Immunomodulatory Properties of Mesenchymal Stem Cells Derived from Dental Pulp and Dental Follicle are Susceptible to Activation by Toll-Like Receptor Agonists

Abstract

Adult mesenchymal stem cells (MSCs) have recently become a potent tool in regenerative medicine. Due to certain shortcomings of obtaining bone marrow MSCs, alternate sources of MSCs have been sought. In this work, we studied MSCs from dental pulp (DP-MSCs) and dental follicle (DF-MSCs), isolated from the same tooth/donor, to define differences in their phenotypic properties, differentiation potential, and immunomodulatory activities. Both cell types showed colony-forming ability and expressed typical MSCs markers, but differed in the levels of their expression. DF-MSCs proliferated faster, contained cells larger in diameter, exhibited a higher potential to form adipocytes and a lower potential to form chondrocytes and osteoblasts, compared with DP-MSCs. In contrast to DF-MSCs, DP-MSCs produced the transforming growth factor (TGF)-β and suppressed proliferation of peripheral blood mononuclear cells, which could be neutralized with anti-TGF-β antibody. The treatment with toll-like receptor 3 (TLR3) agonist augmented the suppressive potential of both cell types and potentiated TGF-β and interleukin-6 secretions by these cells. TLR4 agonist augmented the suppressive potential of DF-MSCs and increased TGF-β production, but abrogated the immunosuppressive activity of DP-MSCs by inhibiting TGF-β production and the expression of indolamine-2,3-dioxygenase-1. Some of these effects correlated with the higher expression of TLR3 and TLR4 by DP-MSCs compared with DF-MSCs. When transplanted in imunocompetent xenogenic host, both cell types induced formation of granulomatous tissue. In conclusion, our results suggest that dental MSCs are functionally different and each of these functions should be further explored in vivo before their specific biomedical applications.

Introduction

A dult mesenchymal stem cells (MSCs) primarily isolated from bone marrow (BM) have become a potent tool in regenerative medicine over the past 2 decades [1]. Besides their capacity for differentiation in cell lineages of mesenchymal origin (adipocytes, osteoblasts, and chondrocytes), a body of evidence suggests their potential to differentiate into nonmesenchymal cell-lineages as well, such as neuron-like cells, hepatocytes, and pancreatic-like cells [2 –5]. Due to certain shortcomings of obtaining the BM-MSCs, including pain, morbidity, and low cell number on harvest, alternate sources of MSCs have been sought. Dental tissue is a potent reservoir of postnatal stem/progenitor cells that were isolated from dental pulp [6], exfoliated deciduous teeth [7], periodontal ligament [8], apical papilla [9], and dental follicle [10]. These postnatal populations possess mesenchymal-stem-cell-like qualities, including the capacity for self-renewal and multilineage differentiation potential [11]. However, the precise relationship among these cell populations remains unclear. Dental MSCs might be more committed, that is, restricted in their differentiation potency in comparison to BM-MSCs due to the interaction of dental mesenchyme with dental epithelium during tooth germ development and the absence of continuous dental tissue remodeling [11].

The characteristics that qualify MSCs as a favorable tool for cellular therapies of autoimmune and chronic diseases manifested both as inflammation and tissue injury is their ability to suppress alloreactive T, B, and NK cell responses [12]. In addition, cotransplantation of BM-MSCs with hematopoetic stem cells may improve the function of these cells and reduce the severity of graft-versus-host disease [13]. The mechanisms by which BM-MSCs suppress immune cells are still controversial and include secretion of transforming growth factor-β (TGF-β) [14], hepatocyte growth factor [15], nitric oxide [16], prostaglandin E2 [17], expression of immunosuppressive minor H antigen [human leukocyte antigen-G (HLA-G)] [18], and interferon (IFN)-γ-induced degradation of tryptophan [14,19].

Toll-like receptors (TLRs) are pattern recognition receptors, broadly distributed on cells throughout the immune system, and are key molecules bridging innate and adaptive immune responses [20]. Beside recognition of microbial molecule patterns, TLRs recognize some host molecules, also involved in the pathogenesis of autoimmune and chronic inflammatory diseases [21,22]. Recent reports suggest that MSCs express TLR1–8 [23] and that TLRs ligation modulates the proliferation and the differentiation of both human and mouse MSCs, indicating that TLRs are functional in MSC biology [23,24]. In addition, TLRs activation influences the immunological and migratory behavior of MSCs [25].

Although some functions of TLRs in MSCs biology are documented for BM-MSCs, similar analyses for dental MSCs are lacking. This aspect of dental MSCs biology is particularly important, because, during the potential transplantation into damaged or inflamed tissue, there is a high probability for transplanted cells to encounter different TLR activating signals [26 –28]. Whether this ligation of TLRs will support or suppress immunosuppressive ability of MSCs is of crucial importance for the positive outcome in cell therapy. Further, the possibility to modulate immunosuppressive properties and other properties of these cells in vitro by TLR ligation before their implantation could enhance the safety of their application. Recent studies showed controversial effects of TLR ligation on the immunosuppressive phenotype of BM-MSCs. Some studies showed that TLR ligation inhibits immunosuppressive properties of BM-MSCs [29], whereas others reported the opposite effects of the same TLR agonists [30]. Finally, one publication recently reported that different TLR agonists can promote either proinflammatory or immunosuppressive phenotype of hMSCs [31].

In this work, we comparatively studied the morphology, phenotype, proliferation rates, and differentiation potential of 2 human dental MSCs subtypes isolated from dental follicle (DF-MSCs) and dental pulp (DP-MSCs). Further, we sought to investigate whether DP-MSCs and DF-MSCs are immunomodulatory and whether these immune functions could be further modulated with TLR agonists. Also, we wondered what mechanisms were responsible for the immunomodulatory properties of these dental MSCs. To exclude the possibility that phenotypic and functional differences between DP-MSCs and DF-MSCs could be masked by individual variations, we compared phenotypic and functional characteristics of these 2 MSC subtypes isolated from the same donor/tooth. We found that DF-MSCs and DP-MSCs are different in terms of morphology, phenotype, differentiation potential, immunomodulatory properties, and response to TLR ligation. Further, some mechanisms by which these dental MSCs respond to TLR ligation seem to be quite different from those published for BM-MSCs.

