Heparin affects the induction of regulatory T cells independent of anti‐coagulant activity and suppresses allogeneic immune responses

This study focused on the effect of heparin on regulatory T cells (Tregs) on the allogeneic immune responses in vitro and in vivo. Heparin‐induced de novo Treg generation and maintained the survival and activation of Tregs, independent of its anti‐coagulant properties.

Summary

Heparin is a widely used anti‐coagulant that enhances anti‐thrombin (AT) activity. However, heparin also suppresses immune and inflammatory responses in various rodent models and clinical trials, respectively. The mechanism by which heparin suppresses immune responses is unclear. The effect of heparin on regulatory T cells (T_regs) in allogeneic immune responses was analysed using an acute graft‐versus‐host disease (aGVHD) mouse model and mixed lymphocyte reactions (MLRs). In‐vitro culture systems were utilized to study the effects of heparin on T_regs. Heparin administration reduced mortality rates and increased the proportion of T_regs in the early post‐transplantation period of aGVHD mice. In both murine and human MLRs, heparin increased T_regs and inhibited responder T cell proliferation. Heparin promoted functional CD4⁺CD25⁺forkhead box protein 3 (FoxP3)⁺ T_reg generation from naive CD4⁺ T cells, increased interleukin (IL)‐2 production and enhanced the activation of pre‐existing T_regs with IL‐2. Heparin‐induced T_reg increases were not associated with anti‐coagulant activity through AT, but required negatively charged sulphation of heparin. Importantly, N‐acetyl heparin, a chemically modified heparin without anti‐coagulant activity, induced T_regs and decreased mortality in aGVHD mice. Our results indicate that heparin contributes to T_reg‐mediated immunosuppression through IL‐2 production and suggest that heparin derivatives may be useful for immunopathological control by efficient T_reg induction.

Introduction

Heparin is a type of glycosaminoglycan and a highly sulphated heparan sulphate. It is a well‐known anti‐coagulant used in the treatment of thromboembolism and disseminated intravascular coagulation, as well as in the prevention of coagulation during artificial dialysis and extracorporeal circulation. In the bloodstream, heparin binds to the enzyme inhibitor anti‐thrombin (AT) through a pentasaccharide sequence in the heparin molecule [1]. Heparin enhances the anti‐coagulant activity of AT, which inhibits the activity of thrombin and coagulation factor Xa (FXa) and inhibits blood coagulation [2]. Heparin combines with blood coagulation‐related factors, such as heparin co‐factor II, platelet factor IV and protein C inhibitor. Additionally, heparin combines with growth factors, adhesion factors and metabolism‐related factors with high affinity [3]. Heparin is synthesized in connective tissue‐type mast cells and stored in secretion granules combined with histamine and serglycin or isolated polysaccharides as a complex [4]. Based on these properties, the physiological roles of heparin include modulation of immune response and inflammatory responses as well as anti‐coagulant activity.

Previous studies have shown that heparin can suppress immune responses in a rodent model of autoimmune encephalomyelitis [5], delayed hypersensitivity [6] adjuvant arthritis, skin allograft [7], heart allograft [8] and acute graft‐versus‐host disease (aGVHD) [9], as well as influence lymphocyte traffic [10]. Based on these studies, heparin is believed to inhibit the expression of heparanase, an enzyme responsible for catalysing site‐selective cleavage of heparan sulphate chains in T cells and improving lymphocyte traffic. However, recent studies using selective heparanase inhibitors have not confirmed the anti‐inflammatory activity of heparin. Endothelial cell heparanase has been found to be important for inflammatory responses [11]. Therefore, the importance of T cell‐based heparanase during inflammatory responses and the mechanism of heparin‐related immune regulation remain unknown. The anti‐inflammatory effects of heparin have been described in a large number of clinical trials [12]. For example, inhaled heparin may represent a useful add‐on therapy in coronary obstructive pulmonary disorder and asthma patient groups [13]. However, these clinical trials have focused upon the effect of heparin on inflammatory responses and not immunosuppression.

CD4⁺CD25⁺forkhead box protein 3 (FoxP3)⁺ regulatory T cells (T_regs) are essential for the maintenance of immune tolerance, as well as moderate inflammation elicited by pathogens and environmental factors [14, 15]. They also contribute to tolerance induction after solid organ transplantation [16, 17] and protect against aGVHD lethality in animal bone marrow transplantation (BMT) models [18, 19, 20]. Furthermore, the adoptive transfer of T_regs is therapeutically effective in aGVHD clinical trials [21]. These studies suggest that when the up‐regulation of T_regs is controlled in vivo, this leads to effective immunoregulatory therapy. We hypothesized that immunosuppressive and anti‐inflammatory effects of heparin are involved, at least in part, in immunosuppression by T_regs. In this study, we examined the role of T_regs in the immunosuppressive effects of heparin. Our results showed that heparin and its non‐anti‐coagulant derivative could promote the differentiation and maintenance of T_regs by increasing interleukin (IL)‐2 production and suppressing aGVHD alloimmune responses and mixed lymphocyte reactions (MLRs).

Materials and methods

Mice

C57BL/6 (H‐2^b) and BALB/c (H‐2^d) mice were purchased from Clea Japan, Inc. (Tokyo, Japan) or bred in our laboratory. B6.129^{Foxp3tm4(YFP/icre)Ayr}/J mice (RRID: MGI_3790499) were purchased from Jackson Laboratory (Bar Harbor, ME, USA). B6.Cg‐Foxp3^sf (scurfy) mice (RRID: MGI_1857034) were provided by Dr Sakaguchi, Osaka University (Osaka, Japan). Mice were between 8 and 12 weeks of age (21–26 days of age in scurfy mice) during the experiment. Mice were housed under specific pathogen‐free conditions with a 12‐h light/dark cycle. Mice were fed pellet food and water ad libitum at the Laboratory Animal Research Centre and handled according to the Guidelines for the Care and Use of Laboratory Animals Research Centre, Dokkyo Medical University (Tochigi, Japan). All efforts were made to minimize suffering of the animals. For intravenous and subcutaneous injections, mice were anaesthetized with isoflurane gas in the air. Mice were monitored daily during all experiments. Mice were euthanized by cervical dislocation under general anaesthesia or by exposure to CO₂. No mice died without euthanasia before meeting the humane endpoint criteria. The animal handling and care protocols used in this study were approved by the Dokkyo Medical University Animal Experiments Committee (animal experiment permit number: 0341) and were in accordance with Dokkyo Medical University’s Regulations for Animal Experiments and with Japanese Governmental Law no. 105.

