β-Catenin Signaling Drives Differentiation and Proinflammatory Function of IRF8-Dependent Dendritic Cells

Sara B. Cohen, Norah L. Smith, Courtney McDougal, Marion Pepper, Suhagi Shah, George S. Yap, Hans Acha-Orbea, Aimin Jiang, Björn E. Clausen, Brian D. Rudd and Eric Y. Denkers
J Immunol January 1, 2015, 194 (1) 210-222; DOI: https://doi.org/10.4049/jimmunol.1402453

Abstract

β-Catenin signaling has recently been tied to the emergence of tolerogenic dendritic cells (DCs). In this article, we demonstrate a novel role for β-catenin in directing DC subset development through IFN regulatory factor 8 (IRF8) activation. We found that splenic DC precursors express β-catenin, and DCs from mice with CD11c-specific constitutive β-catenin activation upregulated IRF8 through targeting of the Irf8 promoter, leading to in vivo expansion of IRF8-dependent CD8α+, plasmacytoid, and CD103+CD11b− DCs. β-Catenin–stabilized CD8α+ DCs secreted elevated IL-12 upon in vitro microbial stimulation, and pharmacological β-catenin inhibition blocked this response in wild-type cells. Upon infections with Toxoplasma gondii and vaccinia virus, mice with stabilized DC β-catenin displayed abnormally high Th1 and CD8+ T lymphocyte responses, respectively. Collectively, these results reveal a novel and unexpected function for β-catenin in programming DC differentiation toward subsets that orchestrate proinflammatory immunity to infection.

This article is featured in In This Issue, p.1

Introduction

Dendritic cells (DCs) critically bridge innate and adaptive immunity through their exquisite capacity to drive Ag-specific T cell activation and effector subset differentiation. Furthermore, DCs are central players in determining tolerance versus immunity during inflammation and infection (1). DCs are a heterogeneous population of cells with varying surface markers and transcription factor requirements. All originate from a common myeloid progenitor (CMP), but they subsequently differentiate into distinct subsets, including monocyte-derived DCs, conventional DCs (cDCs), and plasmacytoid DC (pDCs). Many elegant studies have identified phenotypic and functional differences among these subsets, but identifying factors determining control points of DC subset generation is a continuing focus of intense interest. Several key cytokines and transcription factors have been implicated in controlling DC developmental pathways (2), and recent gene mapping studies have begun to elucidate the order in which these factors become expressed (3, 4). For example, transcription factor Batf3 is involved in generation of splenic CD8α+ DCs, whereas IFN regulatory factor 4 (IRF4) is important in differentiation of CD11b+CD103+ DCs in the intestinal lamina propria (5, 6). Recently, Zbtb46 was identified as a global transcription factor necessary for generation of cDCs (3). Nevertheless, a thorough understanding of the mechanisms of DC differentiation and the signals that direct branch points leading to distinct subsets remains incomplete.

β-Catenin is the primary mediator of the Wnt signaling pathway and is critical for numerous cellular functions, including hematopoietic cell fate determination and proliferation (7, 8). Cytosolic β-catenin levels are normally maintained at low levels through continual phosphorylation by the serine threonine kinases glycogen synthase kinase-3β and casein kinase I-α, which cooperate to promote its ubiquitination and proteasomal degradation. Activating Wnt ligands trigger disassembly of the complex that coordinates these kinases, leaving β-catenin unphosphorylated, in turn enabling nuclear translocation for transcriptional activity in association with T cell factor/lymphoid enhancer factor transcription factors (9). Although normally associated with embryonic development and tumorigenesis (10), β-catenin is increasingly being recognized for its role in immunity (11). This is particularly the case for DCs, where β-catenin signaling was first implicated in cluster disruption-mediated maturation toward a tolerogenic phenotype during in vitro culture (12). Moreover, β-catenin was found to be involved in the generation or maintenance of tolerogenic DC subsets in the intestinal mucosa (13).

In this article, we provide surprising new insight into the role of β-catenin in DC function by using transgenic mice with a CD11c-specific deletion in the third exon of the β-catenin gene. The exon 3 fragment encodes the β-catenin amino acid sequence that is targeted for glycogen synthase kinase-3β–mediated serine threonine phosphorylation and subsequent degradation. Removal of this region through Cre-lox–mediated excision therefore results in phosphorylation-resistant and constitutively active β-catenin (14). We made the unexpected discovery that β-catenin stabilization in DC results in selective expansion of steady-state levels of splenic CD8α+ DCs, pDCs, and peripheral CD103+ DCs. These DC subsets share a dependence on IRF8 for their differentiation, and in accordance with this observation, we show that constitutive β-catenin signaling increases IRF8 expression by these DC subsets via enhanced targeting of the Irf8 promoter. We used infections with the intracellular protozoan Toxoplasma gondii and vaccinia virus (VACV) to determine the in vivo consequences of DC-specific β-catenin stabilization. In accord with the known role of CD8α+ DCs as an IL-12 source and driver of Th1 responses during T. gondii infection (15), the parasite triggered an abnormally strong Th1 response associated with overproduction of IL-12 and IFN-γ. Immunity to VACV is known to require a DC-mediated cross-presentation pathway (16). As such, vaccinia infection in mutant mice triggered enhanced expansion and activation of virus-specific CD8+ T cells. Our results uncover a new role for β-catenin in controlling IRF8 expression in DCs, thereby revealing this transcription factor as a key player regulating IRF8-driven DC differentiation and proinflammatory function.

Materials and Methods

Ethics statement

All experiments in this study were performed strictly according to the recommendations of the Guide for the Care and Use of Laboratory Animals of the National Institutes of Health. The protocols were approved by the Institutional Animal Care and Use Committee at Cornell University (permit no. 1995-0057). All efforts were made to minimize animal suffering during the course of these studies.

Mice and infections

Female Swiss Webster mice (6–8 wk of age) were purchased from The Jackson Laboratory (Bar Harbor, ME), and female C57BL/6 were purchased from Taconic Farms (Germantown, NY). C57BL/6-Tg(TcraTcrb)425Cbn/J (OT-II) mice were obtained from the Jackson Laboratory and maintained as a breeding colony at Cornell University College of Veterinary Medicine. The β-catenin Ex3fl/fl mice were kindly provided by M. M. Taketo (Kyoto University, Kyoto, Japan) and were maintained as breeding colonies crossed to CD11c-cre–expressing mice at the Transgenic Mouse Core Facility at the Cornell University College of Veterinary Medicine. Cre+ offspring (Ex3DC−/− mice) were identified by PCR amplification of the Cre gene from genomic DNA isolated from tail snips. Infections were initiated in 8- to 12-wk-old mice. T. gondii infections were performed by i.p. inoculation of 25 cysts of the type II ME49 strain. Cysts were isolated from chronically infected Swiss Webster mice by homogenization of whole brain in sterile PBS. Alternatively, mice were inoculated with 2 × 105 PFU recombinant VACV expressing MHC class I–restricted HSV glycoprotein B (gB) peptides 498–505 (VACV-gB) by i.p. injection (17). VACV-gB was maintained in 143B cells for the generation of viral stocks.

