International Journal of Molecular Sciences Article Increase of Circulating Monocyte–Platelet Conjugates in Rheumatoid Arthritis Responders to IL-6 Blockage Anaís Mariscal 1,† , Carlos Zamora 2,† , César Díaz-Torné 3 , Ma Àngels Ortiz 2 , Juan José de Agustín 4 , Delia Reina 5 , Paula Estrada 5 , Patricia Moya 3 , Héctor Corominas 3 and Sílvia Vidal 2, * 1 Immunology Department, Hospital de la Santa Creu i Sant Pau, Biomedical Research Institute Sant Pau (IIB Sant Pau), 08041 Barcelona, Spain; amariscal@santpau.cat 2 Laboratory of Inflammatory Diseases, Hospital de la Santa Creu i Sant Pau, Biomedical Research Institute Sant Pau (IIB Sant Pau), 08041 Barcelona, Spain; carlosza@g86mail.com (C.Z.); mortiz@santpau.cat (M.À.O.) 3 Rheumatology Department, Hospital de la Santa Creu i Sant Pau, 08041 Barcelona, Spain; cdiazt@santpau.cat (C.D.-T.); pmoyaa@santpau.cat (P.M.); hcorominas@santpau.cat (H.C.) 4 Rheumatology Department, Hospital Vall d’Hebrón, 08035 Barcelona, Spain; jjagor@hotmail.com 5 Rheumatology Department, Hospital Moisès Broggi, Sant Joan Despí, 08970 Barcelona, Spain; deliareinasanz@gmail.com (D.R.); paulavestradaa@gmail.com (P.E.) * Correspondence: svidal@santpau.cat † These authors contributed equally to this work. Abstract: Platelets (PLT) bind to a significant percentage of circulating monocytes and this im- munomodulatory interaction is increased in several inflammatory and autoimmune conditions. The therapeutic blockage of IL-6 with Tocilizumab (TCZ) alters PLT and the phenotype and function of monocytes in rheumatoid arthritis (RA). However, the relationship between monocyte–PLT conju- gates (CD14+PLT+) and clinical and immunological variables and the regulation of this interaction by IL-6 blockage are still unknown. Here, we compared the presence of monocyte–PLT conjugates Citation: Mariscal, A.; Zamora, C.; (CD14+PLT+) and membrane CD162 expression using flow cytometry, and, by ELISA, the markers of Díaz-Torné, C.; Ortiz, M.À.; Agustín, J.J.d.; Reina, D.; Estrada, P.; Moya, P.; PLT activation (sCD62P and sCD40L) in healthy donors (HD) and patients with long-standing RA Corominas, H.; Vidal, S. Increase of before TCZ (baseline). We found higher percentages and absolute counts of CD14+PLT+, and higher Circulating Monocyte–Platelet plasmatic levels of sCD62P and sCD40L but lower CD162 expression on monocytes from RA patients Conjugates in Rheumatoid Arthritis than those from HD. Additionally, the levels of CD14+PLT+ inversely correlated with inflammatory Responders to IL-6 Blockage. Int. J. parameters. Interestingly, 95% of patients with lower percentages of CD14+PLT+ and only 63% of Mol. Sci. 2022, 23, 5748. https:// patients with higher percentages of CD14+PLT+ achieved a EULAR-defined response at four weeks doi.org/10.3390/ijms23105748 (p = 0.036). After TCZ, the percentage of CD14+PLT+ increased in 92% of RA patients who achieved Academic Editor: Isabella Russo 12 w-remission (p < 0.001). Our results suggest that the binding of PLTs has a modulatory effect, accentuated by the increased binding of PLTs to monocytes in response to the therapeutic blockage Received: 7 April 2022 of IL-6. Accepted: 17 May 2022 Published: 20 May 2022 Keywords: platelets; monocytes; rheumatoid arthritis; tocilizumab; immune modulation; Publisher’s Note: MDPI stays neutral inflammation; immunity with regard to jurisdictional claims in published maps and institutional affil- iations. 1. Introduction Rheumatoid arthritis (RA) is an inflammatory joint disease characterized by persistent Copyright: © 2022 by the authors. systemic inflammation, with extra-articular manifestations that can affect up to 41% of Licensee MDPI, Basel, Switzerland. patients [1]. Neutrophils, mast cells, T and B lymphocytes, and monocytes/macrophages This article is an open access article are involved in the development of inflammation in RA [2]. In particular, monocytes play distributed under the terms and an important role in the initiation, maintenance and the degree of activity of synovial conditions of the Creative Commons inflammation [3]. In RA, activated monocytes possess increased adherent abilities [4]. They Attribution (CC BY) license (https:// massively infiltrate inflammatory sites, where they differentiate into tissue macrophages creativecommons.org/licenses/by/ and type A synoviocytes [5,6]. The activation of these cells can also lead to the production 4.0/). of TNF-α, IL-1, IL-6, and IL-8, which are effector molecules involved in inflammation [4,7]. Int. J. Mol. Sci. 2022, 23, 5748. https://doi.org/10.3390/ijms23105748 https://www.mdpi.com/journal/ijms Int. J. Mol. Sci. 2022, 23, 5748 2 of 13 Platelets (PLTs) are also involved in inflammatory processes through the secretion of inflammatory mediators or through a direct interaction with leukocytes [8–11]. PLTs circu- late in an inactive state and can respond to a wide variety of stimuli, releasing cytokines, chemokines, and growth factors that are stored in granules [12–15]. PLTs from RA patients express higher P-selectin (CD62P) levels and produce greater amounts of soluble CD40 ligand (sCD40L), both of which are markers of PLT activation [16,17]. One of the factors in- volved in this increased PLT activation is anti-citrullinated protein antibodies (ACPA) [13]. Circulating activated PLTs can bind to other PLTs and leukocytes, setting off a cascade of events that contribute to the development, evolution, and resolution of systemic inflam- matory response. PLT adhesion to circulating leukocytes enhances leukocyte rolling on the endothelial wall, facilitating the transport of PLTs to joint space [18,19]. In healthy donors (HD), P-selectin glycoprotein ligand 1 (PSGL-1 or CD162) on leukocytes seems to be the main molecule involved in the interaction of PLTs and leukocytes when it binds to CD62P on activated PLTs [20,21]. However, other ligands can be involved in the binding of PLTs to monocytes: GPIb-CD11b [22], CD40-CD40L [23], GPIIb/IIIa-CD11/CD18 [24], EMMPRIN (CD147/basigin)-CD147/GPIV [25,26], TREM-1-TREM-1 ligand [27] and PADGEM [28]. We previously described an increased percentage of monocytes with bound PLTs in SLE patients. However, SLE monocytes have a decreased expression of CD162, suggesting that other molecules contribute to this binding [29]. Other authors have also reported the down-regulation of CD162 on myeloid innate cells in an inflammatory context or after leukocyte activation [30–33]. In HD, monocyte–PLT interaction decreases monocyte apopto- sis, increases the expression of co-stimulatory molecules and Fc gamma receptors, and leads to a phenotypic shift in CD14+CD16- towards CD14+CD16+ and a monocyte differentiation into macrophages [29,34–36]. Furthermore, monocytes with bound PLTs have increased phagocytosis ability [29]. PLTs can also dampen inflammatory responses by increasing IL-10 and decreasing IL-6 and TNF-α release by monocytes [37,38]. Like other authors, we have reported an increased percentage of monocytes with bound PLTs in several in- flammatory and autoimmune conditions, including RA [19,26,29,32,39–43]. Depending on the disease, these