The XC chemokine receptor 1 is a conserved selective marker of mammalian cells homologous to mouse CD8α⁺ dendritic cells

To assess the cross-presentation capacity of human BDCA3⁺ DCs, we purified these cells together with other types of DCs. BDCA1⁺ DCs, BDCA3⁺ DCs, and pDCs were magnetically enriched from PBMCs by negative selection and then positively sorted by flow cytometry as depicted in Fig. 1 A. The purity of sorted cells was >95% as observed by postsorting FACS reanalysis. Sorted HLA-A2⁺ DC subsets were cultured with apoptotic uninfected, or chronically HIV-infected, H9 cells in the presence of maturation stimuli, poly I:C for BDCA3⁺ DCs, LPS or poly I:C for BDCA1⁺ DCs, and R848 for pDCs. They were then cultured with an HIV-1 Pol-specific CD8⁺ T cell line. To avoid viral replication, experiments were performed in the presence of Saquinavir, an HIV protease inhibitor (Hoeffel et al., 2007). BDCA3⁺ DCs induced 40% more IFN-γ ELISPOTs than poly I:C–stimulated BDCA1⁺ DCs or R848-stimulated pDCs and 60% more as compared with LPS-stimulated BDCA1⁺ DCs (Fig. 1 B). Therefore, as expected from their gene expression profile (Robbins et al., 2008; Crozat et al., 2010), human BDCA3⁺ DCs had higher cross-presentation capacities than other human blood DC subsets and represent functional homologues of mouse CD8α⁺ DCs for cross-presentation.

Based on gene chip analyses, the expression of the XCR1 gene is a selective marker for both human BDCA3⁺ DCs and mouse CD8α⁺ DCs when compared with a large panel of cell types, including other DC subsets, a variety of other leukocytes and of nonhematopoietic cells, and all types of tissue (Fig. 2 A). This contrasts with the markers currently used to identify these DC subsets, which are more broadly expressed as clearly evident for BDCA3 in human or CADM1 in human and mouse (Fig. 2 A). Further extending this striking convergent observation in mouse and human DCs, we found that in a distant mammalian species, i.e., sheep, the XCR1 gene was also selectively expressed in the CD8α⁺-like skin lymph CD26⁺ DC (Fig. 2 B). Thus, among the genes differentially expressed between DC subsets in a conserved manner across the three mammalian species examined, the XCR1 gene stands out as the most discriminating marker for mouse spleen CD8α⁺ DCs, human blood BDCA3⁺ DCs, and sheep lymph CD26⁺ DCs (Fig. 2, A and B). This is also the case for tonsil BDCA3⁺ DCs (Fig. 2 A, orange circle). Xcr1 mRNA was also selectively expressed in CD11c⁺SIRPα⁻CD24⁺ cells from BM FLT3-L cultures (Fig. 3 A), which correspond to a subset of in vitro–derived DCs equivalent to spleen CD8α⁺ DCs which can be referred to as eCD8α⁺ DCs (Naik et al., 2005).

To assess XCR1 protein expression pattern on mouse leukocytes, we resorted to a reporter/knockout mouse strain expressing the β-galactosidase enzyme under the control of the Xcr1 promoter. When total spleen leukocytes were isolated from XCR1^−/− mice and stained for β-galactosidase activity using fluorescein di-β-d-galactopyranoside (FDG) as a substrate, >80% of the cells that stained positive with FDG were CD8α⁺ DCs, and >75% of CD8α⁺ DCs were FDG⁺ (Fig. 2 C). Thus, Xcr1 was confirmed to be expressed selectively by the majority of mouse CD8α⁺ DCs. Altogether, these results showed that XCR1 is a conserved and very discriminating marker for the identification of mouse, ovine, and human CD8α⁺-type DCs.

We then performed transwell migration assays with recombinant XCL1 and mixtures of the three subsets of mouse spleen DCs (Fig. 4 A) or their in vitro equivalents derived in BM FLT3-L cultures (Fig. 3 B). In both instances, XCL1 induced the specific migration of CD8α⁺-type DCs. Moreover, only XCR1^+/+ and XCR1^+/− CD8α⁺ or eCD8α⁺ DCs were responsive to XCL1 (Fig. 3 B and Fig. 4 A), but not their XCR1^−/− counterparts, demonstrating that XCR1 is required for DC responses to XCL1. We then performed transwell assays with enriched DC preparations from human blood and sheep lymph. Only human BDCA3⁺ DCs and sheep CD26⁺ DCs, and none of the other leukocyte subsets tested, were found to migrate in response to XCL1 (Fig. 4 A). Therefore, XCR1 is required for DC responses to XCL1, and its engagement induces the selective motility of mouse, ovine, and human CD8α⁺-type DCs.

