Mesenchymal stromal cells impair the differentiation of CD14++ CD16− CD64+ classical monocytes into CD14++ CD16+ CD64++ activate monocytes

Cytotherapy, 2012; 14: 12–25

Mesenchymal stromal cells impair the differentiation of CD14ⴙⴙ CD16– CD64ⴙ classical monocytes into CD14ⴙⴙ CD16ⴙ CD64ⴙⴙ activate monocytes

BÁRBARA DU ROCHER1,2, ANDRE LUIZ MENCALHA1,2, BERNADETE EVANGELHO GOMES1 & ELIANA ABDELHAY1,2 1Instituto

Nacional de Câncer, Centro de Transplante de Medula Óssea, Laboratório de Célula Tronco, Rio de Janeiro, Brazil, and 2Instituto de Biofísica Carlos Chagas Filho, Universidade Federal do Rio de Janeiro, Rio de Janeiro, Brazil Abstract Background aims. Mesenchymal stromal cells (MSC) possess immunomodulatory activity both in vitro and in vivo. However, little information is available regarding their function during the initiation of immunologic responses through their interactions with monocytes. While many studies have shown that MSC impair the differentiation of monocytes into dendritic cells and macrophages, there are few articles showing the interaction between MSC and monocytes and none of them has addressed the question of monocyte subset modulation. Methods. To understand better the mechanism behind the benefit of MSC infusion for graft-versus-host treatment through monocyte involvement, we performed mixed leucocyte reactions (MLR) in the presence and absence of MSC. After 3 and 7 days, cultures were analyzed by flow cytometry using different approaches. Results. MSC induced changes in monocyte phenotype in an MLR. This alteration was accompanied by an increase in monocyte counting and CD14 expression. MSC induced monocyte alterations even without contact, although the parameters above were more pronounced with cell–cell contact. Moreover, the presence of MSC impaired major histocompatibility complex (MHC) I and II, CD11c and CCR5 expression and induced CD14 and CD64 expression on monocytes. These alterations were accompanied by a decrease in interleukin (IL)-1β and IL-6 production by these monocytes, but no change was observed taking into account the phagocytosis capacity of these monocytes. Conclusions. Our results suggest that MSC impair the differentiation of CD14⫹⫹ CD16– CD64⫹ classical monocytes into CD14⫹⫹ CD16⫹ CD64⫹⫹ activated monocytes, having an even earlier role than the differentiation of monocytes into dendritic cells and macrophages. Key Words: immunosuppression, mesenchymal stromal cells, monocyte subsets

Introduction Mesenchymal stromal cells (MSC) are a heterogeneous population of fibroblast-like cells originally described by Friedenstein et al. in 1968 (1). Although they are distributed to virtually all post-natal organs and tissues (2), MSC are more frequently found in the bone marrow, where they are considered to improve and regulate hematopoiesis (3). Along with this property, MSC are able to differentiate into many different cell types, such as adipocytes, chondroblasts and osteoblasts (4). Because of this ability to differentiate, associate with their tropism for damaged tissue, they have gained attention in regenerative medicine as a promising cellular therapy (5). In fact, many groups have shown that MSC are able to improve patient status with different pathologic conditions, such as acute myocardial infarct and stroke,

and most of them have attributed the therapeutic effect of MSC to their capacity to differentiate into cells and rescue cells from death through paracrine effects such as the release of angiogenic and antiapoptotic molecules and growth factors (6,7). Surprisingly, initial studies performed by two groups have shown that MSC also have an immunomodulatory function, acting as suppressor cells capable of inhibiting immunologic responses (8,9). This discovery has led to two concepts: (a) the possibility of using MSC to control undesirable immunologic responses, such as graft-versus-host disease (GvHD) and autoimmune disease; and (b) the hypothesis that their therapeutic effect in the regenerative medicine field is not only related to their plasticity, angiogenic and anti-apoptotic properties but also their immunomodulatory and anti-inflammatory properties.

Correspondence: Bárbara Du Rocher, Praça da Cruz Vermelha 23, 6° andar, Ala C. Rio de Janeiro, RJ, Brasil. CEP 20.230–130. E-mail: barbaradurocher@ inca.gov.br, [email protected] (Received 7 March 2011; accepted 26 May 2011) ISSN 1465-3249 print/ISSN 1477-2566 online © 2012 Informa Healthcare DOI: 10.3109/14653249.2011.594792

MSC impair CD14⫹⫹ CD16⫺ CD64⫹ monocyte differentiation Supporting this hypothesis, some groups have shown that the beneficial effects of MSC infusion in some pathologic conditions are only achieved during the initial phase, presumably because of the inflammatory status characteristic of this phase (10,11). Despite our unsatisfactory knowledge regarding the physiologic function of MSC and their immunomodulatory properties, these cells have been introduced in the clinical setting with encouraging results and no or minimal side-effects (12–16). The early use of MSC in the clinic has created an immediate need for investigation of their biologic properties to guide their use. For this purpose, particular attention has been given to the interaction of MSC with professional antigen-presenting cells (APC) such as dendritic cells (DC) (17–26) and, more recently, macrophages (27–29). Interestingly, little attention has been given to monocytes (30), which are known to be progenitor cells for both macrophages and myeloid DC. Monocytes originate from myeloid precursors in the bone marrow, and after maturation these cells emigrate from the bone marrow to circulate in the blood for a few days before entering tissues to differentiate into macrophages or DC (31,32). Monocytes are heterogeneous and can be characterized by distinct phenotypes and function. Among these features, CD14 (part of the lipopolysaccharide receptor), CD16 (Fcγ receptor III) and CD64 (Fcγ receptor I) expression has been largely used to define monocyte subsets in humans (33). Monocytes can be divided into at least three subsets based on the expression of CD14 and CD16: a major subset, CD14⫹⫹ CD16– classical monocytes, and a minor population of CD16⫹ monocytes, recently showed to be composed of at least two populations, CD14⫹⫹ CD16⫹ and CD14⫹ CD16⫹, the last one defined as ‘pro-inflammatory’ monocytes (34). These three monocyte subsets exhibit different features and present different biologic properties, such as antigen-presentation capacity and cytokine production. The major subset, CD14⫹⫹ CD16– classical monocytes, shows a typical morphologic phenotype and has higher phagocytic activity and lower activity in stimulating lymphocyte proliferation. In contrast, CD14⫹ CD16⫹ monocytes have lower phagocytic activity and higher activity in stimulating lymphocyte proliferation. The third monocyte subset, CD14⫹⫹ CD16⫹, is thought to be an intermediate phenotype between CD14⫹⫹ CD16– classical monocytes and CD14⫹ CD16⫹ pro-inflammatory monocytes (34,35). Although they can be classified as three different cells, it is believed that CD14⫹⫹ CD16– classical monocytes give rise to CD14⫹ CD16⫹ pro-inflammatory monocytes under homeostatic conditions by a maturation process, and that the latter represent a more

