To generate vaccines that protect mucosal surfaces, a better understanding of the cells required in vivo for activation of the adaptive immune response following mucosal immunization is required. CD11chigh conventional dendritic cells (cDCs) have been shown to be necessary for activation of naive CD8+ T cells in vivo, but the role of cDCs in CD4+ T cell activation is still unclear, especially at mucosal surfaces. The activation of naive Ag-specific CD4+ T cells and the generation of Abs following mucosal administration of Ag with or without the potent mucosal adjuvant cholera toxin were therefore analyzed in mice depleted of CD11chigh cDCs. Our results show that cDCs are absolutely required for activation of CD4+ T cells after oral and nasal immunization. Ag-specific IgG titers in serum, as well as Ag-specific intestinal IgA, were completely abrogated after feeding mice OVA and cholera toxin. However, giving a very high dose of Ag, 30-fold more than required to detect T cell proliferation, to cDC-ablated mice resulted in proliferation of Ag-specific CD4+ T cells. This proliferation was not inhibited by additional depletion of plasmacytoid DCs or in cDC-depleted mice whose B cells were MHC-II deficient. This study therefore demonstrates that cDCs are required for successful mucosal immunization, unless a very high dose of Ag is administered.

Mucosal immunization is often necessary to generate immunity at mucosal surfaces (1). This protection requires generation of Ag-specific T cells and Abs. To activate Ag-specific T cells, peptides must be displayed on MHC molecules on the surface of activated APCs, particularly dendritic cells (DCs)3. Indeed, the superior capacity of DCs to activate naive T cells ex vivo following nasal and oral immunizations has been determined in several studies (2, 3, 4, 5, 6, 7, 8, 9, 10). Additionally, the strong T cell priming capacity of Ag-loaded DCs has not only been shown after i.v. transfer but also after intratracheal injection (11). A transgenic system has been developed where diphtheria toxin (DTx) administration conditionally ablates CD11chigh conventional DCs (cDCs) in vivo (12). Using these CD11c-DTx receptor (CD11c-DTR) mice, the absolute requirement for cDCs to activate naive CD8+ T cells following infections and parenteral immunizations has been demonstrated in several reports (12, 13, 14, 15, 16).

Despite the requirement for cDCs in naive CD8+ T cell activation, the role of cDCs in CD4+ T cell activation is not clear. For example, although depletion of CD11chigh cells significantly reduces the expansion of adoptively transferred vesicular stomatatis virus (VSV)-specific CD4+ T cells following i.v. infection (17), it does not affect the VSV-driven generation of CD4+ T cell-derived cytokines (13). CD4+ T cell activation is delayed but not abrogated in Mycobacterium tuberculosis-infected cDC-depleted mice (18). Additionally, cDCs are not essential for achieving maximal clonal expansion of CD4+ T cells following vaccinia virus infection (19). Finally, a recent study has shown that CD4+ T cells in lymph nodes (LNs), but not spleen, can be activated in vivo following parenteral administration of protein, even in the absence of cDCs (16). This suggests that cells other than cDCs, such as plasmacytoid DCs (pDCs), can prime CD4+ T cells in a tissue- or route-dependent fashion. Despite these data from parenteral immunization systems, the types of APCs required for activation of CD4+ T cells in vivo following mucosal immunization have not been investigated.

A role for DCs in B cell priming has been suggested by both in vitro studies and investigations using adoptively transferred Ag-loaded DCs (20, 21, 22). However, a recent study showed that MHC-II molecules on B cells could be rapidly loaded with peptides despite cDC ablation, illustrating that DCs are not required for loading Ag-specific B cells (23). Furthermore, other experiments using CD11c-DTR mice have shown that differentiation of plasma cells in T-independent immune responses is independent of cDCs (24). T-independent Ab responses to VSV infection do not require CD11chigh cells, despite the fact that these cells are required for induction of Abs to purified VSV-G protein (17).

Ab responses to usually inert protein Ags are very poor unless the Ag is coadministered with an adjuvant. Mucosal immunization, particularly by the oral route, is hampered by the paucity of useful adjuvants. Cholera toxin (CT) is a strong oral adjuvant that, despite its toxicity to humans, is an important tool for gaining insight into the mechanisms of oral immunization and the design of novel mucosal vaccines. To function as a mucosal adjuvant, CT binds ubiquitously expressed GM1 ganglioside receptors (25). CT also causes activation of DCs after mucosal administration (2, 25). However, the extent to which cDCs are required for CT to function as a mucosal adjuvant in vivo has not been addressed.

Here we examine the activation of naive CD4+ Ag-specific T cells and the induction of specific Abs following mucosal administration of OVA and CT in mice depleted of cDCs in vivo. Our results show that cDCs are required for activation of CD4+ T cells after oral and nasal administration of Ag, with or without CT. Additionally, cDC depletion abrogates the induction of Ag-specific intestinal IgA responses as well as Ag-specific serum IgG production following oral immunization. When a very high dose of Ag is given, proliferation of CD4+ T cells is detected in cDC-ablated mice. We demonstrate that this is not due to a compensatory role of B cells, pDCs, or recruited CD11bhighCD11c−/low myeloid cells. These results therefore show that cDCs are required for mucosally induced adaptive immune responses unless a very high dose of Ag is administered.

CD11c-DTR transgenic (B6.FVB-Tg Itgax-DTR/GFP 57Lan/J) mice (12), OT-II (C57BL/6) TCR transgenic mice harboring OVA-specific CD4+ T cells (26), B cell-deficient mice (C57BL/10-IgH-6tm1Cgn) (μMt) (27), and MHC-II-deficient mice (B6.129S2-H2dlAb1-Ea/J) (MHC-II-KO) (28) were all bred and maintained under specific pathogen-free conditions at the Experimental Biomedicine Animal Facility, University of Gothenburg, Göteborg, Sweden. CD11c-DTR Tg:μMt mice were obtained by intercrossing the respective lines, and CD45.1+CD45.2+OT-II mice were obtained from OT-II × C57BL/6-CD45.1 mating. All experiments were performed using protocols approved by the Swedish government’s Animal Ethics Committee and followed institutional animal use and care guidelines.

Femurs and tibias were taken from donor mice and BM was flushed out. Cells were passed through a 70-μm nylon mesh and RBCs were lysed. CD11c-DTR BM donor cells were used to generate CD11c-DTR→wild-type chimeras (CD11c-DTR/WT) (29). Alternatively, a mix of CD11c-DTR Tg:μMt and MHC-II−/− BM donor cells at a 4:1 ratio were used to generate mixed CD11c-DTR/MHC-IIB−/−→WT chimeras (CD11c-DTR/MHC-IIB−/−) (30). C57BL/6 recipient mice were irradiated (1000 rad) before 2–5 × 106 BM cells were transferred i.v. Mice were rested for 7 wk before use. For cDC depletion, mice were injected i.p. with 4 ng/g body weight DTx (Sigma-Aldrich) 3 days and 18 h before immunization. For combined cDC and pDC depletion the mice were, in addition to the DTx injections, injected i.p. with 0.5 mg of Ab 120.G8 (31) (Schering-Plough) for 3 consecutive days before immunization. Chimerism and DC depletions were confirmed at the time of immunization by flow cytometric analysis of splenocytes.

For oral immunizations, food was removed 16–18 h before the mice receiving OVA (Sigma-Aldrich) in 3% NaHCO3 with or without 10 μg of CT (Sigma-Aldrich). For nasal i.p. and i.v. immunizations, mice received LPS-low OVA containing 1.1 endotoxin unit/mg of protein. OVA was given nasally with or without 1 μg of CT or 50 μg of CpG 1826 (Operon) in a total volume of 20 μl and given i.p. with or without 2 μg of CT.