Preparation and purification of leukocytes

Splenocyte single-cell suspensions were prepared by crushing spleens between sterile glass slides and filtering the resulting suspension through 40-μM filters. For lung leukocytes, lung tissue was minced with sterile razor blades and incubated with collagenase type IV (Sigma-Aldrich) in a 37°C water bath for 30 min with frequent agitation. The resulting digest was passed through a 40-μM filter to create a single-cell suspension. A single round of positive selection using CD11c+ magnetic bead sorting was performed for purification of total splenic DCs from single-cell suspensions (Stem Cell Technologies), whereas two-step magnetic bead sorting, with an initial negative selection to enrich for DCs followed by CD8α+ positive selection (Miltenyi Biotec), was performed to isolate CD8α+ splenic DCs.

In vitro culture of bone marrow–derived DCs and MutuDC1940 cells

Bone marrow–derived DCs (BMDCs) were cultured as described previously (18). In brief, femurs of Ex3fl/fl, Ex3DC−/−, or C57BL/6 mice were flushed with PBS and cultured for 9 d in media containing 10% FCS (Hyclone), 100 U/ml penicillin (Life Technologies), 0.1 mg/ml streptomycin (Life Technologies), 50 μM 2-ME (Sigma-Aldrich), and 20 ng/ml GM-CSF (Peprotech). Cells were harvested from the plates with gentle pipetting and cultured as indicated. Flt3L BMDC cultures were performed by flushing femurs with PBS, lysing RBCs with ACK lysis buffer (Life Technologies), and plating cells in RPMI 1640 supplemented with 10% FCS, 25 mM HEPES (Life Technologies), 100 U/ml penicillin (Life Technologies), 0.1 mg/ml streptomycin (Life Technologies), and 100 ng/ml murine Flt3L (PeproTech). Cells were cultured for 9 d at 37°C. MutuDC1940 cells, kindly provided by Dr. Hans Acha-Orbea (University of Lausanne), were grown in a monolayer in media containing 8% FCS (Hyclone), 10 mM HEPES (Life Technologies), 50 μM 2-ME, 100 U/ml penicillin (Life Technologies), and 0.1 mg/ml streptomycin (Life Technologies). Cells were harvested by 10-min incubation with PBS and 5 mM EDTA.

Western blotting

To validate nuclear translocation of β-catenin in Ex3DC−/− mice, we subjected BMDCs to nuclear and cytoplasmic fractionation following the manufacturer’s guidelines (Active Motif). Resulting nuclear and cytoplasmic proteins were diluted in reducing SDS sample buffer and separated by 10% SDS-PAGE. Separated proteins were transferred onto nitrocellulose and blocked for 1 h at room temperature in TBS containing 0.1% Tween 20 and 5% nonfat dry milk (TBST). After three washes in TBST, blots were incubated overnight in primary Ab diluted in TBST containing 5% BSA. Blots were subsequently washed in TBST and incubated for 1 h with anti-rabbit IgG conjugated to HRP-conjugated diluted in TBST containing 5% nonfat dry milk. After five washes in TBST, blots were developed and imaged using a chemiluminescent detection system (Thermo Scientific). Anti–β-catenin and anti-poly (ADP-ribose) polymerase were purchased from Cell Signaling, and anti-Rab5a was purchased from Santa Cruz Biotechnology.

Flow cytometry

Single-cell suspensions were washed in PBS before resuspension in Zombie Aqua viability dye (BioLegend) for 15 min at room temperature to exclude dead cells. Primary Abs (anti-CD11c FITC, eFluor610, or allophycocyanin; anti-CD8α Pacific Blue or allophycocyanin-Cy7; anti-CD4 PerCP-Cy5.5 or FITC; anti–PDCA-1 allophycocyanin or PE; anti-NK1.1 FITC or allophycocyanin; anti-CD11b FITC or allophycocyanin-Cy7; anti-CD24 Pacific Blue; anti-CD103 PE; anti-B220 PE; anti-CD3 FITC; anti-Gr1 FITC; anti-CD127 PerCP-Cy5.5; anti-CD16/32 eFluor 450; anti-CD19 FITC; anti-CD135 PE; anti-Sca1 PE-Cy7; anti-CD117 allophycocyanin-Cy7; anti-MHC class II (anti–MHC II) PE or FITC) resuspended in ice-cold FACS buffer (1% BSA/0.01% NaN3 in PBS) were added directly to the cells for 30 min. Tetramer staining for Tgd057+CD8+ T cells (provided by George Yap, New Jersey School of Medicine and Dentistry), CD4Ag28m+CD4+ T cells (provided by Marion Pepper, University of Washington), and gB-8p:Kb+CD8+ T cells (National Institutes of Health Tetramer Core Facility, Emory University, Atlanta, GA) was performed by labeling at room temperature for 1 h. For intracellular staining, cells were fixed using the Foxp3/transcription factor staining kit fixative (eBioscience) and subsequently incubated with primary Abs resuspended in the Foxp3/transcription factor permeabilization buffer (eBioscience). Abs used for intracellular cytokine staining include anti-IRF8 PerCP and anti-IRF4 eFluor450 (eBioscience), and anti–β-catenin Alexa 647 (Cell Signaling). IgG isotype controls (eBioscience) were used for each fluorophore. For IFN-γ staining post T. gondii infection, cells were incubated for 4 h with brefeldin A (10 μg/ml; eBioscience), PMA (10 ng/ml; Sigma-Aldrich), and ionomycin (1 μg/ml; Sigma-Aldrich), whereas for VACV-gB infection, splenocytes were restimulated with gB-8p peptide (SSIEFARL; 10−7 M) for 5 h in the presence of brefeldin A. Cells were then fixed with the Foxp3/transcription factor staining kit fixative (eBioscience) and subsequently incubated with anti–IFN-γ (PE-Cy7, BioLegend; PE, eBioscience). All samples were run on an LSRII flow cytometer (BD), and the data were analyzed using FlowJo software (FlowJo, Ashland, OR).