Materials and Methods

Tissue samples and cell cultures

A wisdom tooth was extracted for orthodontic reasons from a young adult at the Department for Oral Surgery, Military Medical Academy, after the patient had signed the informed consent. DF-MSCs were isolated from dental follicles as previously described [32]. In brief, a human third molar was surgically removed before its eruption; the dental follicle was separated under sterile conditions from the mineralized tooth and minced with a scalpel. Subsequently, DP-MSCs were isolated from the dental pulp of the same tooth/donor as described elsewhere [6]. The pulp tissue and the dental follicle were digested in a Dulbecco's modified Eagle's medium (DMEM; Sigma, Munich, Germany) solution of type I collagenase (1 mg/mL; Sigma) and DNAase (25 μg/mL; Sigma) for 1 h at 37°C and 5% CO₂ atmosphere. The obtained cell suspensions were then rinsed in DMEM and plated at the density of 6,000 cells per 1 cm² in culture flasks (25 cm²). Nonadherent cells were removed by changing the cultivation medium. BM was obtained from volunteer donors who had provided informed consent, and BM-MSCs were generated as described elswhere [33]. For mixed leukocyte reactions (MLRs), allogenic peripheral blood mononuclear cells (PBMCs) were isolated from healthy donors who had signed informed consent by using a density gradient, as described elsewhere. DF-MSCs, DP-MSCs and BM-MSCs were cultured in the standard medium, composed of DMEM-low glucose (Sigma) supplemented with 10% fetal calf serum (FCS; Sigma), and antibiotics [penicillin/streptomycin (Galenika)/gentamicin (Panfarma)], 1% each), at 37°C, 5% CO₂. On reaching 70% confluence, passaging was performed by incubating the cells with 0.2% trypsin (Sigma) dissolved in 0.02% NaEDTA in phosphate-buffered saline (PBS) (1 mL/25 cm²). The cells were than harvested, rinsed in the standard medium, and plated at the density of 5,000–6,000 per 1 cm² in flasks. On multiplication, some of the cells were frozen in liquid nitrogen in 10% dimethyl sulphoxide (Sigma) in FCS and later used in the experiments. Passages from 4 to 9 were used in all the experiments.

Morphometric measurements

Morphometric measurements of adherent cells are often hampered by various irregular shapes and shape plasticity. Therefore, DP-MSCs and DF-MSCs were collected; cytospins were made using Shandon Cytospin Centrifuge (Thermo Scinetific) and stained with May-Grunwald-Giemsa (MGG) method. For diameter measurements, a total of 150 cells per dental MSC type were measured. Each cell was measured 4 times across different axes (which crossed each other at 45°). The mean diameter was calculated for each cell. The cells whose diameter varied more than 15% were excluded from further analysis. Measurements were performed by using the NIS-element D 2.30 software.

Colony forming units-fibroblasts assay

The colony forming units-fibroblasts assay was preformed as previously described [34]. DP-MSCs and DF-MSCs were harvested; and 25, 50, 100, or 200 cells were seeded in each well of a 6-well plate in 2 mL of standard medium. The cells were cultivated for 14 days while feeding the cells with the fresh medium twice a week. After the cultivation period, colonies were washed with PBS, air-dried, stained with MGG, and counted under the microscope. A colony forming units-fibroblast was defined as a group of at least 50 cells. The frequency of clonogenic cells was calculated as a number of colonies/number of seeded cells ×100.

Population doubling time

DF-MSCs and DP-MSCs from passages 5 and 8 were seeded at 1 × 10³ cells per well of a 6-well plate in standard medium for several intervals (0, 3, 6, 9, and 12 days) in duplicates. The cells were then trypsinazed, as just described, and counted in Tripan blue solution. Population doubling time was calculated between each interval using the formula: T _1/2 = (t _i − t _i − 1) × lg2/lg (N _i/N _i − 1). T_1/2 represents population doubling time, N _i and N _i − 1 represents number of cells harvested at the interval “i” and the previous interval “i − 1” at the time t _i and t _i − 1, respectively. N _i and N _i − 1 were averaged from the duplicates.

In vitro differentiation assays

Adipogenesis

DP-MSCs and DF-MSCs were seeded at the concentration of 2.5 × 10⁴/cm² on presterilized cover slips in 24-well plates containing 1 mL of a commercial mesenchymal stem cell growth medium (MSCG) medium (Lonza, Basel, Switzerland) in an incubator (37°C, 5% CO₂) until they reached 100% confluence (2-3 days). After that, the MSCG medium was replaced with an adipogenic induction medium (Lonza), followed by 3–4 days of incubation. The induction medium was than replaced with an adipogenic maintenance medium (Lonza) for the next 1–3 days of incubation. Differentiation into adipocytes lasted 17–25 days repeating the induction/maintenance cycle. The control cells were cultivated in an adipogenic maintenance medium only, replacing the medium every 3–4 days. The adipogenic differentiation was evaluated by staining the cells with Oil Red O (Sigma). Cover slips were washed in PBS, immediately dried with a vacuum pump, and placed in 60% isopropyl alcohol for 1 min. The cells were then stained with Oil Red O for 15 min and analyzed.

Chondrogenesis

DP-MSCs and DF-MSCs were cultured as pellets in sterile polypropylene cryotubes. The cells were collected, washed twice in an incomplete chondrogenic medium (Lonza) by centrifugation of the suspension at 150 g for 5 min and careful removal of the supernatants followed by vortexing. The cells were then resuspended in 0.5 mL of complete chondrogenic media (consisting of the incomplete medium supplemented with 10 ng/mL TGF-β3; R&D Systems, Minneapolis, MN), centrifuged at 150 g for 5 min, and cultured as pellets for the next 14–28 days, with medium replacement twice a week. The control cells were cultivated likewise in the MSCG medium. After the chondrogenic differentiation, the pellets were embedded in cryostat embedding medium (Bio-Optica) and frozen in liquid nitrogen until the analysis. Cryocuts, 7 μm thick, were made by using a cryomicrotome (Leica Cryocut 1850) and stained with Alcian blue for the light microscopy analysis.

Osteogenesis

DP-MSCs and DF-MSCs (both at 3 × 10³/cm²) were placed in 24-well plates containing 1 mL of the MSCG medium. After the next 24-h incubation, the medium was replaced with an osteogenic commercial medium (Lonza), whereas the control cells continued to reside in the MSCG medium. The cultivation into osteoblasts lasted 3–4 weeks, with medium replacement twice a week. For mineralized nodule assay, the cultured cells were stained with 1% alizarin red (Sigma) in distilled water. The size of calcified nodules and the percentage of alizarin red-positive area over total area were assessed by analyzing at least 10 fields of view under the microscope. The data were processed using NIS-element D 2.30 software. Besides, the cells were trypsinized, and cytospines were prepared. The osteogenic differentiation was evaluated after the alkaline phosphatase (ALP) staining. In brief, an incubation buffer, containing Fast Red (1 mg/mL, Sigma) dissolved in a mixture of (V/V ratio 1/50) naphtol-dimethyl formamide (10 mg/mL, Sigma) and tris(hydroxymethyl) aminomethane (TRIS, pH 8.3; Sigma), was added onto each slide, followed by incubation at 37°C, 5% CO₂ for 1 h. Additionally, the cells were fixed and stained with anti-osteopontin antibody, as described in section Immunocytochemistry. All stained cells were analyzed using a light microscope (Olympus IX51) equipped with a camera (Nikon DXM1200C Microscope Camera).