Single-cell suspensions were washed in FACS buffer (PBS containing 2% FCS and 1 mM EDTA), which was used for all stainings. Cells were stained for 15 min at 4°C with the following Abs conjugated to FITC, PE, allophycocyanin, biotin, Pacific Blue, Alexa Fluor 700, or allophycocyanin-Alexa Fluor 780: anti-CD3 (clone 17A2), anti-CD4 (clone L3T4), anti-CD11b (clone M1/70), anti-CD11c (clones HL3 and N418), anti-CD19 (clone 1D3), anti-CD45.1 (A20), anti-CD69 (H1.2F3), anti-Vα2 (clone B20.1), anti-MHC-II (clone M5/114), anti-Ly6C (clone AL-21), anti-Ly6G (clone 1A8), and anti-PDCA1 (clone JF05-1C2.4.1). All mAbs except anti-CD11c (clone N418) (BioLegend), anti-MHC-II (eBioscience), and mPDCA-1 (Miltenyi Biotec) were purchased from BD Pharmingen. SA-Qdot605 (Invitrogen) was used as secondary reagent to detect biotinylated mAbs. 7-Aminoactinomycin D (7AAD; Sigma-Aldrich) was included in all stainings to define nonviable cells. Cells were acquired on a FACSCalibur or LSR II (both BD Bioscience) and analyzed using FlowJo software (Tree Star).

For histological studies of nasal-associated lymphoid tissue, the head was stripped of facial skin and severed along the line between the upper and lower jaw. Heads were then fixed in 4% paraformaldehyde, decalcified, and embedded in Tissue-Tek OCT compound. All other tissues were collected in Histocon (Histolab), embedded in Tissue-Tek OCT compound, and snap-frozen. Sections (7 μm) were cut on a cryostat (Leica), picked up on microslides, and stored at −70°C until used. All slides, except those for nasal-associated lymphoid tissue studies, were fixed on ice in 50% acetone for 30 s and then in 100% acetone for 5 min before staining. The sections were blocked with normal horse serum in PBS for 20 min in a humidifying chamber. For detection of DCs, sections were incubated with biotinylated anti-CD11c for 30–60 min followed by Alexa Fluor 488-conjugated streptavidin (Invitrogen) for 30–60 min. Sections were double stained with PE-conjugated anti-B220 for visualization of B cells. The B220 signal was further enhanced with Cy3-conjugated anti-rat IgG (Jackson ImmunoResearch Laboratories). Sections were examined using a confocal laser scanning microscope (Zeiss LSM 510 META). Images were further processed using an LSM Image Browser (Zeiss).

Spleens and LNs were taken from OT-II mice, and CD4+ T cells were negatively enriched by magnetic separation using an autoMACS (Miltenyi Biotec) and CD4+ T cell isolation kit (Miltenyi Biotec). Cells were then labeled with CFSE (Invitrogen) by resuspending them at 1 × 107/ml in 5 μM CFSE in serum-free PBS for 5 min at 37°C. The reaction was quenched with an equal volume of FCS. Cells were washed, resuspended in PBS, and 3–5 × 106 were injected i.v. in to mice that had, 70 and 2 h earlier, either been given DTx or not. Eighteen hours later the mice were immunized with OVA with or without CT. Three or 5 days later, mice were sacrificed and then single-cell suspensions from lymphoid tissues were stained and analyzed by flow cytometry to determine the frequency of adoptively transferred cells that had entered division.

To determine OVA-specific Ab titers, 96-well plates (Greiner Bioscience) were coated with OVA (20 μg/ml) and then blocked with PBS/BSA. To determine CT subunit B (CTB)-specific Ab titers, 96-well plates (Nunc) were coated with GM-1 (0.3 nmol/ml) and, after blocking with PBS/BSA, CTB (0.5 μg/ml) was added for 60 min. Serially diluted serum samples or perfusion-extraction samples (32) from intestines or lungs were added to OVA- and CTB-coated plates and then incubated for 90 min. After washing, goat anti-mouse IgA-HRP (SouthernBiotech) or goat anti-mouse IgG-HRP (Jackson ImmunoResearch Laboratories) was added and developed with o-phenylenediamine dihydrochloride before absorbance determination at 450 nm. IgG or IgA titers were defined as the sample dilution giving an OD value of 0.4 above the background value.

Splenocytes from CD11c-DTR mice given DTx i.p. 24, 48, or 72 h previously or PBS-treated controls were seeded 5 × 105 in 96-well plates. Splenocytes were pulsed in six identical wells with titrated amounts of OVA protein or peptide (323–339) for 2 h. Remaining proteins and peptides were washed away and 1 × 105 MACS-purified CD4+ OT-II T cells were added to each well. DTx was added to half of the wells to give a final concentration of 200 ng/ml. Cells were cocultured for 64 h, pulsed for 8 h with [3H]thymidine, and then incorporation into cellular DNA was measured.

Our aim was to determine the role of cDCs in activating CD4+ T cells in vivo following mucosal Ag administration. We thus took advantage of CD11c-DTR Tg mice, which allow conditional ablation of cDCs upon administration of DTx (12). However, injection of DTx leads to death of CD11c-DTR mice within a week (29) in a process mediated by nonhematopoietic cells. The use of CD11c-DTR→WT BM chimeras avoids this problem, and chimeric mice can be given multiple injections of DTx without any adverse effects (29). CD11c-DTR→WT BM chimeras (CD11c-DTR/WT) were generated and used throughout this study unless otherwise stated.

To determine the chimerism of DCs, flow cytometric analysis was performed. A small frequency of GFP-negative resident CD11chigh splenocytes was present in chimeric CD11c-DTR/WT mice (Fig. 1,A, upper left plot). Importantly, an identical population of CD11chighGFP DCs was observed in the spleen and mesenteric LN of (nonchimeric) CD11c-DTR mice (Fig. 1,A, upper middle and right plots), demonstrating that this is background staining or that the penetrance of GFP expression in cDCs of CD11c-DTR mice is not 100%. Furthermore, irradiation experiments using congenic mice confirmed the lack of remaining cDCs of recipient origin in the chimeras (data not shown). To ensure complete ablation of cDCs, the mice received two injections of DTx. This treatment depleted 85–98% of donor GFP+ cDCs from spleen, mucosal tissues, and draining lymphoid tissues (Fig. 1).

FIGURE 1.

Efficient ablation of cDCs in CD11c-DTR and CD11c-DTR/WT following DTx administration. A, Flow cytometric analysis of GFP vs CD11c expression by 7AADCD3CD19 splenocytes or cells from MLN of CD11c-DTR/WT and CD11c-DTR mice treated with or without DTx. B, Flow cytometric analysis of GFP vs CD11c expression by 7AADCD3CD19 cells from cervical LN (CLN), nasal-associated lymphoid tissue, PP, or intestinal lamina propria (LP) of CD11c-DTR/WT mice treated with or without DTx.

FIGURE 1.

Efficient ablation of cDCs in CD11c-DTR and CD11c-DTR/WT following DTx administration. A, Flow cytometric analysis of GFP vs CD11c expression by 7AADCD3CD19 splenocytes or cells from MLN of CD11c-DTR/WT and CD11c-DTR mice treated with or without DTx. B, Flow cytometric analysis of GFP vs CD11c expression by 7AADCD3CD19 cells from cervical LN (CLN), nasal-associated lymphoid tissue, PP, or intestinal lamina propria (LP) of CD11c-DTR/WT mice treated with or without DTx.

Close modal

Histochemical analysis confirmed the depletion of CD11c-expressing cells and showed retention of lymphoid follicles with no gross abnormalities in lymphoid tissue structure following DTx treatment (Fig. 2,A). Additionally, adoptive transfer of OVA-specific OT-II CD4+ T cells resulted in comparable cell recruitment and retention in DTx-treated recipients and control mice with no aberrant cell activation (Fig. 2, B and C). A reduction in lymphoid tissue cellularity (10–20%) was sometimes observed following DTx treatment, but this did not change the frequency of total CD4+ T cells (Fig. 2 C and data not shown). Thus, the DTx depletion of cDCs was very effective but not absolute in DTx-treated chimeras. It also had no detectable effect on baseline recruitment, retention, or activation of adoptively transferred CD4+ T cells.

FIGURE 2.

