Dendritic Cells Differentiated from Human Umbilical Cord Blood-Derived Monocytes Exhibit Tolerogenic Characteristics

Abstract

Human umbilical cord blood (UCB) is rich in diverse hematopoietic stem cells that are competent to differentiate into various cell types with immunological compatibility at transplantation. Thus, UCB is a potential source for the preparation of dendritic cells (DCs) to be used for cell therapy against inflammatory disorders or cancers. However, the immunological properties of UCB-derived DCs are not fully characterized. In this study, we investigated the phenotypes and functions of UCB monocyte-derived DCs (UCB-DCs) in comparison with those of adult peripheral blood (APB) monocyte-derived DCs (APB-DCs). UCB-DCs contained less CD1a⁺ DCs, which is known as immunostimulatory DCs, than APB-DCs. UCB-DCs exhibited lower expression of CD80, MHC proteins, and DC-SIGN, but higher endocytic activity, than APB-DCs. Lipopolysaccharide stimulation of UCB-DCs minimally augmented the expression of maturation markers and production of interleukin (IL)-12 and tumor necrosis factor (TNF)-α, but potently expressed IL-10. When UCB-DCs were cocultured with CD14⁺ cell-depleted allogeneic peripheral blood mononuclear cells, they weakly induced the proliferation, surface expression of activation markers, and interferon (IFN)-γ production of T lymphocytes compared with APB-DCs. UCB possessed higher levels of prostaglandin E2 (PGE2) than APB, which might be responsible for tolerogenic phenotypes and functions of UCB-DCs. Indeed, APB-DCs prepared in the presence of PGE2 exhibited CD1a⁻CD14⁺ phenotypes with tolerogenic properties, including weak maturation, impaired IL-12 production, and negligible T lymphocyte activation as UCB-DCs did. Taken together, we suggest that UCB-DCs have tolerogenic properties, which might be due to PGE2 highly sustained in UCB.

Introduction

Dendritic cells (DCs) are one of the best antigen-presenting cells to bridge innate and adaptive immunities. DCs play a pivotal role in the recognition of invading pathogens and malignant cells before the initiation of antigen-specific adaptive immune responses. Sensing microbial components or apoptotic cancer cells through various pattern recognition receptors, DCs upregulate the expression of costimulatory molecules (CD80, CD86, and CD40), MHC proteins, and cytokines [1]. Activated DCs modulate adaptive immune responses by (i) presenting MHC-associated antigens to naive T cells, (ii) providing costimulatory signals through CD80, CD86, and CD40, and (iii) creating a specific local environmental milieu by producing various cytokines such as interleukin (IL)-12, IL-6, and IL-10 [2]. Through these cellular processes, DCs mediate antigen-specific immunity or tolerance. Possessing diverse immunoregulatory roles and high plasticity, DCs have been considered as a promising cell therapeutic for the treatment of currently incurable diseases, including cancers [3], autoimmune diseases [4], and graft versus host disease (GvHD) [5].

Autologous DCs from cancer patients have been tried as anti-cancer immunotherapy. However, low immunogenicity of the DCs often hinders successful treatment [6]. On the other hand, cell therapy using DCs from unrelated healthy donors may exacerbate allogeneic immune rejection as they can activate alloreactive T lymphocytes by crosspresentation of allogeneic antigens [7]. As DCs in the transplanted tissues highly express endogenous antigens combined with MHC proteins and are capable of providing costimulatory signals to recipient T lymphocytes [5], they are usually considered to be potent initiators of allogeneic immune rejection [8]. Umbilical cord blood (UCB) is a promising source of stem cells for transplantation because it shows a low incidence of GvHD [9] and contains hematopoietic progenitor cells that are competent to differentiate into various immune cells [9]. In addition, the convenience of accessibility and human leukocyte antigen (HLA) matching increases the potential for use of UCB in the transplantation. However, its application is limited to only children since the absolute number of hematopoietic stem cells in a unit of UCB is too low to be applied to adults [9]. Previous studies have demonstrated that UCB-DCs display a low level of costimulatory molecules, low responsiveness to microbial components, and low lymphocyte-activating capacities [10,11]. These tolerogenic properties of UCB-DCs lead to alleviation of immune rejection after the transplantation [12].

