhematopoietic APCs in the liver contribute to the induction of naive CD8⁺ T cell proliferation and differentiation. However, the role of hematopoietic cells, such as DCs, in processing and presenting local Ags to initiate the CD8⁺ T cell response in situ has not been well established. Batf3 is a transcription factor that controls CD103⁺/CD8α⁺ DC development; therefore, Batf3^−/− mice exhibit a selective loss of these subsets. Batf3^−/− mice have been widely used in the study of DC biology and provide a unique and valuable experimental tool to investigate the contribution of CD103⁺ DCs to antiviral immunity in vivo. Although there are potential caveats when using any global gene–deficient transgenic, we are not aware of any report in the literature of any defect beyond the selective deficiency of this DC subset in Batf3^−/− mice. By using Batf3-deficient mice, we demonstrated that liver-resident CD103⁺ DCs play a critical role in the establishment of optimal CD8⁺ T cell responses to hepatic Ad infection. In support of their dominant role as APCs in inducing antiviral CD8⁺ T cell responses, CD103⁺ hepatic DCs displayed a superior ability to process and present virus-derived antigenic peptide to naive CD8⁺ T cells in the steady-state and following infection. Lastly, we demonstrated that liver DC priming of naive CD8⁺ T cells occurs independently of CCR7-mediated DC egress, supporting the view that DCs can activate naive CD8⁺ T cells in situ. Our findings demonstrate for the first time, to our knowledge, that hematopoietic-driven hepatic DCs, in particular the CD103⁺ DC subset, serve as the prominent APC in the liver in response to viral infections and induce Ag-specific CD8⁺ T cell responses within the liver tissue early in infection.

Based on our finding that there is a decrease (3-fold) in accumulating OT-I CD8⁺ T cells in the livers of Batf3^−/− mice that received OT-I cells at 3 dpi of Ad-OVA infection compared with control B6 mice (Fig. 3A), it is likely that liver-resident CD103⁺ DCs play a major role in the induction and accumulation of Ag-specific CD8⁺ T cells at the site of hepatic viral infection. Notably, there is a similar decrease in OT-I CD8⁺ T cells accumulating in the nondraining inguinal LNs (iLNs) and draining celiac/hepatic LNs (cLNs) but a far less significant loss of OT-I cells in the spleens of Batf3^−/− mice. These results are likely due to the contribution of CD11b⁺ DCs to OT-I accumulation in secondary lymphoid tissues. It is also possible that splenic CD11b⁺ DCs and/or additional APCs are better at priming than are their LN counterparts but that they contribute to a lesser degree than do OT-I cells induced by CD8α/CD103⁺ DCs. Moreover, the proliferation index formula to measure OT-I divisions (Fig. 3E) indicates a significant decrease in Batf3^−/− livers, having less accumulation of CFSE^lo OT-I cells, yet is only trending lower in the lymphoid tissues. The proliferation index differences in the livers of control B6 and Batf3^−/− mice suggest that resident CD103⁺ DCs lead to sustained OT-I accumulation and that non-CD103⁺ APCs, such as CD11b⁺ DCs and nonhematopoietic presenting cells, are either less efficient at priming or, more likely, induce unsustainable Ag-specific CD8⁺ T cells. At 1 dpi there is no difference in the number of liver-accumulating OT-I cells between B6 and Batf3^−/− mice, leading us to speculate that initial CD8⁺ T cell priming is independent of CD103⁺ DCs but that, at 3 dpi, any remaining OT-I T cells are greatly outnumbered by OT-I cells primed by resident CD103⁺ DCs.

Infection of peripheral tissues (e.g., lung and skin) triggers the mobilization of tissue-resident DCs to egress out of the sites of pathogen entry and replication. These Ag-bearing migratory tissue-resident DCs arrive at the tissue-draining regional LNs where they participate in instructing naive Ag-specific CD8⁺ T cell activation and differentiation. In contrast, blood-borne pathogens are captured by phagocytic cells (e.g., DCs and macrophages) in the spleen, and Ag-specific CD8⁺ T responses are initiated primarily by lymphoid tissue–resident DCs (i.e., CD8α⁺ DCs). However, there is limited understanding as to how the inductive phase of the CD8⁺ T cell immune response is orchestrated in response to hepatotropic infections. Previously reports described hepatic-resident DCs, suggesting that the CD11b⁺ subset is dominant in the liver (21, 25). However, our data show that CD103⁺ DCs represent the largest population of liver-resident DCs. Having optimized the isolation techniques and gating strategies, we believe that our results are the most accurate representation of liver-resident DC populations. We further demonstrated that the liver displays a distinct DC compartment in which it harbors both CD103⁺ and CD8α⁺ DCs along with the CD11b⁺ subset and pDCs. This finding is in agreement with the view that the liver may inherently possess lymphoid-like features and serves as a tertiary lymphoid organ (38). The unique composition of the DC network within the liver could support T cell activation locally while permitting migratory DC–dependent T cell activation in the regional DLN after migration. Because we noted a decrease in hepatic DCs at 12 hpi, it is possible that these cells are either eliminated or migrate out of the parenchyma. Indeed, it is well established that initial contact between DC and T cells is sufficient to trigger the full activation and differentiation pathways. Thus, we believe that the decrease in CD103⁺ DC numbers postinfection has a minimal impact on the onset of CD8⁺ T cell priming in the liver. Migratory DCs use CCR7 to migrate out of infected tissues and trigger T cell proliferation in the DLN. By limiting DC egress and subsequent homing of the CD8⁺ T cells primed by hepatic migratory DCs in the DLN of CCR7^−/− mice, our findings suggest that intrahepatic DCs are capable of activating CD8⁺ T cell responses to hepatotropic virus infection. Furthermore, preventing naive OT-I T cells from entering secondary lymphoid organs by administering anti-CD62L Ab to splenectomized B6 and Batf3^−/− mice demonstrated that hepatic CD103⁺ DCs are a primary resident APC for induction of naive CD8⁺ T cell responses. It is possible that blocking naive OT-I cell accumulation in the peripheral lymphoid organs prior to infection will increase their availability to hepatic APCs, accounting for the considerable increase in OT-I cells in splenectomized and anti-CD62L–treated livers compared with the sham and anti-IgG controls at 3 dpi. After i.v. administration of the replication-deficient Ad5 virus, the virus is cleared primarily by liver Kupffer cells, with the majority of residual virus entering hepatocytes via factor X/Ad5 complexes (39, 40). CD103⁺ DCs (and possibly CD8α⁺ DCs) are predominantly localized within the marginal areas of the portal veins, where they are ideally poised to take up Ag from circulation and/or neighboring infected cells and present processed Ag to CD8⁺ T cells in situ. The extent to which intrahepatic priming of CD8⁺ T cells is controlled by either CD8α⁺ DCs or CD103⁺ DCs has not been determined. Of note, it is an intriguing possibility that intrahepatic CD8⁺ T cell priming is carried out by CD8α⁺ DCs, whereas CD103⁺ DCs initiate T cell responses in the DLN after migration.