Xenogenic transplantation

The experiment was preformed with the approval of the Military Medical Academy Ethical Committee, which strictly follows European Community Guidelines (EEC Directive 1986; 86/609 EEC). One-day-old BALB/c mice were subcutaneously injected behind the ear with 5 × 10⁵ of DP-MSCs and DF-MSCs as described elsewhere [35]. The implants were harvested after 18 days after euthanasia with ether. The samples were fixed in 3% glutaraldehyde for 3 h and demineralized for 7 days in 13% EDTA (pH 7.2) supplemented with 1% glutaraldehyde and then paraffin embedded. Paraffin cuts, 8 μm thick, were stained using the standard hematoxilin eosine and masson trichrom staining and analyzed by light microscopy.

Flow cytometry

The flow cytometry analysis was performed using the following monoclonal antibodies (mAbs): anti-CD14-fluorescin isocyanate (FITC), anti-CD45-FITC, anti-CD29-FITC, anti-CD105-FITC, anti-CD44-FITC, anti-CD19-FITC, anti-CD34-phycoerithryne (PE) (all from Immunotools), anti-CD146-FITC, anti-CD90-FITC, anti-CD106-FITC, anti-CD56-FITC, mouse IgG1a negative control–FITC, anti-HLA-DR-PE, and mouse IgG1a negative control–PE (all from Serotec). Additionally, the indirect immunofluorescence was performed using primary, anti-CD73 (SantaCruz Biotechnology), and anti-STRO-1 (Millipore/Chemicon) mAbs followed by secondary anti-mouse IgG1-FITC mAb (Serotec). BM-MSCs, DP-MSCs, and DF-MSCs were collected and washed once in PBS containing 2% FCS and 0.1% sodium azide. The cells were then incubated with primary mAbs, using the concentrations prepared in PBS/sodium azide as recommended by the manufacturer, for 1 h at 4°C. The incubation with the secondary mAb, for the indirect immunofluorescence, lasted 15–30 min at 4°C. The cells were then washed in PBS/sodium azide and subsequently analyzed by using a flow cytometer (Coulter XL). The cells were gated according to the cell-specific forward scatter/side scatter parameters, and the isotype control was done for each cell type.

Immunocytochemistry

The immunocytochemistry was performed using the following mAbs: anti-CD14, anti-CD45, anti-CD29, anti-CD105, anti CD34 (all from Dako); anti-CD44, anti-CD19, anti CD56 (all from NovoCastra), anti-CD90, and anti-CD106 (all from Serotec); anti-CD146, anti-STRO-1 (all from Millipore/Chemicon), and anti-CD73 (SantaCruz Biotechnology). Additionally, anti-osteopontin antibody (Dako) was used for staining the cells obtained during osteogenic differentiation. The cells were harvested, and cytospines were prepared from each cell type (0.1–1 × 10⁴ of cells per cytospine). The prepared samples were fixed in 2% pararosaniline [36] for 2 min at the room temperature, washed with PBS for 10 min, and the endogenous peroxydase was blocked with 0.5% H₂0₂ for 10 min. After each subsequent incubation step, the slides were washed with Tris-buffered saline/0.5% bovine serum albumin (Sigma)/0.05% Tween-20 (Sigma) for 5 min. The samples were than incubated with goat serum (Dako) (diluted in PBS 1:2) for 20 min and washed. Subsequently, the samples were incubated with the primary mAbs for 60 min in a water bath. Primary antibodies were detected using a sensitive EnVision + kit (Dako), as recommended by the manufacturer. Finally, DAB + kit (Dako) was used as a chromogen. Appropriate controls included the omission of primary antibodies or the use of irrelevant Abs. Finally, the slides were counterstained with hematoxylin, mounted in Keiser gel, and analyzed by light microscopy.

Mixed leukocyte reactions

The modulatory capacity of DP-MSC and DF-MSCs on the proliferation of PBMCs was evaluated in co-culture. Dental MSCs (1 × 10⁴/well) were seeded in a flat-bottom 96-well plate in RPMI (Sigma)/10% FCS to adhere for 6 h. After that, 25 μg/mL polyinosinic:polycytidylic acid [poly(I:C)] or 200 ng/mL lipopolysaccharide (LPS) (both from Sigma) were added to the appropriate wells for the next 24 h. The control MSCs were left untreated. All cells were then washed twice in PBS with 2% FCS, treated with mitomycin C (Bristol Caribbean) at the final concentration of 25 μg/mL for 30 min, and then washed 4 times with PBS/FCS. Freshly isolated PBMCs (1 × 10⁵/well) were stimulated with 250 ng/mL phytohemagglutinin (PHA; Serva) and immediately added to the wells containing dental MSCs. The effect of MSCs on the proliferation of PBMC was compared with the control cultures containing PHA-stimulated PBMCs without the presence of dental MSCs. The effect of TLR agonists was evaluated by comparing the immunosuppressive effect of MSCs pretreated with TLR agonists with that of untreated MSCs. In some experiments, an anti-TGF-β neutralizing antibody (R&D Systems) was added to MSC/PMBC co-culture in order to evaluate the role of TGF-β in immunosuppression. This was evaluated by comparing the proliferation in MSC/PBMC co-cultures treated with anti-TGF-β and the proliferation of control PHA-stimulated PBMCs treated the same way. PHA-stimulated PBMCs treated with TGF-β (1 ng/mL; R&D Systems) were used as a specific control for the neutralizing activity of the anti-TGF-β antibody. Three-day MLRs were preformed, and the cultures were pulsed with 1 μCi/well [3H] thymidine (6.7 Ci/mmol; Amersham Biosciences) for the last 18 h. The cells were then harvested, and the radionuclide uptake was measured by scintillation counting (Beckman; LS5000TB scintillation counter). The counts of nonstimulated PBMCs and mitomycin-treated dental MSCs, cultured separately, were subtracted from the counts of PBMC/MSC co-cultures. All cultures were done in quadruplicate.