DCs occupy a small proportion (1%–3%) in the peripheral blood and consist of myeloid and plasmacytoid DCs [13]. Since the number of DCs is restricted, blood monocytes, which can differentiate into various DC types, are commonly used in the preparation of DCs [14]. Monocyte-derived DCs are useful models for in vitro studies, because they share many immunological properties with those of blood DCs, despite differences in the specific phenotypes and functions [15,16]. Monocytes consist of two populations, CD14⁺CD16⁻ classical monocytes and CD14⁺CD16⁺ nonclassical monocytes [17], which differentiate into distinct DC subsets [18,19]. The proportion of CD14⁺CD16⁺ monocytes is significantly increased in the blood of patients with systemic inflammatory diseases such as bacterial sepsis [20]. UCB and adult peripheral blood (APB) exhibit difference in the ratio of CD14⁺CD16⁺ monocytes to CD14⁺CD16⁻ monocytes. UCB displays significantly reduced frequency in CD14⁺CD16⁺ monocytes [21]. However, it is yet to be elucidated whether the lower proportions of CD14⁺CD16⁺ monocytes in UCB have a correlation with the tolerogenic immune responses of UCB-DCs and what factors are associated with the distinctive immunological features of UCB-DCs. Therefore, in the present study, we investigated immunological characteristics of UCB-DCs in comparison with those of APB-DCs with respect to their differentiation, activation, and functions.

Materials and Methods

Reagents and chemicals

Ultra-pure lipopolysaccharide (LPS) from Escherichia coli serotype K12 was purchased from InvivoGen (San Diego, CA). Ficoll-Paque PLUS was from GE Healthcare (Uppsala, Sweden). Fetal bovine serum (FBS) and RPMI 1640 media containing 2.05 mM l-glutamine were obtained from HyClone (Logan, UT). The antibiotic–antimycotic solution containing penicillin, streptomycin, and amphotericin B was purchased from Invitrogen (Grand Island, NY). Human recombinant granulocyte-macrophage colony-stimulating factor (GM-CSF) and IL-4 were obtained from PeproTech (Rocky Hill, NJ) and R&D Systems (Minneapolis, MN), respectively. Antibodies used for flow cytometric analysis were as follows: PE-labeled anti-human CD80 antibody (clone L307.4), APC-labeled anti-human CD83 antibody (clone HB15e), FITC-labeled anti-human CD86 antibody [clone 2331 (FUN-1)], PE/Cy5-labeled anti-HLA-A, B, C antibody (clone G46-2.6) for MHC class I, FITC-labeled anti-HLA-DR, DP, DQ antibody (clone Tu39) for MHC class II, AlexaFluor^® 647-labeled anti-human CD205 antibody (clone MMRI-7), and FITC-labeled anti-human CD206 antibody (clone 19.2) were purchased from BD Biosciences (San Jose, CA). PE-labeled anti-human CD1a antibody (clone HI149), APC-labeled anti-human CD14 antibody (clone HCD14), APC-labeled anti-human CD86 antibody (clone IT2.2), PE-labeled anti-human DC-SIGN antibody (clone 9E9A8), APC-labeled anti-human programmed death ligand (PD-L) 1 antibody (clone 29E.2A3), APC-labeled anti-human DC-SIGN antibody (clone 9E9A8), PE-labeled anti-human CD16 antibody (clone 3G8), APC-labeled anti-human CD3 antibody (clone HIT3a), and APC-labeled CD25 antibody (clone 3C7) were purchased from BioLegend (San Diego, CA). R-PE-conjugated anti-EP2 antibody, R-PE-conjugated anti-EP4 antibody, and R-PE-conjugated rabbit IgG negative control antibody were purchased from Cayman Chemical (Ann Arbor, MI). FITC-conjugated dextran (dextran-FITC, MW 40,000) and 5-(and-6)-carboxyfluorescein diacetate succinimidyl ester (CFDA-SE/CFSE) were purchased from Molecular Probes (Eugene, OR). Difco™ Luria-Bertani (LB) broth was purchased from BD Biosciences (Sparks, MD).

Preparation of human monocyte-derived DCs

APB and UCB were provided from the Red Cross and from Allcord, the public cord blood bank of Seoul Metropolitan Boramae Medical Center (Seoul, Korea), respectively. All experiments using human blood were performed under the approval of the Institutional Review Board of the Seoul National University (IRB No. S-D20060001). Both APB and UCB cells were prepared using the same anticoagulant, citrate phosphate dextrose adenine-1. APB and UCB were overlaid on Ficoll, and mononuclear cells (MNCs) were obtained by density gradient centrifugation. CD14⁺ monocytes were isolated using I Mag™ anti-human CD14 particles-DM (BD Biosciences) followed by magnetic separation. The isolated CD14⁺ cells were suspended in RPMI 1640 supplemented with 2.05 mM l-glutamine, 10% FBS, and 1% antibiotic–antimycotic solution (100 U/mL penicillin, 100 μg/mL streptomycin, and 0.25 μg/mL amphotericin B) at a density of 2 × 10⁶ cells/mL, and 3 mL of the cell suspension was seeded in each well of six-well cell culture plates. To differentiate monocytes into immature DCs, the cells were stimulated with human recombinant GM-CSF (800 U/mL) and IL-4 (500 U/mL) for 6 days by fresh addition every 3 days. The DCs (5 × 10⁵ cells/mL) were stimulated with E. coli LPS (100 ng/mL) for 24–48 h in the presence of GM-CSF and IL-4 to induce the maturation.

