Soluble factors released from platelets can modulate the immune response of leukocytes. We and others have recently found that T lymphocytes with bound platelets have reduced proliferation and IFN-γ and IL-17 production. Thus, we speculate that if we induce the binding of platelets to lymphocytes, we will be able to regulate the inflammatory response. When we cocultured platelets with lymphocytes at different ratios, we were able to increase the percentage of lymphocytes with bound platelets. The coculture of platelets with lymphocytes in the presence of stimulation decreased the production of IFN-γ and TNF-α, T cell proliferation, and the expression of CD25, PD-L1, and SLAM. However, this coculture increased CD39 expression. All of these effects were dependent on the dose of platelets and operated indistinctly with platelets from different healthy donors. When platelets were cocultured in the same compartment with lymphocytes, we observed less IFN-γ and TNF-α production and T lymphocyte proliferation than in cultures with platelets separated from lymphocytes by a 0.4-μm pore size filter. The binding of platelets to lymphocytes was blocked with anti–P-selectin Abs, and when this occurred we observed higher IFN-γ and TNF-α production than in nonblocked conditions. The cocultures of platelets with synovial fluid cells from rheumatoid arthritis patients reduced inflammatory cytokine production and increased IL-10 production. These results suggest that platelet binding to lymphocytes effectively regulates T lymphocyte function. This mechanism could be easily applied to reduce inflammatory responses.

Platelets (PLTs) have been recently recognized as immunoregulatory cells (13). Upon activation, PLTs release cytokines, chemokines, growth factors, and PLT-derived microparticles (PMPs) and express activation molecules in the membrane surface as P-selectin (CD62P) or CD40L (47). Soluble factors released by PLTs can modulate leukocyte function. TGF-β modulates IFN-γ and IL-17 production and CD4+ cell proliferation (8). Chemokine PLT factor 4 (PF4) limits the transplantation-induced Th17 differentiation of CD4+ T cells, reducing graft rejection (9, 10). PF4 also inhibits non–regulatory T cell (Treg) proliferation and enhances Treg differentiation and IL-10 production (11). Soluble CD40L (sCD40L) released by PLTs can enhance IL-10 production and decrease TNF-α secretion by monocytes (12). sCD40L can also reduce the inflammatory cytokines produced by CD4+ T cells (13, 14).

It has been shown that PLTs and PMPs can bind to lymphocytes (1518). There are several molecules that PLTs can use to bind to lymphocytes (1921). The interaction of CD62P on PLTs with P-selectin glycoprotein ligand 1 (PSGL-1) on lymphocytes appears to be essential for this binding (17). We and others have shown that CD4+ T lymphocytes with bound PLTs produce lower levels of IFN-γ and IL-17 and proliferate less than CD4+ lymphocytes without bound PLTs (15) (22). Similarly, PMPs released by PLTs decrease IFN-γ and IL-17 production by IL-17+ Tregs in a P-selectin–dependent manner rather than by soluble factors (18). Blocking PLT binding to CD4+ T cells can also have in vivo consequences because it exacerbates the progression of experimental autoimmune encephalitis (22).

PLTs can have also effects on mouse CD8+ T cells. Resting PLTs suppressed cytotoxicity of CD8+ T cells, and allogeneic PLTs prevented cell-mediated immune thrombocytopenia and reduced the ability to reject donor-matched skin grafts (2325). Alternatively, activated PLTs can present peptides in the context of MHC class I, activating Ag-specific CD8+ T cells (26).

PLT therapy with PLT-rich plasma (PRP) is a promising anti-inflammatory and tissue-repairing therapy for osteoarthritis (2731). In a pig model of rheumatoid arthritis (RA), the intra-articular injection of PRP reduced inflammation of the knee joint and attenuated the chondral and synovial changes (32). However, it seems that PLTs in RA could not be always beneficial. PMPs derived from collagen-activated PLTs can amplify inflammation in synovial RA through neutrophil recruitment (33). Therefore, the use of PLT in RA patients needs to be further explored.

In this study, we first analyzed the effects of different doses and different donors on the coculture of PLTs with lymphocytes. In particular, we determined the levels of cytokines, CD4+ and CD8+ T cell proliferation, and the expression of activation markers. Second, we clarified whether these effects were induced by PLT-soluble factors or by the binding of PLTs to lymphocytes. Cytokine production and CD4+ and CD8+ T cell proliferation were found in PLTs cocultured with lymphocytes in the same compartment, separated by a 0.4-μm pore size filter, or after blocking with neutralizing anti-CD62P Abs, which prevent PLT binding to lymphocytes. Finally, we explored whether PLT could be beneficial for RA patients. For this purpose, we cocultured PLTs with synovial fluid (SF) cells to induce PLT binding to lymphocytes and the subsequent immunoregulation.

Ten milliliters of whole blood from healthy donors (n = 22) was collected in BD Vacutainer tubes (BD Biosciences, Franklin Lakes, NJ) containing heparin as an anticoagulant. SF from RA patients (n = 10) was obtained by puncture in a sterile syringe. The diagnosis of RA was based on the diagnostic criteria of the American College of Rheumatology for RA (34). Disease activity was assessed using the disease activity score–C-reactive protein (35). Table I shows the demographics, clinical parameters, and laboratory values of the healthy donors and RA patients enrolled in this study. The study included anonymous consecutive healthy donors and RA patients from the Hospital de la Santa Creu i Sant Pau. The protocol conformed to the Declaration of Helsinki, and guidelines for good clinical practice were approved by the Research Ethics Committee of Sant Pau Researche Institute. All participants received information concerning the participation in the study, and written consent was obtained from all participants.