Detection of indolamine-2,3-dioxygenase-1 expression

DP-MSCs and DF-MSCs were cultivated with TLR agonists as in MLR. The cells were then collected, washed in 10% FCS/RPMI, and counted in 1% Trypan Blue in physiological solution. The samples were fixed with Fix/Perm medium A (Caltag Laboratories) and washed in PBS once, according to the manufacturer's protocol. After that, the cells were incubated with 5% inactivated donkey serum for 3 min followed by 20 min incubation with a goat anti-human indolamine-2,3-dioxygenase-1 (IDO-1) polyclonal antibody (Santa Cruz Biotechnology) diluted in Fix/Perm medium B (Caltag Laboratories). For the control, an isotype antibody was used instead of the goat anti-human IDO-1 Ab. The cells were then washed in PBS/sodium azide once and incubated with a donkey anti-goat IgG:biotin (Serotec) in medium B for 15 min. After the final wash, the cells were incubated with streptavidine:PE (Serotec) in medium B for 15 min and analyzed by flow cytometry. Permeabilized cells were gated according to the predetermined position of permeabilized CD105-FITC positive cells and additionally confirmed by CD105/IDO-1 double staining.

Reverse transcriptase–polymerase chain reaction

To determine the levels of TLR3 and TLR4 mRNA expression, DP-MSCs and DF-MSCs were harvested, washed in PBS, and the pellets were resuspended in an RNA-free solution. The samples were then frozen at −70°C until the RNA isolation. Total RNA isolation was performed using the RNeasy kit (Qiagen), followed by DNase treatment (Fermentas) according the manufacturer's protocols. Reverse transcription was performed with the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems). PCR reaction of 1 μg cDNA was conducted with AmpliTaq Gold DNA Polymerase (Applied Biosystems) using specific primers: TLR3 forward primer 5′-AGC ATA GCA GCC TCT GCT TC-3′ and reverse primer 5′-CCT TCC CCT TCT CTC TTC TTC C-3′; TLR4 forward primer 5′-CCT AAG GAA ACC TGA TTA ACA-3′ and reverse primer 5′-GAT ATT AGC TTA TAG GCA AGA-3′; β-actin forward primer 5′-TCA CCC ACA CTG TGC CCC ATC TAC GA-3′ and reverse primer 5′-CAG CGG AAC CGC TCA TTG CCA ATG G-3′; all bought form MWG-Biotech AG, Ebensburg Germany. The amplification was carried out at 94°C for 30 s/55°C for 30 s/72°C for 45 s for 40 cycles using Eppendorf PCR System. The PCR products, DNA Marker 50 bp Ladder (Sigma) and nontemplate controls, were separated on a 2% agarose and stained with ethidium bromide. Images were taken by ChemiImager 4400 (Alpha Innotech) and analyzed with ImageQuant TL software (Molecular Dynamics). The levels of TLR3 and TLR4 mRNA were expressed as relative values of β-actin expression.

Cytokine detection

Dental MSCs were cultivated with TLR agonists as in MLR. After that, the supernatants were collected and frozen at −40°C for the analysis by spectrophotometry (Behring ELISA Processor II). To determine whether the levels of cytokines secreted by MSCs were changed after the removal of TLR stimuli and the mitomycin treatment, dental MSCs were treated as in MLR, but instead of PBMCs, fresh medium was added to each well. After 24 h of incubation, supernatants were collected and analyzed. Concentrations of TGF-β and interleukin-6 (IL-6) were determined from supernatants using the commercial ELISA kits (R&D Systems) and by calculating the unknown concentrations from the standard curves.

Statistical analysis

All values are given as mean ± standard deviation. The number of samples was 3–6, if not indicated otherwise. The 2-tailed unpaired t test and analysis of variance tests were used for evaluating the differences between the experimental and corresponding control samples. The values P < 0.05 were considered statistically significant. Graphical presentation of the results was prepared in the GraphPad Prism 5.0 software. Flow cytometry data were analyzed using FlowJo 5.7. The images were processed using the NIS-element D 2.30 software and later edited in Photoshop 6.0.

Results

Morphological properties, proliferation, and differentiation potential of DP-MSCs and DF-MSCs

Both DP-MSCs and DF-MSCs showed typical various-shaped fibroblast-like morphology (Fig. 1A), common to adult MSCs isolated from other tissues [37]. Morphometric measurements of adherent cells are often hampered by various irregular shapes and shape plasticity. Therefore, cytospins were made and stained with MGG (Fig. 1B), which enabled easy diameter measurement of MSCs types (Fig. 1C), as described in Materials and Methods. MSCs significantly differed in size, DP-MSCs being 23.95 ± 5.6 μm in diameter (mean ± standard deviation) (N = 150), whereas the diameter of DF-MSCs was 26.58 ± 6.16 μm (N = 150) (P < 0.005).

FIG. 1.

Morphology of DP-MSCs and DF-MSCs. (A) Fibroblast-like shapes of dental MSCs. (B) May-Grunwald-Giemsa staining of dental MSC cytospins. (C) Frequency distribution of dental MSCs' diameter expressed as the percentage of total count (150 cells per dental MSC type). DP-MSCs, dental pulp-mesenchymal stem cells; DF-MSCs, dental follicle-mesenchymal stem cells.

When cultured at a low cell density, DF-MSCs and DP-MSCs both formed adherent clonogenic cell clusters (see Supplementary Fig. S1; Supplementary Data are available online at www.liebertonline.com/scd), similar to those observed for various MSC populations. The frequency of colony-forming cells within DF-MSCs and DP-MSCs population was similar (36.5 ± 15.8 and 34.0 ± 6.8, respectively).

Population doubling time of DF-MSCs was shorter than the one of DP-MSC at both passages 5 (data not shown) and 8 (see Supplementary Fig. S2), suggesting a higher proliferation rate of DF-MSCs.

To determine differentiation potential of DP-MSCs and DF-MSCs, we performed differentiation assays using commercially available mediums. Both cell types were able to differentiate into mesodermal cell lineages that is, osteoblasts, adipocytes, and chondrocytes (Fig. 2).

FIG. 2.

Differentiation potential of DP-MSCs and DF-MSCs. (A) Adipogenic differentiation. Oil Red O staining of dental MSCs cultivated on cover-slips under the adipogenic differentiation conditions. (B) Chondrogenic differentiation. Alcian blue staining of cryocuts of dental MSCs pellets cultivated under the chondrogenic differentiation conditions. (C) and (D) Osteogenic differentiation. Osteogenesis was evaluated by detecting the activity of alkaline phosphatase (C) and by immunocytochemistry, using an anti-osteopontin antibody (D). Representative images from 2 independent experiments are shown. Color images available online at www.liebertonline.com/scd.

