The authors have declared that no competing interests exist.
Conceived and designed the experiments: MZ WZ. Performed the experiments: KZ WS QW DC ZZ YW. Analyzed the data: MZ WS KZ XL CW. Contributed reagents/materials/analysis tools: MZ WS XL CW YW. Wrote the paper: MZ KZ.
The contraction phase of antigen-specific immune responses involves the apoptotic loss of numerous activated lymphocytes. While apoptotic cells are known to induce immune suppression, the mechanisms involved therein are still ambiguous. Some reports have speculated that macrophages can induce regulatory T cells (Tregs) after engulfing apoptotic cells. In this study, we showed that dendritic cells (DCs) that phagocytose apoptotic T cells acquire inhibitory function (named DCapos) toward CD4+ and CD8+ T cells. These inhibitory DCs could not induce the generation of Tregs, but they were found to directly inhibit mDCs that initiate CD4+ and CD8+ T cell proliferation both in vitro and in vivo. Soluble factors including NO play a role in the DCapos-induced suppression of CD4+ and CD8+ T cell proliferation. Further results showed that STAT3 phosphorylation and inducible nitric oxide synthase (iNOS) generation were enhanced when DCs were co-cultured with apoptotic cells. Both iNOS transcription and NO secretion were inhibited in the presence of the specific p-STAT3 inhibitor JSI-124. All the data indicated that apoptotic cells could turn DCs to inhibitory DCs, which might play important roles in the suppression of immune responses. STAT3 activation and the consequent release of NO are responsible for the inhibitory functions of DCapos.
Activation-induced cell death (AICD) of T cells is a key mechanism for downregulating the immune response. For apoptotic cell clearance, antigen-presenting cells (APCs) phagocytose apoptotic T cells to maintain homeostasis of the immune system. These APCs that have phagocytosed apoptotic T cells can also induce immune tolerance, which is considered to be due to the subsequently induced Treg’s effect
DCs are pivotal professional APCs with important functions both in initiating immune response and in maintaining immune tolerance. It has been demonstrated that a distinct group of DCs with suppressive function emerge after exposure to certain cytokines or when the DCs re-differentiation in different stromal microenvironments
Our preliminary data showed that DCs also produce NO after phagocytosing apoptotic T cells, which are similar to regulatory DCs derived from mature DCs co-cultured with stoma, suggesting that after phagocytosing apoptotic T cells, DCs may directly inhibit the immune response. Here, we evaluated the regulatory function of DCs after phagocytosing apoptotic T cells and investigated the relationship between the phagocytosis of apoptotic T cells and NO production in DC.
Six-week-old C57BL/6 (H-2Kb) mice and BALB/c (H-2Kd) mice were purchased from Vitariver (Beijing, China). OVA323–339-specific TCR-transgenic mice DO11.10 (H-2Kd), OVA257–264-specific TCR-transgenic mice OT-1, Foxp3EGFP mice (H-2Kd), and EGFP-transgenic mice C57BL/6-Tg (ACTb-EGFP) (H-2Kb) were obtained from the Jackson Laboratory (Bar Harbor, ME). F1 mice were prepared by crossing C57BL/6 mice with DO11.10 mice (C57BL/6×DO11.10). Another set of F1 mice were prepared by crossing Foxp3EGFP with DO11.10 mice (Foxp3EGFP×DO11.10). All the mice were maintained under specific pathogen-free conditions and used at 6–8 weeks of age. All experimental manipulations were undertaken in accordance with the National Institute of Health Guide for the Care and Use of Laboratory Animals, with the approval of the Tsinghua University Animal Care and Use Committee, Beijing.
CFSE, STAT3 inhibitor JSI-124 (cucurbitacin I), 1,4-phenylene-bis(1,2-ethanediyl)bis-isothiourea dihydrobromide (1,4-PBIT), and the NO donor NOC-18 were all purchased from Sigma-Aldrich (St. Louis, MO). Magnetic beads conjugated with mAbs to CD4, CD11b, and CD11c were purchased from Miltenyi Biotec (Bergisch Gladbach, Germany). Fluorescein-conjugated mAbs to CD4, CD8a, CD11b, CD11c, CD40, CD45.1, CD80, CD86, Ia, KJ1-26, Vβ5.1/5.2, and isotype control mAbs were purchased from BD Pharmingen (San Diego, CA). Specific Abs against STAT3 (Cell Signaling Technology, Beverly, MA), phosphor-STAT3 (Tyr705) (Cell Signaling Technology), iNOS (Cell Signaling Technology), beta-actin (Santa Cruz Biotechnology) were used.