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mature state of CD14⫹⫹ CD16– classical monocytes (36–39). Furthermore, despite the ability of both subsets to generate DC in vitro and in vivo (40,41), CD14⫹ CD16⫹ pro-inflammatory monocytes seem to be more predisposed to become migratory DC (42). Another Fc receptor (FcR) that is differentially expressed by this monocyte subset is the Fcγ receptor I (CD64). This Fcγ receptor was found to be expressed on the majority of monocytes, so the major subset CD14⫹⫹ CD16– classical monocytes are CD64⫹ cells. The CD16⫹ monocytes comprise a more heterogeneous population that can or not express low levels of this molecule (35,43). Because MSC can interfere with the differentiation/maturation status of DC and macrophages, it is presumed that MSC may also interfere with monocytes. Because of this, we have explored the interaction between MSC and monocytes. Our results show that MSC impair the differentiation of CD14⫹⫹ CD16– CD64⫹ classical monocytes into CD14⫹⫹ CD16⫹ CD64⫹⫹ activated monocytes, acting even earlier than previously anticipated by others groups, who have shown that MSC inhibit the differentiation of monocytes into DC and macrophages. As GvHD is triggered by host APC and maintained by donor APC (44), we envisage that MSC infusion would also have its therapeutic effect through inhibition of monocyte subset differentiation, which will lead to the reduction of APC generation and ultimately to the decline of GvHD manifestation. Methods Cell culture Bone marrow samples were obtained from healthy bone marrow donors, and peripheral blood samples were obtained from healthy blood donors. This work was approved by the local ethics committee of Instituto Nacional de Câncer (Rio de Janeiro, Brazil). All samples were collected with written informed consent. MSC primary culture, expansion and characterization MSC were isolated from heparinized bone marrow samples. Briefly, mononuclear cells were obtained by density-gradient centrifugation (Histopaque 1.077 g/ mL; Sigma, St Louis, MO, USA) according to the manufacturer’s protocol. Mononuclear cells were seeded at 2 ⫻ 105 cells/cm2 in MSC complete medium consisting of Dulbecco’s modified Eagle’s mediumlow glucose (DMEM-LG; Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (FBS; HyClone, Waltham, MA, USA), 2 mM glutamine (Invitrogen) and 100 U/mL penicillin with 100 μg/mL streptomycin (Sigma). Cells were

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incubated at 37°C in a humidified 5% CO2 atmosphere and allowed to adhere for 48 h, after which non-adherent cells were removed. One half of the medium was changed twice a week until 80–90% confluence was reached. Afterwards, cells were trypsinized (0.1% trypsin; Invitrogen) at 37°C for 3 min and replated at 4 ⫻ 103 cells/cm2. To achieve a less heterogeneous population, we only used cultures after the second passage. These cells were in accordance with the minimal criteria for defining multipotent MSC as defined by The International Society for Cellular Therapy (45). They were plastic-adherent cells that were able to differentiate into adipocytes, osteoblasts (46) and chondrocytes (data not shown) and expressed CD73, CD90 and CD105 in the absence of lineage commitment markers, such as CD14, CD19, CD34, CD45 and HLA-DR (data not shown). All antibodies were purchased from BD Biosciences (San Jose, CA, USA). Mixed leukocyte reactions Peripheral blood mononuclear cells (PBMC) were isolated from heparinized blood samples of unrelated healthy volunteers by density-gradient centrifugation (1.077 g/mL; Sigma) according to the manufacturer’s protocol. Mixed leukocyte reactions (MLR) were performed by incubating 5 ⫻ 105 PBMC responder cells and 5 ⫻ 105 irradiated (2500 cGy) PBMC stimulator cells in a final volume of 1 mL/well in 24-well, flat-bottomed tissue-culture plates containing MLR complete medium in the absence or presence of 5 ⫻ 104 third-party MSC. MLR complete medium consisted of RPMI-1640 (Invitrogen) supplemented with 10% fetal bovine serum (FBS; HyClone), 2 mM glutamine (Invitrogen) and 100 U/mL penicillin with 100 μg/mL streptomycin (Sigma). Cells were incubated at 37°C in a humidified 5% CO2 atmosphere for 3 or 7 days, depending on the objective of the study, and were analyzed further by flow cytometry. Flow cytometry All experiments were analyzed by flow cytometry using the methodologic strategies outlined below. Data were acquired using a FACScan flow cytometer and analyzed using Paint a Gate or CellQuest software (BD Biosciences). Cell sorting was performed on a FACSAria cell sorter and data analyzed using FACSDiva software (BD Biosciences). Phenotypic characterization by cell-surface markers MLR were incubated in the presence or absence of 10% MSC for 3 or 7 days. At the end of the incubation, cells were harvested and labeled with monoclonal