Identification of bone marrow and splenic DC progenitors

Bone marrow was flushed from the femur and tibia of Ex3fl/fl and Ex3DC−/− mice and broken up with a 21-gauge syringe. Bone marrow and splenocyte pellets were lysed with ACK buffer and passed through a 40-μm filter to generate single-cell suspensions. For precursor staining, a FITC lineage mixture was created in-house (anti-NK1.1, anti-CD11b, anti-Gr1, anti-Ter119, anti-CD3, anti-MHCII, anti-CD19). Gating strategies were adopted from a previous study (3), whereby CMPs were defined as Lin−Sca1−CD127−CD117hiCD11c−CD135+CD16/32−, granulocyte-macrophage progenitor (GMP) as Lin-Sca1−CD127−CD117hiCD11c−CD135−CD16/32+, common dendritic progenitor (CDP) as Lin−Sca1−CD127−CD117intCD11c−CD135+CD16/32−, and preconventional DCs (precDCs) as Lin−Sca1−CD127−CD117loCD16/32−CD11c+CD135+. Pre-CD8α+ DCs were defined as CD11c+CD8α−B220−CD24+ as previously described (19).

Chromatin immunoprecipitation

Recombinant murine Flt3 ligand bone marrow cultures from Ex3DC−/− mice (1 × 107) were cross-linked with 1% formaldehyde (Pierce) at room temperature for 6 min, and chromatin/protein complexes were prepared following the manufacturer’s instructions (Millipore). Shearing was performed using a Bioruptor UCD-200 sonicator (Life Technologies) in an ice slurry, with a program of 30 s on and 60 s off for 30 min. Protein immunoprecipitation was performed overnight with agitation at 4°C using protein G magnetic beads and 2 μg of a rabbit monoclonal β-catenin Ab or rabbit IgG (Cell Signaling). The following day, samples were proteinase K digested at 65°C to free the DNA, and semiquantitative PCR was performed to amplify a region of the Irf8 promoter containing a Wnt/β-catenin binding site (forward, CACACTGGGTGGACATTTG; reverse, ACCTTATAAGCGTATGCAGATT). DNA levels were normalized to 1% input chromatin.

ICG-001 inhibition

BMDCs (1 × 106) or MutuDC1940 cells (2 × 105) were plated and cultured overnight. The following day, the cells were cultured with 5 or 20 μM ICG-001 (Selleck Chemicals), respectively, or DMSO control for 5 h or overnight at 37°C. Cells were surface stained for CD11c and/or CD24; fixed; intracellularly stained for IRF8, IRF4, and β-catenin; and analyzed by flow cytometry. Alternatively, DCs were resuspended in TRIzol (Life Technologies) for quantitative PCR analysis of Irf8 and Axin2 mRNA transcripts.

Measurement of mRNA by quantitative PCR

RNA was isolated from CD11c+ splenocytes magnetically sorted from naive Ex3DC−/− and Ex3fl/fl mice or from BMDCs by resuspension in TRIzol reagent (Life Technologies). RNA was converted to cDNA (Quanta Biosciences, Gaithersburg, MD) and assayed for gene expression by SYBR green technology (Quanta Biosciences). Primers were designed to span exons by Integrated DNA Technologies. The following primer sequences were used: Irf8 forward, TGCCACTGGTGACCGGATAT; reverse, GACCATCTGGGAGAAAGCTGAA; Nfil3 forward, GAACTCTGCCTTAGCTGAGGT; reverse, ATTCCCGTTTTCTCCGACACG; Id2 forward, ATGAAAGCCTTCAGTCCGGTG; reverse, AGCAGACTCATCGGGTCGT; Batf3 forward, CAGACCCAGAAGGCTGACAAG; reverse, CTGCGCAGCACAGAGTTCTC; Axin2 forward, TAGGTTCCGGCTATGTCTTTG; reverse, TGTTTCTTACTCCCCATGCG. GAPDH was used as a housekeeping gene. Gene expression was normalized to Ex3fl/fl samples or DMSO controls.

Cytokine measurement

IFN-γ, IL-12p70, and TNF-α secretion were assayed by ELISA following the manufacturer’s instructions (eBioscience) following culture with media, LPS (100 ng/ml), or soluble tachyzoite Ag (STAg; 50 μg/ml) prepared as previously described (20). IL-12p40 secretion was measured using an in-house ELISA (21).

Parasite burden measurement

Levels of T. gondii DNA were measured as described previously (22). In brief, spleens were homogenized, and DNA was extracted using a tissue extraction kit (Omega Biotech). The T. gondii B1 gene and the host argininosuccinate lyase gene were amplified by quantitative real-time PCR, and resulting threshold cycle (ct) values were compared with standard curves developed from 10-fold serial dilutions of parasite DNA and splenocyte DNA, respectively. The parasite burden is displayed as the ratio of T. gondii DNA to host DNA.

Statistical analyses

Differences between groups were analyzed by Student t test. Expression of β-catenin among splenic DC subsets was analyzed by one-way ANOVA followed by a Newman–Keuls posttest. A Kaplan–Meier curve (log-rank test) was used to calculate differences in survival between Ex3fl/fl and Ex3DC−/− mice. The p values were considered statistically significant at <0.05 and were designated as follows: *p < 0.05, **p < 0.01, ***p < 0.001.

Results

β-Catenin is selectively enriched in splenic DC precursors and mature DCs

CMPs, GMPs, CDPs, and precDCs represent different stages of hematopoiesis toward the DC lineage and can be distinguished by surface marker expression (Supplemental Fig. 1) (3, 23). Although CMP can develop into any cell of the myeloid lineage and GMP can ultimately become macrophages, granulocytes, or DCs, CDPs and precDCs are restricted to the DC lineage, although they remain immature until terminal differentiation within the tissue (23). To investigate the role of β-catenin at these different stages of DC differentiation, we determined β-catenin expression levels among DC precursor populations in the bone marrow and spleens of wild-type (WT) mice by flow cytometry. We were able to detect each precursor population in both tissues, although the overall levels of precursors, particularly CDP, were far lower in the spleen (Supplemental Fig. 1). Interestingly, β-catenin was undetectable in the precursor populations in the bone marrow and in splenic CMPs. However, β-catenin expression was increased in splenic GMPs, CDPs, and precDCs, with GMPs and precDCs displaying the highest levels (Fig. 1A, 1B). These results provide evidence that the β-catenin signaling axis may be active in later stages of DC development.

FIGURE 1.
FIGURE 1.

β-Catenin is upregulated in splenic DC precursors and mature DC subsets. (A and B) Flow cytometric analysis of β-catenin expression by MFI in DC precursors from the bone marrow and spleen of naive WT mice compared with isotype control staining. Data are representative of two independent experiments (n = 4 mice/group). (C) Comparison of β-catenin expression levels among splenic DC subsets by flow cytometry. (D) MFI of β-catenin expression among different DC subsets. Statistics are relative to β-catenin expression in CD4+ DCs. Data are representative of three independent experiments (n = 5 mice/group). *p < 0.05, ***p < 0.001.

