Nucleotide Oligomerization Binding Domain-Like Receptor Signaling Enhances Dendritic Cell-Mediated Cross-Priming In Vivo

Dendritic cells (DCs) present internalized exogenous Ags to CD4⁺ T cells via MHC class II molecules and endogenously synthesized Ags to CD8⁺ T cells through MHC class I molecules. In addition, DCs have the unique ability to deliver exogenous Ag to the MHC class I-restricted Ag presentation pathway and generate CD8⁺ T cell immunity to viral infections and cancer, a phenomenon known as cross-priming (1). Signals mediated by TLRs (2–4), type I IFNs (5), CD4⁺ T cells (6), immune complexes (7), and heat shock proteins (8) augment cross-presentation and the cross-priming of Ag-specific CD8⁺ T cells in vitro and in vivo. In particular, TLR-mediated signals enhance exogenous Ag capture and the cell-surface expression of costimulatory molecules (2–4), which efficiently induces cross-priming.

Two nucleotide oligomerization binding domain (Nod)-like receptor family members, Nod1 and Nod2, recognize bacterial peptidoglycan (PGN)-related molecules. Nod1 recognizes PGN-related dipeptide, γ-d-glutamyl-meso-diaminopimelic acid (iE-DAP), which is produced by most Gram-negative and certain Gram-positive bacteria (9, 10), and Nod2 recognizes muramyldipeptide (MDP), a component of all PGNs (11, 12). Although TLRs are associated with plasma membrane or lysosomal/endosomal vesicles, Nod1 and Nod2 are expressed predominantly in the cytosol of APCs and epithelial cells (13, 14). Nod proteins interact with the receptor-interacting protein-like–interacting caspase-like apoptosis regulatory protein kinase (RICK) (also known as RIP2) and caspase-recruitment domain 9 to activate the MAPK and NF-κB signaling pathways, promoting the production of proinflammatory factors, such as chemokines and cytokines (15–18). Nod1 and Nod2 sense multiple bacteria and are critically involved in protective immune responses in vivo against Helicobacter pylori and Listeria monocytogenes, respectively (19, 20). Dysfunction of NOD proteins is associated with human diseases such as atopic eczema and asthma, Crohn’s disease, graft-versus-host disease, Blau syndrome, and sarcoidosis (13, 14). Recent studies also showed that Nod1 contributes to the initiation of adaptive immune responses [i.e., Th differentiation (21)], and Nod1 and Nod2 are important for host defense, especially in hosts insensitive to TLR stimulation (22).

In this study, we investigated the role played by Nod proteins in inducing the cross-priming of CD8⁺ T cells, and we demonstrate that Nod1 and Nod2 ligands significantly enhance the induction of DC-mediated cross-priming. Together with previous studies, our results indicate that, for the optimization of DC-mediated cross-priming, although TLRs act as sensors on the cell-surface and in the endosome, Nod1/2 act as critical sensors once bacteria invade the cell or escape into the cytosol.

C57BL/6 (B6) mice were purchased from Japan SLC (Hamamatsu, Japan) and Japan Clea (Meguro, Japan). The B6 OT-I Rag2^−/−, B6 Nod1^−/−, B6 Tap1^−/−, and Tlr2^−/−4^−/−9^−/− mice were described previously (2, 9, 23). B6 Nod1^−/−OT-I mice were generated by crossing in-house. All mice were maintained in specific pathogen-free conditions, and all animal experiments were approved by the Institutional Animal Care Committee of Akita University.

FK565, FK156, and iE-DAP compounds were provided by Astellas Pharma (Tsukuba, Japan). MDP, OVA protein (grade VI), propidium iodide, and epoxomicin were purchased from Sigma-Aldrich (St. Louis, MO); FITC-OVA and CFSE were purchased from Invitrogen (Carlsbad, CA); poly(I:C) was purchased from Amersham Biosciences (Piscataway, NJ); and the OVA_257–264 peptide (SIINFEKL) was purchased from Medical and Biological Laboratories (Nagoya, Japan).

