Chronic Immune thrombocytopenia (ITP) is an immune-mediated disorder characterized by thrombocytopenia and mucocutaneous bleeding. Its Pathophysiology is heterogeneous and complex (Stasi et al.) [1]. Glycoprotein had been considered the major etiology in pathogenesis. However cellular immune response has the characteristic hallmarks of breakdown in their tolerance mechanism (Cines et al.) [2]. In addition, Humoral immune response has a significant role in pathogenesis of ITP. It involves a complex interaction among antigen-presenting cells (APCs), T helper (Th) cells, regulatory T (Treg) cells and B cells (Wu et al.) [3].

As a major subset of effecter T lymphocytes, Follicular helper T cells help B cells (Linterman et al.) [4]. They are the key cell type required for the formation of germinal centers (GCs) and the generation of long-lived serological memory (Ma et al.) [5]. TFH, promote maturation B cell and antibody production (Spolski and Leonard, Linterman et al.) [6,7].

TFH cells expressed many effector molecules that are critical for their development and function, including chemokine receptor 5 (CXCR5), inducible co-stimulator (ICOS), programmed death-1 (PD-1), surface receptors of IL-21, IL-6, CD40, as well as two important transcription factors [B-cell lymphoma 6 (Bcl-6) and c-Maf]. So TFH are characterized by their surface phenotype (CD4+CXCR5+ICOS+PD-1+) and their cytokine profile (IL-21, IL-10, IL-17). The over-expression of ICOS is causing CXCR5+CD4+TFH cells over-production, and increased GC reactions with subsequent antibody production and autoimmune disease development [8,9].

Interleukin (IL)-21 is secreted by CD4+ T cells including TFH cells, Th17 cells and natural killer T cells [6]. The IL-21 enhances theproliferation and function of activatedCD4+ and CD8+ T cells [10]. The exposure of CD4+ T cells to IL-21 drives them to differentiate into a TFH cell subset with the expression of CXCR5, CCR7 and upregulation of Bcl-6 [11,12]. THF cells cause B cell proliferation, differentiation and antibody production is via the secretion of IL-21 [13,14]. IL-21 co-stimulation is directly acting on B cells and capable of promoting plasma cells differentiationfrom CD27+ memory B cells, and stimulating naïve blood B cells into IgG-secreting plasma cells in humans (Ettinger et al.) [15]. Moreover, IL-21 is critical for the formation of GCs and the development of TFH cells (Luthje et al.) [16], the absence of IL-21 signaling influences GCs proliferation, transition into memory B cells, and affinity maturation (Zotos D et al.) [17]. Thus, the effect of IL-21 on B cells may contribute to the development of autoimmune diseases.

The previous studies are reported the key role of TFH cells in regulating the humoral immune response that occurs with autoimmune diseases, infectious diseases, and tumors (Xu et al.) [18]. Recently, increased frequencies of circulating TFH cells were detected in many autoimmune diseases such as SLE [19,20], autoimmune thyroid disease (Zhu et al.) [21], rheumatoid arthritis (Liu et al.) [22] and Sjögren's syndrome. Via the secretion of IL-21, TFH can cause B cell proliferation, differentiation and antibody production, which exacerbate the progression of those diseases [18,23]. To date a little is known about role of TFH cells and their cytokine profiles namely IL21, IL-10 and IL-17 in ITP development and their relation to disease outcome. Therefore, our objective was to study the frequency of cTFH cells, together with their cytokine profile (IL-21, IL-10 and IL-17) to show whether they may influence the process of ITP or affecting disease outcome in chronic ITP patients.

Our study involved forty eight ITP patients. Age ranged from 7 to 46 years. Males were 28, while females were 20. The study was conducted from February 2015 to September 2017 in Menoufia and Al Azhar Universities hospitals. Patients were subjected to detailed history, physical examination and lab investigations. Peripheral blood smear and bone marrow aspiration were the most significant investigations. We had used Chinese association of hematology criteria for diagnosis (15). We had also included patients who had bleeding manifestations, e.g. easy bruising, purpura, or mucosal bleeding along with thrombocytopenia, provided that they had no other explainable illnesses.