Ablation of cDCs in CD11c-DTR/WT retains lymphoid follicles and does not effect baseline recruitment, retention, or activation of adoptively transferred CD4+ T cells. A, Histological analysis of tissue sections from mice given DTx (+DTx) or not (−DTx) stained with anti-CD11c (green) and B220 (red) (×20 magnification). B and C, Flow cytometric analysis of CD45.1 and CD4 for detection of adoptively transferred OT-II cells in CLN cells from mice given DTx (+DTx) or not (−DTx). OT-II cells were transferred at the same time as the second DTx treatment and the tissues were taken 18 (B) or 90 h (C) later. B and C, Left panel, The number in the dot plot indicates the frequency of gated cells among total viable cells. C, Right panels, Expression of CD45.1 and CD69 by CD4+ cells gated as indicated by the arrow. The numbers in the dot plot indicate the frequency of CD69 expressing cells among the transferred CD45.1+CD4+ OT-II T cells.

FIGURE 2.

Ablation of cDCs in CD11c-DTR/WT retains lymphoid follicles and does not effect baseline recruitment, retention, or activation of adoptively transferred CD4+ T cells. A, Histological analysis of tissue sections from mice given DTx (+DTx) or not (−DTx) stained with anti-CD11c (green) and B220 (red) (×20 magnification). B and C, Flow cytometric analysis of CD45.1 and CD4 for detection of adoptively transferred OT-II cells in CLN cells from mice given DTx (+DTx) or not (−DTx). OT-II cells were transferred at the same time as the second DTx treatment and the tissues were taken 18 (B) or 90 h (C) later. B and C, Left panel, The number in the dot plot indicates the frequency of gated cells among total viable cells. C, Right panels, Expression of CD45.1 and CD69 by CD4+ cells gated as indicated by the arrow. The numbers in the dot plot indicate the frequency of CD69 expressing cells among the transferred CD45.1+CD4+ OT-II T cells.

Close modal

CFSE-labeled OT-II T cells were adoptively transferred into CD11c-DTR/WT mice 48 h after receiving DTx or PBS. DTx-treated mice received a second injection of DTx shortly after the transfer. Eighteen hours later both groups of mice were fed different doses of OVA, and cells from Peyer’s patches (PP) and MLNs were analyzed for proliferation of the adoptively transferred T cells 3 days later (Fig. 3, A and B). Little, if any, OT-II T cell expansion was detected in either organ of mice receiving DTx before feeding OVA, while significantly more proliferation was readily observed in mice not receiving DTx (Fig. 3, C and D).

FIGURE 3.

cDCs are required for activation of CD4+ T cells following OVA feeding. Flow cytometric analysis of 7AADVα2+CD4+ cells from PP (A) or MLN (B) of CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with DTx or not and fed PBS (control) or OVA as indicated. Cells were analyzed 72 h after OVA feeding. The histograms represent 7AADVα2+CD4+ cells gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division taking into account the expansion of divided cells (undivided cells are indicated with a gate in the histogram). Plots are of three pooled mice and are representative of at least three experiments. The statistical analysis (C and D) was performed using Student’s t test where *, p < 0.05 and **, p < 0.01. The bars show the SEM.

FIGURE 3.

cDCs are required for activation of CD4+ T cells following OVA feeding. Flow cytometric analysis of 7AADVα2+CD4+ cells from PP (A) or MLN (B) of CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with DTx or not and fed PBS (control) or OVA as indicated. Cells were analyzed 72 h after OVA feeding. The histograms represent 7AADVα2+CD4+ cells gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division taking into account the expansion of divided cells (undivided cells are indicated with a gate in the histogram). Plots are of three pooled mice and are representative of at least three experiments. The statistical analysis (C and D) was performed using Student’s t test where *, p < 0.05 and **, p < 0.01. The bars show the SEM.

Close modal

To determine the impact of a mucosal adjuvant on the expansion of transferred T cells, CT was given together with OVA. Again, a significant reduction in the proliferation of labeled CD4+ T cells was found in the draining LNs after oral (Fig. 4, A and C) or nasal (Fig. 4, B and D) administration to DTx-treated mice compared with controls not receiving DTx treatment. Expression of CD69 by the transferred OT-II T cells was detected among undivided OT-II cells in both DTx-treated mice and untreated controls following oral immunization with OVA and CT (Fig. 4,E). In the absence of DTx treatment, CD69 expression was down-regulated upon cell division. The same CD69 expression pattern was detected on OT-II T cells from cervical LN 5 days after nasal immunization with OVA (Fig. 4,F). Restimulation of OT-II T cells from these animals resulted in Ag-specific proliferation irrespective of DTx treatment before immunization (data not shown). This suggests that the observed lack of OT-II T cell division following DTx treatment (Figs. 3 and 4) was not due to a delayed response in the transferred cells and that these cells were not anergic to subsequent stimulation. These results hence show that cDCs are required for priming of CD4+ T cells following nasal and oral administration of protein both in the absence and presence of CT.

FIGURE 4.

cDCs are essential for proliferation of CD4+ T cells following mucosal administration of OVA and CT. Flow cytometric analysis of 7AADVα2+CD4+ cells from (A and E) MLN or (B and F) CLN of CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with or without DTx. Mice were then given OVA plus CT (A and E) orally, (B) nasally, or (F) OVA nasally. Cells were analyzed (A, B, and E) 72 h or (F) 96 h after OVA administration. A and B, Histograms represent 7AADVα2+CD4+ cells gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division taking in to account the expansion of divided cells (undivided cells are indicated with a gate in the histogram). E and F, Expression of CD69 and CFSE by CD45.1+CD4+ OT-II T cells. The numbers in the dot plot indicate the frequency of CD69-expressing cells among the transferred CD45.1+CD4+ OT-II T cells. Plots are of three pooled (A, E, and F) or individual mice (B). Data are representative of three separate experiments. Statistical analysis (C and D) was performed using Student’s t test where **, p < 0.01 and ***, p < 0.001. The bars show the SEM.

FIGURE 4.

cDCs are essential for proliferation of CD4+ T cells following mucosal administration of OVA and CT. Flow cytometric analysis of 7AADVα2+CD4+ cells from (A and E) MLN or (B and F) CLN of CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with or without DTx. Mice were then given OVA plus CT (A and E) orally, (B) nasally, or (F) OVA nasally. Cells were analyzed (A, B, and E) 72 h or (F) 96 h after OVA administration. A and B, Histograms represent 7AADVα2+CD4+ cells gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division taking in to account the expansion of divided cells (undivided cells are indicated with a gate in the histogram). E and F, Expression of CD69 and CFSE by CD45.1+CD4+ OT-II T cells. The numbers in the dot plot indicate the frequency of CD69-expressing cells among the transferred CD45.1+CD4+ OT-II T cells. Plots are of three pooled (A, E, and F) or individual mice (B). Data are representative of three separate experiments. Statistical analysis (C and D) was performed using Student’s t test where **, p < 0.01 and ***, p < 0.001. The bars show the SEM.

Close modal

To investigate the role of cDCs in generating Ag-specific Abs after oral and nasal immunization, CD11c-DTR/WT mice were fed OVA and CT. Before feeding, half of the mice were depleted of cDCs, as described above, and Ag-specific Ab titers were measured. The IgG titers in serum (10–12 days postadministration) and IgA in intestinal tissues (3 wk postadministration) were determined by ELISA. Anti-OVA and anti-CTB IgG titers in serum, following oral (Fig. 5, A and B) and nasal immunization (Fig. 5, D and E), were abrogated in CD11c-DTR/WT given DTx compared with controls. Although no significant intestinal anti-OVA IgA could be detected regardless of the dose of OVA given orally and irrespective of cDC-depletion (data not shown), anti-CTB IgA was clearly detected and completely lost in CD11c-DTR/WT mice given DTx (Fig. 5 C). These results show that cDCs are required for the generation of OVA- and CTB-specific serum IgG after mucosal administration, and of CTB-specific intestinal IgA after feeding mice OVA with CT.