DCs were prepared from the spleens of mice, as described previously (24). In brief, collagenase-digested spleen cells were suspended in a dense 28% BSA solution in PBS (1.080 g/ml; BSA was purchased from MBL), overlaid with FCS-free RPMI 1640 medium (Sigma-Aldrich), and centrifuged in a swing-bucket rotor at 9500 × g for 15 min at 4°C. Cells at the interface were collected, washed, and resuspended in PBS. Then, DCs were positively selected with anti-CD11c MicroBeads and an AutoMacs separation system (Miltenyi Biotec, Bergisch Gladbach, Germany). Purified DCs were routinely >95% CD11c⁺I-A⁺. For CD8α⁺ DC depletion, spleen cells were negatively selected with anti-CD8α⁺ MicroBeads and the AutoMacs separation system. The depleted populations were positively selected with anti-CD11c MicroBeads and AutoMacs. To purify the CD8α⁺ and CD8α⁻ DCs, spleen cells were stained with the following Abs, all conjugated to biotin: anti-pan NK cells (DX5), anti-CD45R (RA3-6B), anti-CD3e (145-2C11), and anti-CD19 (MB19-1) for 30 min at 4°C and then negatively selected with streptavidin MicroBeads (Miltenyi Biotec) and the AutoMacs system. The depleted populations were further stained with FITC–anti-CD11c (N418) and PE–anti-CD8α⁺ (53-6.7) for 30 min at 4°C, and propidium iodide solution was added to exclude dead cells. CD8α⁺ and CD8α⁻ CD11c^high DCs were purified on a MoFlo instrument (Beckman Coulter, Brea, CA). The purity of the cells was ≥98%. MicroBeads and mAbs were purchased from Miltenyi Biotec and eBioscience (San Diego, CA), respectively.

Wild-type (WT) and Nod1^−/− mice were given an i.v. injection of OVA (50 μg) with or without 50 μg FK565, FK156, iE-DAP compounds, MDP, or poly(I:C). Three hours later, splenic DCs (1 × 10⁵) from these mice were cocultured with OT-I cells (1 × 10⁵) in 96-well round-bottom plates at 37°C. After 48 h, the cells were pulsed with ³H-thymidine (0.5 μCi/well) for 12 h and harvested. To examine the effect of a proteasome inhibitor (epoxomicin), splenic DCs were treated for 3 h in vitro with OVA or with OVA plus FK565 with or without epoxomicin (1 μM). Then, the DCs (1 × 10⁵) were cocultured with OT-I cells (1 × 10⁵) for 60 h and pulsed with ³H-thymidine for an additional 12 h. As a control experiment, splenic DCs were treated for 3 h in vitro with or without epoxomicin (1 μM), and the DCs were cocultured with OT-I cells in the presence of OVA_257–264 peptide (0.3 μg/ml) for 48 h and pulsed with ³H-thymidine for an additional 12 h. In some experiments, CD8α⁺ or CD8α⁻ DCs (3 × 10⁴) were cocultured with OT-I cells (1 × 10⁵), which were isolated from the spleen of B6 OT-I Rag2^−/− mice. To evaluate OT-I cell proliferation in vivo, CFSE (5 μM)-labeled OT-I cells (4 × 10⁶) were adoptively transferred into WT and Nod1^−/− mice that had been given an i.v. injection of OVA alone (50 μg) or OVA plus 50 μg FK565, iE-DAP, or MDP 3 h prior to the cell transfer. Two days after the transfer, the proliferation of OT-I cells was determined on a FACS Calibur (BD Biosciences, San Jose, CA) with FlowJo software (Tree Star, Ashland, OR) on the basis of CFSE dilution. The levels of IL-2 in the 48-h culture supernatants, another indicator of OT-I cell proliferation, were determined by ELISA kits (eBioscience) and the Softmax PRO data analysis program (Molecular Devices, Sunnyvale, CA).