The exclusion criteria

All causes of thrombocytopenia must be excluded. The major considerations include pseudo-thrombocytopenia, DIC, viral infections, TTP, and drug-induced thrombocytopenia. Other disease in the differential include pregnancy associated thrombocytopenia, autoimmune disease, systemic lupus erythematosus, acquired immunodeficiency syndrome, other infections, liver disease, hematologic and other malignancies, transfusion reaction.

Inclusion criteria

Thrombocytopenia cases with a normal white blood count and erythrocyte indices with normal morphology, coagulation profile are within normal limits. Normal to increased megakaryocytes in BM with normal morphology was detected and platelet-antibody may be positive. There were no significant illnesses over the preceding 3 months prior to enrollment in study. At time of result analysis, none of patients was on steroid therapy. Twenty five apparently healthy volunteers were our controls. They were 12 males, and 13 females. Age ranged from ten to thirty eight years. They had no history of virus infection in the preceding two weeks and they have normal platelet counts. None of controls or their families had a history of hematologic or autoimmune diseases, nor received blood products therapy.

Treatment protocol

Dexamethazone and then prednisolone were our treatment options. We never transfused platelets or blood.

Clinical examination

Including detailed history about disease onset, bleeding manifestations, history of recent infection or received drugs. Thorough clinical examination with stress on bruising, purpuric eruption on the skin, mucous membrane or gum bleeding, menorrhagia or hematuria, splenic, liver and lymph node examinations.

Laboratory investigations

Blood sampling: Venous blood was collected in 2 sterile vacutainer EDTA tubes for immediate complete blood count, and one for flow cytometric analysis. The remaining sample was put in sterile vacutainer plain tube and centrifuged at 2000 g for 3 min; serum samples were collected and stored at −80°C for subsequent routine serology and cytokine analysis. Bone marrow (BM) aspiration staining and examination revealed normal to increased megakaryocyte numbers reflecting normal to increased BM thrombopoiesis.

Cytokine measurement

Quantitative determination of IL-21, IL-10 and IL-17 was performed using quantitative sandwich enzyme-linked immunosorbent assay (Quantikine; R&D systems, Minneapolis, Minnesota, USA), according to the manufacturer’s instructions. Briefly, the standards and samples were pipetted and incubated into the wells coated by and biotinylated anti-human cytokine measured (IL-21, IL-10 or Il-17). After washing to remove unbound biotinylated antibodies, HPR-conjugated streptavidin is pipetted into the wells followed by second washing. TMB subsequently was added to the wells, resulting in color development, which is proportional to the amount of bound cytokine measured. Stop solution was added. The color intensity was measured by reading optical absorbance at 450 nm using a microplate reader. The digital data of absorbance values were derived from the standard curve of each cytokine concentrations; the final concentration was recorded and put by pg/ml.

Estimation of reticulated platelets (RP)

To obtain platelet-rich plasma (PRP), EDTA blood samples were centrifuged at 120 g for 10 minutes. Five microlitre of PRP were fixed in 1% para-formaldehyde for at least 30 minutes at room temperature. It was then washed for two times and re-suspended at 1 ml of phosphate-buffered saline (PBS) (pH 7.2). Fifty microlitres then were mixed with 10 μl of phycoerythrin-tagged monoclonal antibody against CD41 (Immunotech-Marseille, France).

It was then incubated in dark at room temperature for 10 minutes. Then 1 ml from working solution (thiazole orange) was added to the suspension, followed by incubation at room temperature in the dark for 1 hour. The samples were analyzed on an FACS Calibur (Beckton Dickinson) and analyzed by Cell Quest software (BD Biosciences), by analysis by reading the electronic gate made around platelets according to light scatter electronic. The negative isotopic control was used to exclude all cell autofluoresence or instrument noise.

Flow cytometric analysis of TFH cell frequency and phenotypic markers

Frequency and phenotype of TFH-like cells of patients and controls were assessed by flowcytometry. We used Fluorochrome-conjugated mAbs from (R&D Systems- USA) to examine CD3, CD4, CD8, CXCR5, ICOS and PD-1 expression. The fluorescein isothiocyanate (FITC)-ICOS, phycoerythrin (PE)-CD4, and APC-CXCR5, PE-Cy7- PD-1 monoclonal antibodies were used according to the manufacturers’ protocols. The whole fresh EDTA blood was stained and incubated at normal room temperature for half an hour with surface phenotypical markers antibodies. After being washed with PBS, the cells were subjected to flow cytometry analysis using a FACS Calibur (Beckton Dickinson) and analyzed by Cell Quest software (BD Biosciences). The cells were gated on the forward scatter of living cells and then centered on CD4+T cells. Subsequently, theCD4+ CXCR5+ICOS+Tfh cells were determined by flow cytometric analysis.

