Dendritic cells (DCs) consist of various subsets that play crucial roles in linking innate and adaptive immunity. In the murine spleen, CD8α+ DCs exhibit a propensity to ingest dying/dead cells, produce proinflammatory cytokines, and cross-present Ags to generate CD8+ T cell responses. To track and ablate CD8α+ DCs in vivo, we generated XCR1-venus and XCR1-DTRvenus mice, in which genes for a fluorescent protein, venus, and a fusion protein consisting of diphtheria toxin receptor and venus were knocked into the gene locus of a chemokine receptor, XCR1, which is highly expressed in CD8α+ DCs. In both mice, venus+ cells were detected in the majority of CD8α+ DCs, but they were not detected in any other cells, including splenic macrophages. Venus+CD8α+ DCs were superior to venusCD8α+ DCs with regard to their cytokine-producing ability in response to TLR stimuli. In other tissues, venus+ cells were found primarily in lymph node (LN)-resident CD8α+, LN migratory and peripheral CD103+ DCs, which are closely related to splenic CD8α+ DCs, although some thymic CD8αCD11b and LN CD103CD11b DCs were also venus+. In response to dsRNAs, diphtheria toxin–treated XCR1-DTR mice showed impaired CD8+ T cell responses, with retained cytokine and augmented CD4+ T cell responses. Furthermore, Listeria monocytogenes infection and anti–L. monocytogenes CD8+ T cell responses were defective in diphtheria toxin–treated XCR1-DTRvenus mice. Thus, XCR1-expressing DCs were required for dsRNA- or bacteria-induced CD8+ T cell responses. XCR1-venus and XCR1-DTRvenus mice should be useful for elucidating the functions and behavior of XCR1-expressing DCs, including CD8α+ and CD103+ DCs, in lymphoid and peripheral tissues.

Dendritic cells (DCs) are specialized APCs that play crucial roles in linking innate and adaptive immunity (1). Critical in vivo roles for DCs in various immune responses or the pathogenesis of immune disorders have been clarified by analyzing mutant mice in which diphtheria toxin receptor (DTR) and diphtheria toxin (DT) A subunit are expressed under the control of the CD11c promoter (26). Those mice also enabled us to clarify homeostatic roles for DCs in lymphocyte homing to lymph nodes (LNs) or preventing autoimmunity (57). However, because DCs are heterogeneous and can be divided into several subsets according to function and expression patterns of cell surface molecules, a DC subset–specific ablation system should be generated.

Under steady-state conditions, murine splenic DCs consist of B220+CD11cdull plasmacytoid DCs (pDCs) and B220CD11c+ DCs. In the spleen, all DCs are resident DCs derived from blood precursors, and B220CD11c+ DCs can be divided into CD8α+CD11b and CD8αCD11b+ DCs. In lymphoid tissues, DCs consist of resident and migratory DCs, which can be defined as MHC class II (MHC-II)intCD11c+ and MHC-IIhighCD11c+ DCs, respectively (8, reviewed in Ref. 9). As in the spleen, resident DCs consist mainly of CD8α+CD11b and CD8αCD11b+ cells. Migratory DCs are derived from peripheral tissues, such as skin or lamina propria, and can be divided into several subsets depending on the expression patterns of CD103 and CD11b.

Among these DC subsets, resident CD8α+ DCs and migratory CD103+ DCs are characterized by their ability to take up apoptotic cells and to cross-present soluble and cell-associated Ags for generating CD8+ cytotoxic T cells (1012, reviewed in Ref. 13). This cross-presenting activity is important for establishing antiviral or antitumor immunity. Cross-presentation can be facilitated by targeting Ags to several C-type lectins, such as DEC-205 (CD205), Clec9a, and Langerin (CD207), expressed on these DC subsets (1416). Splenic CD8α+ DCs also show a propensity to produce proinflammatory cytokines in response to a LPS sensor, TLR4, and nucleic acid sensors, TLR3 and TLR9. It is important to clarify the in vivo roles of this DC subset.

Several mutant mice lacking this DC subset have been established and analyzed. Although the mutant mice lacking a transcription factor, IRF-8, show ablation of both CD8α+ DCs and pDCs, spontaneous mutant mice (BXH2 mice) carrying a point mutation on the Irf8 gene lack only CD8α+ DCs (17, 18). CD8α+ DCs are also ablated in the mutant mice lacking a basic leucine zipper transcription factor, ATF-like 3 (BATF3) (19). Both transcription factors are essential for the development of splenic CD8α+ DCs and their equivalent CD103+ DCs in the periphery (13, 20, 21). Batf3-deficient mice have been studied intensively; their in vivo CD8+ T cell responses against virus, bacteria, parasites, and tumors are severely impaired (2224). Ablation of splenic CD8α+ DCs can also be achieved by injection of a proapoptotic reagent, cytochrome C. Cytochrome C–injected mice show impairment of CD8+ T cell responses against soluble or cell-associated Ags and subsequent immunity to tumor challenge (25). These findings suggest that CD8α+ DCs are critical for cross-presentation. However, IRF-8 is also expressed in B cells and myeloid cells, such as monocyte/macrophages (MΦs), and BXH2 mice manifest an expansion of myeloid cells. BATF3 expression is not specific to CD8α+ DCs; it is also detected in a CD11b+ DC subset (19). In addition, cytochrome C may have some detrimental effects on other DC subsets or MΦs, although it does not make them apoptotic. Therefore, it is difficult to rule out the possibility that other cells besides CD8α+ and CD103+ DCs are functionally attenuated in these mice.

A chemokine receptor, XC chemokine receptor 1 (XCR1), is selectively expressed in splenic CD8α+ and peripheral CD103+ DCs but is rare in other DC subsets or lymphocytes (2631). Human XCR1 expression is restricted to CD141+ DCs, which correspond to murine splenic CD8α+ DCs (2729, 32, 33). Thus, XCR1 is a suitable marker for defining this DC subset in both humans and mice. To generate the mutant mice in which this DC subset can be specifically tracked and deleted, we knocked a gene encoding a YFP derivative, venus (34), or a fusion protein consisting of DTR and venus (DTRvenus) into the Xcr1 locus. In these knock-in mice, designated as XCR1-venus or XCR1-DTRvenus mice, venus fluorescence signal was detected mainly in resident CD8α+CD11b and migratory CD103+CD11b DCs in lymphoid tissues and CD103+CD11b DCs in peripheral tissues. Upon DT injection to XCR1-DTRvenus mice, DTRvenus-expressing cells were transiently and efficiently ablated. Using XCR1-DTRvenus mice, we investigated the in vivo roles of XCR1-expressing DCs in CD8+ T cell responses.

Targeting vectors were designed to replace the entire murine Xcr1 coding sequences with a gene encoding venus or DTRvenus (Figs. 1A, 4A). The genes carry a polyadenylation signal derived from bGHpA. A neomycin resistance gene (neo) driven by the MC1 promoter and flanked by bacteriophage P1 loxP and yeast FRT sequence was used as a selection marker. An HSV thymidine kinase gene (HSV-TK) was inserted for negative selection. To construct the DTRvenus cassette, genes for DTR and venus were amplified from HB-EGF cDNA (kindly provided by Dr. Eisuke Mekata, Osaka University) and venus/pCS2 vector (kindly provided by Dr. Atsushi Miyawaki, RIKEN Brain Science Institute, Wako, Japan), respectively, and the 3′ end of human DTR open reading frame was linked in frame with a gene encoding venus. A C57BL/6-derived embryonic stem (ES) cell line, Bruce4 (kindly provided by Drs. Colin L. Stewart, Institute of Medical Biology, Singapore, and Masaki Hikida, Kyoto University, Kyoto, Japan), was transfected with the linearized targeting vector by electroporation and selected with G418 (Nacalai Tesque) and ganciclovir (Mitsubishi Tanabe Pharma). Doubly resistant clones were screened for homologous recombination by PCR and verified by Southern blot analysis. Germline-transmitting chimeras were generated by injection of targeted ES clones to blastocysts from BALB/c mice and bred with C57BL/6J mice. Obtained Xcr1+/venus and Xcr1+/DTRvenus mice were backcrossed with C57BL/6J mice for an additional two to six or two to four generations, respectively. C57BL/6J or Xcr1+/+ littermates were used as wild-type (WT) mice.

FIGURE 1.

Expression of venus in XCR1-venus mice. (A) Schematic diagrams of the mouse Xcr1 WT allele, a targeting vector, and venus knocked-in allele. Filled and open boxes denote coding and noncoding exons of Xcr1, respectively. (B) Southern blot analysis of Xcr1 WT (+/+), Xcr1+/venus (+/v), and Xcr1venus/venus (v/v) mice. Genomic DNAs were isolated from mice tails, digested with EcoRI and EcoRV, electrophoresed, and hybridized with a radiolabeled probe indicated in (A). Southern blot gave a 14.3- and a 7.5-kbp band for WT and knock-in allele, respectively. (C) Splenocytes from WT or XCR1-venus mice were stained with Abs against the indicated cell surface markers. Whole or CD19 splenocytes are shown. C57BL/6J mice were used as WT mice. (D) Venus expression was analyzed in B cells, T cells, NK cells, granulocytes, and monocytes in the spleen of XCR1-venus mice. Each population was defined as indicated. The number in each panel indicates the percentage of venus+ cells. (E) Splenic DCs from XCR1-venus mice were left unstimulated (medium) or were stimulated with 50 μg/ml of poly(I:C) or 1 μM of CpG DNA for 6 h. IL-12p40 production from PDCA-1CD11c+ or PDCA-1CD8α+CD11c+ cells was analyzed by intracellular staining with anti–IL-12p40 Ab and FACS. In (C) and (E), the numbers represent the percentage of the cells within the indicated gates or each quadrant; data are representative of at least two independent experiments. EI, EcoRI; EV, EcoRV.

FIGURE 1.

Expression of venus in XCR1-venus mice. (A) Schematic diagrams of the mouse Xcr1 WT allele, a targeting vector, and venus knocked-in allele. Filled and open boxes denote coding and noncoding exons of Xcr1, respectively. (B) Southern blot analysis of Xcr1 WT (+/+), Xcr1+/venus (+/v), and Xcr1venus/venus (v/v) mice. Genomic DNAs were isolated from mice tails, digested with EcoRI and EcoRV, electrophoresed, and hybridized with a radiolabeled probe indicated in (A). Southern blot gave a 14.3- and a 7.5-kbp band for WT and knock-in allele, respectively. (C) Splenocytes from WT or XCR1-venus mice were stained with Abs against the indicated cell surface markers. Whole or CD19 splenocytes are shown. C57BL/6J mice were used as WT mice. (D) Venus expression was analyzed in B cells, T cells, NK cells, granulocytes, and monocytes in the spleen of XCR1-venus mice. Each population was defined as indicated. The number in each panel indicates the percentage of venus+ cells. (E) Splenic DCs from XCR1-venus mice were left unstimulated (medium) or were stimulated with 50 μg/ml of poly(I:C) or 1 μM of CpG DNA for 6 h. IL-12p40 production from PDCA-1CD11c+ or PDCA-1CD8α+CD11c+ cells was analyzed by intracellular staining with anti–IL-12p40 Ab and FACS. In (C) and (E), the numbers represent the percentage of the cells within the indicated gates or each quadrant; data are representative of at least two independent experiments. EI, EcoRI; EV, EcoRV.

