Continuous retinoic acid induces the differentiation of mature regulatory monocytes but fails to induce regulatory dendritic cells
© VanGundy et al.; licensee BioMed Central Ltd. 2014
Received: 18 July 2013
Accepted: 6 January 2014
Published: 18 February 2014
Myeloid cells (MC) have potent immunoregulatory abilities that can be therapeutically useful to treat inflammatory disease. However, the factors which promote regulatory myeloid cell differentiation remain poorly understood. We have previously shown that estriol (E3) induces mature regulatory dendritic cells in vivo. To determine whether additional steroid hormones could induce mature regulatory myeloid cells, we investigated the effects of retinoic acid (RA) on MCs. Retinoic acid is a steroid hormone important in regulating mucosal immunity in the gut and promoting myeloid differentiation. We hypothesized that the presence of RA during differentiation would promote the formation of mature regulatory myeloid cells (MCregs).
To determine RA’s ability to induce regulatory myeloid cells, we differentiated bone marrow progenitor cells with granulocytic-macrophage colony-stimulating factor (GM-CSF) under the influence of RA. We found that day 7 MCs differentiated in the presence of RA had an increase in the percent positive and relative expression levels of both maturation (CD80, CD86, and MHCII) and inhibitory (PD-L1 and PD-L2) markers compared to control cells. Functionally, these day 7 RA MCs expressed increased intracellular IL-10, induced regulatory T cells in vitro compared to controls and suppressed the proliferation of responder immune cells even after inflammatory challenge with LPS.
RA induced mature regulatory myeloid cells that were suppressive and had a CD11b+ CD11c-Ly6C low/intermediate monocyte phenotype. Surprisingly, RA CD11c+ dendritic cells were not suppressive and could contribute to enhanced proliferation. These results suggest that continuous RA has unique effects on different myeloid populations during monopoeisis and dendropoiesis and promotes a population of regulatory monocytes.
KeywordsRegulatory myeloid cells Dendritic cells Retinoic acid Monocyte
Myeloid cells (MCs) are a diverse population of cells that form during hematopoiesis and play a critical role in host defense. Comprised of granulocytes, mononuclear phagocytes and their precursors, MCs are innate immune cells that have an important role in promoting inflammation and the induction of adaptive immune responses. Inflammatory MCs are induced and increased in numbers following exposure to exogenous (e.g. pathogens) or endogenous “danger” (e.g. post-necrotic release of high-mobility group box 1 or HMGB1) signals b . These and other environmental factors present within peripheral tissue and bone marrow impact granulopoiesis, monopoiesis and dendropoiesis to influence the ultimate fate of inflammatory granulocytes, monocytes/macrophages and DCs, respectively. While inflammatory MCs have been well characterized, within the last several years the potent regulatory abilities of these cells has increasingly been recognized. Such regulatory MCs (MCregs) are a diverse population of cells with the ability to control inflammation and, thus, are a promising target to treat a wide array of inflammatory diseases. To date, however, factors involved in the differentiation of MCreg populations remain poorly understood.