Table I.
Baseline characteristics of healthy donors and RA patients
Healthy DonorsRA Patients
Age, y (mean ± SEM) 49.52 ± 15.22 57 ± 5.36 
Sex, % (n) women 63 (14) 60 (6) 
Years of evolution (mean ± SEM)  8.9 ± 2.81 
DAS28-CRP (mean ± SEM)  4.4 ± 0.34 
ESR, mm/h (mean ± SEM)  35.5 ± 7.54 
CRP, mg/l (mean ± SEM)  33.19 ± 9.94 
ACPAs and/or RF+, % (n 60 (6) 
Healthy DonorsRA Patients
Age, y (mean ± SEM) 49.52 ± 15.22 57 ± 5.36 
Sex, % (n) women 63 (14) 60 (6) 
Years of evolution (mean ± SEM)  8.9 ± 2.81 
DAS28-CRP (mean ± SEM)  4.4 ± 0.34 
ESR, mm/h (mean ± SEM)  35.5 ± 7.54 
CRP, mg/l (mean ± SEM)  33.19 ± 9.94 
ACPAs and/or RF+, % (n 60 (6) 

ACPA, anti-citrullinated peptide Ab; CRP, C-reactive protein; DAS28-CRP, disease activity score 28–C-reactive protein; ESR, erythrocyte sedimentation rate; RF, rheumatoid factor.

SF was treated with hyaluronidase from bovine testes (300 U/ml; Sigma-Aldrich, St. Louis, MO) for 30 min at 37°C. SF cells were washed twice in PBS and filtered through a 50-μm sterile filter (ImmunoTools, Friesoythe, Germany). Then, SF liquids free of cells were collected and kept at −80°C until use.

Mononuclear cells from SF and from peripheral blood were isolated by gradient centrifugation (Lymphoprep; Axis-Shield, Oslo, Norway). PBMCs and SFMCs were washed twice with PBS and resuspended at 1 × 106 cells/ml in complete RPMI 1640 medium (supplemented with 10% FCS; Biological Industries, Kibbutz Beit Haemek, Israel), 2 mM glutamine, 100 U/ml penicillin, and 100 mg/ml streptomycin (BioWhittaker, Verviers, Belgium).

PLTs were isolated from 10 ml of heparin blood. Tubes with blood were centrifuged at 190 × g for 20 min at 22°C. The upper part was PRP, and it was collected and centrifuged at 190 × g for 10 min at 22°C. The new upper part of the centrifuged PRP was collected and centrifuged at 1000 × g for 10 min at 22°C. PLT pellets were washed with PBS containing 5 mM EDTA to avoid PLT activation at 1000 × g for 10 min at 22°C. The PLTs were then resuspended in complete RPMI 1640 medium at 56 × 106 PLTs/ml. Purity (measured as percentage CD41a+ events), activation (measured as percentage P-selectin+ of CD41a+ events), and the number of PLTs were assessed by flow cytometry after PLT isolation: >98% of the events were CD41a+ (specific PLT marker), and 7–15% of PLTs expressed P-selectin with low levels of mean fluorescence intensity (MFI). Activation ability of PLTs after isolation was assessed stimulating PLTs with 10 U/ml thrombin for 5 min (Supplemental Fig. 1).

PBMCs were stimulated with an anti-CD3, CD2, and CD28 T cell activation/expansion kit (Miltenyi Biotec, Bergisch Gladbach, Germany) for 24, 48, and 72 h in 5% CO2 at 37°C in the absence or presence of autologous and heterologous PLTs at different PLT/PBMC ratios (100:1, 50:1, 10:1, and 1:1). These coculturing ratios of PLTs/PBMCs were selected according to normal PLT counts (150 × 109/l to 400 × 109/l) and leukocyte counts (4 × 109/l to 11 × 109/l) in humans (36, 37). In some experiments, PLTs and PBMCs were cultured either in the same well compartment or separated by a 0.4-μm pore size filter for 72 h (Millipore, Billerica, MA). The 0.4-μm pore size filter blocked PLT–PBMC contact but allowed soluble factor diffusion. As a negative control in all experiments, PBMCs were cultured in the same conditions. To prevent PLTs from binding to lymphocytes, 20 μg/ml blocking anti–P-selectin Abs (R&D Systems, Minneapolis, MN) were added to the PLT suspension 15 min prior to the coculture with PBMCs. SFMCs were cultured in the presence or absence of heterologous PLTs from healthy donors at a 100:1 ratio in a medium or with a T cell activation/expansion kit for 72 h in 5% CO2 at 37°C. Cultured cells were collected and stained for flow cytometry analysis. Culture supernatants were collected and kept at −80°C until use.

PBMCs and SFMCs were washed and resuspended in PBS. The PBMCs were then incubated with anti-CD3–PE-Cy5 (UCHT1), CD25-PE (clone M-A251), CD80-PE (clone L307.4), CD86-PE (clone IT2.2), CTLA4-PE (clone BNI3), ICOS-PE (clone DX29), FOXP3-FITC (clone 150D), SLAM-PE (clone A12) (BD Biosciences), CD41a-FITC (clone HIP8), CD41a-PE (clone HIP8), CD54-PE (clone 1H4), granzyme B–PE (clone HC4) (ImmunoTools), CD4–PE-Cy7 (clone RPA-T4), CD39-PE (clone A1), HLA-DR–PE (clone L243), PD-L1–PE (clone 29E.2A3), CCR8-PE (clone L263G8), and perforin-PE (clone B-D48) (BioLegend) mAbs and the corresponding isotype controls. The SFMCs were incubated with anti-CD3–PE-Cy7 (clone SK7) (BD Biosciences), CD4-VioGreen (clone VIT4) (Miltenyi Biotec), CD14-allophycocyanin (18D11), and CD41a-FITC (clone HIP8) (ImmunoTools) mAbs. Samples were incubated for 20 min in the dark, washed, and resuspended with 400 μl of PBS prior to flow cytometry analysis.