Statistical Methods

After data collection, statistical analysis was done using SPSS computer program version 21. Data were expressed as mean ± SD and differences between 2 groups were analyzed by student (t) test for parametric variable distribution or Mann-Whitney test for nonparametric. Pearson's correlation coefficient was used to test the relationship between various variables. We considered P value significant when it was p<0.05.

Patients’ clinical data are listed in Table 1, the study included 48 patients with chronic ITP, 28 of them were males and 20 were females. Among them 46 (95.8%) had petechiae, 43 (89.6%) had easy bruising, 37 (77.1%) had epistaxis, 42 (87.5%) had gingival bleeding, 21 (43.8%) had hematuria, 11 (22.9%) had menorrhagia and 2 (4.2%) had gastrointestinal bleeding (GIT), while and none of the patients have retinal hemorrhage, intracranial hemorrhage, hepatomegaly, splenomegaly, lymph-adenopathy or do splenectomy. After BM, the megakaryocytes were increased in about 36 (75%) of patients and the remaining 12 (25%) have normal megakaryocytes in BM.

Table 1: Clinical and demographic data of all ITP patients.

The patient age and gender were matched to healthy control group; also, the hemoglobin concentration and white blood cell count are not show significant differences between the ITP patients and control group. Platelets counts were significantly decreased (p<0.001) in ITP compared to control group, while, mean platelet volume (MPV) and reticulated platelets (RP) were significantly increased (p<0.05 and p<0.001 respectively). The cytokine levels were significantly increased in ITP patients compared to control group, (p<0.01 for IL-10, p<0.001 for IL-17 and p<0.001 for IL-21) (Table 2 and Figure 1).

Table 2: Comparison between CBC and cytokine levels in ITP patients and control group.

Figure 1: Levels of cytokines in ITP and healthy control groups.

The frequency of circulating TFH cells (CXCR5+CD4+) is significantly higher in chronic ITP patients compared to healthy controls (p<0.001) (Figure 2).

Figure 2: Flowcytometry chart of TFH (CXCR5+CD4+) in ITP.

The phenotype of TFH cells was assessed, the expression of CXCR5+CD4+/ICOS+ (Figure 3) and CXCR5+CD4+/ PD-1+ TFH (Figure 4) cells were significantly increased in ITP patients compared to control group (p<0.01, p<0.01 respectively).

Figure 3: Flowcytometry chart of ICOS+ cells in ITP.

Figure 4: Flowcytometry chart of PD-1+ cells in ITP.

However, the percentages of CD3+, CD4+ and CD8+ cells were not significantly different between ITP patients and control group (Table 3).

Table 3: Comparison between the expression of T cells subsets, TFH cells and its phenotype in ITP patients and control group.

Our study revealed that a positive correlation was detected between the frequency of CXCR5+CD4+TFH cells and each of clinical manifestations, presence of platelet antibody, cytokine levels (IL-10, IL17 and IL-21). In addition, a strong positive correlation was observed between the frequency of the CXCR5+CD4+Tfh cells and each of ICOS+ and PD-1+ on CXCR5+ CD4+TFH cells in chronic ITP patients, but no large difference was found in healthy control group (Table 4).

Table 4: Correlation between the TFH (CXCR5+CD4+) cell percentage and some parameters in ITP patients and control group.

After therapy, the serum levels of IL-10, IL17 and IL-21 are significantly decreased in ITP responding patients (p<0.05, p<0.01, p<0.001 respectively). However, the frequencies of CXCR5+CD4 TFH cells, CXCR5+CD4+/ICOS+ and CXCR5+CD4+/PD-1+ are also significantly decreased (p<0.01 for each), the percentage of CD3+ T lymphocyte, CD4+, CD8+ and the ratio of CD4+/CD8+ T lymphocyte not so differed (Table 5).