Reagents

The reagents used in this study were as follows: fluorescein isothiocyanate (FITC)‐(RRID: AB_312690), phycoerythrin (PE)‐(RRID: AB_312692) and allophycocyanin (APC)‐(RRID: AB_312696) conjugated anti‐CD4 (GK1·5), cyanin 5 (Cy5)‐conjugated anti‐CD8a (53‐6·7, RRID: AB_312750), FITC‐(RRID: AB_312854), APC‐(RRID: AB_312860) and PE/Cy7 (RRID: AB_312864)‐conjugated anti‐CD25 (PC61), FITC‐(RRID: AB_313092) and PE (RRID: AB_313094)‐conjugated anti‐CD62L (MEL‐14), PE‐conjugated anti‐CD90.2 (53‐2·1, RRID: AB_10641145), FITC‐conjugated anti‐H‐2K^d (SF1‐1·1, RRID: AB_313741), purified anti‐CD28 (37.51, RRID: AB_11147170) and purified anti‐CD3ε (145‐2C11, RRID: AB_1877073) were purchased from BioLegend (San Diego, CA, USA). The PE‐(RRID: AB_465936) and APC‐(RRID: AB_469457)‐conjugated anti‐FoxP3 (FJK‐16s) and PE‐conjugated anti‐glycoprotein A repetitions predominant (GARP) (YGIC86, RRID: AB_1963598) were purchased from eBioscience (San Diego, CA, USA). PE‐conjugated anti‐CD25 (7D4) was purchased from Southern Biotech (Birmingham, AL, USA). PE‐conjugated anti‐ Mothers against decapentaplegic homologue 2 (Smad2/3) (O72‐670) was purchased from BD Pharmingen (San Jose, CA, USA). For the human T_reg assay, purified anti‐human CD3ε (OKT3 RRID: AB_11150592) and Alexa647‐conjugated anti‐human FoxP3 (259D, RRID: AB_492985) were purchased from BioLegend. FITC‐conjugated anti‐human CD4 (RPA‐T4, RRID: AB_395751) and PE‐conjugated anti‐human CD25 (M‐A251, RRID: AB_395826) were purchased from BD Pharmingen. A FoxP3/transcription factor staining buffer set (eBioscience) was used for intracellular and intranuclear staining. Carboxyfluorescein succinimidyl ester (CFSE) was purchased from Invitrogen (Carlsbad, CA, USA). PE annexin V was purchased from BD Pharmingen. For the cell cultures, RPMI‐1640 medium was purchased from Wako Pure Chemical Industries (Osaka, Japan). Fetal bovine serum (FBS) was purchased from Biological Industries (Kibbutz Beit‐Haemek, Israel). Sodium pyruvate solution and penicillin–streptomycin solution were purchased from Sigma‐Aldrich Co. (St Louis, MO, USA). L‐glutamine solution and minimum essential medium non‐essential amino acids solution were purchased from Invitrogen. Heparin (unfractionated, heparin sodium) and protamine sulphate were purchased from Mochida Pharmaceutical Co. (Tokyo, Japan). Dalteparin (low‐molecular weight heparin (LMWH), trade name Fragmin) was purchased from Kissei Pharmaceutical Co. (Nagano, Japan). Fondaparinux (trade name Arixtra), a synthetic pentasaccharide FXa inhibitor, was purchased from GlaxoSmithKline plc (Middlesex, UK). Argatroban, a small molecule direct thrombin inhibitor, was purchased from Sawai Pharmaceutical Co. (Osaka, Japan). N‐desulphated heparin was purchased from Iduron (Manchester, UK). N‐acetyl heparin sodium salt was purchased from Sigma‐Aldrich Co.

Mouse model of acute graft‐versus‐host disease

The aGVHD mouse model has been previously reported in detail [19, 22]. Briefly, C57BL/6 or BALB/c recipient mice were exposed to lethal total body irradiation (1100 or 800 cGy) 5 h before allo‐BMT. On day 0, recipient mice were intravenously infused with 5 × 10⁶ T cell‐depleted (TCD) allogeneic bone marrow (BM) cells alone or in combination with 1 × 10⁶ CD90.2⁺‐purified allogeneic splenic T cells. Mice were subcutaneously administered 2 U heparin, 10 U heparin or 80 μg N‐acetyl heparin in phosphate‐buffered saline (PBS, pH 7·5) per mouse from days 0 to 30, every other day. The survival of the mice was assessed as described by Cooke et al. [23]. On day 50 after BMT, peripheral blood mononuclear cell (PBMC) chimerism was analysed in the transplanted mice.

Cell preparation and culture

Spleen cells were isolated from mice and treated with haemolytic alkaline buffer (1 mL 0·17 M Tris‐HCl pH 7·65, 9 ml 0·16M NH₄Cl) then cultured with RPMI‐1640 medium supplemented with 5% (v/v) heat‐inactivated FBS, 2 mM L‐glutamine, 1 mM sodium pyruvate, 100 U/ml penicillin, 0·1 mg/ml streptomycin and 10 mM non‐essential amino acids. Cells were seeded at a concentration of 1 × 10⁶ cells/well in 96‐well flat‐bottomed cell culture plates (AGC Tecno Glass Co., Ltd, Shizuoka, Japan) with primary spleen cell cultures. For the isolation of T_regs, spleen cells were isolated from FoxP3‐yellow fluorescent protein (YFP) mice, and cells expressing B220 or CD8 were removed by panning. After removing adherent cells, YFP⁺ cells were sorted as FoxP3‐expressing cells by an automatic cell sorter (BD FACSAria™; BD Bioscience, Bedford, MA, USA). In purified T_reg cultures, cells were seeded at a concentration of 1–2 × 10⁵ cells/well in 96‐well cell round‐bottomed culture plates (AGC Tecno Glass Co., Ltd). For T_reg induction from naive T cells, CD4⁺CD25⁻CD62L⁺ cells were purified by depletion of cells expressing cell lineage markers B220 (RA3‐6B2; eBioscience, RRID: AB_466451), CD8b (BD Pharmingen, RRID: AB_394576), CD11b (M1/70; Biolegend, RRID: AB_312787), CD25 (PC61; Biolegend, RRID: AB_312853), CD49b (DX5; Biolegend, RRID: AB_313411), NK1.1 (PK136; Biolegend, RRID: AB_313391) and Ter‐119 (Biolegend, RRID: AB_313705; biotin‐conjugated antibodies and streptavidin microbeads). Subsequent positive selection of cells expressing CD62L (FITC‐conjugated antibody and anti‐FITC microbeads or PE‐conjugated antibody and anti‐PE microbeads) utilized a magnetic cell sorter. The CD4⁺ cells used constituted more than 90% of the sorted cells.

Murine MLRs

Responder cells from BALB/c (or C57BL/6) spleens were co‐cultured with stimulators, which were irradiated spleen cells from C57BL/6 (or BALB/c) mice. For the T_reg‐depleted MLRs, CD25⁺ cell‐depleted spleen cells were used as responder cells. For CD25⁺ cell depletion, a magnetic cell sorter (autoMACS™; Miltenyi Biotec, Gladbach, Germany) was used. CFSE‐labelled responder cells were used for the responder cell proliferation assay. Responder cells (1 × 10⁶) were seeded with 4 × 10⁵ stimulator cells in a 96‐well flat‐bottomed cell culture plate.

Human MLRs and T_reg induction

PBMCs from healthy volunteers were obtained from Dokkyo Medical University through approved protocol number 30001 by the Dokkyo Medical University Bioethics Committee. Signed informed consent was obtained from all donors. Human PBMCs were co‐cultured with irradiated PBMCs from another donor as an alloreactive MLR. Responder cells were seeded (1 × 10⁵ cells) with 1 × 10⁵ irradiated stimulator cells in a 96‐well flat‐bottomed cell culture plate. Cells were cultured with RPMI‐1640 medium supplemented with 20% (v/v) heat‐inactivated human serum from the donors of responder cells, 2 mM L‐glutamine, 10 mM HEPES (pH 7·2; Invitrogen), 100 U/ml penicillin, 0·1 mg/ml streptomycin and 10 mM non‐essential amino acids, and analysed by fluorescence‐activated cells sorting (FACS) staining 7 days later. For human T_reg induction, naive CD4⁺ T cells from human PBMCs were purified by positive isolation using an anti‐human CD4 antibody by autoMACS and cultured with plate‐bound anti‐human CD3ε antibody (5 μg/ml) plus transforming growth factor (TGF)‐β (5 ng/ml). Cells were analysed by FACS staining 4 days later.