mRNA extracted from CD8α⁺ and CD8α⁻ DCs was reverse-transcribed into cDNA and amplified by real-time PCR. PCR assays were performed using a Light Cycler System (Roche Applied Science, Indianapolis, IN) to measure SYBR green (Roche) incorporation. Primers for Nod1, Nod2, Tap1, Sec61α1, Canx, Calr, and Cst3 were designed and synthesized by Nihon Gene Research Laboratories (Sendai, Japan) (Table I). The relative amounts of target genes determined the cycle number, which was at the cross-point between the amplification plot and the threshold. Values were normalized to the β-actin mRNA within each sample.

Levels of IFN-α, IFN-β, TNF-α, and IL-6 in the sera were measured by ELISA kits (IFN-α, IFN-β; PBL Biochemical Laboratories [Piscataway, NJ]; TNF-α, IL-6; BioLegend, San Diego, CA) according to the manufacturers’ instructions.

To evaluate the endocytosis of OVA in vitro, DCs were prepared from WT mice that were given an i.v. injection of OVA (500 μg) alone or OVA plus 250 μg FK565, FK156, or MDP. Sixty minutes later, the DCs were isolated from the mice and treated with anti-FcγRIII/II (2.4G2) to block nonspecific Ab binding and then stained with APC-conjugated anti-CD11c (N418) and PE-conjugated anti-CD8α⁺ (53-6.7). Next, FITC-OVA (100 μg/ml) was added to the culture for 30 min at 37°C or 4°C. After three washes with cold PBS, the level of internalized FITC-OVA was determined on a FACS Calibur (BD Biosciences) with FlowJo software (Tree Star).

To detect the H-2K^b-OVA_257–264 peptide (SIINFEKL) complex and costimulatory molecules on CD8α⁺ and CD8α⁻ DCs, splenic cells were prepared from WT and Nod1^−/− mice 48 h after they were given an i.v. injection of OVA (500 μg) with or without 250 μg FK565 or MDP. The plasmacytoid DCs (pDCs) and the NK, T, and B cells were depleted as described above. The cells were treated with anti-FcγRIII/II (2.4G2) to block nonspecific Ab binding and stained with anti-H-2K^b-SIINFEKL complex (25D1.16) or mouse IgG1 isotype control, APC-conjugated anti-CD11c (N418), and PE-conjugated anti-CD8α⁺ (53-6.7), followed by FITC-conjugated rat anti-mouse IgG. To detect costimulatory molecules, cells were stained with biotin–anti-CD40 (1C10), biotin–anti-CD86 (GL1), or biotin-rat IgG2a isotype control, followed by streptavidin-Alexa Fluor 488 (Invitrogen). To detect mannose receptor (MR), cells were fixed and permeabilized with Cytofix/Cytoperm (BD Biosciences) and stained with anti-mouse CD206-Alexa Fluor 488 (MR5D3) (Serotec, Kidlington, U.K.) or isotype-control, rat IgG2a-Alexa 488 (YTH71.3) (Serotec). To detect the OT-I expression of intracellular IFN-γ, spleen cells from the mice prepared for the in vivo OT-I proliferation assay were stimulated with OVA_257–264 peptide (1 μg/ml) in GolgiPlug (BD Biosciences) for 6 h. The cells were then fixed, permeabilized with Cytofix/Cytoperm, and stained with PE anti-mouse IFN-γ (XMG1.2) or PE rat IgG1 isotype control. To detect endogenous T cells expressing OVA_257–264-specific TCRs, Ag-MHC I tetramer staining was performed. WT mice were given injections of OVA alone (500 μg) or OVA (500 μg) plus FK565 (250 μg) on each of three consecutive days, and the spleen cells isolated from these mice were cultured with OVA_257–264-pulsed spleen cells at a 1:1 ratio for 6 d. After the culture, the cells were stained with FITC-anti-CD8 (KT15) (MBL) and PE-H-2K^b-OVA_257–264 tetramer (MBL) or negative tetramer as a control. All mAbs, if not otherwise specified, were from eBioscience. Data were collected on a FACS Calibur and analyzed with FlowJo software.

