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The Journal of Immunology, 2007, 179, 2824 -2831
Copyright © 2007 by The American Association of Immunologists, Inc.
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The Antiproliferative Effect of Mesenchymal Stem Cells Is a Fundamental Property Shared by All Stromal Cells1

Simon Jones*, Nicole Horwood{dagger}, Andrew Cope{dagger} and Francesco Dazzi2,*,{dagger}

* Department of Haematology, Division of Investigative Science, Faculty of Medicine, Imperial College London, Hammersmith Hospital, London, United Kingdom; and {dagger} Kennedy Institute of Rheumatology, Faculty of Medicine, Imperial College London, Hammersmith Hospital, London, United Kingdom


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Although it has been widely demonstrated that human mesenchymal stem cells exert potent immunosuppressive effects, there is little information as to whether more mature mesenchymal stromal cells (SC) share the same property. Accordingly, we set out to test the ability of SC from different human tissues to inhibit the proliferation of PBMC following polyclonal stimuli. Chondrocytes, as well as fibroblasts from synovial joints, lung, and skin, were used as a source of SC. Irrespective of their differentiation potential and/or content of progenitor cells, SC from all tissues exhibited antiproliferative functions. This was in marked contrast to parenchymal cells. Although SC did not interfere with early T lymphocyte activation, they arrested stimulated T cells in the G0/G1 phase of the cell cycle and rescued them from apoptosis. In addition, IFN-{gamma} and TNF-{alpha} production were reduced. We observed that the inhibitory effect is ultimately mediated by soluble factors, the production of which requires SC to be licensed in an inflammatory environment by cell contact. We conclude that the immunosuppressive effect of mesenchymal cells is not confined to multipotent stem cells, but is a fundamental characteristic of all stroma. Our data suggest that SC, appropriately licensed, regulate T cell homeostasis.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Mesenchymal stem cells (MSC)3 are progenitor cells with the ability to differentiate into mesodermal tissues, including osteoblasts, chondrocytes (CH), and adipocytes (1). MSC are typically isolated from the bone marrow (BM), but can also be expanded from a wide variety of adult and fetal tissues (2, 3, 4, 5, 6).

It has been widely demonstrated that MSC are capable of exerting a powerful in vitro and in vivo immunosuppressive effect on virtually all cells of the immune system (reviewed in Ref. 7). Immunosuppression is the result of an arrest of cell division in the G0/G1 phase of the cell cycle, associated with inhibition of cyclin D2 expression (8). The effect, initially described for T cells (9, 10), has also been confirmed for B lymphocytes (8, 11), NK (12), and dendritic cells (13, 14). As a consequence of division arrest, MSC have been shown to affect, to a variable extent, T and NK cell cytokine secretion and cytotoxicity, as well as B cell maturation and Ab secretion. The mechanisms underlying these effects remain to be fully elucidated, but are mediated by direct cell-to-cell interactions in combination with soluble factors. The most prominent candidate molecules are TGFβ-1 (9), IDO (15), NO (16), and PGE2 (13). However, there is no solid evidence that any of these factors is solely and sufficiently responsible for the antiproliferative effect.

Independently of the mechanisms involved in immunosuppression, such properties can be exploited therapeutically. In fact, MSC have been shown to prolong skin-graft survival in nonhuman primates (17) and attenuate disease in an animal model of multiple sclerosis (18). More importantly, infusion of MSC has beneficial effects in the treatment of steroid-resistant graft-vs-host disease in patients with hematological malignancies undergoing allogeneic hemopoietic stem cell transplantation (19).

We have recently demonstrated that the MSC-mediated immunosuppressive effect is the result of an antiproliferative effect that can be exerted not only on cells of the immune system, but on a variety of cells derived from different tissues (20). This questions the physiological significance of an immunomodulatory role for MSC, especially considering their presence in the BM is estimated to be ~1 x 10–4 among nucleated cells (1) and large numbers of MSC are required to exert their clinical effects (21). However, MSC also contribute to the progeny of cells that are responsible for the hemopoietic niche (22) and other types of stroma (23, 24).