Differentiation into adipocytes was estimated after 3 weeks of cultivation in an adipogenic medium by Oil Red O staining. DF-MSCs had a larger capacity to differentiate into adipocytes, as assessed by the higher number of accumulated lipid droplets and their larger diameter, compared with the ones accumulated in DP-MSCs (Fig. 2A).

Chondrogenesis was estimated after staining the pellet culture kept in chondrogenic media for 3 weeks, with Alcian blue, which stains proteoglycan-rich extracellular matrix. As shown in Fig. 2B, both DF-MSCs and DP-MSCs were able to differentiate into chondrocytes. DP-MSCs were more efficient in that process, as judged by stronger Alcian blue staining.

The potential of dental MSCs to form osteoblasts was estimated by evaluating the endogenous ALP activity and the expression of osteopontin, a prominent component of the mineralized extracellular matrices of bones and teeth. Twenty to forty percent of both dental MSCs subtypes showed endogenous ALP activity after a 3-week cultivation period in osteogenic medium. A higher percentage of DP-MSCs possessed ALP activity in comparison to DF-MSCs after the differentiation, although DF-MSCs contained individual cells that showed strong positivity for ALP (Fig. 2C). Similar results were obtained by analyzing the expression of osteopontin. Nearly all DP-MSCs and 85% of DF-MSCs showed strong/moderate membranous and intracellular staining with an anti-osteopontin mAb after osteogenesis induction (Fig. 2D). In contrast, control cells showed weak expression of osteopontin, predominantly localized perinucleary (Fig. 2D). Alizarin Red-positive nodule formation in DP-MSCs and DF-MSCs was analyzed after the osteogenesis differentiation, which confirmed calcium accumulation in vitro (see Supplementary Fig. S3). Alizarin-positive area in DP-MSCs cultures was significantly larger compared with the area observed in DF-MSCs cultures (P < 0.05).

Cumulatively, we concluded that DP-MSCs were more potent to form osteoblast- and chondrocyte-like cells after the differentiation in appropriate media, whereas DF-MSCs had a stronger potential to form adipocyte-like cells.

Next, we wanted to evaluate the differentiation potential of DP-MSCs and DF-MSCs in vivo by implanting these cells subcutaneously behind the ear of 1-day old BALB/c mice. After 18 days, the implants were shown to induce granulomatouse tissue formation characteristic for xenotransplant rejection. At the center of the implants that was predominantly composed of death cells, no sign of differentiated structure was observed. At the periphery, cellular infiltrate predominantly composed of macrophages, giant cells, and various inflammatory cells (see Supplementary Fig. S4) could be seen. The implants were largely enclosed in a fibrous capsule of connective tissue rich in collagen, as detected by Masson trichrome staining (data not shown).

Phenotypic characteristics of DP-MSCs and DF-MSCs

To assess whether the phenotypes of DP-MSCs and DF-MSCs correspond to the phenotype of other MSCs, we analyzed the expression of molecules crucial for the identification of MSCs by immunocytostaining and flow cytometry (Fig. 3). Neither cell type expressed CD34, CD45, HLA-DR, CD14, and CD19 (data not shown). Similar expression of CD44, CD29, and CD46 was observed between DP-MSCs and DF-MSCs; and the percentage of positive cells for these molecules was 75%–100%. The expression of CD106 could be seen on less than 0.3% of cells in both cell types. DF-MSCs were more prominently positive for CD73, CD90, and CD105 in comparison to DP-MSCs. In contrast to those markers, a higher percentage of DP-MSCs was positive for CD146 and CD56 compared with DF-MSCs. The expression of STRO-1 and CD271 was detected only by immunocytochemistry; and DF-MSCs contained a higher percentage of positive cells compared with DP-MSCs. BM-MSCs were used as a positive control for all tested markers (see Supplementary Fig. S5).

FIG. 3.

Phenotypic properties of (A) DP-MSCs and (B) DF-MSCs. Immunocytochemistry and flow cytometry data of dental MSCs labeled with primary or immunoglobulin isotype control antibodies were shown. The indicated percentage of positive cells corresponds to the data obtained by flow cytometry. The percentage of positive cells obtained by light microscopy analysis is given in the text unless it was different from the data obtained by flow cytometry. Scale bars represent 50 μm. Representative images are shown from 2 independent experiments.

Immunosuppressive properties of DP-MSCs and DF-MSCs in vitro

The immunosuppressive potential of DP-MSCs and DF-MSCs was studied in a co-culture with allogenic PHA-stimulated PBMCs. As shown in Fig. 4, DP-MSCs slightly suppressed the proliferation of PBMCs at 1/10 (MSC/PBMC) ratio (P < 0.05), whereas under the same conditions, DF-MSCs were nonmodulatory.

FIG. 4.

Immunomodulatory properties of DP-MSCs and DF-MSCs. Dental MSCs (1 × 10⁴ per well) either pretreated with TLR agonists [25 μg/mL poly(I:C) or 200 ng/mL LPS] for 24 h (black bars) or untreated (gray bars) were cultivated for 3 days with PBMCs (1 × 10⁵/well) stimulated with 250 ng/mL phytohemagglutinin, as described in Materials and Methods. Controls were the cultures of phytohemagglutinin-stimulated PBMCs cultivated without MSCs (white bars). All cultures were done in quadruplicates calculating the mean ± SD. The representative results, out of 4 experiments with similar results, are shown. *P < 0.05; **P < 0.01 compared with corresponding controls. SD, standard deviation. TLR, toll-like receptor; poly(I:C), polyinosinic:polycytidylic acid; LPS, lipopolysaccharide; PBMCs, peripheral blood mononuclear cells.

To determine whether TLR activation changes the MSCs' immunomodulatory phenotype, dental MSCs were pretreated with poly(I:C), a TLR3 agonist, or LPS, a TLR4 agonist, for 24 h; thoroughly washed; and then co-cultivated with PBMCs. Additionally, we confirmed that these dental MSCs expressed mRNA for both TLR-3 and TLR-4. Compared with DF-MSCs, DP-MSCs expressed 1.3 and 2.6 times more mRNA for TLR3 and TLR4, respectively (see Supplementary Fig. S6). The pretreatment of MSCs with 25 μg/mL poly(I:C) for 24 h augmented the suppressive effect of DP-MSCs on the proliferation of PBMCs and made DF-MSCs immunosuppressive. DF-MSCs, pretreated with LPS, also suppressed the proliferation of PBMCs, whereas, surprisingly, LPS pretreatment abrogated the suppressive effect of DP-MSCs (Fig. 4). Cumulatively, the pretreatment of MSCs with either poly(I:C) or LPS significantly influenced the modulatory properties of MSCs on the proliferation of PBMCs.