The in vivo CTL assay was carried out as described (4). In brief, WT and Nod1^−/− mice were given an i.v. injection of OVA (50 μg) alone, OVA plus FK565 (50 μg), OVA plus FK156 (500 μg), or OVA plus MDP (500 μg), respectively. Three hours later, OT-I cells (1 × 10⁶) were transferred into the mice. Four days after the transfer, a 1:1 mixture of OVA peptide (10 μg/ml)-pulsed and CFSE (0.5 mM)-labeled spleen cells (1 × 10⁷) and nonpulsed and CFSE (5 mM)-labeled spleen cells (1 × 10⁷) was transferred. Twenty-four hours after the target cell transfer, flow cytometry was used to measure the CTL activity by monitoring the specific deletion of the CFSE^low target cells. The percentage of specific lysis was calculated on the basis of the following formula: [1 − (ratio in unprimed mice/ratio in primed mice)] × 100; the ratio in unprimed and primed mice represents the percentage of CFSE^high cells/percentage of CFSE^low cells. To evaluate the antitumor CTL activity in vivo, WT mice were given one injection of OVA (100 μg) with or without FK565 (100 μg) on each of two consecutive days. Three hours after the second injection, OT-I cells (1 × 10³) were transferred into the mice. Four days after the transfer, OVA-expressing EG-7 cells (1 × 10⁷) were implanted intradermally to evaluate the antitumor effect of the CTL, which was determined by measuring the tumor’s perpendicular diameter. Tumor volume was calculated by the following formula: tumor volume = 0.4 × length (mm) × [width (mm)] (25). To evaluate the antibacterial CTL activity in vivo, WT mice were given an i.v. injection of OVA (50 μg) with or without FK565 (50 μg). Three hours later, OT-I cells (2 × 10⁴) were transferred into the mice. Four days after the transfer, the mice were infected with OVA-expressing L. monocytogenes (2 × 10⁷ CFU), provided by Y. Yoshikai of Kyushu University, and the bacterial titers in the spleens of the infected mice were determined 48 h later.

Femurs and tibias were flushed with a syringe, and the suspension was passed through a 70-mm nylon mesh. The RBCs were lysed, and the T cells were removed. To make chimeras, mice were lethally irradiated (two doses of 600 rad, separated by 3 h) and were subsequently reconstituted with bone marrow cells (5 × 10⁶). Mice were allowed to ‘rest’ for 6–8 wk before use.

The statistical significance was evaluated by the Student t test.

The DC-mediated cross-presentation of OVA is enhanced by simultaneous stimulation with TLRs (2–4). To examine the roles of Nod1 and Nod2 in DC-mediated cross-presentation, WT and Nod1^−/− mice were given an injection of OVA alone, OVA plus iE-DAP or FK565 as a Nod1 ligand, or OVA plus MDP as a Nod2 ligand. Three hours later, DCs were isolated from the spleen of these mice and cocultured with OVA-specific OT-I cells from B6 OT-I Rag2^−/− mice. Notably, DCs from WT mice given an injection of OVA plus these Nod ligands significantly enhanced the proliferation of OT-I cells compared with DCs from WT mice given OVA alone (Fig. 1A). The elevated OT-I cell proliferation was minimal when the DCs were from Nod1^−/− mice given OVA plus iE-DAP or FK565; however, the elevation was the same as in WT mice when the Nod1^−/− mice were given OVA plus MDP, indicating that the effect of the Nod1 ligands depended strictly on Nod1 signaling. Similar Nod1-dependent OT-I proliferation was observed when the DCs were from WT mice given an injection of OVA plus other iE-DAP derivatives, including FK156 (Fig. 1B) and (Supplemental Fig. 1). In this context, FK565, FK156, and iE-DAP derivatives induced cross-presentation more efficiently than iE-DAP, probably, at least in part, because these molecules contain acyl residues conjugated to the core iE-DAP, which enhance host cell membrane permeability and, thereby, the ability to stimulate Nod1 (26, 27). Consistent with previous findings (2–4), poly(I:C)-stimulated DCs also enhanced the cross-presentation of OT-I cells (Supplemental Fig. 2A). As expected from previous studies, poly(I:C) administration substantially induced both type I IFNs and proinflammatory cytokines, such as TNF-α and IL-6, in the sera of WT mice. Interestingly, compared with poly(I:C), FK565 administration induced few type I IFNs and TNF-α, as well as IL-6 at a much reduced level (Supplemental Fig. 3).