In contrast, OT-I CD8⁺ T cells in B6 wild-type mice appear to be activated in the spleen and iLNs with similar kinetics to T cells activated in the liver and cLNs, but no activated OT-I T cells are observed in the spleen or iLNs of CCR7^−/− mice. These results suggest that nondraining peripheral secondary lymphoid tissues depend on a CCR7⁺ APC entering those sites (possibly coming from the liver via the blood). Moreover, fewer activated OT-I T cells in CCR7^−/− liver-DLNs compared with those in wild-type mice suggest that either activated liver DCs use additional chemokine receptors (possibly CXCR4) when trafficking through their draining lymphatics or that Ag is brought to the draining nodes via alternative mechanisms.

The immunologically hyporesponsive (tolerogenic) microenvironment of the liver is necessitated by its continuous exposure to food and bacterially derived Ag. Liver-resident DCs localized in the vicinity of the portal veins are likely actively encountering material draining from the gut and circulation. However, unwarranted activation in response to nonpathogenic Ags requires DCs to be under tight immune control to continuously maintain the “normal” state. Our studies demonstrate that resident DCs express significant amounts of PD-L1. Of note, the CD11b⁺ subset preferentially expresses the greatest amount of this coinhibitory molecule. Our data argue that, immediately following Ad-OVA infection, hepatic CD103⁺ DCs significantly upregulate CD80, CD86, and MHC-II and are capable of inducing antiviral CD8⁺ T cell immune responses, whereas CD11b⁺ DCs may provide counter-regulatory signals to CD103⁺ DC–mediated T cell activation. This potential cross-talk between CD103⁺ DCs and CD11b⁺ DCs may control the threshold in the liver to induce antiviral immunity or to induce tolerogenic immune responses. Consistent with our previously published studies, intrahepatic CD8⁺ T cells primed in Batf3^−/− mice exhibit equally poor effector function as do those primed in B6 mice. This impaired CD8⁺ T cell effector function might be reflected by additional factors controlling hepatic CD8⁺ T cell responses in the liver compartment. Thus, interventions using anti–PD-L1 blockade agents could lead to an increase in T cell activation by these hepatic DCs (41). Collectively, these data suggest that liver-resident CD103⁺ DCs may serve as critical regulators of immunity to intrahepatic antigenic challenges.

Although there have been substantial therapeutic agents aimed at curing chronic infection, such as HCV, the current medical cost for treating chronic HCV patients is unsustainable. Therefore, vaccine development is necessary to protect the host from microbial infection. In particular, hepatotropic pathogens (i.e., HCV, malaria) exploit the immunosuppressive liver microenvironment and often establish persistent infection, resulting in severe liver diseases, including cirrhosis and hepatocellular carcinoma. Thus, understanding liver immunology and studying ways to generate effective immune responses is crucial for the development of efficient liver-disease therapies. Given the ability of CD103⁺ DCs to mount sustainable CD8⁺ T cell responses specific to Ag in situ, as described in this report, targeting of Ag to liver-resident DCs (i.e., CD103⁺ DCs) would be helpful in designing vaccines against hepatotropic pathogens and boost host immune responses to combat infection. In summary, our study highlights the prominent contribution of liver-resident CD103⁺ DCs to the initiation of antiviral CD8⁺ T cell immune responses to hepatotropic infections. Further, this work opens up the possibility of targeting liver-resident CD103⁺ DCs for potential vaccination strategies.

We thank the members of the Hahn laboratory for providing advice and constructive criticism on this work. In particular, we thank Sowmya Narayanan and Albert H.S. Nieh for critical reading and comments during the preparation of this manuscript.

The authors have no financial conflicts of interest.