Mechanisms of immunosuppressive properties of DP-MSCs and DF-MSCs in vitro

To determine the mechanisms responsible for the observed induction of the immunomodulatory phenotype of dental MSCs after TLR activation, the effect of poly(I:C) and LPS on the secretion of TGF-β, an immunosuppressive cytokine, by DP-MSCs and DF-MSCs, was studied. In the absence of TLR agonists, DF-MSCs did not secrete detectable amounts of TGF-β in contrast to DP-MSCs, which produced 325 ± 46 pg/mL per 5 × 10⁴ cells of this cytokine (Fig. 5A). Both poly(I:C) and LPS stimulated the secretion of TGF-β by DF-MSCs. Poly(I:C) almost doubled TGF-β secretion by DP-MSCs, but the production of this cytokine was reduced after the LPS treatment, in comparison to the untreated DP-MSCs. These dental MSCs were able to secrete comparable levels of TGF-β even after the stimulus withdrawal and the mitomycin treatment for 24 h (data not shown).

FIG. 5.

The effect of TLR agonists on the production of (A) TGF-β and (B) IL-6 by DP-MSCs and DF-MSCs. The levels of TGF-β and IL-6 were determined by enzyme-linked immunosorbent assay from culture supernatants of dental MSCs (1 × 10⁴/well) that were either treated with TLR agonists [25 μg/mL poly(I:C) or 200 ng/mL LPS] for 24 h (black bars) or left untreated (gray bars). All cultures and measurements were done in quadruplicates calculating the mean ± SD. The representative results, out of 4 experiments with similar results, are shown. *P < 0.05; **P < 0.01; ***P < 0.005 compared with corresponding controls. TGF-β, transforming growth factor-β; IL-6, interleukin-6.

To examine the relationship between the production of TGF-β and cellular proliferation, a neutralizing anti-TGF-β antibody was added to PBMC cultures. The antibody completely restored the proliferation of PBMC suppressed by DP-MSCs as well as TGF-β-induced inhibition of cellular proliferation, used as a positive control. However, the inhibition caused by poly(I:C)-treated DP-MSCs and DF-MSCs was not completely normalized (Table 1).

Table 1.

Effect of Anti-Transforming Growth Factor-β Neutralizing Antibody on the Proliferation of Peripheral Blood Mononuclear Cells in Presence of Dental Pulp-Mesenchymal Stem Cells and Dental Follicle-Mesenchymal Stem Cells

	Control	+ anti-TGF-β mAb
PBMCs	100.0% ± 14.1%	106.1% ± 9.2%
PBMCs + TGF-β (1 ng/mL)	42.1% ± 6.2%^a	89.1% ± 4.9%
PBMC/DP-MSCs	72.5% ± 9.3%^b	103.4% ± 11.3%
PBMC/DP-MSCs poly(I:C)	54.2% ± 16.2%^c	76.4% ± 8.5%^b
PBMCs/DF-MSCs	123.2% ± 10.24%	116.4% ± 18.2%
PBMCs/DF-MSCs poly(I:C)	58.33% ± 10.5%^c	80.4% ± 12.2%^b

Dental MSCs (1 × 10⁴/well) either pretreated with poly(I:C) (25 μg/mL) for 24 h or left untreated were cultivated for 3 days with phytohemagglutinin-stimulated PBMCs (1 × 10⁵/well) in presence of anti-TGF-β antibody (10 ng/mL), as described in Materials and Methods. Phytohemagglutinin-stimulated PBMCs treated with TGF-β (1 ng/mL) were used as a specific control for the neutralizing activity. The cultures were done in quadruplicates calculating the mean ± standard deviation. The representative results, out of 2 experiments with similar results, are shown.

P < 0.005;^b P < 0.05;^c P < 0.01 compared with corresponding controls.

TGF-β, transforming growth factor β; mAb, monoclonal antibodies; PBMC, peripheral blood mononuclear cells; DP-MSCs, dental pulp-mesenchymal stem cells; poly(I:C), polyinosinic:polycytidylic acid; DF-MSCs, dental follicle-mesenchymal stem cells.

These results suggest that TGF-β could not be the only factor responsible for the immunosuppressive activity of dental MSCs. Therefore, we studied the expression of IDO-1 in the dental MSCs types. In the absence of TLR activation, a higher percentage of DP-MSCs expressed IDO-1 in comparison to DF-MSCs (Fig. 6). Poly(I:C) did not alter the expression of IDO-1 by these cells. In contrast, LPS decreased the percentage of IDO-1 expressing DF-MSCs and DP-MSCs, 1.4 and 3.2 times, respectively.

FIG. 6.

The effect of TLR agonists on the expression of IDO-1 by DP-MSCs and DF-MSCs. Dental MSCs (1 × 10⁴/well) that were either treated with TLR agonists [25 μg/mL poly(I:C) or 200 ng/mL LPS] for 24 h or left untreated were labeled with anti-IDO-1 antibody or immunoglobulin isotype control and analyzed by flow cytometry, as described in Material and Methods. The representative results, out of 4 experiments with similar results, are shown. IDO-1, indolamine-2,3-dioxygenase-1; PE, phycoerithryne.

Some articles suggest that the TLR treatment of BM-MSCs can inhibit the immunosuppressive phenotype of these cells and trigger the proinflammatory phenotype [38]. Therefore, beside the levels of TGF-β, we determined the level of IL-6 (Fig. 5B.). Nonstimulated and LPS-stimulated DP-MSCs and DF-MSCs secreted hardly detectable levels of IL-6. In contrast, poly(I:C) induced a sudden burst of IL-6 secretion by both dental MSCs in spite of the inhibited cellular proliferation.

Discussion

To date, most cell-based therapies were conducted using BM-MSCs [2] and, more recently, using adipose-derived stem cells [39]. A growing body of evidence suggests that dental tissue is another potent source of alternate, postnatal MSCs. The present study comparatively evaluates 2 human dental MSCs subtypes, DF-MSCs and DP-MSCs, in terms of morphology, clonogenic capacity, proliferation rates, differentiation potential, phenotype, and immunomodulatory properties, to encourage their potential application in cell-based therapies. To exclude the possibility that differences between DP-MSCs and DF-MSCs are due to individual variations, we compared these 2 subtypes isolated from the same donor/tooth.