XCL1 is produced and secreted specifically by activated mouse and human NK and CD8⁺ T cells (Müller et al., 1995; Dorner et al., 2002), whereas no XCL1 protein is detected by intracellular staining in quiescent mouse NK cells (Dorner et al., 2002). The Xcl1 gene is specifically expressed in NK and CD8⁺ T cells already at steady state not only in mouse and human (Fig. 4 B) but also in sheep (Fig. 4 C). Moreover, similar to what we previously reported for CCL5 (Walzer et al., 2003), we show that XCL1 mRNA is selectively expressed to high levels in antiviral central memory CD8⁺ T lymphocytes (T_CMs) and, to a lesser extent, in memory T cells induced by sterile inflammatory conditions (inflammatory memory CD8⁺ T cells [T_IMs]; Fig. 4 D). Cytokine secretion profiling of the mouse CD8⁺ T cell subsets demonstrated a rapid and higher level secretion of XCL1 by T_IM after antigen-specific reactivation in vitro compared with naive CD8⁺ T cells (unpublished data). The selective expression of Xcl1 in mouse T_CM was confirmed independently by mining published mouse microarray datasets retrieved through the GEO database (Sarkar et al., 2008; Fig. S1 A). Therefore, XCL1 mRNA is specifically expressed at steady state in NK and, to a lesser extent, CD8⁺ T cells, further induced in these cell types upon activation, including during viral infections in mouse and human (Fig. S1; Hyrcza et al., 2007; Sarkar et al., 2008), and stored in T_CM. Hence, XCR1 expression is likely integral to the unique potency of CD8α⁺-type DCs for interaction with NK and CD8⁺ T cells.

Sequences similar to the Xcr1 locus are available from many mammals including all the species with a full genome assembly. Only one gene per species was found, located in the same genomic region as several CC chemokine receptors (CCRs). A unique Xcr1 sequence was also identified from genomes of birds and the Anole lizard. Several sequences similar to Xcr1 were present in teleost fish, such as zebrafish and stickleback, but only one in others like Fugu. The Xcr1 sequence is well conserved in all eutherian species examined (>75% identity with the human sequence), in marsupials (∼70% identity), in birds and reptiles (60–65% identity), and in fish (50–60% identity; Fig. S2). These sequences were aligned with those of several human CCRs, and phylogenetic analyses were performed using different methods. Trees reconstructed were congruent and demonstrated that the Xcr1-like sequences found in the different vertebrate taxa are true orthologues of the mammalian Xcr1 gene (Fig. 5 A and Fig. S2), which was confirmed by synteny (Fig. 5, B and C). The multiple genes found in fish probably derive from the complex history of fish genomes that have been marked by large duplication events, fast evolution, and extensive genome rearrangements (Ravi and Venkatesh, 2008). It is remarkable that both stickleback Xcr1 co-orthologues are closely linked to paralogues of the marker FYCO1, which is also encoded in the close neighborhood of mammalian Xcr1 (Fig. 5 B-C). This conserved synteny indicates that the stickleback Xcr1-like genes indeed constitute valid co-orthologues of the mammalian receptor. Hence, XCR1 is a promising candidate marker to identify, isolate, study, and possibly target CD8α⁺-type DCs in all vertebrates.