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FIGURE 4.

Depletion of XCR1+ cells in XCR1-DTRvenus mice. (A) Schematic diagrams of the mouse Xcr1 WT allele, a targeting vector, and venus knocked-in allele. Filled and open boxes denote coding and noncoding exons of Xcr1, respectively. (B) Southern blot analysis of XCR1 WT (+/+) and Xcr1+/DTRvenus mice. Genomic DNAs were isolated from mice tails, digested with HincII and PstI, electrophoresed, and hybridized with a radiolabeled probe indicated in (A). Southern blot gave a 4.3- and a 3.1-kbp band for WT and knock-in allele, respectively. (C) XCR1-DTRvenus mice were injected i.p. with DT and analyzed on the indicated days. Whole splenocytes were analyzed for the expression of B220 and CD11c, and the percentages of pDCs (B220+CD11cdull) and conventional DCs (B220CD11c+) are shown (upper panels). Expression of CD8α and DTRvenus in B220CD11c+ DCs was analyzed, and the percentages of cells within the indicated quadrants are shown (lower panels). (D) Numbers and percentages of DTRvenus+ cells in the MHC-II+CD11c+ population from the spleen or in migratory and resident DCs are shown. Cells were prepared from the indicated lymphoid tissues at the indicated days after DT injection. The line connects the mean of each day (n = 6 for days 0, 1, 2; n = 5 for day 4; n = 3 for day 8). (E) Spleen or inguinal LN cryosections from PBS- or DT-treated XCR1-DTRvenus mice were stained with Abs against GFP (DTRvenus, green) and the indicated molecules. Scale bar, 100 μm. (F) LN cells from PBS- or DT-treated XCR1-DTRvenus mice were stained with Abs against the indicated surface molecules. Whole LN cells were gated on CD169+CD11c (R1), CD169+CD11clow (R2), and CD169+CD11chigh (R3) and analyzed for DTRvenus expression. Numbers in the dot plots and the graphs indicate the percentages of cells in each gate and DTRvenus+ cells, respectively. Data in (C)–(F) are representative of at least two independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001. H, HincII; P, PstI.

FIGURE 4.

Depletion of XCR1+ cells in XCR1-DTRvenus mice. (A) Schematic diagrams of the mouse Xcr1 WT allele, a targeting vector, and venus knocked-in allele. Filled and open boxes denote coding and noncoding exons of Xcr1, respectively. (B) Southern blot analysis of XCR1 WT (+/+) and Xcr1+/DTRvenus mice. Genomic DNAs were isolated from mice tails, digested with HincII and PstI, electrophoresed, and hybridized with a radiolabeled probe indicated in (A). Southern blot gave a 4.3- and a 3.1-kbp band for WT and knock-in allele, respectively. (C) XCR1-DTRvenus mice were injected i.p. with DT and analyzed on the indicated days. Whole splenocytes were analyzed for the expression of B220 and CD11c, and the percentages of pDCs (B220+CD11cdull) and conventional DCs (B220CD11c+) are shown (upper panels). Expression of CD8α and DTRvenus in B220CD11c+ DCs was analyzed, and the percentages of cells within the indicated quadrants are shown (lower panels). (D) Numbers and percentages of DTRvenus+ cells in the MHC-II+CD11c+ population from the spleen or in migratory and resident DCs are shown. Cells were prepared from the indicated lymphoid tissues at the indicated days after DT injection. The line connects the mean of each day (n = 6 for days 0, 1, 2; n = 5 for day 4; n = 3 for day 8). (E) Spleen or inguinal LN cryosections from PBS- or DT-treated XCR1-DTRvenus mice were stained with Abs against GFP (DTRvenus, green) and the indicated molecules. Scale bar, 100 μm. (F) LN cells from PBS- or DT-treated XCR1-DTRvenus mice were stained with Abs against the indicated surface molecules. Whole LN cells were gated on CD169+CD11c (R1), CD169+CD11clow (R2), and CD169+CD11chigh (R3) and analyzed for DTRvenus expression. Numbers in the dot plots and the graphs indicate the percentages of cells in each gate and DTRvenus+ cells, respectively. Data in (C)–(F) are representative of at least two independent experiments. *p < 0.05, **p < 0.01, ***p < 0.001. H, HincII; P, PstI.

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C57BL/6J mice were purchased from CLEA Japan. β2m-deficient (B6.129P2-B2mtm1Unc/J) mice and transgenic mice ubiquitously expressing membrane-bound OVA (C57BL/6-Tg(CAG-OVA)916Jen/J [mOVA-Tg]) were purchased from The Jackson Laboratory and crossed to obtain mOVA-Tg:β2m−/− mice, which carry OVA-expressing MHC class I–deficient cells. All mice were bred and maintained in the Animal Facility of RIKEN Research Institute for Allergy and Immunology (Yokohama, Japan) and the Animal Resource Center for Microbial Diseases, Research Institute for Infectious Diseases, Osaka University (Suita, Japan) under specific pathogen–free conditions and were used at 7–14 wk of age under institutional guidelines of RIKEN Institute and Osaka University. All animal experiments were approved by the Animal Research Committees of RIKEN Yokohama Research Institute and Osaka University.

DT was purchased from Sigma-Aldrich. LPS-free OVA protein and OVA 257–264 peptide (SIINFEKL; OVA-I peptide) were purchased from Worthington Biochemical and Toray Research Center, respectively. Polyinosinic-polycytidylic acid [poly(I:C)] was purchased from GE Healthcare Life Sciences. ODN1668 (5′-TCCATGACGTTCCTGATGCT-3′, with phosphorothioate backbone) was synthesized by Hokkaido System Science and used as a TLR9 ligand, CpG DNA. LPS (O55:B5) was purchased from Sigma-Aldrich.

DT was injected i.p. to the mice at a dose of 25 ng/g body weight. PBS was used as control vehicle. The dose of DT was determined based on the deletion efficiency after injection of graded doses of DT (data not shown).

Spleen, thymus, skin-draining LNs (SDLNs; including inguinal, axillary, and brachial LNs), and mesenteric LNs (mLNs) were digested with 400 Mandl units/ml Collagenase D (Roche) at 37°C for 20 min. EDTA (5 mM) was added for the last 5 min. To prepare DCs in the skin, epidermal sheets were separated from ear skin by digestion with 2 mg/ml Dispase II (Roche) for 2.5 h at 37°C. Epidermal sheets and dermal tissues were cut into small pieces and digested with 400 U/ml Collagenase D for 30 min at 37°C. EDTA (5 mM) was added for the last 5 min.

For isolation of lamina propria DCs, fat and Peyer’s patches were removed from the small intestine, which were opened longitudinally and stirred in RPMI 1640 containing 2% FCS, 2 mM EDTA for 20 min at 37°C to remove epithelial cells and intraepithelial lymphocytes. After additional stirring in RPMI 1640 supplemented with 2% FCS, intestinal tissues were minced and stirred in 170 U/ml Collagenase (Wako Pure Chemical Industries) and 10 μg/ml DNase I (Sigma-Aldrich) for 20 min at 37°C, and floating cells were collected. Collagenase digestion was repeated three times. Low-density cells were enriched by centrifugation with 40 and 75% Percoll (GE Healthcare) density gradient.

To obtain monocyte-derived DCs, bone marrow (BM) monocytes (MHC-IILy6GCD11cLy6C+CD11b+) were sorted and cultured with RPMI 1640 containing 5% FCS, 100 μM 2-ME, penicillin, streptomycin, 20 ng/ml recombinant mouse GM-CSF (R&D Systems), and 20 ng/ml recombinant mouse IL-4 (BD Biosciences) (35, 36). To obtain BM-derived DCs, BM cells were cultured with 10 ng/ml recombinant mouse GM-CSF or 100 ng/ml recombinant human Flt3 ligand (Flt3L; PeproTech), as described previously (37, 38).

Single-cell suspensions were incubated with anti-CD16/32 Ab (BD Biosciences or eBioscience) to block nonspecific binding of Abs. The cells were stained with fluorochrome-conjugated Abs and biotinylated Abs against mice CD3ε (145-2C11), CD4 (RM4-5), CD8α (53-6.7), CD11b (M1/70), CD11c (N418 or HL3), CD24 (M1/69), B220 (RA3-6B2), CD62L (MEL-14), CD49b (DX5), CD103 (M290), CD206 (C068C2), CD207 (eBioL31), Ly6C (HK1.4), Ly6G (1A8), I-A/I-E (M5/114.15.2), PDCA-1/BST-2 (JF05-1C2.4.1 or eBio927), IFN-γ (XMG1.2), and IL-12p40 (C15.6). Biotinylated Abs were visualized by fluorochrome-conjugated streptavidin. Abs and fluorochrome-conjugated streptavidin were purchased from BD Biosciences, eBioscience, BioLegend, and Miltenyi Biotec. CD206 and CD207 were stained intracellularly using a Cytofix/Cytoperm kit (BD Bioscience). In Figs. 2A, 3, 4D, and 7B, dead cells were excluded by staining with a LIVE/DEAD Fixable Dead Cell Stain Kit (Invitrogen).

FIGURE 2.

Expression of venus in lymphoid and peripheral DCs from XCR1-venus mice. Cells from the indicated lymphoid (A) and peripheral (B, C) tissues in WT or XCR1-venus mice were stained with Abs against the indicated markers. (B) Epidermal LCs and dermal DCs from ear skin were defined as CD207+CD11b+MHC-II+CD11c+ and MHC-II+CD11c+ cells, respectively. (C) MHC-II+CD11c+ cells are shown as lamina propria DCs in the small intestine. The number in each graph indicates the percentage of venus+ cells. The numbers in the dot plots indicate the percentages of the cells within the indicated gates or each quadrant. C57BL/6J mice were used as WT mice. Data are representative of at least two independent experiments.

FIGURE 2.

Expression of venus in lymphoid and peripheral DCs from XCR1-venus mice. Cells from the indicated lymphoid (A) and peripheral (B, C) tissues in WT or XCR1-venus mice were stained with Abs against the indicated markers. (B) Epidermal LCs and dermal DCs from ear skin were defined as CD207+CD11b+MHC-II+CD11c+ and MHC-II+CD11c+ cells, respectively. (C) MHC-II+CD11c+ cells are shown as lamina propria DCs in the small intestine. The number in each graph indicates the percentage of venus+ cells. The numbers in the dot plots indicate the percentages of the cells within the indicated gates or each quadrant. C57BL/6J mice were used as WT mice. Data are representative of at least two independent experiments.

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FIGURE 3.

Expression of venus in DCs induced by bacterial infection or LPS injection. (A) WT or XCR1-venus mice were infected with 5 × 103L.m.-OVA. Two days later, splenocytes were stained with Abs against the indicated molecules. Tip-DCs, defined as Ly6C+CD11b+MHC-II+CD11c+ cells, and Ly6CMHC-II+CD11c+ DCs were monitored for venus expression. (B) WT or XCR1-venus mice were injected i.v. with PBS or 10 μg of LPS. Twenty-four hours later, cells from SDLNs were stained with Abs against the indicated molecules. Mo-DCs, defined as CD206+CD11c+ cells, and CD206CD11c+ cells were monitored for venus expression. (C) MHC-II+CD11c+ in vitro Mo-DCs were analyzed for venus expression. (D) BM-derived DCs were analyzed for venus expression. In Flt3L-induced BM-derived DCs, pDCs, CD24high DCs, and CD11bhigh DCs were identified as B220+CD11clow, B220CD24highCD11blowCD11c+, and B220CD24highCD11blowCD11c+ cells, respectively. Numbers in the dot plots and the graphs represent the percentage of cells within the indicated gates and venus+ cells, respectively. Xcr1+/+ littermates (A, C, D) or C57BL/6J (B) mice were used as WT mice. Data are representative of two independent experiments.