FIGURE 5.

cDCs are essential for efficient generation of Ag-specific Abs following mucosal administration of OVA and CT. The indicated amounts of OVA and CT were administered (A–C) orally or (D and E) nasally to CD11c-DTR/WT mice. The graphs show log10 titers of anti-OVA- (A and D), anti-CTB- (B and E) specific serum IgG or anti-CTB-specific intestinal IgA (C) in CD11c-DTR/WT mice treated with DTx (open bars) or not (filled bars). DTx was administered 72 and 18 h before the immunizations. Serum samples were collected 10–12 days postimmunization and intestines were collected 3 weeks postimmunization. The IgG or IgA titer was defined as the sample dilution giving an OD value of 0.4 above the background value in ELISA. Unimmunized controls showed a titer <10 for IgA and IgG. Error bars show the SEM and statistical analysis was performed using Student’s t test where *, p < 0.05, **, p < 0.01, and ***, p < 0.001.

FIGURE 5.

cDCs are essential for efficient generation of Ag-specific Abs following mucosal administration of OVA and CT. The indicated amounts of OVA and CT were administered (A–C) orally or (D and E) nasally to CD11c-DTR/WT mice. The graphs show log10 titers of anti-OVA- (A and D), anti-CTB- (B and E) specific serum IgG or anti-CTB-specific intestinal IgA (C) in CD11c-DTR/WT mice treated with DTx (open bars) or not (filled bars). DTx was administered 72 and 18 h before the immunizations. Serum samples were collected 10–12 days postimmunization and intestines were collected 3 weeks postimmunization. The IgG or IgA titer was defined as the sample dilution giving an OD value of 0.4 above the background value in ELISA. Unimmunized controls showed a titer <10 for IgA and IgG. Error bars show the SEM and statistical analysis was performed using Student’s t test where *, p < 0.05, **, p < 0.01, and ***, p < 0.001.

Close modal

A very high dose of protein Ag is sometimes required to initiate immune responses at mucosal surfaces. The abundance of Ag has also been suggested to affect activation of the adaptive immune response, possibly because additional APC populations access the Ag in peripheral or lymphoid tissues (17). To test this, we mucosally immunized cDC-depleted mice with a very high dose of OVA, 30-fold more than that required to detect proliferation of Ag-specific CD4+ T cells in WT mice after mucosal immunization (Fig. 6, A and B). OVA administration at 300 mg orally or 3 mg nasally in the presence of CT resulted in extensive proliferation of adoptively transferred OT-II T cells in control mice not given DTx. However, treatment with DTx had no significant effect on the frequency of cells that entered division (Fig. 6, A, B, D, and E). To determine whether this was due to the route of immunization, a very high dose of OVA (0.3 mg) was injected i.v. (Fig. 6,C) and titrated amounts of OVA were given i.p. (Fig. 6,G) and then the proliferation of the transferred cells in the spleen was measured. As observed with mucosal immunization, increasing the dose of OVA to a very high dose resulted in similar proliferation of the transferred cells in cDC-depleted mice and in animals not treated with DTx (Fig. 6, F, G, and H). This is in contrast to cDC-dependent proliferation where lower doses of Ag are used (Fig. 6 H). These results show that immunizing mice depleted of cDCs with a very high dose of Ag results in proliferation of CD4+ T cells.

FIGURE 6.

Priming of CD4+ T cells after mucosal and parenteral administration of a very high dose of OVA. Flow cytometric analysis of 7AADVα2+CD4+ cells from (A) MLN, (B) cervical LN, or (C and G) spleen of CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with or without DTx and given OVA plus CT orally (A), OVA plus CT nasally (B), OVA i.v. (C), or OVA i.p (G). Histograms represent 7AADVα2+CD4+ cells, gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division taking into account the expansion of divided cells (undivided cells are indicated with a gate in the histogram). Plots are of (A) three pooled or (B, C, and G) individual mice and the data are representative of three or more separate experiments. Statistical analysis was performed (D–F) using Student’s t test or (H) one-way-ANOVA where **, p < 0.01 and n.s, not significant. The bars show the SEM.

FIGURE 6.

Priming of CD4+ T cells after mucosal and parenteral administration of a very high dose of OVA. Flow cytometric analysis of 7AADVα2+CD4+ cells from (A) MLN, (B) cervical LN, or (C and G) spleen of CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with or without DTx and given OVA plus CT orally (A), OVA plus CT nasally (B), OVA i.v. (C), or OVA i.p (G). Histograms represent 7AADVα2+CD4+ cells, gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division taking into account the expansion of divided cells (undivided cells are indicated with a gate in the histogram). Plots are of (A) three pooled or (B, C, and G) individual mice and the data are representative of three or more separate experiments. Statistical analysis was performed (D–F) using Student’s t test or (H) one-way-ANOVA where **, p < 0.01 and n.s, not significant. The bars show the SEM.

Close modal

A possible explanation for CD4+ T cell activation in DTx-treated chimeric mice given a very high dose of Ag is that other APCs accumulate in tissues following DTx-induced cell death of DTR+ DCs. The cellular composition of lymphoid organs from CD11c-DTR/WT mice treated twice with DTx was therefore analyzed. After giving one or two injections of DTx, CD11bhighCD11c−/low cells accumulated in lymphoid tissues (Fig. 7,A). Multicolor flow cytometry of the splenic CD11bhighCD11c−/low cells revealed a decrease in frequency of Ly6Chigh6G−/low and an increase in Ly6Chigh6Ghigh cells, representing monocytes and neutrophils, respectively (33) (Fig. 7 B). The increase of Ly6Chigh6Ghigh cells was detected very rapidly and at all time points when cDCs were efficiently ablated. In contrast, no increase of these cells was observed in DTx-treated WT mice (data not shown). Recruitment of neutrophils is thus an effect of the DTx-induced ablation in CD11c-DTR/WT mice.

FIGURE 7.

CD11bhighCD11c−/low cells with very poor CD4+ T cell activation capacity accumulate in DTx-treated CD11c-DTR mice. A, Flow cytometric analysis of CD11b vs CD11c expression by 7AADCD3CD19 splenocytes or cells from MLN of CD11c-DTR/WT mice treated with or without DTx. B, Flow cytometric analysis of Ly6C/Ly6G expression by 7AADCD3CD19CD11bhighCD11c−/low splenocytes. C, In vitro proliferation of OT-II cells cocultured with CD11c-DTR splenocytes pulsed for 2 h with titrated amounts of OVA protein (left) or peptide (right) in the presence (•) or absence (○) of DTx. After 72 h the cocultures were pulsed with [3H]thymidine and incorporation was measured after 6 h. D, In vitro proliferation of OT-II cells cocultured with splenocytes from CD11c-DTR mice given PBS (○) or DTx i.p. 24 (•), 48 (♦), or 72 h (♦) earlier and subsequently pulsed for 2 h with titrated amounts of OVA protein (left) or peptide (right). After 72 h the cocultures were pulsed with [3H]thymidine and incorporation was measured after 6 h. One representative experiment out of two is shown.

FIGURE 7.

CD11bhighCD11c−/low cells with very poor CD4+ T cell activation capacity accumulate in DTx-treated CD11c-DTR mice. A, Flow cytometric analysis of CD11b vs CD11c expression by 7AADCD3CD19 splenocytes or cells from MLN of CD11c-DTR/WT mice treated with or without DTx. B, Flow cytometric analysis of Ly6C/Ly6G expression by 7AADCD3CD19CD11bhighCD11c−/low splenocytes. C, In vitro proliferation of OT-II cells cocultured with CD11c-DTR splenocytes pulsed for 2 h with titrated amounts of OVA protein (left) or peptide (right) in the presence (•) or absence (○) of DTx. After 72 h the cocultures were pulsed with [3H]thymidine and incorporation was measured after 6 h. D, In vitro proliferation of OT-II cells cocultured with splenocytes from CD11c-DTR mice given PBS (○) or DTx i.p. 24 (•), 48 (♦), or 72 h (♦) earlier and subsequently pulsed for 2 h with titrated amounts of OVA protein (left) or peptide (right). After 72 h the cocultures were pulsed with [3H]thymidine and incorporation was measured after 6 h. One representative experiment out of two is shown.