Nod1 is expressed in immune cells, such as DCs, T cells, and B cells, as well as in epithelial tissues (13, 14). Interestingly, Nod1 expression by nonhematopoietic cells is required for the Nod1-induced Th2 polarization of OVA-specific naive CD4⁺ T cells (21). To determine whether the effect of Nod1 ligands was attributable to the activation of immune cells or epithelial cells, we made bone marrow chimeras of WT and Nod1^−/− mice (Fig. 1C), by transferring the bone marrow of each genotype into a lethally irradiated mouse of the other genotype. Interestingly, little enhanced OT-I proliferation was observed when DCs from chimeras that did not express Nod1 in their hematopoietic cells were used (Fig. 1C), indicating that Nod1 expression by hematopoietic cells was required. To determine which of the hematopoietic cells effected the enhancement, we cocultured Nod1^−/− OT-I cells with DCs from WT mice treated with OVA plus FK565. The OT-I proliferation was enhanced (Supplemental Fig. 2B), indicating that Nod1 must be expressed by DCs rather than T cells for it to induce enhanced cross-presentation.

There are two known pathways for Ag cross-presentation: one is TAP dependent (28–32), and the other is TAP independent (33–36). We investigated the effect of DCs from Tap1^−/− mice treated with OVA and FK565 on OT-I cells and found that OT-I proliferation was not enhanced, indicating that OVA is cross-presented TAP-dependently (Fig. 1D). Based on these results, we examined the role of proteasomal degradation in the TAP-dependent cross-presentation pathway. DCs were isolated from the spleen of WT mice and treated with OVA alone or OVA plus FK565 in the absence or presence of the proteasome inhibitor epoxomicin in vitro. After washing, the DCs were cocultured with OT-I cells. Compared with DCs treated with OVA alone, DCs treated with OVA plus FK565 significantly enhanced the proliferation of OT-I cells. This enhanced proliferation was completely blocked when epoxomicin-treated DCs were used in the cocultures with OT-I cells (Fig. 1E, left panel). In contrast, the same epoxomicin treatment did not affect the OVA peptide-induced proliferation of OT-I cells (Fig. 1E, right panel). Collectively, these results indicate that FK565 enhances TAP- and proteasome-dependent cross-presentation by DCs.

CD8α⁺ DCs are a DC subset specialized for potent Ag cross-presentation (37–42). To identify the DC subset critical for the Nod1-dependent enhanced cross-presentation, CD8α⁺ DCs and CD8α⁺-depleted (CD8α⁻) DCs from WT mice treated with OVA plus FK565 were tested for cross-presentation to OT-I cells in vitro. OT-I cell proliferation was not increased by CD8α⁻ DCs (Fig. 2A, 2B), but it was efficiently enhanced by purified CD8α⁺ DCs (Fig. 2B), which also correlated well with the IL-2 production by the OT-I cells (Fig. 2D, 2E). Similar results were obtained using MDP (Fig. 2C). These results demonstrated that the CD8α⁺ DC subset, rather than the CD8α⁻ DC subset, is critical for the Nod-dependent enhancement of cross-presentation. As a control for contamination of the OVA preparation by TLR ligands PGN and LPS, we repeated this experiment with DCs from OVA- and FK565-injected Tlr2^−/−4^−/−9^−/− mice and found no reduction in the effect (Supplemental Fig. 4A).