We showed that both of these dental MSCs subtypes possessed fibroblast-like morphology typical for MSCs [37], colony-forming ability, and were able to differentiate into osteoblast-, adipocyte- and chondrocyte-like cells. These results were confirmed by others as well (reviewed in [11]). Although chondrogenic potential of DF-MSCs is still in doubt [32,40], our study clearly shows that DF-MSCs also have the multilineage differentiation capacity in vitro. When comparing the 2 subtypes, DP-MSCs exhibited a higher potential to form chondrocyte-like cells, whereas DF-MSCs exhibited a higher potential to form adipocyte-like cells. Both DP-MSCs and DF-MSCs are comprised of heterogeneous cell subpopulations shown to have different proliferation rates, morphology, or differentiation potential [11,41]. Recently, Battula et al. showed that a CD271^bright BM-MSC subpopulation can be sorted, according to the specific CD56 epitop, in 2 subpopulations having different potential to form adipocytes (CD56⁻ cells) or chondrocytes (CD56⁺ cells). In addition, a higher percentage of the CD56⁺ subpopulation was positive for ALP after induction of osteogenesis [42]. These results are in line with our results regarding dental MSCs, showing that DP-MSCs contained more CD56⁺ cells compared with DF-MSCs. To prove this hypothesis, a separation into CD56⁺ and CD56⁻ subpopulations of both dental MSC types and the subsequent evaluation of their differentiation potential is necessary in further studies. Although DP-MSCs and DF-MSCs were able to differentiate in vitro, no signs of differentiated structure could be seen in vivo after the xenogenic transplantation of these cells. Xenogenic transplant reaction, characterized by cellular infiltrates and encapsulation of the implanted tissue, could be a reason for this lack of differentiation in vivo. Several studies also reported that human BM-MSCs or ADPCs transplanted into imunocompetent rodents induce similar immunologic reaction as observed in our study [43,44]. Therefore, differentiation potential of DP-MSCs and DF-MSCs should be explored in immunodeficient animal model in future studies.

DF-MSCs and DP-MSCs were identified as CD29, CD44, CD46, CD90, CD105, CD73, and STRO-1 positive cells that do not express CD34, CD45, HLA-DR, CD19, and CD14. Thus, together with the multilieage differentiation potential of these cells, the minimal criteria for defining multipotent MSCs are fulfilled [45]. This is also in accordance with the previously described phenotype of dental MSCs [11]. Additionally, we showed some specificity regarding the expression of CD146 and CD271. Namely, 57% of DP-MSCs and 15% of DF-MSCs were positive for the CD146 molecule, although its expression was previously attributed only to DP-MSCs [11]. Also, the immunocytochemistry analysis showed that 56% of DP-MSCs and 63% of DF-MSCs expressed CD271 (low-affinity nerve growth factor receptor, p75 neurotrophin receptor). To our knowledge, this is the first time to describe the CD271 positive subpopulation within DF-MSCs. Quirici et al. showed that CD271 sorted cells are a highly homogeneous subpopulation of cells within BM-MSCs with a high proliferative capacity and the potential for multilineage differentiation [46]. The role of CD271-positive cells in DP-MSCs and DF-MSCs is still unclear and needs further investigation. Isolation of CD271-positive subpopulations from these dental MSCs could be compromised by its low membranous expression (around 1%). Low membranous expression of CD271 is in agreement with that of Soncini et al., who showed that the membranous expression of CD271 on BM-MSCs, chorionic membrane-, and amnion membrane-derived MSCs is down-regulated during culture passaging [47]. To determine whether CD271-positive sorting could enrich dental MSCs from the starting cell population, ex vivo isolated cells or the cells at early passages should be used. The finding that DF-MSCs proliferated faster and expressed higher levels of CD90, CD105, CD73, CD271, and STRO-1 compared with DP-MSCs suggests that DF-MSCs contain more undifferentiated cells. This is in line with the knowledge that Notch-1, which affects self-renewal and lineage-specific differentiation [48] of both stem cells and precursor cells, is expressed by DF-MSCs [10]. Further, a finding that the constitutive active Notch-1 in mesenchymal cells suppressed osteoblastic differentiation in vitro [49] could further explain the lower potential of DF-MSCs for differentiation into osteoblasts, compared with DP-MSCs.

A growing body of evidence has demonstrated that BM-MSCs have profound immunomodulatory and anti-inflammatory effects both in vitro and in vivo via inhibiting the proliferation and function of several major types of innate and adaptive immune cells. Recent reports suggest that dental MSCs, such as those isolated from human gingival tissue [50], periodontal ligament [51] and dental pulp [52] can suppress mitogen- or allogenic-stimulated proliferation of PBMCs or T cells in vitro at a MSCs/PBMC ratio higher than 1/10. A lower MSCs number in culture was nonsuppressive. Further, it has been shown that hMSCs could even be allostimulatory in an allogenic MLR and induce IL-2 and IFN-γ production by PBMCs at a 1/10 cell ratio [53]. We used a lower MSC/PBMC ratio (1/10), which could be more relevant in vivo [54], and showed that DP-MSCs are immunosuppressive, whereas DF-MSCs did not significantly change cellular proliferation.

The effect of TLR ligation on immunosuppressive properties has been previously studied using BM-MSCs [23,25,29,30]; but to our knowledge, this is the first time it is studied on dental MSCs. For the first time we showed and comparatively evaluated the expression of mRNA encoding for TLR3 and TLR4 in DP-MSCs and DF-MSCs. During potential transplantation of MSCs into damaged or inflamed tissue, there is a high probability for transplanted cells to encounter additional signals, such as endogenous/host and infection agents that can activate TLR pathways [26 –28]. Therefore, the role of these receptors in immunomodulatory properties of MSCs is particularly important to evaluate before their application in cell therapies. Recent studies showed controversial effects of TLR ligation on the immunosuppressive phenotype of BM-MSCs. Some of the studies demonstrated that TLR3 and TLR4 signaling inhibits the immunosuppressive effect of BM-MSCs on the proliferation of CD4⁺ T cells [29], whereas others showed that activation of the same TLRs enhanced the immunosuppressive phenotype of BM-MSCs [30]. In accordance with the latter study, we showed that both poly(I:C)- and LPS-pretreated DF-MSCs suppressed the proliferation of PBMCs at the cell ratios that were not suppressive under basal conditions. The suppressive potential of DP-MSCs was significantly augmented after the pretreatment with poly(I:C), whereas LPS treatment had no such effect. The difference between DP-MSCs and DF-MSCs in response to LPS pretreatment could be a consequence of different TLR4 expression obtained in this study. The response to LPS depends on the dosage of LPS [55] and on the level of TLR4 expression [56]. In addition, TLR4 is the only TLR that signals through both MyD88 and TRIF signaling pathway and these signaling pathways are tightly regulated by a number of regulatory proteins [57]. Consequently, different signaling outcomes, on the treatment with the same concentration of LPS, could be expected between DF-MSCs and DP-MSCs.