Stromal cells (SC), often referred to generically as fibroblasts, exhibit specific functions related to their anatomical site (25, 26, 27). The traditional view that the stroma contributes merely to the structural integrity of a tissue has been revised with the realization that SC participate in a dynamic interplay with other cells, particularly cells of the immune system (reviewed in Ref. 28). For example, the interactions between stroma and lymphocytes in the thymus direct the development and repertoire of T cells (23, 24). The stromal microenvironment has also been strongly implicated in the initiation and persistence of inflammation in autoimmune diseases such as rheumatoid arthritis and Grave’s disease (29, 30).

In this study, we explore whether the immunosuppressive properties uniquely attributed to MSC are shared by SC of different tissue origin. We demonstrate that, independently of their differentiation state and origin, fibroblasts and mature, terminally differentiated mesenchymal cells inhibit the proliferation and prevent apoptosis of activated T cells. This activity was comparable to that of MSC and required licensing in an inflammatory environment through cell contact-dependent pathways. Our findings suggest a major role for stroma in immune homeostasis during tissue injury.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Cell isolation and culture

To isolate adult human MSC, samples of 5–20 ml of BM were taken from BM aspirates of normal donors during BM donation for transplant procedures. Mononuclear cells were separated through density gradient separation by Ficoll-Paque (Amersham Biosciences) and seeded at 1 x 106 cells/ml in 75-cm2 flasks (Costar) with SC medium. SC medium consisted of high glucose DMEM (Invitrogen Life Technologies) supplemented with 10% preselected FBS (Cambrex), 1 x 105 U/ml penicillin G sodium, 1 x 105 µg/ml streptomycin sulfate, and 25 µg/ml amphotericin B (PAA). Cells were allowed to adhere for 48–72 h, nonadherent cells were removed, and the medium was changed. Thereafter, the medium was changed every 3 days. When 70–80% confluent, adherent cells were detached with 0.05% trypsin-EDTA (Invitrogen Life Technologies) at 37°C for 5 min and reseeded at 1000 cells/cm2. All MSC were expanded to at least passage three, to remove contaminating monocytes and hemopoietic cells, before being used in experiments. MSC were also routinely tested for their ability to differentiate into adipocytes and osteoblasts, as previously described (8). A control sample of MSC was also purchased commercially (Cambrex).

Synovial fibroblasts (SF) were isolated from synovial membrane tissue obtained at the time of knee or hip replacement surgery or during synovectomy or synovial biopsy from patients with rheumatoid arthritis and osteoarthritis. All patients were seen at rheumatology clinics within the Hammersmith Hospitals Trust. Synovial membrane samples were first digested with 500 µg/ml collagenase type VIII (Sigma-Aldrich). The resulting cell suspension was then separated by density gradient separation using Ficoll-Paque (Amersham Biosciences), and the mononuclear cells were seeded at 1 x 106 cells/ml in 75-cm2 flasks (Costar) in SC medium.

Human dermal fibroblasts (DF) were generated from explant skin obtained from normal donors during cosmetic surgery. All patients were seen at dermatology clinics within the Hammersmith Hospitals Trust. Explant skin was cut into thin shallow strips and digested with 2 mg/ml dispase II neutral protease (Sigma-Aldrich). Samples were incubated overnight at 4°C, after which the epidermis was removed and the dermal layer was placed directly into 75-cm2 flasks (Costar). Human skin-derived and human lung-derived fibroblast (LF) primary cell lines were also obtained commercially (BJ and MRC-5, respectively; American Type Culture Collection).

Primary human articular CH were a gift from C. Murphy (Kennedy Institute of Rheumatology, Imperial College London, London, U.K.). CH used in experiments were between passage one and three. All SC were maintained and cultured as outlined above for MSC. All samples were provided with written, informed consent in accordance with the Hammersmith Hospital and Queen Charlotte’s Hospital ethical committee requirements. HUVEC were a gift from J. Mongkolsapaya (Department of Immunology, Imperial College London, London, U.K.). A sample of HUVEC was also obtained commercially (Promo Cell). Neural progenitor cells were a gift from S. Chandran (Department of Clinical Neurosciences, Centre for Brain Repair, Cambridge University, Cambridge, U.K.). Neurons were obtained from neural progenitor cells, as described previously (31), by culturing them as an adherent layer in T75 flasks (Costar) precoated with glycine and 0.5% laminin (both Sigma-Aldrich).