monocyte–PLTs aggregates can play pro- [44], or anti-inflammatory roles [29,32,45]. Specifically, PLTs from RA patients can induce HD monocytes to synthesize pro-inflammatory cytokines (TNFα and IL-6) through CD147 engagement [26]. Although RA methotrexate (MTX), a conventional synthetic disease-modifying anti- rheumatic drug (DMARDs), is the first-line treatment, not all patients achieve remission. Tocilizumab (TCZ) is indicated in patients who have failed two courses with conventional synthetic DMARDs or who have adverse prognostic markers [46]. TCZ is a humanized anti-human IL-6 receptor antibody that blocks the binding of IL-6 to its receptor, thereby preventing signalling. It has been shown that TCZ induces apoptosis, downregulates the expression of CD80 and suppresses the expression of IL-6 on stimulated monocytes from HD [47]. Additionally, in more than 10% of patients, PLTs have been affected by TCZ [48,49]. Therefore, the main goal of this study is to decipher whether a new mechanism of action of TCZ on RA regulates the binding of PLT to monocytes. First, we analysed monocyte–PLT conjugates, as well as markers of PLT activation, in a cohort of long-lasting RA patients that are refractory to standard treatment with DMARDs. Second, we studied the association between the frequency of monocytes with bound PLTs and the clinical and laboratory features of RA patients before TCZ treatment. Third, we investigated whether monocytes with bound PLTs can function as biomarkers of TCZ treatment response. Finally, we clarified the functional consequences of both IL-6 signalling and TCZ treatment over PLTs binding to monocytes in in vitro cultures. 2. Results 2.1. Circulating CD14+PLT+ in RA Patients We observed higher percentages and absolute counts of circulating CD14+PLT+ in RA than in HD (Figure 1A,B and Figure S1B). However, we did not find differences in absolute counts of monocytes between RA patients and HD (201.08 ± 77.12 CD14+/µL in 2. Results 2.1. Circulating CD14+PLT+ in RA Patients Int. J. Mol. Sci. 2022, 23, 5748 We observed higher percentages and absolute counts of circulating CD14+PLT+ 3 ofin 13 RA than in HD (Figure 1A,B and S1B). However, we did not find differences in absolute counts of monocytes between RA patients and HD (201.08 ± 77.12 CD14+/µL in RA pa- tients vs. 208.44 RA patients ± 54.23 vs. 208.44 CD14+/µL ± 54.23 CD14+/µLin HD; p = 0.8). in HD; We We p = 0.8). also found also founda adirect directcorrelation correlation between between the the percentage percentage andand absolute absolute counts counts of of CD14+PLT+ CD14+PLT+in inboth both RA RA patients patients and and HD HD (Figure (Figure 1C). 1C). When When we we compared compared membrane membrane CD162 CD162 (mCD162) (mCD162) expression, expression, we we found found that that monocytes monocytes from from RARA patients patients had had aa lower lower expression expression of of mCD162 mCD162 thanthan HD HD (Figure (Figure 1D1D and and S1B). FigureInS1B). addition, mCD162 In addition, expression mCD162 directly expression correlated directly correlatedwith withthe thepercentage percentage of of CD14+PLT+ CD14+PLT+ in in HD HD(Figure (Figure1E)1E) butbut correlated correlated inversely inversely in RAin RA patients patients (Figure (Figure 1F). 1F). mCD162 mCD162 expressionexpression did notwith did not correlate correlate withcounts absolute absolute counts of CD14+PLT+ of circulating circulating CD14+PLT+ in RA patientsin RA −0.199; p(r== 0.284) (r =patients −0.199;orpin HD (r or = 0.284) = 0.34; in HDp =(r0.306). = 0.34;Monocytes with or without p = 0.306). Monocytes withbound PLTs or without had similar bound PLTsmCD162 had similarexpressions mCD162(Figure 1G). There expressions (Figurewere noThere 1G). differences were noin soluble CD162 differences in (sCD162) soluble between CD162 RA patients (sCD162) between andRA HD (Figureand patients 1H). HD (Figure 1H). Figure 1. Monocytes with Figure with bound boundplatelets plateletsand andCD162 CD162expression onon expression monocytes monocytes from healthy from donors healthy do- nors (HD)(HD) and rheumatoid and rheumatoid arthritis arthritis (RA) patients. (RA) patients. (A) Percentage (A) Percentage and (B) numbers and (B) absolute absoluteofnumbers monocytesof monocytes with boundwith bound platelets platelets (CD14+PLT+) (CD14+PLT+) in HD and RA in patients. HD and (C) RACorrelation patients. (C) Correlation between between percentage and percentage and absolute absolute number number in of CD14+PLT+ of HD CD14+PLT+ in HD and and RA patients. (D)RA patients. CD162 (D) CD162 expression expression on monocytes on from monocytes from HD and RA patients. (E) Correlation between CD162 expression and percentage of HD and RA patients. (E) Correlation between CD162 expression and percentage of CD14+PLT+ in CD14+PLT+ in HD and (F) RA patients. (G) CD162 expression on CD14+PLT- or CD14+PLT+ in HD HD and (F) RA patients. (G) CD162 expression on CD14+PLT- or CD14+PLT+ in HD and RA patients. and RA patients. (H) Quantification of soluble CD162 (sCD162) in plasma from HD and RA pa- (H) Quantification tients. White circlesof soluble CD162 represent HD, and (sCD162) in plasma black circles fromRA represent HDpatients. and RAStatistical patients. analysis White circles was represent HD, and black circles represent RA patients. Statistical analysis was performed using the Mann–Whitney test for comparisons between HD and RA and Spearman’s correlation for correlation analysis. ** p < 0.01, *** p < 0.001, **** p < 0.0001. We did not find any association between the percentage or absolute counts of CD14+PLT+ and the plasmatic concentrations of IL-6, sIL-6R, IL-10, IL-22, LBP and VEGF (data not CD14+PLT+ and the plasmatic concentrations of IL-6, sIL-6R, IL-10, IL-22, LBP and VEGF (data not shown). However, we found that the percentage, but not the absolute counts, of CD14+PLT+ correlated with plasmatic IL-17 levels in RA patients but not in HD (r = 0.505; p = 0.006 and r = 0.163; p = 0.407 respectively). Int. J. Mol. Sci. 2022, 23, 5748 We next analysed surrogate soluble markers of PLT activation: sCD40L and sCD62P 4 of 13 [16]. Although we found that both molecules were higher in RA patients than in HD (Figure S2), no association was found between sCD40L or sCD62P and the percentage of CD14+PLT+ in RA patients or in HD (Figure S2C,D). In addition, we did not observe any shown). However, we found that the percentage, but not the absolute counts, of CD14+PLT+ association between sCD40L and sCD62P in RA patients or in HD (Figure S2E). correlated with plasmatic IL-17 levels in RA patients but not in HD (r = 0.505; p = 0.006 and r = 0.163; p = 0.407 respectively). 