MSC‐sEV inhibited the differentiation of CD14⁺ monocytes into DCs

To demonstrate the effects of MSC‐sEV on the differentiation of CD14⁺ monocytes, iDCs and sEV‐immature DCs (sEV‐iDCs) were generated as stated in the absence or presence of MSC‐sEV for 5 days. MSC‐sEV altered the phenotype of iDCs by reducing the expression of cell surface markers. The gating strategies are shown in Supporting Information Fig. S2. sEV‐iDCs expressed a significantly lower level of CD11c, HLA‐DR, CD40, CD80, and CD86 compared to their iDCs counterpart in different dosages (Fig. 2 and Supporting Information Fig. S3A). Representative pictures are also shown in Supporting Information Fig. S3A. These results were consistent with the potency of their parent cells as our previous report [5]. However, no differences were observed regarding the expression of CD14, CD1a, and CD141 (Supporting Information Fig. S3B). Both iDCs and sEV‐iDCs exhibited low expression of CD14, suggesting that MSC‐sEV suppressed the differentiation of iDCs rather than blocking DC differentiation from monocytes at the beginning.

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Figure 2

MSC‐sEV inhibited the differentiation of CD14⁺ monocytes into DCs. Sorted CD14⁺ monocytes were cultured in the presence or absence of MSC‐sEV (5, 10 or 20 μg/mL) for 5 days. (A) DC markers of CD11c, HLA‐DR, CD40, CD80, and CD86 were determined by flow cytometry. MFI values for different conditions were indicated top left in the figures. (B) Mean fluorescence intensity of the markers. The total amount of donors was n = 10, as human blood buffy coats from “anonymous donors.” Data are presented as mean ± SEM, and are from three independent experiments with three to four donors per experiment. Significance was determined comparing all sEV‐iDC samples to iDC without MSC‐sEV treatment. *p < 0.05, **p < 0.01, and ***p < 0.001. Statistical comparisons were performed using Tukey's multiple comparisons test. DCs: dendritic cells; FMO: fluorescence minus one, MSC: mesenchymal stromal cells, sEV: small extracellular vesicles.

MSC‐sEV promote the mDC phagocytic ability but do not affect the expression of maturation markers

To further confirm the effects of MSC‐sEV on DC maturation, we induced the generation of sEV‐mDCs in vitro and determined the expression of DC surface markers. However, we did not find any differences in the phenotypic maturation between mDCs and sEV‐mDCs (Fig. 3A and B), which is consistent with our previous study about the effects of MSCs on DCs [5]. Here, sEV‐mDCs acquired a typical phenotype of mature DCs, by expressing CD11c, HLA‐DR, CD40, CD80, and CD86, regardless different concentrations of the MSC‐sEV added. These findings suggested that MSC‐sEV did not affect the expression of maturation markers from iDCs to mDCs.

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Figure 3

MSC‐sEV promoted the phagocytic ability of mDCs but did not affect the maturation surface markers. iDCs were treated with LPS in the presence or absence of MSC‐sEV for an additional 2 days. (A) DC markers of CD11c, HLA‐DR, CD40, CD80, and CD816 were determined by flow cytometry. MFI values for different conditions were indicated top left in the figures. (B) Bar graphs of the (MFI) of the markers. (C) Phagocytic ability of the iDCs, mDCs, and sEV‐mDCs was analyzed by flow cytometry, and the black line presents the expression of FITC‐Dextran at 4°. (D) Bar graph of the ΔMFI (37‐4°) of FITC‐dextran. Data are representative of collated data of four donors (A, B) or three donors (C, D) (mean + SEM). Statistical comparisons were performed using Tukey's multiple comparisons test. iDCs: immature dendritic cells, LPS: lipopolysaccharide, MFI: mean fluorescence intensity, mDCs: mature dendritic cells, MSC: mesenchymal stromal cells, sEV: small extracellular vesicles.