PBMC were obtained from buffy coats purchased from the National Blood Service (Colindale). Buffy coats were diluted 1/1 with RPMI 1640 (Invitrogen Life Technologies) and layered on Ficoll-Paque (Amersham Biosciences) for density gradient separation. The mononuclear cell layer was removed and cryopreserved in freezing medium (90% heat-inactivated FBS (Cambrex) and 10% DMSO (Sigma-Aldrich)) at 2 x 107 cells/ml.

Osteogenic and adipogenic differentiation

SC were plated at a density of 4 x 103 cells/cm2 in 25-cm2 culture flasks (Costar) and cultured in SC medium until confluent. At day 0, either osteogenic conditioned medium, containing 100 nM dexamethasone, 10 mM β-glycerol phosphate, and 50 µM ascorbic acid (all Sigma-Aldrich), or adipogenic induction medium (Cambrex) was added to cultures. Control flasks with normal SC medium were cultured in parallel. Medium changes were performed every 3 days. After 21 days, all cultures were rinsed with PBS and fixed in 10% formaldehyde (Sigma-Aldrich). For alizarin red staining, fixed cells were rinsed with water and 40 mM alizarin red solution was added to the flasks. Monolayers were incubated for 20 min at room temperature and then rinsed thoroughly with water to remove excess alizarin red solution. For oil red O (ORO) staining, fixed cells were rinsed twice with water, and 60% isopropanol was briefly added to the flasks. ORO working solution was prepared by dissolving 300 mg of ORO powder (Sigma-Aldrich) in 100 ml of 99% isopropanol. This stock solution was then mixed at a 3:2 ratio with water, filtered, and added to fixed cells for 5 min. Stained monolayers were counterstained with hematoxylin and visualized under a phase-contrast light microscope.

Proliferation assay and cell cycle analysis

PBMC were thawed and washed twice in RPMI 1640 (Invitrogen Life Technologies) supplemented with 10% FBS (Cambrex) and 1 x 105 U/ml penicillin G sodium, 1 x 105 µg/ml streptomycin sulfate, and 25 µg/ml amphotericin B (PAA). Cells were then trypan blue (Sigma-Aldrich) tested for viability and adjusted to a concentration of 1 x 106 cells/ml. PBMC were cultivated in triplicate, and 1 x 105 cells were incubated in 96-well U-bottom microtiter cell plates (Costar) with either 5 µg/ml PHA (Sigma-Aldrich) or 0.2 µl/well anti-CD3/CD28-coated T cell beads (Dynal Biotech). The cells were incubated in a total volume of 200 µl/well in the presence or absence of SC for 3 days. SC were preplated overnight before addition of PBMC. A total of 18.5 MBq (0.5 µCi) [3H]TdR (Amersham) was added to each well for the last 15 h of culture. Plates were harvested onto glass fiber filter mats (Wallac) using a 96-well automated cell harvester (Molecular Devices). Scintillation fluid was added, and [3H]TdR incorporation was measured using a liquid scintillation counter (Betaplate; Wallac). For cell cycle analysis, PBMC were removed from a 3-day proliferation culture, in the presence or absence of SC, and fixed in ice-cold 70% ethanol. Cells were incubated overnight at 4°C and then washed twice. A total of 0.1 µg/ml FITC, 40 µg/ml propidium iodide (PI), and 25 µg/ml RNase A (all Sigma-Aldrich) was added to the cells for 30 min at room temperature in the dark. Cells were immediately analyzed by flow cytometry.

Flow cytometry (FACS)

PBMC or SC were stained with a selection of the following conjugated mouse anti-human mAbs and corresponding isotype controls for 30 min at 4°C: FITC-conjugated CD4, CD14, CD45, MHC class II, CD80, CD90, and annexin V; PE-conjugated mouse anti-human CD69, CD73, CD106, and CD166; Cychrome-5-conjugated CD86 and MHC class I; and allophycocyanin-conjugated mouse anti-human CD3, CD4, and CD25. Unconjugated mouse anti-human Abs to CD55 (Serotec) and CD105 were detected using an allophycocyanin-conjugated rat anti-mouse secondary Ab (all BD Biosciences, unless otherwise stated). FACS was performed using FACSCalibur (BD Biosciences) and analyzed with FlowJo software (Tree Star).