2.2. Relationship between CD14+PLT+ and mCD162 Expression on Monocytes with Clinical RA We next analysed surrogate soluble markers of PLT activation: sCD40L and sCD62P [16]. Parameters Although we found that both molecules were higher in RA patients than in HD (Figure S2), We found no association was no association found betweenorthe between sCD40L percentages sCD62P or the absolute and the percentage counts of of CD14+PLT+ in CD14+PLT+ RA patients andor inmCD162 HD (Figureexpression S2C,D). on monocytes In addition, weand didDAS28, SDAI, not observe anyor association CDAI scores before TCZ treatment. Since those disease activity indexes between sCD40L and sCD62P in RA patients or in HD (Figure S2E). integrate different parameters, we then analysed each parameter independently. We found that absolute counts of CD14+PLT+ 2.2. Relationshipwere inversely between correlated CD14+PLT+ with inflammatory and mCD162 Expression on markers MonocytesCRPwith(Figure 2A) and Clinical RAESRParameters (Figure 2B), but CD14+PLT+ percentages did not correlate with CRP and ESR (r = −0.4; Wepfound = 0.054nowith CRP and association r = −0.33; between p = 0.09 withorESR). the percentages mCD162 the absolute expression counts on mono- of CD14+PLT+ andcytes correlated mCD162 with joint expression US scores and on monocytes (Figure 2C) and DAS28, SDAI,health assessment or CDAI questionnaire scores before TCZ treatment. Since(Figure scores (HAQ) those disease 2D). activity indexes integrate different parameters, we then analysed Weeach alsoparameter observed aindependently. We found correlation between sCD40L that absolute and counts RF titers of CD14+PLT+ (r = 0.544; p = 0.013). In were inversely correlated with inflammatory markers CRP (Figure 2A) and addition, we found that sCD40L levels between ACPA+ and ACPA- patients (839 (389.1– ESR (Figure 2B), but1794) CD14+PLT+ percentages did not correlate with CRP and ESR (r = − 0.4; pg/mL in ACPA+ vs. 332.9 (64.7–1551) pg/mL in ACPA-; p 0.08) were not signifi- p = 0.054 with CRP r = −0.33; p = 0.09 with ESR). mCD162 expression on monocytes correlated with anddifferent. cantly joint US scores (Figure 2C) and health assessment questionnaire scores (HAQ) (Figure 2D). Figure 2. Association of monocytes with bound platelets and CD162 expression on monocytes with Figure 2. Association of monocytes with bound platelets and CD162 expression on monocytes with baseline clinical and laboratory parameters in rheumatoid arthritis (RA) patients. (A) Absolute num- baseline clinical and laboratory parameters in rheumatoid arthritis (RA) patients. (A) Absolute ber of monocytes with bound platelets (CD14+PLT+) correlation with CRP and (B) ESR. (C) CD162 expression on monocytes correlation with joint ultrasound (US) scores and (D) health assessment questionnaire (HAQ). Statistical analysis was performed using Spearman’s correlation. We also observed a correlation between sCD40L and RF titers (r = 0.544; p = 0.013). In addition, we found that sCD40L levels between ACPA+ and ACPA- patients (839 (389.1–1794) pg/mL in ACPA+ vs. 332.9 (64.7–1551) pg/mL in ACPA-; p = 0.08) were not significantly different. 2.3. Relationship between CD14+PLT+ and mCD162 Expression on Monocytes with Clinical Outcome in RA Patients after TCZ Treatment We found that the percentage of CD14+PLT+ increased, while mCD162 expression on monocytes decreased 4 and 12 weeks after initiating TCZ treatment (Figure 3A,B). Int. J. Mol. Sci. 2022, 23, 5748 5 of 13 sCD62P Int. J. Mol. Sci. 2022, 23, x FOR also decreased at four weeks (Figure S3A), while sCD40L did not change PEER REVIEW 6 of 14 after TCZ treatment (data not shown). FigureFigure 3. Changes 3. Changes in inmonocytes monocytes with withboundbound platelets and CD162 platelets expression and CD162 after tocilizumab expression after tocilizumab treatment and association with clinical response in rheumatoid arthritis (RA) patients. (A) Per- treatment and association with clinical response in rheumatoid arthritis centage of monocytes with bound platelets (CD14+PLT+) and (B) CD162 expression on monocytes (RA) patients. (A) Per- centage of monocytes before (Baseline) and with four bound weeks (4 platelets w) and twelve (CD14+PLT+) andtocilizumab weeks (12 w) after (B) CD162(TCZ) expression treatment.on monocytes (C) Kaplan-Meier curve for EULAR response. Percentage of patients with low or high before (Baseline) and four weeks (4 w) and twelve weeks (12 w) after tocilizumab (TCZ) treatment. %CD14+PLT+ achieving 1 (moderate) or 2 (good) EULAR response (days). (D) Changes in the (C) Kaplan-Meier percentage of curve CD14+PLT+for EULAR response. and (E) CD162 Percentage expression after 4 w ofofpatients with low TCZ treatment or high %CD14+PLT+ in rheumatoid arthritis achieving patients with no 1 (moderate) or remission 2 (good)or EULAR remission response at 12 w (left). Black circles (days). (D) represent Changes RAin patients the percentage of before TCZ treatment, and grey circles RA patients after TCZ treatment. Statistical analysis was CD14+PLT+ and (E) CD162 expression after 4 w of TCZ treatment in performed using Wilcoxon test for comparisons between baseline and 4 w or 12 w after TCZ rheumatoid arthritis patients with notreatment remission and or remission log-rank at 12 w Mantel–Cox test(left). for theBlack circles analysis represent of EULAR RA response patients during before follow-up. * p TCZ < treatment, 0.05, ** p < 0.01, *** p < 0.001. and grey circles RA patients after TCZ treatment. Statistical analysis was performed using Wilcoxon test for comparisons between We also analysed thebaseline associationandbetween 4 w or changes 12 w after TCZ in the treatment percentage ofand log-rank Mantel–Cox CD14+PLT+, test formCD162 the analysis of EULAR expression responseand on monocytes during sCD62Pfollow-up. * p < 0.05, at four weeks ** p < 0.01, with changes *** p < 0.001. in clinical and laboratory parameters at four weeks. We found that changes in CD14+PLT+ corre- latedsegregation The inversely with ofchanges in DAS28, patients ESR,with into those and with lowswollen or highjoint count (SJC) or percentages (Figure absolute counts 4A,C). We did not find any association between changes in mCD162 of CD14+PLT+ was established by calculating cut-offs with a confidence intervalexpression on of 95% monocytes or plasmatic sCD62P levels and changes in clinical and laboratory parameters for HD values, as explained (data not shown). in the Material and Methods section. A higher percentage of patients with lower percentages or absolute counts of CD14+PLT+ at baseline achieved a good (2) or moderate (1) EULAR response compared to patients with higher percentages or absolute counts of CD14+PLT+ (%CD14+PLT+ low: HR = 2.68 when compared to %CD14+PLT+ high) (Figure 3C and data not shown). A decreased percentage of CD14+PLT+ at four weeks after TCZ was found in patients who did not achieve remission at 12 weeks. In contrast, an increased percentage of CD14+PLT+ at four weeks after TCZ was found in patients who achieved remission at 12 weeks (Figure 3D). A decreasing mCD162 expression four weeks after TCZ was found only in patients who did not achieve remission (Figure 3E). Regarding sCD62P, there were no significant changes in patients who did not achieve remission, while decreasing changes were found in patients who achieved remission (Figure S3B). Interestingly, in the group of patients who achieved remission, changes in CD14+PLT+ were inversely correlated with changes in sCD62P (r = −0.582, p = 0.04). No correlation