MCreg subsets are particularly diverse, both in terminology and in function. Regulatory, tolerogenic, type II or steady-state are terms applied to regulatory populations of DCs, macrophages, monocytes, and their precursors [1–3]. By and large, the regulatory abilities of macrophages, and more recently, DCs have been most thoroughly studied. First described over 30 years ago, alternatively activated macrophages are able to promote wound healing and resolve inflammation [4, 5]. Over the last 10 years the regulatory abilities of DCs and their therapeutic potential have been the focus of many studies [2, 6]. Monocytes are circulating myeloid cells that give rise to tissue macrophages and DCs. Monocytes have been recognized as a contributor to the inflammatory responses, and are now known to contribute to immune regulation . MCregs can regulate immune responses through the production of soluble regulatory factors (e.g. IL-10, TGF-beta, indoleamine 2,3 deoxygenase (IDO), arginase, nitric oxide (NO), etc.), expression of inhibitory or regulatory cell surface molecules (e.g. PD-L1, PD-L2) and induction other regulatory cells (e.g. regulatory T cells; Tregs) or enhance regulatory feed-back loops [8, 9]. At present, MCregs are identified based on combination of phenotype and function, with no equivalent to Treg FoxP3 marker being as yet identified [10–13]. Through cell-cell interactions and the production of soluble immunoregulatory molecules, MCregs have very potent and diverse means of inducing immune regulation. However, much remains to be characterized about factors controlling MCreg induction and how different MCreg subsets regulate immune responses. Given that MCreg therapy has the potential to diminish disease in the 100+ millions of individuals impacted by immune-mediated, chronic inflammatory and autoimmune diseases worldwide, it is critical to determine the factors which govern the induction and function of these cells [14–16]. The therapeutic potential of MCregs, has been described in several experimental models of inflammatory and autoimmune disease. Specifically, MCregs, including MDSC, conventional DCs, lung-resident tissue macrophages, monocytes, and plasmacytoid DCs have all been shown to impact disease course in animal models of diabetes , colitis , allergic asthma , experimental autoimmune disease , and rheumatoid arthritis  respectively.
For many MCregs, an arrest in immature and/or altered functionality contributes to their regulatory abilities [22, 23]. Glucocorticoids, vitamin D and IL-10 are the most common means to induce these immature MCregs. These altered MCregs cells have decreased expression levels of maturation/activation markers CD80, CD86 and MHC class II [2, 24–28]. Additionally, these immature MCregs can have reduced inflammatory cytokine expression [29, 30], overall blunted function, induce Tregs and suppress the action of other immune cells. However, a primary concern with using immature MCregs for therapy is that they may mature into inflammatory MCs under inflammatory disease conditions. Such inflammatory MCs could then actually exacerbate the very inflammatory disease they were used to treat [2, 22, 23, 31]. Thus, mature (and stable) MCregs may avoid such concerns but, to date only a handful of studies have significantly explored the induction of such mature MCregs[18, 22, 32]. Typically, mature MCregs have been induced by combining traditional immature MCreg induction protocols with the addition of inflammatory stimuli such as LPS or TNF-alpha [33, 34]. Our laboratory has focused on identifying non-inflammatory systems to induce mature MCregs and we have previously found that estriol (E3), a steroid hormone of pregnancy, produce mature activated DCregs. These E3 DCregs maintained their regulatory abilities within an inflammatory environment and protected mice against the inflammatory autoimmune disease, experimental autoimmune encephalomyelitis (EAE) . Although E3 shows promise, the fact that there are limitations on using estrogens broadly in the human patient population necessitated investigating alternative means of inducing mature stable MCreg populations.
All-trans retinoic acid (RA) is a steroid hormone metabolite of vitamin A that plays both an important role during embryonic development and has recently been identified as the key metabolite regulating immune responses at mucosal sites [36–38]. RA is a logical candidate for inducing mature MCregs given its defined role in both mucosal immunoregulation and its ability to promote myeloid cell differentiation and maturation. Within the gut, RA influences the balance between Tregs and Th17 cells, B cell isotype switching, antibody production and mucosal homing of numerous immune cells [6, 37, 39–43]. Mucosal myeloid cells are largely responsible for producing local RA which acts in a paracrine and autocrine manner to regulate mucosal immune responses [6, 37]. Although mucosal DCs produce much of the RA required for immune regulation at mucosal sites, much less is known about RA’s direct impact on MC populations at both mucosal and non-mucosal sites [9, 19, 39, 40].
RA regulates myeloid cell survival and promotes the differentiation of immature myeloid cells into mature populations of DCs, macrophages and granulocytes [18, 44–46]. Additionally, RA appears to be required for the production of mature phagocytes in the bone marrow through its effects on MHC class II and co-stimulatory molecule expression . Therapeutically, RA has long been used to treat myeloid leukemia given that it promotes myeloid cell differentiation and maturation [48, 49]. More recently, it has been used to promote the differentiation of immature myeloid cells (i.e. myeloid derived suppressor cells; MDSCs) in cancer patients to diminish immunosuppressive MDSC effects [36, 44, 50–53]. Given RA’s important roles in both mucosal immunoregulation and myeloid cell differentiation we hypothesized that RA would induce mature MCregs.