Table 5: Comparison between cytokine levels, expression of TFH cells and its phenotypes in ITP patients before and after therapy.

The exact cascading factor initiating ITP is still not well known. It is unclear whether immune defects play roles in the disease or not. Circulating TFH cells are a subset of T cells which help B cells to secrete autoantibody (Cooper et al.) [24]. still no much is known about role of TFH cells in patients with ITP. The present study was carried on 73 participates, 25 of them were a healthy control and 48 patients with chronic ITP to evaluate the frequency of cTFH cells, together with cytokine profile (IL21, IL10 and IL-16) and to be correlated with disease outcome.

In our study, clinical features such as petechiae, easy bruising, epistaxis, gingival bleeding, hematuria, menorrhagia and GIT in ITP patients had direct relationship with degree of thrombocytopenia. That was accordant with previous reports (Stasi et al.) [25,26]. The megakaryocytes were increased in the BM in about 75% of patients and were normal count in 25% of patients. WBCs or Hb concentration are not show significant differences between the ITP patients and control group. The platelet counts were significantly decreased in ITP patients, while mean platelet volume (MPV) and reticulated platelets (RP) were significantly increased. This may be due to the fact that even if the number of circulating platelets is decreased by platelet destruction, it cannot be affected by platelet production. Same results were obtained by previous studies by Pons et al. [27] on the contrary Del Vecchio et al. [28] did not find a correlation between platelet count and reticulated platelets. However lack of standardized technique for determining reticulated platelets might explain the inconsistency of our results with those reported in literature.

In the current study, serum IL-10 was measured at the disease onset. Some authors had previously described high IL-10 in some chronic ITP (Mouzaki et al. Del Vecchio et al. [28,29]). Raised IL-10 levels have been also described in children with chronic ITP [30]. IL10 stimulates the production of immunoglobulins andthe elevated cytokines, IL-2, IL-4, IL-6, IL-7 and IL-10 are all critical for B cell survival and differentiation [18,30,31]. On the contrary, reduced levels of IL-10 have also been reported in patients with active disease, which was not the case in healthy controls or patients in remission [32]. This controversy may be due to our case selection criteria of chronic ITP.

The results of this study revealed increasing IL-17 level in ITP patients. Level was directly proportionate to TFH. Increased IL-17 cytokine was also reported in ITP patients in other studies [33,34]. This many denote a possible role for Th17 cells in ITP immunopathology. On the other hand some reports did not detect any difference [34,35]. Cytotoxic T cells may indirectly play a role in lysis of platelets and megakaryocytes in the bone marrow [19,36]. Interleukin-17 secreting T-cells are pro-inflammatory cells. It is of interest in ITP pathogenesis partly due to evidences implicating IL-17 in autoimmunity (Johnsen et al.) [37]. Immune thrombocytopenic patients have decrease in immune compartment regulation (Arroyo- Villa et al.) [38], in addition to an increase in the effector T cell arm of the immune response (Th1, Th17 and CD8 cells).

In our study, the serum IL-21 level was significantly increased and its level was positively correlated with the frequency of TFH cells in the ITP patents, however, there were no correlation between IL-21 levels and TFH cells in the healthy control, suggesting IL-21 was able to directly promote B cell activation and auto-antibody production. This was in agreement with the finding of Cooper and Bussel, Eto et al., and Xie et al. [23,39].

Antibody production is a key component in B cell differentiation and generation of protective humoral immune responses. It indicates that the responsibility of IL-21 for Ig production may be stronger in ITP patients. IL-21 is prone to inducing B cell activation, expansion and differentiation. It has been implicated in autoimmunity disease via the IL-21R pathway (Liu et al.) [22]. Increased IL-21 levels might play a predominant role in the generation and differentiation of TFH cells with ICOS high molecule compared to PD-1 molecule, and the productions of platelet antibodies in ITP patients. This suggests a role in the pathogenesis of ITP patients. This is in accordance with previous report (Wang et al.) [40,41]. Contrary, other study by Zhu et al. [42] reported elevated IL-21 expression on T cells, in untreated newly diagnosed ITP patients, although circulating IL-21 was unchanged.

In our study, TFH cells frequency of circulation was markedly increased in ITP patients compared to control group, irrespective to normal levels of the T cell subtypes (CD4+ and CD8+) and ratio. In contrast, other studies (Wang et al.) [43] revealed high Th1/Th2 (“helper” CD4-cells) ratios and high Tc1/Tc2 (“cytotoxic” CD8 cells) ratios in acute ITP patients. They added that, the Th1/Th2 ratio imbalance is inversely correlated with disease severity [42,44] i.e. the higher is Th1/Th2 ratio, the lower the platelet count would be there. Increasing evidence shows a role for Cytotoxic CD8+T cells in platelet destruction (Nishimoto and Kuwana) [45]. This discrepancy in the results may be due to our cases is of chronic ITP type. TFH cell migration to follicular areas of B cell in GC such as spleen requires CXCR5 expression. It is also needed for the formation of GC. In the absence of CXCR5 GC will be impaired https://youtu.be/ CJHmdEWuPQw?t=14 [20,46,47]. The up-regulation of CXCR5 enhances the probability of antigen-specific contact between TFH cells and B cells in draining lymphoid tissue (Fazilleau et al.) [48]. Our results revealed that the high percentages of TFH (CXCR5+CD4+) cells were positively correlated with presence of platelet antibody in those patients. This would suggest a role of TFH cells in the pathogenesis of ITP and assisting function of FFH cells for B cells to secret antibodies. This was in agreement with (Xie et al.) [40] and (Simpson et al.) [19], findings, who demonstrated a higher frequency of TFH cells in the chronic ITP patients than in healthy control group. The function of TFH cells is closely related to the expression of ICOS and PD-1. Thus, it was important to determine the expression of TFH phenotype to show if typically associated with TFH cells altering in ITP cases. Our study revealed that, expressions of CXCR5+CD4+/ICOS+ and CXCR5+CD4+/PD-1+ TFH cells were significantly increased in ITP patients compared to control group. In addition, to further analyze whether the increased TFH cells had also high expression of ICOS or PD-1 molecules, the correlation study revealed a strongly positive correlation between the frequency of the CXCR5+CD4+TFH cells and each of ICOS+ and PD-1+ on CXCR5+ CD4+TFH cells in chronic ITP patients, but not in control group. These results are in accordance of (Xie et al.) [40] The expression of ICOS and PD-1 was increased in TFH cells in chronic ITP patients compared to healthy control. They added that, the frequencies of circulating TFH cells with ICOS high and PD-1 high were highest in ITP with platelet antibody (+) patients, although there was not differences of these parameters in between the ITP platelet negative (-) patients and healthy controls. Nishimoto and Kuwana reported involvement of TFH cells in B cell recruitment and differentiation in ITP patient’s spleens [46]. Therefore, the autoimmune attack on platelets (and megakaryocytes) is amplified by multiple mechanisms [39,49]. The increased mRNA expressions of IL-21 by PCR analysis in patients with ITP were possible explanations for the increased number of circulating CXCR5+CD4+TFH cells with ICOS high and PD-1 high in these ITP patients, particularly in those ITP (+) patients. These findings are consistent with prior studies [21,47,50]. After therapy, the serum levels of IL-10, IL17 and IL-21 are significantly decreased in ITP responding patients. However, the frequencies of TFH cells, CXCR5+CD4+/ICOS+ and CXCR5+CD4+/PD-1+ are also significantly decreased, the percentage of CD3+ T lymphocyte, CD4+, CD8+ and the ratio of CD4+/CD8+ T lymphocyte not so differed. This result is in agreement with Feng et al. and Qian et al. [51]. Likewise, Li et al. [52] study, is partially agreed with our study, they reported that, after the treatment of immunosuppressive therapy, IL-17 were downregulated while IL-10 not so different among ITP patients before or after immunosuppressive therapy and healthy controls.

Finally we conclude that, the increasing frequencies of circulating TFH cells with associated phenotypes ICOS+ and PD-1+ expressions of TFH cell might play a critical role in the pathogenesis of ITP via associated markers and secreted cytokines. Steroid treatment in ITP may significantly reduce the frequencies of TFH cells and its cytokines which will reduce illness severity. Further studies are still needed to explore the exact pathogenesis of ITP. This will help identify disease features and predictors of treatment response, thus facilitating the decision-making process for ITP management in the future.