IFN-γ, IL-10, TNF-α (BD Biosciences), IL-17, PF4, and sCD40L (PeproTech, London, U.K.) levels were determined using specific ELISA kits according to the manufacturers’ instructions. All cytokines were quantified using specific standard curves of recombinant cytokines provided by the corresponding ELISA kit. The limits of detection were as follows: 32 pg/ml for IFN, 10 pg/ml for IL-17 and IL-10, 20 pg/ml for TNF, 46.87 pg/ml for PF4, and 31.25 pg/ml for sCD40L.

All samples were acquired using the MACSQuant Analyzer 10 cytometer. CD41a+ PLTs/μl were quantified in the PLT suspension by flow cytometry. PLT activation was assessed using anti–CD62P-PE mAb (ImmunoTools). CD4+ T lymphocytes were identified and gated as CD3+CD4+ cells (CD4+). Owing to limitations in the number of parameters for flow cytometry analysis, anti-CD8 was not included in the mix. Instead, in an additional tube, we analyzed T cell subsets according to the markers anti-CD3, anti-CD4, and anti-CD8. Less than 0.5% of T cells were CD3+CD4CD8 (data not shown). We could therefore infer than >99.5% of CD3+CD4 cells were CD8+. To determine the percentage of CD4+ and CD8+ T cells with bound PLTs, CD41a+CD3+CD4+ and CD41a+CD3+CD4 (CD8+) T cells were analyzed. Surface expression of markers was analyzed on CD4+, CD8+, CD4+CD41a+ (CD4+PLT+), CD8+CD41a+ (CD8+PLT+), CD4+CD41a (CD4+PLT), and CD8+CD41a (CD8+PLT) cell subsets. To assess the proliferation of CD4+ and CD8+ T cells in the presence or absence of PLTs, we quantified CD4+ and CD8+ cells per microliter. In the SFMC cultures, macrophages were gated according to side scatter and CD14+ parameters. CD4+ and CD8+ T cells were gated according to side scatter, CD3+, and CD4+ cells or CD4 cells, respectively. The percentage of positive cells and MFI of each individual marker were obtained using FlowJo version 10 (FlowJo, Ashland, OR).

Statistical analyses were performed using paired t tests, t tests, one-way ANOVA, and a Spearman correlation on GraphPad Prism 5 software. Data are presented as mean ± SEM. A p value <0.05 was considered statistically significant.

We first studied whether the addition of PLTs, at the physiological blood rate (100 PLTs/1 leukocyte), influenced the stimulation of PBMCs with anti-CD3, CD2, and CD28 Abs. After 72 h of culture, we evaluated the cytokine production in supernatants and the proliferation and expression of activation markers of CD4+ and CD8+ T cells. We observed that the levels of Th1 cytokines, IFN-γ, and TNF-α in the cultures with PLTs were lower than in the cultures without PLTs (Fig. 1A, 1B). There were no significant differences in the IL-10 and IL-17 levels between the two cultures (IL-10, 414.32 ± 42.56 pg/ml without PLTs versus 457.45 ± 102.9 pg/ml with PLTs; IL-17, 753.2 ± 102.2 pg/ml without PLTs versus 613.5 ± 102.2 pg/ml with PLTs). Interestingly, the PBMCs from donors producing less IFN-γ in the presence of PLTs were the PBMCs that also produced less IL-17 (Supplemental Fig. 2A, 2B). Less CD4+ and CD8+ T cell proliferation was observed in the cultures with PLTs than in the cultures without PLTs (Fig. 1C, 1D). The expressions of CD25, PD-L1, and SLAM were lower on the CD4+ and CD8+ T cells cultured with PLTs than on those cultured without PLTs, whereas CD39 expression was higher on the CD4+ T cells cultured with PLTs but not on CD8+ T cells (Fig. 1E–I).

FIGURE 1.

Platelets inhibit T cell function. PBMCs from healthy donors were stimulated for 72 h with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of PLTs (ratio, 100 PLTs/1 PBMC). Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, CD25-PE, CD39-PE, SLAM-PE, and PD-L1–PE mAbs for flow cytometry analysis. (A) IFN-γ and (B) TNF-α were analyzed in culture supernatants. (C) The proliferation of CD4+ cells and (D) CD8+ cells was analyzed as CD4+ or CD8+ cells per microliter by flow cytometry, respectively. (E) A representative expression of markers analyzed on CD4+ T cells is shown. The dot plot shows the percentage of (F) CD25, (G) PD-L1, (H) SLAM, and (I) MFI of CD39 on CD4+ and CD8+ T lymphocytes. Data are presented as mean. Statistical analysis was performed using a paired t test. *p < 0.05, **p < 0.01, ***p < 0.0001.

FIGURE 1.