CD8α⁺ DCs, but not CD11b⁺ DCs or pDCs, isolated from mice infected with Lm present bacterial antigens for the priming of naive CD8⁺ T cells (Belz et al., 2005). Treatment of Lm-infected mice with pertussis toxin compromises the later activation of adoptively transferred antibacterial CD8⁺ T cells (Aoshi et al., 2008). Pertussis toxin is a nonspecific inhibitor of G proteins that is widely used to inhibit chemokine receptor signaling. Thus, it was suggested that a chemokine receptor expressed on DCs is required to enable them to induce CD8⁺ T cell responses to Lm infection. We reasoned that XCR1 could contribute to this process. Therefore, we challenged XCR1^−/− and XCR1^+/+ mice with Lm to evaluate the impact of XCR1 deficiency on the control of the bacteria load and on the induction of CD8⁺ T cell responses (Fig. 6). As compared with XCR1^+/+ mice, XCR1^−/− animals infected with WT Lm displayed a log increase in their bacteria loads measured in livers and spleens at day 3 after inoculation (Fig. 6 A). There was still a trend toward higher bacterial loads in the spleen and liver of XCR1^−/− mice at day 4 after inoculation, although it was weaker than at day 3 (unpublished data). Because anti-Lm CD8⁺ T cell responses are readily detectable at 3–4 d after infection (Khanna et al., 2007), their alteration in XCR1^−/− animals could contribute to the higher bacterial replication in these animals. To compare the levels of anti-Lm CD8⁺ T cell responses between XCR1^−/− and XCR1^+/+ mice, they were infected with a recombinant Lm strain expressing the ovalbumin antigen (Lm-OVA), and anti-OVA CD8⁺ T cells were enumerated by MHC class I tetramer staining at day 4 after inoculation. XCR1^−/− mice exhibited significantly fewer anti-Lm CD8⁺ T cells than XCR1^+/+ animals (Fig. 6 B). Thus, these data suggest that XCR1 activity in CD8α⁺ DCs allows them to more efficiently induce antigen-specific CD8⁺ T cell responses and, hence, promote bacteria control early after infection with Lm.

The demonstration that human blood BDCA3⁺ DCs isolated ex vivo are more potent than their BDCA1⁺ counterparts or pDCs for antigen cross-presentation and CD8⁺ T cell activation fulfills our prediction, based on our comparative genomics studies, that human BDCA3⁺ DCs represent, in this respect, a functional homologue of mouse CD8α⁺ DCs and an interesting target for the design of innovative vaccination or immunotherapeutic strategies against infections by intracellular pathogens or cancer (Robbins et al., 2008; Crozat et al., 2010). More generally, our results demonstrate a common specialization of mouse CD8α⁺, human BDCA3⁺, and sheep CD26⁺ DCs for CD8⁺ T cell activation through cross-presentation of exogenous antigens and identical high selective expression of XCR1 by these cells that is consistent with the role of this molecule in the promotion of the interactions between CD8α⁺-like DCs and CD8⁺ T cells. These results establish a definitive proof of concept that validates our comparative genomics approach to hunt for conserved functions based on conserved molecular pathways for DC subsets across several mammalian species (Robbins et al., 2008; Crozat et al., 2010).This work exemplifies how critical knowledge can be gained to understand important immunological processes by using comparisons between species as a tool and by developing tight collaborations between teams working with different animal models. In particular, this work illustrates the utility of nonprimate animal models for understanding human immunology, provided that a rational approach is followed to help focusing on conserved biological processes and molecular functions.

Based on gene chip analyses, we previously reported that mouse CD8α⁺ DCs specifically expressed the Xcr1 gene, to high levels, whereas none of the other leukocyte subsets examined did. Indeed, in the Supplemental material of our paper (Robbins et al., 2008), we reported Xcr1 as the third most specific gene for mouse CD8α⁺ DCs in our analyses. This result was confirmed and significantly extended by Dorner et al. (2009). Indeed, these authors additionally reported that XCR1 engagement induces calcium mobilization specifically in CD8α⁺ DCs and that XCL1 responses are required for the induction of optimal primary CD8⁺ T cell responses by CD8α⁺ DCs in response to soluble model antigens in vitro and in vivo. No functional ProbeSet specific for the human XCR1 gene had been identified on the Affymetrix Human Genome U133 Plus 2.0 Array until very recently. XCR1 was, therefore, not reported by us (Robbins et al., 2008) or by Dorner et al. (2009) to be specifically expressed in human BDCA3⁺ DCs. However, by mapping on the human genome all individual orphan ProbeSets that were giving a high and specific signal only in human BDCA3⁺ DCs as compared with other blood leukocyte subsets, we identified that the 1561226_at ProbeSet mapped to the human XCR1 gene, which was later confirmed by the November 2009 update of the Ensembl genome browser. Hence, we demonstrate in this paper, for the first time, to the best of our knowledge, a conserved specific expression and activity of XCR1 on CD8α⁺-type DCs derived in vitro in mouse BM FLT3-L cultures or isolated ex vivo from human and sheep. In our hands, XCL1 did not induce any maturation of DCs, either alone or in combination with IFN-γ, and did not seem to affect TLR-induced maturation of DCs in vitro (unpublished data). We confirm the specific expression of XCR1 ligands in NK cells and CD8⁺ T lymphocytes in mouse and human and extend it to sheep. We show that Xcl1 mRNA is stored in quiescent NK cells and antiviral memory CD8⁺ T lymphocytes. We show that XCR1 promotes the induction of CD8⁺ T cell responses and the control of bacterial load in vivo early during Lm infection. Our phylogenetic analyses show the presence of a unique and well conserved XCR1 orthologue in all higher vertebrates from reptiles to human. Altogether, these observations strongly suggest that the XCL1→XCR1 chemotactic pathway constitutes one of the critical and conserved molecular bases promoting selective and efficient cross talk between CD8α⁺-type DCs and CD8⁺ T lymphocytes as well as NK cells.