The capacity of DCs to take up and process antigens is highly dependent on the stage of differentiation of DCs. Notably, iDCs have more strong ability to phagocytose dextran compared to mDCs. Therefore, we conducted the phagocytosis assay to study the influence of MSC‐sEV on the phagocytic ability of mDCs. As expected, iDC‐D7, the iDCs without the treatment of lipopolysaccharide (LPS) were harvested on day 7, was more powerful in the uptake of FITC‐dextran than mDCs. Furthermore, MSC‐sEV improved the ability of mDCs to take up FITC‐dextran but without significant difference (Fig. 3C and D, p = 0.0579). Generally, it indicated that sEV‐mDC acquired a capability of immune tolerance similar to iDCs but without any change in the phenotype.

Additionally, to test the lymphocyte‐stimulating ability of sEV‐mDCs, we conducted a mixed lymphocyte culture (MLC) system. sEV‐mDCs induced T‐cell proliferation in parallel to mDCs (Supporting Information Fig. S4), suggesting that MSC‐sEV had no effects on the ability of mDCs on T‐cell proliferation.

MSC‐sEV impair the priming of mDCs on type 2 immune response from patients with AR in vitro

To investigate whether MSC‐sEV affect the function of mDCs, human CD4⁺ T cells were isolated and co‐cultured with allogeneic sEV‐mDCs for 5 days. As we did not observe any dose‐dependent differences of MSC‐sEV on the maturational markers of DCs, we used the lowest dosage of MSC‐sEV with 5 μg/mL in the following functional experiments. We found that T cells produced lower IL‐13 but not IL‐9 and IFN‐γ when co‐cultured with sEV‐mDC compared with those co‐cultured with mDCs (Fig. 4A). We further observed a significant decrease in the percentage of IL‐13⁺CD4⁺ T cells and IL‐9⁺CD4⁺ T cells after co‐cultured with sEV‐mDCs compared to their mDC counterpart (Fig. 4B, Fig. S2B). Recently, IL‐9‐producing T cells were considered as a new T helper (Th) cell subset, instead of a subgroup of Th2 cells, namely Th9 cells [23]. In addition, IL‐9 was reported as the most enriched gene in allergen‐specific T cells from patients with asthma [24]. Taken together, these findings suggested that the treatment of MSC‐sEV to mDCs impaired their type 2 immunity enhancing ability, but did not affect the development of type 1 immune response in vitro. Moreover, MSC‐sEV might have the potential to suppress the function of Th9 cells in parallel.

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Figure 4

MSC‐sEV impaired the type 2 immunity priming capacity of mDCs in patients with AR in vitro. sEV‐mDCs were co‐cultured with sorted CD4⁺ T cells from buffy coat (A and B) or PBMCs from patients with AR (C and D) for 3 days. (A) The levels of IL‐13, IL‐9, and IFN‐γ in the supernatants of the co‐cultures. (B) The percentage of intracellular IL‐13, IL‐9, and IFN‐γ in T cells after co‐cultures with conditioned DCs (n = 6). (C and D) The percentages of IL‐13⁺ T cells, IL‐4⁺ T cells, and IL‐9⁺ T cells (n = 5). Data are representative of collated data of six donors for T cells isolation and three donors for DC generation, all from buffy coat (A, B), or five donors from patients with AR for PBMC and three donors for DC generation from buffy coat (C, D) (mean + SEM). *p < 0.05 and **p < 0.01, Tukey's multiple comparisons test. mDCs: mature dendritic cells, MSC: mesenchymal stromal cells, sEV: small extracellular vesicles.

mDCs were recognized to present the allergens to the naïve T cells and induce them into Th2 cells or Th9 cells in the pathogenesis of allergic diseases [25, 26]. To further examine the function of sEV‐mDCs, human PBMCs from patients with AR were co‐cultured with allogeneic sEV‐mDCs. Similar results were found that the levels of intracellular IL‐13 and IL‐4 in CD4⁺ T cells but not IL‐9⁺CD4⁺ T cells were decreased after the treatment of sEV‐mDCs (Fig. 4C and D). Overall, these findings suggest that MSC‐sEV could mitigate the function of mDCs in promoting Th2 cells from patients with AR in vitro.