Phenotypic analysis

Monocytes and DCs were washed with phosphate-buffered saline (PBS) once and then stained with fluorescence-conjugated monoclonal antibodies for 30 min on ice. The antibodies specific for human antigens that were used in the present study were CD80, CD83, CD86, MHC class I, MHC class II, CD14, CD1a, CD16, DC-SIGN, CD205, and CD206. Then the cells were washed three times with PBS containing 2% FBS and subjected to flow cytometric analysis. Ten thousand cells were acquired for each sample, and dead cells or debris were gated out. All flow cytometric data were analyzed using FACSCalibur (BD Biosciences) and FlowJo software (Tree star, San Carlos, CA).

Determination of endocytic capacity

Endocytic capacity of monocytes, unstimulated DCs, and LPS-stimulated DCs was measured using a dextran-FITC uptake assay. Briefly, the cells were washed with PBS and suspended in complete RPMI 1640 containing dextran-FITC (1 mg/mL) and incubated for 1 h at 4°C or 37°C to measure nonspecific or specific uptake, respectively. Then, the cells were washed three times with ice-cold PBS containing 2% FBS, and the geometric mean fluorescence intensity (MFI) of the cells was analyzed by flow cytometry as described above. The actual uptake (net MFI) was calculated by subtracting the MFI at 4°C from the MFI at 37°C.

Preparation of heat-killed E. coli

E. coli BL21 (DE3) was obtained from Stratagene (La Jolla, CA). Bacteria were cultured in LB broth media at 37°C until mid-log phase and harvested by centrifugation. The bacterial cells were suspended in PBS and killed by heating at 60°C for 1 h. To confirm complete killing, heat-treated bacteria were plated onto LB agar plates and cultured overnight at 37°C. No bacterial colony was observed (data not shown).

Quantification of cytokines

DCs (5 × 10⁵ cells/mL) were stimulated with E. coli LPS (100 ng/mL) for 48 h, and the amount of IL-12p40, tumor necrosis factor (TNF)-α, and IL-10 in the cell culture supernatant was measured by enzyme-linked immunosorbent assay (ELISA) (R&D Systems). DCs were stimulated with heat-killed E. coli (HKEC) (1 × 10⁶ CFU/mL) for 24 h, and the concentration of IL-12p70 was analyzed by ELISA (BioLegend). All procedures were conducted according to the manufacturer's instructions.

Analysis of T lymphocyte proliferation and activation

DCs (2.5 × 10⁵ cells/mL) were treated with HKEC (1 × 10⁶ CFU/mL) for 24 h and washed with PBS. Unstimulated and HKEC-stimulated DCs (5 × 10⁴ cells) were cocultured with CFSE-labeled allogeneic peripheral blood MNCs (PBMC, 1 × 10⁶ cells) for 3–5 days. On day 3, the culture supernatant was collected and subjected to ELISA for measuring the amount of interferon (IFN)-γ and IL-10 (R&D Systems). On day 5, CFSE-labeled PBMCs were stained with anti-human CD3 antibodies to detect T lymphocytes among the PBMCs, and proliferation of CD3⁺ cells and CD3⁻ cells was analyzed by flow cytometry. To examine the expression of activation markers on T lymphocytes, PBMCs were stained with anti-human CD3 antibodies, anti-human CD25 antibodies, and anti-human HLA-DR, DP, DQ antibodies, followed by flow cytometry as described above.

Prostaglandin E2 quantification

To obtain plasma, heparinized UCB and APB were centrifuged at 1,320× g at 4°C for 10 min. The plasma was used to determine PGE2 levels using a PGE2 Competitive ELISA Kit from Arbor Assays (Ann Arbor, MI) according to the manufacturer's instruction.

Statistical analysis

Experimental data were compared using Student's t-test or ANOVA test (Tukey's multiple comparison test). P values under 0.05 were considered statistically significant. Results are indicated as mean ± standard error of the mean (SEM).

Results

UCB-derived monocytes successfully differentiate into immature DCs

Enrichment of pure and functional monocytes is crucial for successful differentiation into DCs. In this study, we examined purity of UCB- and APB-derived CD14⁺ monocytes before and after the magnetic separation. Flow cytometric analyses showed that UCB exhibited significantly lower frequencies of CD14⁺ cells than APB (Fig. 1A). After magnetic separation, the enriched CD14⁺ cells from both sources displayed 96%–99% purities on average, but the expression of CD14 on the UCB-derived monocytes was lower than that of APB-derived monocytes (Fig. 1B). When the monocytes differentiated into immature DCs, UCB-DCs displayed similar morphologies to those of APB-DCs (Fig. 1C, F). In addition, both UCB- and APB-DCs showed significant changes in the phenotypes and endocytic abilities upon differentiation. Although UCB-DCs exhibited CD14⁺ and CD1a⁻ phenotypes, unlike CD14⁻ and CD1a⁺ APB-DCs, more than 88% of the cells expressed DC-SIGN and PD-L1, which are other indicative markers for DCs (Fig. 1D, G). Furthermore, UCB-DCs and APB-DCs displayed enhanced endocytic abilities compared to their precursors (Fig. 1E, H). These results suggest that highly pure monocytes isolated from UCB successfully differentiated into immature DCs.

