functional consequences of platelet binding to t lymphocytes in inflammation
TRANSCRIPT
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TITLE: Functional consequences of platelet binding to T lymphocytes in 1
inflammation 2
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Carlos Zamora*, Elisabet Cantó*, Juan C. Nieto*, M. Angels Ortiz*, Cesar 4
Diaz-Torné†, Cesar Diaz-Lopez†, Josep M. Llobet†, Candido Juarez‡, Sílvia 5
Vidal* 6
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* Dep. Immunology of Institut Recerca Hospital S. Pau, IIB-Sant Pau, 8
Barcelona, Spain 9
† Unit of Rheumatology of Hospital S. Pau, Barcelona, Spain 10
‡ Dep. Immunology of Hospital S. Pau, Barcelona, Spain 11
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Summary sentence: Platelet binding is a new regulatory mechanism of T 13
cell function. Higher frequency of platelet binding to lymphocytes is 14
associated with low severe inflammatory profile 15
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Running title: Platelet binding to T lymphocytes 17
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Corresponding author: Sílvia Vidal 19
Avda. Antoni M.Claret, 167; Barcelona-08025, Spain; 20
Telf. +(34) 935537544 21
Fax +(34) 935537598 22
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Key words: Human, T cell activation, Rheumatoid arthritis 25
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Total character count: 18000 27
Total number of figures: 5 28
Total number of references: 43 29
Total number of words in abstract: 191 30
Total number of words in summary sentence: 25 31
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Abbreviations 35
Anti-CCP Anti-cyclic citrullinated peptide antibodies 36
CFSE 5(6)-carboxyfluorescein diacetate succinimidylester 37
CRP C-reactive protein 38
DAS28-CRP Disease activity score-C reactive protein 39
ESR Erythrocyte sedimentation rate 40
iGMFI Integrated geometric fluorescence mean 41
Plt Platelet 42
RA Rheumatoid arthritis 43
RF Rheumatoid factor44
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Abstract 45
Expression of the scavenger receptor CD36 on lymphocytes is intriguing. We 46
observed that a minor subpopulation of lymphocytes expressed CD36 on cell 47
surface. We investigated the source of CD36 and also the proliferation and 48
cytokine production of these CD36+ CD4+ lymphocytes. Flow cytometry 49
analysis and immunofluorescence microscopy showed that CD36+ platelets 50
were responsible for CD36 detection on lymphocytes. CD36 was then used as a 51
tool to characterize lymphocytes with bound platelets. Activation-induced 52
proliferation was lower in CD4+ lymphocytes with bound platelets than 53
lymphocytes without bound platelets. IL-17 and IFNγ production was also 54
reduced in lymphocytes with bound platelets. We then studied the presence of 55
CD36+ CD4+ lymphocytes in rheumatoid arthritis (RA) patients. We observed 56
that percentage of CD4+ lymphocytes with bound platelets was higher on RA 57
patients than in healthy donors. RA patients with higher titers of anti-cyclic 58
citrullinated peptide antibodies (anti-CCP), rheumatoid factor levels (RF) and 59
cardiovascular risk index presented a lower percentage of CD4+ lymphocytes 60
with bound platelets. These patients also had higher IL-17 and IFNγ production. 61
These results suggest that platelet binding modifies lymphocyte function. This 62
binding could be a regulatory mechanism in RA that confers a less severe 63
phenotype. 64
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Introduction 72
CD36 is a transmembrane glycoprotein member of the class B scavenger 73
receptor family. This receptor has been found on macrophages/monocytes, 74
microglia, microvascular endothelium, cardiac and skeletal muscle, adipocytes 75
and platelets [1]. In these cells, CD36 recognizes a wide variety of ligands that 76
support its scavenger role. We have recently found a minor subpopulation of 77
lymphocytes that express CD36 (unpublished results). The presence of such a 78
scavenger receptor on a lymphocyte is intriguing. Vallejo et al. were not able to 79
detect CD36 in T lymphocytes by western blot [2]. However, recent reports 80
showed that B and γδ T cells can express CD36 [3-4]. Other authors described 81
that after mitogenic stimulation, CD36 was upregulated on peripheral blood 82
lymphocytes [5]. 83
The subpopulation of positive CD36 lymphocytes could have a characteristic 84
function. In the recent literature, the description and analysis of subpopulations 85
provide useful markers to predict disease development or activity. Th17 86
lymphocytes, for instance, are involved in the pathogenesis of rheumatoid 87
arthritis (RA), partially through IL-17 production [6-7]. IL-17 is highly expressed 88
in RA synovium [8] where it contributes to angiogenesis [9]. IL-17 has also 89
become a predictor of microvascular function and arterial compliance in RA 90
[10]. CD161 has been described as a marker of a T lymphocyte subpopulation 91
producing IL-17 [11]. The analysis of CD4+ CD161+ by flow cytometry on whole 92
blood has been associated to RA disease activity [12]. Other T lymphocyte 93
subpopulations, like Th1, produce IFNγ and TNFα. These two inflammatory 94
mediators are also associated with RA, supporting the microenvironment in RA 95
joint [13]. Another type of relevant cells in inflammatory diseases is Treg that 96
produced IL-10 and inhibits T cell expansion in vivo [14] and regulates Th1 and 97
Th2 response [15]. Tregs can be identified by flow cytometry based on their 98
CD4+ CD25 (high) CD127 (low/-) phenotype. The percentage of cells with Treg 99
phenotype has been associated with RA disease activity [16]. 100
To clarify the controversy about CD36 expression on T lymphocytes and the 101
function of CD36+ cell subset: Firstly, we analyzed whether CD36 expression 102
was extrinsic or intrinsic to lymphocytes. Secondly, we studied the functional 103
capability of this subpopulation selecting T lymphocytes with this marker. 104
Finally, we analyzed their role in autoimmune pathologies such as RA. 105
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Materials and methods 106