Close modal

The depleted splenic cDC population in DTx-treated mice is restored 6 days following treatment, during which time the observed neutrophil accumulation occurs. Therefore, we wondered whether cells (precursor or inflammatory cells) recruited to the tissue during this period of cDC absence could contribute to the activation of CD4+ T cells. To address this, depletion of CD11chigh cells from the splenocytes was conducted in vitro (12) so that no recruitment of cells was possible. Initial experiments showed that 0.2 μg/ml DTx efficiently depleted CD11c+ cells from splenocyte cultures (0.4% compared with 23% CD11c+ cells among nonlymphocytes with or without DTx, respectively). There was no increase in CD11bhighCD11c−/low cells (data not shown). Addition of DTx, followed by a pulse with OVA peptide or protein, abrogated the proliferative response of cocultured OT-II cells unless very high peptide or protein concentrations were used (Fig. 7 C).

Having shown that cDCs were required to induce proliferation of CD4+ T cells in vitro, we next assessed whether the cells recruited to the spleen after DTx treatment could activate CD4+ T cells. CD11c-DTR mice were therefore given DTx before sacrifice. Splenocytes from CD11c-DTR mice given DTx 72, 48, or 24 h earlier were pulsed with OVA protein or peptide and cultured with OT-II cells. While proliferation was observed in PBS-treated controls, little proliferation was detected when splenocytes from CD11c-DTR mice given DTx in vivo were used (Fig. 7 D). Taken together, these results show that CD11chigh cells are efficiently ablated by DTx treatment in lymphoid tissues of CD11c-DTR/WT mice. Concomitantly, CD11bhighCD11c−/low neutrophils are recruited to tissues, but these cells have a very poor capacity to present peptides to CD4+ T cells in vitro.

Having shown that the recruited CD11bhighCD11c−/low cells are not capable of inducing proliferation of CD4+ T cells after Ag exposure, we next addressed if other APCs could be responsible for the observed activation of OVA-specific CD4+ T cells after administration of a high dose of OVA to cDC-ablated mice. To determine whether B cells were capable of priming T cells in the absence of cDCs, we crossed the DTR Tg mice with B cell-deficient μMT mice, generating DTR Tg:μMt mice. Similar to μMT mice, these mice display a multitude of architectural defects in the spleen, including absence of follicular DCs, marginal zone macrophages, and metallophilic macrophages (34), as well as differences in DC function (35). We therefore made mixed BM chimeras (CD11c-DTR/MHC-IIB−/−) in which the B cell compartment is normal in number but MHC-II is deficient, as described by Crawford et al. (30). Seven weeks after engraftment, ∼80% of splenic cDCs (CD11c+B220 cells) from CD11c-DTR/MHC-IIB−/− mice were MHC-II+ and expressed DTR (i.e., were GFP+) (Fig. 8,A). Importantly, the remaining DCs (GFP) and all B cells were MHC-II. These mice were then adoptively transferred with CFSE-labeled OT-II cells and 1 day later given OVA at a high dose nasally or at titrated amounts of OVA i.p. Although expansion of the transferred T cells was cDC-dependent when lower doses of Ag were administered i.p. (Fig. 8,C, left panel), cDCs were not essential for OT-II T cell division when a high dose of Ag was given nasally or i.p. (Fig. 8, B and C, middle panel). Because the DTx-treated mice had a MHC-II-deficient B cells, the observed T cell proliferation must have been initiated by cells other than B cells.

FIGURE 8.

Expansion of CD4+ T cells following administration of a very high dose of OVA to mice with MHC-II-deficient B cells and ablated cDCs. A, Flow cytometric analysis of MHC-II and GFP expression by CD11c+B220 cells (cDCs) and CD11cB220+ cells (B cells) from CD11c-DTR/MHC-IIB−/− mixed BM chimeric mice at the time of immunization with OVA. B and C, Flow cytometric analysis of 7AADVα2+CD4+ cells from cervical LN or spleen taken from CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with or without DTx and given OVA or PBS nasally (B) or i.p. (C). Histograms represent 7AADVα2+CD4+ cells, gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division (determined from two separate experiments with a total of four mice per group) taking in to account the expansion of divided cells (undivided cells are indicated with a gate in the histogram).

FIGURE 8.

Expansion of CD4+ T cells following administration of a very high dose of OVA to mice with MHC-II-deficient B cells and ablated cDCs. A, Flow cytometric analysis of MHC-II and GFP expression by CD11c+B220 cells (cDCs) and CD11cB220+ cells (B cells) from CD11c-DTR/MHC-IIB−/− mixed BM chimeric mice at the time of immunization with OVA. B and C, Flow cytometric analysis of 7AADVα2+CD4+ cells from cervical LN or spleen taken from CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with or without DTx and given OVA or PBS nasally (B) or i.p. (C). Histograms represent 7AADVα2+CD4+ cells, gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division (determined from two separate experiments with a total of four mice per group) taking in to account the expansion of divided cells (undivided cells are indicated with a gate in the histogram).

Close modal

Our data so far show that neither B cells nor recruited myeloid cells are responsible for CD4+ T cell proliferation in DTx-treated mice given a very high dose of OVA. We thus determined if pDCs contribute to this CD4+ T cell expansion by administering the pDC-depleting Ab 120.G8 (31) to DTx-treated CD11c-DTR/WT mice before administering OVA (Fig. 9). This combined treatment resulted in a complete loss of cDCs (CD11chighmPDCA-1) and pDCs (CD11cintmPDCA-1+) in the spleen (Fig. 9,A). As shown above, cDC depletion had a relatively minor effect on the proliferation of transferred OT-II CD4+ T cells in CD11c-DTR/WT mice given 3 mg of OVA nasally (Fig. 9,B) or i.p. (Fig. 9 C). Depleting both pDCs and cDCs had little additional effect, and significant expansion could still be observed in the pDC/cDC-depleted mice.

FIGURE 9.

Activation of CD4+ T cells and induction of serum Ab responses in mice depleted of both cDCs and pDCs after administration of a very high dose of Ag. A, Flow cytometric analysis of mPDCA-1 and CD11c expression by 7AADCD3CD19 splenocytes from CD11c-DTR/WT treated with PBS, DTx, or DTx plus 120.G8. The mAb 120.G8 was given for 3 consecutive days before immunization. The numbers indicate the frequency of cells among gated 7AADCD3CD19 cells. B, Flow cytometric analysis of 7AADVα2+CD4+ cells from the cervical LN or spleen from CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with DTx, DTx plus 120.G8, or untreated and then given OVA nasally (B) or i.p. (C). Analysis was performed 72 h after OVA administration. Histograms represent 7AADVα2+CD4+ cells, gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division (determined from three separate experiments) taking into account the expansion of divided cells (undivided cells are indicated with a gate in the histogram). Plots are of individual mice and are representative of three separate experiments. D, Log10 titers of anti-OVA-specific IgG in serum from CD11c-DTR/WT mice treated with DTx, DTx plus 120.G8, or untreated before nasal immunization with OVA plus CT or OVA plus CpG. Ten to 14 days later these mice and unimmunized controls (filled gray bars) received OVA i.p. Serum was collected 1 wk later and analyzed for OVA-specific IgG titers by ELISA. Titers were defined as the sample dilution giving an OD value of 0.4 above the background value. Error bars show the SEM and statistical analysis was performed using Student’s t test where *, p < 0.05 and n.s., not significant.

FIGURE 9.