IL‐10 is responsible for the immunomodulatory effects of sEV‐mDCs

DCs can exert their function on T cells by the secretion of a series of soluble inflammatory cytokines. Therefore, to address the potential mechanism underlying the immunomodulatory effects of sEV‐mDCs, we examined the mRNA levels of the main functional cytokines after the treatment of MSC‐sEV. We observed an increased mRNA level of IL‐10 in sEV‐mDC compared to mDCs (Fig. 5A). Meanwhile, no difference was observed in the mRNA levels of IL‐12p70, TNF‐α, IL‐1β, and OX‐40L (encoded by tnfsf4) (Supporting Information Fig. S5). The production of IL‐10 is a certain characteristic of the regulatory DCs. Then we first identified that there was higher level of IL‐10 in the supernatants of sEV‐mDCs co‐cultured with T cells compared to those of mDCs (Fig. 5B). To confirm the activity of DCs to produce IL‐10, we next examined the intracellular level of IL‐10 in DCs in the co‐culture system. We found that there was higher level of IL‐10⁺mDCs after T cells were co‐cultured with sEV‐mDCs but not mDCs, but without significant difference (p = 0.0811, Fig. 5C and D), which indicated that MSC‐sEV promoted mDCs to get the function of regulatory DCs, and induced the production of IL‐10 in sEV‐mDCs. We then checked the effects of rhIL‐10 on the activation of mDCs on CD4⁺ T cells. We found that rhIL‐10 significantly reversed the activate effects of mDCs on T cells as inhibiting the production of IL‐13 and IL‐9 (Fig. 5E and Supporting Information Fig. S6A). Taken together, these data suggest that MSC‐sEV treatment promoted the production of IL‐10 from mDCs, resulting in the suppression of type 2 immune response.

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Figure 5

IL‐10 was responsible for the immunomodulatory effects of sEV‐mDCs. (A) The relative expression of IL‐10 mRNA in mDCs treated with or without MSC‐sEV. (B) The level of IL‐10 in the supernatants of DC‐T cell co‐cultures. (C and D) The level of intracellular IL‐10 in mDCs after co‐cultured with sorted CD4⁺ T cells from buffy coat. (E) The levels IL‐13⁺CD4⁺T cells and IL‐9⁺CD4⁺T cells after treated with rhIL‐10 (10 ng/mL). (F) The levels of intracellular IL‐9 and IL‐13 in T cells in the addition of anti‐IL‐10 antibody to the co‐cultures. (G) The levels of IL‐9 and IL‐13 in the supernatants of co‐cultures with the administration of anti‐IL‐10 antibody. Data are representative of collated data of nine donors (A), or eleven donors for T cell isolation and seven donors for DC generation (B–G), all from buffy coat (mean ± SEM). *p < 0.05, **p < 0.01, and ****p < 0.0001, Wilcoxon matched‐pairs signed‐rank test (A), paired t‐test (D) and Tukey's multiple comparisons test (B, E, F, and G). mDCs: mature dendritic cells; MSC: mesenchymal stromal cells; sEV: small extracellular vesicles.

To further confirm the function of IL‐10 secreted by sEV‐mDCs in inhibiting Th2 immunity, we conducted the co‐culture experiments of T cells with conditioned DCs by the addition of the IL‐10‐neutralizing antibodies, and the levels of IL‐13 and IL‐9 were examined. As expected, the treatment of neutralizing anti‐IL‐10 antibodies led to a significant reversal in levels of IL‐13⁺CD4⁺T cells (Fig. 5F), and IL‐9 and IL‐13 protein compared to the sEV‐mDCs alone (Fig. 5G). Moreover, by blocking IL‐10Rα using blocking antibody, the suppressive effects of sEV‐mDC on the levels of IL‐13⁺CD4% but not IL‐9⁺CD4% were significantly reversed (Supporting Information Fig. S6B). Generally, these data demonstrated the inhibitory effects of sEV‐mDCs on the type 2 responses partially by acting on the IL‐10/IL‐10Rα axis, further indicating the potential application of the cell‐free therapy of MSC‐sEV in the treatment of allergic rhinitis.

