B6.MRL-Tnfrsf6^lpr mice were from The Jackson Laboratory. C57BL/6 mice were from Shanghai SLAC Laboratory Animal Center (Shanghai, China). All mice were kept in a specific pathogen-free (SPF) barrier facility maintained by Shanghai Jiao Tong University School of Medicine. All the experimental mice were used at 6–10 weeks. Animal protocols were approved by Institutional Animal Care and Use Committee of Shanghai Jiao Tong University School of Medicine.

BMDCs were cultured as previously described (16). Briefly, bone marrow cells were collected by perfusing mouse femur and tibia. After red blood cell lysis, cells were cultured in RPMI 1640 (Invitrogen Corp.) supplemented with 10% fetal calf serum (FCS, Gibco), penicillin and streptomycin (Invitrogen), 2-mercaptoethanol (Sigma-Aldrich), supernatant from J5 cells (provided by Dr. Qibin Leng, Institut Pasteur of Shanghai, China) expressing GM-CSF (1:50) and 10 ng/ml IL-4 (R&D). On day 3, the entire medium was removed and replaced with fresh differentiation medium. On day 7, the cells were harvested for analyses. The purity of CD11c⁺ BMDCs was >80%.

In BMDC adoptive transfer experiment, wild-type or Fas-deficient BMDCs were pulsed with 50 μg/ml HDM (Greer Laboratories, Lenoir, NC) for 12 h, washed, and then 1 × 10⁶ HDM-pulsed BMDCs were administered intravenously into naïve C57BL/6 recipients. Recipients transferred with un-pulsed wild-type or Fas-deficient BMDCs as control. On day 10–12, all the recipients were lightly anesthetized and challenged intranasally with 10 μg HDM in 40 μl PBS and the mice were sacrificed for analysis on day 13.

Anti-mouse CD11c (N418), MHC-II (M5/114.15.2), CD11b (M1/70), Ly6G (RB6-8C5), siglec F (E50-2440), CD4 (RM4-5), TCRβ (H57-597), IL-4 (11B11), IL-13 (eBio13A), IL-17A (eBio17B7), CD25 (PC61.5), CD44 (IM7), CD62L (MEL-14), CD69 (H1.2F3), Ki-67 (SolA15), IL-12p40 (C17.8), CD45 (30-F11), CD178 (MFL3) antibodies were obtained from eBioscience. Anti-mouse IL-5 (TRFK5) was obtained from BD Biosciences. Anti-mouse CCR3 (J073E5) was obtained from Biolegend. For surface staining, cells were stained with antibodies in PBS containing 1% FCS (Hyclone) on ice for 30 min. For intracellular staining, cells were stimulated with PMA (Sigma-Aldrich) and ionomycin (Sigma-Aldrich) for 5 h in the presence of Golgistop (BD Biosciences) before being stained according to the manufacturer's instructions (eBioscience). CD4⁺T cell proliferation was detected by anti-Ki-67 staining according to the manufacturer's instructions (eBioscience) 3 days after co-culture. Labeling OTII CD4⁺ T cells with CFSE (carboxyfluorescein diacetate succinimidyl diester; Invitrogen), cell proliferation was detected by flow cytometry 4 days after co-culture. For cell apoptosis analysis, cells were stained with CaspACE™ FITC-VAD-FMK in situ Marker (Promega). The samples were acquired on a FACSCantoII (BD) or LSRFortessa^TM X-20 (BD) and analyzed with FlowJo software (Treestar).

For BALF collection, lung tissues were lavaged with 1 ml cold PBS for 3 times and the supernatant was collected. Lung mononuclear cells were prepared as previously described (17). Briefly, lung tissues were removed, minced and digested with 1 mg/ml Collagenase IV (Life Technologies) in RPMI-1640 (Hyclone) with 5% FCS (Hyclone) for 45 min at 37°C. Cells were enriched by using 38% Percoll gradient (GE Healthcare Life Sciences). Red blood cells were lyzed with ACK lysis buffer (R&D Systems). Cells were harvested for analyses.

Left lobe of lung tissues were removed from mice after BALF collection, fixed with 4% paraformaldehyde (PFA) at room temperature for 24 h and embedded in paraffin, cut into 5-μm sections for hematoxylin and eosin (HE) or periodic acid-Schiff (PAS) staining. The lung inflammation was blindly quantified using HE-stained sections according to the criteria previously published (18). The quantification of the goblet cell hyperplasia in the airway was done as previously described (19).