Splenic CD4+ T cells from DO11.10×C57BL/6 F1 hybrid mice were obtained by magnetic cell sorting and then co-cultured in 24-well plates with mDCs (DCs that were stimulated by LPS 1 ng/ml for more than 24 hours) for 5 days at a ratio of 1∶10 in the presence of OVA323–339. The CD4+ T cells were sufficiently activated and used as activated CD4+ T cells. Thereafter the CD4+ T cells were obtained by negative selection with CD11b magnetic microbeads and subsequent cultured without mDCs for another 2 days to induce apoptosis of more than 70% of the CD4+ T cells. The CD4+ T cells were stained by Hoechst 33342, and then centrifuged at 100
Bone marrow-derived DCs were prepared from male or female mice in the presence of GM-CSF and IL-4 as described previously
DCs and DCapos were incubated with Alexa 488-conjugated OVA at a final concentration of 100 µg/ml, and the geometric mean fluorescence was detected using FACSAria
As described previously
CFSE-labeled OVA323–339-specific TCR-transgenic splenic CD4 T cells (5×106 cells) were freshly isolated from DO11.10×C57BL/6 F1 hybrid mice, with OVA323–339-loaded mDCs and/or DCapos injected intraperitoneally into each F1 (BALB/c×C57BL/6) hybrid mouse. After 4 days, mononuclear cells from peripheral blood were prepared and double stained with CD4-APC and KJ1-26-PE. The CD8 T cells were transferred in the same way. CFSE-labeled OVA257–264-specific TCR-transgenic splenic CD8 T cells (1×106 cells) were freshly isolated from OT1 mice, with OVA257–264-loaded mDCs and/or DCapos injected intraperitoneally into each C57BL/6 mouse. After 3 days, mononuclear cells from the peripheral blood were prepared and double stained with CD8-APC and Vβ5.1/5.2-PE. To investigate the development of Tregs in vivo, apoptotic cells from BALB/c mice were prepared and injected intraperitoneally into Foxp3EGFP or (Foxp3EGFP×DO11.10) F1 mice.
We collected supernatants for detection of cytokines. Cytokines were detected on a commercial ELISA kit (Bender Medsystems) according to the manufacturer’s instructions. NO production was determined by measuring the nitrite concentration by Griess assay, as described previously
Total cellular RNA from DCs or DCapos was isolated by using TRIzol reagent (Invitrogen). The RNA was reverse transcribed using a TransScript II First-Strand cDNA Synthesis SuperMix (TransGen Biotech, Beijing, China). The RT-PCR was carried out using the following mouse-specific primer pairs: iNOS (forward:
Western blotting was carried out as described
Data were obtained from 3 independent experiments. All statistical analyses were performed with ANOVA using SPSS version 11.0. A value of p < 0.05 was considered statistically significant.
Naïve ovalbumin (OVA)-specific TCR transgenic CD4+ T cells can be activated by DCs in the presence of OVA peptide (OVA323–339). Approximately 70–80% of activated T cells undergo apoptosis when cultured for an additional 2 days without DCs; which can be verified by the morphological changes observed in the T cells (
A) Apoptosis of CD4+ T cells. Left panel, top left (×400): CD4+ T cells stained by Hoechst 33342 after activation by mDCs and OVA323–339 for 2 days; bottom left (×400): after activation by mDCs and OVA323–339 for 5 days, CD4+ T cells were cultured for another 2 days in the absence of mDCs and then stained by Hoechst 33342. Numbers showed the percentage of apoptotic cells in high power field. Upper right panel, control cells. Lower right panel, apoptotic CD4+ T cells were assessed by flow cytometry with Annexin V and PI. (B) Purification of DCapos. Top panel, DCs were co-cultured with apoptotic CD4+ T cells prelabeled with Hoechst 33342. Then the CD11b+Hoechst+ cells were sorted by flow cytometry. Numbers showed the purity of the sorted cells. Bottom panel, DCs incubated with apoptotic CD4+ T cells prelabeled with Hoechst 33342 (blue with red arrows) and incubated for 2 hours. The DCs were then stained with CD11b-FITC (green) and observed under a light microscope (left) and a confocal immunofluorescence microscope (right). Bar represent 15 µm. (C) Morphologies and phenotypes of mDCs and DCapos. Left panel, left top (×400), the morphology of mDCs; left bottom (×400), the morphology of DCapos. Right panel, different groups of DCs were stained with fluorescence-conjugated anti-CD11b, anti-CD11c, anti-CD80, anti-CD86, anti-CD40, and anti-Ia mAbs for flow cytometric analysis. Numbers represented the mean fluorescence intensity. (D) Endocytic abilities of mDCs and DCapos were assessed by flow cytometry of Alexa 488-OVA uptake. Numbers in the histograms indicate the geometric mean fluorescence of each DC population. Ctrl, control (cells incubated with Alexa 488-OVA at 4°C). Data represent results of 1 of the 3 experiments performed to yield similar results.