antibodies (MAb) according to the manufacturer’s protocol, followed by fixation with 1% paraformaldehyde. In some cases, we also performed phenotypic characterization of fresh isolated monocytes. The antibodies used were as follows: fluorescein isothiocyanate (FITC)-labeled anti-CD14, anti-CD69 and anti-HLA-ABC; phycoerythrin (PE)-labeled anti-CD4, anti-CD8, anti-CD11c, anti-CD14, anti-CD16, anti-CD25, anti-CD38, anti-CD56 and anti-HLA-DR; peridinin-chlorophyll protein (PerCP)-labeled anti-CD3 and anti-CD19 (all from BD Biosciences); PE-labeled anti-CD195 (eBioscience, San Diego, CA, USA); and RPhycoerythrin (RPE)-labeled anti-CD64 (DAKO, Glostrup, Denmark). Twenty-thousand events were collected from each sample, except for the samples examining the monocyte phenotype, from which seven- to eight-thousand CD14 ⫹ events were collected. Results were expressed as a percentage of positive cells or as median relative fluorescence intensity (MRFI), which was calculated by subtracting the median fluorescence intensity (MFI) for specific MAb by the MFI of the respective isotype control (47). Proliferation by carboxyfluorescein diacetate succinidyl ester detection For detection of proliferation in a one-way MLR, only PBMC used as responder cells were labeled with 0.3 μM carboxyfluorescein diacetate succinidyl ester (CFSE; Invitrogen) according to the manufacturer’s protocol, prior to their use in the MLR. MLR were incubated in the presence or absence of 10% MSC for 7 days, after which cells were harvested and fixed with 1% paraformaldehyde or labeled for phenotypic characterization. We incubated the cells for 7 days after in-house optimization because we used an allogeneic stimulus, which is a discrete stimulus compared with a polyclonal stimulus such as anti-CD3 and anti-CD28, and CFSE to detect proliferation, which needs the division of cells. In this case, lymphocytes were stained with CFSE, which is a green fluorescent cell-staining dye, and upon division the dye was equally distributed to the daughter cells. The number of days also reflects previous studies in the field. To evaluate the mechanism of immunosuppression mediated by MSC, a transwell insert membrane with a 0.4-μm pore size (Corning, New York, NY, USA) was used in some experiments to prevent cell-to-cell contact and MSC were seeded into the lower chamber. Another adaptation in some experiments was the use of conditioned medium (CM) from MSC primary culture instead of the MSC themselves. In this case, primary culture under confluent conditions, approximately 16 000 cells/cm2 were incubated during 3 days in

MSC impair CD14⫹⫹ CD16⫺ CD64⫹ monocyte differentiation fresh medium and then supernatant was collected, centrifuged to obtain a cell-free CM, and immediately used in the assay in a final volume of 1 mL/well. Twenty-thousand CFSE⫹ events were collected from each sample. Results were expressed as a percentage of positive cells for CFSE (hi and low) in combination or not with specific(s) MAb for identification of lymphocyte subsets. Cell cycle distribution by Propidium iodide (PI) staining MLR were incubated for 7 days in the presence or absence of 10% MSC, after which cells were harvested, washed in phosphate-buffered saline (PBS) and reconstituted in hypotonic buffer (0.1% sodium citrate, 0.1% Triton-X, 100 μg/mL RNase and 50 μg/mL PI). Cells were immediately acquired and ten-thousand events were collected from each sample. Results were expressed as a percentage of cells in each cell cycle phase. Monocyte morphology by cell sorting and May– Grünwald–Giemsa staining MLR were incubated for 7 days in the presence or absence of 10% MSC. At the end of the incubation, cells were harvested, labeled with anti-CD14–PE (BD Biosciences) and sorted. Sorting was performed using CD14 expression, forward-scatter and side-scatter as discriminating parameters. Sorted cells were then centrifuged onto glass slides by using a cytospin cytocentrifuge (Shandon, Pittsburgh, PA, USA) stained with May–Grünwald–Giemsa solution and analyzed by light microscopy on an Olympus BX41 microscope. The purity of sorted cells was above 90%. Monocyte counting by the flow rate calibration method MLR were incubated for 7 days in the presence or absence of 10% MSC. At the end of the incubation, cells were harvested and labeled with anti-CD14–PE (BD Biosciences) followed by fixation with 1% paraformaldehyde. The flow rate calibration method was used as described previously (48). For this method, microbeads were added to the sample just prior to acquisition. A FACScan flow cytometer was set up to save five-thousand beads. Results were expressed as cells per minute (c.p.m.). Cytokine production by intracellular detection MLR were incubated for 3 days in the presence or absence of 10% MSC, and 4 h prior to intracellular cytokine detection 1 μL Brefeldin A (BD Biosciences) was added to each well to inhibit cytokine