Total RNAs of lung tissues were isolated using Trizol regent (Invitrogen). Total RNAs of cells were isolated using RNeasy mini Kit (QIAGEN) according to the manufacturer's instructions. 1 μg total RNA was used for reverse transcription with PrimeScript RT Master Mix (TAKARA) according to the manufacturer's instructions in a total volume of 20 μl. qPCR was carried out with SYBR Green PCR Master Mix (Applied Biosystems) in a Vii7 Real-Time PCR system (Applied Biosystems). mRNA expression of genes was normalized to Hprt. The primers shown below were from Primerbank: Hprt, forward primer: TCAGTCAACGGGGGACATAAA, reverse primer: GGGGCTGTACTGCTTAACCAG; Il4, forward primer: GGTCTCAACCC CCAGCTAGT, reverse primer: GCCGATGATCTCTCTCAAGTGAT; Il17a, forward primer: TCAGCGTGTCCAAACACTGAG, reverse primer: CGCCAAGGGAG TTAAAGACTT; Ifng, forward primer: GCCACGGCACAGTCATTGA, reverse primer: TGCTGATGGCCTGATTGTCTT; Il6, forward primer: CTGCAAGAGACTTCCATCCAG, reverse primer: AGTGGTATAGACAGGTATGTTGG; Il12p35, forward primer: CAATCACGCTACCTCCTCTTT, reverse primer: CAGCAGTGCAGGAATAATGTTTC; Il12p40, forward primer: GTCCTCAGAAGCTAACCATCTC, reverse primer: CCAGAGC CTATGACTCCATGTC; Il10, forward primer: CTTACTGACTGGCATGAGGATCA, reverse prime: GCAGCTCTAGGAGCATGTGG. The other primers were used as described, such as Il5, Il13 (20), Gata3 (21), Il9 (22), Tnfsf4 (8), Cd86 (23), Tslp (24).

BMDCs were stimulated with HDM in the presence or absence of Fas agonistic antibody Jo2 (1 μg/ml, BD) or Fas antagonistic antibody kp7-6 (1 mM, Merk) for 5 h for RNA analysis. For drug inhibitor treatments, cells were incubated with vehicle (DMSO) or U0126 (10 μM) (from Calbiochem) for 0.5–1 h before adding other stimuli. For BMDC–T cell co-culture, BMDCs and flow cytometry-sorted naïve OT-II CD4⁺ T cells (CD4⁺25⁻CD44⁻CD62L⁺, purity >99%) were mixed at a ratio of 1:10 in the presence of OVA_323−339 peptide and HDM, and then the CD4⁺ T cells were harvested at 48 h for mRNA analysis or supernatant was harvested at 72 h for ELISA. For cytokine treatment, cultures were supplemented with 1 ng/ml IL-12p70 (R&D).

Concentrations of IL-4 and IL-13 were measured by ELISA according to the manufacturer's instructions (R&D; eBioscience). Read the OD values at 450 nm on the MultiSKAN GO microplate reader (Thermo). Immunoblot analysis was performed as described (25) with antibody to ERK phosphorylated at Thr202 and Tyr204, antibody to p38 phosphorylated at Thr180 and Tyr182, antibody to JNK phosphorylated at Thr183 and Tyr185 and antibody to ERK (all from Cell Signaling Technology), antibody to alpha tubulin (Proteintech).

All statistical analyses were performed by unpaired Student's t-tests or ANOVA using GraphPad Prism software (version 5.0). P < 0.05 was considered significant. Results represent means ± SEM.