MSC‐sEV preparation

The MSC‐sEV‐enriched medium in this study was prepared as described in our previous study [21]. iPSC‐MSCs were plated at a density of 1 × 10⁶ cells/dish in 150 mm diameter cell culture dishes with complete culture medium. After the cells reached 70%–80% confluency (about 1 × 10⁸ cells in total), the complete culture medium was replaced with chemically defined and protein‐free medium after three washes with phosphate‐buffered saline (PBS). Chemically defined and protein‐free medium was discarded and then refreshed after 6 h of incubation. After another 42‐h incubation, the sEV‐enriched supernatants were harvested and centrifuged immediately at 2000 g for 20 min at 4°C to exclude detached cells and debris.

Isolation of MSC‐sEV was conducted using anion exchange chromatography as our previous report [21]. An Econo‐Pac column (Bio‐Rad Laboratories, CD63 Hercules, CA, USA) was packed with anion exchange resin (GE Health Care Life Science, Pittsburgh, PA, USA) and equilibrated with equilibration buffer. Then the column was loaded with MSC supernatants, washed with wash buffer to remove the proteomic impurities, and eluted continuously with elution buffer for eight times. The fractions with peak concentration of MSC‐sEV were pooled for dialysis overnight at 4°C. Then, we further concentrated the MSC‐sEV using Pierce™ Protein Concentrator (Thermo Fisher Scientific, Rockford, IL, USA), which were used for further analyses and studies. Collected MSC‐sEV were characterized based on their morphology, particle size, and surface protein expression. The final protein and particle concentrations of MSC‐sEV preparations were quantified by Bradford protein assay and nanoparticle tracking analysis (NanoSight NS300; Malvern, UK).

Transmission electron microscopy for MSC‐sEV

The MSC‐sEV preparations were fixed with 2% glutaraldehyde (Sigma, Saint Louis, MO, USA) for 30 min. The exact procedures could be referred in our previous paper [21, 32]. Micrographs of MSC‐sEV were obtained with a transmission electron microscopy (H7650; HITACHI, Tokyo, Japan) at Guangdong Institute of Microbiology.

Western blot analysis

The protein concentration of MSC‐sEV and MSC lysates were quantified using the Pierce™ BCA protein assay (Thermo Fisher Scientific, Rockford, IL, USA). Western blot was performed as described previously [21, 32]. CD9 (1:2000), CD63 (1:2000), CD81 (1:2000), Alix (1:5000), TSG101 (1:1000), and Calnexin (1:1000) were used as primary antibodies, all from Abcam, Cambridge, UK. All secondary antibodies were purchased from Jackon ImmunoResearch, West Grove, PA, USA. Finally, the figures were obtained using an imaging analysis system (ImageQuant LAS 4000, GE Health Care Life Science, Uppsala, Sweden).

Generation of sEV‐immature DCs and sEV‐mature DCs in vitro

Like our previous study, CD14⁺ monocytes from PBMCs in the buffy coat “anonymous donors” provided by Guangzhou Blood Center were isolated using the MACS CD14 MicroBeads (Miltenyi Biotec, Bergisch Gladbach, Germany) [5, 33]. And then the CD14⁺ monocytes were stimulated with recombinant human granulocyte‐macrophage colony‐stimulating factor (GM‐CSF; 50 ng/mL; PeproTech Inc., Rocky Hill, NJ, USA) and IL‐4 (10 ng/mL; R&D Systems, Minneapolis, MN, USA) with or without the administration of MSC‐sEV (5 μg/mL, 10 μg/mL or 20 μg/mL) for 5 days to get iDCs or sEV‐iDCs. On the other hand, the iDCs on day 5 were then stimulated with LPS (100 ng/mL; Sigma‐Aldrich, St Louis, MO, USA) and MSC‐sEV (5–20 μg/mL) for an additional 2 days to be sEV‐mDCs. And the iDCs without the treatment of LPS were harvested on day 7 as iDC‐D7. The DCs were used in the examination of the surface markers by flow cytometry or co‐cultures.