Flow cytometry

To prevent non‐specific monoclonal antibody staining, an anti‐CD16/32 antibody (clone 2.4G2) was used. Flow cytometry were performed using a FACSCalibur (BD Bioscience) and data were acquired using CellQuest software (BD Bioscience). Data analysis was performed using FlowJo software (FlowJo LLC, Ashland, OR, USA).

Enzyme‐linked immunosorbent assay (ELISA)

An ELISA kit [mouse interleukin (IL)‐2 ELISA MAX™ standard, no. 431001] for murine IL‐2 was purchased from BioLegend. ELISAs were performed according to the manufacturer’s instructions.

Immunoblotting

Purified splenic naive T cells (CD4⁺CD8⁻CD25⁻CD62L⁺ cell sorting by BD FACSAria™; BD Bioscience) from BALB/c mice were stimulated with an anti‐CD3 antibody plus TGF‐β (5 ng/ml), heparin (5 U/ml) and protamine sulphate (50 μg/ml) for 24 h. Cells were lysed in 10 mM HEPES (pH 7·9), 1·5 mM MgCl₂ (Wako), 10 mM KCl (Wako), 0·5 mM dithiothreitol (DTT; Sigma), containing 0.6 % (v/v) Nonidet P‐40 (Nacalai Tesque, Kyoto, Japan) with protease inhibitors (aprotinin; Sigma, pepstatin; Sigma, leupeptin; Roche Diagnostics K.K., Tokyo, Japan) and phenylmethylsulphonyl fluoride (PMSF; Invitrogen) and isolated with a cytoplasmic extract solution. Nuclear extracts were clarified by centrifugation at > 12 000 g for 1 min at 4ºC. Nuclear extracts were lysed in 20 mM HEPES (pH 7·9), 25% (v/v) glycerol (Wako), 0·42 mM NaCl (Wako), 1·5 mM MgCl₂ (Wako), 0·2 mM ethylenediamine tetraacetic acid (Wako), 0·5 mM PMSF and 0·5 mM DTT containing protease inhibitors, and clarified by centrifugation at > 12 000 g for 5 min at 4ºC. Nuclear proteins were quantified and separated by sodium dodecyl sulphate–polyacrylamide gel electrophoresis and transferred onto nitrocellulose membranes (0·45 μm pore size; Bio‐Rad, Hercules, CA, USA) for immunoblotting. The amount of intranuclear nuclear factor of activated T cells, cytoplasmic 1 [anti‐nuclear factor of activated T cells (NFATc1, 7A6; BioLegend, RRID: AB_10680239] and TATA‐binding protein (TBP) (anti‐TBP, 1TBP18; BioLegend, RRID: AB_2721754) was assessed by immunoblotting with horseradish peroxidase‐conjugated anti‐mouse immunoglobulin (Ig) G from sheep (GE Healthcare Bio‐sciences, Pittsburgh, PA, USA). The immunoblots were developed by electrochemiluminescence (Bio‐Rad, Hercules, CA, USA). Band intensities were measured by LuminoGraph I and analysed by CS Analyser 4 (ATTO Corporation, Tokyo, Japan).

Immunofluorescence microscopy

Murine T cells were stained for NFAT, as described previously [24]. Cells cultured with indicated stimuli were washed, placed at 1–2 × 10⁵ cells on a MAS‐coated glass slide (Matsunami Glass Ltd, Osaka, Japan) and allowed to adhere for 10 min at 20ºC. The cells were washed, fixed with cold 4% (w/v) paraformaldehyde (Nacalai Tesque) in PBS (pH 7·5), permeabilized with 0·1% (v/v) polyoxyethylene (10) octylphenyl ether (Wako) in PBS, and then blocked with 1% (v/v) normal goat serum (Sigma) in PBS containing 1% (w/v) BSA (Sigma). Cells were stained with a primary antibody against NFATc1, washed and incubated with FITC‐conjugated anti‐mouse IgG1 (Caltag Laboratories, Carlsbad, CA, USA). Finally, slides were washed with PBS and mounted in ProLong Diamond Antifade Mountant with 4′,6‐diamidino‐2‐phenylindole (DAPI; Thermo Fisher Scientific, Waltham, MA, USA) and analysed using an LSM 780 confocal microscope system (Carl Zeiss, Oberkochen, Germany) to acquire fluorescence images. Nuclear localization of NFATc1 as determined by the co‐localization of DAPI with FITC in images was analysed by ZEN microscope software (Carl Zeiss, Oberkochen, Germany).

Statistical analysis

Statistical analysis was performed using GraphPad Prism software version 6.0 for Mac (GraphPad Software, San Diego, CA, USA). Statistical significance was taken as P < 0·05.

Results

Heparin reduces the mortality rate and increases the proportion of splenic T_regs in aGVHD mice

Low‐dose heparin has been shown to inhibit aGVHD in a hemi‐allo BMT mouse model, attenuate the severity of aGVHD, and reduce the mortality rate without abrogating the graft‐versus‐leukaemia effect induced by the allograft or impairing marrow engraftment [9]. We hypothesized that this heparin‐induced amelioration of aGVHD was mediated, at least in part, by T_regs. To test our hypothesis, we first examined the effect of heparin on the mortality rate and CD4⁺CD25⁺FoxP3⁺ T_reg count in the spleen of allo‐BMT mice. Heparin 2 U/mouse is an almost equivalent dose to that of the previous study [9]. Depending on the body weight, 10 U of heparin/mouse is the maximum clinical preventive dose for pulmonary emboli and deep venous thrombosis. Heparin reduced the mortality rate and the effect was more pronounced in the high‐dose group (10 U/mouse; Fig. 1a). Moreover, modest subcutaneous haemorrhages were observed in some C57BL/6 recipient mice compared to the low‐dose group (2 U/mouse). One week after the BMT, heparin administration significantly increased the proportion of CD25⁺FoxP3⁺ T_regs in the CD4⁺ spleen cells (P = 0·0254 in the 2 U/mouse group; P = 0·0379 in the 10 U/mouse group; Fig. 1b). These results are in agreement with those of the previous study, indicating that the improvement of aGVHD mice was due to small increases in the percentages of T_regs [25]. These results suggest that the effects of heparin on T_regs may ameliorate GVHD in mice. The total amount of T_regs and donor T_regs tended to increase upon heparin administration (Supporting information, Fig. S1a,b). Conversely, in the experiments involving the exchange of recipient and donor mice, the low‐dose group was effective in improving GVHD in BALB/c recipient mice (Fig. 1c). However, heparin administration significantly increased the proportion of CD25⁺FoxP3⁺ donor T_regs in the CD4⁺ spleen cells in a dose‐dependent manner (P = 0·0441 in 2 U/mouse group; P = 0·0054 in 10 U/mouse group; Fig. 1d). The contradiction between the increase in T_reg proportion and mortality of aGVHD mice was attributed to the anti‐coagulant activity of heparin. Considerable haemorrhages and haematoma were observed subcutaneously in the high‐dose group of BALB/c recipient mice (Supporting information, Fig. S2). In the low‐dose group, only mild haemorrhages were observed. This contradiction may have caused the lower platelet count and fibrinogen levels in BALB/c mice compared to those of the C57BL/6 mice [26]. It may thus be possible that haemorrhages increase the mortality rate of the BALB/c mice high‐dose group. To confirm the effects of heparin on T_regs, anti‐inflammation and anti‐coagulation in the early post‐transplantation period of aGVHD mice models, donor T cells from T_reg‐deficient‐Foxp3^sf (scurfy) mice (C57BL/6 background) were co‐transferred with BM cells from wild‐type mice to the recipient mice. Heparin administration did not protect against aGVHD in recipient mice transferred with the Foxp3^sf T cells (Fig. 1e). This result demonstrated that heparin’s anti‐inflammatory activity, associated with an anti‐coagulant effect, did not protect aGVHD mice. Donor‐derived T_regs may be crucial for the improvement of acute GVHD upon heparin administration. Taken together, these results suggested that heparin increases the risk of haemorrhages but improves aGVHD by increasing the proportion of T_regs.