FIG. 1.

Umbilical cord blood (UCB)-derived monocytes successfully differentiate into immature dendritic cells (DCs). Mononuclear cells (MNCs) were isolated from UCB and adult peripheral blood (APB), respectively, by density gradient centrifugation. (A) Frequencies of CD14⁺ cells in UCB- and APB-derived MNCs were analyzed by flow cytometry. Percentages of the CD14⁺ cells in MNCs from both sources were shown in a scatter plot (right panel). (B) After magnetic separation, the expression of CD14 on UCB- and APB-derived monocytes was analyzed by flow cytometry. Percentage (middle panel) and mean fluorescence intensity (MFI) (right panel) of CD14⁺ cells in the enriched monocytes were shown in a scatter plot. (C–H) The enriched CD14⁺ monocytes (2 × 10⁶ cells/mL) were differentiated into immature DCs in the presence of granulocyte-macrophage colony-stimulating factor (GM-CSF) (800 U/mL) and interleukin (IL)-4 (500 U/mL) for 6 days. (C, F) Morphologies of the monocytes and immature DCs were observed by optical microscopy (40× magnification). A scale bar in each picture indicates 30 μm. (D, G) The expression of CD14, CD1a, DC-SIGN, and PD-L1 on the monocytes and immature DCs was analyzed by flow cytometry. (E, H) Monocytes and immature DCs were suspended in 50 μL of phosphate-buffered saline (PBS) containing dextran-FITC (1 mg/mL) for 1 h at 4°C (non-specific uptake) and 37°C (specific uptake). Endocytic activity of the monocytes and DCs was examined by flow cytometric analysis. The vertical axis indicates the net MFI values of the DCs, obtained by subtracting MFI at 4°C from MFI at 37°C. Statistical differences between the two experimental groups were analyzed by Student's t-test. The results are shown as mean ± standard error of the mean (SEM). N.S., not significant.

Immature UCB-DCs display distinctive features from immature APB-DCs in phenotypes and endocytic capacity

Next, we comparatively examined the expression of phenotypic markers and endocytic activity of immature DCs differentiated from UCB- or APB-derived monocytes. Cell populations gated for the phenotypic and functional analyses were presented in the upper panel of Fig. 2A. When the monocytes were differentiated into immature DCs, UCB-DCs displayed less CD1a⁺ cells, but more CD14⁺ cells than APB-DCs (Fig. 2A, lower panel, B). In addition, UCB-DCs showed significantly lower levels of CD80, MHC class I and II, and DC-SIGN than APB-DCs, whereas the expression of CD86, CD205, and CD206 was not significantly different from those of APB-DCs (Fig. 2C–I). UCB-DCs displayed better endocytic activity than APB-DCs (Fig. 2J). These results indicate that immature UCB-DCs exhibited lower expression of CD1a, CD80, MHC proteins, and DC-SIGN, but higher CD14 expression and endocytic capacity than immature APB-DCs.

FIG. 2.

UCB-DCs display distinctive features from APB-DCs in phenotypes and endocytic capacity. CD14⁺ monocytes (2 × 10⁶ cells/mL) isolated from UCB and APB were differentiated into DCs in the presence of human GM-CSF (800 U/mL) and IL-4 (500 U/mL) for 6 days. (A) Dead cells or debris were gated out by forward scatter and side scatter (upper panel). The expression of CD14 and CD1a on DCs was analyzed by flow cytometry (lower panel). (B) Percentages of CD14⁺ cells (left) or CD1a⁺ cells (right) on UCB-DCs and APB-DCs were shown in scatter plots. Expression of (C) CD80, (D) CD86, (E) MHC class I, (F) MHC class II, (G) DC-SIGN, (H) CD205, and (I) CD206 on the DCs were analyzed by flow cytometry and MFI of the cells was shown in scatter plots. (J) The cells (1 × 10⁵ cells) were suspended in 50 μL of PBS containing dextran-FITC (1 mg/mL) for 1 h at 4°C (nonspecific uptake) and 37°C (specific uptake). Endocytic activity of the DCs was examined by flow cytometric analysis. The vertical axis indicates net MFI values of the DCs subtracting MFI at 4°C from MFI at 37°C. Statistical differences between the two experimental groups were analyzed by Student's t-test. The results are shown as mean ± SEM.