107
Samples and isolation of peripheral blood mononuclear cells 108
Whole blood from healthy donors (n=16) and RA patients (n=20) was collected 109
in BD Vacutainer tubes (BD Pharmingen, Franklin Lakes, NJ), containing 110
heparin as anticoagulant. Diagnosis was based on the American College of 111
Rheumatology diagnostic criteria for rheumatoid arthritis [17]. Disease activity 112
was assessed using the Disease activity score - C reactive protein (DAS28-113
CRP) [18]. The mean DAS28-CRP was 2.81 (range 1.13-5.08) and the mean 114
disease duration was 12.6 years (range, 2-40). Sixteen RA patients were 115
receiving methotrexate (7.5-22.5 mg), 6 of them were receiving prednisone too 116
(1.25-5 mg), and 4 RA patients were receiving leflunomide (20 mg). We 117
obtained clinical parameters of RA patients, C reactive protein (CRP) and 118
erythrocyte sedimentation rate (ESR). The cardiovascular risk index was 119
calculated according to age at the time of the study, gender, smoking history, 120
systolic arterial blood pressure, and total serum cholesterol level [19]. Written 121
consent was obtained and ethical approval for the study was granted by the 122
Institutional Ethics Committee. 123
Human peripheral blood mononuclear cells (PBMC) were isolated from 124
peripheral blood by gradient centrifugation on Ficoll-Hypaque (Lymphoprep, 125
AXIS-SHIELD PoCAs, Oslo, Norway). After centrifugation, cells were collected, 126
washed twice in PBS and resuspended in complete medium (RPMI 127
supplemented with 10% fetal calf serum, Biological Industries, Kibbutz Beit 128
Haemek, Israel, 2 mM glutamine, 100 U/ml penicillin and 100 μg/ml 129
streptomycin ,Biowhittaker, Verviers, Belgium). 130
131
Peripheral blood mononuclear cells and whole blood staining 132
PBMC (3 x 105 cells) were washed and resuspended in 100 μl of staining buffer 133
(PBS supplemented with 0.5 % BSA, Calbiochem Merk KGaA, Darmstadt, 134
Germany). PBMC were incubated with anti CD4-PECy7, CCR5-PE ,CCR6-PE, 135
CCR7-FITC, HLA-DR-PE, (BioLegend, San Diego, CA), CD8-PEDy647, 136
CD14-PEDy647, CD36-FITC (Clone TR9), CD41a-FITC, CD45RA-PEDy647 137
(Immunotools, Friesoythe, Germany), CD3-PECy7, CD25-PE, CD36-PE (clone 138
CDB38), CD45RA-FITC, CD45RO-FITC, CCR4-PE, CXCR3-PE (BD 139
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Pharmingen, Frankin Lakes, NJ) monoclonal antibodies and the corresponding 140
isotype controls for 20 minutes in the dark. Samples were then washed with 141
staining buffer and resuspended in PBS. 142
Whole blood cells (100 μl) were incubated for 20 minutes at room temperature 143
in the dark with anti CD4-PECy7, CD8-PEDy647 and CD36-FITC monoclonal 144
antibodies. Red blood cells were lysed and white cell fixed using TQ-prep 145
System (Coulter Corp., Miami, Fla) to be analyzed by flow cytometry. 146
147
Immunofluorescence microscopy analysis 148
To assess CD36 expression on monocyte and lymphocyte surface, PBMC (2 x 149
105 cells) were stained with anti CD3-FITC (Immunotools) or anti CD14-FITC 150
(Immunotools) with anti CD36-PE (BD Pharmingen), CD41a-PE (Immunotools) 151
or corresponding isotype control. A drop of cells in PBS was then left to dry on a 152
glass slide at room temperature. Expression of these markers was analyzed 153
using NIKON ECLIPSE 80i microscopy. 154
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Proliferation assay of lymphocyte 156
To determine CD4+ lymphocyte proliferation, PBMC were labeled with 5(6)-157
carboxyfluorescein diacetate succinimidylester (CFSE) (Sigma, S. Louis, 158
Missouri). Adjusted 107 cells/ml were stained with 7 μM of CFSE and incubated 159
for 15 minutes at 37ºC. After incubation, cells were washed with pre-warmed 160
medium and incubated for 30 minutes at 37ºC to ensure complete modification 161
of the probe. Cells were then washed with pre-warmed medium, resuspended 162
with complete medium at 5 x 106 cells/ml and incubated with anti-CD3, CD28 163
and CD2 T cell activation/expansion kit for 72 hours and 7 days according to the 164
manufacturer’s instructions (Milteny Biotec, CA, USA). After culturing, PBMC (3 165
x 105 cells) were collected, and stained with anti CD4-PECy7 and anti CD36-PE 166
monoclonal antibody (BD Pharmingen). Cells were then washed with staining 167
buffer and resuspended in PBS. CD4+ CD36- and CD4+ CD36+ history cycles 168
of proliferation were analyzed by flow cytometry. 169
170
Determination of intracellular cytokine production 171
PBMC were cultured with anti-CD3, CD28 and CD2 T cell activation/expansion 172
kit for 48 hours according to the manufacturer’s instructions. Cytokine 173
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production on healthy controls was assessed on CD3+ CD36- and CD3+ 174
CD36+ cells by intracellular staining of IL-17-PE (eBioscience, San Diego CA, 175
USA), IFNγ-PE (BD Pharmingen) IL-10-PE (eBioscience) and TNFα-PE (BD 176
Pharmingen) using a Cytofix/Cytoperm and Perm/Wash kit from BD 177
biosciences. Five hours before analysis, PMBC were cultured with PMA (50 178
ng/ml)/ Ionomycin (1 μg/ml) in the presence of BD GolgiStop TM Protein 179
Transport Inhibitor. Cells were collected and stained. Intracellular cytokine 180
production was analyzed on gated subpopulations by flow cytometry. 181
182
Determination of cytokine plasma levels 183
Plasma samples from healthy donors and RA patients were recollected and 184
kept at -80ºC until use. IL-17 (Peprotech, London, UK), TNFα, IFNγ, and IL-10 185
(BD Pharmingen) levels were tested using a specific ELISA kit according to the 186
manufacturer’s instructions. All cytokines were quantified with standard curves 187
provided by the corresponding ELISA kit. The limits of detection were: 10 pg/ml 188
for IL-17 and IL-10, 20 pg/ml for TNFα, and 32 pg/ml for IFNγ. 189
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Flow cytometry analysis 191
Surface expression of CD36 and CD14 was analyzed on monocytes and 192
lymphocytes (gated according to morphology by forward and side scatter 193
parameters). Other markers were analyzed on CD3+, CD4+ and CD8+ positive 194
cells. Naïve and memory T cells were selected according to CD3+ CD45RA+ 195