CFSE labeling and Transwell experiments

PBMC were washed twice in cold PBS and resuspended at 1 x 106 cells/ml in PBS/0.1% BSA. CFSE was added at a final concentration of 2.5 µM and incubated at 37°C for 10 min. Staining was then quenched by the addition of at least 5 volume of ice-cold culture medium. Cells were washed a total of three times and then resuspended at 1 x 106 cells/ml, and 1 ml was added to the bottom of a 24-well tissue culture plate (Costar) in the presence or absence of 5 µg/ml PHA. SC were preseeded in the lower or upper chamber (with the presence of unstained PBMC in some conditions) of a 6.5-mm diameter, 3.0-µm-pore-size Transwell insert (Costar). After 3 days, CFSE-stained cells were harvested, labeled with allophycocyanin-conjugated anti-CD3 (BD Biosciences), and analyzed by FACs.

Intracellular cytokine-staining and apoptosis assay

Six hours from the end of a 3-day PBMC culture in the presence or absence of SC, PMA (100 ng/ml) and calcium ionophore (500 ng/ml) (both Sigma-Aldrich) were added to cultures. For the final 3 h, 0.7 µl/ml monensin (GolgiStop; BD Biosciences) was added to wells. PBMC were labeled with allophycocyanin-conjugated anti-CD3 (BD Biosciences) for 30 min at 4°C. Cells were then fixed in 2% paraformaldehyde for 30 min and permeabilized in 0.5% saponin (both Sigma-Aldrich) for 10 min at room temperature. PBMC were finally stained with FITC-conjugated anti-IFN-{gamma} (BD Biosciences) for 30 min at 4°C and then analyzed by FACS. For quantification of apoptotic cells, PBMC were stimulated with PHA (5 µg/ml) in low serum condition (2% FBS) for 3 days in the presence or absence of SC. PBMC were removed, washed, and surface stained with either anti-CD3 or anti-CD4 and anti-CD8 Abs (all BD Biosciences). Cells were washed twice in PBS and resuspended at 1 x 106 cells/ml in annexin V-binding buffer (BD Biosciences). FITC-conjugated mouse anti-human annexin V was then added at a concentration of 5 µl/105 cells along with 5 µg/ml PI, and incubated for 15 min in the dark at room temperature. Cells were analyzed immediately by FACS.

Human TNF-{alpha} ELISA

Cell-free supernatants from SC and PBMC proliferation assays were analyzed for TNF-{alpha} content. Ninety-six-well plates were coated with 4 µg/ml capture Ab for human TNF-{alpha} (BD Biosciences) overnight at 4°C. Plates were washed and blocked with 2% BSA for 2 h. Supernatant was added in triplicate and incubated with 0.5 µg/ml biotinylated detection Ab for 1 h at room temperature. Streptavadin HRP conjugate was then added for 1 h, after which tetramethylbenzidine was added and the reaction stopped with sulfuric acid. Plates were read at 450 nm.

Statistics

Data were analyzed using GraphPad Prism 4 software for Macintosh. Results are expressed as mean ± SEM. Differences between experimental conditions were analyzed using two-tailed Student’s t test. A p value of <0.05 was considered statistically significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Characterization of SC

MSC, SF, DF, LF, and CH were evaluated for the expression of a number of surface molecules. All cell types lacked MHC class II, CD14, CD45, CD80, CD86, and CD106 (VCAM-1), whereas they were all positive for MHC class I, CD55, CD73 (Src homology 3/4), CD90 (Thy-1), and CD105 (SH2). There was some variation in the expression of CD166 (activated leukocyte cell adhesion molecule 1) between cell types (Table I).


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  		Table I. Mature and immature SC share a common cell surface marker phenotypea

 
The osteogenic and adipogenic differentiation ability of MSC, SF, DF, and CH was also tested. MSC were consistently able to differentiate into osteoblasts and adipocytes. Although SF displayed some multilineage differentiation potential, the proportion of cells differentiating between samples varied, thus suggesting the presence of variable numbers of progenitor cells. DF failed to differentiate along both osteogenic and adipogenic lineages. CH did not differentiate into osteoblasts, but did show some adipogenic differentiation potential (Fig. 1).