To ask whether β-catenin expression was maintained in mature tissue-resident DCs, we measured levels in splenic cDC subsets and pDCs. Compared with the CD4+ splenic cDC subset, immediate CD8α+ DC precursors (pre-CD8α+ DCs), which are defined by CD24 expression (19), CD8α+ DCs, and pDCs were all significantly enriched for β-catenin expression, whereas pre-CD8α+ DCs and pDCs expressed the highest levels (Fig. 1C, 1D). These data demonstrate that β-catenin is selectively induced in particular DC subsets and that, based upon protein expression levels, β-catenin signaling is more active in tissue-resident DC progenitors and mature DCs than in bone marrow cells.

β-Catenin stabilization directs splenic DC progenitors toward CD8α+ DCs

Because β-catenin was clearly expressed by DC progenitors, we next investigated the effect of β-catenin stabilization on the outcome of DC differentiation. To address this, we crossed mice floxed for exon 3 of the β-catenin gene with CD11c-cre animals, resulting in Cre-positive progeny whose CD11c+ cells possessed an exon 3–deleted β-catenin form resistant to phosphorylation-induced degradation (14). Flow cytometric analysis of CD11c+ splenic DCs from Cre-positive offspring (Ex3DC−/− mice) demonstrated high β-catenin expression levels compared with Cre-negative littermate controls (Ex3fl/fl mice), indicating accumulation of β-catenin protein upon exon 3 deletion (Fig. 2A). This was confirmed to be specific to CD11c+ cells, because CD4+ T cells from Ex3DC−/− mice did not display upregulation of β-catenin compared with CD4+ T cells from Ex3fl/fl mice (Fig. 2B). Furthermore, upon cytoplasmic and nuclear fractionation, Ex3DC−/− BMDCs were found to be enriched for a truncated form of nuclear β-catenin compared with Ex3fl/fl DCs, confirming enhanced nuclear translocation of protein (Fig. 2C). Thus, CD11c-directed exon 3 deletion results in β-catenin accumulation and nuclear translocation.

FIGURE 2.
FIGURE 2.

β-Catenin stabilization directs splenic DC progenitors toward CD8α+ DC development. (A) Intracellular β-catenin expression in naive Ex3fl/fl and Ex3DC−/− splenic CD11c+ cells. (B) Intracellular β-catenin levels in splenic CD4+ T cells isolated from Ex3fl/fl and Ex3DC−/− mice. Data show results from an individual mouse that is representative of at least three experiments with three to five mice per group. (C) Western blot analysis of β-catenin in BMDCs from Ex3fl/fl and Ex3DC−/− mice after cytoplasmic (C) and nuclear (N) fractionation. Abs against poly (ADP-ribose) polymerase (PARP) and Rab5 were used for nuclear and cytoplasmic loading controls, respectively. Data are from one independent trial. (D and E) Comparison of precDC populations from the (D) bone marrow and (E) spleen of Ex3fl/fl and Ex3DC−/− mice by flow cytometry. Numbers in representative plots represent percentages of relevant populations within the indicated gate. Bar graphs show mean percentages + SE of relevant populations. Data represent the combination of two independent experiments (n = 10 mice/group). (F) Levels of splenic pre-CD8α+ DCs, defined as CD11c+CD8α−B220−CD24+, in Ex3fl/fl and Ex3DC−/− mice by flow cytometry. Data are representative of three independent experiments, each involving four to five mice per group. **p < 0.01, ***p < 0.001.

To determine the effect of β-catenin stabilization on DC precursor levels, we quantified bone marrow and splenic precDCs from Ex3fl/fl and Ex3DC−/− mice by flow cytometry. Consistent with the failure to detect β-catenin in bone marrow precursors (Fig. 1B), no differences were observed in the levels of DC progenitors in the bone marrow after β-catenin stabilization (Fig. 2D). However, β-catenin stabilization in Ex3DC−/− mice led to significantly fewer splenic precDCs compared with Ex3fl/fl littermate controls (Fig. 2E), suggesting that constitutive β-catenin signaling was depleting this precursor pool. We next asked whether this decrease in progenitors influenced later stages of DC development, in particular, by quantifying levels of pre-CD8α+ DCs. Indeed, DC-specific β-catenin activation resulted in a 2-fold increase in splenic pre-CD8α+ DCs compared with WT controls (Fig. 2F). These data suggest that β-catenin signaling drives the differentiation of splenic DC progenitors into the immediate precursors of mature CD8α+ DCs.

β-Catenin stabilization expands splenic and peripheral DC populations

Because β-catenin signaling influenced the levels of splenic DC progenitors, we next focused on the outcome of β-catenin stabilization on steady-state levels of mature tissue-resident DCs. We first observed that the percentage and total number of CD11c+MHCII+ cells were unaffected in Ex3DC−/− mice (Fig. 3A, 3B). However, further analysis revealed a striking expansion of the CD8α+ DC subset and a concomitant decrease in CD11b+ DCs (Fig. 3C, 3E). Furthermore, pDCs, as defined by expression of B220 and PDCA-1, were also expanded in the spleens of Ex3DC−/− mice (Fig. 3F, 3G). These collective data suggest that β-catenin exerts major effects on the generation of specific splenic DC populations and are consistent with the finding that these particular subsets upregulate β-catenin during development (Fig. 1D).

FIGURE 3.
FIGURE 3.

β-Catenin stabilization expands splenic CD8α+ and pDC populations. (A–G) Mature DC subset analysis of naive Ex3fl/fl and Ex3DC−/− splenocytes by flow cytometry. (A and B) Percentage and total number of CD11c+ cells in Ex3fl/fl and Ex3DC−/− spleens. (C–E) Percentage and total number of (C and D) CD8α+ DCs and (C and E) CD11b+ DCs in naive Ex3fl/fl and Ex3DC−/− spleens. Data are representative of at least three independent experiments (n = 3–5 mice/group). (F and G) Percentage and total number of B220+PDCA-1+ pDCs in naive Ex3fl/fl and Ex3DC−/− spleens. (H and I) Mature DC subset analysis of naive Ex3fl/fl and Ex3DC−/− lung tissue by flow cytometry. (H) Plots from representative mice and (I) percentage of CD103+CD11b− lung DCs for multiple mice are shown. (J–L) Mature DC subset analysis of naive Ex3fl/fl and Ex3DC−/− intestinal lamina propria by flow cytometry. (J) Plots from representative mice and percentages of (K) CD103+CD11b− and (L) CD103+CD11b+ intestinal DCs for multiple mice are shown. Dots in relevant graphs represent results from individual mice. Bar graphs display means and SEs of individual mice. Data are representative of at least two independent experiments (n = 3–5 mice per group). *p < 0.05, **p < 0.01, ***p < 0.001.