Activation of CD4+ T cells and induction of serum Ab responses in mice depleted of both cDCs and pDCs after administration of a very high dose of Ag. A, Flow cytometric analysis of mPDCA-1 and CD11c expression by 7AADCD3CD19 splenocytes from CD11c-DTR/WT treated with PBS, DTx, or DTx plus 120.G8. The mAb 120.G8 was given for 3 consecutive days before immunization. The numbers indicate the frequency of cells among gated 7AADCD3CD19 cells. B, Flow cytometric analysis of 7AADVα2+CD4+ cells from the cervical LN or spleen from CD11c-DTR/WT mice adoptively transferred with CFSE-labeled OT-II cells treated with DTx, DTx plus 120.G8, or untreated and then given OVA nasally (B) or i.p. (C). Analysis was performed 72 h after OVA administration. Histograms represent 7AADVα2+CD4+ cells, gated as shown. The number in the histograms indicates the frequency of transferred T cells that have entered division (determined from three separate experiments) taking into account the expansion of divided cells (undivided cells are indicated with a gate in the histogram). Plots are of individual mice and are representative of three separate experiments. D, Log10 titers of anti-OVA-specific IgG in serum from CD11c-DTR/WT mice treated with DTx, DTx plus 120.G8, or untreated before nasal immunization with OVA plus CT or OVA plus CpG. Ten to 14 days later these mice and unimmunized controls (filled gray bars) received OVA i.p. Serum was collected 1 wk later and analyzed for OVA-specific IgG titers by ELISA. Titers were defined as the sample dilution giving an OD value of 0.4 above the background value. Error bars show the SEM and statistical analysis was performed using Student’s t test where *, p < 0.05 and n.s., not significant.

Close modal

We next measured the OVA-specific IgG response in CD11c-DTR/WT mice primed with OVA plus CT nasally in the presence or absence of DCs (cDCs or cDCs/pDCs) and boosted with OVA i.p. with neither DTx nor 120.G8 treatment (Fig. 9,D). This immunization regime led to a high anti-OVA IgG titer in the serum of CD11c-DTR/WT mice that was significantly reduced by DTx treatment. However, there was no significant effect of additional pDC depletion. To confirm that the pDC depletion was functional, depleted mice were immunized with OVA and CpG, as pDCs are essential for CpG-induced activation of cDCs (36). Only the combined cDC and pDC ablation led to a significant reduction in serum anti-OVA IgG titers upon OVA plus CpG immunization (Fig. 9 D). These results show that a high dose of OVA results in CD4+ T cell proliferation even in mice depleted of both cDCs and pDCs. Finally, depletion of pDCs in addition to cDCs at the time of priming with OVA and CT via the nasal route did not further reduce the anti-OVA Ab response compared with cDC depletion alone.

To generate vaccines that protect mucosal surfaces, a better understanding of the cells required in vivo for activation of the adaptive immune response following mucosal immunization is required. To determine the role of DCs in this regard, we analyzed the activation of CD4+ T cells following oral and nasal immunization of mice ablated of cDCs in vivo. Our results show that cDCs are required for the activation of naive OVA-specific CD4+ T cells in vivo and for the generation of mucosal and systemic Ag-specific Abs after both oral and nasal administration of OVA and CT, unless a very high dose of OVA is used.

Our results with high doses of Ag show that an APC population present in, or recruited to, the tissue of DTx-treated CD11c-DTR mice possesses the capacity to present peptides, but only when Ag is abundant. DC depletion resulted in an immediate recruitment of cells to lymphoid tissues, most of which express a high level of CD11b, Ly6C, and Ly6G, but no, or very low level of, CD11c. These cells are likely neutrophils, but the recruited population could also contain other cells potentially capable of differentiating into APCs (33). However, this appears unlikely, as splenocytes from CD11c-DTR mice treated with DTx 3 days earlier (a time frame that should allow for differentiation) could not activate CD4+ T cells. This result is identical to that observed with splenocytes following DC depletion in vitro where no recruitment was possible.

The appearance of neutrophils in tissues of CD11c-DTR mice after DTx-induced ablation of DCs has not previously been reported, even though this Tg mouse has been used in several studies (12, 13, 14, 15, 16, 17, 19, 24, 29, 37, 38, 39). Although our results show that the recruited cells do not have the capacity to efficiently present peptides on MHC-II ex vivo, these results emphasize the need for a cautious approach when using DTx-treated CD11c-DTR mice to study the role of inflammatory cells and secretion of soluble factors in the absence of DCs.

Furthermore, we could not detect a significant role for B cells in the activation of T cells following immunization with a high dose of Ag in the absence of cDCs. Our results therefore support a previous report showing that CD4+ T cell activation in LNs could still be observed in a B cell-deficient animal where cDCs had been ablated (16). Importantly, our results also extend this observation, as we used mice in which B cells are present but are MHC-II deficient. Using this system, we ensured that the observed MHC-II presentation was not due to changes in LN cellularity or altered structure of peripheral lymphoid tissues, which is a caveat when using B cell-deficient mice (34, 35). Our results could seem contradictory to a study using mice with MHC-II−/− B cells, which showed that B cells provide extra Ag presentation capacity above that provided by DCs (30). This could be because different doses of Ag were used in that study compared with the doses used by us, and/or that Crawford et al. performed all immunizations in the presence of alum (30). It remains possible, however, that B cells may collaborate with cDCs during CD4+ T cell activation.

Our study shows that immunization of mice depleted of both cDCs and pDCs with a very high dose of OVA still resulted in activation of CD4+ T cells, comparable to that seen when only cDCs were depleted. It has been reported that OVA-specific CD4+ T cells can be activated by pDC-specific Abs carrying OVA, and also in LNs of cDC-depleted mice following s.c immunization with OVA (16). Sapoznikov et al. (16) found no significant depletion of pDCs after DTx treatment, while we consistently observe a partial depletion, which has also been reported by others (13, 17). It is therefore possible that this partial depletion is sufficient to deplete mucosal pDCs that have a capacity to activate CD4+ T cells. However, only after combining DTx treatment with the pDC-depleting Ab could we significantly inhibit the generation of OVA-specific Abs in mice that had been given CpG and OVA mucosally. This shows that pDCs remaining after DTx treatment are indeed fully functional, since CpG-induced immune activation is dependent on IL-15-mediated cross-talk between pDCs and cDCs (36). Although Sapoznikov et al. (16) showed that specific targeting of Ag to pDCs resulted in activation of CD4+ T cells, they did not address whether removal of pDCs abrogated the observed activation. As emphasized by our study, the dose of Ag administered will also have a significant impact. This makes comparison between different routes of immunization difficult unless the Ag is titrated. Finally, it is also possible that pDCs in skin are functionally different from those in the spleen and mucosal tissues. For example, we have not been able to detect pDCs in afferent lymph from mucosal tissues (40), while this has been reported in lymph draining the skin (41).

CT holotoxin is the most potent oral adjuvant, but the mechanism behind its adjuvanticity is not fully known. Ab responses to both OVA and CT following mucosal coadministration have been shown to be completely dependent on CD4+ T cell help (42). The effect of CT could therefore be to make DCs more potent activators of CD4+ T cells. Alternatively, CT could act on B cells, rendering them capable of activating naive T cells. Our results show that coadministering CT and OVA mucosally did not overcome the essential role of cDCs for activation of CD4+ T cells and induction of intestinal IgA and serum IgG. Detection of anti-OVA IgA responses in mucosal tissues required that the mice be immunized a second time with OVA and CT. However, ablation of cDCs before both immunizations creates a problem, as plasma cells in the spleen express CD11c and thereby become sensitive to DTx treatment (24). Indeed, in preliminary experiments we have found that the number of CT- and OVA-specific IgA-secreting cells in the intestine are reduced after DTx treatment (OVA, 120 ± 50, CT, 3515 ± 1889 per million cells (−DTx) compared with OVA, 18 ± 13, CT, 275 ± 129 per million cells (+DTx)). In these experiments DTx was only given 1 wk after the second immunization with OVA and CT, making it unlikely that the effect of DTx was on DCs.