Platelets inhibit T cell function. PBMCs from healthy donors were stimulated for 72 h with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of PLTs (ratio, 100 PLTs/1 PBMC). Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, CD25-PE, CD39-PE, SLAM-PE, and PD-L1–PE mAbs for flow cytometry analysis. (A) IFN-γ and (B) TNF-α were analyzed in culture supernatants. (C) The proliferation of CD4+ cells and (D) CD8+ cells was analyzed as CD4+ or CD8+ cells per microliter by flow cytometry, respectively. (E) A representative expression of markers analyzed on CD4+ T cells is shown. The dot plot shows the percentage of (F) CD25, (G) PD-L1, (H) SLAM, and (I) MFI of CD39 on CD4+ and CD8+ T lymphocytes. Data are presented as mean. Statistical analysis was performed using a paired t test. *p < 0.05, **p < 0.01, ***p < 0.0001.

Close modal

We next stimulated PBMCs with anti-CD3, CD2, and CD28 for 72 h with different PLT/PBMC ratios (0:1, 1:1, 10:1, 50:1, and 100:1). Supernatant IFN-γ and TNF-α concentrations, CD4+ and CD8+ T cell proliferation, and CD4+CD25+ and CD8+CD25+ percentages decreased in a PLT dose-dependent manner (Fig. 2). Supernatant IL-10 and IL-17 concentrations were not significantly different in the cultures with different doses of PLTs (IL-10, 466.2 ± 113.6 pg/ml for 0:1, 351.1 ± 48.68 pg/ml for 1:1, 311.4 ± 84.39 pg/ml for 10:1, 266.3 ± 76.72 pg/ml for 50:1, and 259.1 ± 28.7 pg/ml for 100:1; IL-17, 780.4 ± 313.9 pg/ml for 0:1, 671.2 ± 321.8 pg/ml for 1:1, 589.5 ± 265.4 pg/ml for 10:1, 522.7 ± 276 pg/ml for 50:1, and 227.3 ± 78.28 pg/ml for 100:1). Similar results were observed in 24- and 48-h cultures (Supplemental Fig. 3). We next stimulated PBMCs with anti-CD3, CD2, and CD28 for 72 h in the presence of autologous and heterologous PLTs at the 100:1 ratio. The decrease in IFN-γ and TNF-α concentration, CD4+ and CD8+ T cells proliferation, and the changes in CD25, PD-L1, SLAM, and CD39 expression induced by autologous and heterologous PLTs were similar (Fig. 3). Interestingly, we observed that PLT effects on IL-17 production were PBMC donor-dependent but PLT donor-independent (Supplemental Fig. 2C).

FIGURE 2.

Inhibition of T cell functions depending on PLT doses in cocultures. PBMCs from healthy donors (n = 5) were stimulated with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of autologous PLTs at different PLT/PBMC ratios (1:1, 10:1, 50:1, and 100:1) for 72 h. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, and CD25-PE mAbs for flow cytometry analysis. (A) IFN-γ and (B) TNF-α in culture supernatants and (C) CD4+ or (D) CD8+ cells per microliter and (E) percentage of CD25 on CD4+ and (F) CD8+ T cells from cocultures at different PLT/PBMC ratios are shown. Multiple comparisons were analyzed by one-way ANOVA (IFN-γ and TNF-α, p < 0.0001; CD4+ cells per microliter, p = 0.0056; percentage CD4+CD25+, 0.049; and percentage CD8+CD25+, 0.038). Data are presented as mean ± SEM. Comparisons between the 0:1 ratio and each of the other ratios were analyzed by a paired t test. *p < 0.05, **p < 0.01.

FIGURE 2.

Inhibition of T cell functions depending on PLT doses in cocultures. PBMCs from healthy donors (n = 5) were stimulated with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of autologous PLTs at different PLT/PBMC ratios (1:1, 10:1, 50:1, and 100:1) for 72 h. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, and CD25-PE mAbs for flow cytometry analysis. (A) IFN-γ and (B) TNF-α in culture supernatants and (C) CD4+ or (D) CD8+ cells per microliter and (E) percentage of CD25 on CD4+ and (F) CD8+ T cells from cocultures at different PLT/PBMC ratios are shown. Multiple comparisons were analyzed by one-way ANOVA (IFN-γ and TNF-α, p < 0.0001; CD4+ cells per microliter, p = 0.0056; percentage CD4+CD25+, 0.049; and percentage CD8+CD25+, 0.038). Data are presented as mean ± SEM. Comparisons between the 0:1 ratio and each of the other ratios were analyzed by a paired t test. *p < 0.05, **p < 0.01.

Close modal
FIGURE 3.

Inhibition of T cell function by autologous or heterologous PLTs. PBMCs from healthy donors (n = 6) were stimulated with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of autologous (100:1 A) and heterologous (100:1 H) PLTs at a ratio of 100:1 for 72 h. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, CD25-PE, CD39-PE, CD41a-FITC, SLAM-PE, and PD-L1–PE mAbs for flow cytometry analysis. (A) IFN-γ and (B) TNF-α production, (C) CD4+ and CD8+ cells per microliter, and (D) CD4+ and CD8+ PLT+ cells are shown. Percentage of expression of (E) CD25, (F) PD-L1, (G) SLAM, and (H) MFI of CD39 on CD4+ and CD8+ T cells is shown. Data are presented as mean ± SEM. Statistical analysis was performed using a t test. *p < 0.05, **p < 0.01, ***p < 0.0001.

FIGURE 3.