Phagocytosis assay

To compare the phagocytic ability of DCs, iDC‐D7, mDCs, and sEV‐mDCs were incubated with FITC‐conjugated dextran (mol wt 4000, 100 μg/mL; Sigma‐Aldrich) for 1 h at 37°C, or at 4°C as a negative control. The cells were then washed twice and re‐suspended in ice‐cold PBS for immediate analysis by flow cytometry. The results were shown as mean MFI_(37‐4°C) ± SEM.

Co‐culture experiments of sEV‐mDCs with PBMCs or T cells

Human PBMCs from patients with AR or buffy coat from “anonymous donors” were isolated by density centrifugation with Ficoll‐Paque Plus (MP Biomedicals, Santa Ana, CA, USA). CD4⁺ T cells were sorted from PBMCs of buffy coat of human volunteers using the CD4 Microbeads (Miltenyi Biotec, Bergisch Gladbach, Germany). sEV‐mDCs were harvested and washed twice, and then were co‐cultured with allogeneic PBMCs isolated from patients with AR or freshly sorted CD4⁺ T cells (1:10) for 5 days in RPMI 1640 (Hyclone, Pittsburgh, PA, USA) supplemented with 10% FBS and IL‐2 (100 U/mL). Cell counting was performed manually using a hemocytometer and the cell viability was over 95%. In some experiments, CD4⁺ T cells were treated with the rhIL‐10 (10 ng/mL; PeproTech Inc.) together with mDC, or incubated with anti‐IL‐10 monoclonal antibody (75 ng/mL) or IL‐10Rα blocking antibody (5 μg/mL; both from R&D systems, Minneapolis, MN, USA) for 30 min before the co‐culture. The PBMCs or the T cells were collected for the analysis of intracellular Th2 cytokines using flow cytometry, and the supernatants were used for the evaluation of IL‐13, IL‐9, IL‐10, or IFN‐γ. T cells were also evaluated for IL‐10⁺Foxp3⁺ T cells or Treg cells.

T‐cell proliferation

For the analysis of cell proliferation, freshly sorted CD4⁺ T cells stained with carboxyfluorescein diacetate succinimidyl diester (Invitrogen/Molecular Probes, Eugene, Ore) were cocultured with mDCs or sEV‐mDCs for 5 days, and the proliferation rate was evaluated on a CytoFLEX S flow cytometer. The data were analyzed with CytExpert (Beckman Coulter).

Flow cytometry

DCs were stained with the following specific mAbs: FITC‐conjugated lineage cocktail: anti‐CD2 (RPA‐2.10), anti‐CD3 (OKT3), anti‐CD14 (61D3), anti‐CD16 (CB16), anti‐CD19 (HIB19), anti‐CD56 (TULY56), anti‐CD235a (HIR2), eFluor 450‐conjugated anti‐CD45 (HI30), PE‐conjugated anti‐CD11c (BU15), APC‐conjugated anti‐CD40 (5C3), FITC‐conjugated anti‐CD80 (2D10.4), PE‐Cyanine7‐conjugated anti‐CD86 (IT2.2, all from eBioscience, San Diego, CA, USA), and APC‐H7‐conjugated anti‐HLA‐DR (G46‐6; BD Pharmingen, Bergen, NJ, USA). T cells and Treg cells were assessed using the antibodies of Fixable Viability Dye eFluor™ 506, PerCP‐Cyanine5.5‐conjugated anti‐CD4 (RPA‐T4), PE‐Cyanine7‐conjugated anti‐CD25 (BC96; all from eBioscience). Detailed gating strategies are shown in Supporting Information Fig. S2.