DTx treatment of CD11c-DTR mice has also been reported to ablate marginal zone macrophages and their sinusoidal counterparts in LN when using the same dose of DT used by us in this study (43). This makes it unlikely that these cells contribute to activation of CD4+ T cells following immunization with a very high dose of Ag. Additionally, removal of macrophages using chlodronate-containing liposomes increases, rather than reduces, the amount of Ab-secreting cells following immunization with a T cell-dependent Ag (44). The mucosal epithelium has also been suggested to present Ags on MHC-II (45). However, in preliminary experiments using chimeric mice, we have been unable to find a role for nonhematopoietic cells (including epithelial cells) for CD4+ T cell activation in the absence of cDCs (data not shown).

It is still possible that a hematopoietic cell type that we have been unable to define could be responsible for the activation of CD4+ T cells when a very high dose of Ag is administered. During the revision of this manuscript it has been shown that basophils purified from blood can, when loaded with peptides, activate CD4+ T cells in vitro (46). However, in order for significant numbers of basophils to enter lymphoid tissues a parasitic stimulus is required (46). Another possibility is that, although only few cDCs remain after the DTx-induced ablation, these cDCs are sufficient to activate CD4+ T cells when the Ag is abundant. The functional status of CD4+ T cells that proliferate in cDC-depleted mice following immunizations with very high doses is currently being investigated and preliminary results suggest that these cells are not anergized.

Our study therefore shows the crucial and irreplaceable role of cDCs for nasal and oral immunizations unless a very high dose of protein is used. Whether mucosally applied Ags are directly taken up by DCs that are in contact with the lumen, or DCs acquire the Ag after transport over the epithelium or following transport in lymph (cell associated or not), or if all of these pathways operate is not known. Additionally, if the same rules apply for Ag and different adjuvants after mucosal coadministration also remain to be addressed. Answering these questions will be crucial to improving the efficacy of mucosal vaccinations. However, the essential role of cDCs shown in this study suggests that targeting of the vaccine to DCs would be a useful way to ensure successful mucosal vaccination.

We thank medical physicist Elin Haglund and the Department of Radiation Physics, Sahlgrenska University Hospital, for excellent assistance in the generation of chimeric mice.

The authors have no financial conflicts of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This study was supported by grants from the Swedish Research Council, Swedish Foundation for Strategic Research through its support of the Mucosal Immunobiology and Vaccine Center, Marianne and Marcus Wallenberg Foundation, Jeansson Foundation, Åke Wiberg Foundation, Clas Grochinsky Foundation, Magnus Bergvall Foundation and the Royal Arts and Society of Arts and Science in Göteborg.

3

Abbreviations used in this paper: DC, dendritic cell; DTx, diphtheria toxin; cDC, conventional dendritic cell; DTR, diphtheria toxin receptor; VSV, vesicular stomatatis virus; LN, lymph node; pDC, plasmacytoid dendritic cell; CT, cholera toxin; Tg, transgenic; BM, bone marrow; WT, wild type; 7AAD, 7-aminoactinomycin D; CTB, cholera toxin subunit B; MLN, mesenteric LN; PP, Peyer’s patches.