Fig. 1.

Heparin protects against acute graft‐versus‐host disease (aGVHD) and increases the proportion of peripheral regulatory T cells (T_regs). Recipient mice received lethal total body irradiation (TBI) (a,b, 1100 cGly to C57BL/6 recipients; c–e, 800 cGly to BALB/c recipients) followed by 5 × 10⁶ allogeneic T cell depleted‐bone marrow cells (TCD‐BMs; a–c, BALB/c; d, C57BL/6) alone or with 1 × 10⁶ allogeneic spleen T cells (a,b, BALB/c; c–e, C57BL/6, B6. forkhead box protein 3 (FoxP3)‐yellow fluorescent protein (YFP) or B6.Foxp3^sf) on day 0. Heparin was subcutaneously administered to mice from days 0 to 30, every other day. (a) C57BL/6 recipient mice‐transplanted BALB/c splenic T cells. Survival curves for the different groups. **P = 0·0066, by the log‐rank (Mantel–Cox) test, between phosphate‐buffered saline (PBS) and heparin 10 U/mouse [T cell‐depleted bone marrow (TCD‐BM) control; n = 3, PBS; n = 17, 2 U; n = 18, 10 U; n = 17]. (b) Mean ± standard error of the mean (s.e.m.) percentages of CD4⁺CD25⁺FoxP3⁺ T_regs among splenic CD4⁺ T cells 1 week after allogeneic bone marrow transplant (BMT) in C57BL/6 recipient mice (n = 7–8). *P < 0·05, by unpaired Student’s t‐test. (c) BALB/c recipient mice transferred C57BL/6 spleen T cells. Survival curves for the different groups. *P = 0·0117, by log‐rank (Mantel–Cox) test, between PBS and 2 U/mice (TCD‐BM control; n = 7, PBS; n = 10, 2 U; n = 10, 10 U; n = 10). (d) BALB/c recipient mice transferred B6.FoxP3‐YFP spleen T cells. Mean ± s.e.m. percentages of CD4⁺CD25⁺FoxP3⁺ donor T_regs among splenic CD4⁺ T cells 1 week after allogeneic BMT in BALB/c recipient mice (n = 6). *P < 0·05, **P < 0·01, by unpaired Student's t‐test. (e) BALB/c recipient mice transplanted Foxp3^sf spleen T cells. Survival curves for the different groups. P = 0·9518, by the log‐rank (Mantel–Cox) test, between PBS and 2 U/mouse (TCD‐BM control; n = 7, PBS; n = 12, 2 U; n = 12, 10 U; n = 12).

Heparin induces T_regs and suppresses alloimmune responses in MLR

Next, we investigated the ability of heparin to induce T_regs and suppress the alloimmune response in vitro. To this end, we analysed the allogeneic MLRs in the presence of heparin. We found that heparin significantly increased the proportion of CD4⁺CD25⁺FoxP3⁺ T_regs in MLRs (C57BL/6 responder cells; Fig. 2a). This effect was abolished by protamine sulphate, an antidote of heparin. Similar results were obtained from the MLRs with a reciprocal combination of stimulator and responder cells (P = 0·0068 in BALB/c responder cells; P = 0·0092 in C57BL/6 responder cells; Fig. 2b). Taken together, the above data demonstrated that heparin increases T_regs in alloimmune responses both in vitro and in vivo. To evaluate whether heparin can suppress the alloimmunoreactions of effector T cells, spleen cells labelled with CFSE were used as responder cells in the MLRs. We showed that heparin suppressed the proliferation of both CFSE‐labelled CD8⁺ and CD4⁺ responder T cells and decreased their cell number (Fig. 2c,d and Supporting information, Fig. S3). As the CFSE levels in the presence of heparin in CD8⁺ cells are higher than those of control, heparin may also suppress the spontaneous CD8⁺ cell division in the absence of stimulator cells (Fig. 2c). However, heparin did not increase the cell number of T_regs or decrease the cell number of non‐T_regs in the MLR (Fig. 2e). To remove the effect of pre‐existing T_regs in responder cells, CD25⁺ T_reg‐depleted responder cells were used in the MLR. Heparin decreased the cell numbers of CD8⁺ and CD4⁺ T cells. It also significantly increased the proportion (P = 0·0006) and cell numbers (P = 0·0188) of T_regs, which were induced by heparin, in the MLRs with T_reg‐depleted responder cells (Fig. 2f). These results suggest that heparin suppresses effector T cells in alloimmune responses by inducing functional T_regs. However, these results indicated that heparin may decrease the number of pre‐existing T_regs. To demonstrate the clinical relevance of our findings, we examined T_reg induction by heparin with human PBMCs isolated from healthy volunteers. Even in the allogenic MLRs of human PBMCs, heparin significantly increased the proportion of CD4⁺CD25⁺FoxP3⁺ T_regs (P = 0·0375) and increased the ratio index of T_regs (P = 0·0081; Fig. 3a, Supporting information, Table S1). Heparin enhanced the induction of CD25⁺FoxP3⁺ cells after the stimulation of human CD4⁺ T cells and PBMCs with anti‐CD3ε antibody in the presence of TGF‐β, and its effect was inhibited by the heparin antidote, protamine sulphate (Fig. 3b,c). These results suggested the possibility that heparin may induce T_regs and suppress the immune responses in both mouse and human peripheral tissues.

Fig. 2.

Heparin suppresses immune reactions by the induction of regulatory T cells (T_regs) in allogeneic mixed lymphocyte reactions (MLRs). Spleen cells were harvested from BALB/c (or C57BL/6) mice and co‐cultured with irradiated‐spleen cells from C57BL/6 (or BALB/c) mice as an alloreactive MLR. (a) Five days after culturing with C57BL/6 responder cells, cells were harvested and expression levels of CD25 and forkhead box protein 3 (FoxP3) were analysed. Representative data showing CD25⁺FoxP3⁺ cells among CD4⁺ cells at the indicated heparin and protamine concentrations. (b) Individual animal and group mean ± standard error of the mean (s.e.m.) percentages of CD25⁺FoxP3⁺ T_regs among CD4⁺ cells (left: BALB/c responder cells, right: C57BL/6 responder cells; n = 4–6). (c) Carboxyfluorescein succinimidyl ester (CFSE)‐labelled BALB/c spleen cells were used as responder cells in MLRs with heparin. Representative plots of CFSE levels in CD8⁺ or CD4⁺ T cells 5 days after MLR culture at the indicated concentrations of heparin. (d) Individual group mean ± s.e.m. number of CD8⁺ and CD4⁺ T cells in MLRs with BALB/c responder cells (n = 7). (e) Individual group mean ± s.e.m. number of CD4⁺CD25⁺FoxP3⁺ T_regs and CD4⁺ non‐T_regs in MLRs with BALB/c responder cells (n = 6). (f) Individual group mean ± s.e.m. number of CD8⁺, CD4⁺ T cells and CD4⁺CD25⁺FoxP3⁺ T_regs, or percentages of CD25⁺FoxP3⁺ T_regs among CD4⁺ cells in MLRs with CD25⁺ T_reg‐depleted BALB/c responder cells (n = 6). *P < 0·05, **P < 0·01, ***P < 0·005, ****P < 0·0001, by paired Student’s t‐test.