FIGURE 3.

Expression of venus in DCs induced by bacterial infection or LPS injection. (A) WT or XCR1-venus mice were infected with 5 × 103L.m.-OVA. Two days later, splenocytes were stained with Abs against the indicated molecules. Tip-DCs, defined as Ly6C+CD11b+MHC-II+CD11c+ cells, and Ly6CMHC-II+CD11c+ DCs were monitored for venus expression. (B) WT or XCR1-venus mice were injected i.v. with PBS or 10 μg of LPS. Twenty-four hours later, cells from SDLNs were stained with Abs against the indicated molecules. Mo-DCs, defined as CD206+CD11c+ cells, and CD206CD11c+ cells were monitored for venus expression. (C) MHC-II+CD11c+ in vitro Mo-DCs were analyzed for venus expression. (D) BM-derived DCs were analyzed for venus expression. In Flt3L-induced BM-derived DCs, pDCs, CD24high DCs, and CD11bhigh DCs were identified as B220+CD11clow, B220CD24highCD11blowCD11c+, and B220CD24highCD11blowCD11c+ cells, respectively. Numbers in the dot plots and the graphs represent the percentage of cells within the indicated gates and venus+ cells, respectively. Xcr1+/+ littermates (A, C, D) or C57BL/6J (B) mice were used as WT mice. Data are representative of two independent experiments.

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FIGURE 7.

Immune responses against L.m.-OVA infection in XCR1-DTRvenus mice. XCR1-DTRvenus mice received PBS or DT at days −1 and 0 of L.m.-OVA infection. (A) Mice were infected i.v. with the indicated numbers of L.m.-OVA. Three days postinfection, bacterial load in the spleen or liver was evaluated. (B) Depletion of liver DTRvenus-expressing cells. XCR1-DTRvenus mice were injected with PBS or DT at days −1 and 0. One day after the last injection, liver leukocytes were prepared and stained with the indicated Abs. MHC-II+CD11c+ cells were gated, and their expression profiles of CD103 and DTRvenus are shown (left panels). Numbers represent the percentages of the cells in each quadrant. Percentages of DTRvenus+ cells in MHC-II+CD11c+ cells are also shown (right panels). PBS- or DT-treated XCR1-DTRvenus mice were infected i.v. with 1 × 103 (C, D) or the indicated numbers (E, F) of L.m.-OVA and analyzed for CD8+ T cell responses 7 d later. (C and E) H-2Kb/OVA-I tetramer+ cells in CD49bDTRvenusCD8+ splenocytes were monitored by FACS. (E) The percentages of H-2Kb/OVA-I tetramer+CD62L cells in CD49bDTRvenusCD8+ splenocytes, gated as in (C), are shown. (D) Whole splenocytes were further restimulated with (+) or without (-) OVA-I peptide and subjected to intracellular cytokine staining. (F) After in vitro restimulation with OVA-I peptide, IFN-γ production was analyzed by intracellular staining. The percentages of IFN-γ–producing CD62L cells in CD49bDTRvenusCD8+ splenocytes, gated as in (D), are shown. Data in (C) and (D) are representative of six PBS-treated and three DT-treated mice, respectively. Data were combined from three (A, E, F) or two (B) independent experiments. In (A), (B), (E), and (F), each symbol represents an individual mouse, and the horizontal lines represent the means. *p < 0.05, **p < 0.01, ***p < 0.001.

FIGURE 7.

Immune responses against L.m.-OVA infection in XCR1-DTRvenus mice. XCR1-DTRvenus mice received PBS or DT at days −1 and 0 of L.m.-OVA infection. (A) Mice were infected i.v. with the indicated numbers of L.m.-OVA. Three days postinfection, bacterial load in the spleen or liver was evaluated. (B) Depletion of liver DTRvenus-expressing cells. XCR1-DTRvenus mice were injected with PBS or DT at days −1 and 0. One day after the last injection, liver leukocytes were prepared and stained with the indicated Abs. MHC-II+CD11c+ cells were gated, and their expression profiles of CD103 and DTRvenus are shown (left panels). Numbers represent the percentages of the cells in each quadrant. Percentages of DTRvenus+ cells in MHC-II+CD11c+ cells are also shown (right panels). PBS- or DT-treated XCR1-DTRvenus mice were infected i.v. with 1 × 103 (C, D) or the indicated numbers (E, F) of L.m.-OVA and analyzed for CD8+ T cell responses 7 d later. (C and E) H-2Kb/OVA-I tetramer+ cells in CD49bDTRvenusCD8+ splenocytes were monitored by FACS. (E) The percentages of H-2Kb/OVA-I tetramer+CD62L cells in CD49bDTRvenusCD8+ splenocytes, gated as in (C), are shown. (D) Whole splenocytes were further restimulated with (+) or without (-) OVA-I peptide and subjected to intracellular cytokine staining. (F) After in vitro restimulation with OVA-I peptide, IFN-γ production was analyzed by intracellular staining. The percentages of IFN-γ–producing CD62L cells in CD49bDTRvenusCD8+ splenocytes, gated as in (D), are shown. Data in (C) and (D) are representative of six PBS-treated and three DT-treated mice, respectively. Data were combined from three (A, E, F) or two (B) independent experiments. In (A), (B), (E), and (F), each symbol represents an individual mouse, and the horizontal lines represent the means. *p < 0.05, **p < 0.01, ***p < 0.001.

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To monitor cytokine production after in vitro stimulation of DCs, CD11c+ splenic DCs were enriched by MACS sorting with anti-CD11c beads (Miltenyi Biotec). Then the cells were stimulated with 50 μg/ml poly(I:C) or 1 μM CpG DNA (ODN1668). After 30 min, GolgiStop (BD Bioscience) was added, and the cells were cultured for another 6 h. The cells were stained with mAbs to CD11c, CD8α, and PDCA-1, fixed with fixation/permeabilization solution (Cytofix/Cytoperm Kit), and stained with anti-IL12p40 mAb, according to the manufacturer’s instructions. PDCA-1CD11c+ cells were analyzed for IL-12p40 production.

To evaluate OVA-specific CD8+ T cell responses, splenocytes were first analyzed by H-2Kb/OVA-I peptide tetramer (MBL) staining. For intracellular IFN-γ staining, splenocytes were restimulated with 1 μg/ml OVA-I peptide for 6 h in the presence of GolgiStop or brefeldin A (10 μg/ml; Sigma-Aldrich), as described previously (38). Subsequently, the cells were stained with mAbs to CD8α, CD62L, and CD49b, fixed with fixation/permeabilization solution (Cytofix/Cytoperm Kit), and stained with anti-IFN-γ mAb. CD49bDTRvenusCD8+ cells were analyzed for IFN-γ production. For CD4+ T cell responses, splenocytes were restimulated with 100 μg/ml OVA protein for 12 h. Brefeldin A (10 μg/ml) was added to the culture for the last 6 h. Subsequently, the cells were stained with mAbs to CD4, CD62L, and CD49b, fixed with fixation/permeabilization solution, and stained with anti–IFN-γ mAb. CD49bDTRvenusCD4+ cells were analyzed for IFN-γ production. The cells were analyzed by FACSCalibur, FACSCanto II, FACSAria II, or FACSVerse (BD Biosciences). Data were analyzed with FlowJo software (TreeStar).

Spleen and inguinal LNs were embedded in OCT compound (Sakura Finetek) or FSC22 frozen section compound (Leica Microsystems), and 5-μm cryosections were prepared. Spleen sections were stained with rabbit anti-GFP polyclonal Ab (MBL), allophycocyanin-conjugated anti-F4/80 (BM8; eBioscience), and biotinylated anti-CD169 (MOMA-1; Abcam) and then developed with Alexa Fluor 480–conjugated anti-rabbit IgG (H+L) (Invitrogen) and streptavidin–Pacific Blue. LN sections were stained with rabbit anti-GFP polyclonal Ab (MBL), Pacific Blue–conjugated anti-B220, and biotinylated anti-CD169 and then developed with Alexa Fluor 480–00conjugated anti-rabbit IgG (H+L) and streptavidin-APC. Stained sections were analyzed by FV10i (Olympus).

XCR1-DTRvenus mice were injected i.p. with DT at days −2 and −1 of poly(I:C) injection. At day 0, PBS or 150 μg poly(I:C) was injected i.v. into the mice, and serum samples were collected at the indicated times. Serum cytokine levels were analyzed by Bio-Plex Suspension Array System (Bio-Rad) or ELISA. ELISA kits for IFN-α and IFN-β were from PBL InterferonSource.

Mice were injected with PBS or DT on days −1 and 0 of immunization. At day 0, each hind footpad was immunized with soluble OVA (50 μg) with 10 μg poly(I:C). OVA-specific CD4+ and CD8+ T cell responses were monitored 7 d later. For cross-presentation of cell-associated Ags, OVA-expressing MHC class I–deficient (mOVA-Tg:β2m−/−) thymocytes and splenocytes were subjected to gamma radiation (15 Gy) to induce apoptosis. Irradiated dying cells (1.3–2 × 107 cells) were immediately injected i.v. into the mice with 50 μg poly(I:C) at day 0. OVA-specific CD8+ T cell responses were monitored 7 d later.

L. monocytogenes expressing OVA (L.m.-OVA) was originally established by Dudani et al. (39) and was purchased from DMX. DT or PBS was injected i.p. at days −1 and 0 of infection. L.m.-OVA was grown in brain–heart infusion broth to an OD600 of 0.1–0.2. An OD600 of 0.1 is equivalent to 2 × 108 bacteria/ml. To measure bacterial loads, mice were infected i.v. with the indicated numbers of L.m.-OVA and sacrificed 3 d postinfection to collect spleens and livers, which were lysed in PBS containing 0.05% Triton X-100. Bacterial titers were determined by plating serial dilutions on brain–heart infusion agar plates containing 1 μg/ml erythromycin (Sigma-Aldrich). To assess L.m.-OVA–specific CD8+ T cell responses, mice were analyzed 7 d after i.v. infection. To isolate liver leukocytes, the liver was perfused with PBS via the portal vein and digested with 400 U/ml Collagenase D for 35 min at 37°C. EDTA (5 mM) was added for the last 5 min. Low-density cells were enriched by centrifugation with 40 and 80% Percoll density gradient.

The data were analyzed using the two-tailed unpaired Student t test, with or without a Welch correction, depending on whether the data had possible unequal or equal variances, respectively. For differences in CFU, data were analyzed by the Mann–Whitney U test. The p values <0.05 were considered statistically significant. All statistical analyses were performed using GraphPad Prism software (GraphPad).

We first replaced the coding region of Xcr1 with a gene encoding venus (Fig. 1A). The replacement was confirmed by Southern blot analysis (Fig. 1B). Xcr1+/venus mice were born at an expected Mendelian ratio and did not show any obvious developmental abnormalities. Throughout this study, Xcr1+/venus mice were used as XCR1-venus mice.