Peripheral DCs can be subdivided into CD103+CD11b−, CD103+CD11b+, and CD103−CD11b+ DCs, and much like splenic DC subsets, these DCs display differential transcription factor requirements and functions (6, 24). Importantly, CD103+CD11b− DCs found in nonlymphoid tissues, such as the intestine and the lung, exhibit a similar dependence on Batf3 as the CD8α+ DC subset, leading to the conclusion that these two subsets are developmentally related (24). Examination of these DC subsets in Ex3DC−/− mice revealed a dramatic expansion of resident lung CD103+CD11b− DCs compared with Ex3fl/fl littermate controls (Fig. 3H, 3I). Furthermore, intestinal CD103+CD11b− DCs, but not CD103+CD11b+ DCs, were also expanded in Ex3DC−/− mice (Fig. 3J–L). These data demonstrate that constitutive DC β-catenin signaling promotes a developmental pathway that is shared by splenic CD8α+ DCs, pDCs, and peripheral CD103+ DCs.

β-Catenin signaling controls Irf8 expression

Genetic knockout studies have identified several transcription factors involved in CD8α+ DC differentiation, including Id2, Nfil3, Batf3, and Irf8 (5, 25–27). Therefore, we determined expression levels of these transcription factors among splenic CD11c+ cells from Ex3fl/fl and Ex3DC−/− mice. Although there was no significant difference in Id2 expression, Nfil3 and Batf3 transcripts were slightly increased albeit in a statistically nonsignificant manner (Fig. 4A). However, there was a striking increase in Irf8 expression in the CD11c compartment upon β-catenin stabilization (Fig. 4A). We also assessed IRF8 protein expression in CD8α− and CD8α+ splenic DCs in Ex3fl/fl and Ex3DC−/− mice by flow cytometry. Levels of IRF8 were relatively low in CD8α− DCs from both mouse strains (Fig. 4B, 4C). However, there was an increase in IRF8 mean fluorescence intensity (MFI) in CD8α+ DCs in both mouse strains. Furthermore, IRF8 MFI and percent positive populations were increased when comparing CD8α+ DCs from Ex3DC−/− relative to Ex3fl/fl strains (Fig. 4B–D). To confirm that the effect of β-catenin stabilization was specific for IRF8, we examined expression of IRF4 and found it to be unchanged in Ex3DC−/− relative to Ex3fl/fl CD8α+ splenic DCs (Fig. 4E). Consistent with this finding, levels of splenic CD4+ DCs and intestinal CD11b+CD103+ DCs, both of which are known to depend on IRF4 for development, were unchanged between Ex3fl/fl and Ex3DC−/− mice (6, 26) (Fig. 3J, 3L; data not shown).

FIGURE 4.
FIGURE 4.

β-Catenin signaling controls Irf8 expression. (A) Semiquantitative PCR analysis of Nfil3, Batf3, Id2, and Irf8 mRNA in CD11c+ splenocytes magnetically purified from naive Ex3fl/fl and Ex3DC−/− mice. mRNA levels were normalized to GAPDH. Data are representative of two independent experiments (n = 2–3 mice/group). (B) Representative flow cytometric plots of IRF8 expression by Ex3fl/fl and Ex3DC−/− CD8α− and CD8α+ splenic DCs. (C) MFI of IRF8 within Ex3fl/fl and Ex3DC−/− CD8α− and CD8α+ DCs, and (D) the percent of CD8α+ DCs expressing IRF8 are shown. Dots represent results from individual mice. Data are the combined results of two experiments, and the experiment was independently performed at least three times (n = 4–5 mice/group). (E) Representative FACS plot of IRF4 expression and IRF4 MFI in Ex3fl/fl and Ex3DC−/− CD11c+ splenocytes. Data are representative of three independent experiments (n = 4 mice/group). (F) Chromatin immunoprecipitation of naive Ex3DC−/− Flt3L DC cultures with control IgG or β-catenin Ab followed by quantitative PCR to determine Irf8 promoter occupancy. DNA levels were normalized to 1% input chromatin. Data are representative of two independent experiments. (G) Quantitative PCR analysis of Axin2 and Irf8 gene expression in BMDCs after 5-h culture with DMSO or ICG-001. Fold change is relative to DMSO control. Data are from one independent trial. (H and I) Intracellular expression of IRF8 and β-catenin after ICG-001 treatment of BMDCs (H) or MutuDC1940 cells (I). Data are representative of two (MutuDC1940 cells) and four (BMDC) independent experiments with three replicates per treatment per experiment. *p < 0.05, **p < 0.01, ***p < 0.001.

To ask whether β-catenin targets the Irf8 promoter in vivo, we performed chromatin immunoprecipitation assays on Flt3 ligand cultures of bone marrow DCs derived from Ex3DC−/− mice. β-Catenin displayed a >6-fold enrichment in Irf8 promoter occupancy over IgG control, suggesting a direct role for this signaling axis in Irf8 transcription (Fig. 4F). To ask whether Irf8 transcription could be blocked by inhibiting β-catenin, we cultured WT BMDCs with the Wnt/β-catenin inhibitor ICG-001, which competes with β-catenin for binding to its cofactor Creb-binding protein (CBP) (28). As a control, 5 h of ICG-001 treatment led to a significant reduction in transcript levels of the known β-catenin target gene Axin2 (29). Importantly, Irf8 transcripts were also significantly reduced after ICG-001 treatment (Fig. 4G). This downregulation was observed at the protein level, because IRF8 expression in BMDCs was markedly decreased compared with DMSO-treated control cells by flow cytometry (Fig. 4H). Furthermore, ICG-001 treatment of MutuDC1940 cells, a DC-derived cell line that has many characteristics of CD8α+ DCs (30), also resulted in strong downregulation of IRF8 (Fig. 4I). IRF4 levels were unchanged by ICG-001 treatment, confirming the specificity of β-catenin signaling for IRF8 (data not shown). These data establish a functional link between β-catenin and IRF8 expression that controls differentiation of CD8α+ DCs, pDCs, and peripheral CD103+ DCs.