Fig. 3.

Heparin enhances the induction of human regulatory T cells (T_regs) from peripheral blood mononuclear cells (PBMCs) in vitro. (a) Human PBMCs were collected from healthy volunteers and co‐cultured with irradiated PBMCs from another donor as an alloreactive mixed lymphocyte reaction (MLRs). Seven days after culturing, cells were harvested and expression levels of CD25 and forkhead box protein 3 (FoxP3) were analysed. Left: representative data showing CD25⁺FoxP3⁺ cells among CD4⁺ cells at the indicated heparin and protamine concentrations. Individual group mean ± standard error of the mean (s.e.m.) percentages of CD25⁺FoxP3⁺ T_regs among CD4⁺ cells (n = 12). Right: increasing ratio index of T_reg proportions summarised as bar graphs. T_reg percentages from the heparin 0 U/ml group were assigned a value of 1 (n = 12, Supporting information, Table S1). (b) CD4⁺ peripheral T cells or (c) PBMCs were cultured with plate‐coated anti‐CD3ε antibody (OKT‐3: 5 μg/ml) in the presence of transforming growth factor (TGF)‐β (5 ng/ml). Four days after culture, cells were harvested and expression levels of CD25 and FoxP3 were analysed. Representative data showing CD25⁺FoxP3⁺ cells among CD4⁺ cells at the indicated heparin and protamine concentrations. Individual group mean ± s.e.m. percentages of CD25⁺FoxP3⁺ T_regs among CD4⁺ cells (CD4⁺ T cells, n = 5; PBMCs, n = 5). *P < 0·05, **P < 0·01, by paired Student’s t‐test.

Heparin promotes induced regulatory T cell (iT_reg) differentiation by up‐regulation of IL‐2 production

T_reg differentiation from naive CD4⁺ T cells in the periphery is induced by antigen stimulation with an appropriate combination of cytokines, such as IL‐2 and TGF‐β. FoxP3⁺ cells generated in vitro in this way are called iT_regs. Heparin potentiated the induction of FoxP3⁺ cells after stimulation of purified CD4⁺CD25⁻CD62L⁺ naive T cells with anti‐CD3ε antibody and TGF‐β, and its effect was inhibited by the antidote protamine sulphate (Fig. 4a). However, heparin did not drastically enhance the induction of T_regs from naive T cells by additional stimulation of CD28 co‐stimulation and IL‐2 (Supporting information, Fig. S4). The data in Supporting information, Fig. S4 indicate an increase in the proportion of T_regs by heparin and protamine, although the total cell number was actually remarkably reduced. The addition of heparin‐induced iT_regs to MLRs resulted in the inhibition of responder T cell proliferation (Fig. 4b), thereby indicating that heparin induces functional iT_regs. TGF‐β receptor signalling is essential for FoxP3‐expressing T_regs. Phosphorylation of Smad3 is required for the TGF‐β mediated‐induction of FoxP3 in naive T cells. However, we found that heparin did not up‐regulate the level of phospho‐Smad2/3 in naive T cells (Supporting information, Fig. S5). IL‐2 is indispensable for the induction of FoxP3 expression and T_reg differentiation [27]. T_regs also highly express IL‐2Rα (CD25), but are unable to produce IL‐2. Therefore, T_reg induction is dependent upon IL‐2 produced by other T cells. More importantly, heparin significantly augmented IL‐2 production (P = 0·0015) from naive T cells stimulated with anti‐CD3ε antibody in the presence of TGF‐β (Fig. 4c). This result was further supported by the immunoblotting and immunofluorescence microscopy data. Heparin increased the NFATc1 in the nucleus during the induction of T_regs from naive T cells (P = 0·0158; Fig. 4d). Microscopic co‐localization analysis revealed that NFATc1 translocation from the cytosol to the nucleus was significantly increased by heparin (P = 0·0018; Fig. 4e). These results clearly indicate that heparin induces IL‐2 production from naive T cells and promotes iT_reg differentiation.

Fig. 4.

Heparin enhances the induction of induced regulatory T cells (iT_regs) from naive T cells by interleukin (IL)‐2 production. (a) Naive T cells were harvested from the spleen of BALB/c mice and cultured with plate‐coated anti‐CD3ε antibody in the presence of transforming growth factor (TGF)‐β. Three days after culture, the expression levels of CD25 and FoxP3 in harvested cells were analysed. Left: representative data; right: percentages of CD25⁺forkhead box protein 3 (FoxP3)⁺ cells at the indicated heparin and protamine concentrations. Individual group mean ± standard error of the mean (s.e.m.) percentages of CD25⁺FoxP3⁺ T_regs (n = 5). (b) Naive T cells were harvested from the spleens of B6.FoxP3‐yellow fluorescent protein (YFP) mice and cultured with plate‐coated anti‐CD3ε antibody in the presence of TGF‐β and heparin. Three days after culture, YFP‐FoxP3⁺‐induced T_regs were sorted from the culture. Next, inducible T_regs (iT_regs) were added into the mixed lymphocyte reactions (MLRs). Data are representative plots of carboxyfluorescein succinimidyl ester (CFSE) levels among CD8⁺ T cells 3 days after MLRs. (c) Culture supernatants of naive T cells from BALB/c mice under T_reg‐inducing conditions were harvested at day 2. Data are mean ± s.e.m. values of IL‐2 production by naive T cells under the indicated conditions (n = 3–6). (d) Naive T cells from BALB/c mice under T_reg‐inducing conditions were harvested on day 1. Nuclear extracts were subjected to Western blot analysis for NFATc1. Upper: representative data; lower: relative value analysed using spot intensity at indicated heparin and protamine concentrations. Individual group mean ± s.e.m. value of nuclear factor of activated T cells, cytoplasmic 1 (NFATc1) protein normalized to TATA‐binding protein (TBP) (n = 5). (e) Naive T cells from BALB/c mice under T_reg‐inducing conditions were harvested on day 1. Left: representative merged images of day 1 cells stained with anti‐NFATc1 (green). Nuclei were stained with 4′,6‐diamidino‐2‐phenylindole (DAPI) (blue). Right: number of co‐localized pixels of DAPI with fluorescein isothiocyanate (FITC) (NFATc1; ×20 objective lens) in day 1 culture. Individual and mean ± s.e.m. group data (n = 30). *P < 0·05, **P < 0·01, ***P < 0·001, by Student’s t‐test.