was found in the non-responder group (data not shown). Int. J. Mol. Sci. 2022, 23, 5748 6 of 13 We also analysed the association between changes in the percentage of CD14+PLT+, Int. J. Mol. Sci. 2022, 23, x FOR PEER REVIEW 7 of 14 mCD162 expression on monocytes and sCD62P at four weeks with changes in clinical and laboratory parameters at four weeks. We found that changes in CD14+PLT+ correlated inversely with changes in DAS28, ESR, and with swollen joint count (SJC) (Figure 4A,C). We Int. J. Mol. Sci. 2022, 23, x FORdid PEERnot find any association between changes in mCD162 expression on monocytes REVIEW 7 of 14 or plas- matic sCD62P levels and changes in clinical and laboratory parameters (data not shown). Figure 4. Association of changes in monocytes with bound platelets and CD162 expression on monocytes after 4 weeks of tocilizumab treatment with clinical and laboratory parameters. (A) Changes in percentage of monocytes with bound platelets (CD14+PLT+) correlation with changes in DAS28, (B) ESR and (C) swollen joint count (SJC). Statistical analysis was performed using the Spearman’s correlation. 4. Association FigureFigure of changes 4. Association in monocytes of changes in monocyteswith withbound bound platelets andCD162 platelets and CD162expression expression on on mono- 2.4. IL-6 and TCZ Effects over PLT Binding to Monocytes, mCD162 Expression on Monocytes cytes after 4 weeks of tocilizumab treatment with clinical and laboratory parameters. monocytes after 4 weeks of tocilizumab treatment with clinical and laboratory parameters. (A) (A) Changes in and PLT Activation Changes in percentage of monocytes with bound platelets (CD14+PLT+) correlation with percentage of monocytes with bound platelets (CD14+PLT+) correlation with changes in DAS28, (B) ESR changes To analyse in DAS28, (B) ESR the and mechanism of TCZ (C) swollen joint on(SJC). count PLT Statistical binding toanalysis monocytes, we stimulated was performed using the and (C) swollen whole Spearman’s joint blood count from HD(SJC). correlation. Statistical with LPS, rhIL-6,analysis was We or/and TCZ. performed found anusing the Spearman’s increased percentage correlation. of CD14+PLT+ when we stimulated whole blood with LPS or with rhIL-6, without syn- 2.4. IL-6 2.4. andeffects ergic IL-6 TCZwhen Effects and TCZ bothover Effects PLTwere stimuli over PLT Binding to(data present Binding Monocytes, mCD162 not shown). to Monocytes, mCD162When Expression we added Expression onafter onTCZ Monocytes and Monocytes PLT Activation LPS or rhIL-6 stimulus, the percentage of CD14+PLT+ was not different from LPS or and PLT Activation TorhIL-6 analyseconditions To analyse (data not shown). the mechanism the mechanism of of TCZHowever, TCZononPLT when we added PLTbinding bindingto TCZ before LPS tomonocytes, monocytes, we weorstimulated rhIL-6 stimulated whole stimulus, the percentage of CD14+PLT+ was significantly lower than LPS or rhIL-6 con- bloodwhole frombloodHDfrom withHD LPS, with rhIL-6, or/and TCZ. We found an increased percentage of ditions (Figure 5A, Table S1).LPS, rhIL-6, or/and TCZ. We found an increased percentage CD14+PLT+ We when of CD14+PLT+ we that when also found stimulated we mCD162 stimulated whole wholeblood expression blood with with LPS on monocytesLPS orwith or within decreased rhIL-6, rhIL-6, without without the presence ofsyn-synergic ergic effectsLPS effects when(Figure when both both stimuli 5B,stimuli Table were S1). Nowere present present differences in(data (data not shown). not mCD162 shown). When expression on we When added TCZ we added monocytes wereafter TCZ after LPS LPS or or between found rhIL-6 rhIL-6 stimulus, LPS alone stimulus, the theand percentage LPS when we percentage ofofhad CD14+PLT+ added TCZ CD14+PLT+ was not different beforehand was not(Figure from 5B, LPS different Ta- fromor LPS or rhIL-6S1)conditions (data not shown). However, shown).when we added TCZ TCZ before before LPS or de- rhIL-6 rhIL-6ble or after LPS conditions (datastimulus (data not not shown). However, The addition when we ofadded TCZrhIL-6 before LPS or rhIL- stimulus, the percentage creased mCD162 of CD14+PLT+ expression on monocytes was significantly (Figure 5B, Tablelower than S1). We didLPS or rhIL-6 not find differ-con- 6 stimulus, ences in ditions the percentage the percentage (Figure 5A, Table of S1).of activated free PLTs (CD41a+CD62P+) between the different or rhIL-6 CD14+PLT+ was significantly lower than LPS conditions (Figure stimulations 5A, (data not We also found Table S1). expression on monocytes decreased in the presence of shown). that mCD162 LPS (Figure 5B, Table S1). No differences in mCD162 expression on monocytes were found between LPS alone and LPS when we had added TCZ beforehand (Figure 5B, Ta- ble S1) or after LPS stimulus (data not shown). The addition of TCZ before rhIL-6 de- creased mCD162 expression on monocytes (Figure 5B, Table S1). We did not find differ- ences in the percentage of activated free PLTs (CD41a+CD62P+) between the different stimulations (data not shown). 5. Effect Figure Figure of IL-6 5. Effect and of IL-6 andtocilizumab tocilizumab onon platelet platelet binding binding and expression and CD162 CD162 expression on monocytes. on monocytes. Peripheral Peripheral whole whole bloodblood fromhealthy from healthy donors donors waswascultured without cultured stimulusstimulus without or stimulated or with stimulated with lipopolysaccharide (LPS), recombinant human IL-6 or tocilizumab (TCZ). When TCZ stimulus is lipopolysaccharide (LPS), recombinant human IL-6 or tocilizumab (TCZ). When TCZ stimulus is prior to LPS or IL-6 treatment is indicated as pre-. Differences between each condition and basal prior to LPS or IL-6 treatment is indicated as pre-. Differences between each condition and basal condition were calculated from 7 independent experiments. X axis represents percentage of increase or decrease from basal condition. Data are expressed as mean with standard error of mean. Stars insideFigure the bars indicate 5. Effect comparison of IL-6 withonbasal and tocilizumab condition. platelet (A)CD162 binding and Percentage of monocytes expression with bound on monocytes. platelets or (B) whole Peripheral CD162blood expression on monocytes from healthy donors waswas evaluated. cultured without Statistical stimulus oranalysis was stimulated performed with usinglipopolysaccharide Wilcoxon test. * p(LPS), recombinant < 0.05, human IL-6 or tocilizumab (TCZ). When TCZ stimulus is ** p < 0.01. prior to LPS or IL-6 treatment is indicated as pre-. Differences between each condition and basal Int. J. Mol. Sci. 2022, 23, 5748 7 of 13 We also found that mCD162 expression on monocytes decreased in the presence of LPS (Figure 5B, Table S1). No differences in mCD162 expression on monocytes were found between LPS alone and LPS when we had added TCZ beforehand (Figure 5B, Table S1) or after LPS stimulus (data not shown). The addition of TCZ before rhIL-6 decreased mCD162 expression on monocytes (Figure 5B, Table S1). We did not find differences in the percentage of activated free PLTs (CD41a+CD62P+) between the different stimulations (data not shown). 