Using an in vitro model to induce differentiation of MC populations (i.e. DCs, macrophages and monocytes), we evaluated the ability of RA to generate mature MCregs[42, 54]. We demonstrated that bone marrow cells differentiated with GM-CSF for 7 days in the presence of RA had an activated regulatory phenotype (i.e. increased CD80, CD86, MHC class II, PD-L1 and PD-L2), produced increased IL-10, increased the induction of Treg and suppressed the proliferation of responder immune cells. We found that the suppressive population was a small but potent CD11b+ CD11c- Ly6Clow/intermediate population whose phenotype is consistent with a regulatory monocyte. Surprisingly the CD11c+ DCs were not suppressive. Taken together these results demonstrate a differential effect of RA during monopoiesis and dendropoiesis which results in the induction of regulatory monocytes but not regulatory DCs.
Differentiation with retinoic acid induced mature activated regulatory myeloid cells
Given that increased IL-10 is seen in E3 DCregs and other MCreg populations [50, 55] we next evaluated whether RA induced an increase number of IL-10+ cells. Figure 1C shows that RA differentiated cells had an increased percentage of IL-10-producing cells compared to either media or E3 control cells. We next evaluated whether RA differentiated cells could increase Treg numbers. We found that RA differentiated cells were able to induce a significant increased percentage of FoxP3+ cells following a 5 day culture with naïve immune cells (Figure 1D). Cells differentiated in vitro in the presence of E3 failed to significantly increase either IL-10+ cells or induce Treg cells (Figures 1C, D). These results show that RA differentiated cells suppressed the proliferative abilities of responder immune cells and induced FoxP3+ (Treg) cells.
CD11b+ but not CD11c+ cells were the suppressive population
CD11b+ CD11c-Ly6Clow/intermediate were the primary population responsible for suppression
The principal objective of this study was to determine whether RA, a steroid hormone known to play important roles in regulating both mucosal immune responses and differentiation of myeloid cells could generate an activated (or mature) MCreg population. We demonstrate that RA influences myelopoiesis to a regulatory MCreg (monocyte) with the phenotype of CD11b+ CD11c-Ly-6Clow/intermediate but fails to induce DCregs. These cells can influence both CD4+ and CD8+ responses and promote FoxP3+ (Treg) cell induction. Our data suggest that RA has distinctly different effects on monopoiesis and dendropoiesis to promote the generation of regulatory monocytes.
MCregs are a diverse population of cells and much attention has focused on the in vitro generation and clinical application of MCregs. While the in vitro generation of such MCreg populations has great therapeutic potential, much remains to be learned regarding the factors which contribute to MCreg induction. The majority of in vitro generated MCregs are arrested in an immature or hypo-functional state. An emerging concern is that these immature MCregs populations may mature to become inflammatory DCs or macrophages and, thus, contribute to inflammatory disease pathology [2, 22, 23, 29–31, 64]. A more recent approach is to induce mature MCregs which would be stable and maintain regulatory potential in an inflammatory environment [22, 32, 65, 66]. Anderson and colleagues have demonstrated that human DCregs (generated with dexamethasone, vitamin D and LPS) maintain tolerogenic activity and actually induce significantly higher levels of IL-10 production by resultant T cells . However, the relative stability and ability of MCregs (such as DCregs) to maintain regulatory abilities during inflammation may still be in question. For example, a study by Voigtlander et al. suggests that DCregs induced by TNF-alpha do not maintain their regulatory abilities upon a secondary stimulation with TNF-alpha in vivo. Obviously, this is of considerable concern given that TNF-alpha is present in a large array of inflammatory conditions where such DCregs (or other MCreg populations) may be applied therapeutically. Much work remains to determine critical factors important in generating mature MCregs for anti-inflammatory therapies but we have focused on non-inflammatory pathways to induce mature MCregs.