Heparin maintains the survival and activation of T_regs in co‐operation with IL‐2

To ascertain the effect of heparin on pre‐existing T_regs, which are natively present in peripheral lymphoid organs, such as naturally occurring T_regs and peripherally derived T_regs, purified CD4⁺FoxP3⁺ T_regs from the spleen were analysed after culturing in the presence or absence of heparin. The inhibitory activity of heparin‐treated T_regs and heparin‐untreated T_regs was similar (Fig. 5a). The expression level of FoxP3 and number of T_regs decreased in cultures lacking stimulation and cytokines. Interestingly, heparin prevented a decrease in FoxP3 expression but enhanced the decrease in T_reg numbers in the cultures (Fig. 5b,c). These results were expected based on the data regarding the number of T_regs in the allogeneic MLRs (Fig. 2e). However, the addition of IL‐2 to the cultures prevented heparin‐induced decreases in T_regs (Fig. 5d). Therefore, heparin alone maintains FoxP3 expression and the suppressive activity of pre‐existing T_regs but does not support their survival. In contrast, heparin maintains their survival and up‐regulates FoxP3 and CD25 expression in the presence of IL‐2 (Fig. 5e). This is also supported by the fact that heparin increases FoxP3^high GARP^high T_regs in the presence of IL‐2 (Fig. 5f). The expression of GARP, a T_reg activation marker, was up‐regulated in mouse T_regs following IL‐2 exposure in the absence of T cell receptor (TCR) signalling [28]. Taken together, these results indicate that heparin induces de‐novo iT_reg generation by IL‐2 production from naive T cells and that heparin maintains the survival and activation of previously differentiated T_regs in co‐operation with IL‐2.

Fig. 5.

Interleukin (IL)‐2‐dependent heparin‐induced forkhead box protein 3 (FoxP3) expression and activation of regulatory T cells (T_regs). CD4⁺FoxP3⁺ T_regs were harvested from the spleens of B6.FoxP3‐yellow fluorescent protein (YFP) mice and cultured in the presence of heparin alone or in combination with protamine. (a) One day after culture, T_regs were added into the mixed lymphocyte reaction (MLR). Representative plots of Carboxyfluorescein succinimidyl ester (CFSE) levels in CD8⁺ T cells 5 days after MLRs with T_regs. (b) Left: representative plots of FoxP3 expression in T_regs collected at the indicated times after culture. Right: mean ± standard error of the mean (s.e.m.) values of FoxP3 fluorescence intensity in cells (n = 3–4). (c) Ratio of T_reg numbers calculated at the indicated times after culture (n = 3–4). (d) Individual group mean ± s.e.m. ratio of T_reg numbers calculated 5 days after culture with heparin and IL‐2 (100 ng/ml; n = 3). (e) Left: representative plots of FoxP3 and CD25 expression in T_regs collected 5 days after culture with heparin and IL‐2. Right: mean ± s.e.m. values of geometric fluorescence intensity in cells cultured in the presence of heparin and IL‐2 at day 5 (n = 5). (f) Upper: representative profiles of FoxP3 and glycoprotein A repetitions predominant (GARP) expression levels in T_regs collected 5 days after culture with heparin and IL‐2. Lower: mean ± s.e.m. percentages of FoxP3^highGARP^high T_regs cultured in the presence of heparin and IL‐2 on day 5 (n = 5). *P < 0·05, **P < 0·01, ***P < 0·005, by paired Student’s t‐test.

Sulphation, rather than anti‐coagulant ability, of heparin is important for T_reg induction

Heparin forms a complex with AT to enhance the anti‐coagulant activity that inhibits thrombin and FXa. To determine the role of the anti‐coagulant activity of heparin in T_reg induction, we examined the effects of heparin analogues and direct coagulation inhibitors with the inhibition of thrombin (using argatroban) or FXa (using dalteparin or fondaparinux) on T_reg increase in spleen cell cultures without stimulation. Heparin significantly increased the proportion (P = 0·0039 at 5 U/ml) of CD4⁺CD25⁺FoxP3⁺ T_regs (up to 35%) dose‐dependently (Fig. 6a). Furthermore, the increase was not accompanied by cell death of conventional T cells as annexin V binding levels of CD4⁺ T cells did not increase in spleen cell cultures with heparin (Supporting information, Fig. S6a). Protamine sulphate inhibited the increase in T_reg proportion induced by heparin (Fig. 6b and Supporting information, Fig. S6b). Conversely, the proportion of CD4⁺CD25⁺FoxP3⁺ T_regs in the spleen after heparin injection in vivo and the proportion of T_regs among spleen cell cultures after heparin injection in vivo did not change significantly (in vivo, P = 0·3502; cultures after injection in vivo, P = 0·3903) (Fig. 6c). These results suggested that inflammatory or immune responses are necessary to induce T_regs by heparin in vivo. Dalteparin increased CD4⁺CD25⁺FoxP3⁺ T_regs in spleen cell cultures (Fig. 6d). However, no such increase was noted in the presence of fondaparinux or argatroban (Fig. 6d). Dalteparin is a LMWH and, similar to heparin, LMWHs inhibit the coagulation process by binding to AT via a pentasaccharide sequence. Unlike heparin‐activated AT, LMWH‐activated AT does not inhibit thrombin, although it inhibits FXa [29]. Therefore, the anti‐coagulant activity of heparin does not always influence T_reg induction, as FXa inhibition by fondaparinux did not increase T_reg proportion.

Fig. 6.

The stable negative charge of heparin by the sulphate group is important for enhancing regulatory T cell (T_reg) induction. (a,b,d–f) Spleen cells were harvested from BALB/c mice and cultured. (a) Day 5 analysis in murine spleen cells cultured with unfractionated heparin. Left: representative data of CD25⁺forkhead box protein 3 (FoxP3)⁺ T_regs in CD4⁺ cells at indicated heparin concentrations. Right: percentage of CD25⁺FoxP3⁺ T_regs at indicated heparin concentrations. Data are mean ± standard error of the mean (s.e.m.) (n = 4–9). (b) Percentages of CD25⁺FoxP3⁺ T_regs in the cultures with heparin and protamine sulphate. Individual group mean ± s.e.m. (n = 12). (c) Percentage of CD25⁺FoxP3⁺ T_regs in the spleen or in spleen cell cultures after heparin injection in vivo. Individual group mean ± s.e.m. (n = 7–9, n = 5). (d–f) Day 5 analysis in murine spleen cells cultured with Factor Xa (FXa) inhibitor, a thrombin inhibitor or chemically modified heparins. (d) Mean ± s.e.m. percentages of CD25⁺FoxP3⁺ T_regs cultured with FXa or thrombin inhibitors (n = 4–8). (e) Mean ± s.e.m. percentages of CD25⁺FoxP3⁺ T_regs cultured with chemically modified heparins (n = 4). (f) Mean ± s.e.m. percentages of CD25⁺FoxP3⁺ T_regs cultured with heparins and protamine sulphate (n = 12). (g) Day 5 analysis in mixed lymphocyte reaction (MLR) with N‐acetyl heparin. Mean ± s.e.m. percentages of CD25⁺FoxP3⁺ T_regs (n = 6). (h) Day 3 analysis in the culture with N‐acetyl heparin of naive T cells with a plate‐coated anti‐CD3ε antibody with transforming growth factor (TGF)‐β. Mean ± s.e.m. percentages of CD25⁺FoxP3⁺ T_regs (n = 6). *P < 0·05, **P < 0·01, ***P < 0·001, ****P < 0·0001, by Student’s t‐test.