In summary, our results demonstrate a functional homology between mouse CD8α⁺, sheep CD26⁺, and human BDCA3⁺ DCs and identify XCR1 as the best candidate to date for the identification of CD8α⁺-type DCs in all vertebrates, with a very narrow expression pattern highly restricted to these cell types. In addition, we show that, contrary to the molecules classically used to identify CD8α⁺-like DCs, XCR1 is not a mere phenotypical marker of these cells but actually contributes to confer on them their exquisite ability for CD8⁺ T cell activation. Thus, our study brings significant advances toward the generation of a unifying phenotypic and functional view of mammalian DC subsets, with major implications both for basic and applied immunology. Our findings open a way for the general survey by immunohistofluorescence of the anatomical location of CD8α⁺-type DCs and for their specific targeting in vivo for vaccination against intracellular pathogens or tumors threatening human or domestic animal health.

For each experiment, >500 × 10⁶ PBMCs were isolated by Ficoll-Hypaque density gradient centrifugation (GE Healthcare) from peripheral blood cytapheresis residues obtained from normal healthy volunteer donors through the Etablissement Français du Sang according to the French ethics committee on human experimentations, within a convention with Institut National de la Santé et de la Recherche Médicale. Primary DCs (BDCA1⁺ DCs, BDCA3⁺ DCs, and pDCs) were isolated from PBMCs depleted of CD3⁺ and CD14⁺ cells using CD3⁺ and CD14⁺ MACS microbeads and an AutoMACS (Miltenyi Biotec). Enriched cells were labeled and DCs were sorted on a FACSAria flow cytometer (BD) as events negative for CD14, CD16, and CD19 and positive for HLA-DR and BDCA1 for BDCA1⁺ DCs, HLA-DR and BDCA3 for BDCA3⁺ DCs, or HLA-DR and CD123 for pDCs. The following antibodies were used: HLA-DR-PerCp (G46-6), CD14-PE-Cy7 (M5E2), CD16-APC-H7 (3G8; all obtained from BD), CD19-ECD (J3-119; Beckman Coulter), BDCA1-Pacific Blue (L161; BioLegend), BDCA3-APC (AD5-14H12), and CD123-PE (AC145; both obtained from Miltenyi Biotec). Cell sorting gave a purity >95%. CD8⁺ primary T cell lines specific for the HLA-A2–restricted HIV-Pol_476-84 epitope were generated as previously described (Hoeffel et al., 2007) using PBMCs from HLA-A2⁺ HIV⁺ individuals from a cohort study with the approval of Cochin Hospital’s ethics committee. They were cultured in the presence of 1 µM Pol_476-484 in complete medium (CM; RPMI 1640–GlutaMAX containing 100 U/ml penicillin, 100 mg/ml streptomycin, 1% nonessential amino acids, 1 mM sodium pyruvate, and 10 mM Hepes buffer; Life Technologies) + 5% human serum (BioWest) and maintained for 4 wk at 0.7 × 10⁶–10⁶ cells/ml, adding 10 U/ml IL-2 (Roche) twice a week. Uninfected (American Type Culture Collection; HTB-176) or chronically infected (American Type Culture Collection; CRL-8543) HLA-A2⁻ H9 cells were UV-irradiated (400 mJ/cm² at 312 nm) to induce their apoptosis and incubated at 10⁶ cells/ml for 2 h in CM + 10% FCS (Invitrogen). Purified BDCA-1⁺ DCs, BDCA-3⁺ DCs, and pDCs were cultured with apoptotic cells at a 1:1 ratio for 16 h in the presence of a maturation signal, 100 ng/ml LPS (Sigma-Aldrich), or 10 µg/ml poly I:C (InvivoGen) or 10 µM R848 (InvivoGen) in CM + 5% human AB serum. After loading, DCs were extensively washed and used (5,000/well) in a 16-h IFN-γ ELISPOT assay with HIV-specific CD8⁺ T cells lines as effectors (15,000/well). All cultures were performed in the presence of 1 µM Saquinavir (Roche), an HIV-protease inhibitor, to avoid HIV replication (Hoeffel et al., 2007).