and CD3+ CD45RA- respectively. These analyses were performed on Beckman 196
Coulter F500 flow cytometer. We calculated the percentages of positive cells (% 197
cells) and geometric mean fluorescence intensity (GMFI) of each individual 198
marker using EXPO™ 32 MiltiCOMP Software (Beckman coulter). The 199
integrated geometric mean fluorescence intensity (iGMFI) determines the 200
amount of cytokines produced in each subpopulation and was calculated by 201
multiplying the percentage of positive cytokine-producing cells by the GMFI [20]. 202
Stimulation index in CFSE culture was measured calculating the ratio of 203
proliferating vs non-proliferating cells [21]. 204
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Quantification of anti-CCP and Rheumatoid factor levels 208
Anti-cyclic citrullinated peptide antibodies (anti-CCP) were determined in 209
plasma using ELIA Unicap (Phadia Laboratory Systems, Uppasala, Sweden) 210
and Rheumatoid factor (RF) was determined using Nephelometry system 211
(Beckman Coulter). Levels of anti-CCP antibodies and RF were expressed as 212
ELIAunits/ml and IU/ml respectively. Anti-CCP antibodies were positive with 213
levels > 20 ELIAunits/ml and RF were positive with levels >26 IU/ml. 214
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Statistics 216
Statistical analyses were performed using the probes t-test, Mann-Whitney and 217
Fisher exact test on Graph Pad Prism 5 software. Non-parametric correlation 218
Spearman test was applied to the analysis. Data were presented as mean and 219
± SEM deviation. P values <0.05 were considered significant. 220
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Results 223
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CD36 expression on lymphocytes 225
Most CD14+ monocytes from Ficoll-isolated peripheral blood expressed CD36 226
on their cell surface (90.8 ± 1.83%). Unexpectedly, 6.23 ± 1.23% of 227
lymphocytes (CD14 negative) were also CD36+. To rule out the possibility of 228
unspecific staining of the monoclonal antibody (clone CB38), the analysis was 229
repeated with another clone (TR9: 89.5 ± 1.74% on gated CD14+ monocytes 230
and 6.59 ± 1.55% on lymphocytes) (Fig. 1A). Monocytes and platelets express 231
CD36 on their surface. Since lymphocytes can interact with circulating platelets 232
[22] we speculated that the CD36 expression on lymphocytes was derived from 233
bound CD36+ platelets. Our results showed that those CD36+ lymphocytes 234
were also CD41a+ (a specific marker of platelets) (Fig. 1B). We observed that 235
both CD4+ CD36+ and CD8+ CD36+ lymphocytes were CD41a+. 236
Immunofluorescence microscopy finally confirmed that CD36 expression on 237
lymphocytes was due to bound platelets. In contrast to the whole surface CD36 238
staining of monocytes, the CD36 staining of the CD36+ lymphocyte minority 239
was in a small protruding area of the surface. The image was compatible with a 240
platelet bound to a lymphocyte. A comparable image was observed staining 241
with CD41a+. These observations confirmed that the source of this marker is 242
the presence of platelets bound to lymphocytes (Fig. 1C). We observed that 243
stoichiometry between platelet and T cells were 1 to 1 in the majority of cells 244
(73%). 245
We next analyzed CD36 expression on CD4+ and CD8+ cells from whole blood. 246
We observed a comparable CD36 expression on CD4+ and CD8+ cells from 247
whole blood and Ficoll isolated mononuclear cells (data not shown). This result 248
confirms that the binding between lymphocytes and platelets was neither a 249
separation artifact nor dependent on the proportion of platelets per lymphocyte. 250
CD36 expression on lymphocytes was then a convenient marker to study 251
lymphocytes with bound platelets. 252
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Phenotyping of CD3 naive and CD3 memory lymphocytes with bound 256
platelets 257
We firstly investigated whether the presence of bound platelets on naïve and 258
memory lymphocytes is restricted to a characteristic phenotype. The 259
percentage of naïve CD3+ lymphocytes with bound platelets was not different 260
from memory CD3+ lymphocytes without platelets (naïve CD3+: 61.1 ± 3.33 of 261
Plt-ng vs 55.4 ± 6.05 % of Plt-pos; memory CD3+: 38.9 ± 5.29 % of Plt-ng vs 262
44.6 ± 6.37 of Plt-pos). A significant higher percentage of CXCR3+ and CCR4+ 263
cells was observed in the naïve CD3+ cells with platelets than in naïve CD3+ 264
cells without platelets (CCR4: 2.33 ± 0.53 of Plt-ng vs 4.71 ± 0.85 of Plt-pos, 265
p=0.02; CXCR3: 8.81 ± 1.93 of Plt-ng vs 13.98 ± 3.47 of Plt-pos, p=0.04). In 266
contrast, a significant higher percentage of CD25+, CCR5+, CCR4+ and HLA-267
DR+ cells were observed in memory CD3+ cells with bound platelets than in 268
memory CD3+ without platelets (CD25: 3.43 ± 0.65 of Plt-ng vs 7.47 ± 1.61 of 269
Plt-pos, p=0.01; HLA-DR: 1.98 ± 0.23 of Plt-ng vs 4.83 ± 0.55 of Plt-pos, 270
p=0.01) and chemokine receptor CCR5 (32.4 ± 3.05 of Plt-ng vs 44.11 ± 4.69 of 271
Plt-pos, p=0.005) and CCR4 (15.37 ± 2.24 of Plt-ng vs 20.86 ± 3.04 of Plt-pos, 272
p=0.04) (Table 1). 273
We next evaluated whether the presence of bound platelets was associated to 274
Th1 (CCR5 and CXCR3) or Th2 (CCR4) markers. Significant higher 275
percentages of CCR5+ were observed in the CD4+ cells with bound platelets 276
than in CD4+ cells without platelets, suggesting that platelet are predominantly 277
bound to cells expressing certain Th1 markers (CCR5: 11.99 ± 1.01 of Plt-ng vs 278
21.59 ± 1.81 of Plt-pos, p=0.0004; CXCR3: 20.44 ± 2.09 of Plt-ng vs 25.76 ± 279
2.74 of Plt-pos, p=0.14). 280
We finally observed no differences in the percentage of naïve, effector and 281
central memory in CD4+ T cells with or without platelets (data not shown). 282
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Reduced TCR-induced proliferation of CD4+Plt-pos lymphocytes 284
We compared the TCR-induced proliferation of CD4+ Plt-pos and CD4+ Plt-ng 285
cells. PBMC from healthy donors were labelled with CFSE and stimulated with 286
anti-CD3, CD28 and CD2 T cell activation/expansion for 72 hours and 7 days 287