3. Discussion Our present findings suggest that, as a result of exposure to an inflammatory envi- ronment, more monocytes bound to PLTs but had decreased mCD162 expression. After the administration of TCZ, the percentage of CD14+PLT+ increased in RA patients who achieved remission after 12 weeks post-TCZ treatment. Interestingly, patients with a lower percentage of CD14+PLT+ before TCZ treatment achieved EULAR-defined response to a greater extent. Our work showed that RA patients had increased percentages and absolute counts of CD14+PLT+ cells and increased levels of soluble markers of PLT activation. We also found an inverse association between CD14+PLT+ and the inflammatory parameters CRP and ESR. These findings suggest that an inflammatory environment contributes to the binding of PLT to monocytes with a regulatory effect. Whereas, like other authors, we have not found any statistical association between PLT activation markers and disease activity [50], Lee et al. found a correlation between sCD62P levels and disease activity, CRP and ESR [51]. This discrepancy between studies may be due to differences in cohort characteristics. In fact, our cohort had a longer disease duration and higher disease activity but a lower percentage of RF- and ACPA-positive patients, as well as a lower percentage of patients in MTX treatment. We found a significantly lower expression of mCD162 on monocytes in RA patients, despite them having more CD14+PLT+ cells. This finding suggests that, in addition to mCD162, other molecules on the surface of monocytes participate in PLT–monocyte in- teraction during inflammation. This fact is not exclusive to RA, and we previously found an increased percentage of CD14+PLT+ and a lower expression of mCD162 on monocytes in other chronic inflammatory diseases, such as ulcerative colitis and SLE [29,32]. Some authors have described a mCD162 downregulation on monocytes in the presence of endo- toxin in HIV patients or after leukocyte activation [4,30,31,33,52]. There are three possible explanations for these findings. First, the lower expression of mCD162 may have been due to a misdetection by the binding of PLTs that hide this molecule. This is unlikely because RA monocytes with or without bound PLTs had a similar mCD162 expression in our ex- periments. Second, the lower expression of mCD162 may be related to the activation state of monocytes [4,52] and the shedding of this molecule from their cell surface. Particularly, IL-8 and GM-CSF induce ADAM10 to shed CD162 on the cell-surface [33,53]. However, despite the lower mCD162 expression on RA monocytes, we did not find differences in sCD162 levels, the product of mCD162 shedding, between RA and HD. Since we measured sCD162 on plasma samples and supernatants from whole-blood cultures, we cannot rule out sCD162 consumption and/or degradation in our experimental system. Third, the lower expression of mCD162 on RA monocytes may have been due to reduced transcription. Therefore, we cannot discard the possibility that mCD162 downregulation is secondary to activation and shedding or to reduced transcription in RA monocytes. The follow-up analysis of patients during our study showed that TCZ treatment exacerbated the differences between RA patients and HD, increasing the percentage of CD14+PLT+ and decreasing plasmatic sCD62P and the mCD162 expression on monocytes. Again, these results support the idea that the binding of PLT to monocytes is a regulatory mechanism. In line with this, we found that, in patients in remission at 12 weeks, the percentage of CD14+PLT+ increased and correlated with the decrease in sCD62P. This result seems to indicate that the binding of non-activated PLT to monocytes has beneficial Int. J. Mol. Sci. 2022, 23, 5748 8 of 13 consequences. However, it is possible that other DMARDs have different effects on mono- cyte and PLT interaction due to characteristic target cells in the therapy mechanisms of action. In line with this, Veale et al. reported a decrease in sCD62P after sulphasalazine. Although Bunescu et al. reported a decrease in monocyte activation after MTX but not after TNF blocker treatment, they did not find changes in monocyte–PLT aggregates after either of the treatments [19,50]. Some studies have revealed different consequences after PLT–monocyte interaction. Thus, the specific binding of PLTs from RA patients to HD monocytes through CD147 increases TNFα and IL-6 expression on monocytes [26]. In contrast, we have reported that the co-cultures of PLT from HD with synovial fluid cells from RA patients reduced inflammatory cytokine production and increased IL-10 production [54]. Thus, it is likely that PLT can exert different effects, depending on the environment in which they have been “educated” and the cellular target for their binding. We speculate that the binding of PLT to monocytes after TCZ, with less inflammation and activation of PLT and monocytes, may have different consequences in comparison to binding before TCZ treatment. When we stimulated in vitro whole blood from HD with LPS or rhIL-6, the percentage of CD14+PLT+ increased. These results would explain ex vivo observations in patients before TCZ therapy. It is possible that IL-6 modifies the activation status of monocytes. Another possibility is that IL-6 induces CD16 expression on monocytes and PLTs would therefore preferentially bind to CD16+ monocytes [34,37,51]. This is unlikely in our settings because we did not find differences in CD16 expression on monocytes cultured for 28 h with or without rhIL-6 (data not shown). A detailed kinetics with different times of culture will clarify this point. IL-6 might also activate PLTs in RA patients. However, we observed a higher binding of PLT to monocytes without higher PLT activation, suggesting that both monocyte activation and phenotype changes are crucial in the interaction with PLT. Further conclusions regarding the involvement of IL-6 in PLT–monocyte interaction were obtained when we blocked in vitro IL-6 signaling with TCZ. The addition of TCZ to the culture prior to, but not after, LPS or rhIL-6 prevented the increase in CD14+PLT+, suggesting that TCZ has a preventive effect. Nevertheless, we are aware of the difficulty of extrapolating the results of in vitro 28 h experiments to in vivo RA patients after 12 weeks of follow-up [4,16,55]. Additionally, differences in both monocytes and PLTs between RA patients and HD could also explain this difference, as monocytes and PLTs from RA patients are activated [4,16,55]. Further experiments with RA samples will reveal if the in vitro effects of TCZ over monocyte–PLT conjugates are equivalent in RA and HD. Interestingly, in RA patients, despite the high levels of CD14+PLT+ in half of the patients before TCZ, these increased further after initiating TCZ treatment. A possible explanation for the high levels at baseline in this group is that CD14+PLT+ is an induced regulatory mechanism to try to control inflammation. In line with our findings with TCZ treatment, mCD162 expression decreased when we added TCZ prior to rhIL-6 culture. However, we did not reproduce this effect when TCZ was added before or after LPS culture. These results support the idea that other molecules, in addition to IL-6, regulate mCD162 in an inflammatory context. Hashizume et al. showed that the stimulation of whole blood with IL-6 for two hours downregulated mCD162 expression [53]. We did not reproduce the results of Hashizume et al. when stimulating with rhIL-6. Differences in culture timing could explain this discrepancy, as our experimental system has a 28-h duration. To avoid the interference of multiple mechanisms involved in different phases of RA, we have only included patients with a long disease duration. This restriction may have biased the cohort to a group with more systemic inflammation and refractoriness to previous DMARDs. This restriction has also limited the number of patients. To generalize our findings, a larger cohort, including early-onset patients and patients without previous treatments, should be included. Furthermore, our observations should be completed with additional experiments that could explain the mechanism by which TCZ treatment increases CD14+PLT+, as well as the functional consequences of this rise. Nevertheless, in addition Int. J. Mol. Sci. 2022, 23, 5748 9 of 13 to the description of a new mechanism in the pathology of RA, the role of monocyte–PLT conjugates in the modulation of inflammation, our results suggest that the percentage of CD14+PLT+ can be applied as a biomarker of early remission to TCZ treatment. 4. Materials and Methods 4.1. Patients Peripheral blood samples from 35 RA patients and 15 HD were collected in BD vacutainer CPT tubes containing heparin (BD, Franklin Lakes, NJ). RA diagnosis was based on the American College of Rheumatology diagnostic criteria for RA [56]. All RA patients were refractory to standard treatment with DMARDs, including MTX. TCZ treatment was started following European and Spanish guidelines [57,58]. The demographic, clinical and laboratory data of RA patients enrolled in this study are shown in Table 1. This study was approved by the ethics committee of Hospital de la Santa Creu i Sant Pau. In accordance with the Declaration of Helsinki, written consent was obtained from all patients. Table 1. Demographic, clinical, and laboratory characteristic data of study patients. RA (n = 35) HD (n = 15) p Sex, % (n), women 80.6 (28) 80 (12) 0.96 Age in years, mean ± sd 53.82 ± 10.72 52.48 ± 8.1 0.72 Years of evolution, median (IQR) 12.5 (7–17) Corticoids, % (n) 67.7 (21) Monotherapy, % (n) 29 (9) MTX, % (n) 42 (13) Previous biological therapies, % (n) 64.5 (20) HAQ, mean ± sd 1.4 ± 0.67 DAS28, mean ± sd 5.7 ± 1.15 ESR (mm/h), mean ± sd 51.4 ± 32 CRP (mg/L), median (IQR) 24.6 (4–68) ACPA+, % (n) 61.3 (19) ACPA (UI/mL), median (IQR) 140.7 (54–318) RF+, % (n) 71 (22) RF (UI/mL), median (IQR) 161 (32.5–333.5) Patients were treated with 8 mg/kg of TCZ every four weeks. Response to TCZ therapy was evaluated at 12 weeks. Blood samples and clinical data were collected one hour prior to the first infusion (baseline) and one hour prior to the respective infusions at weeks four (4 w) and 12 (12 w) after initiating treatment. Laboratory analyses for all visits included a hemogram, erythrocyte sedimentation rate (ESR), C-reactive protein (CRP), rheumatoid factor (RF), and anti-cyclic citrullinated peptide antibodies (ACPAs). Joint ultrasound (US) scores for all visits were evaluated using power doppler and grey scale [59]. Clinical data were collected at each visit. The disease activity score (DAS) 28 was calculated with ESR values. Simplified Disease Activity Index (SDAI), and Clinical Disease Activity Index (CDAI) were also calculated [60]. EULAR response criteria were used to classify patients as good, moderate or non-responders after 12 weeks of TCZ [61]. A good response patient (EULAR 2) had a ∆DAS28 > −1.2 and DAS28 final ≤ 3.2. A non-response patient (EULAR 0) had a ∆DAS28 ≤ −0.6 or −1.2 ≤ ∆DAS28 ≤ −0.6 and DAS28 final > 5.1. The rest of the patients (EULAR 1) had a moderate response. 4.2. Peripheral Blood Staining and Flow Cytometry Analysis Peripheral blood cells (100 µL) were incubated for fifteen minutes at room temperature in the dark with anti-CD14-APC (clone 18D11), -CD41a-FITC (clone HIP8) (ImmunoTools, Friesoythe, Germany), and -CD162-PE (clone KPL-1) (Biolegend, San Diego, CA, USA) mAb. Red blood cells were lysed and white cells were fixed using BD FACS lysing solution (BD Biosciences, San Jose, CA, USA) to be analysed by flow cytometry. Int. J. Mol. Sci. 2022, 23, 5748 10 of 13 Monocyte–platelet conjugates were identified as CD14+CD41a+ cells (CD14+PLT+). Surface expression of CD162 was analysed on gated CD14+ monocytes, as well as on CD14+PLT- and CD14+PLT+. Samples were acquired with the MACSQuant Analyzer 10 flow cytometer (Miltenyi Biotec GmBh, BergischGaldbach, Germany). We calculated the percentage of positive cells and mean fluorescence intensity (MFI) of each individual marker using MACSQuant Analysis Software (v.2.13.1). 4.3. Plasma Analysis of RF, ACPAs, and Cytokines RF titration was determined by nephelometry (Beckman ICS II, Beckman Coulter). ACPA titre was determined by EliA test using UniCAP (Phadia Laboratory Systems, Up- pasala, Sweden). Plasma was tested for IL-17, IL-22, VEGF, sCD40L (Peprotech, London, UK), soluble P-Selectin (sCD62P), soluble CD162 (sCD162) (R&D Systems, Minneapolis, MN), Lipopolysaccharide binding protein (LBP) (Hycult Biotech, Uden, The Netherlands), IL-10, IL-6 and soluble IL-6 receptor (sIL-6R) (ImmunoTools), using specific ELISAs accord- ing to the manufacturer’s instructions. All cytokines were quantified with standard curves provided by manufacturer. The detection limits were: 31.2 pg/mL for IL-17, 15.6 pg/mL for IL-22, 31.2 pg/mL for VEGF, 15.6 pg/mL for sCD40L, 125 pg/mL for sCD62P, 31.3 pg/mL for sCD162, 4.4 ng/mL for LBP, 16 pg/mL for IL-10, 3 pg/mL for IL-6 and 230 pg/mL for sIL-6R. 4.4. Peripheral Blood Cultures Heparinized blood was diluted with RPMI 1640 medium (supplemented with 2 mM glutamine, 100 U/mL penicillin, and 100 mg/mL streptomycin (BioWhittaker, Verviers, Belgium)) at a proportion of 1:1.4. Blood cells were stimulated with ultrapure TLR4 agonist lipopolysaccharide (LPS) (Invivogen, San Diego, CA, USA) at 10 pg/mL for four hours before adding 50 ng/mL of recombinant human IL-6 (rhIL-6) (ImmunoTools) for 12 h. TCZ was added at 100 µg/mL 30 min prior to LPS (conditions pre) or after rhIL-6 (conditions post). Culture conditions were then as follows: TCZ (pre) + LPS + rhIL-6 and LPS + rhIL-6 + TCZ (post). As control conditions, we included: (a) absence of stimulation, (b) LPS alone, (c) rhIL-6 alone, (d) TCZ alone, (e) rhIL-6+ TCZ (post), (f) TCZ (pre) + LPS, (g) TCZ (pre) + rhIL-6, (h) LPS + TCZ (post), (i) LPS + rhIL-6. After culture, supernatants were collected and kept at −20 ◦ C until used. A total of 100 µL of cultured whole blood from each condition was stained with anti-CD14-PECy7 clone M5E2 (BD Biosciences), anti-CD16 AF647 clone 3G8, anti-CD162 (BioLegend), anti-CD41a-FITC, anti-CD62P-APC clone HI62P (Immunotools). Red blood cells were lysed and white cells were fixed using BD FACS lysing solution (BD Biosciences) to be analysed by flow cytometry. Activated PLTs were identified as CD41a+CD62P+. Samples were acquired with the MACS-Quant Analyzer 10 flow cytometer (Miltenyi Biotec). We calculated the percentage of positive cells and the expression (MFI) of each individual marker using MACSQuant Analysis Software. 