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secretion. For detection of cytokine expression inside monocytes, cells were harvested and labeled with anti-CD14–FITC (BD Biosciences) followed by fixation (1% paraformaldehyde) and permeabilization (0.5% Tween) to allow intracellular cytokine staining. All procedures were performed according to the manufacturer’s protocol. The antibodies used were PE-labeled anti-interleukin (IL)-1β and anti-IL-6 (eBioscience). Seven- to eight-thousand CD14⫹ events were collected from each sample. Results were expressed as the MRFI of each cytokine within the monocyte gate defined by CD14 expression. Phagocytosis assay by zymosan internalization MLR were incubated for 3 days in the presence or absence of 10% MSC and then 5 ⫻ 105 cells from MLR were transferred to polystyrene tubes. Zymosan–FITC (Molecular Probes, Eugene, OR, USA) was mixed with opsonizing reagent (Molecular Probes) at a 1:1 ratio and transferred to the polystyrene tubes at a final concentration of 0.125 mg/mL. Cells were incubated for 15 min at 37°C to allow zymosan phagocytosis, uptake was stopped by adding ice-cold PBS containing 1% bovine serum albumin (BSA) and 0.2% sodium azide, and cells were stained with anti-CD14–PE (BD Biosciences) followed by fixation with 1% paraformaldehyde. In order to discriminate between attachment versus internalization of zymosan particles, extracellular fluorescence was quenched by trypan blue during 5 min at 1 mg/ mL (final concentration), as described previously (49). This approach enabled the internalized target to remain fluorescent while inhibiting extracellular FITC fluorescence. After trypan blue treatment, cells were fixed with 1% paraformaldehyde. Sevento eight-thousand CD14⫹ events were collected from each sample. Results were expressed as the percentage of positive cells for zymosan–FITC within the monocyte gate defined by CD14 expression. Real-time quantitative polymerase chain reaction analysis MSC were incubated alone (5 ⫻ 104 cells/well in a 24-well plate) or in the presence of MLR in the upper chamber of a transwell insert for 2, 4 or 7 days. After that total RNA from MSC was extracted with TRIzol LS reagent (Invitrogen) according to the manufacturer’s protocol and stored at –70°C. Analysis of mRNA levels was carried out by real-time quantitative polymerase chain reaction (RT-qPCR). A total of 2 μg RNA was reverse transcribed with Superscript II Reverse Transcriptase® (Invitrogen). cDNA dilutions (1:100) were mixed with SYBR Green PCR Master Mix® (Applied Biosystems, Carlsbad, CA,

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USA) and COX-2 primers (forward primer 5′-CAG ACG CCC TCA GAC AGC AAA-3′, reverse primer 5′-ATG GGT GGG AAC AGC AAG GAT-3′). RT-qPCR was performed in a Rotor Gene 6000 thermocycler (Corbett, Sydney, New South Wales, Australia) with 50 cycles of 20 s at 95°C, 30 s at 60°C and 30 s at 72°C. For each sample, the expression of target genes was normalized to GAPDH mRNA levels (forward primer 5′-ACCACAGTCCATGCCATCAC-3′, reverse 5′-CCACCACCCTGTTGCTGTA-3′). All primers demonstrated equal amplification efficiency and specific PCR product through dissociation curve analysis. Fold-expression was calculated using the Delta delta cycle threshold (DDCt) method. Statistical analysis A student’s t-test, with one-way or two-way analysis of variance (ANOVA), was performed. P-values less than 0.05 were considered as statistically significant (∗P ⬍ 0.05, ∗∗P ⬍ 0.01, and ∗∗∗P ⬍ 0.001). Statistical analyzes and graphical representations were performed using GraphPad Prism™ software (GraphPad, La Jolla, CA, USA) ns (not significant). Results MSC do not inhibit lymphocyte activation MSC failed to inhibit lymphocyte activation (Figure 1A). Even in the presence of MSC, the percentage of lymphocytes expressing CD25, CD38 and CD69 was unchanged compared with MLR alone, reflecting the idea that MSC need to be activated before becoming suppressive. The percentage in MLR versus MLRMSC was, respectively: CD25, 18 ⫾ 7% ⫻ 15 ⫾ 6%; CD38, 57.2 ⫾ 14.8% ⫻ 57.5 ⫾ 15%; and CD69, 5.6 ⫾ 5.4% ⫻ 5.8 ⫾ 4.4%. MSC inhibit the proliferation of T and B lymphocytes and natural killer cells As described previously, MSC failed to elicit lymphocyte proliferation when cultured with unmatched mononuclear cells (data not shown). Moreover, when added to MLR, MSC were able to inhibit lymphocyte proliferation in a dose-dependent manner (data not shown). Because responder cells constitute a heterogeneous population, we analyzed which subsets of cells were susceptible to MSC modulation. Our results showed that MSC were able to inhibit the proliferation of helper T lymphocytes, (Th) cytotoxic T lymphocytes (Tc), B lymphocytes and natural killer (NK) cells (Figure 1B). The percentage of proliferative cells in MLR versus MLR-MSC