We next examined the expression levels of Tap1, Sec61α1, calnexin (Canx), calreticulin (Calr), and cystatin C (Cst3) molecules directly or indirectly involved in cross-presentation (43, 44). Compared with CD8α⁻ DCs, the expression levels of these molecules were efficiently upregulated in CD8α⁺ DCs upon FK565 and MDP stimulation (Fig. 3). Under steady-state conditions, these proteins were expressed at relatively higher levels in CD8α⁺ DCs than in CD8α⁻ DCs, as previously reported (41). Of note, the expression of Nod2, but not Nod1, was upregulated in both DC subsets (Fig. 3). Our data indicate that Nod signaling enhances the expression of molecules involved in cross-presentation, predominantly in CD8α⁺ DCs.

LPS, a representative TLR ligand, enhances the DC-mediated endocytosis of OVA and cross-presentation (3). Therefore, we examined whether Nod ligands also enhanced the DC-mediated endocytosis of OVA in vitro. Under steady-state conditions, the internalization of FITC-labeled soluble OVA by CD8α⁺ DCs was higher than that by CD8α⁻ DCs, but it was not enhanced by stimulation with FK565, FK156, or MDP (Fig. 4A). In this context, MR-mediated endocytosis of soluble OVA is essential for cross-presentation (42). However, the expression of the MR was not altered in the CD8α⁺ or CD8α⁻ DC subsets by exposure to the same stimuli (Fig. 4B), indicating that the Nod ligand-mediated promotion of DC cross-presentation is probably not caused by increased OVA uptake by DCs.

To identify the mechanisms responsible for the Nod ligand-mediated promotion of DC cross-presentation, DCs were prepared from WT and Nod1^−/− mice after an injection of OVA with or without FK565, and the expression levels of Ag presentation and costimulatory molecules were examined (Fig. 5A, 5B). We found that the levels of the H-2K^b-OVA peptide complex [detected by an mAb (25D1.16) specific for it (45)] and costimulatory molecules, including CD40 and CD86, were substantially increased on the CD8α⁺ DCs, but minimally on the CD8α⁻ DCs, of WT mice after an injection of FK565 with OVA, which correlated well with the relative upregulation in intracellular components for cross-presentation of CD8α⁺ DCs (Fig. 3). In contrast, no increased expression of the H-2K^b-OVA peptide complex or costimulatory molecules was detected on DCs from Nod1^−/− mice following FK565 stimulation. The same phenotypical changes were observed on the DCs of WT mice injected with OVA plus MDP (Fig. 5C). Thus, Nod signaling probably enhances the CD8α⁺ DC-mediated cross-presentation, at least in part, by upregulating Ag presentation and costimulatory molecule expression in vitro.

To examine whether Nod ligands enhance the cross-presentation and proliferation of Ag-specific CD8⁺ T cells in vivo, 5,6-CFSE–labeled OT-I cells were adoptively transferred into WT and Nod1^−/− mice that had been given an injection of OVA plus FK565, iE-DAP, or MDP. Two days after the transfer, we examined the OT-I cell proliferation, which was evaluated from the CFSE dilution, and the number of IFN-γ⁺ OT-I cells, and we noted that they were significantly and Nod1 dependently greater in WT mice given OVA plus FK565 or iE-DAP and Nod1 independently (Nod2 dependently) in mice given OVA plus MDP, compared with WT mice given OVA alone (Fig. 6A).