As anti‐coagulation is not involved in T_reg induction, N‐desulphated heparin and N‐acetyl heparin, which are chemically modified heparins lacking anti‐coagulant activity, were used to test their ability to increase the proportion of T_regs in spleen cell cultures. The results showed that, similar to heparin, N‐acetyl heparin (and not N‐desulphated heparin) induced T_regs (Fig. 6e). The effects of dalteparin and N‐acetyl heparin on T_reg induction were inhibited by protamine sulphate (Fig. 6f). Protamine, a positively charged protein, inhibits the effects of heparin by competing with AT and forming an ionic bond with these negatively charged heparins. The negative charge in the N‐desulphated heparin is unstable because the free amino group (NH₃ ⁺) on the N‐sulphate of glucosamine interacts with other sulphate groups. Accordingly, the negatively charged sulphation on heparin is important for the induction of T_regs by these heparins. N‐acetyl heparin also increased the T_reg proportion in the MLRs and the induction of iT_reg from naive CD4⁺ T cells (Fig. 6g,h). These results suggest that the stable negative charge as a result of heparin sulphation is important for the promotion of heparin‐induced T_reg induction.

A chemically modified heparin without anti‐coagulant activity enhances T_reg induction and suppresses allogeneic immune responses

Finally, we examined the improvement in the survival rate of aGVHD mice by administration of N‐acetyl heparin without anti‐coagulant activity or risk of haemorrhage. As expected, the administration of N‐acetyl heparin significantly increased the proportion of splenic T_regs (P = 0·04; Fig. 7a) as well as the number of donor T_regs (P = 0·0478; Fig. 7b) and improved the mortality rate of aGVHD mice (P = 0·0261; Fig. 7c). These results strongly suggest that negatively charged heparin promotes the induction of functional T_regs through the binding of the sulphate group on heparin with certain proteins other than AT and suppresses the immune responses.

Fig. 7.

N‐acetyl heparin enhances the induction of regulatory T cells (T_regs) and suppresses the allogeneic immune responses. (a–c) C57BL/6 recipient mice received lethal total body irradiation (TBI) followed by 5 × 10⁶ allogeneic T cell‐depleted bone marrow (TCD‐BM) (BALB/c) alone or with 1 × 10⁶ allogeneic spleen T cells (BALB/c) on day 0. N‐acetyl heparin was subcutaneously administered to mice from days 0 to 30, every other day. (a) Mean ± s.e.m. number of CD4⁺ cells in the spleen and numbers or percentages of CD25⁺forkhead box protein 3 (FoxP3)⁺ T_regs among splenic CD4⁺ T cells 1 week after allogeneic BMT (n = 6). (b) Mean ± s.e.m. number of H‐2K^d+ donor or H‐2K^d‐ recipient T_regs in the spleen 1 week after allogeneic bone marrow transplant (BMT) (n = 6). (c) Survival curves of allogeneic BMT calculated by the log‐rank (Mantel–Cox) test (*P = 0·0261) in phosphate‐buffered saline (PBS0 versus N‐acetyl heparin (TCD‐BM control; n = 7, PBS; n = 12, N‐acetyl heparin; n = 12). *P < 0·05, by Student’s t‐test.

Discussion

In addition to the anti‐coagulant effect of heparin, anti‐inflammatory and immunosuppressive effects of heparin have been reported. We examined whether T_regs play a part in heparin’s immunosuppressive effects. We demonstrated that heparin treatment reduces aGVHD mouse mortality rates following BMT and increases the proportion of splenic T_regs. The survival rate improved in the high‐dose group of C57BL/6 recipient mice and the low‐dose group of BALB/c recipient mice. These results considered the difference in sensitivity of heparin to the anti‐coagulant effect of mouse strains; for example, differences in the level of coagulation factors and the number of platelets [26]. In BALB/c recipient mice 1 week after transplantation severe haematomas were found in the high‐dose group, but there were higher proportions of donor T_regs in the spleen of the high‐ than in the low‐dose group. In the high‐dose group of BALB/c recipient mice, the mortality due to frequent lethal haemorrhage might increase. As the anti‐coagulant effect of heparin has previously been thought to improve GVHD in previous studies, we examined the effect of heparin on T_regs in GVHD mice. The presence of donor T_regs is essential for improving the survival of GVHD mice by heparin administration, because improvement in GVHD was not observed when T cells from scurfy mice were used as donor cells. These results suggest that the anti‐coagulant effect and exertion of T_reg‐mediated immunosuppression are two separate functions of heparin. Several studies have demonstrated that CD4⁺CD25⁺FoxP3⁺ T_regs control CD4⁺ and CD8⁺ conventional T cell (T_conv) proliferation, limiting aGVHD lethality yet retaining anti‐viral and graft‐versus‐tumour activity and promoting animal survival [18, 19, 22]. Donor T_regs have also been shown to be the most effective protection against aGVHD in allogeneic haematopoietic stem cell transplantation [30]. In our study, heparin induced T_regs, including donor T_regs, in aGVHD mice and inhibited T_conv proliferation in MLRs.

In addition, heparin can inhibit the activation of various inflammatory cells, including neutrophils and eosinophils, as well as platelets [31, 32, 33]. Heparin binds to and neutralizes various inflammatory mediators and enzymes released during the inflammatory process. It also binds directly to several adhesion molecules expressed during inflammation [34, 35, 36]. Therefore, it may be possible that the heparin‐induced promotion of T_reg induction involves IL‐2 as a heparin‐binding protein. Previous studies have reported that the binding ability of heparin to IL‐2 does not interfere with IL‐2/IL‐2 receptor interactions [37]. It is thought that binding to glycosaminoglycans, such as heparin, may be the mechanism responsible for the local active form cytokine retention close to the tissue secretion site. In the present study, we found that negatively charged heparin promotes the induction of T_regs. The addition of protamine sulphate is thought to inhibit heparin’s effects on T_regs by neutralizing the negatively charged heparin. We are also examining the possibility of inhibiting T_reg induction and the induction of T helper type 17 (Th17) cells with protamine. Accordingly, the negatively charged sulphation on heparin is essential for T_reg induction by heparin. Our splenocyte culture results demonstrated that heparin sulphation plays a critical role in enhancing T_reg induction, and that N‐acetyl heparin, a chemically modified heparin with reduced haemorrhage risk, has potential immunoregulatory functions via T_reg induction. Importantly, our study suggests that heparin did not induce T_regs in steady‐state mice in vivo and induced de‐novo T_reg generation in response to inflammatory or immune responses.

Several studies have shown that heparin has inhibitory effects on T cell proliferation in MLRs [4, 38] and natural killer cell cytotoxicity [39]. Reduced differentiation of Th1, Th2, and Th17 in the spleen by both heparin and LMWH treatment has also been reported [40]. The results of these studies support the hypothesis that heparin promotes T_reg induction. Indeed, in the present study, heparin enhanced FoxP3 expression and maintained differentiated T_reg survival in cooperation with IL‐2. As IL‐2 is produced by T cells during the immune response, the action of heparin alone may cancel out the immunosuppressive effects of T_regs in steady‐state in the absence of IL‐2. The magnitude of T_reg functionality may be up‐regulated in the IL‐2 existing site, even if the number of T_regs does not significantly increase in aGVHD mice or in allogeneic MLRs upon heparin administration. Activated T_regs expressing GARP/latent TGF‐β1 complexes are potent inducers of Th17 cell differentiation in the presence of exogenous IL‐6 and inducers of T_reg differentiation in the presence of IL‐2 [28]. It is possible that heparin can indirectly inhibit Th17 cell differentiation by IL‐2 production. However, it is still unclear whether heparin promotes TGF‐β activation. T_reg induction by heparin is dependent upon TGF‐β and independent of co‐stimulatory signals in both murine and human cells. Heparin did not influence SMAD2/3 phosphorylation, but increased the production of IL‐2 in stimulated naive T cells and nuclear translocation of NFATc1. The IL‐2 signalling pathway (and not the TGF‐β signalling pathway) is assumed to be critical for heparin‐induced T_regs. In this study, we analysed the early period of T_reg induction. Considering ex vivo T_reg therapy, the effect of heparin in T_reg expansion should be examined. To confirm the effects of heparin on T_reg expansion with CD28 co‐stimulation and IL‐2, it should be analysed after culturing for more extended periods.