Next, we investigated venus expression in the spleen of XCR1-venus mice. Less than 1% of splenocytes showed venus expression (Fig. 1C). More than 85% of venus+ cells expressed CD8α and CD11c, suggesting that venus was selectively expressed in CD8α+ DCs (Fig. 1C). Meanwhile, venus expression was not observed in CD11c cells, including T, B, and NK cells, granulocytes, and monocytes (Fig. 1D). The expression pattern of venus in CD8α+ DCs (Fig. 1C) suggested that CD8α+ DCs consisted of two subsets: XCR1+ and XCR1 DCs. We then asked whether venus expression correlates with the cytokine-producing abilities of CD8α+ DCs. Splenic DCs were stimulated with a TLR3 agonist [poly(I:C)] or a TLR9 agonist (CpG DNA), and IL-12p40 production was monitored by FACS (Fig. 1E). When stimulated with poly(I:C), IL-12p40 production was strongly induced in CD8α+ DCs but not in CD8α DCs. Among CD8α+ DCs, IL-12p40 production was observed primarily in venus-expressing cells. When stimulated with a TLR9 agonist, both CD8α+ and CD8α DCs produced IL-12p40. Among CD8α+ DCs, venus+ cells mainly responded to CpG DNA. Thus, venus is dominantly expressed in mature CD8α+ DCs with a high capacity to produce cytokines.

We further analyzed venus expression in various lymphoid tissues, such as spleen, thymus, SDLNs, and mLNs (Fig. 2A). In these organs, similar ratios of DC subsets were observed between WT and XCR1-venus mice (Fig. 2A), and venus was expressed only in MHC-II+CD11c+ cells (data not shown). In the spleen, venus was expressed in 70–90% of CD8α+CD11b DCs. Venus was not expressed in CD8αCD11b+ DCs, although a small fraction of CD8αCD11b DCs was positive for venus. In the thymus, venus expression was observed in 70% of CD8α+CD11b DCs and 50% of CD8αCD11b DCs. Although only resident DCs are present in the spleen, SDLNs carry both resident MHC-IIintCD11c+ and migratory MHC-IIhighCD11c+ DCs. Migratory DCs were characterized by the expression profile of CD103 and CD11b. In the SDLNs, venus+ cells were found mainly in CD8α+CD11b and CD103+CD11b cells in the resident and migratory DCs, respectively. In the mLNs, migratory CD103+ DCs contained CD11b+ DCs, as well as CD11b DCs. As in SDLNs, venus+ cells were found primarily in resident CD8α+CD11b DCs and migratory CD103+CD11b DCs, whereas 30% of CD8αCD11b cells were positive for venus. We further assessed venus expression in the peripheral tissues, such as the skin and small intestine lamina propria (Fig. 2B, 2C). In the skin, epidermal Langerhans cells (LCs) did not express venus but dermal CD103+CD11b DCs did. In the small intestinal lamina propria, venus was expressed only in CD103+CD11b DCs, but not in CD103+CD11b+ or CD103CD11b+ DCs, as in the case of migratory DCs in the mLNs (Fig. 2A). Thus, venus is abundantly expressed in resident CD8α+CD11b DCs and migratory CD103+CD11b DCs in lymphoid organs and peripheral CD103+CD11b DCs. In the thymus or LNs, venus was also expressed in a subset of CD8αCD11b or CD103CD11b DCs.

Distinct populations of DCs are generated transiently and accumulate in the inflamed tissues and lymphoid organs in response to microbial infection or inflammatory stimuli. We analyzed whether venus was expressed in TNF-α/inducible NO synthase–producing DCs (Tip-DCs) detected in the spleen after systemic infection of L. monocytogenes and in monocyte-derived DCs (Mo-DCs) observed in LNs after LPS injection or Gram-negative bacterial infection (35, 40). To analyze Tip-DCs, mice were infected with L.m.-OVA. Two days postinfection, splenocytes were analyzed for venus expression. L.m.-OVA infection induced the generation and accumulation of Tip-DCs (Ly6C+CD11b+MHC-II+CD11c+) in the spleen (Fig. 3A). Ly6C DCs contained venus-expressing cells, >90% of which were CD8α+CD11b DCs (data not shown). However, the majority of Tip-DCs was negative for venus. To analyze Mo-DCs, mice were injected i.v. with LPS. After 24 h, Mo-DCs, which can be defined as CD206+CD11c+ cells, were increased in the LNs. Although CD206CD11c+ cells contained venus+ cells, which should correspond to resident CD8α+ DCs and migratory CD103+ DCs, Mo-DCs were negative for venus (Fig. 3B). Thus, the majority of inflammatory DCs, such as Tip-DCs and Mo-DCs, did not express XCR1.

We further confirmed venus expression in in vitro Mo-DCs or BM-derived DCs (Fig. 3C, 3D). In vitro Mo-DCs were generated by culturing purified BM monocytes with GM-CSF and IL-4. Venus expression was not observed in in vitro Mo-DCs (Fig. 3C). In vitro BM-derived DCs were generated in the presence of GM-CSF or Flt3L. GM-CSF–induced BM DCs failed to show venus expression (Fig. 3D). Flt3L-induced BM DCs include B220+CD11c+ and B220CD11c+ cells. The former population corresponds to pDCs. B220CD11c+ cells can be further divided into CD24highCD11blow DCs (CD24high DCs) and CD24lowCD11bhigh DCs (CD11bhigh DCs), and CD24high DCs are thought to be the equivalents of splenic CD8α+ DCs. Venus expression was detected in CD24high DCs but not in CD11bhigh DCs or pDCs (Fig. 3D).

To achieve conditional ablation of XCR1+ DCs in vivo, we next replaced the coding region of Xcr1 with the DTRvenus cassette (Fig. 4A). The replacement was confirmed by Southern blot analysis (Fig. 4B). Xcr1+/DTRvenus mice were born at an expected Mendelian ratio and did not show any obvious developmental abnormalities. Throughout this study, Xcr1+/DTRvenus mice were used as XCR1-DTRvenus mice.

In XCR1-DTRvenus mice, subsets of CD11c+ DCs and other cellularities were similar to those in WT mice, suggesting that the expression of DTRvenus itself was not toxic to cells or mice. Although the fluorescence intensity of venus in XCR1-DTRvenus mice was lower than that in XCR1-venus mice, the venus-expressing cell population was similar between these two mutant mice (data not shown).

We next assessed the ablation of DTRvenus-expressing cells after DT injection. XCR1-DTRvenus mice were injected i.p. with DT (25 ng/g body weight) and monitored for the presence of DTRvenus-expressing cells in the spleen (Fig. 4C). DTRvenus-expressing CD8α+ DCs were rapidly depleted at days 1 and 2 after DT injection. Then, DTRvenus-expressing cells began to increase around day 4 and had nearly recovered at day 8. Both resident and migratory DTRvenus-expressing DCs were depleted with similar kinetics in SDLNs and mLNs, as shown by the frequency and absolute cell numbers (Fig. 4D). Other cells, such as B, T, and NK cells, granulocytes, monocytes, and DTRvenus DC subsets, including pDCs and CD11b+ DCs, were unaffected (Fig. 4C, Table I, data not shown).

Table I.
Analysis of XCR1 cell population in the spleen of XCR1-DTRvenus mice injected with PBS or DT
% of Whole Splenocytes
Cell No.
CellsPBSDTPBSDT
T cells 32.5 ± 2.79 35.6 ± 2.18 3.22 ± 0.39 × 107 3.26 ± 0.36 × 107 
B cells 45.0 ± 4.31 49.1 ± 1.89 4.44 ± 0.49 × 107 4.49 ± 0.39 × 107 
NK cells 2.93 ± 0.31 2.43 ± 0.22 2.95 ± 0.54 × 106 2.22 ± 0.27 × 106 
Granulocytes 1.44 ± 0.15 1.87 ± 0.31 1.54 ± 0.50 × 106 1.65 ± 0.12 × 106 
Monocytes 1.20 ± 0.42 0.90 ± 0.06 0.95 ± 0.49 × 106 1.10 ± 0.24 × 106 
CD8αCD11b+ DCs 0.53 ± 0.10 0.60 ± 0.09 5.59 ± 0.83 × 105 5.57 ± 0.29 × 105 
pDCs 0.57 ± 0.03 0.90 ± 0.22 6.81 ± 0.26 × 105 9.36 ± 2.24 × 105 
% of Whole Splenocytes
Cell No.
CellsPBSDTPBSDT
T cells 32.5 ± 2.79 35.6 ± 2.18 3.22 ± 0.39 × 107 3.26 ± 0.36 × 107 
B cells 45.0 ± 4.31 49.1 ± 1.89 4.44 ± 0.49 × 107 4.49 ± 0.39 × 107 
NK cells 2.93 ± 0.31 2.43 ± 0.22 2.95 ± 0.54 × 106 2.22 ± 0.27 × 106 
Granulocytes 1.44 ± 0.15 1.87 ± 0.31 1.54 ± 0.50 × 106 1.65 ± 0.12 × 106 
Monocytes 1.20 ± 0.42 0.90 ± 0.06 0.95 ± 0.49 × 106 1.10 ± 0.24 × 106 
CD8αCD11b+ DCs 0.53 ± 0.10 0.60 ± 0.09 5.59 ± 0.83 × 105 5.57 ± 0.29 × 105 
pDCs 0.57 ± 0.03 0.90 ± 0.22 6.81 ± 0.26 × 105 9.36 ± 2.24 × 105 

XCR1-DTRvenus mice were injected with PBS or DT. One day after injection, splenocytes were stained with CD3ε, CD11b, CD19, CD115, DX5, and Ly6G. T cells (CD3ε+CD19), B cells (CD3εCD19+), NK cells (CD3εDX5+), granulocytes (Ly6G+CD11b+), monocytes (Ly6GCD115+CD11b+), CD8αCD11b+ DCs (CD8αCD11b+MHC-II+CD11c+), and pDCs (PDCA-1+CD11clow) were analyzed by FACS. Data represent the mean ± SEM (n = 3 or 4). No values were significantly different between PBS- and DT-injected mice.

In the murine spleen, MΦ subsets include CD169+ marginal zone metallophilic MΦs (MZMMΦs) and F4/80+ red pulp MΦs. CD169+ MZMMΦs are known to contribute to cross-presentation of blood-borne Ags by splenic DCs (41). In the LNs, CD169+ cells are found in the sinus and have the ability to cross-present Ags from dead cells (42). CD169+ cells in the spleen and LNs were ablated after DT injection to CD11c-DTR mice (43). We then asked whether MΦs in the spleen and sinus CD169+ cells in the LNs of XCR1-DTRvenus mice were affected by DT injection. DTRvenus was detected in the T cell area and red pulp of the spleen and T cell area, sinus, and medulla of the inguinal LNs (Fig. 4E). After DT injection, these DTRvenus+ cells were depleted. In contrast, the majority of CD169+ MZMMΦs and F4/80+ red pulp MΦs in the spleen and CD169+ cells in the inguinal LNs was present in XCR1-DTRvenus mice after DT injection. According to Asano et al. (42), the CD169+CD11c+ cell fraction in LNs shows a high capacity to ingest dead cells and is responsible for cross-presenting Ags to T cells. Therefore, we further analyzed venus expression in subsets of CD169+ cells in LNs (Fig. 4F). CD169+CD11c and CD169+CD11cdull cells did not express DTRvenus, but about a half of CD169+CD11chigh cells expressed DTRvenus. The DTRvenus+ cells included MHC-IIhigh and MHC-IIint cells (data not shown), which should correspond to migratory and resident cells, respectively. All of the DTRvenus+ cells were ablated in DT-treated XCR1-DTRvenus mice (Fig. 4F).