β-Catenin stabilization enhances IL-12 production by CD8α+ DCs

CD8α+ DCs are a potent IL-12 source during infection with the Th1 pathogen Toxoplasma gondii (15). Furthermore, it was recently shown that this cytokine activity requires IRF8 (31). In addition, LPS has been shown to upregulate Irf8 expression, resulting in binding to the IL-12 promoter (32). Therefore, we wanted to determine whether increased IRF8 expression in CD8α+ DCs from Ex3DC−/− mice would impact their functional activity, in particular as related to IL-12 production.

To examine this, we cultured whole splenocytes from Ex3fl/fl and Ex3DC−/− mice in the presence of media, LPS, or STAg, an antigenic preparation of T. gondii that stimulates CD8α+ DC IL-12 through the interaction between TLR11/12 and parasite profilin (31). Indeed, upon LPS or STAg stimulation, supernatants from Ex3DC−/− splenocytes contained significantly increased levels of IL-12p40 compared with Ex3fl/fl controls (Fig. 5A). CD11c+ cells were magnetically purified (∼90% purity) from Ex3fl/fl and Ex3DC−/− splenocytes and cultured with LPS and STAg to confirm that the source of IL-12 was DCs. As expected, increased IL-12 secretion was again observed in Ex3DC−/− cells (Fig. 5B). These results clearly show that DCs from Ex3DC−/− produce more IL-12 than cells from WT littermates. However, they leave open to question whether this is a result of an increase in the proportion of CD8α+ DCs or whether Ex3DC−/− CD8α+ DCs produce increased IL-12 on a cell-to-cell basis relative to corresponding cells from Ex3fl/fl mice. Therefore, CD8α+ and CD8α− DCs were magnetically purified from Ex3fl/fl and Ex3DC−/− spleens and stimulated in vitro with STAg. Although IL-12 production was restricted to the CD8α+ subset, the Ex3DC−/− CD8α+ DCs secreted enhanced IL-12 levels compared with Ex3fl/fl CD8α+ DCs (Fig. 5C). Next, WT splenocytes were treated with the β-catenin inhibitor ICG-001 and then stimulated with STAg or LPS overnight. Inhibitor-treated cells secreted lower levels of IL-12p40 compared with DMSO-treated cells without negatively affecting viability (Fig. 5D, data not shown). Furthermore, the elevated IL-12 secretion displayed by Ex3DC−/− splenocytes over Ex3fl/fl splenocytes could be inhibited by treatment with ICG-001, further indicating that ICG-001 treatment suppresses β-catenin–dependent responses (Fig. 5E). To further implicate CD8α+ DCs in these findings, we replicated these experiments in vitro using MutuDC1940 cells. Stimulation of MutuDC1940 cells with STAg resulted in extremely high IL-12 levels, consistent with their origin from splenic CD8α+ DCs. Furthermore, pretreatment of MutuDC1940 cells with ICG-001 significantly impaired the IL-12 response to STAg (Fig. 5F). Thus, β-catenin stabilization promotes both differentiation and IL-12–secreting capacity of CD8α+ DCs by promoting increased Irf8 expression.

FIGURE 5.
FIGURE 5.

β-Catenin stabilization enhances IL-12 production by CD8α+ DCs. (A) IL-12p40 production by naive Ex3fl/fl and Ex3DC−/− splenocytes stimulated in vitro with LPS, STAg, or media control measured by ELISA. (B) IL-12p40 production by splenic CD11c+ DCs magnetically purified from Ex3fl/fl and Ex3DC−/− mice stimulated in vitro with LPS, STAg, or media control measured by ELISA. (C) IL-12p40 production by CD8α+ and CD8α− DCs purified from naive Ex3fl/fl and Ex3DC−/− splenocytes after in vitro stimulation with media or STAg for 48 h measured by ELISA. (D) IL-12p40 secretion by Ex3fl/fl splenocytes pretreated with ICG-001 for 5 h and then stimulated overnight with LPS or STAg measured by ELISA. (E) IL-12p40 production by splenocytes (106) from Ex3DC−/− mice cultured for 5 h with 5 μM ICG-001 or DMSO and then stimulated with media, LPS (100 ng/ml), or STAg (25 μg/ml) overnight. (F) IL-12p40 production by MutuDC1940 cells (105) pretreated with 20 μM ICG-001 or DMSO for 2 h and then stimulated with media or STAg (25 μg/ml) overnight. Data are representative of at least three (A and F) and four (B–E) independent experiments, each involving three to five mice per group, except (C), which used pooled samples from three mice per experiment. *p < 0.05, **p < 0.01, ***p < 0.001.

Constitutive DC β-catenin signaling promotes Th1 immunity during Toxoplasma gondii infection

As a source of IL-12 that drives Th1 activation, CD8α+ DCs are required to control infection with Toxoplasma, yet overexpression of IL-12 and downstream proinflammatory cytokines is also lethal (15, 33). Therefore, we used i.p. infection with T. gondii to evaluate the impact of DC β-catenin stabilization on host immunity. Ex3DC−/− mice began to succumb within 9 d of low-dose T. gondii i.p. infection, whereas the Ex3fl/fl littermate controls fully survived acute infection (Fig. 6A). Parasite levels in the spleen (Fig. 6B) and peritoneal cavity (data not shown) were equivalent between the genotypes, arguing that susceptibility of Ex3DC−/− mice was not due to defective control of Toxoplasma.

FIGURE 6.
FIGURE 6.

Constitutive DC β-catenin signaling increases the proinflammatory cytokine response to Toxoplasma. (A) Survival of Ex3fl/fl and Ex3DC−/− mice post i.p. infection with Toxoplasma type II strain ME49 (25 cysts; n = 4–6 mice/group). Data are representative of at least three experiments. (B) Quantitative PCR amplification of parasite (B1 gene) and host DNA (argininosuccinate lyase gene) isolated from Ex3fl/fl and Ex3DC−/− spleens 9 d postinfection. Parasite load is displayed as a ratio of parasite genomes to host genomes (n = 3–4 mice/group). (C) IL-12p40 production by CD11c+ DCs magnetically separated from day 6 postinfection Ex3fl/fl and Ex3DC−/− splenocytes and cultured overnight without additional stimulation (n = 3 mice/group). (D) IL-12p70 production by bulk splenocytes from day 6 postinfection Ex3fl/fl and Ex3DC−/− mice (n = 3 mice/group). Data are representative of two independent experiments. (E) IL-12p40 and IFN-γ production by splenocytes from day 10 postinfection Ex3fl/fl and Ex3DC−/− mice cultured for 72 h without additional stimulation (n = 3–5 mice/group). Data are representative of three independent experiments. (F) IL-12p40, IFN-γ, and TNF-α levels in serum collected from day 9 postinfection Ex3fl/fl and Ex3DC−/− mice (n = 3–5 mice/group). Data are representative of two independent experiments. Means and SE of individual mice are shown. *p < 0.05, **p < 0.01.