LPS weakly induces the maturation of UCB-DCs and their expression of inflammatory cytokines in comparison with those of APB-DCs

Since maturation of DCs is an important process in the mediation of immune responses, we examined the phenotypic changes and cytokine expression of DCs. Stimulation with LPS hardly or slightly induced costimulatory molecules, including CD80, CD83, and CD86 and MHC class I and II on UCB-DCs, whereas APB-DCs exhibited a remarkable increase in the expression of these molecules (Fig. 3A, B). UCB-DCs still sustained high endocytic activity even after LPS stimulation, while LPS-matured APB-DCs showed remarkably decreased endocytic activity (Fig. 3C). The expression of phagocytosis-related receptors (CD205 and CD206) was minimally changed in LPS-treated UCB-DCs (Fig. 3D). UCB-DCs produced significantly lower IL-12p40 and TNF-α, but higher IL-10 in response to LPS than APB-DCs did (Fig. 3E). HKEC-stimulated UCB-DCs hardly induced IL-12p70 in comparison with HKEC-stimulated APB-DCs (Fig. 3F). Taken together, these results indicate that UCB-DCs weakly elicit phenotypic and functional maturation in response to LPS or HKEC.

FIG. 3.

LPS weakly induces the maturation of UCB-DCs and their expression of inflammatory cytokines in comparison with those of APB-DCs. (A, B) DCs (5 × 10⁵ cells/mL) were stimulated with LPS (100 ng/mL) for 48 h. Expression of DC maturation markers (CD80, CD83, CD86, and MHC class I and II) on the cell surface was analyzed by flow cytometry. (C) Unstimulated and LPS-stimulated DCs (1 × 10⁵ cells) were suspended in PBS containing dextran-FITC (1 mg/mL, 50 μL) for 1 h at 4°C and 37°C and subjected to flow cytometric analysis. The vertical axis indicates the net MFI values of the DCs subtracting MFI at 4°C from MFI at 37°C. (D) The expression of phagocytosis-related receptors (CD205 and CD206) was analyzed by flow cytometry. (E) DCs (5 × 10⁵ cells/mL) were stimulated with LPS (100 ng/mL) for 48 h, and the amount of IL-12p40, TNF-α, and IL-10 in the culture supernatant was measured by enzyme-linked immunosorbent assay (ELISA). (F) The DCs (2.5 × 10⁵ cells/mL) were stimulated with heat-killed E. coli (HKEC) (1 × 10⁶ CFU/mL), and the amount of IL-12p70 in the culture supernatant was determined by ELISA. The statistical differences among the experimental groups were analyzed by ANOVA or Student's t-test. P values under 0.05 were considered statistically significant. The results are shown as mean ± SEM.

UCB-DCs weakly induce proliferation and activation of allogeneic T lymphocytes

One of the major functions of DCs is to stimulate naive T lymphocytes to differentiate into various effector T cells [22]. To examine T lymphocyte-activating capacities of DCs, unstimulated or HKEC-stimulated DCs were cocultured with CD14⁺ cell-depleted allogeneic PBMCs. HKEC-stimulated UCB-DCs induced less proliferation of CD3⁺ T lymphocytes in PBMCs than those of APB-DCs (Fig. 4A, B). Interestingly, APB-DCs stimulated with HKEC markedly augmented the proliferation of CD3⁻ cells. Given the results that most of the proliferated CD3⁻ cells expressed CD56 (data not shown), APB-DCs appear to preferentially induce proliferation and activation of CD56⁺ natural killer cells besides CD3⁺ T lymphocytes. Concomitantly, UCB-DCs poorly induced CD25⁺ MHC class II⁺ T lymphocytes, whereas APB-DCs increased those cell populations (Fig. 4C, D). In addition, HKEC-stimulated UCB-DCs weakly induced IFN-γ production (Fig. 4E), whereas that of HKEC-stimulated APB-DCs by PBMCs was potently induced. However, the level of IL-10 in the supernatants of PBMCs cocultured with HKEC-stimulated DCs from both sources was not significantly different (Fig. 4F) although slightly more potent IL-10-producing capacities were observed in UCB-DCs than in APB-DCs (Fig. 3E). These results indicate that UCB-DCs weakly induced activation of allogeneic T lymphocytes in comparison with APB-DCs.

FIG. 4.

UCB-DCs weakly induce proliferation and activation of T lymphocytes. DCs (5 × 10⁴ cells) were stimulated with HKEC (1 × 10⁶ CFU/mL) for 24 h, and the allogeneic peripheral blood MNCs (PBMCs) (5 × 10⁵ cells) were cocultured with the stimulated DCs for 3–5 days. (A) Cells were stained with anti-human CD3 antibodies to detect T lymphocytes from the PBMCs. Proliferation of T lymphocytes among the PBMCs was analyzed by the CFSE dilution assay followed by flow cytometric analysis. Numbers on the histograms indicate the percentage of cells on each quadrant. (B) DC-mediated proliferation of CD3⁺ T lymphocytes was shown in a scatter plot. (C) Expression of T lymphocyte activation markers (CD25 and MHC class II) on CD3⁺ cells was analyzed by flow cytometry. The number in each quadrant indicates the percentage of cells. (D) DC-mediated induction of CD25 on CD3⁺ T lymphocytes was presented in a scatter plot. (E, F) The amount of interferon (IFN)-γ and IL-10 in the supernatant of PBMCs cocultured with UCB-DCs or APB-DCs was measured by ELISA. Statistical differences among the experimental groups were analyzed by ANOVA. P values under 0.05 were considered statistically significant. The results are shown as mean ± SEM.