We have shown that mature MCregs can be generated with the use of steroid hormones alone . Our previous work has shown that the sex steroid hormone estriol (E3) induces a mature activated MCreg population of CD11c+ DCregs that protects against inflammatory challenge in vitro and in an in vivo disease model . In the present study, we have extended our research of pathways involved in normal homeostatic induction of mature MCregs by investigating the ability of the steroid hormone RA to induce mature MCregs that are resistant to inflammatory challenge. Our results show that RA is more effective than E3 in vitro in generating MCregs and that these MCregs are resistant to LPS inflammatory challenge.
RA is known for its ability to promote the differentiation, and maturation, of myeloid cell populations. This ability, along with its known immunoregulatory role at mucosal sites, made it a logical candidate for these studies [44, 52]. RA is present in relatively large concentrations within mucosal sites and is largely produced by local antigen presenting cells (APCs) residing within these mucosal sites. Specifically, mucosal CD103+ DCs are the primary immunoregulatory myeloid cells within the gut. These DCs have up-regulated raldh2 gene expression, constitutively produce RA, and produce increased TGF-beta. They also have a significant ability to induce Foxp3+ Tregs, mucosal homing receptors CCR9 and α4β7 expression on lymphocytes and enhance antibody production and Ig isotype switching [6, 9, 36, 57]. These mucosal DCs are the most common MCs investigated regarding RA biology and induced mucosal DCs have been generated from monocytes or splenic DCs with GM-CSF with IL-4 [8, 43] or bone marrow precursors with RA [18, 41, 57, 67, 68]. Increasingly, the non-mucosal and therapeutic applications of RA (i.e. in cancer) are being investigated [9, 19, 43, 44, 53] and this study focused on RA’s ability to induce mature activated MCregs that are able to suppress responder immune cell proliferation [8, 35, 41, 43, 57, 69].
Given RA’s critical role in DC-mediated immunoregulation within the gut, it was quite surprising that RA CD11c+ cells were not suppressive. One possibility is that DC’s differentiated with RA could generate mucosal DCs but wouldn’t generate mature activated DCregs that could suppress proliferation as seen with E3 DCregs. While induction of mucosal DCs can be accomplished with RA [18, 43], the immunomodulatory abilities of these DCs as described in these studies was not the focus of this study. Alternatively, timing of RA administration may have resulted in the lack of DCreg induction as described by Feng and colleagues . Specifically, their studies showed that the presence of 1 μM RA from day 0 throughout differentiation failed to induce mucosal DCs. Although different dosages and criteria were used to generate and identify DCs as mucosal (versus DCregs in our study), the continuous presence of RA during differentiation may have resulted in the inability to induce DCregs in our study. Similarly, Wada’s group showed that the use of a synthetic RARα and β agonist (AM-80) could differentiate human peripheral blood monocytes into dendritic cells that have a tolerogenic phenotype and function . The use of AM-80 versus ATRA in our study or the differentiation of human monocytes versus murine myeloid progenitors could explain the differences in DCregs versus MCregs in our study.
It could be argued that CD11c- DC precursors existed within the population of CD11b+ CD11c- suppressive cells. Given the described effects of RA in promoting differentiation and maturation, in conjunction with our data demonstrating an activated phenotype, we believe this to be unlikely [57, 70–72]. Rather, our data on Ly6C expression strongly support that the suppressive cells were regulatory monocytes with an activated regulatory phenotype (increased CD80, CD86, MHC class II and PD-L1) consistent with previous work within our lab. Given that the CD11b+ CD11c- population is comprised less than 20% of the entire population, the ability of these cells to suppress both CD4+ and CD8+ responses is noteworthy. The specific contributions of cell contact-dependent (i.e. PD-L1) versus cell contact-independent (i.e. IL-10, TGF-beta, etc.) mechanisms responsible for the regulatory abilities of these cells was beyond the scope of this study. However, we did see increases in regulatory markers including PD-L1, IL-10 and the percentage of FoxP3+ cells with RA MCregs.