Splenic DCs from infected Ex3DC−/− mice secreted dramatically more IL-12 compared with Ex3fl/fl controls as measured by both p40 (Fig. 6C) and p70 (Fig. 6D) subunits, and at nearly 50 ng/ml, the level of IL-12p40 detected in Ex3DC−/− mice was ∼5-fold over WT controls. To examine Th1 responses in these animals, we cultured Ex3fl/fl and Ex3DC−/− splenocytes from infected mice in vitro, and the supernatants were assayed for both IL-12 and IFN-γ. Accompanying increased IL-12 secretion by Ex3DC−/− splenocytes, IFN-γ levels were enhanced, suggesting elevated T and possibly NK cell activation in Ex3DC−/− mice in response to T. gondii infection (Fig. 6E). Furthermore, serum collected from infected mice revealed significantly increased levels of IFN-γ, IL-12, and TNF-α in Ex3DC−/− mice compared with Ex3fl/fl controls (Fig. 6F), suggesting systemic hyperproduction of proinflammatory cytokines upon DC β-catenin stabilization. By contrast, IL-17 was undetectable or expressed at very low levels in splenocyte cultures or serum prepared from infected Ex3fl/fl or Ex3DC−/− mice (data not shown).

IFN-γ production by splenic CD4+ T cells, CD8+ T cells, and NK cells was assayed 8 d postinfection to determine the source of elevated Th1 cytokines in Ex3DC−/− mice during T. gondii infection. Both CD4+ T cells and NK cell populations in Ex3DC−/− mice displayed a marked increase in IFN-γ responses compared with Ex3fl/fl littermate controls (Fig. 7A, 7C). Interestingly, given previous studies implicating CD8α+ DCs in cross-presentation (5, 34), IFN-γ from CD8+ T cells was not significantly changed between the two genotypes (Fig. 7B).

FIGURE 7.
FIGURE 7.

CD4+ T cells and NK cells, but not CD8+ T cells, overproduce IFN-γ post Toxoplasma infection in mice with constitutive DC β-catenin signaling. (A–C) Intracellular flow cytometric analysis of IFN-γ production by (A) CD4+ T cells, (B) CD8+ T cells, and (C) NK cells from day 9 T. gondii–infected Ex3fl/fl and Ex3DC−/− spleens after 4 h of PMA and ionomycin stimulation in the presence of brefeldin A. Shown are representative contour plots of individual mice and the mean ± SE of IFN-γ levels from multiple mice. Data are representative of three independent experiments (CD4 and CD8) and one (NK) experiment. (D and E) Quantification of Toxoplasma-specific (D) Tgd057+CD8+ T cells and (E) CD4Ag28m+CD4+ T cells. Means and SEs of individual mice are shown, and each dot represents a single mouse. (F and G) IFN-γ levels expressed by tetramer+ CD8+ (F) and CD4+ (G) T cells after 4 h of culture with PMA, ionomycin, and brefeldin A. Data are representative of three independent experiments (n = 3 mice/group). *p < 0.05, ***p < 0.001.

We next determined whether the increase in IFN-γ could be explained by increased activation and expansion of Toxoplasma-specific T cells. We used MHCI tetramers that bind CD8+ T lymphocytes specific for the endogenous T. gondii epitope Tgd057 and MHCII tetramers that bind CD4+ T cells specific for the Toxoplasma epitope CD4Ag28m (35, 36). The frequency of parasite-specific CD4+ and CD8+ T cells was equivalent in Ex3DC−/− and Ex3fl/fl mice (Fig. 7D, 7E). Although IFN-γ produced by Ag-specific CD8+ T cells was equivalent in Ex3fl/fl and Ex3DC−/− mice, levels of cytokine produced by tetramer+ CD4+ T lymphocytes were increased in Ex3DC−/− relative to Ex3fl/fl mice (Fig. 7F, 7G). Therefore, in the context of T. gondii infection, DC β-catenin stabilization impacts the intensity of the CD4 and NK cell IFN-γ response but does not have a measurable influence on IFN-γ production by CD8+ T cells.

DC β-catenin signaling enhances CD8+ T cell priming during VACV infection

DC-mediated cross-presentation of exogenous Ag through the cytosolic pathway is crucial for the generation of a CD8+ T cell response against VACV infection (16, 37). Thus, we finally asked whether DC β-catenin signaling influenced DC cross-presentation in the context of VACV infection. To address this, we infected Ex3fl/fl and Ex3DC−/− mice i.p. with recombinant VACV expressing the HSV glycoprotein B (gB) peptide (VACV-gB) (17). On day 6 postinfection, spleens were harvested, and gB-specific CD8+ T cells were quantified by flow cytometry using MHCI-restricted gB-8p:Kb tetramers. We found a striking increase in the percentage and total number of gB-specific CD8+ T cells in Ex3DC−/− mice compared with littermate controls, suggesting increased cross-presentation by β-catenin–stabilized DCs (Fig. 8A–C). Furthermore, restimulation of splenocytes with gB peptide revealed significantly increased IFN-γ+CD8+ T cells in Ex3DC−/− mice, demonstrating a functional impact of DC β-catenin activation on CD8+ T cell activity (Fig. 8D–F). These results demonstrate that stabilization of DC β-catenin promotes development of CD8+ T cell responses in the context of viral infection.

FIGURE 8.
FIGURE 8.

β-Catenin stabilization promotes activation of Ag-specific CD8+ T cells during viral infection. (A–C) Quantification of gB-8p–specific CD8+ T cells in Ex3fl/fl and Ex3DC−/− spleens 6 d post VACV-gB infection. (A) Representative plots of MHCI-restricted gB-8p:Kb tetramer+ CD8+ T cells, and (B) percentage and (C) total number of gB-8p:Kb+CD8+ T cells. (D–F) IFN-γ production by gB-8p–specific CD8+ T cells in Ex3fl/fl and Ex3DC−/− spleens 6 d post VACV-gB infection. (D) Representative plots of CD8+CD44+IFN-γ+ T cells after gb-8p restimulation of splenocytes, and (E) percentage and (F) total number of gB-8p–specific CD8+CD44+IFN-γ+ T cells. Data shown are the combination of two independent experiments (n = 9–13). **p < 0.01.