We next examined dsRNA-induced immune responses in XCR1-DTRvenus mice. In mice, there are two systems to detect dsRNAs, including poly(I:C) and viral RNAs. One is endosomal recognition by TLR3, and the other is cytoplasmic recognition by RIG-I–like receptors, including RIG-I and MDA5 (4447). TLR3 and RIG-I–like receptors activate the distinct signaling pathways through the adaptors TRIF and IPS-1, respectively (48). dsRNA-mediated induction of proinflammatory cytokines, such as IL-12p40, TNF-α, and IL-6, and type I IFNs is dependent on TLR3–TRIF and RLR–IPS-1 pathways, respectively (46, 49) (data not shown). TLR3 is expressed only in CD8α+ DCs among DC subsets (50), and venus+ cells were potent IL-12p40–producing cells in response to poly(I:C) stimulation (Fig. 1D). Therefore, we measured serum cytokine levels in XCR1-DTRvenus mice after poly(I:C) injection. The serum cytokine levels were similar in DT-treated XCR1-DTRvenus mice and PBS-treated XCR1-DTRvenus mice (Fig. 5). Thus, XCR1-expressing cells are dispensable for in vivo cytokine induction by dsRNA.

FIGURE 5.

dsRNA-induced cytokine production in XCR1-DTRvenus mice. XCR1-DTRvenus mice were treated with PBS or DT on days −2 and −1 and then poly(I:C) (150 μg/mouse) was injected i.p. Sera samples were collected at the indicated times, and the concentration of IFN-α, IFN-β, IL-12p40, IL-12p70, TNF-α, and IL-6 was measured. Data are mean ± SEM (n = 6).

FIGURE 5.

dsRNA-induced cytokine production in XCR1-DTRvenus mice. XCR1-DTRvenus mice were treated with PBS or DT on days −2 and −1 and then poly(I:C) (150 μg/mouse) was injected i.p. Sera samples were collected at the indicated times, and the concentration of IFN-α, IFN-β, IL-12p40, IL-12p70, TNF-α, and IL-6 was measured. Data are mean ± SEM (n = 6).

Close modal

We next checked dsRNA-induced cross-presentation activity in XCR1-DTRvenus mice. DT-treated WT mice and PBS- or DT-treated XCR1-DTRvenus mice were immunized with soluble OVA protein in the presence or absence of poly(I:C). Seven days postimmunization, CD8+ T cell responses in the spleen were monitored by H-2Kb/OVA-I peptide tetramer staining and IFN-γ production (Fig. 6A–C). In both DT-treated WT and PBS-treated XCR1-DTRvenus mice, H-2Kb/OVA-I peptide tetramer+ and IFN-γ–producing CD8+ T cells were clearly observed in a CD62L activated fraction after immunization with OVA and poly(I:C) but not with OVA only. However, these responses were severely impaired in DT-treated XCR1-DTRvenus mice. On the contrary, after immunization with OVA and poly(I:C), IFN-γ production from CD4+ T cells after restimulation with OVA protein in vitro was enhanced in DT-treated XCR1-DTRvenus mice compared with PBS-treated XCR1-DTRvenus mice (Fig. 6D). Thus, XCR1-expressing cells were essential for dsRNA-induced CD8+, but not CD4+, T cell responses against soluble Ags.

FIGURE 6.

dsRNA-induced immune responses against soluble (AD) or cell-associated (EG) Ags in XCR1-DTRvenus mice. WT or XCR1-DTRvenus mice were injected with PBS or DT on days −1 and 0, immunized s.c. in the footpads with soluble OVA in the presence or absence of 10 μg of poly(I:C), and then analyzed for CD8+ (A–C) or CD4+ (D) T cell responses 7 d later. (A) Numbers indicate the percentages of H-2Kb/OVA-I tetramer+CD62L cells in CD49bvenusCD8+ splenocytes from DT-treated WT, PBS-treated XCR1-DTRvenus, or DT-treated XCR1-DTRvenus mice immunized with soluble OVA plus poly(I:C). (B) Whole splenocytes were further restimulated with (+) or without (-) OVA-I peptide and subjected to intracellular cytokine staining. (C) After in vitro restimulation with OVA-I peptide, IFN-γ production was analyzed by intracellular staining. Percentages of IFN-γ–producing CD62L cells in CD49b-DTRvenus-CD8+ splenocytes gated as in (B) are shown. (D) Whole splenocytes were restimulated with OVA protein and subjected to intracellular cytokine staining. Percentages of IFN-γ–producing CD62L cells in CD49bDTRvenusCD4+ splenocytes are shown. (E–G) XCR1-DTRvenus mice were injected with PBS or DT on days −1 and 0 and immunized i.v. with cell-associated OVA (Cell-OVA) derived from dying mOVA-Tg:β2m−/− cells in the presence or absence of 50 μg of poly(I:C). Seven days later, CD8+ T cell responses were analyzed. (E) Numbers indicate the percentages of H-2Kb/OVA-I tetramer+CD62L cells in CD49bvenusCD8+ splenocytes from PBS- or DT-treated XCR1-DTRvenus mice immunized with cell-OVA plus poly(I:C). (F) Whole splenocytes were further restimulated with (+) or without (-) OVA-I peptide and subjected to intracellular cytokine staining. (G) After in vitro restimulation with OVA-I peptide, IFN-γ production was analyzed by intracellular staining. Percentages of IFN-γ–producing CD62L cells in CD49bvenusCD8+ splenocytes, gated as in (F), are shown. In (C), (D), and (G), each symbol represents an individual mouse, and the horizontal lines represent the means. Data in (A), (B), (E), and (F) are representative of at least three mice. In (C), (D), and (G), data were combined from three independent experiments. *p < 0.05, **p < 0.01. Imm., Immunization.

FIGURE 6.

dsRNA-induced immune responses against soluble (AD) or cell-associated (EG) Ags in XCR1-DTRvenus mice. WT or XCR1-DTRvenus mice were injected with PBS or DT on days −1 and 0, immunized s.c. in the footpads with soluble OVA in the presence or absence of 10 μg of poly(I:C), and then analyzed for CD8+ (A–C) or CD4+ (D) T cell responses 7 d later. (A) Numbers indicate the percentages of H-2Kb/OVA-I tetramer+CD62L cells in CD49bvenusCD8+ splenocytes from DT-treated WT, PBS-treated XCR1-DTRvenus, or DT-treated XCR1-DTRvenus mice immunized with soluble OVA plus poly(I:C). (B) Whole splenocytes were further restimulated with (+) or without (-) OVA-I peptide and subjected to intracellular cytokine staining. (C) After in vitro restimulation with OVA-I peptide, IFN-γ production was analyzed by intracellular staining. Percentages of IFN-γ–producing CD62L cells in CD49b-DTRvenus-CD8+ splenocytes gated as in (B) are shown. (D) Whole splenocytes were restimulated with OVA protein and subjected to intracellular cytokine staining. Percentages of IFN-γ–producing CD62L cells in CD49bDTRvenusCD4+ splenocytes are shown. (E–G) XCR1-DTRvenus mice were injected with PBS or DT on days −1 and 0 and immunized i.v. with cell-associated OVA (Cell-OVA) derived from dying mOVA-Tg:β2m−/− cells in the presence or absence of 50 μg of poly(I:C). Seven days later, CD8+ T cell responses were analyzed. (E) Numbers indicate the percentages of H-2Kb/OVA-I tetramer+CD62L cells in CD49bvenusCD8+ splenocytes from PBS- or DT-treated XCR1-DTRvenus mice immunized with cell-OVA plus poly(I:C). (F) Whole splenocytes were further restimulated with (+) or without (-) OVA-I peptide and subjected to intracellular cytokine staining. (G) After in vitro restimulation with OVA-I peptide, IFN-γ production was analyzed by intracellular staining. Percentages of IFN-γ–producing CD62L cells in CD49bvenusCD8+ splenocytes, gated as in (F), are shown. In (C), (D), and (G), each symbol represents an individual mouse, and the horizontal lines represent the means. Data in (A), (B), (E), and (F) are representative of at least three mice. In (C), (D), and (G), data were combined from three independent experiments. *p < 0.05, **p < 0.01. Imm., Immunization.

Close modal

We further analyzed dsRNA-induced cross-presentation of cell-associated Ags. PBS- or DT-treated XCR1-DTRvenus mice were immunized with dying cells from mOVA-Tg:β2m−/− mice in the presence or absence of poly(I:C). Seven days postimmunization, CD8+ T cell responses in the spleen were monitored by H-2Kb/OVA-I peptide tetramer staining and IFN-γ production. In PBS-treated XCR1-DTRvenus mice, H-2Kb/OVA-I tetramer+ and IFN-γ–producing CD62LCD8+ T cells were increased when immunized with poly(I:C). However, the responses were severely diminished in DT-treated XCR1-DTRvenus mice (Fig. 6E–G). Thus, XCR1-expressing cells were crucial for dsRNA-induced CD8+ T cell responses against dead/dying cell–associated Ags.

Analyses of various mutant mice revealed that splenic CD8α+ DCs are crucial for L. monocytogenes to enter into and replicate in the spleen (23, 51). We then examined the roles of XCR1-expressing cells in L. monocytogenes infection in XCR1-DTRvenus mice. DT- or PBS-treated XCR1-DTRvenus mice were infected i.v. with the indicated numbers of L.m.-OVA, and bacterial titers were measured at day 3 postinfection. Bacterial burden in the spleen was severely diminished in DT-treated mice compared with PBS-treated mice (Fig. 7A). Meanwhile, bacterial burden in the liver was comparable between PBS- and DT-treated XCR1-DTRvenus mice. We then evaluated DTRvenus expression and depletion of DTRvenus+ cells in the liver after DT injection to XCR1-DTRvenus mice (Fig. 7B). DTRvenus was primarily expressed in CD103+MHC-II+CD11c+ cells, and DTRvenus+ cells were efficiently ablated after DT injection, excluding the possibility that comparable bacterial loads in the liver of DT-injected XCR1-DTRvenus mice was due to incomplete deletion of hepatic DTRvenus+ cells, which could be target cells of L. monocytogenes infection.

We further evaluated OVA-specific T cell responses (Fig. 7C–F). Expansion of Ag-specific activated T cells was clearly impaired in DT-treated mice compared with PBS-treated mice (Fig. 7C, 7E). IFN-γ production from OVA-I peptide–stimulated CD8+ T cells was also severely impaired in DT-treated mice (Fig. 7D, 7F). Thus, XCR1-expressing cells were essential for the establishment of L. monocytogenes infection in the spleen and for the development of L. monocytogenes–specific CD8+ T cell responses.