(Fig. 2). After 72 hours and 7 days of culture, we observed that fewer CD4+ Plt-288
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pos than CD4+ Plt-ng lymphocytes reached a higher number of proliferation 289
cycles (72h, cycle 3: 8.18 ± 3.24% of CD4+ Plt-pos vs 17.67 ± 1.89% of CD4+ 290
Plt-ng, p<0.05; 7d, cycle 4: 7.87 ± 2.9% of CD4+ Plt-pos vs 28 ± 0.33 of CD4+ 291
Plt-ng, p<0.05) (Fig. 2B and 2C). Stimulation index (SI) was also calculated in 292
CD4+ Plt-ng and CD4+ Plt-pos. CD4+ with platelets had a lower SI than CD4+ 293
without platelets (72 hours: 1.49 ± 0.17 for CD4+ Plt-ng vs 0.55 ± 0.17 for CD4+ 294
Plt-pos, p<0.01; 7d: 4.22 ± 0.37 for CD4+ Plt-ng vs 1.74 ± 0.05 for CD4+ Plt-295
pos, p<0.01) (Supplemental figure 1). The binding of platelets to lymphocytes 296
can therefore reduce the proliferation capacity of the CD4+ cells. 297
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Binding of platelets modified the cytokine production of CD3+ 299
lymphocytes 300
PBMC from healthy donors were stimulated with anti-CD3, CD28 and CD2 T 301
cell activation/expansion kit for 48h. Five hours before cytokine production 302
analysis, PMA/Ionomycin and BD GolgiStop TM Protein Transport Inhibitor was 303
added to the lymphocyte culture. We then compared the intracellular cytokine 304
production of CD3+ Plt-pos and CD3+ Plt-ng lymphocytes. 305
Although there were no differences in the percentage of cells producing IL-17, 306
IFNγ, IL-10 and TNFα, binding platelets modified the amount of cytokine levels 307
in lymphocytes (iGMFI). CD3+ Plt-ng cells expressed higher levels of IL-17 and 308
IFNγ than CD3+ Plt-pos cells (iGMFI IL-17: 67.94 ± 25.89 in CD3+ Plt-ng vs 309
19.5 ± 10.73 in CD3+ Plt-pos, p<0.05; iGMFI IFNγ: 340 ± 97.53 in CD3+ Plt-ng 310
vs 172 ± 57.66 in CD3+ Plt-pos, p<0.05). No differences were observed in IL-10 311
or TNFα levels between the two lymphocyte populations (iGMFI IL-10: 1.36 ± 312
0.49 on CD3+ Plt-ng vs 8.54 ± 3.62 on CD3+ Plt-pos; iGMFI TNFα: 245 ± 137.1 313
on CD3+ Plt-ng vs 153 ± 35.34 on CD3+ Plt-pos) (Fig 3). These results suggest 314
that the binding of platelets decreased the production of Th1 and Th17 315
cytokines. 316
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CD4+ Plt-pos and cytokine plasma levels from healthy donors and RA 318
patients 319
Activation molecule P-selectin has a significantly higher expression on platelets 320
from active RA patients than from remission RA patients or healthy donors [23]. 321
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P-selectin can bind to its ligand PSGL1 expressed on leukocytes [24-25]. We 322
observed that platelets bound to lymphocytes in RA patients present higher 323
levels of P-selectin than healthy donors (37.11 ± 6.18 for RA patients vs.13.35 ± 324
1.76 for healthy donors p<0.05) (data not shown). We then compared the 325
percentage of T lymphocytes with bound platelets in the peripheral blood of RA 326
patients and healthy donors. The percentage of CD8+ Plt-pos was similar in 327
healthy donors and RA patients (data not shown). However, the percentage of 328
CD4+ Plt-pos in RA patients was significantly higher than in healthy donors 329
(7.81 ± 0.84% and 4.46 ± 0.51% respectively, p<0.01). According to the CD4+ 330
Plt-pos percentage, we distinguished two groups of RA patients. One group 331
(group-H) presented higher CD4+Plt-pos percentage (10.61 ± 0.68%) than 332
healthy donors. The other group (group-L) presented CD4+ Plt-pos levels (3.79 333
± 0.44%) comparable to healthy donors (4.46 ± 0.51%) (Fig. 4A and 4B). 334
Clinical and laboratory parameters were compared between these two groups 335
of RA patients (Supplemental table 1). Cardiovascular risk index were 336
significantly different between these groups. Plasma IL-17 and IFNγ levels in 337
group-L of RA patients presented higher plasma levels of IL-17 (189.7 ± 70.83 338
pg/ml) and IFNγ (147.3 ± 36.7 pg/ml) than healthy donors and group-H RA 339
patients (IL-17: 38.28 ± 18.24 pg/ml for healthy donors and 51.13 ± 11.26 pg/ml 340
for group-H RA patients, p<0.05; IFNγ: 43.25 ± 20.33 pg/ml for healthy donors 341
and 67.76 ± 15.55 pg/ml for group-H RA patients, p<0.05). These results 342
suggest that increased levels of CD4+ lymphocytes with bound platelets have a 343
regulatory effect in the excessive Th1 and Th17 cytokine levels of RA patients. 344
We did not observe a significant correlation between the proportion of CD4+ Plt-345
pos with serum levels of cytokines in healthy donors. However, a negative 346
correlation was observed between CD4+ Plt-pos and IFNγ in RA patients (r=-347
0.47, p<0.05). 348
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Percentage of CD4+Plt-pos lymphocytes in RA patients segregated 350
according to clinical parameters 351
We analyzed the association between CD4+ Plt-pos and antibody levels and 352
clinical parameters in RA patients (anti-CCP, RF, CRP, ESR, DAS 28 and score 353
cardiovascular risk index). Negative or moderate-low anti-CCP titer (<340) RA 354
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patients showed a higher percentage of CD4+ Plt-pos than high anti-CCP titer 355
RA patients (10.01 ± 0.93% for titers <20, 9.98 ± 1.72% for 20-340 titers and 356
5.64 ± 0.64% for titers >340, p<0.05) (Fig 5A). Using a rank based correlation, 357
anti-CCP antibodies inversely correlated with CD4+ Plt-pos (R= -0.443, 358
p=0.05). RA patients with lower RF levels (<26) showed a higher percentage of 359
CD4+ Plt-pos than RA patients with RF levels >160 RF (10.08 ± 1.42% and 360
5.49 ± 1.11% respectively, p<0.05)(Fig 5B). RA patients with lower CRP levels 361
(≤10 mg/L) showed a higher percentage of CD4+ Plt-pos than RA patients with 362
CRP>10 mg/L (9.74 ± 1.5% and 4.19 ± 0.54% respectively, p<0.05) (Fig 5C). 363
No differences in CD4+ Plt-pos percentages were observed when patients were 364
segregated according to ESR levels and DAS28 (Fig 5D and 5E). RA patients 365
with a lower cardiovascular risk index (≤3) presented a higher percentage of 366
CD4+ Plt-pos lymphocytes than patients with higher cardiovascular risk index 367