Figure 1
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  		FIGURE 1. Mature and immature SC possess variable differentiation abilities. The appropriate osteogenic and adipogenic conditioned medium was added to confluent monolayers of MSC, SF, DF, and CH for 21 days. Control cultures in normal growth medium were also maintained in parallel (data not shown). Osteogenic differentiation was assessed by alizarin red staining for calcium mineralization (top panel). Adipogenic differentiation was visualized using ORO staining for lipid vacuoles (bottom panel). Photomicrographs were taken at x200 magnification. Results are representative of at least three experiments per SC type.

 
SC inhibit PBMC proliferation

We then assessed the ability of the various types of SC to inhibit T cell proliferation in response to polyclonal stimuli and compared it with BM-derived MSC. The addition of MSC or SF, DF, LF, and CH to PBMC stimulated with PHA or anti-CD3/CD28 inhibited their proliferation in a dose-dependent fashion (*, p < 0.05) (Fig. 2A). On the contrary, the addition of endothelial cells or neurons did not inhibit T cell proliferation (Fig. 2B). This indicates that the immunosuppressive effect on T cells can be exerted by several types of immature and mature SC, but not by the parenchymal cells tested.


Figure 2
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  		FIGURE 2. SC inhibit lymphocyte proliferation. A, PBMC (1 x 105) were stimulated with either PHA (solid lines) or anti-CD3/CD28 beads (dashed lines) in the presence of increasing ratios of different stromal and parenchymal cells (B). MSC (PHA, n = 7; CD3, n = 10; 9 donors tested), SF (PHA, n = 4; CD3, n = 10; 7 donors tested), DF (PHA, n = 11; CD3, n = 12; 3 donors tested), LF (PHA, n = 7; CD3, n = 8; 1 donor tested), CH (n = 4; 2 donors tested), HUVEC (n = 4; 2 donors tested), and neurons (n = 6; 2 donors tested). Proliferation assays were incubated for 3 days, and for the final 15 h of culture [3H]TdR was added to wells. Results are expressed as percentage of proliferation, with the positive control at 100% (data not shown). Percentage of proliferation was calculated by dividing the cpm in the presence of stromal or parenchymal cells, by the cpm of the positive control (in the absence of stromal or parenchymal cells) and multiplying by 100. Results show mean ± SEM (*, p < 0.05).

 
SC do not inhibit the early phase of T cell activation

To determine at what stage SC exerted their inhibitory effect, we assessed the surface expression of activation markers on CD3+ lymphocytes. CD69 and CD25 expression was up-regulated on T cells upon stimulation with PHA for 24 h and remained unchanged in the presence of MSC or any of the other SC types tested, including SF, DF, and CH. Among the samples analyzed, there was no significant difference in the expression of both CD69 (p = 0.6712) and CD25 (p = 0.1479) on CD3+ lymphocytes stimulated in the presence or absence of SC (Fig. 3).


Figure 3
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  		FIGURE 3. SC do not inhibit the early activation of stimulated lymphocytes. PBMC were stimulated with PHA in the presence of MSC, SF, DF, or CH at a 1:5 ratio of SC:PBMC. After 24 h, PBMC were stained with mAbs to CD3, CD69, and CD25. Results show percentage of CD3+CD69+ and CD3+CD25+ lymphocytes in cultures of PBMC only (Figure 3, n = 3), PBMC + SC (Figure 3, n = 4), PBMC + PHA (Figure 3, n = 3), and PBMC + PHA + SC (Figure 3, n = 8).

 
SC inhibit the production of IFN-{gamma} and secretion of TNF-{alpha}

We then evaluated the ability of SC to interfere with TNF-{alpha} and IFN-{gamma} production by T cells in response to stimuli. The coculture of PBMC with MSC, SF, DF, or CH for 3 days before stimulation with PMA and ionomycin resulted in a significant down-regulation of IFN-{gamma} production. When PBMC were cultured with HUVEC, there was no inhibition of IFN-{gamma} production (Fig. 4A). The capacity to inhibit IFN-{gamma} production was comparable between the SC tested, and IFN-{gamma} levels were significantly reduced when compared with cells stimulated in the absence of SC (p < 0.05).