Monocytes are circulating myeloid cells which give rise to tissue DCs and macrophages, and their regulatory abilities have recently been recognized . Although numerous markers can be present on mouse monocytes (e.g. CD11b, CD115, CCR2, CX3CR1 and Ly-6C), we chose to investigate Ly6C expression levels given that they have been correlated with monocyte function [7, 63, 71, 73]. Specifically, Ly-6Chigh represents an inflammatory monocytes while, Ly-6Clow/intermediate monocytes have been shown to play important roles in patrolling the vasculature and potentially resolving inflammation and tissue repair [7, 63, 74–76]. Ly6C is also down regulated following differentiation which is consistent with our findings where RA, a molecule known to promote differentiation and maturation, increases the percentage of cells that are Ly-6Clow/intermediate (Figure 5A) [3, 44]. Our data suggest that Ly-6C levels correlate with suppressive abilities with the lowest Ly-6C expression associated with the most suppressive ability. Given that Ly-6Chigh monocytes are typically inflammatory monocytes, it is not surprising that proliferation is actually enhanced following LPS stimulation in this cell population (Figure 5C). Taken together, these data showed a progression from Ly-6Chigh to Ly-6Clow associated with increasing regulatory abilities. These results are consistent with the association seen between Ly-6C expression and blood monocyte function described by others [7, 63, 71, 77]. Currently, the mechanisms and pathways by which RA maturation of monocytes imparts them with increased regulatory abilities remain undefined. Whether a specific signal during differentiation drives monocytes to become regulatory in an active process or whether differentiation under homeostatic or regulatory (i.e. RA) conditions in the absence of inflammatory stimuli is a default mechanism for regulatory monocyte induction is unknown. Additionally, whether these RA Ly-6Clow/intermediate monocytes have the potential to further differentiate into DCreg or regulatory macrophage populations remains to be determined and is the subject of ongoing studies within the laboratory .
Our findings show that continuous RA exposure during myelopoiesis promotes the induction of MCregs. Specifically, RA induced CD11b+ CD11c-Ly-6Clow/intermediate regulatory monocytes which suppressed the proliferation of immune cells but RA failed to induce DCregs. Our data suggests that RA has unique effects on different myeloid populations during differentiation that may influence the regulatory abilities of monocytes and DCs. A more thorough understanding of how RA mediates these differential effects has important implications in our understanding of MCreg biology and the potential application of these cells to treat a wide variety of inflammatory diseases.
C57BL/6 (H-2b) mice (4–8 wk old), C57BL/6-Tg (TcraTcrb)425Cbn/J, C57BL/6-Tg(Tcra2D2,Tcrb2D2)1Kuch/J and reporter Foxp3EGFP (B6.Cg-Foxp3 tm2Tch) ) were purchased from Jackson Laboratories (Bar Harbor, ME, USA) or bred in-house. Mice were housed five per cage and maintained on a 12 hr. light/dark cycle, maintained under specific pathogen-free conditions and were housed and cared for according to the institutional guidelines of the Ohio State University’s Institute for Animal Care and Use Committee.
EG7 and EL7 (kindly provided by P. Boyaka, Ohio State University) were used to study the MHC class I-restricted response of CTLs in mice. The EG7 cells have been transfected with plasma to synthesize and constitutively secrete OVA 257–264 peptide and should be cultured in 10% RPMI. The EL4 cells are the non-OVA secreting duplicate of the EG7. Both are commonly found at ATCC but were acquired through Dr. Boyaka. The DC2.4 cell line was kindly provided by K. Rock, University of Massachessetts and as a DC antigen-presenting cell.