was, respectively: Th cells, 46.4 ⫾ 19% ⫻ 8 ⫾ 4.2% (∗∗∗P ⬍ 0.001); Tc cells, 44.1 ⫾ 16.4% ⫻ 13.3 ⫾ 5.2% (∗∗P ⬍ 0.01); B cells, 27.9 ⫾ 13.3% ⫻ 8.26 ⫾ 6.6% (∗P ⬍ 0.05); and NK cells, 51.6 ⫾ 16.5% ⫻ 12.9 ⫾ 10.5% (∗∗∗P ⬍ 0.001). MSC inhibit lymphocyte proliferation through mechanisms dependent and independent of contact In accordance with previous studies, MSC had inhibitory activity even in a transwell system, suggesting that the observed inhibition is mediated by soluble factors (Figure 1C, D). The percentage of proliferative cells in MLR was 40.7 ⫾ 15%, and in the presence of MSC 6.3 ⫾ 1.7% (∗∗P ⬍ 0.01); when contact between cells was prevented, we observed 17.8 ⫾ 4.2% (∗P ⬍ 0.05). Moreover, the percentage of cells in G0–G1 phase was 93.7 ⫾ 3.3% in MLR, 99 ⫾ 0.5% in MLR-MSC (∗∗P ⬍ 0.01) and 97.5 ⫾ 0.9% in a transwell system (∗P ⬍ 0.05). This results showed that, although MSC inhibits through soluble factors, cell-to-cell contact improves their immunosuppression capacity. To evaluate the mechanism of inhibition, CM from primary culture was used instead of MSC and no inhibitory activity was observed (Figure 1E; 41.3 ⫾ 6.7% in MLR ⫻ 47 ⫾ 6.6% in MLR-CM). These results suggested that, although MSC inhibit lymphocyte proliferation through soluble factors (Figure 1C, D), they do not produce these molecules under steady-state conditions or at least not enough to suppress the immune response (Figure 1E). As conditined medium were not able to inhibit lymphocyte proliferation but MSC in a transwell system were, these results suggested that MSC need to receive some sign from mononuclear cells to act as suppressor cells. MSC induce changes in monocyte phenotype The presence of MSC in the MLR induced morphologic changes, such as size and granulosity, in the monocyte population, as evidenced by flow cytometry analysis (Figure 2A–C). Regarding size, we observed, using arbitrary units, 806 ⫾ 57 in MLR ⫻ 604 ⫾ 91 in MLR-MSC, a decrease of 20% (∗∗P ⬍ 0.01), and for granulosity we observed 611 ⫾ 123 in MLR ⫻ 396 ⫾ 94 in MLR-MSC, a decrease of 35% (∗∗∗P ⬍ 0.001). To confirm this further, monocytes were sorted and their morphology analyzed by light microscopy (Figure 2D). These alterations were accompanied by an increase in monocyte numbers (Figure 2E; 1319 ⫾ 787 in MLR ⫻ 6125 ⫾ 1790 in MLR-MSC; ∗P ⬍ 0.05) and in approximately 50% of CD14 expression (Figure 2F; 121 ⫾ 67.7 in MLR ⫻ 251 ⫾ 151.6 in MLRMSC; ∗P ⬍ 0.05). These observations prompted us to investigate whether monocyte modulation

MSC impair CD14⫹⫹ CD16⫺ CD64⫹ monocyte differentiation

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Figure 1. Characterization of MSC-mediated immunosuppression. (A) MSC do not inhibit lymphocyte activation but inhibit Th, Tc, B and NK cell proliferation (B). MSC were able to inhibit MLR even in a transwell (t) system (C, D); however, CM was ineffective in the control of lymphocyte proliferation (E). The activation profile of the lymphocyte population was assessed in MLR incubated for 3 days and the lymphocyte proliferation and cell cycle phase were assessed in MLR incubated for 7 days. Experiments were performed in the absence (MLR) or presence of 10% MSC, with (MLR-MSC) or without contact (MLR-MSC t) or in the presence of CM (MLR-CM). Data were analyzed within lymphocyte gates defined by CD3 expression (A–E), except for cell-cycle analyzes. For proliferation assessment, the gate was defined based on CD3 and CFSE, and, when appropriate, CD4, CD8, CD19 and CD56 were employed (B). Results are expressed as mean ⫾ SD of four independent experiments (A–C), six independent experiments (D) and seven independent experiments (E) (∗∗∗P ⬍ 0.001, ∗∗P ⬍ 0.01, ∗P ⬍ 0.05).

could be involved in MSC-mediated immunosuppression and how monocytes could contribute to MSC-mediated modulation. To investigate whether monocytes could be involved in the immunosuppression mediated by MSC, we performed the experiments using a 3-day culture (see Supplementary Figure 1). This incubation period was enough to induce the same alterations that were observed after the 7-day culture

(Figure 2A–F). We also performed transwell experiments, because lymphocyte proliferation was abrogated even without cell–cell contact. The results showed that MSC induced monocyte alteration even without contact, although these alterations were more pronounced with cell–cell contact (see Supplementary Figure 1). Regarding size, we observed 771 ⫾ 111 in MLR, 651.8 ⫾ 89.6 in MLR-MSC (∗∗∗P ⬍ 0.001) and 625.2 ⫾ 143.5 in MLR-MSC

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Figure 2. MSC induced a range of alterations in the monocyte population. (A) Cell size measured by forward scatter (FSC; arbitrary unit). (B) Cell granulosity measured by side scatter (SSC; arbitrary unit). (C) Representative assay showing an FSC ⫻ SSC dot-plot of the monocyte population (CD14 ⫹ ). (D) Light microscopic analysis of sorted monocytes (⫻100). (E) Monocyte counting (c.p.m.). (F) CD14 expression (MRFI). Monocytes were assessed in MLR incubated for 7 days performed in the absence (MLR) or presence of 10% MSC (MLR-MSC). Data were analyzed within a monocyte gate defined by CD14 expression. Results are expressed as mean ⫾ SD of six independent experiments (A–C, F) and four independent experiments (E) (∗∗∗P ⬍ 0.001, ∗∗P ⬍ 0.01, ∗P ⬍ 0.05).

t (∗P ⬍ 0.05), regarding granulosity 487 ⫾ 73 in MLR, 444.6 ⫾ 66 in MLR-MSC (∗∗P ⬍ 0.01) and 448.5 ⫾ 50.5 in MLR-MSC t, and regarding CD14 expression 62.12 ⫾ 29.5 in MLR, 150.4 ⫾ 87.6 in MLR-MSC (∗∗P ⬍ 0.01) and 104.7 ⫾ 4.4 in MLRMSC t. These results indicated that monocytes can be involved in the mechanism of immunosuppression induced by MSC. MSC impair monocyte maturation CD14⫹ CD16⫹ pro-inflammatory monocytes appear to be more mature monocytes that are generated by

the maturation of CD14⫹⫹ CD16– classical monocytes under homeostatic conditions, as indicated by previous studies (36–39). As MSC impair immunologic responses, we hypothesized that MSC could be interfering in the balance between these subsets. Initially, we evaluated the phenotype of preculture monocytes (Pre-culture Mo) and compared this with the phenotype of the same monocytes after 3 days in the MLR. Pre-culture monocytes were composed mainly of CD14⫹⫹ CD16– CD64⫹ classical monocytes (approximately 90–95% of all monocytes), and 3 days were sufficient to induce their differentiation into a unique population of CD14⫹⫹ CD16⫹