To explore the role of Fas signaling in DCs in the regulation of HDM-induced allergic inflammation in mice, we used a BMDC adoptive transfer protocol to induce lung inflammation (Figure 1A). We transferred HDM-pulsed or un-pulsed wild-type or Fas-deficient BMDCs into naïve wild-type recipient mice. After HDM re-challenged, mice received HDM-pulsed BMDCs showed higher total cell number in the bronchoalveolar lavage (BAL) compared to mice received un-pulsed BMDCs (Figure 1B). A significantly increased total cell number was also observed in the BAL of recipients transferred with HDM-pulsed Fas-deficient BMDCs (Figure 1B). We also observed higher inflammatory cell infiltration and mucus production in lung tissues of recipients transferred with HDM-pulsed Fas-deficient BMDCs than those transferred with HDM-pulsed wild-type BMDCs (Figures 1C,D). Flow cytometry showed that a dramatically increased eosinophil infiltration both in the BAL and lung tissues of recipients transferred with HDM-pulsed Fas-deficient BMDCs compared to those transferred with HDM-pulsed wild-type BMDCs (Supplementary Figures 1A,B). We also analyzed the inflammatory eosinophils (iEos) (CD45⁺Siglec F^intCCR3⁺CD62L⁻) and resident eosinophils (rEos) (CD45⁺Siglec F^intCCR3⁺CD62L⁺) in lung tissues (26). The cell number of iEos was increased in recipients transferred with HDM-pulsed Fas-deficient BMDCs compared to those transferred with HDM-pulsed wild-type BMDCs, but rEos was comparable between recipients transferred with HDM-pulsed wild-type BMDCs and those transferred with Fas-deficient BMDCs (Figure 1E). However, the neutrophil infiltration had no difference between the two groups (Supplementary Figures 1C,D). Together, these data indicate that HDM-pulsed Fas-deficient BMDCs can induce more severe allergic airway inflammation than HDM-pulsed wild-type BMDCs.

In addition to a role of innate immune cells in the development of allergic inflammation, the adaptive immune system also play an important role in driving and sustaining this inflammation (27). A comparable CD4⁺ T cell activation was observed in lung tissues of the recipients transferred with HDM-pulsed wild-type or Fas-deficient BMDCs (Figure 2A). We also examined whether Fas-deficient in BMDCs could affect CD4⁺ T cell proliferation or apoptosis in vivo, we performed Ki-67 and VAD staining assay, respectively. Our results showed that the proportion of Ki-67⁺CD4⁺ T cells and the median fluorescence intensity (MFI) value of VAD⁺CD4⁺ T cell in lung tissues and mediastinal lymph nodes (mLN) had no differences between the recipients transferred with HDM-pulsed wild-type BMDCs and those transferred with Fas-deficient BMDCs (Figures 2B–E). Taken together, these results show that Fas signaling in BMDCs is not required for CD4⁺ T cell activation, proliferation and apoptosis upon HDM treatment.

Given that there was no difference in the proliferation and apoptosis of CD4⁺ T cells in the recipients transferred with HDM-pulsed wild-type or Fas-deficient BMDCs, we reasoned that the difference in inflammatory response might be due to the potential of these cells to produce inflammatory cytokines. Thus, we first measured cytokine expression in lung tissues of these two group mice. The qPCR results showed that the recipients transferred with HDM-pulsed Fas-deficient BMDCs had higher mRNA expression of Il4 (encoding IL-4), Il5 (encoding IL-5), and Il13 (encoding IL-13), but comparable expression of Ifng (encoding IFNγ) and Il17a (encoding IL-17A) in lung tissues than those transferred with HDM-pulsed wild-type BMDCs (Figure 3A). Although a comparable percentage and cell number of CD4⁺ T cells was observed in lung tissues of the two HDM-pulsed groups (Figure 3B), intracellular staining showed that recipients transferred with HDM-pulsed Fas-deficient BMDCs had higher percentage of IL-4⁺, IL-5⁺, and IL-13⁺CD4⁺ T cells in lung tissues than those transferred with wild-type BMDCs, along with higher cell number of IL-4⁺, IL-5⁺, and IL-13⁺CD4⁺ T cells (Figures 3C,D), whereas the percentage and cell number of IL-17⁺CD4⁺ T cells were similar in the two groups (Supplementary Figures 2A,B). However, the percentage and cell number of IL-4⁺, IL-5⁺, and IL-13⁺CD4⁺ T cells had no difference between the recipients transferred with un-pulsed wild-type and Fas-deficient BMDCs (Supplementary Figures 2C,D). Taken together, these results demonstrate that HDM-pulsed Fas-deficient BMDCs promote Th2 responses in vivo.