Naïve TCR-transgenic CD4+ T cells obtained from DO11.10×C57BL/6 F1 mice (
For CD4+ T cells, purified CD4+ T cells from DO11.10×C57BL/6 F1 hybrid mice (2×105 cells) stained with CFSE were co-cultured in vitro with OVA323–339-loaded mDCs (2×104 cells) in the presence or absence of DCapos (2×104 cells) for 4 days. For CD8+ T cells, purified OT1-CD8+ T cells (2×105 cells) stained with CFSE were co-cultured in vitro with OVA257–264-loaded mDCs (2×104 cells) in the presence or absence of DCapos (2×104 cells) for 3 days. (A) The cells were collected at 4 days, and both CD4- and KJ1-26-positive cells were counted using flow cytometry. (B) The cells were collected at 3 days, and both CD8- and Vβ5.1/5.2-positive cells were counted using flow cytometry. Histograms showed the mean ± SEM of triplicate wells counted using flow cytometry, and results represent 3 separate experiments. *
To further confirm the inhibitory function of DCapos, we adoptively transferred naïve transgenic CD4+ and CD8+ T cells together with the corresponding peptide-loaded mDCs and DCapos into donor mice to test T cell proliferation. In our previous study, we obtained consistent results for the blood and spleen T cells. Therefore, the transferred CD4+ and CD8+ T cells in the blood can be recognized by the corresponding TCR-specific monoclonal antibodies. Direct cell counting (
A) CD4+ T cells (5×106 cells) from DO11.10×C57BL/6 F1 hybrid mice stained with CFSE, OVA323–339-loaded mDCs (1×106 cells) with or without DCapos (1×106 cells) were transferred together into the peritoneal cavity of normal mice for 4 days. Mononuclear cells from the blood were separated and stained for flow cytometric analysis of the frequency of both CD4- and KJ1-26-positive T cells. (B) The CFSE dilutions of CD4+KJ1-26+ T cells were analyzed using flow cytometry. (C) OT1 CD8+ T cells (1×106 cells), OVA257–264-loaded mDCs (3×105 cells) with or without DCapos (3×105 cells) were transferred together into the peritoneal cavity of normal mice for 3 days. Then, the mononuclear cells were separated from the blood and stained for flow cytometric analysis of the frequency of both CD8- and Vβ5.1/5.2-positive T cells. (D) The CFSE dilutions of CD8+ Vβ5.1/5.2+ T cells were analyzed by flow cytometry. Data are presented as mean ± SD of 5 mice, and results represent 1 of the 2 experiments with similar results.