MSC impair CD14⫹⫹ CD16⫺ CD64⫹ monocyte differentiation CD64⫹⫹ monocytes (Figure 3A, B; pre-culture Mo 8.9 ⫾ 4.1% ⫻ MLR 100%, ∗∗∗P ⬍ 0.001). Although under homeostatic conditions monocytes seem to differentiate from classical monocytes to pro-inflammatory monocytes, this does not seem to be the case in our study. Our results clearly showed that, under allogeneic stimulation, the pattern of monocyte activation was not the same as under homeostatic conditions. This result prompted us to hypothesize that MSC could be impairing the differentiation of CD14⫹⫹ CD16– CD64⫹ classical monocytes into CD14⫹⫹ CD16⫹ CD64⫹⫹ activated monocytes, favoring the CD14⫹⫹ CD16– CD64⫹ phenotype. Based on this prediction, we analyzed CD14, CD16 and CD64 expression in the monocyte population before culture and in the presence and absence of MSC. Our results showed that pre-culture monocytes expressed high levels of CD14. After 3 days, CD14 expression decreased slowly in the MLR, but in the presence

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of MSC CD14 expression increased considerably (Figure 4A; see also Supplementary Figure 2; 69.6 ⫾ 21.5 in pre-culture Mo, 60.8 ⫾ 35.1 in MLR and 126.2 ⫾ 56.1 in MLR-MSC). In contrast, CD16 expression in pre-culture monocytes was low, being almost undetectable. After MLR, in the presence or absence of MSC, CD16 expression increased significantly (Figure 4B; see also Supplementary Figure 2; 0.02 ⫾ 0.05 in pre-culture Mo, 7.9 ⫾ 6.5 in MLR and 12.7 ⫾ 12.8 in MLR-MSC). Finally, pre-culture monocytes expressed CD64 as expected, and in MLR CD64 increased. In the presence of MSC this effect was even more prominent (Figure 4C; see also Supplementary Figure 2; 6.2 ⫾ 4.6 in pre-culture Mo, 46.2 ⫾ 11.2 in MLR and 81 ⫾ 19 in MLR-MSC). Summarizing, pre-culture monocytes were predominantly composed of CD14⫹⫹ CD16– CD64⫹ classical monocytes; during MLR they shifted towards a unique phenotype of CD14⫹⫹

Figure 3. Phenotypic characterization of the monocyte population before and after MLR. (A) CD16⫹ monocyte frequency (%). (B) Representative assay showing CD14 ⫻ CD16 and CD14 ⫻ CD64 dot-plots of the monocyte population (CD14⫹). Monocyte populations were assessed in pre-culture monocytes (pre-culture Mo) and after their activation by incubation in MLR for 3 days. Data were analyzed within a monocyte gate defined by CD14 expression. Results are expressed as mean ⫾ SD of 11 independent experiments (∗∗∗P ⬍ 0.001).

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Figure 4. MSC impair the differentiation of CD14⫹⫹ CD16– CD64⫹ classical monocytes into CD14⫹⫹ CD16⫹ CD64⫹⫹ activated monocytes. (A) CD14 expression (MRFI); (B) CD16 expression (MRFI); (C) CD64 expression (MRFI); (D) HLA-ABC (class I) expression (MRFI); (E) HLA-DR (class II) expression (MRFI); (F) CD11c expression (MRFI); (G) CCR5 (CD195) expression (MRFI). Monocyte populations were assessed in pre-culture monocytes (pre-culture Mo) and after their activation in MLR for 3 days performed in the absence (MLR) or presence of 10% MSC (MLR-MSC). Data were analyzed within a monocyte gate defined by CD14 expression. Results are expressed as mean ⫾ SD of nine independent experiments (A, B), six independent experiments (C) and four independent experiments (D–G) (∗∗∗P ⬍ 0.001, ∗∗P ⬍ 0.01, ∗P ⬍ 0.05).

CD16⫹ CD64⫹⫹ monocytes, but in the presence of MSC they were CD14⫹⫹⫹ CD16⫹ CD64⫹⫹⫹. As during differentiation, monocytes are thought to decrease CD14 and CD64 expression, these results suggested that MSC inhibits monocyte differentiation by inducing CD14 and CD64 expression. These results suggest that pre-culture monocytes undergo differentiation in MLR, and it is this differentiation that is impaired by the presence of MSC.