We generated mutant mice by knocking the venus or DTRvenus gene into the Xcr1 locus. In these mice, venus was selectively expressed in splenic CD8α+ DCs and related subsets in lymphoid organs, such as thymus and LNs, and in the peripheral tissues, although it was also expressed in a significant fraction of CD8αCD11b or CD103CD11b DCs in the thymus or LNs. The percentages of venus-expressing cells among CD8αCD11b DCs were highest in the thymus. This might be due to the lower expression of CD8α in the thymic DCs and the difficulty in discriminating CD8α+CD11b DCs from CD8αCD11b DCs. Venus expression was also detected in a CD24high DC subset, which can be generated in vitro with Flt3L from BM cells and is equivalent to splenic CD8α+ DCs. Meanwhile, the other DC subsets among Flt3L-induced BM DCs, in vitro and in vivo Mo-DCs, as well as Tip-DCs, failed to show venus expression. Furthermore, venus was not expressed in CD11c cells, including T, B, and NK cells, granulocytes, and monocytes. These expression patterns are consistent with the previous findings based on the expression pattern of LacZ knocked into the Xcr1 locus or analysis with anti-XCR1 Ab (26, 27, 30, 31). Twenty to thirty percent of splenic CD8α+ DCs did not express venus. We found that venus+CD8α+ DCs produced much greater amounts of cytokines than did venusCD8α+ DCs in response to TLR agonists, which indicates that XCR1+ DCs are more mature than XCR1 DCs and that functional CD8α+ DCs can be tracked in our mutant mice. It is also notable that the XCR1 expression level was unchanged after injection of LPS or poly(I:C) (Fig. 3B, data not shown).

The XCR1+ DCs were efficiently ablated in XCR1-DTRvenus mice. Conditional ablation of CD8α+ DCs was also achieved in several DTR knock-in mice (52, 53). Clec9A, also known as DC NK lectin group receptor-1, is expressed dominantly in CD8α+ DCs (15). Clec9A-DTR mice were generated by transfecting ES cells with recombineered bacterial artificial chromosome clones carrying insertion of human DTR cDNA with its polyA site into the Clec9a locus (52). Upon injection of DT in Clec9A-DTR mice, splenic CD8α+ DCs can be efficiently depleted, but pDCs are also partially depleted. This is consistent with the low expression of Clec9A in pDCs. CD205, also called DEC-205, is a C-type lectin–like molecule that is expressed abundantly in CD8α+ DCs (9, 54). CD205-DTR enhanced GFP (EGFP) mice were generated by inserting an internal ribosome entry site followed by cDNA encoding human DTR fused to EGFP into the Cd205 locus (53). Although DT injection in CD205-DTREGFP mice depleted splenic CD8α+ DCs efficiently, it also depleted CD205-expressing cells in CD8α DCs. Furthermore, the irradiated WT mice reconstituted with BM of CD205-DTREGFP mice had to be analyzed, because CD205-DTREGFP mice die within 10 d after DT injection, probably as a result of DTR expression on certain radioresistant cells (53). Meanwhile, the XCR1-DTRvenus mice were viable after injection of up to 100 ng/g body weight of DT (data not shown). Langerin is expressed in splenic CD8α+ DCs and subsets of resident/migratory LNs and dermal DCs, as well as LCs in the epidermis (8, 5558). Langerin+ LN and dermal DCs contain both CD103+ and CD103 DCs (59). A cDNA encoding a fusion protein consisting of DTR and EGFP was knocked into the Langerin locus to generate Langerin-DTREGFP mice (8, 55). Langerin+ DCs, including both XCR1+ DCs (dermal and LN CD103+ DCs) and XCR1 DCs (dermal and LN CD103 DCs and LCs), can be depleted upon DT injection in Langerin-DTREGFP mice (8, 55, 56). Thus, each mouse shows a distinct pattern of cell ablation. In terms of cell tracking, fluorescence protein expression can be monitored in XCR1-DTRvenus and Langerin-DTREGFP mice but not in Clec9A-DTR or CD205-DTREGFP mice. Thus, XCR1-DTRvenus mice are useful mutant mice for analyzing splenic CD8α+ DCs and their relatives.

Using XCR1-DTRvenus mice, we clarified the roles of XCR1-expressing cells in dsRNA-induced immune responses. Induction of type I IFNs by dsRNA was retained in DT-treated XCR1-DTRvenus mice. This is consistent with the finding that splenic CD8α+ DCs show faint expression of cytosolic dsRNA sensors that can lead to type I IFN production (50). Furthermore, induction of proinflammatory cytokines, including IL-12p40, which depends on a TLR3-TRIF axis, was also preserved in DT-treated XCR1-DTRvenus mice, although DTRvenus+CD8α+ DCs are mainly responsible for dsRNA-induced IL-12p40 production among DC subsets. It can be assumed that TLR3-expressing DTRvenus cells compensate for dsRNA-induced production of proinflammatory cytokines in vivo. Thus, XCR1-expressing cells were dispensable for dsRNA-induced cytokine induction in vivo.

Type I IFN is required for optimal induction of cross-priming (24, 37, 60). In DT-treated XCR1-DTRvenus mice, dsRNA-induced CD8+ T cell responses were impaired under the conditions in which the cytokines are normally induced. This suggests that dsRNA-stimulated XCR1 DCs fail to induce CD8+ T cell responses, even in the presence of cytokines, including type I IFNs. Notably, dsRNA-induced CD4+ T cell responses were enhanced in DT-treated XCR1-DTRvenus mice. CD4+ T cell responses should be provoked by other APCs that can be activated by cytosolic sensors or type I IFNs. CD8α DCs should be the candidate APCs, according to previous reports on the division of labor between CD8α+ and CD8α DCs (61, 62). The absence of XCR1-expressing DCs can make Ags more accessible to CD8α DCs and might have led to augmented CD4+ T cell responses (Fig. 6D).

Among CD169+ cells in LNs, CD11c+ cells were reported to be involved in incorporating dead tumor cells and cross-presenting their Ags (42). In XCR1-DTRvenus mice, a fraction of CD169+CD11chigh cells in LNs was depleted by DT injection. Therefore, it is possible that this cell population (i.e., XCR1-expressing CD169+CD11chigh cells) also contributes to dsRNA-induced cross-presentation of soluble or dying cell–associated Ags.

In L. monocytogenes infection, bacterial replication was severely decreased in the spleen, but not in the liver, of DT-treated XCR1-DTRvenus mice. DTRvenus-expressing cells in the liver were efficiently depleted in DT-treated XCR1-DTRvenus mice, indicating that comparable bacterial loads in the liver cannot be ascribed to inefficient depletion of DTRvenus-expressing cells (Fig. 7B). It is possible that, in the liver of DT-treated XCR1-DTRvenus mice, L. monocytogenes infects and expands in the cells that do not express XCR1, although further studies are required to confirm this possibility. However, consistent with our findings, bacterial loads in DT-treated CD11c-DTR mice were decreased in the spleen but remained stable in the liver (51). Crozat et al. (27) reported that XCR1-deficient mice showed increased bacterial loads in both the spleen and the liver. Target cells for L. monocytogenes infection are present in Xcr1-deficient mice, and diminished CD8+ T cell responses likely led to increased bacterial loads. We can assume that Xcr1-expressing cells are the main targets of L. monocytogenes infection in the spleen and that diminished CD8+ T cell responses result from decreased infection. However, we cannot exclude the possibility that defective cross-presentation contributes to impairment of anti–L. monocytogenes CD8+ T cell responses in DT-treated XCR1-DTRvenus mice, because infection in the liver was not diminished.

In this study, we demonstrated critical roles for XCR1-expressing DCs by analyzing XCR1-venus and XCR1-DTRvenus mice. These mice are suitable for ablating and tracking splenic CD8α+ DCs and their relatives. Because the expression pattern of XCR1 and its ligand, XCL1, is quite similar between humans and mice (2729, 32, 33), information on the function and behavior of XCR1-expressing DCs in mice could be applicable to humans. The mutant mice generated in this study should be useful for revealing the expected or unexpected roles of XCR1-expressing DCs in various immune responses or disease models.

We thank A. Kato, I. Ogahara, Y. Tanaka, and E. Haga for technical assistance and S. Haraguchi and Y. Matsuhisa for secretarial assistance. We also thank Drs. E. Mekata and A. Miyawaki for providing human HB-EGF cDNA and venus/pCS2 vector, respectively, as well as Drs. C.L. Stewart and M. Hikida for providing an ES cell line (Bruce4).

This work was supported by the Kishimoto Foundation, a Grant-in-Aid for Scientific Research (B, C), a Grant-in-Aid for Challenging Exploratory Research, a Grant-in-Aid for Scientific Research on Priority Areas, a Grant-in-Aid for Scientific Research on Innovative Areas, the Uehara Memorial Foundation, and a Grant-in-Aid for Young Scientists. C.Y., M.S., and I.S. were supported by a RIKEN Junior Research Associate grant.

Abbreviations used in this article:

     
  • DC

    dendritic cell

  •  
  • DT

    diphtheria toxin

  •  
  • DTR

    diphtheria toxin receptor

  •  
  • EGFP

    enhanced GFP

  •  
  • ES

    embryonic stem

  •  
  • Flt3L

    Flt3 ligand

  •  
  • LC

    Langerhans cell

  •  
  • L.m.-OVA

    Listeria monocytogenes expressing OVA

  •  
  • LN

    lymph node

  •  
  • macrophage

  •  
  • MHC-II

    MHC class II

  •  
  • mLN

    mesenteric lymph node

  •  
  • Mo-DC

    monocyte-derived dendritic cell

  •  
  • mOVA-Tg

    C57BL/6-Tg(CAG-OVA)916Jen/J

  •  
  • MZMMΦ

    marginal zone metallophilic macrophage

  •  
  • pDC

    plasmacytoid dendritic cell

  •  
  • poly(I:C)

    polyinosinic-polycytidylic acid

  •  
  • SDLN

    skin-draining lymph node

  •  
  • Tip-DC

    TNF-α/inducible NO synthase–producing dendritic cell

  •  
  • WT

    wild-type

  •  
  • XCR1

    XC chemokine receptor 1.