(>5) (9.01 ± 1.16% and 3.84 ± 0.58% respectively, p<0.05). CD4+ Plt-pos in RA 368
patients segregated according to medication were not significantly different 369
(data not shown) 370
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Discussion 390
Our results demonstrated that the CD36 expression on lymphocytes was 391
derived from bound platelets. Platelet binding reduced the capability of 392
lymphocytes to proliferate and to produce Th1/Th17 cytokines. Less IL-17, IFNγ 393
and cardiovascular risk was observed in a group of RA patients with a higher 394
percentage of CD4+ cells with bound platelets. 395
Vallejo et al. demonstrated that T lymphocytes do not express CD36 [2]. 396
However, other reports described a subpopulation of B, T and NK lymphocytes 397
expressing CD36 [3-4]. Some authors have already suggested the presence of 398
platelet-leukocyte complexes [26-27]. Accordingly, we observed that these 399
platelets were the source of the unexpected CD36 expression on this 400
subpopulation of lymphocytes. We were able to finally confirm this result by 401
microscopy. Presence of CD36 on lymphocytes was not due to trogocytosis of 402
membrane chunks of antigen presenting cells because platelet-bound 403
lymphocytes also expressed CD41a, a specific marker of platelets, absent on 404
antigen presenting cells. Most of the CD36+ lymphocytes had one bound 405
platelet (73%). Undergoing analysis of bound-platelet characteristics will provide 406
some explanation to this stoichiometry. 407
Staining lymphocytes with anti-CD36 antibodies proved to be a good tool to 408
analyze the functional consequences of platelet binding. Significant phenotype 409
differences were observed between CD3+ Plt-pos and CD3+ Plt-ng in naïve 410
and memorty T cells. CD3+ Plt-pos memory cells present a higher expression of 411
DR+, CD25+, CCR5+ and CCR4+, while CD3+ Plt-pos naïve present a higher 412
expression of CXCR3+ and CCR4+. We are currently analyzing a predominant 413
subpopulation of CCR5+ CD4+ T lymphocytes among those with bound 414
platelets. 415
Multiple ligands could be involved in this platelet-lymphocyte binding. P-selectin 416
(CD62P) on activated platelets can bind to its ligand PSGL-1 (CD162), 417
expressed on most leukocytes [25]. GPIb-CD11b [28] and GPIIb/IIIa-418
CD11/CD18 [29] could also be involved in platelet-leukocyte binding. It has 419
been shown that platelet-lymphocyte binding, initiated by platelet activation, can 420
be abolished by P-selectin blockade and reduced by inhibition of GPIIb/IIIa, 421
15
CD11b or CD40L [22]. We have found that <15% of platelets bound to 422
lymphocytes are P-selectin positive and this seem contradictory. However, we 423
can not rule out that bound platelet were activated when bound to lymphocyte 424
and then CD62P were released for the surface of platelets due to shedding of 425
this molecule [30]. 426
Only 3% of circulating lymphocytes carried bound platelets. This low percentage 427
could be due to the binding of few free activated platelets on circulation [27]. 428
Accordingly, in vitro activation of platelets increases the lymphocyte-platelet 429
binding (12% for Th lymphocytes and 15% by Tc lymphocytes) [22]. Activation 430
of T lymphocytes also increases the binding of platelets [22] probably because 431
activation upregulates PSGL-1 expression. This upregulation could explain the 432
superior capacity of platelet binding by activated lymphocytes [24]. Accordingly, 433
we could speculate that a higher percentage of CCR5+ CD4+ lymphocytes 434
have bound platelets, because they express more PSGL-1. Previous reports 435
with juvenile idiopathic arthritis patients showed that CCR5+ CD4+ lymphocytes 436
from synovial fluid have higher PSGL-1 expression than CCR4+ and CXCR3+ 437
CD4+ lymphocytes [31]. 438
After activation, lymphocytes with bound platelets have a reduced proliferation 439
and IFNγ and IL-17 production. Previous reports showed that co-culture with 440
activated platelets reduces the activation-induced proliferation of lymphocytes 441
[32-33] and IL-2 and IL-6 release [33]. However, other authors have shown that 442
co-culturing activated platelets with lymphocytes increases the release of IFNγ, 443
TNFα and IL-10 [32]. These reports suggest that soluble molecules released by 444
platelets could be activating or inhibiting factors [34]. One of these molecules, 445
PF4, was able to modulate Treg and non-Treg cells but in opposite manners. 446
PF4 inhibits non-Treg cell proliferation and also reduced Treg ability to inhibit 447
non-Treg proliferation [35]. In our system, we cannot exclude the presence of 448
inhibitory soluble factors released by activated platelets. However, the inhibitory 449
effects were more evident in lymphocytes with bound platelets. Furthermore, 450
our experiments were performed with isolated PBMC after eliminating most free 451
platelets through the Ficoll gradient. 452
Our results suggest that platelet binding to lymphocytes could be relevant in the 453
pathology of inflammatory diseases. We distinguished two groups of RA 454
16
patients according to the percentage of CD4+ Plt-pos. One group with CD4+Plt-455
pos was comparable to healthy donors and the other group had a higher 456
CD4+Plt-pos percentage. This was not surprising because platelets in active 457
RA patients show higher expression of P-selectin than those in remission or 458
healthy controls [23], observing in our experiment that platelets bound to 459
lymphocyte in RA patients present higher percentage of activated platelet than 460
healthy donors. The plasma levels of IL-17 and IFNγ were significantly higher in 461
patients with normal CD4+ Plt percentage. This result was not observed in 462
patients with higher CD4+ Plt-pos percentage, concordant with the lower 463
production of IL-17 and IFNγ by lymphocytes with bound platelets. It has been 464
described that RA pathology is caused partially by an increased Th1/Th17 465
response, with high levels of inflammatory cytokines on serum [36-37]. We 466
suggest that binding of platelets could then be a regulatory mechanism to 467
correct the altered production of inflammatory cytokines in RA patients. 468