4.5. Statistical Analysis Statistical analyses were performed using Graph Pad Prism 7 software (v.7.00). The Kolmogorov–Smirnov test was applied to test the normal distribution of the data. Variables with a normal distribution were reported as mean ± standard deviation (SD) and the variables with a non-normal distribution were reported as median (interquartile range) (IQR). The t-test and paired t-test were used for the comparison of independent and related variables with normal distribution, respectively. The Mann–Whitney test and Wilcoxon test were used for the comparison of independent and related variables with non-normal distribution, respectively. The long-rank Mantel–Cox test was used to analyse differences in EULAR response during the follow-up period. p values lower than 0.05 were considered significant. Int. J. Mol. Sci. 2022, 23, 5748 11 of 13 Supplementary Materials: The following supporting information can be downloaded at: https:// www.mdpi.com/article/10.3390/ijms23105748/s1. Author Contributions: Conceptualization, C.Z. and S.V.; methodology, A.M., C.Z. and M.À.O.; soft- ware, A.M. and C.Z.; validation, A.M., C.Z. and M.À.O.; formal analysis, A.M. and C.Z.; investigation, A.M.; resources, S.V., C.D.-T., J.J.d.A., D.R., P.E., P.M. and H.C.; data curation, A.M.; writing—original draft preparation, A.M.; writing—review and editing, C.Z. and S.V.; visualization, C.Z. and S.V.; supervision, C.Z. and S.V.; project administration, S.V.; funding acquisition, S.V. All authors have read and agreed to the published version of the manuscript. Funding: This study is supported by Instituto de Salud Carlos III and Fondos FEDER (PI17/00072 and PI20/00184). Institutional Review Board Statement: The study was conducted in accordance with the Declaration of Helsinki, and approved by the Ethics Committee of Hospital de la Santa Creu I Sant Pau (protocol codes IIBSP-PLT-2018-1 and IIBSP-PLA-2021-36 approved 03/14/2018 and 06/15/2021 respectively). Informed Consent Statement: Informed consent was obtained from all subjects involved in the study. Data Availability Statement: Not applicable. Acknowledgments: Silvia Vidal was supported by Fondo de Investigaciones Sanitarias (FIS) and is a participant in the Program for the Stabilization of Investigators of the Direcció d’Estrategia i Coordinació del Departament de Salut, Generalitat de Catalunya. This work has been performed within the Ph.D. Immunology Program of Universitat Autònoma de Barcelona (UAB). Conflicts of Interest: The authors declare no conflict of interest. References 1. Muravyev, Y.V. Extra-articular manifestations of rheumatoid arthritis. Nauchno-Prakt. Revmatol. 2018, 56, 356–362. [CrossRef] 2. McInnes, I.B.; Schett, G. Mechanism of Disease the Pathogenesis of Rheumatoid Arthritis. N. Engl. J. Med. 2011, 365, 2205–2219. [CrossRef] [PubMed] 3. Gordon, S.; Taylor, P.R. Monocyte and macrophage heterogeneity. Nat. Rev. Immunol. 2005, 5, 953–964. [CrossRef] [PubMed] 4. Lioté, F.; Boval-Boizard, B.; Weill, D.; Kuntz, D.; Wautier, J.L. Blood monocyte activation in rheumatoid arthritis: Increased monocyte adhesiveness, integrin expression, and cytokine release. Clin. Exp. Immunol. 1996, 106, 13–19. [CrossRef] 5. Swirski, F.K.; Nahrendorf, M.; Etzrodt, M.; Wildgruber, M.; Panizzi, P.; Figueiredo, J.; Kohler, R.H.; Chudnovskiy, A.; Waterman, P.; Aikawa, E.; et al. Identification Monocytes Inflammatory of Splenic Reservoir and Their Deployment Sites. Science 2009, 325, 612–616. [CrossRef] 6. Boutet, M.A.; Courties, G. Novel insights into macrophage diversity in rheumatoid arthritis synovium. Autoimmun. Rev. 2021, 20, 102758. [CrossRef] 7. Nakashima, T.; Takayanagi, H. The dynamic interplay between osteoclasts and the immune system. Arch. Biochem. Biophys. 2008, 473, 166–171. [CrossRef] 8. Semple, J.W.; Freedman, J. Platelets and innate immunity. Cell. Mol. Life Sci. 2010, 67, 499–511. [CrossRef] 9. Smith, T.L.; Weyrich, A.S. Platelets as central mediators of systemic inflammatory responses. Thromb. Res. 2011, 127, 4. [CrossRef] 10. Tunjungputri, R.N.; van der Ven, A.J. Differential effects of platelets and platelet inhibition by ticagrelor on TLR2- and TLR4- mediated inflammatory responses. Thromb. Haemost. 2015, 113, 1035–1045. 11. Zamora, C.; Cantó, E. Functional consequences of platelet binding to T lymphocytes in inflammation. J. Leukoc. Biol. 2013, 94, 521–529. [CrossRef] [PubMed] 12. Cognasse, F.; Nguyen, K.A. The inflammatory role of platelets via their TLRs and Siglec receptors. Front. Immunol. 2015, 6, 83. [CrossRef] [PubMed] 13. Habets, K.L.L.; Trouw, L.A. Anti-citrullinated protein antibodies contribute to platelet activation in rheumatoid arthritis. Arthritis Res. Ther. 2015, 17, 209. [CrossRef] [PubMed] 14. Estevez, B.; Du, X. New concepts and mechanisms of platelet activation signaling. Physiology 2017, 32, 162–177. [CrossRef] 15. Cafaro, G.; Bartoloni, E. Platelets: A potential target for rheumatoid arthritis treatment? Expert Rev. Clin. Immunol. 2019, 15, 1–3. [CrossRef] 16. Yun, S.H.; Sim, E.H. Platelet activation: The mechanisms and potential biomarkers. Biomed Res. Int. 2016, 2016, 10–15. [CrossRef] 17. Pamuk, G.E.; Vural, Ö. Increased platelet activation markers in rheumatoid arthritis: Are they related with subclinical atheroscle- rosis? Platelets 2008, 19, 146–154. [CrossRef] 18. Boilard, E.; Blanco, P. Platelets: Active players in the pathogenesis of arthritis and SLE. Nat. Rev. Rheumatol. 2012, 8, 534–542. [CrossRef] 19. Bunescu, A.; Seideman, P. Enhanced Fc γ Receptor I, α M ß 2 Integrin Receptor Expression by Monocytes and Neutrophils in Rheumatoid Arthritis: Interaction with Platelets. J. Rheumatol. 2004, 31, 2345–2355. Int. J. Mol. Sci. 2022, 23, 5748 12 of 13 20. Stephen, J.; Emerson, B. The uncoupling of monocyte-platelet interactions from the induction of proinflammatory signaling in monocytes. J. Immunol. 2013, 191, 5677–5683. [CrossRef] 21. da Costa Martins, P.A.; van Gils, J.M. Platelet binding to monocytes increases the adhesive properties of monocytes by up- regulating the expression and functionality of beta1 and beta2 integrins. J. Leukoc. Biol. 2006, 79, 499–507. [CrossRef] [PubMed] 22. Simon, D.I.; Chen, Z. Platelet glycoprotein Ibα is a counterreceptor for the leukocyte integrin Mac-1 (CD11b/CD18). J. Exp. Med. 2000, 192, 193–204. [CrossRef] [PubMed] 23. Lindmark, E.; Tenno, T. Role of platelet P-selectin and CD40 ligand in the induction of monocytic tissue factor expression. Arterioscler. Thromb. Vasc. Biol. 2000, 20, 2322–2328. [CrossRef] [PubMed] 24. Silverstein, R.L.; Asch, A.S. Glycoprotein IV mediates thrombospondin-dependent platelet-monocyte and platelet-U937 cell adhesion. J. Clin. Investig. 