Figure 4
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  		FIGURE 4. SC inhibit the production and secretion of IFN-{gamma} and TNF-{alpha}. A, PBMC were cultured for 3 days in the absence or presence of MSC, SF, DF, or HUVEC at a 1:5 ratio. During the last 6 h of culture, PBMC were stimulated with PMA and ionomycin. PBMC were then surface stained for CD3, fixed, permeabilized, and stained for intracellular IFN-{gamma}. Results show the percentage of IFN-{gamma}-producing CD3+ lymphocytes in cultures of stimulated PBMC alone (Figure 4, n = 9) in the presence of MSC (Figure 4, n = 6), SF (Figure 4, n = 6), DF (Figure 4, n = 9), or HUVEC (Figure 4, n = 3). IFN-{gamma} production was significantly inhibited in the cultures containing SC when compared with the positive control (*, p < 0.05). B, Stimulated PBMC were cultured in triplicate in the absence or presence of an adherent layer of DF. Supernatants were removed from the wells at 24 and 48 h. TNF-{alpha} concentration (pg/ml) was measured in the supernatants by ELISA. Solid lines represent PHA, and dashed lines anti-CD3/CD28 stimlulation. Results show mean of triplicate values ± SEM.

 
Because our results did not show any difference between the inhibitory activities of the various SC tested, we selected DF as a source of SC for the following experiments. Cell-free supernatants from cultures in which DF were in contact with PHA or anti-CD3/CD28-stimulated PBMC for 24 and 48 h were examined for the concentration of TNF-{alpha} by ELISA. DF exerted a dose-dependent inhibition of TNF-{alpha} production, which was particularly evident at 48 h (Fig. 4B). TNF-{alpha} levels in cultures with DF and unstimulated PBMC, DF alone, or PBMC alone were below the detection limit of the assay (data not shown).

SC cause cell cycle arrest of activated PBMC

Because BM-derived MSC arrest T cells in the G0/G1 phases of the cell cycle (8), we decided to dissect the effect of different SC on the cell cycle of PHA-stimulated PBMC. After 3 days of PHA stimulation in the presence of DF, PBMC were arrested in the G0/G1 phase, with only a small proportion of cells found in S and M phases (Fig. 5). On the contrary, a higher number of PBMC stimulated in the absence of DF had entered S and M phases. We observed the same inhibitory effect using MSC and SF, whereas HUVEC did not affect cell cycle progression of stimulated PBMC.


Figure 5
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  		FIGURE 5. SC arrest the cell cycle of stimulated PBMC. A total of 1 x 106 PBMC/well was stimulated for 3 days with PHA in the absence or presence of DF or HUVEC at a 1:5 ratio. PBMC were fixed, stained for protein synthesis (FITC) and DNA synthesis (PI), and analyzed by FACs. Values show percentage of cells in each cell cycle phase. Results are representative of at least three experiments with similar result as the DF condition obtained using MSC and SF.

 
SC protect T cells from apoptosis and activation-induced cell death

The effect of SC on cell proliferation may or may not also translate into cell death. When unstimulated PBMC were cultured for 3 days in low serum concentration to induce apoptosis, ~7% of the cells stained positive for annexin V. PBMC cultured in the same conditions with MSC, SF, DF, or CH showed a trend toward reduced apoptosis in the presence of SC; however, this was not significant. When PBMC were stimulated with PHA at low serum concentration in the presence of MSC, SF, DF, or CH, the proportion of apoptotic cells was significantly reduced, as compared with the cultures in which activation-induced cell death was measured in the absence of SC (p = 0.0173). Such a protective effect is shown for CD4+ cells (Fig. 6A) with similar findings observed for CD8+ lymphocytes (data not shown).


Figure 6
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  		FIGURE 6. SC rescue lymphocytes from apoptosis. A, PBMC were cultured in low serum medium (2% FBS) either unstimulated in the absence (Figure 6, n = 5) or presence of SC (Figure 6, n = 5) or stimulated with PHA in the absence (Figure 6, n = 5) or presence of SC (Figure 6, n = 10). Cultures were incubated for 3 days at a 1:5 ratio of SC:PBMC, where appropriate. The percentage of CD4+/annexin V+ T cells was enumerated for each culture condition by FACS analysis. Similar results were obtained for all SC tested (MSC, SF, DF, and CH). Results show mean ± SEM (*, p = 0.0173). B, Representative plots showing apoptosis of CD3+ lymphocytes in cultures of unstimulated or PHA-stimulated PBMC, in the presence or absence of DF or HUVEC. The bottom left quadrant in each plot depicts live cells; bottom right quadrant, early apoptotic cells; and top right quadrant, late apoptotic cells. Values show percentage of cells in each quadrant.