An in vivo killing assay was performed to determine whether the IFN-γ⁺ OT-I cells exhibited OVA-specific cytotoxic activity in vivo. Four days after the transfer of OT-I cells into WT and Nod1^−/− mice given OVA plus FK565, a 1:1 mixture of OVA-peptide–pulsed (CFSE^lo) and -unpulsed (CFSE^hi) spleen cells was transferred into the same mice as target cells. Twenty-four hours later, the CTL activity, estimated from the reduction in target cell number, was significantly augmented in WT, but not Nod1^−/−, mice given an injection of OVA plus FK565, compared with that of control mice given OVA alone (Fig. 6B, 6C). Furthermore, the OVA-specific in vivo cytotoxic activity was augmented by injected FK156 or MDP (Fig. 6D). In addition, the FK565-augmented OVA-specific cytotoxic activity was observed in Tlr2^−/−4^−/−9^−/− mice (Supplemental Fig. 4B). Our findings indicate that Nod signaling induces elevated cross-priming of Ag-specific CD8⁺ T cells in vivo.

To evaluate the physiological significance of the Nod-dependent elevation of DC cross-priming in vivo, OT-I cells alone, FK565 alone, FK565 plus OT-I cells, OVA alone, OVA plus FK565, OVA plus OT-I cells, or OVA, FK565, and OT-I cells were injected into WT mice. OVA and FK565 were injected once per day for two consecutive days. Four days after the second injection, OVA-expressing EG-7 tumor cells were implanted intradermally into the mice of each group. Tumor growth was rapid in the untreated mice and those treated with FK565 alone, and there was no significant difference in the tumor volumes between these two groups (Fig. 7A, 7B, 7D). Compared with these groups, tumor growth was slightly slower in mice given OVA alone, OT-I cells alone, FK565 plus OT-I cells, or OVA plus OT-I cells (Fig. 7A–F, 7H), and it was markedly slower in the mice treated with OVA plus FK565 (Fig. 7A, 7G), suggesting that this treatment induced endogenous T cell responses against the EG-7 cells that led to a substantial reduction of tumor cell growth. In support of this finding, injections of OVA and FK565 on three consecutive days clearly induced H-2K^b-OVA tetramer⁺ CD8⁺ T cells in a subsequent culture with OVA peptide-pulsed spleen cells (Fig. 7J). The most effective tumor suppression was observed in mice treated with OVA, FK565, and OT-I cells (Fig. 7A, 7I).

To further examine the relevance of the Nod-dependent elevation of DC cross-priming in vivo, we also used a bacterial infection model, in which mice treated with OVA and OT-I cells or OVA, FK565, and OT-I cells were subsequently infected with OVA-expressing L. monocytogenes. The bacterial titers of the spleens were compared among the treatment groups 48 h later and were significantly lower in the spleens of the mice treated with OVA, FK565, and OT-I cells (Supplemental Fig. 5).

These results indicate that FK565 potentiates DC-mediated cross-priming, leading to effective antitumor and antibacterial CTL activity in vivo.

In this study, we showed that Nod1 and Nod2 signals significantly augment CD8α⁺ DC-mediated cross-presentation and the cross-priming of Ag-specific CD8⁺ T cells in vivo, through the upregulation of the MHC class I-dependent Ag presentation pathway and costimulation. As a result, considerably enhanced Ag-specific CTL activity was induced, leading to the effective elimination of tumor cells (Fig. 8).

Three models of cross-presentation are currently proposed (46). The canonical model proposes that endocytosed or phagocytosed Ag is exported into the cytosol and degraded into antigenic peptides in the proteasome. The peptides are then transported by TAP into the endoplasmic reticulum (ER) and loaded onto MHC class I molecules for presentation at the cell surface. A revised version of the canonical model is the ER-phagosome fusion model, which is based on the evidence that ER-associated proteins (e.g., TAP, Sec61, and calnexin) are present in phagosomes (29–31, 47). Third, the early endosome model proposes that cross-presentation occurs in early endosomes in a TAP-dependent (48) or TAP-independent (33–36) manner. In the TAP-dependent early endosome model, recruitment of TAP to the early endosomes, an essential step for the cross-presentation of soluble Ag, is induced by TLR4-MyD88–mediated signals (48). In the TAP-independent model, human pDCs cross-present exogenous Ags to memory CD8⁺ T cells through the early endocytic compartment (36). However, Nod signaling did not significantly increase this early endosome-dependent cross-presentation in pDCs (data not shown). Because soluble OVA is taken up by MR and selectively supplied into the early endosome (42), and Nod1 signals TAP-dependently augmented the DC-mediated cross-presentation of soluble OVA, Nod-RICK–mediated signals might mimic the TLR4-MyD88 signals.