RA patients with the highest levels of CRP, rheumatoid factor and anti-CCP 469
antibodies are those with the lowest CD4+Plt-pos levels. Binding of platelets to 470
CD4+ lymphocytes might therefore be associated with a less severe phenotype 471
of RA disease. These results are intriguing because platelets have been shown 472
to participate in the pathogenesis of RA and atherogenesis, a complication of 473
RA [38]. Free activated platelets shed vesicles called microparticles; during 474
inflammatory arthritis, these microparticles are abundant in synovial fluid and 475
can exacerbate inflammation via IL-1. This cytokine can promote release of 476
inflammatory cytokines such as IL6 and IL8 [39]. Moreover, platelets also 477
produce prostaglandins. PGI2 is increased in synovial fluid from the joints of RA 478
patients and contributes to arthritis in animal models [40]. 479
Patients with the highest cardiovascular risk also had the lowest levels of CD4+ 480
Plt-pos. However, it has been described that platelets are associated with 481
cardiovascular risk. CD62P levels on platelets increase in cardiovascular 482
ischemia associated with atherosclerotic complications [41-42]. This apparent 483
contradiction could be explained by a different function between free and bound 484
activated platelets. We should also consider other factors apparently unrelated 485
to platelets, like anti-CCP antibodies and IL-17, which contribute to the 486
development of endothelial dysfunction and cardiovascular disease [10, 43]. In 487
these cases, our patients with the highest levels of CD4+ Plt-pos could have 488
17
had a lower cardiovascular risk because they had a lower production of IL-17 489
and anti-CCP antibodies. 490
Further experiments are needed to decipher the mechanisms of platelet binding 491
and determine how and when these platelets can regulate lymphocyte 492
functions. Similarly, we need to study the influence of platelets on other 493
subpopulations. Ongoing experiments will describe whether this mechanism is 494
RA specific or whether it could be extended to other inflammatory pathologies. 495
The platelet-lymphocyte interaction could open new therapeutic approaches to 496
regulate the chronic inflammation. 497
498
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18
523
524
Authorship 525
CZ Flow cytometry, ELISAs, cell culture and data analysis. Manuscript 526
preparation 527
EC Flow cytometry 528
JCN Flow cytometry 529
MAO ELISAs 530
CDT Collection of samples and clinical parameters 531
CDL Collection of samples and clinical parameters 532
JL Collection of samples and clinical parameters 533
CJ Analysis of results 534
SV Analysis of results, conception and design. Manuscript preparation 535
536
Acknowledgements 537
This research was supported by the Ministerio Educacion y Ciencia 538
(BFU06/15063) and Fondo de Investigaciones Sanitarias (PS09/132). S.V. was 539
supported by “Fondo Investigaciones Sanitarias” and participant in the Program 540
for Stabilization of Investigators of the “Direcció d'Estrategia i Coordinació del 541
Departament Salut de la Generalitat de Catalunya.” 542
We thank Carolyn Newey for helpful editorial assistance and Ignasi Gich for his 543
statistical assistance. 544
This work has been performed within the Ph.D Immunology program of 545
Universitat Autònoma de Barcelona (UAB). 546
547
Conflicts of Interest disclosure 548
The authors declare no financial or commercial conflicts of interest associated 549
with this study. 550
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559
References 560
1. Febbraio, M., Hajjar, D. P., Silverstein, R. L. (2001) CD36: a class B scavenger 561
receptor involved in angiogenesis, atherosclerosis, inflammation, and lipid 562
metabolism. J Clin Invest 108, 785-91. 563
2. Vallejo, A. N., Mugge, L. O., Klimiuk, P. A., Weyand, C. M., Goronzy, J. J. 564
(2000) Central role of thrombospondin-1 in the activation and clonal expansion 565
of inflammatory T cells. J Immunol 164, 2947-54. 566
3. McDonnell, M., Liang, Y., Noronha, A., Coukos, J., Kasper, D. L., Farraye, F. 567
A., Ganley-Leal, L. M. (2011) Systemic Toll-like receptor ligands modify B-cell 568
responses in human inflammatory bowel disease. Inflamm Bowel Dis 17, 298-569
307. 570
4. Lubick, K. and Jutila, M. A. (2006) LTA recognition by bovine gammadelta T 571
cells involves CD36. J Leukoc Biol 79, 1268-70. 572
5. Tassone, P., Tuccillo, F., Bonelli, P., Turco, M. C., Cecco, L., Cerra, M., Bond, 573
H. M., Barbieri, V., Venuta, S. (1998) CD36 is rapidly and transiently 574
upregulated on phytohemagglutinin (PHA)-stimulated peripheral blood 575
lymphocytes. Analysis by a new monoclonal antibody (UN7). Tissue Antigens 576
51, 671-5. 577
6. Harrington, L. E., Hatton, R. D., Mangan, P. R., Turner, H., Murphy, T. L., 578
Murphy, K. M., Weaver, C. T. (2005) Interleukin 17-producing CD4+ effector T 579
cells develop via a lineage distinct from the T helper type 1 and 2 lineages. Nat 580
Immunol 6, 1123-32. 581
7. Bettelli, E., Carrier, Y., Gao, W., Korn, T., Strom, T. B., Oukka, M., Weiner, H. 582
L., Kuchroo, V. K. (2006) Reciprocal developmental pathways for the 583
generation of pathogenic effector TH17 and regulatory T cells. Nature 441, 235-584
8. 585
8. Kotake, S., Udagawa, N., Takahashi, N., Matsuzaki, K., Itoh, K., Ishiyama, S., 586
Saito, S., Inoue, K., Kamatani, N., Gillespie, M. T., Martin, T. J., Suda, T. (1999) 587
IL-17 in synovial fluids from patients with rheumatoid arthritis is a potent 588
stimulator of osteoclastogenesis. J Clin Invest 103, 1345-52. 589
9. Pickens, S. R., Volin, M. V., Mandelin, A. M., 2nd, Kolls, J. K., Pope, R. M., 590
Shahrara, S. (2010) IL-17 contributes to angiogenesis in rheumatoid arthritis. J 591
Immunol 184, 3233-41. 592
20
10. Marder, W., Khalatbari, S., Myles, J. D., Hench, R., Yalavarthi, S., Lustig, S., 593
Brook, R., Kaplan, M. J. (2011) Interleukin 17 as a novel predictor of vascular 594
function in rheumatoid arthritis. Ann Rheum Dis 70, 1550-5. 595
11. Maggi, L., Santarlasci, V., Capone, M., Peired, A., Frosali, F., Crome, S. Q., 596
Querci, V., Fambrini, M., Liotta, F., Levings, M. K., Maggi, E., Cosmi, L., 597