Reduced frequency of CD14⁺CD16⁺ monocytes in UCB correlates with increased level of blood PGE2

CD14⁺CD16⁻ and CD14⁺CD16⁺ monocytes exhibit different immunological properties when they differentiate into DCs [19]. To unravel the mechanism responsible for the distinct phenotypes and functions of UCB-DCs, monocyte subtypes of UCB and APB were analyzed. Frequency of CD14⁺CD16⁺ monocytes was significantly lower in UCB than in APB (Fig. 5A). PGE2 is a potent immunomodulator that regulates differentiation and activation of various immune cells and is known to exist at a high level in UCB [23]. Thus, we measured PGE2 concentration in the plasma of UCB and APB and examined whether it was related to the frequencies of CD14⁺CD16⁺ monocytes. Higher amounts of PGE2 were detected in the plasma of UCB than that of APB (Fig. 5B). It was ensured that PGE2 was not additionally synthesized during the plasma preparation process even in the absence of cyclooxygenase inhibitors (data not shown). The PGE2 level of UCB plasma was inversely correlated with the percentage of CD14⁺CD16⁺ monocytes (R ² = 0.8) (Fig. 5C), but a significant correlation between two parameters was not observed in APB (data not shown). To examine whether PGE2 is directly involved in the regulation of CD14⁺CD16⁺ monocyte differentiation, CD14⁺ monocytes isolated from APB were cultured with or without PGE2. An increase of CD14⁺CD16⁺ cells was observed in the monocytes cultured in the absence of PGE2 (Fig. 5D), which is concordant with the previous report [24]. In contrast, CD14⁺CD16⁺ cells were less augmented in the monocytes cultured in the presence of PGE2 (Fig. 5D). Furthermore, CD14⁺ monocytes enriched from UCB-derived MNCs showed lower levels of E-prostanoid (EP) 2 and EP4 than those of APB (Fig. 5E, F). These results suggest that the reduced frequency of CD14⁺ CD16⁺ monocyte in the UCB is associated with the levels of PGE2 in the plasma.

FIG. 5.

Reduced frequencies of CD14⁺CD16⁺ monocytes in UCB correlate with increased level of blood prostaglandin E2 (PGE2). (A) MNCs were isolated from UCB and APB, and the subtypes of monocytes were analyzed by flow cytometry. The scatter plot shows frequencies of CD14⁺CD16⁺ monocytes in the UCB and APB. (B) The amount of PGE2 in the plasma of UCB and APB was quantified by ELISA. Statistical differences were analyzed by Student's t-test. The results are presented as mean ± SEM. (C) The correlation between plasma PGE2 levels and frequencies of the CD14⁺CD16⁺ monocytes in UCB was analyzed. (D) Monocytes isolated from APB were cultured in RPMI 1640 containing 10% fetal bovine serum (FBS) with or without PGE2 (1 μM) for 24 h. Changes in the expression of CD16 on CD14⁺ monocytes were analyzed by flow cytometric analysis. A scatter plot shows the changes in the frequency of CD14⁺CD16⁺ cells upon treatment with PGE2 (right panel). (E, F) The expression of EP2 and EP4 on CD14⁺ monocytes enriched from UCB- and APB-derived MNCs was presented in scatter plots. Statistical differences were analyzed by Student's t-test.

PGE2 induces differentiation of APB monocytes into UCB-DC-like cells

To further investigate the role of PGE2 in phenotypic and functional characteristics of DCs, APB monocytes were differentiated into immature DCs in the absence (control-DCs) or presence of PGE2 (PGE2-DCs). PGE2-DCs preferentially expressed CD14 rather than CD1a, whereas control-DCs expressed CD1a, but not CD14 (Fig. 6A). PGE2-DCs displayed higher endocytic activity than the control-DCs (Fig. 6B). Unlike control-DCs, PGE2-DCs weakly augmented the expression of costimulatory molecules in response to LPS (Fig. 6C). PGE2-DCs expressed low IL-12p70, but remarkably high IL-10 upon stimulation with LPS. In contrast, the LPS-treated control-DCs efficiently upregulated IL-12p70, but weakly induced IL-10 (Fig. 6D). HKEC-stimulated PGE2-DCs poorly induced allogeneic T lymphocyte proliferation when compared to that of control-DCs (Fig. 6E). Furthermore, less IFN-γ, but more IL-10 was detected in the allogeneic PBMCs cocultured with PGE2-DCs than in PBMCs cocultured with control-DCs (Fig. 6F). These results show that monocytes exposed to PGE2 differentiate into DCs similar to UCB-DCs.