C57BL/6Ncrl mice were provided by Charles River. XCR1^−/− mice (B6.129P2-Xcr1^<tm1Dgen>/J mice; MGI:3604538) were generated by Deltagen. Frozen XCR1^+/− embryos were provided by The Jackson Laboratory for strain reviviscence in the Centre d’Immunologie de Marseille-Luminy (CIML). In these mice, the β-galactosidase is expressed under the Xcr1 promoter and in place of Xcr1, which permits a detection of the natural expression pattern of Xcr1. All mice were kept under pathogen-free conditions in the CIML animal care facility. Mouse experiments were conducted in accordance with institutional guidelines for animal care and use (French Provence Ethical Committee Protocol no. 04/2005 and US Office of Laboratory Animal Welfare Assurance A5665-01). Sheep were cannulated at the Center of Research in Interventional Imaging, Institut National de La Recherche Agronomique-Assistance Publique Hôpitaux de Paris (Jouy en Josas, France) for collecting prescapular skin afferent lymph as previously described (Schwartz-Cornil et al., 2006), and the sheep were housed in the Domestic Animal Experimental Unit INRA (Paris-Sud ethic commitee no. 08002A, Jouy en Josas, France).

T_IMs were generated by intraperitoneal injection of 50 nmol influenza A nucleoprotein NP366-374 in PBS in naive thymectomized transgenic F5 mice (Mbitikon-Kobo et al., 2009). Antiviral T_CMs were generated by immunizing F5 mice with a recombinant vaccinia virus including the NP68 epitope as previously described (Cottalorda et al., 2009). Spleens and lymph nodes were collected 6 wk after infection for cell preparation as described in the next section.

Mouse splenocytes were prepared and DC subsets sorted as described previously (Robbins et al., 2008). CD8⁺ T cells purified from mouse spleen and lymph nodes were stained for surface expression of CD44, CD8, and F5 TCR (Walzer et al., 2002). Naive CD8⁺ T cells, T_IM, and antiviral T_CM were sorted by flow cytometry as CD8⁺/CD44^low, CD8⁺/CD44^int, and CD8⁺/CD44^high/F5 TCR⁺, respectively. β-Galactosidase enzyme activity in splenocytes from XCR1^−/− mice was revealed using FDG as a substrate according to the manufacturer’s instructions (Invitrogen). In brief, FDG was diluted in warm distilled water at 2 mM, and cells were mixed 1:1 with the substrate. After an incubation of one minute at 37°C, cells were placed on ice, and the enzymatic reaction was terminated by adding ice-cold PBS. Cells were stained with a marker of viability and promptly acquired by flow cytometry. Sheep CD4⁺ T, CD8⁺ T, NK, and B cells were enriched from afferent lymph, and monocytes were enriched from blood. Sheep leukocytes were stained with mouse mAbs against CD1b (clone TH97A; Parsons et al., 1991), CD26 (CC69; Gliddon and Howard, 2002), CD8α (CACT80C; MacHugh and Sopp, 1991), CD4 (ST4; Hopkins, 1991), CD14 (CAM36A; Berthon and Hopkins, 1996), and NKp46 (AKS4). AKS4 was produced against bovine NKp46 according to a previously described protocol (Storset et al., 2004), and it cross reacts with ovine NK cells defined as CD16⁺/CD14⁻ blood leukocytes (Elhmouzi-Younes et al., 2010). NKp46 has been shown to be the best discriminating and conserved marker for NK cell identification in mammals (Storset et al., 2004; Walzer et al., 2007). The DU2-104 mAb is used to characterize all B cells in sheep (Press et al., 1993). Sheep cells were reacted with 2 µg/ml of primary antibodies, stained with fluorochrom-conjugated purified Ig directed to specific mouse isotypes, and sorted by FACS with >95% purity.