Romagnani, S., Annunziato, F. (2010) CD161 is a marker of all human IL-17-598
producing T-cell subsets and is induced by RORC. Eur J Immunol 40, 2174-81. 599
12. Chalan, P., Abdulahad, W., Huitema, M. G., Kroesen, B. J., Brouwer, E., Boots, 600
A. M. H. (2011) Rheumatoid arthritis patients shown a decline in peripheral 601
blood CD4+CD161+T lymphocytes ANNALS OF THE RHEUMATIC DISEASES 602
70, A6-A6 603
13. Ruschpler, P. and Stiehl, P. (2002) Shift in Th1 (IL-2 and IFN-gamma) and Th2 604
(IL-10 and IL-4) cytokine mRNA balance within two new histological main-types 605
of rheumatoid-arthritis (RA). Cell Mol Biol (Noisy-le-grand) 48, 285-93. 606
14. Sundstedt, A., O'Neill, E. J., Nicolson, K. S., Wraith, D. C. (2003) Role for IL-10 607
in suppression mediated by peptide-induced regulatory T cells in vivo. J 608
Immunol 170, 1240-8. 609
15. Xu, D., Liu, H., Komai-Koma, M., Campbell, C., McSharry, C., Alexander, J., 610
Liew, F. Y. (2003) CD4+CD25+ regulatory T cells suppress differentiation and 611
functions of Th1 and Th2 cells, Leishmania major infection, and colitis in mice. J 612
Immunol 170, 394-9. 613
16. Kawashiri, S. Y., Kawakami, A., Okada, A., Koga, T., Tamai, M., Yamasaki, S., 614
Nakamura, H., Origuchi, T., Ida, H., Eguchi, K. (2011) 615
CD4+CD25(high)CD127(low/-) Treg cell frequency from peripheral blood 616
correlates with disease activity in patients with rheumatoid arthritis. J 617
Rheumatol 38, 2517-21. 618
17. Arnett, F. C., Edworthy, S. M., Bloch, D. A., McShane, D. J., Fries, J. F., 619
Cooper, N. S., Healey, L. A., Kaplan, S. R., Liang, M. H., Luthra, H. S., et al. 620
(1988) The American Rheumatism Association 1987 revised criteria for the 621
classification of rheumatoid arthritis. Arthritis Rheum 31, 315-24. 622
18. Aletaha, D. and Smolen, J. S. (2006) The definition and measurement of 623
disease modification in inflammatory rheumatic diseases. Rheum Dis Clin North 624
Am 32, 9-44, vii. 625
19. Conroy, R. M., Pyorala, K., Fitzgerald, A. P., Sans, S., Menotti, A., De Backer, 626
G., De Bacquer, D., Ducimetiere, P., Jousilahti, P., Keil, U., Njolstad, I., 627
Oganov, R. G., Thomsen, T., Tunstall-Pedoe, H., Tverdal, A., Wedel, H., 628
Whincup, P., Wilhelmsen, L., Graham, I. M. (2003) Estimation of ten-year risk of 629
21
fatal cardiovascular disease in Europe: the SCORE project. Eur Heart J 24, 630
987-1003. 631
20. Darrah, P. A., Patel, D. T., De Luca, P. M., Lindsay, R. W., Davey, D. F., Flynn, 632
B. J., Hoff, S. T., Andersen, P., Reed, S. G., Morris, S. L., Roederer, M., Seder, 633
R. A. (2007) Multifunctional TH1 cells define a correlate of vaccine-mediated 634
protection against Leishmania major. Nat Med 13, 843-50. 635
21. Milovanova, T. N. (2007) Comparative analysis between CFSE flow cytometric 636
and tritiated thymidine incorporation tests for beryllium sensitivity. Cytometry B 637
Clin Cytom 72, 265-75. 638
22. Li, N., Ji, Q., Hjemdahl, P. (2006) Platelet-lymphocyte conjugation differs 639
between lymphocyte subpopulations. J Thromb Haemost 4, 874-81. 640
23. Wang, F., Wang, N. S., Yan, C. G., Li, J. H., Tang, L. Q. (2007) The 641
significance of platelet activation in rheumatoid arthritis. Clin Rheumatol 26, 642
768-71. 643
24. Moore, K. L., Stults, N. L., Diaz, S., Smith, D. F., Cummings, R. D., Varki, A., 644
McEver, R. P. (1992) Identification of a specific glycoprotein ligand for P-645
selectin (CD62) on myeloid cells. J Cell Biol 118, 445-56. 646
25. Furie, B. and Furie, B. C. (1995) The molecular basis of platelet and endothelial 647
cell interaction with neutrophils and monocytes: role of P-selectin and the P-648
selectin ligand, PSGL-1. Thromb Haemost 74, 224-7. 649
26. Furman, M. I., Benoit, S. E., Barnard, M. R., Valeri, C. R., Borbone, M. L., 650
Becker, R. C., Hechtman, H. B., Michelson, A. D. (1998) Increased platelet 651
reactivity and circulating monocyte-platelet aggregates in patients with stable 652
coronary artery disease. J Am Coll Cardiol 31, 352-8. 653
27. Joseph, J. E., Harrison, P., Mackie, I. J., Isenberg, D. A., Machin, S. J. (2001) 654
Increased circulating platelet-leucocyte complexes and platelet activation in 655
patients with antiphospholipid syndrome, systemic lupus erythematosus and 656
rheumatoid arthritis. Br J Haematol 115, 451-9. 657
28. Simon, D. I., Chen, Z., Xu, H., Li, C. Q., Dong, J., McIntire, L. V., Ballantyne, C. 658
M., Zhang, L., Furman, M. I., Berndt, M. C., Lopez, J. A. (2000) Platelet 659
glycoprotein ibalpha is a counterreceptor for the leukocyte integrin Mac-1 660
(CD11b/CD18). J Exp Med 192, 193-204. 661
29. Silverstein, R. L., Asch, A. S., Nachman, R. L. (1989) Glycoprotein IV mediates 662
thrombospondin-dependent platelet-monocyte and platelet-U937 cell adhesion. 663
J Clin Invest 84, 546-52. 664
30. Dole, V. S., Bergmeier, W., Patten, I. S., Hirahashi, J., Mayadas, T. N., Wagner, 665
D. D. (2007) PSGL-1 regulates platelet P-selectin-mediated endothelial 666
22
activation and shedding of P-selectin from activated platelets. Thromb Haemost 667
98, 806-12. 668
31. Issekutz, A. C., Quinn, P. J., Lang, B., Ramsey, S., Huber, A. M., Rowter, D., 669
Karkada, M., Issekutz, T. B. (2011) Coexpression of chemokine receptors 670
CCR5, CXCR3, and CCR4 and ligands for P- and E-selectin on T lymphocytes 671
of patients with juvenile idiopathic arthritis. Arthritis Rheum 63, 3467-76. 672
32. Gerdes, N., Zhu, L., Ersoy, M., Hermansson, A., Hjemdahl, P., Hu, H., 673
Hansson, G. K., Li, N. (2011) Platelets regulate CD4(+) T-cell differentiation via 674
multiple chemokines in humans. Thromb Haemost 106, 353-62. 675
33. Wang, Y. and Niu, J. (2008) Platelets inhibit in vitro response of lymphocytes to 676
mitogens. Immunol Lett 119, 57-61. 677
34. Fleischer, J., Grage-Griebenow, E., Kasper, B., Heine, H., Ernst, M., Brandt, E., 678
Flad, H. D., Petersen, F. (2002) Platelet factor 4 inhibits proliferation and 679
cytokine release of activated human T cells. J Immunol 169, 770-7. 680
35. Liu, C. Y., Battaglia, M., Lee, S. H., Sun, Q. H., Aster, R. H., Visentin, G. P. 681
(2005) Platelet factor 4 differentially modulates CD4+CD25+ (regulatory) versus 682