Inhibition of T cell function by autologous or heterologous PLTs. PBMCs from healthy donors (n = 6) were stimulated with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of autologous (100:1 A) and heterologous (100:1 H) PLTs at a ratio of 100:1 for 72 h. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, CD25-PE, CD39-PE, CD41a-FITC, SLAM-PE, and PD-L1–PE mAbs for flow cytometry analysis. (A) IFN-γ and (B) TNF-α production, (C) CD4+ and CD8+ cells per microliter, and (D) CD4+ and CD8+ PLT+ cells are shown. Percentage of expression of (E) CD25, (F) PD-L1, (G) SLAM, and (H) MFI of CD39 on CD4+ and CD8+ T cells is shown. Data are presented as mean ± SEM. Statistical analysis was performed using a t test. *p < 0.05, **p < 0.01, ***p < 0.0001.

Close modal

The percentages of CD4+ and CD8+ T cells that bound PLTs were comparable in cultures with autologous and heterologous PLTs (Fig. 3D). However, the binding of PLTs to CD4+ and CD8+ T cells increased in the PBMC cultures with the highest ratios of added PLTs (Fig. 4A, 4B) and those with longer incubation times (Supplemental Fig. 3B). We measured two PLT-derived factors that regulate T cell function, PF4 and sCD40L. We observed that the levels of both factors increased with the ratios of added PLTs (PF4, 33,726 ± 8,855 pg/ml for 0:1, 52,681 ± 13,145 pg/ml for 1:1, 154,387 ± 41,253 pg/ml for 10:1, 325,026 ± 79,293 pg/ml for 50:1, and 641,392 ± 112,822 pg/ml for 100:1, p < 0.0001 by ANOVA; sCD40L, 160.8 ± 34.2 pg/ml for 0:1, 195.9 ± 40.46 pg/ml for 1:1, 274.2 ± 59.65 pg/ml for 10:1, 303.8 ± 44.29 for 50:1, and 369.5 ± 48.16 for 100:1, p < 0.05 by ANOVA). When we selected the highest ratio of PLTs/lymphocytes, IFN-γ production in cultures without PLTs correlated with the percentage of CD4+ and CD8+ T cells with bound PLTs in cultures with PLTs (r = 0.45, p = 0.039 and r = 0.51, p = 0.013, respectively) (Fig. 4C, 4D). We next compared the phenotypes of CD4+ and CD8+ T cells with and without bound PLTs without adding PLTs. Activated T cell markers, CD80, CD86, CTLA-4, CD39, ICOS, HLA-DR, CCR8, granzyme B, perforin, FOXP3, CD54, CD25, SLAM, and PD-L1, were upregulated on CD4+ and CD8+ PLT+ compared with CD4+ and CD8+ PLT cells (Supplemental Fig. 4). However, CD28, CD62L, CD69, CD123, GITR, CD44, and CD95 expression did not differ on CD4+PLT+ and CD4+PLT cells (data not shown). After coculturing PLTs and lymphocytes for 72 h, the CD4+ T cells that bound PLTs also showed higher levels of CD25, SLAM, PD-L1, and CD39 compared with the CD4+PLT cells (Fig. 4E).

FIGURE 4.

Binding of PLTs to CD4+ and CD8+ T cells in cocultures with PLTs and phenotype of CD4+PLT+. PBMCs from healthy donors (n = 5) were stimulated with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of autologous PLTs (ratios 1:0, 10:1, 50:1, and 100:1) for 72 h. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, CD25-PE, CD39-PE, CD41a-FITC, and PD-L1–PE for flow cytometry analysis. (A) Percentage of CD4+ and (B) CD8+PLT+ cells in cocultures of PLTs/PBMCs at different ratios is shown. Multiple comparisons were analyzed by one-way ANOVA (CD4+PLT+ and CD8+PLT+, p < 0.0001). (C) A correlation between IFN-γ levels in 0:1 cultures (without PLTs) and the percentage of CD4+ and (D) CD8+ PLT+ in cocultures with PLTs at the 100:1 ratio are shown. (E) Representative expression of CD25, SLAM, PD-L1, and CD39 on CD4+PLT+ and CD4+PLT from 0:1 and 100:1 cocultures are shown. Data are presented as mean ± SEM. Comparisons between 0:1 ratio and each of the other ratios were analyzed by paired t test. For analysis of correlation we used a Spearman correlation. *p < 0.05, **p < 0.01, ***p < 0.0001.

FIGURE 4.

Binding of PLTs to CD4+ and CD8+ T cells in cocultures with PLTs and phenotype of CD4+PLT+. PBMCs from healthy donors (n = 5) were stimulated with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of autologous PLTs (ratios 1:0, 10:1, 50:1, and 100:1) for 72 h. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, CD25-PE, CD39-PE, CD41a-FITC, and PD-L1–PE for flow cytometry analysis. (A) Percentage of CD4+ and (B) CD8+PLT+ cells in cocultures of PLTs/PBMCs at different ratios is shown. Multiple comparisons were analyzed by one-way ANOVA (CD4+PLT+ and CD8+PLT+, p < 0.0001). (C) A correlation between IFN-γ levels in 0:1 cultures (without PLTs) and the percentage of CD4+ and (D) CD8+ PLT+ in cocultures with PLTs at the 100:1 ratio are shown. (E) Representative expression of CD25, SLAM, PD-L1, and CD39 on CD4+PLT+ and CD4+PLT from 0:1 and 100:1 cocultures are shown. Data are presented as mean ± SEM. Comparisons between 0:1 ratio and each of the other ratios were analyzed by paired t test. For analysis of correlation we used a Spearman correlation. *p < 0.05, **p < 0.01, ***p < 0.0001.