The generation and analysis of microarray data for mouse and human DC subsets were described elsewhere (Robbins et al., 2008; Zucchini et al., 2008). All datasets used are public and their references for download from databases are given in the legends of figures. A compendium of human hgu133plus2 Affymetrix.CEL files was established, using Bioconductor (release 2.5) in the R statistical environment (version 2.10.1). This consisted of the quality control assessment using the AffyPLM package (Bolstad et al., 2004; e.g., visual examination of hybridization chip images, relative log expression, and normalized unscaled standard errors plots) and the robust multichip analysis normalization (Irizarry et al., 2003) with the Affy package. Normalized log₂ scaled values were then averaged among replicate arrays for each probe, and standard deviations were calculated. For real-time PCR experiments, total RNA was prepared with either the RNA NOW reagent (Biogentex) for mouse CD8⁺ T cells or the RNeasy Isolation kit (QIAGEN) for mouse DCs. RNA was reverse transcribed into cDNA with the QuantiTect reverse transcription kit (QIAGEN), and the cDNA was analyzed by real-time PCR using the Power SYBR green PCR master mix (Applied Biosystems). Primers were as follows: mouse Xcr1, 5′-CTCAGCCTTGTGGGTAACAGC-3′ and 5′-ACAGGCAGTAGACAGGAGAAC-3′; mouse Ccl5, 5′-GCACCTGCCTCACCATATGG-3′ and 5′-AGCACTTGCTGCTGGTGTAG-3′; mouse Ifng, 5′-CCAGCGCCAAGCATTCAATGAG-3′ and 5′-GACAATCTCTTCCCCACCCCG-3′; mouse ubiquitin, 5′-AAGAATTCAGATCGGATGACACACT-3′ and 5′-GCCACTTGGAGGTTGACACTTT-3′; sheep XCR1, 5′-TGCCATCTTCCACAAGGTGTT-3′ and 5′-ACGGAGGCGAGGAACCA-3′; sheep XCL1, 5′-TGAGCCAGAGCAAGCCTACA-3′ and 5′-TCACTACCCAGTCAGGGTCACA-3′; and sheep GAPDH, 5′-CACCATCTTCCAGGAGCGAG-3′ and 5′-CCAGCATCACCCCACTTGAT-3′. Relative gene expression was calculated using the ΔΔCt method with b-actin for mouse DCs, ubiquitin for mouse CD8⁺ T cells, and GAPDH for sheep leukocytes as the endogenous control housekeeping genes.

Mouse DCs were obtained by density gradient enrichment from splenocytes or derived in vitro from BM Flt3-L cultures as previously described (Sjölin et al., 2006). Human DCs were enriched using the human Blood Dendritic Cell Isolation kit II (Miltenyi Biotec). Sheep DCs were enriched by density gradient. Cells were resuspended in migration medium (0.5% BSA in RPMI) and distributed in the upper chamber of a 24 Transwell system insert (5-µm pore size; Corning). The lower chamber was filled with migration medium alone or containing 300 ng/ml of mouse recombinant XCL1 (R&D Systems) for mouse cells, 500 ng/ml of human recombinant XCL1 (R&D Systems) for human cells, and 100 ng/ml of recombinant mouse XCL1 for sheep cells. Cells were incubated for 3–4 h at 37°C in 5% CO₂. A constant number of polybead polystyrene microspheres (Polyscience) was added in each lower chamber before harvesting migrated cells. Migrated cells were then stained and analyzed by flow cytometry. A constant event of microspheres for each well was acquired to evaluate the absolute number of cells recovered after migration. The percentage of migrated cells was calculated by the following formula for each subset: [number of cells having migrated in response to XCL-1 − number of cells having migrated spontaneously]/[total number of input cells] × 100. All experiments were performed with duplicate wells.