To support our hypothesis further, we investigated other molecules that were clearly associated with the phenotype of cells under activation and that were differentially expressed by CD14⫹⫹ CD16– classical and CD14⫹ CD16⫹ pro-inflammatory monocytes. CD14⫹ CD16⫹ pro-inflammatory monocytes expressed higher levels of HLA class I and II, CD11c and CCR5 compared with CD14⫹⫹ CD16– classical monocytes; therefore we decided

MSC impair CD14⫹⫹ CD16⫺ CD64⫹ monocyte differentiation to analyze the expression of these molecules. Preculture monocytes expressed lower levels of all of these molecules compared with monocytes in MLR and in MLR-MSC. After activation, all these molecules were up-regulated, but to a lesser extent when MSC were present (Figure 4D–G; see also Supplementary Figure 2). For HLA-ABC, mean values were 101.8 ⫾ 49 in pre-culture Mo, 311 ⫾ 33.5 in MLR and 222.8 ⫾ 33.4 in MLR-MSC; for HLA-DR, 21.72 ⫾ 9.5 in pre-culture Mo, 498.3 ⫾ 227.2 in MLR and 193 ⫾ 172.5 in MLR-MSC, for CD11c, 60.2 ⫾ 25.6 in pre-culture Mo, 271.8 ⫾ 48.7 in MLR and 208.4 ⫾ 40 in MLR-MSC; and for CCR5, 0.03 ⫾ 0.05 in pre-culture Mo, 3.96 ⫾ 1.25 in MLR and 0.84 ⫾ 0.44 in MLR-MSC.

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MSC reduce the production of inflammatory cytokines by monocytes and do not interfere in phagocytosis activity We analyzed cytokine production and phagocytosis activity as a way to evaluate the functional properties of these monocytes. Because monocytes are the major producers of IL-1β and IL-6, and these cytokines play a key role in the initiation of immunologic responses, we decided to quantify them. In the presence of MSC, the amount of IL-1β and IL-6 production by monocytes was decreased (Figure 5A–C). For IL-1β, we observed 206.2 ⫾ 83.1 in MLR versus 144.4 ⫾ 53.3 in MLR-MSC (∗P ⬍ 0.05), and for IL-6, 252.2 ⫾ 78 in MLR versus 144.4 ⫾ 41.7 in MLR-MSC (∗∗P ⬍ 0.01). Monocytes

Figure 5. IL-1β and IL-6 production by monocytes decreases in the presence of MSC but phagocytosis activity was unchanged. (A) IL-1β production by monocytes; (B) IL-6 production by monocytes; (C) representative assay showing cytokine ⫻ SSC dot-plots of the monocyte population (CD14⫹; isotype controls from both MLR and MLR-MSC were omitted); (D) phagocytosis capacity of the monocyte population; (E) representative phagocytosis assay showing an overlay histogram of monocytes from MLR and MLR-MSC. Monocyte populations were assessed in MLR incubated for 3 days performed in the absence (MLR) or presence of 10% MSC (MLRMSC). Data were analyzed within a monocyte gate defined by CD14 expression. Results are expressed as mean ⫾ SD of eight independent experiments (A–C) and four independent experiments (D, E) (∗∗P ⬍ 0.01, ∗P ⬍ 0.05).

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generated after MLR, in the presence or absence of MSC, were also tested for their ability to phagocytose FITC–zymosan particles. No difference was observed between them (Figure 5D, E; 91.8 ⫾ 7.5 in MLR and 92.6 ⫾ 5.3 in MLR-MSC). MSC up-regulate COX-2 mRNA after co-culture with MLR in a contact-independent manner In search for insights regarding the mechanism of immunosuppression induced by MSC, we cultured MSC alone or in the presence of MLR in the upper chamber of a transwell insert for 2, 4 or 7 days (D2, D4 and D7). MSC dramatically increased COX-2 mRNA levels upon co-culture with MLR (Figure 6A). After 4 days we observed 1.76 ⫾ 1.73 in MSC versus 204.64 ⫾ 127.12 in MSC t (∗∗P ⬍ 0.01) This effect was time dependent, as COX-2 mRNA relative levels were 78.8-fold increased in D2, 291.0 in D4 and 664.0 in D7 in comparison with MSC cultured alone (Figure 6B). Discussion It is well established that MSC induce immunosuppression and that this effect is mediated by soluble factors (5). In accordance with previous studies, we also observed that MSC induced immunosuppression via soluble factors. However, these factors are not produced by MSC under steady-state conditions, or at least not in amounts large enough to induce immunosuppression. This finding indicates that a crosstalk between these populations is essential, and, as proposed by other groups, MSC need to be ‘licensed’ in an inflammatory environment through their activation, mainly by interferon (IFN)-γ (50–52).

Once activated, MSC act as immunosuppressor cells through many mechanisms. Of these mechanisms, particular attention has been given to the interaction between MSC and DC. During DC differentiation and maturation, DC up-regulate the expression of many important molecules: major histocompatibility complex (MHC) II and CD1a antigen-presenting molecules; co-stimulatory molecules, such as CD40, CD80 and CD86; and maturation antigens, such as CD83 and CD11c adhesion molecules. Despite heterogeneous results between groups, in general, the acquisition of all of these molecules has been shown to be impaired by MSC. Moreover, functional assays show that, after exposure to MSC, DC diminish their capacity to produce pro-inflammatory cytokines, such as IL-12 and tumor necrosis factor (TNF)-α, to present antigen to T lymphocytes and to stimulate allogeneic T-cell proliferation and naive T cells. Interestingly, the phagocytic activity of DC in the presence of MSC did not increase as expected. Taken together, these results suggest that MSC interfere with DC differentiation and maturation, inducing an intermediate phenotype between immature DC and mature DC (17–26). It has been shown that MSC also interfere with the maturation of macrophages, inducing an alternatively activated phenotype with anti-inflammatory properties, such as high IL-10 production (27–29). In addition, it has been observed that these macrophages produce low amounts of IL-12 and TNF-α, pro-inflammatory cytokines that are associated with high phagocytic ability (28,29). It is clear that monocytes are precursor cells for DC, at least those of myeloid origin, and macrophages. Because MSC interfere with these populations, we hypothesized that they could also interact with

Figure 6. MSC up-regulate COX-2 mRNA after co-culture with MLR in a contact-independent manner (A); this induction was timedependent (B). RT-qPCR analysis was assessed in MSC incubated alone or in the presence of MLR in the upper chamber of the transwell insert for 4 days (A) or 2, 4 and 7 days (B). Raw expression values were normalized to GAPDH expression. Analyzes of COX-2 mRNA expression changes were performed in four independent experiments (A) and one independent experiment (B) (∗∗P ⬍ 0.01).