 
By combining annexin V and PI staining, CD3+ lymphocytes were defined into populations of live cells (annexin V–/PI–), early apoptotic cells (annexin V+/PI–), and late apoptotic cells (annexin V+/PI+). The addition of DF to stimulated PBMC increased the percentage of cells in the live gate and decreased the proportion of both early and late apoptotic T cells, whereas the addition of HUVEC did not rescue PBMC from apoptosis (Fig. 6B).

Cell-cell contact is important for the SC antiproliferative effect

To determine whether cell-cell contact is required for the antiproliferative effect, cell-free supernatants were taken from different SC cultures (MSC, SF, and DF) and tested for their ability to inhibit PBMC proliferation. The supernatant from cultures of SC alone had no significant effect on PBMC proliferation. Therefore, we evaluated the supernatant derived from cultures in which SC were cultivated with unstimulated or PHA-stimulated PBMC. Only supernatant in which SC were in contact with activated PBMC produced a significant inhibition of T cell proliferation (p = 0.0079) (Fig. 7A). We observed similar findings for supernatant taken from cultures of SC and anti-CD3/CD28-stimulated PBMC (data not shown).


Figure 7
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  		FIGURE 7. The SC antiproliferative effect is mediated by cell-cell contact. A, Cultures of either SC alone (hatched bars, n = 7), SC + unstimulated PBMC (Figure 7, n = 7), or SC + PBMC + PHA (Figure 7, n = 6) were incubated for 3 days. SC and PBMC were cultured at a ratio of 1:5, where appropriate. Medium was then removed, the SC monolayer was rinsed twice with serum-free medium, and fresh culture medium was replaced. Cells were cultured for an additional 3 days, after which the resulting supernatant was removed and added, at 75% to a secondary, anti-CD3/CD28-stimulated, 3-day PBMC proliferation assay. For the final 15 h of culture, [3H]TdR was added to wells. Percentage of proliferation was calculated by dividing the cpm in the presence of supernatant by the cpm of the positive control (in the absence of supernatant) and multiplying by 100. Results show mean ± SEM (*, p < 0.05). B, 1 x 106 CFSE-labeled PBMC (solid round cells) were stimulated with PHA in the lower chamber of a Transwell (I). A total of 2 x 105 DF (shaded oval cells) was cocultured with the stimulated PBMC either in direct contact (II) or separated by a Transwell membrane (III). In some conditions, stimulated PBMC (hollow round cells) were also added on top of the DF monolayer (IV). Values show the percentage of proliferating cells. Plots are representative of at least three experiments with similar results obtained using MSC and SF.

 
These results were confirmed in a Transwell system (Fig. 7B). CFSE-labeled PBMC were plated in the bottom chamber and activated with PHA. DF were cultivated in the same well or physically separated in the upper chamber. When DF were in contact with PBMC, T cell proliferation was abrogated (Fig. 7BII). Proliferation remained unaffected when DF were physically separated by the Transwell membrane (Fig. 7BIII). However, if stimulated PBMC were also added to the upper chamber, in contact with the DF monolayer, cell proliferation in the bottom chamber was strongly inhibited (Fig. 7BIV). HUVEC did not exert any effect on PBMC proliferation, even when in contact with stimulated PBMC (data not shown). These results indicate that SC mediate their antiproliferative effect ultimately via soluble factors, but they first require a licensing step that involves cell contact.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Although it has been widely documented that MSC have potent immunomodulatory functions in vitro and in vivo (reviewed in Ref. 7) and clinical trials are already underway to evaluate their therapeutic efficacy, the physiological significance of such an effect remains poorly defined. The immunosuppressive activity of MSC is frequently cited as a unique function of the cells. However, until now, the possibility that other SC exhibited a similar effect was not clear. In this study, we demonstrate that mature SC from other tissues, at various stages of differentiation or containing variable proportions of progenitor cells (Fig. 1), are capable of exerting a powerful dose-dependent immunosuppressive activity on T cells, comparable, if not superior, to that of MSC (Fig. 2). Previous studies have proposed that MSC, differentiated in vitro, maintained their immunosuppressive effect (32, 33), but the contamination of residual undifferentiated MSC could not be excluded in those experiments. More recently, it was described that primary articular human CH were able to exert an inhibitory effect on anti-CD3-stimulated T lymphocytes, comparable to that of MSC, strongly supporting the data reported in this study (34). Therefore, the immunosuppressive effect of MSC is not a property confined to mesenchymal progenitor cells, but is also shared by terminally differentiated mesenchymal cells.