1
Holmgren, J., C. Czerkinsky.
2005
. Mucosal immunity and vaccines.
Nat. Med.
11
:
S45
-S53.
2
Anjuere, F., C. Luci, M. Lebens, D. Rousseau, C. Hervouet, G. Milon, J. Holmgren, C. Ardavin, C. Czerkinsky.
2004
. In vivo adjuvant-induced mobilization and maturation of gut dendritic cells after oral administration of cholera toxin.
J. Immunol.
173
:
5103
-5111.
3
Chirdo, F. G., O. R. Millington, H. Beacock-Sharp, A. M. Mowat.
2005
. Immunomodulatory dendritic cells in intestinal lamina propria.
Eur. J. Immunol.
35
:
1831
-1840.
4
Chung, Y., J. H. Chang, M. N. Kweon, P. D. Rennert, C. Y. Kang.
2005
. CD8α-11b+ dendritic cells but not CD8α+ dendritic cells mediate cross-tolerance toward intestinal antigens.
Blood
106
:
201
-206.
5
del Rio, M. L., J. I. Rodriguez-Barbosa, E. Kremmer, R. Forster.
2007
. CD103 and CD103+ bronchial lymph node dendritic cells are specialized in presenting and cross-presenting innocuous antigen to CD4+ and CD8+ T cells.
J. Immunol.
178
:
6861
-6866.
6
Jaensson, E., H. Uronen-Hansson, O. Pabst, B. Eksteen, J. Tian, J. L. Coombes, P. L. Berg, T. Davidsson, F. Powrie, B. Johansson-Lindbom, W. W. Agace.
2008
. Small intestinal CD103+ dendritic cells display unique functional properties that are conserved between mice and humans.
J. Exp. Med.
205
:
2139
-2149.
7
Porgador, A., H. F. Staats, Y. Itoh, B. L. Kelsall.
1998
. Intranasal immunization with cytotoxic T-lymphocyte epitope peptide and mucosal adjuvant cholera toxin: selective augmentation of peptide-presenting dendritic cells in nasal mucosa-associated lymphoid tissue.
Infect. Immun.
66
:
5876
-5881.
8
Sung, S. S., S. M. Fu, C. E. Rose, Jr, F. Gaskin, S. T. Ju, S. R. Beaty.
2006
. A major lung CD103 (αE)-β7 integrin-positive epithelial dendritic cell population expressing Langerin and tight junction proteins.
J. Immunol.
176
:
2161
-2172.
9
Vermaelen, K. Y., I. Carro-Muino, B. N. Lambrecht, R. A. Pauwels.
2001
. Specific migratory dendritic cells rapidly transport antigen from the airways to the thoracic lymph nodes.
J. Exp. Med.
193
:
51
-60.
10
Wikstrom, M. E., E. Batanero, M. Smith, J. A. Thomas, C. von Garnier, P. G. Holt, P. A. Stumbles.
2006
. Influence of mucosal adjuvants on antigen passage and CD4+ T cell activation during the primary response to airborne allergen.
J. Immunol.
177
:
913
-924.
11
Lambrecht, B. N., M. De Veerman, A. J. Coyle, J. C. Gutierrez-Ramos, K. Thielemans, R. A. Pauwels.
2000
. Myeloid dendritic cells induce Th2 responses to inhaled antigen, leading to eosinophilic airway inflammation.
J. Clin. Invest.
106
:
551
-559.
12
Jung, S., D. Unutmaz, P. Wong, G. Sano, K. De los Santos, T. Sparwasser, S. Wu, S. Vuthoori, K. Ko, F. Zavala, 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.
13
Ciavarra, R. P., A. Stephens, S. Nagy, M. Sekellick, C. Steel.
2006
. Evaluation of immunological paradigms in a virus model: are dendritic cells critical for antiviral immunity and viral clearance?.
J. Immunol.
177
:
492
-500.
14
Kassim, S. H., N. K. Rajasagi, X. Zhao, R. Chervenak, S. R. Jennings.
2006
. In vivo ablation of CD11c-positive dendritic cells increases susceptibility to herpes simplex virus type 1 infection and diminishes NK and T-cell responses.
J. Virol.
80
:
3985
-3993.
15
Probst, H. C., M. van den Broek.
2005
. Priming of CTLs by lymphocytic choriomeningitis virus depends on dendritic cells.
J. Immunol.
174
:
3920
-3924.
16
Sapoznikov, A., J. A. Fischer, T. Zaft, R. Krauthgamer, A. Dzionek, S. Jung.
2007
. Organ-dependent in vivo priming of naive CD4+, but not CD8+, T cells by plasmacytoid dendritic cells.
J. Exp. Med.
204
:
1923
-1933.
17
Scandella, E., K. Fink, T. Junt, B. M. Senn, E. Lattmann, R. Forster, H. Hengartner, B. Ludewig.
2007
. Dendritic cell-independent B cell activation during acute virus infection: a role for early CCR7-driven B-T helper cell collaboration.
J. Immunol.
178
:
1468
-1476.
18
Tian, T., J. Woodworth, M. Skold, S. M. Behar.
2005
. In vivo depletion of CD11c+ cells delays the CD4+ T cell response to Mycobacterium tuberculosis and exacerbates the outcome of infection.
J. Immunol.
175
:
3268
-3272.
19
Hagymasi, A. T., A. M. Slaiby, M. A. Mihalyo, H. Z. Qui, D. J. Zammit, L. Lefrancois, A. J. Adler.
2007
. Steady state dendritic cells present parenchymal self-antigen and contribute to, but are not essential for, tolerization of naive and Th1 effector CD4 cells.
J. Immunol.
179
:
1524
-1531.
20
Bergtold, A., D. D. Desai, A. Gavhane, R. Clynes.
2005
. Cell surface recycling of internalized antigen permits dendritic cell priming of B cells.
Immunity
23
:
503
-514.
21
Qi, H., J. G. Egen, A. Y. Huang, R. N. Germain.
2006
. Extrafollicular activation of lymph node B cells by antigen-bearing dendritic cells.
Science
312
:
1672
-1676.
22
Wykes, M., A. Pombo, C. Jenkins, G. G. MacPherson.
1998
. Dendritic cells interact directly with naive B lymphocytes to transfer antigen and initiate class switching in a primary T-dependent response.
J. Immunol.
161
:
1313
-1319.
23
Pape, K. A., D. M. Catron, A. A. Itano, M. K. Jenkins.
2007
. The humoral immune response is initiated in lymph nodes by B cells that acquire soluble antigen directly in the follicles.
Immunity
26
:
491
-502.
24
Hebel, K., K. Griewank, A. Inamine, H. D. Chang, B. Muller-Hilke, S. Fillatreau, R. A. Manz, A. Radbruch, S. Jung.
2006
. Plasma cell differentiation in T-independent type 2 immune responses is independent of CD11chigh dendritic cells.
Eur. J. Immunol.
36
:
2912
-2919.
25
Kawamura, Y. I., R. Kawashima, Y. Shirai, R. Kato, T. Hamabata, M. Yamamoto, K. Furukawa, K. Fujihashi, J. R. McGhee, H. Hayashi, T. Dohi.
2003
. Cholera toxin activates dendritic cells through dependence on GM1-ganglioside which is mediated by NF-κB translocation.
Eur. J. Immunol.
33
:
3205
-3212.
26
Robertson, J. M., P. E. Jensen, B. D. Evavold.
2000
. DO11.10 and OT-II T cells recognize a C-terminal ovalbumin 323–339 epitope.
J. Immunol.
164
:
4706
-4712.
27
Kitamura, D., K. Rajewsky.
1992
. Targeted disruption of mu chain membrane exon causes loss of heavy-chain allelic exclusion.
Nature
356
:
154
-156.
28
Madsen, L., N. Labrecque, J. Engberg, A. Dierich, A. Svejgaard, C. Benoist, D. Mathis, L. Fugger.
1999
. Mice lacking all conventional MHC class II genes.
Proc. Natl. Acad. Sci. USA
96
:
10338
-10343.
29
Zammit, D. J., L. S. Cauley, Q. M. Pham, L. Lefrancois.
2005
. Dendritic cells maximize the memory CD8 T cell response to infection.
Immunity
22
:
561
-570.
30
Crawford, A., M. Macleod, T. Schumacher, L. Corlett, D. Gray.
2006
. Primary T cell expansion and differentiation in vivo requires antigen presentation by B cells.
J. Immunol.
176
:
3498
-3506.
31
Asselin-Paturel, C., G. Brizard, J. J. Pin, F. Briere, G. Trinchieri.
2003
. Mouse strain differences in plasmacytoid dendritic cell frequency and function revealed by a novel monoclonal antibody.
J. Immunol.
171
:
6466
-6477.
32
Villavedra, M., H. Carol, M. Hjulstrom, J. Holmgren, C. Czerkinsky.
1997
. “PERFEXT”: a direct method for quantitative assessment of cytokine production in vivo at the local level.
Res. Immunol.
148
:
257
-266.
33
Daley, J. M., A. A. Thomay, M. D. Connolly, J. S. Reichner, J. E. Albina.
2008
. Use of Ly6G-specific monoclonal antibody to deplete neutrophils in mice.
J. Leukocyte Biol.
83
:
64
-70.
34
Crowley, M. T., C. R. Reilly, D. Lo.
1999
. Influence of lymphocytes on the presence and organization of dendritic cell subsets in the spleen.
J. Immunol.
163
:
4894
-4900.
35
Moulin, V., F. Andris, K. Thielemans, C. Maliszewski, J. Urbain, M. Moser.
2000
. B lymphocytes regulate dendritic cell (DC) function in vivo: increased interleukin 12 production by DCs from B cell-deficient mice results in T helper cell type 1 deviation.
J. Exp. Med.
192
:
475
-482.
36
Kuwajima, S., T. Sato, K. Ishida, H. Tada, H. Tezuka, T. Ohteki.
2006
. Interleukin 15-dependent crosstalk between conventional and plasmacytoid dendritic cells is essential for CpG-induced immune activation.
Nat. Immunol.
7
:
740
-746.
37
Kool, M., T. Soullie, M. van Nimwegen, M. A. Willart, F. Muskens, S. Jung, H. C. Hoogsteden, H. Hammad, B. N. Lambrecht.
2008
. Alum adjuvant boosts adaptive immunity by inducing uric acid and activating inflammatory dendritic cells.
J. Exp. Med.
205
:
869
-882.
38
van Rijt, L. S., S. Jung, A. Kleinjan, N. Vos, M. Willart, C. Duez, H. C. Hoogsteden, B. N. Lambrecht.
2005
. In vivo depletion of lung CD11c+ dendritic cells during allergen challenge abrogates the characteristic features of asthma.
J. Exp. Med.
201
:
981
-991.
39
Zaft, T., A. Sapoznikov, R. Krauthgamer, D. R. Littman, S. Jung.
2005
. CD11chigh dendritic cell ablation impairs lymphopenia-driven proliferation of naive and memory CD8+ T cells.
J. Immunol.
175
:
6428
-6435.
40
Yrlid, U., V. Cerovic, S. Milling, C. D. Jenkins, J. Zhang, P. R. Crocker, L. S. Klavinskis, G. G. MacPherson.
2006
. Plasmacytoid dendritic cells do not migrate in intestinal or hepatic lymph.
J. Immunol.
177
:
6115
-6121.
41
Pascale, F., V. Contreras, M. Bonneau, A. Courbet, S. Chilmonczyk, C. Bevilacqua, M. Epardaud, V. Niborski, S. Riffault, A. M. Balazuc, et al
2008
. Plasmacytoid dendritic cells migrate in afferent skin lymph.
J. Immunol.
180
:
5963
-5972.
42
Hornqvist, E., T. J. Goldschmidt, R. Holmdahl, N. Lycke.
1991
. Host defense against cholera toxin is strongly CD4+ T cell dependent.
Infect. Immun.
59
:
3630
-3638.
43
Probst, H. C., K. Tschannen, B. Odermatt, R. Schwendener, R. M. Zinkernagel, M. Van Den Broek.
2005
. Histological analysis of CD11c-DTR/GFP mice after in vivo depletion of dendritic cells.
Clin. Exp. Immunol.
141
:
398
-404.
44
Delemarre, F. G., N. Kors, N. van Rooijen.
1990
. The in situ immune response in popliteal lymph nodes of mice after macrophage depletion: differential effects of macrophages on thymus-dependent and thymus-independent immune responses.
Immunobiology
180
:
395
-404.
45
Hershberg, R. M., L. F. Mayer.
2000
. Antigen processing and presentation by intestinal epithelial cells: polarity and complexity.
Immunol. Today
21
:
123
-128.
46
Perrigoue, J. G., S. A. Saenz, M. C. Siracusa, E. J. Allenspach, B. C. Taylor, P. R. Giacomin, M. G. Nair, Y. Du, C. Zaph, N. van Rooijen, et al
2009
. MHC class II-dependent basophil-CD4+ T cell interactions promote TH2 cytokine-dependent immunity.
Nat. Immunol.
10
:
697
-705.