Although the appropriate stimulation can induce CD8α⁻ DC-mediated cross-presentation (7), CD8α⁺ DCs are the predominant subset involved in cross-presentation because of their efficient phagocytosis of dead cells (38) and the predominant expression of C-type lectin receptors, such as DEC205 (CD205) (41) and MR (42), which correlate well with the constitutive expression of critical intracellular components of the cross-presentation pathway (41). MR is of particular interest because it takes up soluble OVA (42). However, our data suggested that neither the expression nor the function of MR is upregulated by Nod-mediated signals.

Because microbial components at the cell surface and endolysosome are recognized by TLRs, whereas those in the cytosol are sensed by Nod-like receptors and RIG-I–like helicases (RLHs), distinct and complementary surveillance systems apparently occur in these compartments and seem to be necessary for the potentiation of cross-presentation in vivo. Interestingly, certain bacteria, such as L. monocytogenes, contain Nod1 and Nod2 ligands and use a strategy to escape from the endolysosome into the cytosol, which protects them from the protease-dependent degradation process. Bacillus spp. have Nod1- and Nod2-stimulatory activities, but lack TLR4-stimulatory activity (49). We propose a model whereby once Nod-dependent cross-priming is effectively induced during such bacterial infection, Ag-specific CD8⁺ cells secrete IFN-γ, which, in turn, stimulates DCs and macrophages to produce NO, oxygen radicals, and TNF-α, which are critical for eliminating these bacteria. In support of this scenario, peritoneal exudate macrophages of WT mice secrete NO in response to tracheal cytotoxin, a Nod1 ligand, in an IFN-γ–dependent manner (50). In addition, Nod1- and Nod2-mediated signals are particularly important in TLR-tolerized macrophages, and upon systemic infection with L. monocytogenes, bacterial clearance is strictly dependent on Nod1 and Nod2 when mice are pretreated with TLR ligands, which make them refractory to TLR signals (22).

In viral infections, TLR- and RLH-mediated signals also effectively induce DC cross-priming. However, the viral structure contains various ligands recognized by the TLRs and RLHs, such as RIG-I and MDA5; thus, many viral proteins suppress TLR- and RLH-mediated signaling pathways to evade the immune surveillance system (51). For example, vaccinia virus A46R and A52R inhibit multiple TLR pathways, including TLR3; hepatitis C virus NS3/4A cleaves Trif and IFNB-promoter stimulator 1, thereby abrogating TLR3/4 and RLH pathways; paramyxovirus V proteins and influenza virus NS1 bind to MDA5 and RIG-I, respectively, and inhibit RLH pathways; and Ebola virus VP35 inhibits the RIG-I pathway. Given that the viral particle does not contain Nod ligands and, thus, the Nod-dependent pathway must be relatively intact, our findings may suggest a new avenue of approach toward the development of novel preventive and therapeutic applications for viral infections as well as cancer.

We thank K. Yamashita, Y. Abe, and A. Onai for animal care and experimental support; J. Imai and M. Maruya for helpful suggestions; W.R. Heath, The Walter and Eliza Hall Institute of Medical Research, for the B6 OT-I Rag2^−/− mice; Toru Abo, Niigata University, for the B6 Tap1^−/− mice; R.N. Germain, National Institutes of Health, for the 25D1.16 hybridoma; Astellas Pharma for the FK565, FK156, and iE-DAP compounds; Y.F. and K.F., Osaka University, for the iE-DAP; and T. Nishimura, Hokkaido University, for the EG-7 cells.

Disclosures The authors have no financial conflicts of interest.