1
Steinman
R. M.
2012
.
Decisions about dendritic cells: past, present, and future.
Annu. Rev. Immunol.
30
:
1
22
.
2
Jung
S.
,
Unutmaz
D.
,
Wong
P.
,
Sano
G.
,
De los Santos
K.
,
Sparwasser
T.
,
Wu
S.
,
Vuthoori
S.
,
Ko
K.
,
Zavala
F.
, et al
.
2002
.
In vivo depletion of CD11c+ dendritic cells abrogates priming of CD8+ T cells by exogenous cell-associated antigens.
Immunity
17
:
211
220
.
3
Zammit
D. J.
,
Cauley
L. S.
,
Pham
Q. M.
,
Lefrançois
L.
.
2005
.
Dendritic cells maximize the memory CD8 T cell response to infection.
Immunity
22
:
561
570
.
4
Hochweller
K.
,
Striegler
J.
,
Hämmerling
G. J.
,
Garbi
N.
.
2008
.
A novel CD11c.DTR transgenic mouse for depletion of dendritic cells reveals their requirement for homeostatic proliferation of natural killer cells.
Eur. J. Immunol.
38
:
2776
2783
.
5
Birnberg
T.
,
Bar-On
L.
,
Sapoznikov
A.
,
Caton
M. L.
,
Cervantes-Barragán
L.
,
Makia
D.
,
Krauthgamer
R.
,
Brenner
O.
,
Ludewig
B.
,
Brockschnieder
D.
, et al
.
2008
.
Lack of conventional dendritic cells is compatible with normal development and T cell homeostasis, but causes myeloid proliferative syndrome.
Immunity
29
:
986
997
.
6
Ohnmacht
C.
,
Pullner
A.
,
King
S. B.
,
Drexler
I.
,
Meier
S.
,
Brocker
T.
,
Voehringer
D.
.
2009
.
Constitutive ablation of dendritic cells breaks self-tolerance of CD4 T cells and results in spontaneous fatal autoimmunity.
J. Exp. Med.
206
:
549
559
.
7
Moussion
C.
,
Girard
J. P.
.
2011
.
Dendritic cells control lymphocyte entry to lymph nodes through high endothelial venules.
Nature
479
:
542
546
.
8
Kissenpfennig
A.
,
Henri
S.
,
Dubois
B.
,
Laplace-Builhé
C.
,
Perrin
P.
,
Romani
N.
,
Tripp
C. H.
,
Douillard
P.
,
Leserman
L.
,
Kaiserlian
D.
, et al
.
2005
.
Dynamics and function of Langerhans cells in vivo: dermal dendritic cells colonize lymph node areas distinct from slower migrating Langerhans cells.
Immunity
22
:
643
654
.
9
Liu
K.
,
Nussenzweig
M. C.
.
2010
.
Origin and development of dendritic cells.
Immunol. Rev.
234
:
45
54
.
10
den Haan
J. M.
,
Lehar
S. M.
,
Bevan
M. J.
.
2000
.
CD8(+) but not CD8(-) dendritic cells cross-prime cytotoxic T cells in vivo.
J. Exp. Med.
192
:
1685
1696
.
11
Iyoda
T.
,
Shimoyama
S.
,
Liu
K.
,
Omatsu
Y.
,
Akiyama
Y.
,
Maeda
Y.
,
Takahara
K.
,
Steinman
R. M.
,
Inaba
K.
.
2002
.
The CD8+ dendritic cell subset selectively endocytoses dying cells in culture and in vivo.
J. Exp. Med.
195
:
1289
1302
.
12
Bedoui
S.
,
Whitney
P. G.
,
Waithman
J.
,
Eidsmo
L.
,
Wakim
L.
,
Caminschi
I.
,
Allan
R. S.
,
Wojtasiak
M.
,
Shortman
K.
,
Carbone
F. R.
, et al
.
2009
.
Cross-presentation of viral and self antigens by skin-derived CD103+ dendritic cells.
Nat. Immunol.
10
:
488
495
.
13
Shortman
K.
,
Heath
W. R.
.
2010
.
The CD8+ dendritic cell subset.
Immunol. Rev.
234
:
18
31
.
14
Bonifaz
L. C.
,
Bonnyay
D. P.
,
Charalambous
A.
,
Darguste
D. I.
,
Fujii
S.
,
Soares
H.
,
Brimnes
M. K.
,
Moltedo
B.
,
Moran
T. M.
,
Steinman
R. M.
.
2004
.
In vivo targeting of antigens to maturing dendritic cells via the DEC-205 receptor improves T cell vaccination.
J. Exp. Med.
199
:
815
824
.
15
Sancho
D.
,
Mourão-Sá
D.
,
Joffre
O. P.
,
Schulz
O.
,
Rogers
N. C.
,
Pennington
D. J.
,
Carlyle
J. R.
,
Reis e Sousa
C.
.
2008
.
Tumor therapy in mice via antigen targeting to a novel, DC-restricted C-type lectin.
J. Clin. Invest.
118
:
2098
2110
.
16
Idoyaga
J.
,
Lubkin
A.
,
Fiorese
C.
,
Lahoud
M. H.
,
Caminschi
I.
,
Huang
Y.
,
Rodriguez
A.
,
Clausen
B. E.
,
Park
C. G.
,
Trumpfheller
C.
,
Steinman
R. M.
.
2011
.
Comparable T helper 1 (Th1) and CD8 T-cell immunity by targeting HIV gag p24 to CD8 dendritic cells within antibodies to Langerin, DEC205, and Clec9A.
Proc. Natl. Acad. Sci. USA
108
:
2384
2389
.
17
Schiavoni
G.
,
Mattei
F.
,
Sestili
P.
,
Borghi
P.
,
Venditti
M.
,
Morse
H. C.
 III
,
Belardelli
F.
,
Gabriele
L.
.
2002
.
ICSBP is essential for the development of mouse type I interferon-producing cells and for the generation and activation of CD8α(+) dendritic cells.
J. Exp. Med.
196
:
1415
1425
.
18
Tailor
P.
,
Tamura
T.
,
Morse
H. C.
 III
,
Ozato
K.
.
2008
.
The BXH2 mutation in IRF8 differentially impairs dendritic cell subset development in the mouse.
Blood
111
:
1942
1945
.
19
Hildner
K.
,
Edelson
B. T.
,
Purtha
W. E.
,
Diamond
M.
,
Matsushita
H.
,
Kohyama
M.
,
Calderon
B.
,
Schraml
B. U.
,
Unanue
E. R.
,
Diamond
M. S.
, et al
.
2008
.
Batf3 deficiency reveals a critical role for CD8α+ dendritic cells in cytotoxic T cell immunity.
Science
322
:
1097
1100
.
20
Ginhoux
F.
,
Liu
K.
,
Helft
J.
,
Bogunovic
M.
,
Greter
M.
,
Hashimoto
D.
,
Price
J.
,
Yin
N.
,
Bromberg
J.
,
Lira
S. A.
, et al
.
2009
.
The origin and development of nonlymphoid tissue CD103+ DCs.
J. Exp. Med.
206
:
3115
3130
.
21
Edelson
B. T.
,
KC
W.
,
Juang
R.
,
Kohyama
M.
,
Benoit
L. A.
,
Klekotka
P. A.
,
Moon
C.
,
Albring
J. C.
,
Ise
W.
,
Michael
D. G.
, et al
.
2010
.
Peripheral CD103+ dendritic cells form a unified subset developmentally related to CD8alpha+ conventional dendritic cells.
J. Exp. Med.
207
:
823
836
.
22
Mashayekhi
M.
,
Sandau
M. M.
,
Dunay
I. R.
,
Frickel
E. M.
,
Khan
A.
,
Goldszmid
R. S.
,
Sher
A.
,
Ploegh
H. L.
,
Murphy
T. L.
,
Sibley
L. D.
,
Murphy
K. M.
.
2011
.
CD8α(+) dendritic cells are the critical source of interleukin-12 that controls acute infection by Toxoplasma gondii tachyzoites.
Immunity
35
:
249
259
.
23
Edelson
B. T.
,
Bradstreet
T. R.
,
Hildner
K.
,
Carrero
J. A.
,
Frederick
K. E.
,
Kc
W.
,
Belizaire
R.
,
Aoshi
T.
,
Schreiber
R. D.
,
Miller
M. J.
, et al
.
2011
.
CD8α(+) dendritic cells are an obligate cellular entry point for productive infection by Listeria monocytogenes.
Immunity
35
:
236
248
.
24
Fuertes
M. B.
,
Kacha
A. K.
,
Kline
J.
,
Woo
S. R.
,
Kranz
D. M.
,
Murphy
K. M.
,
Gajewski
T. F.
.
2011
.
Host type I IFN signals are required for antitumor CD8+ T cell responses through CD8{alpha}+ dendritic cells.
J. Exp. Med.
208
:
2005
2016
.
25
Lin
M. L.
,
Zhan
Y.
,
Proietto
A. I.
,
Prato
S.
,
Wu
L.
,
Heath
W. R.
,
Villadangos
J. A.
,
Lew
A. M.
.
2008
.
Selective suicide of cross-presenting CD8+ dendritic cells by cytochrome c injection shows functional heterogeneity within this subset.
Proc. Natl. Acad. Sci. USA
105
:
3029
3034
.
26
Dorner
B. G.
,
Dorner
M. B.
,
Zhou
X.
,
Opitz
C.
,
Mora
A.
,
Güttler
S.
,
Hutloff
A.
,
Mages
H. W.
,
Ranke
K.
,
Schaefer
M.
, et al
.
2009
.
Selective expression of the chemokine receptor XCR1 on cross-presenting dendritic cells determines cooperation with CD8+ T cells.
Immunity
31
:
823
833
.
27
Crozat
K.
,
Guiton
R.
,
Contreras
V.
,
Feuillet
V.
,
Dutertre
C. A.
,
Ventre
E.
,
Vu Manh
T. P.
,
Baranek
T.
,
Storset
A. K.
,
Marvel
J.
, et al
.
2010
.
The XC chemokine receptor 1 is a conserved selective marker of mammalian cells homologous to mouse CD8α+ dendritic cells.
J. Exp. Med.
207
:
1283
1292
.
28
Yamazaki
C.
,
Miyamoto
R.
,
Hoshino
K.
,
Fukuda
Y.
,
Sasaki
I.
,
Saito
M.
,
Ishiguchi
H.
,
Yano
T.
,
Sugiyama
T.
,
Hemmi
H.
, et al
.
2010
.
Conservation of a chemokine system, XCR1 and its ligand, XCL1, between human and mice.
Biochem. Biophys. Res. Commun.
397
:
756
761
.
29
Contreras
V.
,
Urien
C.
,
Guiton
R.
,
Alexandre
Y.
,
Vu Manh
T. P.
,
Andrieu
T.
,
Crozat
K.
,
Jouneau
L.
,
Bertho
N.
,
Epardaud
M.
, et al
.
2010
.
Existence of CD8α-like dendritic cells with a conserved functional specialization and a common molecular signature in distant mammalian species.
J. Immunol.
185
:
3313
3325
.
30
Crozat
K.
,
Tamoutounour
S.
,
Vu Manh
T. P.
,
Fossum
E.
,
Luche
H.
,
Ardouin
L.
,
Guilliams
M.
,
Azukizawa
H.
,
Bogen
B.
,
Malissen
B.
, et al
.
2011
.
Cutting edge: expression of XCR1 defines mouse lymphoid-tissue resident and migratory dendritic cells of the CD8α+ type.
J. Immunol.
187
:
4411
4415
.
31
Bachem
A.
,
Hartung
E.
,
Güttler
S.
,
Mora
A.
,
Zhou
X.
,
Hegemann
A.
,
Plantinga
M.
,
Mazzini
E.
,
Stoitzner
P.
,
Gurka
S.
, et al
.
2012
.
Expression of XCR1 characterizes the Batf3-dependent lineage of dendritic cells capable of antigen cross-presentation.
Front. Immunol.
3
:
214
.
32
Bachem
A.
,
Güttler
S.
,
Hartung
E.
,
Ebstein
F.
,
Schaefer
M.