The finding that DP-MSCs, unlike DF-MSCs, suppressed proliferation of PBMCs in the absence of TLR ligation correlates with the ability of DP-MSCs to secrete TGF-β. This suggests that TGF-β could be an important mediator of immunosuppression by dental MSCs. Further, the proliferation of PBMCs in the presence of unstimulated DP-MSCs was completely restored by using a neutralizing anti-TGF-β antibody, suggesting the dominant role of this cytokine in the immunosuppressive properties of DP-MSCs. The study of Di Nicola et al. (2002), which identified TGF-β1 and HGF as potential mediators of MSC immunosuppressive properties, further supports this observation [15]. The augmented suppressive potential after the treatment with poly(I:C) was followed by a significantly higher production of TGF-β by both dental MSCs subtypes. This effect was even more obvious after the treatment of DF-MSCs with LPS. Our results contradict those of Opitz et al. [30], who showed that TLR activation did not modulate TGF-β production by BM-MSCs. In contrast, DP-MSCs responded differently to LPS and lowered the TGF-β production compared with control, which correlated with lower suppression of these cells on PBMC proliferation. The observation that DP-MSCs and DF-MSCs were able to secrete comparable levels of TGF-β even after removal of TLR agonists suggests that the effects caused by TLR activation remained when these cells were placed in the culture with PBMCs. Since anti-TGF-β antibody could not completely restore the suppression of PBMCs proliferation triggered by activated dental MSCs, we presumed that, under such conditions, TGF-β is not the sole mediator responsible for the observed effect.

IDO-1-mediated tryptophan degradation is reported to be an important T-cell inhibitory mechanism of BM-MSCs, which is regulated via IFN-γ receptors and TLRs [14,30]. Additionally, IDO-1 is involved in synthesis of kynurenine, a cytotoxic mediator responsible for immunesuppression mediated by BM-MSCs, which can be blocked with 1-methyl-L-tryptophan [30]. Our results, showing that untreated DP-MSCs had a higher expression of IDO-1 compared with DF-MSCs, are in line with the previous observation. Further, LPS decreased its expression in both subsets of dental MSCs, which correlated with the abrogated suppression of PMBC proliferation mediated by LPS-pretreated DP-MSCs, but not by DF-MSCs treated similarly. Therefore, decrease in the IDO-1 expression, besides the reduced secretion of TGF-β, could additionally explain the abrogated effect of LPS on the immunosuppressive phenotype of DP-MSCs. In contrast, a small decrease in IDO-1 expression by LPS-treated DF-MSCs was probably compensated by the enormous increase in the production of TGF-β after LPS treatment. In addition, lower decrease in IDO-1 expression in DF-MSCs on LPS treatment, compared with DP-MSCs, could be due to lower TLR4 expression by these cells and, thus, lower responsiveness to LPS and/or activation of different signaling pathways. This could also mean that LPS could act predominantly to reduce immunosuppressive capacity of dental MSCs by reducing IDO-1 expression, thus enabling the surrounding lymphocytes more available tryptophan and increased survival rate. This hypothesis is supported by a recent study suggesting that TLR4-primed human MSCs exhibit proinflammatory behavior, whereas those primed with TLR3 agonist exhibit immunosuppressive behavior [31]. Our results contradict those of Opitz et al., who showed that the enhanced immunosuppressive activity of BM-MSCs on TLR3 and TLR4 activation correlated with induction of IDO-1 via IFN-β and protein kinase R signaling [30]. We showed that poly(I:C) did not induce IDO-1 and that LPS down-regulated IDO-1 in DF-MSCs and DP-MSCs. Therefore, these findings suggest that the regulation of IDO-1 expression, on TLR3 and TLR4 activation, in dental MSCs might be different from that in BM-MSCs. Romieu-Mourez et al. showed that TLR3 or TLR4 activation in BM-MSCs resulted in formation of an inflammatory site, attracting inflammatory cells due to increased IL-1β, IL-6, IL-8, and CCL-5 production [38]. This finding correlates with the effect of TLR3 activation on increased IL-6 production by dental MSCs showed in our study and may contradict the observed suppression of PBMCs proliferation in co-culture with poly(I:C)-pretreated dental MSCs. However, increased IL-6 production was followed by increased production of TGF-β by these cells. McGeachy et al. found that TGF-β and IL-6 treatment of myelin-reactive Th17-cells completely abrogated the production of proinflammatory cytokines by these cells and their pathogenic function in EAE model. Further, such treatment increased the production of IL-10, an anti-inflammatory cytokine, by these cells [58]. Therefore, although poly(I:C) did not increase IDO-1 expression by dental MSCs, it could be possible that in our study, IL-6 and TGF-β in combination contributed to the observed suppressive effect of poly(I:C) treated dental MSCs on PBMC proliferation. Since DF-MSCs and DP-MSCs pretreated with poly(I:C) exhibited an immunosuppressive behavior in vitro, it should be further explored whether such pretreatment of dental MSCs could reduce or even omit the granulomatous tissue formation that was observed when the untreated cells were implanted into imunocompetent mice.

In summary, we showed that DF-MSCs and DP-MSCs, isolated from the same tooth/donor, differed in morphology, proliferation, levels of MSC marker expression, and functions. The functional differences are related to their differentiation potential, immunomodulatory properties, and the response to TLR signaling. These findings may be relevant for their biomedical applications for which further explorations of their functions in vivo are necessary. In line with our results obtained in vitro, DP-MSCs, exhibiting a higher potential to form chondrocytes and osteoblasts, may be useful for the treatment of osteo-chondral diseases. On the other hand, the properties of DF-MSCs are desirable for the treatment of diseases caused by chronic inflammation accompanied by tissue injuries.

Footnotes

Acknowledgments

This work was financially supported by The Ministry of Science and Technological Development of The Republic of Serbia (project: 156017) and by The Military Medical Academy, Belgrade, Serbia (project: VMA/06/10/B.2). The authors thank Prof. M. Micev for providing expertise in field of Histochemistry; Dr. B. Bozic and M. Jovic, M.Sc., for helpful assistance; and L. Rubinjoni and M. Mitic for proofreading the manuscript.

Author Disclosure Statement

No competing financial interests exist.

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