FIG. 6.

PGE2 induces differentiation of monocytes into UCB-DC-like cells. Monocytes (2 × 10⁶ cells) isolated from APB were differentiated into immature DCs in the absence or presence of exogenous PGE2 (0.1, 1, 10, or 100 nM) for 6 days. (A) The expression of CD14 and CD1a on the DCs was analyzed by flow cytometry. The numbers on the histogram indicate the percentage of cells in each quadrant. (B) The endocytic activity of the DCs was examined by dextran-FITC uptake assay followed by flow cytometric analysis. DCs (2.5 × 10⁵ cells/mL) were stimulated with LPS (100 ng/mL) for 24 h. (C) The expression of CD83 and CD86 on DCs treated with or without LPS was analyzed by flow cytometry. (D) The amount of IL-12p70 and IL-10 in the culture supernatant was measured by ELISA. DCs (5 × 10⁴ cells) were stimulated with HKEC (1 × 10⁶ CFU/mL) for 24 h, and the unstimulated or HKEC-stimulated DCs were cocultured with allogeneic PBMC (5 × 10⁵ cells) for 3–5 days. (E) Proliferation of T lymphocytes in the PBMCs was analyzed by flow cytometry. (F) The amount of IFN-γ and IL-10 in the culture supernatant was measured by ELISA. The results shown are representative of three independent experiments.

Discussion

In the present study, we investigated immunological characteristics of UCB-DCs. UCB-DCs exhibited unique phenotypes and functions as distinct from those of APB-DCs. UCB-DCs expressed lower CD1a, CD80, MHC proteins, and DC-SIGN than APB-DCs and minimally induced maturation markers and inflammatory cytokines. Moreover, UCB-DCs elicited weaker T cell proliferation and activation than APB-DCs. All of these results are in keeping with those of other groups [23,25,26] and are also coincident with the immunological characteristics of in vitro-generated tolerogenic DCs [27 –29]. The phenotypes and functions of UCB-DCs seem to have relevance to PGE2 level in the plasma in that (i) the UCB monocytes were exposed to a higher level of PGE2 than the APB monocytes, (ii) the higher level of PGE2 was correlated with reduced frequency of CD14⁺CD16⁺ monocytes in UCB, and (iii) APB-DCs differentiated in the presence of PGE2 acquired similar phenotypes and functions with those of UCB-DCs.

CD1a⁺ DC is a typical immunostimulatory DC subtype that produces a high level of IL-12 and has a potent lymphocyte-activating capacity, while CD1a⁻ DC displays tolerogenic functions [30 –32]. Particularly, a high endocytic ability is a distinctive property of CD1a⁻ DCs and it more efficiently takes up dextran, lipid molecules, and formalin-fixed bacteria than CD1a⁺ DCs [32]. In this study, we found that UCB-DCs mainly consisted of CD1a⁻ DCs and APB-DCs prepared in the presence of PGE2 showed the decreased CD1a expression. In addition, UCB-DCs containing high number of CD1a⁻ cells exhibited higher endocytic capacity than APB-DCs. Furthermore, the APB-DCs differentiated in the presence of PGE2 showed potentiated endocytic ability as well compared to control-DCs differentiated in the absence of PGE2. Thus, the enhanced endocytic ability seems to be a common feature of CD1a⁻ DCs, which were rich in UCB-DCs.

UCB-DCs weakly expressed the maturation markers and inflammatory cytokines in response to LPS or HKEC that are crucial for the activation of subsequent adaptive immunity. These results are in agreement with the previous report that UCB-DCs showed lower expression level of costimulatory molecules, MHC proteins, and IL-12p70 in response to LPS or dying cells, including apoptotic cells and necrotic cells [26]. Therefore, UCB-DCs might be less potent than APB-DCs in the induction of adaptive immunity due to inadequate provision of costimulatory signals and immunostimulatory cytokines to T lymphocytes necessary for the generation of effector T lymphocytes. Indeed, we observed that UCB-DCs weakly induced the proliferation and activation of T lymphocytes at the coculture of UCB-DCs and allogeneic PBMCs. All these properties of UCB-DCs seem to be associated with those of CD1a⁻ DCs, in light of the fact that CD1a⁻ DCs even in response to LPS displayed a decrease in the expression of CD83, CD86, and IL-12p70 and activation of T lymphocytes [32,33].