For intracellular cytokine detection, PBMCs or T cells were stimulated with the cell stimulation cocktail consisting of phorbol 12‐myristate 13‐acetate (50 ng/mL), ionomycin (1000 ng/mL; both from Sigma), and monensin (1:1000; eBioscience) for 5 h prior the staining. Thereafter, the cells were fixed, permeabilized with intracellular fixation and permeabilization buffer set (eBioscience), and stained for intracellular IL‐13 (V450‐conjugated anti‐IL‐13; JES10‐5A2), IL‐9 (Alexa Fluor 647‐conjugated anti‐IL‐9; MH9A3; both from BD Pharmingen, Bergen, NJ, USA), and IL‐4 (APC‐conjugated anti‐4; 8D4‐8; eBioscience). And the intracellular IL‐10 in DCs or T cells was determined using the antibody of PE‐Dazzle594‐conjugated anti‐IL‐10 (JES3‐9D7, BioLegend, San Diego, CA, USA).

T cells were processed with the FoxP3/Transcription Factor Staining Buffer Set (eBioscience) before staining with the antibody of PE‐conjugated anti‐Foxp3 (PCH101; eBioscience) for the evaluation of Treg cells. Flow cytometry were performed and presented, adhered to the guidelines [34].

ELISA

The cell supernatants were analyzed by using IL‐13, IL‐9, and IL‐10 ELISA kits (Invitrogen, Waltham, MA, USA), and IFN‐γ ELISA kits (RayBiotech, Guangzhou, Guangdong, China), according to the manufacturer's instruction.

RT‐qPCR

Total RNA was isolated from mDCs with RNAiso Plus reagent and the 5X PrimeScript™ RT Master Mix kit (all from TaKaRa, Shiga, Kusatsu, Japan) was used for converting 1 μg of total RNA to the first‐strand cDNA following the manufacturer's instructions. The quantitative PCR of IL‐10 (sense primers, 5′‐GACTTTAAGGGTTACCTGGGTTG‐3′, and reverse primer, 5′‐TCACATGCGCCTTGATGTCTG‐3′) were performed using the FastStart Universal SYBR Green Master kit (Roche, Mannheim, Germany). β‐Actin (sense primers, 5′‐AGAGCTACGAGCTGCCTGAC‐3′, and reverse primer, 5′‐ AGCACTGTGTTGGCGTACAG‐3′) was used as an endogenous reference. The PCR was performed as 10 min initial denaturation at 95°C, 40 cycles consisted of 10 s at 95°C and 30 s at 60°C carried out on the CFX96™ Real‐Time PCR cycler (Bio‐Rad, Hercules, CA, USA). The expressions of the target genes were expressed as fold increase relative to the expression of β‐actin. The mean value of the replicates for each sample was calculated and expressed as cycle threshold (CT). The amount of gene expression was then calculated as the difference (ΔCT) between the Ct value of the target genes and the Ct value of β‐actin. Fold changes in target genes mRNA were determined as 2^−ΔCT.

Statistical analyses

Differences between two groups were analyzed by the paired t‐test for the data with normal distribution, and the Wilcoxon matched‐pairs signed‐rank test was performed to compare the data with abnormal distribution. Three or more groups were compared using one‐way analysis of variance (ANOVA) with Tukey's multiple comparisons test. Kolmogorov–Smirnov test was used to test the normality of all the data. p < 0.05 was considered statistically significant. Analyses were performed using GraphPad Prism 8.0 software (GraphPad Software, La Jolla, CA, USA).

We sincerely thank Guangzhou Blood Center for kindly providing us with buffy coats of human volunteers. Also, we want to give heartfelt thanks to the nurses in Otorhinolaryngology Hospital, The First Affiliated Hospital, Sun Yat‐sen University for collecting the blood samples. This study was supported by grants from National Natural Science Foundation of China (81770984 and 81970863).