Close modal

We next compared the cytokine production and CD4+ and CD8+ T cell proliferation when we cocultured T cells and PLTs in the same compartment (PLTw), when PLTs and T cells were separated by a 0.4-μm pore size filter (PLTtw), and when we cultured T cells without PLTs (PLTwo). As expected, we only observed an increase in the percentage of CD4+PLT+ and CD8+PLT+ cells in the PLTw cultures (Fig. 5A). The lowest IFN-γ concentration was observed in the PLTw cultures (Fig. 5B). The TNF-α concentration was significantly lower in the PLTw but not in the PLTtw cultures when compared with that in the PLTwo cultures (Fig. 5C). IL-10 and IL-17 concentrations were similar in the PLTw and PLTtw cultures (IL-10, 266.2 ± 53.45 pg/ml for PLTwo, 280.2 ± 122.3 pg/ml for PLTw, and 315.2 ± 162 pg/ml for PLTtw; IL-17, 451.1 ± 117.8 pg/ml for PLTwo, 485.5 ± 120.5 pg/ml for PLTw, and 487 ± 147.6 pg/ml for PLTtw). CD4+ and CD8+ T cell proliferation only decreased significantly when PLTs were cultured in the same compartment (Fig. 5D).

FIGURE 5.

Effect of soluble factor release by PLTs and binding of PLTs to CD4+ and CD8+ T cells on T cell function. PBMCs from healthy donors were stimulated for 72 h with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of PLTs at a PLT/PBMC ratio of 100:1 in the same compartment (100:1 PLTw) or in 0.4-μm pore size filter–separated compartments (100:1 PLTtw), which enable soluble factor diffusion but prevent PLT–cell contact. Moreover, the binding of PLT to T cells was blocked using anti-CD62P blocking Ab. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, and CD41a-FITC for flow cytometry analysis. (A) Percentage of CD4+ and CD8+ PLT+ (B) levels of IFN-γ, (C) TNF-α, and (D) CD4+ and CD8+ cells per microliter in cultures of 0:1, 100:1 PLTs/PBMCs in the same well (PLTw), and 100:1 PLT/PBMCs separated by transwell (PLTtw) is shown. (E) Percentage of CD4+ and CD8+ PLT+ (F) levels of IFN-γ, (G) TNF-α, and (H) CD4+ and CD8+ cells per microliter in cultures with a PLT/PBMC ratio of 0:1 or with anti-CD62P blocking Ab is shown. Data are presented as mean. Statistical analysis was performed using a paired t test. *p < 0.05, **p < 0.01, ***p < 0.0001.

FIGURE 5.

Effect of soluble factor release by PLTs and binding of PLTs to CD4+ and CD8+ T cells on T cell function. PBMCs from healthy donors were stimulated for 72 h with anti-CD3, CD28, and CD2 in the absence (0:1) or presence of PLTs at a PLT/PBMC ratio of 100:1 in the same compartment (100:1 PLTw) or in 0.4-μm pore size filter–separated compartments (100:1 PLTtw), which enable soluble factor diffusion but prevent PLT–cell contact. Moreover, the binding of PLT to T cells was blocked using anti-CD62P blocking Ab. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy5, CD4–PE-Cy7, and CD41a-FITC for flow cytometry analysis. (A) Percentage of CD4+ and CD8+ PLT+ (B) levels of IFN-γ, (C) TNF-α, and (D) CD4+ and CD8+ cells per microliter in cultures of 0:1, 100:1 PLTs/PBMCs in the same well (PLTw), and 100:1 PLT/PBMCs separated by transwell (PLTtw) is shown. (E) Percentage of CD4+ and CD8+ PLT+ (F) levels of IFN-γ, (G) TNF-α, and (H) CD4+ and CD8+ cells per microliter in cultures with a PLT/PBMC ratio of 0:1 or with anti-CD62P blocking Ab is shown. Data are presented as mean. Statistical analysis was performed using a paired t test. *p < 0.05, **p < 0.01, ***p < 0.0001.

Close modal

To confirm the importance of PLT binding to T lymphocytes for regulating the immune response, we next blocked this binding with an anti–P-selectin Ab. The blocking decreased the percentage of CD4+ and CD8+ PLT+ cells (Fig. 5E) and consequently increased IFN-γ and TNF-α levels (Fig. 5F, 5G). However, it did not alter cellular proliferation (Fig. 5H) or IL-10 levels (635.4 ± 85.45 pg/ml for 0:1 versus 624.1 ± 109.9 pg/ml for anti–P-selectin cultures). IL-17 levels also tended to increase after the binding of PLTs to T lymphocytes was blocked (1231 ± 360.2 pg/ml for PLTwo versus 1644.1 ± 556.5 pg/ml for anti–P-selectin cultures, p = 0.12).

We next analyzed the effect of adding heterologous PLTs from healthy donors to mononuclear cells from SF (SFMCs) of patients with rheumatoid arthritis (Table I). We did not observe bound PLTs to CD14+ macrophages and T lymphocytes after isolation of SFMCs. After culturing SFMCs with PLTs at the ratio of 100:1, we detected that PLTs were bound to CD14+ macrophages, CD4+, and CD8+ T lymphocytes from SF (84.51 ± 2.75% of CD14+PLT+ macrophages, 36.76 ± 4.47% of CD4+PLT+, and 38.08 ± 6.13% of CD8+PLT+). When PLTs were added to SFMCs, spontaneous IFN-γ, IL-17, and TNF-α production decreased (Fig. 6A–C) and IL-10 production increased (Fig. 6D).

FIGURE 6.