Discussion

DC differentiation is a complex process involving an array of transcription factors and growth factor cytokines whose details are continuing to be elucidated. In this study, we identify an unexpected role for β-catenin in controlling differentiation of CD8α+ DCs, pDCs, and developmentally related nonlymphoid CD103+ DCs. This pattern of transcriptional control precisely matches that of IRF8 (24, 26, 38). In support of a functional connection between β-catenin and IRF8, overexpression of β-catenin increased steady-state IRF8 levels in CD8α+ DCs. In addition, IL-12 production by CD8α+ DCs, a known function of this cell subset, was prevented by β-catenin inhibition concomitant with IRF8 downmodulation, and CD8α+ DCs overexpressing β-catenin produced enhanced IL-12 in response to microbial stimulation. During infection with T. gondii, a prototypic type 1–inducing pathogen, mice overexpressing DC-specific β-catenin developed exacerbated Th1 responses culminating in early death. In addition, infection with VACV increased expansion and IFN-γ production by virus-specific CD8+ T lymphocytes.

Wnt/β-catenin signaling has a well-studied role in embryogenesis and tumorigenesis (10). It is now also clear that differentiation of T cells and NK cells requires LEF-1/TCF β-catenin cofactors. Furthermore, β-catenin regulates proliferation of pro-B cells (39). Using a β-catenin stabilization approach similar to that used in this study, it has been found that this signaling molecule plays a fundamental role in regulating hematopoietic stem cells in the bone marrow (40). Furthermore, β-catenin has been recently shown to target Irf8 during hematopoiesis to alter granulocyte development (41). We now show that stabilization of β-catenin at the CD11c-expressing stage has unanticipated effects on generation of mature DC subsets whose differentiation depends upon IRF8.

Our data are notable because other recent studies have implicated β-catenin signaling in promoting tolerogenic DC phenotypes. For example, in contrast with activation by microbial stimulation, cluster disruption of BMDCs activates β-catenin, endowing the cells with the ability to promote regulatory T cells that protect against experimental autoimmune encephalitis (12). In the intestine, β-catenin expression by CD11c+ cells was shown to be required for regulatory T cell induction and production of anti-inflammatory factors, including retinoic acid–metabolizing enzymes, TGF-β, and IL-10 (13). In this case, absence of β-catenin in DCs increased sensitivity to dextran sodium sulfate–mediated colitis. Our data uncover an important new facet of β-catenin signaling because they reveal a role in promoting the differentiation of IRF8-dependent CD8α+ DCs with enhanced proinflammatory activity.

IRF8 controls cDC formation by actively suppressing granulocyte differentiation while simultaneously promoting commitment to the DC lineage (42, 43). This is best illustrated by the observation that IRF8-deficient mice suffer from chronic myeloid leukemia-like syndrome with massive expansion of granulocytic cells (44). As such, IRF8 expression is enriched in CDPs over CMPs and is nearly absent in GMPs (43), which is consistent with our finding that splenic DC precursors express β-catenin. Mice expressing constitutive β-catenin in the hematopoietic stem cell compartment, either through exon 3 deletion or transgenic expression of a phosphorylation-resistant β-catenin, display a differentiation block at the granulocyte progenitor stage, which leads to lethality of the host (40, 45). It is tempting to speculate, therefore, that overactivation of β-catenin may lead to excessive IRF8 expression as early as the progenitor stage, resulting in suppression of granulocytic precursors and the subsequent direct promotion of DC development. This is supported by earlier findings that showed impaired granulocyte formation after β-catenin activation in hematopoietic stem cells (41). Further, constitutive DC β-catenin signaling in our study appeared to push differentiation toward mature DC subsets by depleting early progenitor precDC pools. Future studies should identify the activating Wnt ligand responsible for driving DC differentiation and suppressing granulocyte formation during hematopoiesis.

Several lines of evidence indicate a role for CD8α+ DCs in Ag cross-presentation and activation of CD8+ T lymphocytes (5, 34). Toxoplasma is well-known for its ability to elicit potent CD8+ T cell responses, and cross-presentation has previously been found to play a role in MHC class I presentation to CD8+ T cells during infection with this intracellular protozoan (46, 47). Therefore, it was initially surprising that there was no indication of abnormally strong CD8+ T cell responses in T. gondii–infected Ex3DC−/− mice that overexpress CD8α+ DCs. However, the recent discovery that the parasite directly injects a subset of secretory proteins into the host cell cytoplasm (48), as well as evidence that the majority of T cell activation is stimulated by actively infected DCs (49), argues that conventional presentation rather than cross-presentation may be the dominant mechanism for CD8+ T cell priming during T. gondii infection.

In contrast with normal CD8+ T cell responses post Toxoplasma infection in mice with DC-specific β-catenin activation, there was a clear increase in CD4+ T cell, and to a lesser extent NK cell, IFN-γ production in the mutant mice. This is of interest because CD4+ and CD11b+ DCs, which are believed to be the most adept at activating CD4+ T cells (50), were unchanged or even reduced in Ex3DC−/− mice. Therefore, it is most likely that increased CD4+ T and NK cell IFN-γ resulted from increased splenic CD8α+ DC activity, and that failure to activate CD8+ T cells was due to the lack of cross-presentation. This is supported by the finding that virus-specific CD8 T cell responses were enhanced during VACV infection, which has been shown to require the cross-presentation pathway for CD8+ T cell activation.

To our knowledge, our findings uncover for the first time a role for stabilized β-catenin signaling in promoting DC subset differentiation and activity. Based upon previous findings, it has been suggested that exploiting strategies that activate β-catenin signaling in DCs might be useful in the control of inflammatory and autoimmune diseases (12, 13). Our study throws a cautionary light on this approach, showing that constitutive β-catenin signaling promotes differentiation of IRF8-dependent DCs resulting in increased proinflammatory responses during protozoan and viral infection.

Disclosures

The authors have no financial conflicts of interest.

Acknowledgments

We thank M. Hossain for technical assistance.

Footnotes

  • This work was supported by National Institutes of Health Grants AI109061 (to E.Y.D.), AI083405 (to G.S.Y.), and AI110613 (to B.D.R.) and a scholarship from the Cornell Vertebrate Genomics group (to S.B.C.).

  • The online version of this article contains supplemental material.

  • Abbreviations used in this article:

    BMDC
    bone marrow–derived DC
    cDC
    conventional DC
    CDP
    common dendritic progenitor
    CMP
    common myeloid progenitor
    DC
    dendritic cell
    gB
    glycoprotein B
    GMP
    granulocyte-macrophage progenitor
    IRF
    IFN regulatory factor
    MFI
    mean fluorescence intensity
    pDC
    plasmacytoid DC
    precDC
    preconventional DC
    STAg
    soluble tachyzoite Ag
    VACV
    vaccinia virus
    VACV-gB
    VACV expressing MHC class I–restricted HSV glycoprotein B (gB) peptides 498–505
    WT
    wild-type.

  • Received September 25, 2014.
  • Accepted October 31, 2014.

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