Differentiation into CD1a⁺ DCs or CD1a⁻ DCs appears to be differently regulated depending on the types of precursors and microenvironment. For example, unlike our findings, some of the previous studies have demonstrated that CD14⁺CD16⁺ monocytes differentiate into CD1a⁻ DCs exhibiting low immunostimulatory potencies, compared to CD1a⁺ DCs differentiated from CD14⁺CD16⁻ monocytes [18,19]. In addition, monocytes consisting of mostly CD14⁺CD16⁻ cells could differentiate into CD1a⁻ DCs upon exposure to inflammatory mediators such as microbial molecular patterns [34] and phospholipid metabolites, including PGE2 [29,32]. In keeping with the previous evidence, we have also shown that monocytes stimulated with PGE2 differentiate into CD1a⁻ DCs. However, under the present experimental condition, PGE2 was not able to induce CD14⁺CD16⁺ monocytes as the precursor of CD1a⁻ DCs, because PGE2 treatment suppressed the in vitro culture-induced CD16 on the CD14⁺CD16⁻ monocytes. Specific mechanisms by which PGE2 modulates immunological characteristics of DCs should be further studied.

PGE2 levels are highly sustained in the human placenta and UCB to maintain pregnancy and help in uterine contractions during parturition [35]. Considering that PGE2 levels soar transiently in both maternal and fetal circulation during periparturition [36], exposure of immune cells and their precursors to high amounts of PGE2 seems to be inevitable. Cells exposed to PGE2 have been reported to desensitize and downregulate the expression of EP receptors [37,38]. In the present study, we found that UCB-derived monocytes expressed significantly lower levels of EP2 and EP4 than APB-derived monocytes, which may be due to PGE2-rich environment of UCB. Additionally, it was observed that the precursors of UCB-DCs were exposed to high concentrations of PGE2, which would enhance tolerogenic properties during differentiation. Remarkably, PGE2 is likely to differentially regulate immune responses of DCs depending on the differentiation stage of the DCs. At the early phase of differentiation, PGE2 suppresses differentiation of monocytes into inflammatory DCs, inducing Th1 cell responses [39]. In contrast, the presence of PGE2 at the late phase of differentiation promotes final maturation of DCs [40]. Therefore, exposure of DC precursors to PGE2 at an early differentiation stage might be critical in the determination of immunological properties of UCB-DCs.

UCB contained a lower number of CD14⁺CD16⁺ monocytes, which are known as activated monocytes than APB. Concomitant with our findings, reduction of inflammatory CD14⁺CD16⁺ monocytes in UCB and its implications for weak immune responses of newborns to infections has been demonstrated [21]. In this study, we observed that PGE2 level in the UCB plasma has inverse correlation with the frequencies of CD14⁺CD16⁺ cells. Indeed, treatment with PGE2 suppressed the differentiation of CD14⁺CD16⁻ monocytes into CD14⁺CD16⁺ monocytes. In addition, PGE2 has been previously reported to suppress activation and differentiation of T and B lymphocytes [41,42]. Taken together, these data suggest that sustaining high PGE2 levels in UCB might be a plausible strategy for protecting the immunocompromised neonates from excessive inflammation. Conclusively, UCB-DCs would be worthy of being developed into immune cell therapeutics against chronic inflammatory disorders, including GvHD and autoimmune diseases.

Footnotes

Acknowledgments

This work was supported by grants from the National Research Foundation of Korea funded by the Korean government (MISIP) (2010-0029116) and the Korea Health Technology R&D Project through the Korea Health Industry Development Institute (KHIDI), funded by the Ministry of Health & Welfare (HI14C0469), Republic of Korea. Umbilical cord blood was kindly provided from Allcord, SMG-SNU Boramae Medical Center.

Author Disclosure Statement

No competing financial interests exist.

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Dendritic Cells Differentiated from Human Umbilical Cord Blood-Derived Monocytes Exhibit Tolerogenic Characteristics

Abstract

Introduction

Materials and Methods

Reagents and chemicals

Preparation of human monocyte-derived DCs

Phenotypic analysis

Determination of endocytic capacity

Preparation of heat-killed E. coli

Quantification of cytokines

Analysis of T lymphocyte proliferation and activation

Prostaglandin E2 quantification

Statistical analysis

Results

UCB-derived monocytes successfully differentiate into immature DCs

Immature UCB-DCs display distinctive features from immature APB-DCs in phenotypes and endocytic capacity

LPS weakly induces the maturation of UCB-DCs and their expression of inflammatory cytokines in comparison with those of APB-DCs

UCB-DCs weakly induce proliferation and activation of allogeneic T lymphocytes

Reduced frequency of CD14+CD16+ monocytes in UCB correlates with increased level of blood PGE2

PGE2 induces differentiation of APB monocytes into UCB-DC-like cells

Discussion

Footnotes

Acknowledgments

Author Disclosure Statement

References

Reduced frequency of CD14⁺CD16⁺ monocytes in UCB correlates with increased level of blood PGE2