To acquire antiproliferative functions, SC require a licensing step. In fact, only the supernatants obtained from cultures in which SC were incubated with activated T cells displayed an immunosuppressive effect when added to secondary cultures in which T cells were induced to proliferate (Fig. 7). No effect was detectable using supernatants from cultures of SC alone. It is likely that monocytes play a primary role in licensing SC, as suggested by a recent study using MSC (35). We examined the role of IFN-{gamma} and IL-1 in the licensing effect on SC, because both cytokines have been suggested to facilitate the inhibitory effect of MSC (35, 36). We were unable to see any effect of blocking IFN-{gamma} in our system (data not shown). This would concur with findings in this study that the presence of SC inhibits the percentage of IFN-{gamma}-producing cells (Fig. 4). Groh et al. (35) demonstrated that by adding exogenous IL-1β, the inhibitory effect of MSC was enhanced. We observed that the addition of IL-1RA to stimulated PBMC cultures inhibited proliferation in the absence of SC (data not shown). This makes interpreting any possible effects of neutralizing IL-1 on licensed SC problematic.

The multitude of inhibitory soluble factors produced by MSC (9, 13, 15, 16, 37) suggests that the cellular and molecular composition of the environment, in which SC make contact with T cells, is of fundamental importance. This may explain the persistence of pathogenic T cells in the presence of a large number of SC, as is the case in inflammatory joint diseases (38, 39, 40).

Like their immature progenitors, SC induced in T cells a split anergy phenotype, whereby inhibition of proliferative responses is the major characteristic (10). We further observed that the effect of SC on cell cycle was associated with an enhanced survival of T cells when exposed to low serum conditions or after induction of activation-induced cell death (Fig. 6). Similarly, fibroblasts have previously been shown to rescue T lymphocytes from apoptosis induced by cytokine deprivation (41). The majority of studies using MSC also support our findings in this study. Although MSC inhibited TCR-dependent and independent proliferation, they did not induce apoptosis of T cells (9, 18, 36). In fact, T cells were still able to proliferate when rechallenged with Ag after the removal of MSC (9).

We propose that such antiproliferative and prosurvival effects are unique characteristics of cells of stromal origin, which may have evolved to regulate tissue inflammatory responses (42). Although, under particular conditions, parenchymal cells may impose reduced immunogenicity and even Ag-specific tolerance (43, 44), we did not detect any immunosuppressive activity of cells of endodermal (HUVEC) and ectodermal (neurons) origin (Figs. 2, 4, and 5). We have recently shown that the MSC-induced antiproliferative effect can also be exerted on nonimmune cells of various tissue origin (20). Therefore, it is plausible that, in parallel to their function in the BM (22, 45), SC in other tissues may provide a niche that regulates not only immune responses, but also the expansion, survival, and self-renewal of resident parenchymal cells.


   Acknowledgment
 
We thank Alexis Joannides (Department of Clinical Neurosciences, Centre for Brain Repair, Cambridge University, Cambridge, U.K.) for his assistance with the culture and use of neurons in experiments.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 S.J. is supported by a Medical Research Council studentship. Back

2 Address correspondence and reprint requests to Prof. Francesco Dazzi, Head of Stem Cell Biology, Department of Haematology, Division of Investigative Science, Faculty of Medicine, Imperial College London, Hammersmith Hospital, Du Cane Road, London. W12 0NN. E-mail address: f.dazzi{at}imperial.ac.uk Back

3 Abbreviations used in this paper: MSC, mesenchymal stem cell; BM, bone marrow; CH, chondrocyte; DF, dermal fibroblast; LF, lung fibroblast; ORO, oil red O; PI, propidium iodide; SC, stromal cell; SF, synovial fibroblast. Back

Received for publication March 8, 2007. Accepted for publication June 28, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

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