1989, 84, 546–552. [CrossRef] [PubMed] 25. Schulz, C.; Von Brühl, M.L. EMMPRIN (CD147/basigin) mediates platelet-monocyte interactions in vivo and augments monocyte recruitment to the vascular wall. J. Thromb. Haemost. 2011, 9, 1007–1019. [CrossRef] [PubMed] 26. Rong, M.; Wang, C. Platelets induce a proinflammatory phenotype in monocytes via the CD147 pathway in rheumatoid arthritis. Arthritis Res. Ther. 2014, 16, 478. [CrossRef] [PubMed] 27. van Gils, J.M.; Zwaginga, J.J. Molecular and functional interactions among monocytes, platelets, and endothelial cells and their relevance for cardiovascular diseases. J. Leukoc. Biol. 2008, 85, 195–204. [CrossRef] 28. Larsen, E.; Celi, A. PADGEM protein: A receptor that mediates the interaction of activated platelets with neutrophils and monocytes. Cell 1989, 59, 305–312. [CrossRef] 29. Mariscal, A.; Zamora, C. Phenotypic and Functional Consequences of PLT Binding to Monocytes and Its Association with Clinical Features in SLE. Int. J. Mol. Sci. 2021, 22, 4719. [CrossRef] 30. Liang, H.; Duan, Z. Higher levels of circulating monocyte-platelet aggregates are correlated with viremia and increased sCD163 levels in HIV-1 infection. Cell. Mol. Immunol. 2015, 12, 435–443. [CrossRef] 31. Marsik, C.; Mayr, F. Endotoxin down-modulates P-selectin glycoprotein ligand-1 (PSGL-1, CD162) on neutrophils in humans. J. Clin. Immunol. 2004, 24, 62–65. [CrossRef] [PubMed] 32. Zamora, C.; Canto, E. Inverse Association Between Circulating Monocyte-Platelet Complexes and Inflammation in Ulcerative Colitis Patients. Inflamm. Bowel Dis. 2018, 24, 818–828. [CrossRef] [PubMed] 33. Davenpeck, K.L.; Brummet, M.E. Activation of Human Leukocytes Reduces Surface P-Selectin Glycoprotein Ligand-1 (PSGL-1, CD162) and Adhesion to P-Selectin In Vitro. J. Immunol. 2000, 165, 2764–2772. [CrossRef] [PubMed] 34. Passacquale, G.; Vamadevan, P. Monocyte-platelet interaction induces a pro-inflammatory phenotype in circulating monocytes. PLoS ONE 2011, 6, e25595. [CrossRef] [PubMed] 35. Lang, D.; Dohle, F. Down-Regulation of Monocyte Apoptosis by Phagocytosis of Platelets: Involvement of a Caspase-9, Caspase-3, and Heat Shock Protein 70-Dependent Pathway. J. Immunol. 2002, 168, 6152–6158. [CrossRef] 36. Ammon, C.; Kreutz, M. Platelets induce monocyte differentiation in serum-free coculture. J. Leukoc. Biol. 1998, 63, 469–476. [CrossRef] 37. Gudbrandsdottir, S.; Hasselbalch, H.C. Activated platelets enhance IL-10 secretion and reduce TNF-α secretion by monocytes. J. Immunol. 2013, 191, 4059–4067. [CrossRef] 38. Schwartzkopff, F.; Petersen, F. CXC chemokine ligand 4 (CXCL4) down-regulates CC chemokine receptor expression on human monocytes. Innate Immun. 2012, 18, 124–139. [CrossRef] 39. Sarma, J.; Laan, C.A. Increased Platelet Binding to Circulating Monocytes in Acute Coronary Syndromes. Circulation 2002, 105, 2166–2171. [CrossRef] 40. Harding, S.A.; Sommerfield, A.J. Increased CD40 ligand and platelet—monocyte aggregates in patients with type 1 diabetes mellitus. Atherosclerosis 2004, 176, 321–325. [CrossRef] 41. Joseph, J.E.; Harrison, P. Increased circulating platelet-leucocyte complexes and platelet activation in patients with antiphospholipid syndrome, systemic lupus erythematosus and rheumatoid arthritis. Br. J. Haematol. 2001, 115, 451–459. [CrossRef] [PubMed] 42. Pamuk, G.E.; Turgut, B. Increased circulating platelet-leucocyte complexes in patients with primary Raynaud’s phenomenon and Raynaud’s phenomenon secondary to systemic sclerosis: A comparative study. Blood Coagul. Fibrinolysis 2007, 18, 297–302. [CrossRef] 43. Pamuk, G.E.; Pamuk, Ö.N. Elevated platelet-monocyte complexes ın patıents wıth psorıatıc arthrıtıs. Platelets 2009, 20, 493–497. [CrossRef] [PubMed] 44. von Hundelshausen, P.; Weber, K.S.C. RANTES Deposition by Platelets Triggers Monocyte Arrest on Inflamed and Atherosclerotic Endothelium. Circulation 2001, 103, 1772–1777. [CrossRef] 45. Linke, B.; Schreiber, Y. Activated Platelets Induce an Anti-Inflammatory Response of Monocytes/Macrophages through Cross- Regulation of PGE2 and Cytokines. Mediat. Inflamm. 2017, 2017, 1463216. [CrossRef] [PubMed] 46. Kiely, P.; Walsh, D. Outcome in rheumatoid arthritis patients with continued conventional therapy for moderate disease activity- the early RA network (ERAN). Rheumatology 2011, 50, 926–931. [CrossRef] 47. Tono, T.; Aihara, S. Effects of anti-IL-6 receptor antibody on human monocytes. Mod. Rheumatol. 2015, 25, 79–84. [CrossRef] 48. Lee, J.S.; Oh, J.S. Tocilizumab-induced Thrombocytopenia in Patients with Rheumatoid Arthritis. J. Rheum. Dis. 2019, 26, 186. [CrossRef] 49. Gibiansky, L.; Frey, N. Linking interleukin-6 receptor blockade with tocilizumab and its hematological effects using a modeling approach. J. Pharmacokinet. Pharmacodyn. 2012, 39, 5–16. [CrossRef] Int. J. Mol. Sci. 2022, 23, 5748 13 of 13 50. Veale, D.J.; Maple, C. Soluble cell adhesion molecules—P-selectin and ICAM-1, and disease activity in patients receiving sulphasalazine for active rheumatoid arthritis. Scand. J. Rheumatol. 1998, 27, 296–299. 51. Lee, S.J.; Yoon, B.R. Activated Platelets Convert CD14+CD16- Into CD14+CD16+ Monocytes With Enhanced FcγR-Mediated Phagocytosis and Skewed M2 Polarization. Front. Immunol. 2021, 11, 3363. [CrossRef] [PubMed] 52. Torsteinsdóttir, I.; Arvidson, N.G. Monocyte activation in rheumatoid arthritis (RA): Increased integrin, Fcγ and complement receptor expression and the effect of glucocorticoids. Clin. Exp. Immunol. 1999, 115, 554–560. [CrossRef] [PubMed] 53. Hashizume, M.; Higuchi, Y. IL-6 plays an essential role in neutrophilia under inflammation. Cytokine 2011, 54, 92–99. [CrossRef] [PubMed] 54. Zamora, C.; Cantó, E. Binding of Platelets to Lymphocytes: A Potential Anti-Inflammatory Therapy in Rheumatoid Arthritis. J. Immunol. 2017, 198, 3099–3108. [CrossRef] [PubMed] 55. Cloutier, N.; Paré, A. Platelets can enhance vascular permeability. Blood 2012, 120, 1334–1343. [CrossRef] 56. Matsumoto, Y.; Uemura, M. Clinical usefulness of the 1987 revised criteria for rheumatoid arthritis by American Rheumatism Association. Nihon Naika Gakkai Zasshi 1988, 77, 742–743. [CrossRef] 57. Sanmartí, R.; García-Rodríguez, S. Actualización 2014 del Documento de Consenso de la Sociedad Española de Reumatología sobre el uso de terapias biológicas en la artritis reumatoide. Reumatol. Clin. 2015, 11, 279–294. [CrossRef] 58. Smolen, J.S.; Landewé, R. EULAR recommendations for the management of rheumatoid arthritis with synthetic and biological disease-modifying antirheumatic drugs: 2016 update. Ann. Rheum. Dis. 2017, 76, 960–977. [CrossRef] 59. Botar-Jid, C.; Bolboaca, S. Gray scale and power Doppler ultrasonography in evaluation of early rheumatoid arthritis. Med. Ultrason. 2010, 12, 300–305. 60. Futó, G.; Somogyi, A. Visualization of DAS28, SDAI, and CDAI: The magic carpets of rheumatoid arthritis. Clin. Rheumatol. 2014, 33, 623–629. [CrossRef] 61. Van Gestel, A.M.; Prevoo, M.L.L. Development and validation of the European League Against Rheumatism response criteria for rheumatoid arthritis. Arthritis Rheum. 1996, 39, 34–40. [CrossRef] [PubMed]