As the heparin dosage used in our study was a prophylactic dose for thrombosis in clinical sites, we believe that the obstacles to the clinical application of immunomodulatory therapy using heparin are low. Non‐anti‐coagulant heparins, such as N‐acetyl heparin, can also be used for patients who are considered at risk of bleeding. Here, we show that the promotion of T_reg induction by heparin in a patient can be confirmed for the MLRs or iT_reg induction using the patient’s PBMCs. Consequently, it is possible to examine in advance the effect of heparin in promoting the induction of T_regs in patients to whom heparin treatment is applied. In addition, previous studies have shown that human leucocyte antigen‐matching influenced the production of T_regs in human MLRs [41] and that T_regs were up‐regulated in LMWH‐treated thrombophilic pregnancies [42], which implies the necessity for further detailed research on heparin‐induced T_regs in human immunological diseases. Recently, the effects of heparin on the behaviour of stromal cells were elucidated [43]. Moreover, we have previously reported that stromal cells support T_reg survival via CD2 [44]. Therefore, the effects of heparin on stromal cells may impact T_reg survival, which would affect the proportion of T_regs. Further studies to elucidate the detailed mechanism by which heparin contributes to T_reg induction and activation are required. In conclusion, we have demonstrated that heparin promotes both de‐novo generation and activation of T_regs by IL‐2 production and that administration of heparin and derivative substances ameliorates lethal aGVHD by T_reg induction independently of anti‐coagulative properties. These results are promising, and readily translatable to clinical trials where the induction of T_regs by heparin analogues could result in the prevention and treatment of various immunopathological conditions, such as aGVHD, graft rejection, inflammatory disease and autoimmune diseases.

Disclosures

The authors declare no conflicts of interest.

Author contributions

Y. K. and H. K. designed and performed the experiments, analysed the results and wrote the manuscript; Y. K. performed the experiments; M. H. analysed the results; and T. K. designed the experiments, analysed the results and wrote the manuscript.

Supporting information

Fig. S1. Cell number of Tregs and chimerism one week after allogeneic bone marrow transplantation in C57BL/6 recipient mice. C57BL/6 recipient mice received lethal TBI (1100 cGly) followed by 5 × 106 allogeneic T cell depleted‐bone marrow cells (TCD‐BMs; BALB/c) alone or with 1 × 106 allogeneic spleen T cells (BALB/c) on day 0. Heparin was subcutaneously administered to mice from day 0 to 7, every other day. (a) Mean ± SEM numbers of CD4+ cells in the spleen, and CD4+CD25+Foxp3+ Tregs among splenic CD4+ T cells one week after allogeneic BMT in C57BL/6 recipient mice (n = 7‐8). (b) Mean ± SEM percentages and number of H‐2Kd+ donor and H‐2Kd‐ recipient Tregs in the spleen one week after allogeneic BMT in C57BL/6 recipient mice (n = 7‐8). p values by paired Student's t‐test.

Fig. S2. Heparin confers a severe risk of haemorrhage in the high‐dose group of BALB/c recipient aGVHD mice. BALB/c recipient mice received lethal TBI (800 cGly) followed by 5 × 106 allogeneic TCD‐BMs (B6.Foxp3‐YFP) alone on day 0. Heparin was subcutaneously administered to mice from day 0 to 30, every other day. (a) Effect on the dorsal subcutis by subcutaneous injection at the indicated dose of heparin one week after bone marrow transplantation in BALB/c recipient mice. Red circle: subcutaneous haemorrhage and hematoma.

Fig. S3. Heparin suppresses the proliferation of CD8+ T cells in allogeneic‐MLRs. Spleen cells were harvested from BALB/c mice and co‐cultured with irradiated spleen cells from C57BL/6 mice as alloreactive MLRs. (a) CFSE‐labelled BALB/c spleen cells were used as responder cells in MLRs with heparin. Representative plots of CFSE levels in CD8+ or CD4+ T cells five days after MLR culture at the indicated concentrations of heparin. (b) Mean ± SEM geometric MFI of CFSE in CD8+ and CD4+ T cells in the MLRs with BALB/c responder cells (n = 5). p values by paired Student's t‐test.

Fig. S4. Heparin does not drastically enhance Treg induction with CD28 co‐stimulation and IL‐2. Naïve T cells were harvested from the spleen of BALB/c mice and cultured with plate‐coated anti‐CD3ε antibody (1μg/mL) and anti‐CD28 antibody (1μg/mL) in the presence of TGF‐β (5 ng/mL) and IL‐2 (10 ng/mL). Three days after culture, the expression levels of CD25 and Foxp3 in harvested cells were analysed. Data are from three independent experiments with similar results.

Fig. S5. Heparin does not enhance TGF‐βR/Smad2/3 signalling. (a) Naïve splenic T cells were cultured with TGF‐β (5 ng/mL). Representative plots of pSmad2/3 expression in cells were collected at the indicated times after culture with heparin. Data are from three independent 4 experiments with similar results. (b) Naïve T cells from the spleen were cultured with platecoated anti‐CD3ε mAb (2C11: 5 μg/mL) with TGF‐β (5 ng/mL). Left: Representative plots of pSmad2/3 expression in cells collected 30 min after culture with heparin. Right: Mean ± SEM values of geometric fluorescence intensity in cells cultured with heparin at 30 min (n = 5). p values by paired Student's t‐test.

Fig. S6. Treg induction by heparin and protamine in the spleen cell cultures. (a) Spleen cells were harvested from C57BL/6‐Foxp3‐YFP mice cultured in the presence of heparin. Top panels: Representative profiles of CD25 and Foxp3 expression levels in CD4+ cells collected two days after culture. Lower panels: Representative plots of annexin V binding levels in CD25‐Foxp3‐, CD25+Foxp3‐, CD25‐Foxp3+, and CD25+Foxp3+ cells collected at the indicated times after culture. Data are from three independent experiments with similar results. (b) Day 5 analysis in murine spleen cell cultures with protamine sulphate. Left: Percentage of CD25+Foxp3+ Tregs at indicated protamine sulphate concentrations. Right: Percentages of CD25+Foxp3+ Tregs in the cultures with protamine sulphate or protamine sulphate. Data are mean ± SEM (n = 6). p values by paired Student's t‐test.

Table S1. Heparin increases regulatory T cells (Tregs) in alloreactive mixed lymphocyte reactions (MLRs) of human peripheral blood mononuclear cells (PBMCs). Human PBMCs collected from healthy volunteers were co‐cultured with irradiated‐PBMCs from another donor as an alloreactive MLR. Seven days after culture, cells were harvested and expression levels of CD25 and Foxp3 were analysed. Raw percentage data of CD25+Foxp3+Treg in CD4+ cells and calculated data of the increasing ratio index are shown.