To determine whether the increased Th2 response in vivo is due to the direct interaction between DCs and T cells, we co-cultured wild-type or Fas-deficient BMDCs with naïve OT-II CD4⁺ T cells in the presence of OVA_323−339 peptide with or without HDM. The polarization of CD4⁺ T cells was determined by qPCR or ELISA, respectively. OT-II CD4⁺ T cells activated with HDM-stimulated Fas-deficient BMDCs had higher IL-4 and IL-13 expression both in mRNA and in protein levels than those stimulated with wild-type BMDCs (Figures 4A,B). GATA3, the master transcription factor for Th2 cell differentiation, was also found increased in T cells activated by HDM-pulsed Fas-deficient BMDCs, while the expression of Il9 and Il10 was comparable (Figure 4A). The activation and proliferation of OT-II CD4⁺ T cells activated by HDM-pulsed both wild-type and Fas-deficient BMDCs were comparable (Figures 4C–E). Collectively, these data indicate that Fas signaling mediates the direct crosstalk between BMDCs and Th2 cells upon HDM stimulation in vitro.

Next we explored the molecular mechanism by which Fas signaling in BMDCs to shape Th2 differentiation upon HDM stimulation. We stimulated wild-type BMDCs with Fas agonistic antibody Jo2, the expression of Cd86, Tslp, Il10, Il6, and Tnfsf4, which had been reported to regulate Th2 polarization, had no difference compared to control group (Figure 5A and Supplementary Figure 3A). HDM stimulation could dramatically increase the expression of Il6 and Tnfsf4, while Fas agonistic antibody Jo2 did not affect the expression of these two genes (Figure 5A), indicating that FAS signaling is not required for Il6 and Tnfsf4 expression in DCs upon HDM stimulation. IL-12 has been reported to affect Th2 differentiation (28). Fas agonistic antibody Jo2 stimulation could not affect the expression of Il12p35 and Il12p40 in wild-type BMDCs without HDM stimulation (Figure 5A). Upon HDM stimulation, the expression of Il12p35 and Il12p40 was significantly increased compared with non-HDM stimulation, and Fas agonistic antibody Jo2 could further enhance the expression of Il12p35 and Il12p40 in mRNA level and IL-12p70 in protein level (Figures 5A,B). Given that HDM-stimulated BMDCs had increased FasL, Fas antagonistic antibody kp7-6-treated HDM-pulsed wild-type BMDCs produced lower Il12p40 in mRNA level and IL-12p70 in protein level than those of BMDCs stimulated with HDM alone, but the expression of Il12p35 had no difference between HDM and HDM plus kp7-6 stimulated BMDCs (Figures 5C,D and Supplementary Figure 3B). Fas-deficient BMDCs had decreased expression of Il12p35 and Il12p40 compared to wild-type BMDCs upon HDM stimulation in the presence of Fas agonistic antibody Jo2 (Figure 5E). We next examined whether the lower expression of IL-12 in Fas-deficient BMDCs could contribute to the increased Th2 differentiation. We added exogenous IL-12 into the BMDC–T cell co-culture system and found that the IL-12 supplement could significantly decrease the expression of Th2-related cytokines, such as IL-4 and IL-13 in OT-II CD4⁺ T cells activated by Fas-deficient BMDCs. The expression of IFNγ and IL-17 was comparable between wild-type and Fas-deficient BMDC activated T cells (Figure 5F). Altogether, these results indicate that Fas-deficient BMDCs promote Th2 differentiation through downregulation of IL-12 expression.

We next examined the downstream signaling of Fas involved in the regulation of IL-12 expression by analyzing the activation of p38, JNK, and ERK in BMDCs. We found that the phosphorylation of ERK was decreased in wild-type BMDCs treated with Fas agonistic antibody Jo2 and HDM compared to those treated with HDM alone (Figure 6A). Accordingly, the activation of ERK in Fas-deficient BMDCs was increased compared to that of wild-type BMDCs upon HDM and Fas agonistic antibody Jo2 stimulation (Figure 6B). To determine whether the increased ERK activation was contributed to the decreased IL-12 expression in Fas-deficient BMDCs, we treated Fas-deficient BMDCs with specific ERK inhibitor U0126, which resulted in a dramatically increased IL-12p70 expression in Fas-deficient BMDCs (Figure 6C). Consequently, U0126-treated Fas-deficient BMDCs completely restored the increased IL-4 and IL-13 expression in T cells to the level of T cells activated with wild-type BMDCs (Figure 6D). These results indicate that Fas signaling regulates IL-12 expression and Th2 differentiation through modulating the ERK activity in BMDCs.

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.