CD4+CD25- (nonregulatory) T cells. J Immunol 174, 2680-6. 683
36. Tukaj, S., Kotlarz, A., Jozwik, A., Smolenska, Z., Bryl, E., Witkowski, J. M., 684
Lipinska, B. (2010) Cytokines of the Th1 and Th2 type in sera of rheumatoid 685
arthritis patients; correlations with anti-Hsp40 immune response and diagnostic 686
markers. Acta Biochim Pol 57, 327-32. 687
37. Ziolkowska, M., Koc, A., Luszczykiewicz, G., Ksiezopolska-Pietrzak, K., 688
Klimczak, E., Chwalinska-Sadowska, H., Maslinski, W. (2000) High levels of IL-689
17 in rheumatoid arthritis patients: IL-15 triggers in vitro IL-17 production via 690
cyclosporin A-sensitive mechanism. J Immunol 164, 2832-8. 691
38. Boilard, E., Blanco, P., Nigrovic, P. A. (2012) Platelets: active players in the 692
pathogenesis of arthritis and SLE. Nat Rev Rheumatol 8, 534-42. 693
39. Boilard, E., Nigrovic, P. A., Larabee, K., Watts, G. F., Coblyn, J. S., Weinblatt, 694
M. E., Massarotti, E. M., Remold-O'Donnell, E., Farndale, R. W., Ware, J., Lee, 695
D. M. (2010) Platelets amplify inflammation in arthritis via collagen-dependent 696
microparticle production. Science 327, 580-3. 697
40. Honda, T., Segi-Nishida, E., Miyachi, Y., Narumiya, S. (2006) Prostacyclin-IP 698
signaling and prostaglandin E2-EP2/EP4 signaling both mediate joint 699
inflammation in mouse collagen-induced arthritis. J Exp Med 203, 325-35. 700
41. McCabe, D. J., Harrison, P., Sidhu, P. S., Brown, M. M., Machin, S. J. (2004) 701
Circulating reticulated platelets in the early and late phases after ischaemic 702
stroke and transient ischaemic attack. Br J Haematol 126, 861-9. 703
23
42. Yamazaki, M., Uchiyama, S., Iwata, M. (2001) Measurement of platelet 704
fibrinogen binding and p-selectin expression by flow cytometry in patients with 705
cerebral infarction. Thromb Res 104, 197-205. 706
43. Lopez-Longo, F. J., Oliver-Minarro, D., de la Torre, I., Gonzalez-Diaz de 707
Rabago, E., Sanchez-Ramon, S., Rodriguez-Mahou, M., Paravisini, A., 708
Monteagudo, I., Gonzalez, C. M., Garcia-Castro, M., Casas, M. D., Carreno, L. 709
(2009) Association between anti-cyclic citrullinated peptide antibodies and 710
ischemic heart disease in patients with rheumatoid arthritis. Arthritis Rheum 61, 711
419-24. 712
713
714
715
716
24
Figure Legends 717
718
Fig. 1. CD36 expression on monocytes and lymphocytes after Ficoll 719
isolation. PBMC from healthy donors (n=10) were Ficoll gradient isolated. Cells 720
were then stained with anti CD3, CD4, CD8, CD14, CD36 (clone CB38,and 721
clone TR9), and CD41a. (A) Expression of CD14-PEDy647, CD36-PE and 722
CD36-FITC were analyzed on monocytes and lymphocytes gated according to 723
FS and SS parameters and (B) Expression of CD41a-FITC and CD36-PE were 724
analyzed on CD4+ and CD8+ lymphocytes by flow cytometry gated on CD3+ 725
lymphocytes. (C) CD36 expression was analyzed on CD3+ and CD14+ cells 726
and CD41a on CD3+ cells using fluorescence microscopy. A representative 727
experiment is shown. Data are expressed as percentage ± SEM of analyzed 728
marker. 729
730
Fig. 2. Influence of binding platelet to CD4+ lymphocytes on activation-731
induced proliferation. PBMC from healthy donors were labelled with 7 μM of 732
CFSE. PBMC were then cultured with anti-CD3 plus CD2 and CD28 or medium 733
for 72h and 7d. After culturing, PBMC were collected and stained with anti CD4-734
PECy7 and CD36-PE. (A) Representative experiment of history proliferation 735
cycles on CD4+ Plt-ng and CD4+ Plt-pos cells after 72h and 7d. (B) Bar graph 736
shown percentage of cells in each proliferation cycle after 72h and (C) 7 days of 737
culture with anti-CD3 plus CD2 and CD28 on CD4+ Plt-ng and CD4+ Plt-pos 738
cells (n=5). Mann-Whitney test was used for statistical analysis. * P <0.05 and 739
** P<0.01. 740
741
Fig. 3. Influence of binding platelet to CD3+ lymphocytes on cytokine 742
production. PBMC from healthy donors were cultured with anti-CD3 plus CD2 743
and CD28 or medium. After 48h, cells were cultured with PMA (50 744
ng/ml)/Ionomycin (1 μg/ml) in presence of BD GolgiStopTM Protein Transport 745
Inhibitor for five hours before analysis of cytokine production. (A) IL-17, (B) IL-746
10 (C) IFNγ and (D) TNFα iGMFI production cells was calculated gating on 747
CD3+ Plt-ng and CD3+ Plt-pos lymphocytes (n=5). Data are expressed as 748
iGMFI ± SEM. Mann-Whitney test was used for statistical analysis. * P<0.05. 749
25
Fig. 4. CD4+ Plt-pos and cytokine plasma levels in RA patients. PBMC 750
from healthy donors and RA patients were stained with anti CD4-PECy7 and 751
CD36-PE. (A) Graph shown percentage of Plt-pos ± SEM on CD4+ 752
lymphocytes from healthy donors (n=16) and RA patients (n=20). (B) 753
Representative experiment of CD36 expression on CD4+ lymphocytes from 754
healthy controls and groupH and groupL RA patients. (C) Bar graph showing IL-755
17, IFNγ and IL-10 plasma levels from healthy donors and RA patients (pg/ml). 756
T-test was used for statistical analysis. * P<0.05 ** P<0.01. 757
758
Fig. 5. CD4+ Plt-pos percentage according to levels of CCP, rheumatoid 759
factor, C-reactive protein, ESR, DAS28 and cardiovascular risk in RA 760
patients. We analyzed the percentage of CD4+ Plt-pos in those RA patients 761
segregated according to (A) anti-CCP, (B) RF, (C) CRP, (D) ESR, (E) DAS28 762
and (F) cardiovascular risk index. Data are expressed as mean of percentage ± 763
SEM. T-test was used for statistical analysis * P<0.05 764
CD3 naive* CD3 memory** Plt- Plt+ Plt- Plt+
Marker Mean ± SEM Mean ± SEM p value Mean ± SEM Mean ± SEM p value
CD25 0.86 ± 0.34 0.79 ± 0.24 0.86 3.43 ± 0.65 7.47 ± 1.61 0.01
CCR5 10.48 ± 1.76 17.16 ± 4.36 0.22 32.4 ± 3.05 44.11 ± 4.69 0.005
CCR6 4.06 ± 1.14 4.92 ± 1.01 0.28 35.42 ± 7.44 32.37 ± 6.53 0.09
CXCR3 8.81 ± 1.93 13.98 ± 3.47 0.04 35.32 ± 6.76 34.97 ± 6.96 0.86
CCR4 2.33 ± 0.53 4.71 ± 0.85 0.02 15.37 ± 2.24 20.86 ± 3.04 0.04
HLA-DR 1.11 ± 0.55 2.63 ± 1.06 0.08 1.98 ± 0.23 4.83 ± 0.55 0.01 *Percentage of each marker in the CD3+ naive cells with or without platelets (n=5) ** Percentage of each marker in the CD3+ memory with or without platelets (n=5) p value was calculated with Mann Whitney test
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