2 × 10⁵ sorted naive CD8⁺ T cells and T_IM were stimulated for 6 h with the NP68 peptide at a concentration of 10 nM in 200 µl of medium. Supernatants were analyzed using Rodent MAP 1.0 (Rules Based Medicine).

The Lm WT strain 10403S and the Lm strain that had been engineered to express OVA (Lm-OVA; gift from Grégoire Lauvau, Albert Einstein College of Medicine, Bronx, NY) were prepared from clones grown from organs of infected mice. Stocks of bacteria were kept frozen at −80°C. For infections, a sample of the frozen stock was grown to a logarithmic phase (OD₆₀₀ = 0.05–0.15) in broth heart infusion (BHI) medium (Sigma-Aldrich), diluted in PBS, and injected into the retroorbital vein. Bacterial loads in organs were evaluated after infection of mice with one LD₅₀ of WT Lm (5 × 10⁴ bacteria). CD8⁺ T cell responses were measured in animals infected with 0.1 LD₅₀ of Lm-OVA (5 × 10³ bacteria). The dose of Lm injected was confirmed by CFU counting of the inoculum. To measure bacterial titers, spleens and livers were harvested at day 3 and dissociated on nylon meshes in 5 ml of dissociation buffer (0.2% NP-40). Serial dilutions were performed and plated onto BHI agar plates. For analysis of Lm-specific CD8⁺ T cell responses at day 4, OVA-specific CD8⁺ T cells were readily detected using OVA-iTag-H-2K^b-SIINFEKL PE-conjugated MHC class I tetramers (Beckman Coulter).

12 representatives of XCR1 sequences were retrieved from Ensembl genome assemblies of relevant vertebrate species. No good candidate for XCR1 counterpart was found in other phyla. Chromosomal locations were obtained from Ensembl (November 2009 release). Conserved syntenies were searched manually in Ensembl and with the Genomicus tool (http://www.dyogen.ens.fr/genomicus/cgi-bin/search.pl). The amino acid sequences of XCR1 domain were aligned using ClustalW and Mega4 programs. Any site at which the alignment introduces a gap in any sequence was excluded, and a comparable set of sites was therefore used for the phylogenetic analyses. The phylogenetic analysis was made using two methods: NJ and parsimony using the MEGA 4 program (Tamura et al., 2007).

Fig. S1 documents the expression pattern of XCL1 in mouse NK cells, mouse T_CM, and human CD8⁺ T cells during viral infections. Fig. S2 shows the alignment of XCR1 protein sequences and the Maximum Parsimony phylogenetic analysis of Xcr1 evolution in vertebrates.

We thank Michel Bonneau (Centre de Recherche en Imagerie Interventionnelle, Jouy en Josas, France) for the sheep surgery, Céline Urien for excellent technical support, the Domestic Animal Experimental Unit (UCEA, Institut National de La Recherche Agronomique, Jouy en Josas, France) for sheep care, the Centre d’Immunologie de Marseille-Luminy (CIML) staff of the mouse care facilities and the flow cytometry core facility for assistance, and Grégoire Lauvau (Albert Einstein College of Medicine, New York, USA) for providing the Lm strains. We thank Lene Vimeux (Centre National de la Recherche Scientifique [CNRS] UMR 8104) for generating T cell lines, and Antonio Cosma, Sabrina Guenounou, Thibault Andrieu, and Roger Le Grand (flow cytometry cell sorting core facility of the Commissariat à l’Énergie Atomique [CEA] in Fontenay aux Roses, France). We thank Bernard Charley and Bernard Malissen for critical reading of the manuscript.

This work was supported by Association pour la recherche sur le cancer (postdoctoral fellowship to K. Crozat and research grants to M. Dalod and to J. Marvel), CNRS (postdoctoral fellowship to R. Guiton), Agence nationale de recherches sur le SIDA et les hépatites virales (fellowship to C-A. Dutertre), Institut National de la Santé et de la Recherche Médicale (Jeune Chercheur fellowship to V. Feuillet), Ligue nationale contre le cancer (fellowship to E. Ventre), institutional grants to CIML, and the Agence nationale de la recherche program Genanimal 2006 (research grant to I. Schwartz-Cornil).

The authors declare no competing financial interests.