Our previous studies on stroma-educated regulatory DCs (DCregs) suggested that NO is an important soluble factor released by DCregs, and is involved in their immunosuppression
Purified DO11.10 CD4+ T cells (2×105 cells) stained with CFSE were co-cultured with OVA323–339-loaded mDCs (2×104 cells) in the presence or absence of DCapos (2×104 cells) for 4 days in vitro. (A) The NO donor NOC-18 and the NOS inhibitor PBIT were added to the co-culture system to evaluate their effect on the inhibitory function of DCapos. (B) The supernatants of different groups were collected and the NO secretions were tested by Griess assay. *
Tregs are known to play an indispensable role in maintaining immunological unresponsiveness to self-antigen and in suppressing excessive immune responses that are deleterious to the host. We then evaluated whether Tregs were involved in the DCapos-mediated suppression of the specific immune responses. We used hybrid mice co-expressing enhanced green fluorescent protein (EGFP) and the regulatory T cell-specific transcription factor Foxp3 on OVA323–339-specific TCR-transgenic CD4+ T cells. As compared to the CD4+ group, the percentage of Foxp3+CD25+ cells in the DCapo/CD4 group slightly increased but without statistically significant difference. Surprisingly, in the mDC/CD4 group, the Foxp3+CD25+ cells proliferated according to the total cell number. There was no evidence that DCapos induced the generation of Tregs (
(A) Left, DO11.10 CD4+ T cells (2×105 cells) with Foxp3EGFP were purified, and the CD4+CD25+Foxp3+ cells were measured in a FACS system (numbers indicate percentages). These cells were then co-cultured with OVA323–339-loaded mDCs (2×104) in the presence or absence of DCapos (2×104 cells) for 5 days in vitro. On the fifth day, the cells were harvested and the numbers of CD4- and KJ1-26-positive cells were determined by FACS analysis (black). The proportion of CD4+KJ1-26+CD25+Foxp3+ cells was determined by FACS (grey). Data are presented as mean ± SD of triplicate wells, and the results represent 2 separate experiments. (B, C) Apoptotic cells (AC), DCapos, or phosphate-buffered saline (PBS) was injected into the peritoneal cavity of Foxp3EGFP mice (PBS,
We further explored the mechanisms of AC-induced inhibition by DCapos. Recently, the tyrosine kinase Mer (MerTK) receptor was implicated in the blockade of NF-kappaB activation by apoptotic cells in DCs
(A) Apoptotic cells (AC) induced STAT3 activation in DCapos. DCs incubated with or without different concentrations of JSI-124 for 1 h were treated with or without ACs for the indicated times, then stimulated with 50 ng/ml LPS for 0.5 hours. The p-STAT3, STAT3, iNOS, and β-actin proteins were measured in whole-cell lysates by western blotting using the same membranes. (B) DCs were incubated with or without different doses of JSI-124 for 1 h, co-cultured with or without ACs for 2 h, then cultured for 24 hours before the cells were used for RT-PCR assay of mRNA expression of iNOS, TGF-β, and β-actin. Data represents 1 of 3 experiments with similar results. (C) The supernatants of different groups were collected and NO was analyzed by the Griess assay. *
Previous data have demonstrated that DCapos exert inhibitory functions via NO. Since STAT3 signaling was involved in blocking DC activation
Our previous study had demonstrated that mature DCs in stromal microenvironment could re-differentiate into DCregs after antigen presentation, but not undergo apoptosis
In this study, we constructed a model in which most lymphocytes underwent AICD after the stimulation of mature DCs. Some molecules displayed on the surface of apoptotic cells (AC), particularly phosphatidylserine, are reportedly recognized as “eat-me” signals by DCs
DCapos could significantly inhibit the proliferation of CD4+ or CD8+ T cells stimulated by mature DCs, indicating that DCapos are inhibitory cells. Various immunosuppressive factors have been proposed to be responsible for the inhibitory functions of DCs, such as TGF-beta, IL-10, IL-13, and NO
Some studies have shown that DCs that phagocytose apoptotic cells could induce the differentiation of naive T cells into Tregs, especially into Foxp3+ Tregs, which contribute to the suppression of immune responses and immune tolerance
We put NO at the center of the immunosuppression mediated by apoptotic cells. While NO secretion by DC depends on iNOS, iNOS activation in the DCs after phagocytosing the apoptotic cells remains to be investigated. Previous studies showed that STAT3 is a cell intrinsic negative regulator of DC activity
The mechanisms of STAT3 signaling activation in DC after apoptotic cells pahgotosis remain to be elucidated. Some molecules displayed on the surface of apoptotic cells (AC), including phosphatidylserine, are recognized as “eat-me” signals by DCs. Research has indicated that both macrophages and DCs require MerTK binding of apoptotic cells to mediate phagocytosis through phosphatidylserine
In summary, we demonstrated that myeloid DCapos could directly inhibit the immune reponses via NO which is induced after activation of the STAT3 signaling pathway by phagocytosis activation-induced apoptotic CD4+ T cells. These findings shed more insights into the mechanism by which regulatory DCs negatively regulate antigen-specific immune responses. Targeting these molecules may provide an approach to induce or eliminate the tolerogenic DCs in the immunotherapy of autoimmune diseases.
We thank Jinwen Liu and Zhigang Yang for their excellent technical assistance; Zhijie Chang for reagents and Bin Wang for the Foxp3EGFP mouse. We are also grateful to Hua Tang for the helpful discussion.