BM-MC differentiation and development of regulatory MC differentiation model
Bone marrow (BM) cells were collected from C57Bl/6 mice femurs and tibias. After erythrocyte lysis (AKC or in-house lysis buffer), cells were cultured with RPMI 1640 (Invitrogen) supplemented with 10% FBS, 25 mM HEPES, 2 mM L-glutamine, 50 U/ml penicillin, 50 mg/ml streptomycin, 5 × 10-5 M 2-mercaptoethanol and 200U/ml recombinant murine GM-CSF (R&D Systems) ± 100 nM of either estriol (E3) or all-trans retinoic acid (RA) (Sigma-Aldrich) for 6–7 days at a density of 2 × 106 cells/ml. Day 6–7 cells were considered differentiated BM-MCs (media control) and BM-MCregs(RA and E3). Cells were challenged with inflammatory stimulus LPS (1 μg/ml, 055:B5, Sigma-Aldrich) during culture as indicated at day 6 or later for BM-DCs.
Functional immunosuppressive assays: T cell proliferation assay
Myeloid cells (BM-MCs or BM-MCregs) were cultured with responder spleen cells from antigen-specific T cell receptor transgenic (TCR Tg; where antigen was either OVA323-339 or MOG35-55) or Foxp3EGFP mice as indicated. To assess T cell proliferation co-cultures were stimulated with anti-CD3 (BD Bioscience), T cell-receptor specific antigen MOG35-55 (Bio Matic) or T cell-receptor specific antigen OVA 323–339 (Anaspec). To assess the effects of myeloid cell activation, co-cultures were stimulated with LPS from Escherichia coli, 055:B5 (Sigma-Aldrich) for 96 hours, pulsed with (H3 thymidine) (Perkin Elmer Life Sciences or MP Biomedicals) in the last 18 hours, harvested and counted, data is expressed as counts per million (cpm) ± SEM .
Functional immunosuppressive assay: CD8+ cytotoxic assay
To generate CTL, spleen and lymph nodes (LN) were removed from OT-1 mice and co-cultured with OVA (257–264) pulsed DC2.4 cells (kindly provided by Kenneth Rock, University of Massachussets) for 4 days, removed and cultured with mrIL-2 (R&D Systems) for 2 days. OVA-expressing (EG7) and non-transfected control cells (EL4) were seeded at 2 × 104 cells per well and co-cultured with CTLs (1 × 105) and control or RA treated monocytes (2 × 104) for 6–18 hours [73, 78]. The MTT assay (Sigma-Aldrich) was used to determine quantity of live cells. Briefly, after incubation, cells were centrifuged (1500 RPM for 5 min) media was decanted and 100 ul of fresh media was added. 10ul of 5 mg/ml thiazolyl blue tetrazolium bromide (MTT) was added to each well for 2 hours at 37°C. After incubation cells were centrifuged (1500 RPM for 7 min) and media was decanted, cells were allowed to dry for 15–30 min before 100 ul of DMSO was added, mixed well and read at 570 nm on a Spectra Max 2. The absorbance levels were calculated by averaging the non-specific and specific absorbance levels of five separate data sets. Media control is compared to RA treated cells.
In vitro Treg induction
Bone marrow (BM) cells were collected from C57Bl/6 mice femurs and tibias. After erythrocyte lysis, BM cells were cultured with supplemented RPMI (as previously described) for 7 days at a density of 2x106 cells/ml +/− RA. Spleens from mice with reporter Foxp3EGFP (B6.Cg-Foxp3 tm2Tch) ) were harvested, passed through cell strainers (70 μm, BD Falcon), collected by centrifugation (1500 RPM for 7 Min at 4°C) and subjected to erythrocyte lysis. Responder cells and MCs or CD11c- MCs were cultured for 4–6 days and aliquots from cultures assessed for Foxp3 expression by flow cytometry.