Effect of PLTs in SF cell function. SFMCs and SF were isolated as described in 2Materials and Methods. SFMCs were cultured in the absence (0:1) or presence of healthy donor PLTs (100:1) in medium or with anti-CD3, CD2, and CD28 (beads) for 72 h. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy7, CD4-VioGreen, CD14-allophycocyanin, and CD41a-FITC for flow cytometry analysis. (A) IFN-γ, (B) TNF-α, (C) IL-17, and (D) IL-10 levels in SF and cultures with medium at 0:1 and 100:1 are shown. (E) IFN-γ, (F) TNF-α, (G) IL-17, (H) IL-10, and (I) CD4+ and CD8+ cells per microliter levels in cultures of beads at 0:1 and 100:1 are shown. Data are presented as mean. Statistical analysis was performed using a paired t test. *p < 0.05, **p < 0.01.

FIGURE 6.

Effect of PLTs in SF cell function. SFMCs and SF were isolated as described in 2Materials and Methods. SFMCs were cultured in the absence (0:1) or presence of healthy donor PLTs (100:1) in medium or with anti-CD3, CD2, and CD28 (beads) for 72 h. Culture supernatants were collected for ELISA analysis, and cells were stained with anti-CD3–PE-Cy7, CD4-VioGreen, CD14-allophycocyanin, and CD41a-FITC for flow cytometry analysis. (A) IFN-γ, (B) TNF-α, (C) IL-17, and (D) IL-10 levels in SF and cultures with medium at 0:1 and 100:1 are shown. (E) IFN-γ, (F) TNF-α, (G) IL-17, (H) IL-10, and (I) CD4+ and CD8+ cells per microliter levels in cultures of beads at 0:1 and 100:1 are shown. Data are presented as mean. Statistical analysis was performed using a paired t test. *p < 0.05, **p < 0.01.

Close modal

When SFMCs were stimulated with anti-CD3, CD2, and CD28, the addition of PLTs increased the PLT binding to SFMCs (78.55 ± 3.11% of CD14+PLT+ macrophages, 43.59 ± 5.81% of CD4+PLT+, and 43.06 ± 6.05% of CD8+PLT+) and reduced the CD4+ and CD8+ synovial T cell proliferation, decreased the induced IFN-γ, IL-17, and TNF-α production, but increased IL-10 production (Fig. 6E–I).

Our results suggest that PLTs regulate T cell function, decreasing the levels of inflammatory cytokines and inhibiting cellular proliferation. These changes take place in a PLT dose-dependent manner, indistinctly of the PLT donor. Although soluble factors released by PLTs can induce these changes, the binding of PLTs to CD4+ and CD8+ T lymphocytes seems to substantially contribute to the regulation of T cell function.

We have shown that coculturing PLTs with PBMCs significantly decreased TNF-α and IFN-γ but not IL-17 production. Other authors have shown that the presence of PLTs or PMPs decreases the production of IFN-γ and IL-17 through TGF-β and PF4 release (810, 18). The difference between our findings and these previous reports in regard to IL-17 production could be due to the different cellular content of the cultures, because our experiments were conducted with PBMCs instead of isolated CD4+ T cells. The IL-17 production in our experiments was heterogeneous, observing samples where IL-17 production increased or decreased with the addition of PLTs. Moreover, following our strategy, we were able to observe that the effects of PLTs on IL-17 were PBMC donor-dependent but PLT donor-independent. Different cellular content, such as Th1 IL-17+ or CD8+IL-17+ cells, in PBMCs from healthy donors could explain these two observations (38, 39). Another interesting observation was that the PBMC donors with a higher decrease of IFN-γ with PLT in coculture were the ones with a PLT-induced IL-17 decrease.

However, PLTs do not always have a suppressor effect. In some diseases, such as immune thrombocytopenia, PLTs enhance IFN-γ and IL-2 production by CD4+ T cells through the presentation of specific autoantigens (40).

We observed that decreases of IFN-γ, TNF-α levels, and CD4+ and CD8+ proliferation in cocultures with PLTs were PLT-dose dependent. In parallel, we observed the increase of two soluble factors, sCD40L and PF4, released by PLTs (11, 13, 14). Then, the increase of these two factors could be partially responsible for the inhibition of T cell function. In the cocultures in which transwells were used to physically separate PLTs and lymphocytes, we observed how soluble factors released by PLTs immunoregulate T cell function. In the cocultures with PLTs and lymphocytes in the same compartment, we observed the strongest reduction of IFN-γ and CD4+ and CD8+ T cell proliferation. In the transwells, only the inhibitory soluble factors were not able to reduce the TNF-α production. In the same compartment with lymphocytes, two regulatory mechanisms were operating: soluble factors released by PLTs (such as sCD40L and PF4) as well as the binding of PLTs to CD4+ and CD8+ cells through P-selectin (817). This finding is concordant with the observation that CD4+ T lymphocytes with bound PLTs produce fewer inflammatory cytokines and proliferate less than do those without bound PLTs (15, 22). A similar conclusion was reached by Dinkla et al. (18) who showed that PMPs bind to Tregs and inhibit IFN-γ and IL-17 production. Blocking PLT–lymphocyte binding in vivo can also exacerbate the progression of experimental autoimmune encephalitis (22). Moreover, we observed that binding of PLTs to T lymphocytes is PLT dose-dependent. We have also shown that autologous and heterologous PLTs similarly bind to T lymphocytes. This could explain the comparable reduction IFN-γ, TNF-α, and CD4+ and CD8+ proliferation by PLTs from both origins.