MSC impair CD14⫹⫹ CD16⫺ CD64⫹ monocyte differentiation monocytes. Our observation that monocytes stimulated under allogeneic reactions in co-culture with MSC acquired a different phenotype from those cultured without MSC, together with reports regarding the existence of different monocyte subsets, prompted us to investigate whether MSC could be impairing monocyte differentiation from one subset to the other. Moreover, observations that CD14⫹ CD16⫹ pro-inflammatory monocytes are drastically expanded in a number of diseases such as sepsis, diabetes, atherosclerose and bacterial infections, notable inflammatory conditions (31,32,34), suggest that, under pathologic conditions, cytokine deregulation and other inflammatory stimuli can trigger monocyte activation, which is evidence that monocyte subsets can be regulated and that a plasticity between these subsets is feasible. In an attempt to address this question, we analyzed CD14, CD16 and CD64 expression on the monocyte membrane surface along with other molecules that are differentially expressed by these monocyte subsets. Interestingly, we found that monocytes under allogeneic stimulation did not differentiate into the classical pathway, from CD14⫹⫹ CD16– CD64⫹ to CD14⫹ CD16⫹ CD64– monocytes; instead they differentiated into the phenotype CD14⫹⫹ CD16⫹ CD64⫹⫹. This could be because of the allogeneic stimulation used, as the classical pathway of differentiation was reported under steady-state conditions, which is clearly not the case here. Stec et al. (53) also observed the appearance of CD14⫹ CD16– and CD14⫹⫹ CD16⫹ monocytes from CD34⫹ hematopoietic progenitors cells under non-homeostatic conditions with 1.25(OH)2 vitamin D3 treatment. As in a homeostatic conditions CD14⫹ CD16– cells are not produced, these observations corroborate our results and show that non-homeostatic stimuli may give rise to different monocytic populations. These results indicate that more studies regarding the physiologic process of monocyte subset differentiation under homeostatic and non-homeostatic conditions, such as inflammation, are needed in order to understand the modulation of these populations by MSC. In our hands, We found that MSC were able to impair the differentiation of CD14⫹⫹ CD16– CD64⫹ classical monocytes into CD14⫹⫹ CD16⫹ CD64⫹⫹ activated monocytes, maintaining these cells with an intermediate phenotype. Moreover, this process was a clear maturation arrest, as evidenced by the lower levels of MHC I, MHC II, CD11c, CCR5, IL-1β and IL-6 in comparison with monocytes from MLR. The comparable phagocytosis activity between monocytes in the presence and absence of MSC, not an increase as expected, could be because of the incomplete inhibition of monocyte maturation exerted by MSC.

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While many studies have shown that MSC impair the differentiation of monocytes into DC and macrophages (17–29), there are few articles showing that MSC impair the differentiation of monocytes into more mature monocytes (54,55), and none of them has shown that MSC interfere with monocyte subsets. This means that MSC not only inhibit monocyte–DC and macrophage differentiation, but also act at a more upstream step, impairing the maturation of monocyte within the monocyte stages. The mechanisms by which MSC induce monocyte subset modulation are currently under investigation by our group. It is possible that some chemokines and derivative eicosanoids from COX-2 are involved, because we observed high levels of different chemokines (unpublished observations) and COX-2 mRNA in MSC upon their interaction with MLR. COX-2 is an enzyme involved in lipid metabolism. Using aracdonic acid as a substrate, this enzyme produces as many as seven different prostaglandins (PGG2 and its derivatives, PGI2, PGF1A PGH2, PGD2, 15-dPGJ2, PGE2 and PGF2) and two thromboxanes (TXA2 and TXB2) (56,57). Some groups, using COX inhibitors such as indomethacin, acetyl sialic acid and NS-398, have observed that the immunosuppression induced by MSC is reverted. Simultaneously, they observed an increase in PGE2 levels. Because of this, these groups have suggested that the immunosuppression induced by MSC is because of PGE2 production (54,55). It is well know that PGE2 can exert immunosuppressive activity, mainly through the prostaglandin EP2 and EP4 receptors, in mononuclear cells (54,55). However, this is an oversimplification because COX-2 produces other prostanoids that can also exert some effect. Among them, 15-d-PGJ2 has emerged as a potent immunosuppressive prostaglandin because it inhibits NFkB signaling in lymphocytes through multiple mechanisms (58,59). In this way, more studies regarding the downstream pathway governing MSC inhibition through COX-2 derivatives are necessary. Finally, because MSC act at one of the earliest stages of the developing immune response, it is possible that these inhibitory properties may have a significant effect on the immunosuppression induced by MSC. In the bone marrow transplantation setting this would be of great importance because GvHD is thought to be initiated by host APC and perpetuated by donor APC (44). As MSC inhibit the differentiation and activation of monocyte subsets, we envisage that this property has the potential to limit the generation of both host and donor APC, limiting the GvHD injury observed in clinical practice.

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Acknowledgments This work was supported by grants from CNPq (Conselho Nacional de Desenvolvimento Científico), FAPERJ (Fundação de Amparo à Pesquisa do Estado do Rio de Janeiro) and FINEP (Financiadora de Estudos e Projetos). Bárbara Du Rocher and Andre Luiz Mencalha are supported by a PhD studentship from MS (Ministério da Saúde).

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Declaration of interest: The authors declare no financial or commercial conflict of interest. 17.

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