In vivo and in vitro derived DCs and MCs were labeled and evaluated by three-color flow cytometry using combination of the following conjugated directly antibodies (clone): CD11c (HL3), CD11b (M1/70), Gr-1 (RB6-8C5), Ly-6G (1A8), Ly-6C (AL-21), MHC class II (AF6-120.1), CD80 (16-10A1), CD86 (IT 2.2), PD-L1 (MIH5), and PD-L2 (YT25) with appropriate isotype controls. (BD Bioscience, eBiosciences or Miltenyi Biotec). Cells were stained with fluorochrome-labeled antibodies or isotype controls for 20 min in the dark at 4°C, washed twice in FACS buffer and re-suspended in 300 μl FACS buffer for flow cytometry analysis.
Intracellular IL-10, IL-4, IL-17 and INF-γ levels were measured after incubating myeloid cells with 1 μg/ml LPS overnight (IL-10) or Ionomyocin (1 mg/ml) and PMA (25 ng/ml) for 4 hours (IL-4, IL-17, and INF-γ). 2 μM of Monensin (eBioscience) added 2–4 hrs. before harvesting cells. Cells were removed from culture, washed with 2 ml of supplemented RPMI and blocked with 0.5 μg/ml Fc block (anti-CD16/CD32) for 15 minutes. Cells that required extracellular markers were re-suspended in FACS buffer and stained with anti-CD4 (0.2 mg/ml) and incubated in the dark at 4°C for 20 min. Cells were washed with FACS buffer (2x with 1 ml) and then fixed and permeabilized using FIX/PERM solution (BD Bioscience), briefly vortexed and incubated in the dark at 4°C for 20 min. Cells were then washed twice with 1 ml of PERM/WASH buffer (BD Bioscience), re-suspended in PERM/WASH buffer and stained with 0.2 mg/ml anti-IL-10 (BD Bioscience) for 30 min. in the dark at 4°C. All flow samples were processed on an Accuri C6 flow cytometer and results analyzed using the Accuri C6 Flow software (BD Biosciences).
Myeloid cell purification
Day 6–7 differentiated BM cells were incubated with manufacturer suggested amounts of CD11c/CD11b microbeads (Miltenyi Biotec) for 15 minutes in the dark at 4°C. Cells were washed with running buffer (10% FBS in PBS with 900 mg of NaN2 per 1 L of PBS), and centrifuged (1500 RPM, 7Min). Cell separation was performed using either the Auto Macs (Miltenyi Biotec) magnetic separation instrument or the FACS Aria III 12 color, 4 laser cell sorter. The Auto Macs was used according to the manufacturer’s instructions. Cell sorting with the FACS Aria III was performed at the OSU Flow Cytometry Core and isotype control antibodies were included to determine detection levels. CD11b+ CD11c- Ly-6Clow monocyte populations were serially gated on CD11c- cells, followed by CD11b+ with gates set around distinct populations of Ly-6C low, intermediate and high. The purity of the cell populations was ≥95%.
Data are represented as mean +/− SEM or fold change. Statistical significance was determined using a Student’s t-test or 1 way ANOVA with a significance level (p-value) < 0.05 and the Wilcoxen signed-rank test. All analyses were performed using Excel and/or GraphPad Prism software (La Jolla, CA).
Antigen presenting cell
Granulocyte-macrophage colony-stimulating factor
PD-L2, Program death ligand 1 and 2
Tolerogenic/Regulatory Dendritic cell.
This work was supported by the National Institutes of Health (K01RR022198 and R21 AI095773-01A1) and the Department of Veterinary Biosciences, College of Veterinary Medicine. The authors thank Drs. Caroline Whitacre, Prosper Boyaka and Kenneth Rock for providing the FoxP3+ EGFP, EG7/EL4 and DC2.4 cell lines, respectively. The authors thank Dr. Blake Eason Hildreth III for his contributions to statistical analysis and Drs. Prosper Boyaka and Junbae Jee for their intellectual discussions. The authors would like to acknowledge the Analytical Cytometry Shared Resource at the OSUCCC-James for all flow sorting.
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