,
Tannert
A.
,
Salama
A.
,
Movassaghi
K.
,
Opitz
C.
,
Mages
H. W.
, et al
.
2010
.
Superior antigen cross-presentation and XCR1 expression define human CD11c+CD141+ cells as homologues of mouse CD8+ dendritic cells.
J. Exp. Med.
207
:
1273
1281
.
33
Haniffa
M.
,
Shin
A.
,
Bigley
V.
,
McGovern
N.
,
Teo
P.
,
See
P.
,
Wasan
P. S.
,
Wang
X. N.
,
Malinarich
F.
,
Malleret
B.
, et al
.
2012
.
Human tissues contain CD141hi cross-presenting dendritic cells with functional homology to mouse CD103+ nonlymphoid dendritic cells.
Immunity
37
:
60
73
.
34
Nagai
T.
,
Ibata
K.
,
Park
E. S.
,
Kubota
M.
,
Mikoshiba
K.
,
Miyawaki
A.
.
2002
.
A variant of yellow fluorescent protein with fast and efficient maturation for cell-biological applications.
Nat. Biotechnol.
20
:
87
90
.
35
Cheong
C.
,
Matos
I.
,
Choi
J. H.
,
Dandamudi
D. B.
,
Shrestha
E.
,
Longhi
M. P.
,
Jeffrey
K. L.
,
Anthony
R. M.
,
Kluger
C.
,
Nchinda
G.
, et al
.
2010
.
Microbial stimulation fully differentiates monocytes to DC-SIGN/CD209(+) dendritic cells for immune T cell areas.
Cell
143
:
416
429
.
36
Satpathy
A. T.
,
Kc
W.
,
Albring
J. C.
,
Edelson
B. T.
,
Kretzer
N. M.
,
Bhattacharya
D.
,
Murphy
T. L.
,
Murphy
K. M.
.
2012
.
Zbtb46 expression distinguishes classical dendritic cells and their committed progenitors from other immune lineages.
J. Exp. Med.
209
:
1135
1152
.
37
Sugiyama
T.
,
Hoshino
K.
,
Saito
M.
,
Yano
T.
,
Sasaki
I.
,
Yamazaki
C.
,
Akira
S.
,
Kaisho
T.
.
2008
.
Immunoadjuvant effects of polyadenylic:polyuridylic acids through TLR3 and TLR7.
Int. Immunol.
20
:
1
9
.
38
Hoshino
K.
,
Kaisho
T.
,
Iwabe
T.
,
Takeuchi
O.
,
Akira
S.
.
2002
.
Differential involvement of IFN-β in Toll-like receptor-stimulated dendritic cell activation.
Int. Immunol.
14
:
1225
1231
.
39
Dudani
R.
,
Chapdelaine
Y.
,
Hv Hv
Faassen
,
Smith
D. K.
,
Shen
H.
,
Krishnan
L.
,
Sad
S.
.
2002
.
Multiple mechanisms compensate to enhance tumor-protective CD8(+) T cell response in the long-term despite poor CD8(+) T cell priming initially: comparison between an acute versus a chronic intracellular bacterium expressing a model antigen.
J. Immunol.
168
:
5737
5745
.
40
Serbina
N. V.
,
Salazar-Mather
T. P.
,
Biron
C. A.
,
Kuziel
W. A.
,
Pamer
E. G.
.
2003
.
TNF/iNOS-producing dendritic cells mediate innate immune defense against bacterial infection.
Immunity
19
:
59
70
.
41
Backer
R.
,
Schwandt
T.
,
Greuter
M.
,
Oosting
M.
,
Jüngerkes
F.
,
Tüting
T.
,
Boon
L.
,
O’Toole
T.
,
Kraal
G.
,
Limmer
A.
,
den Haan
J. M.
.
2010
.
Effective collaboration between marginal metallophilic macrophages and CD8+ dendritic cells in the generation of cytotoxic T cells.
Proc. Natl. Acad. Sci. USA
107
:
216
221
.
42
Asano
K.
,
Nabeyama
A.
,
Miyake
Y.
,
Qiu
C. H.
,
Kurita
A.
,
Tomura
M.
,
Kanagawa
O.
,
Fujii
S.
,
Tanaka
M.
.
2011
.
CD169-positive macrophages dominate antitumor immunity by crosspresenting dead cell-associated antigens.
Immunity
34
:
85
95
.
43
Probst
H. C.
,
Tschannen
K.
,
Odermatt
B.
,
Schwendener
R.
,
Zinkernagel
R. M.
,
Van Den Broek
M.
.
2005
.
Histological analysis of CD11c-DTR/GFP mice after in vivo depletion of dendritic cells.
Clin. Exp. Immunol.
141
:
398
404
.
44
Alexopoulou
L.
,
Holt
A. C.
,
Medzhitov
R.
,
Flavell
R. A.
.
2001
.
Recognition of double-stranded RNA and activation of NF-kappaB by Toll-like receptor 3.
Nature
413
:
732
738
.
45
Kato
H.
,
Sato
S.
,
Yoneyama
M.
,
Yamamoto
M.
,
Uematsu
S.
,
Matsui
K.
,
Tsujimura
T.
,
Takeda
K.
,
Fujita
T.
,
Takeuchi
O.
,
Akira
S.
.
2005
.
Cell type-specific involvement of RIG-I in antiviral response.
Immunity
23
:
19
28
.
46
Kato
H.
,
Takeuchi
O.
,
Sato
S.
,
Yoneyama
M.
,
Yamamoto
M.
,
Matsui
K.
,
Uematsu
S.
,
Jung
A.
,
Kawai
T.
,
Ishii
K. J.
, et al
.
2006
.
Differential roles of MDA5 and RIG-I helicases in the recognition of RNA viruses.
Nature
441
:
101
105
.
47
Gitlin
L.
,
Barchet
W.
,
Gilfillan
S.
,
Cella
M.
,
Beutler
B.
,
Flavell
R. A.
,
Diamond
M. S.
,
Colonna
M.
.
2006
.
Essential role of mda-5 in type I IFN responses to polyriboinosinic:polyribocytidylic acid and encephalomyocarditis picornavirus.
Proc. Natl. Acad. Sci. USA
103
:
8459
8464
.
48
Kawai
T.
,
Akira
S.
.
2008
.
Toll-like receptor and RIG-I-like receptor signaling.
Ann. N. Y. Acad. Sci.
1143
:
1
20
.
49
Longhi
M. P.
,
Trumpfheller
C.
,
Idoyaga
J.
,
Caskey
M.
,
Matos
I.
,
Kluger
C.
,
Salazar
A. M.
,
Colonna
M.
,
Steinman
R. M.
.
2009
.
Dendritic cells require a systemic type I interferon response to mature and induce CD4+ Th1 immunity with poly IC as adjuvant.
J. Exp. Med.
206
:
1589
1602
.
50
Luber
C. A.
,
Cox
J.
,
Lauterbach
H.
,
Fancke
B.
,
Selbach
M.
,
Tschopp
J.
,
Akira
S.
,
Wiegand
M.
,
Hochrein
H.
,
O’Keeffe
M.
,
Mann
M.
.
2010
.
Quantitative proteomics reveals subset-specific viral recognition in dendritic cells.
Immunity
32
:
279
289
.
51
Neuenhahn
M.
,
Kerksiek
K. M.
,
Nauerth
M.
,
Suhre
M. H.
,
Schiemann
M.
,
Gebhardt
F. E.
,
Stemberger
C.
,
Panthel
K.
,
Schröder
S.
,
Chakraborty
T.
, et al
.
2006
.
CD8α+ dendritic cells are required for efficient entry of Listeria monocytogenes into the spleen.
Immunity
25
:
619
630
.
52
Piva
L.
,
Tetlak
P.
,
Claser
C.
,
Karjalainen
K.
,
Renia
L.
,
Ruedl
C.
.
2012
.
Cutting edge: Clec9A+ dendritic cells mediate the development of experimental cerebral malaria.
J. Immunol.
189
:
1128
1132
.
53
Fukaya
T.
,
Murakami
R.
,
Takagi
H.
,
Sato
K.
,
Sato
Y.
,
Otsuka
H.
,
Ohno
M.
,
Hijikata
A.
,
Ohara
O.
,
Hikida
M.
, et al
.
2012
.
Conditional ablation of CD205+ conventional dendritic cells impacts the regulation of T-cell immunity and homeostasis in vivo.
Proc. Natl. Acad. Sci. USA
109
:
11288
11293
.
54
Jiang
W.
,
Swiggard
W. J.
,
Heufler
C.
,
Peng
M.
,
Mirza
A.
,
Steinman
R. M.
,
Nussenzweig
M. C.
.
1995
.
The receptor DEC-205 expressed by dendritic cells and thymic epithelial cells is involved in antigen processing.
Nature
375
:
151
155
.
55
Bennett
C. L.
,
van Rijn
E.
,
Jung
S.
,
Inaba
K.
,
Steinman
R. M.
,
Kapsenberg
M. L.
,
Clausen
B. E.
.
2005
.
Inducible ablation of mouse Langerhans cells diminishes but fails to abrogate contact hypersensitivity.
J. Cell Biol.
169
:
569
576
.
56
Poulin
L. F.
,
Henri
S.
,
de Bovis
B.
,
Devilard
E.
,
Kissenpfennig
A.
,
Malissen
B.
.
2007
.
The dermis contains langerin+ dendritic cells that develop and function independently of epidermal Langerhans cells.
J. Exp. Med.
204
:
3119
3131
.
57
Ginhoux
F.
,
Collin
M. P.
,
Bogunovic
M.
,
Abel
M.
,
Leboeuf
M.
,
Helft
J.
,
Ochando
J.
,
Kissenpfennig
A.
,
Malissen
B.
,
Grisotto
M.
, et al
.
2007
.
Blood-derived dermal langerin+ dendritic cells survey the skin in the steady state.
J. Exp. Med.
204
:
3133
3146
.
58
Bursch
L. S.
,
Wang
L.
,
Igyarto
B.
,
Kissenpfennig
A.
,
Malissen
B.
,
Kaplan
D. H.
,
Hogquist
K. A.
.
2007
.
Identification of a novel population of Langerin+ dendritic cells.
J. Exp. Med.
204
:
3147
3156
.
59
Henri
S.
,
Poulin
L. F.
,
Tamoutounour
S.
,
Ardouin
L.
,
Guilliams
M.
,
de Bovis
B.
,
Devilard
E.
,
Viret
C.
,
Azukizawa
H.
,
Kissenpfennig
A.
,
Malissen
B.
.
2010
.
CD207+ CD103+ dermal dendritic cells cross-present keratinocyte-derived antigens irrespective of the presence of Langerhans cells.
J. Exp. Med.
207
:
189
206
.
60
Diamond
M. S.
,
Kinder
M.
,
Matsushita
H.
,
Mashayekhi
M.
,
Dunn
G. P.
,
Archambault
J. M.
,
Lee
H.
,
Arthur
C. D.
,
White
J. M.
,
Kalinke
U.
, et al
.
2011
.
Type I interferon is selectively required by dendritic cells for immune rejection of tumors.
J. Exp. Med.
208
:
1989
2003
.
61
Schnorrer
P.
,
Behrens
G. M.
,
Wilson
N. S.
,
Pooley
J. L.
,
Smith
C. M.
,
El-Sukkari
D.
,
Davey
G.
,
Kupresanin
F.
,
Li
M.
,
Maraskovsky
E.
, et al
.
2006
.
The dominant role of CD8+ dendritic cells in cross-presentation is not dictated by antigen capture.
Proc. Natl. Acad. Sci. USA
103
:
10729
10734
.
62
Dudziak, D., A. O. Kamphorst, G. F. Heidkamp, V. R. Buchholz, C. Trumpfheller, S. Yamazaki, C. Cheong, K. Liu, H. W. Lee, C. G. Park, et al. 2007. Differential antigen processing by dendritic cell subsets in vivo. Science 315: 107–111. PubMed doi:10.1126/science.1136080

The authors have no financial conflicts of interest.