P-selectin plays a major role in the binding of PLT to CD4+ and CD8+ lymphocytes. The P-selectin on activated PLTs can bind to the PSGL-1 expressed on leukocytes (17, 41). There are other ligands that can be involved in the binding of PLT to leukocytes: GPIb-CD11b (20), CD40-CD40L (17), and GPIIb/IIIa-CD11/CD18 (21). However, the binding of PLTs to T lymphocytes was essentially abolished by the P-selectin blockade, whereas the inhibition of GPIIb/IIIa, CD11b, or CD40 ligands only led to a reduction of this binding (17). We observed that PLT binding to CD4+ and CD8+ T cells decreased after blocking their interaction with P-selectin Abs. Consequently, IFN-γ and TNF-α production increased. This is in line with the suggested immunoregulatory role of PLT binding to lymphocytes.

It is intriguing that only CD4+ T lymphocytes with a Th1 cytokine profile were able to bind PLTs through PSGL-1 because activated Th1 and Th2 CD4+ T lymphocytes express similar levels of PSGL-1 (42, 43). However, it has been shown that selectively Th1-overexpressed α3-fucosyltransferases have fucosylated PSGL-1, and this modification is associated with the ability to bind to P-selectin (44). Indeed, the fucosylation of O-glycans is necessary for the binding of PSGL-1 and P-selectin. After the interaction with P-selectin, PSGL1 signaling seems to affect T cell phenotype and function (45) because the deletion of PSGL-1 increases T cell proliferation and exacerbates inflammation in vivo (46).

We showed that CD4+ and CD8+ T lymphocytes with bound PLTs and CD4+ and CD8+ T lymphocytes without PLTs expressed different surface markers. One explanation for this finding is that PLTs selected cells with which to bind to regulate their function. In this line, Dinkla et al. (18) observed that PMPs bound selectively to CCR6+HLA-DR+ Tregs. Another possibility is that the binding of PLTs modified the expression of surface markers on lymphocytes. This is unlikely because when we added PLTs in coculture to induce the binding to leukocytes, the percentage of different subsets did not change. The original CD4+PLT+ cells (before we added PLTs) and the CD4+ cells that were recently bound to PLTs in our cocultures showed similarly increased levels of CD25, SLAM, PD-L1, and CD39. As Dinkla et al. (18) suggested with PMP on Tregs, PLTs could bind to specific activated CD4+ T cells to downregulate their proinflammatory functions. A similar mechanism could be operating on CD8+ T cells. Accordingly, CD8+PLT+ and CD8+PLT T cells showed a different phenotype. In this case, we can speculate that PLTs directly regulate CD8+ T cells through the binding and indirectly through the regulation of CD4+ T cells.

By adding PLTs, we were able to decrease the amount of inflammatory cytokines and increase the amount of anti-inflammatory cytokines produced by SFMCs from RA patients in vitro. RA is a chronic autoimmune disease characterized by polyarticular synovitis (47). In the RA joint, synovium is infiltrated by macrophages and lymphocytes, and these infiltrating cells produce inflammatory cytokines such as TNF-α, IL-1, IL-6, IL-12, IL-15, IL-18, IFN-γ, IL-17, and GM-CSF, contributing to joint destruction (48, 49). The objective of most therapies in RA is to decrease these infiltrates and their cytokines (50). PLT therapy has not been applied to RA patients, but it is currently used in osteoarthritis (2731). In these patients, PLT therapy decreases the destruction of joints and stimulates cartilage regeneration. PLT therapy has only been examined in a pig model of RA. Intra-articular injection of PRP in these pigs reduced inflammation of the knee joint and attenuated the chondral and synovial changes observed during disease progression (32). According to our findings, PRP could also be beneficial for RA patients. We observed not only a decrease in IFN-γ, TNF-α, and IL-17 production but also an increase in IL-10 production by SFMCs in RA patients. The effect of bound PLTs and their soluble factors on cells of SFMCs could be similar to the effect on PBMCs. The preactivated state of SFMCs in RA patients can explain the IL-10 increase induced by the PLT binding to lymphocytes from RA SF as well as the same binding to lymphocytes from healthy donor blood (48, 51). PLTs and their soluble factors could increase Treg function (8, 11, 13, 14) that are present in SFMCs (52). This fact could explain the IL-10 increase induced by PLTs in RA SFMCs. We are currently comparing the effect of PLTs on target cells when they are in a preactivated state and during the differentiation to a Th1, Th2, or Th17 profile. This could have a significant impact on the development of PLTs as an anti-inflammatory therapy for several chronic inflammatory diseases.

This work was supported by Fondos Feder-Instituto de Salud Carlos III (PI14/741). S.V. was supported by Fondo Investigaciones Sanitarias and was a participant in the Program for Stabilization of Investigators of the Direcció i d’Estrategia i Coordinació del Departament Salut de la Generalitat de Catalunya.

The online version of this article contains supplemental material.

Abbreviations used in this article:

MFI

mean fluorescence intensity

PF4

PLT factor 4

PLT

platelet

PLTtw

PLTs and T cells separated by a 0.4-μm pore size filter

PLTw

cocultured T cells and PLTs in the same compartment

PLTwo

T cells cultured without PLTs

PMP

platelet-derived microparticle

PRP

PLT-rich plasma

PSGL-1

P-selectin glycoprotein ligand 1

RA

rheumatoid arthritis

sCD40L

soluble CD40L

SF

synovial fluid

SFMC

SF mononuclear cell

Treg

regulatory T cell.

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The authors have no financial conflicts of interest.

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