Heparin enhances uptake of platelet factor 4/heparin complexes by monocytes and macrophages

Summary

Background

Heparin-induced thrombocytopenia (HIT) is an iatrogenic complication of heparin therapy caused by antibodies to a self-antigen, platelet factor (4) and heparin. The reasons why antibodies form to PF4/heparin, but not to PF4 bound to other cellular glycosaminoglycans are poorly understood.

Objective

To investigate differences in cellular responses to cell-bound PF4 and PF4/heparin complexes, we studied the internalization of each by peripheral blood-derived monocytes, dendritic cells and neutrophils.

Methods and Results

Using unlabeled, fluorescently-labeled antigen and/or labeled monoclonal antibody to PF4/heparin complexes (KKO), we show that PF4/heparin complexes are taken up by monocytes in a heparin-dependent manner and are internalized by human monocytes and dendritic cells, but not by neutrophils. Complexes of PF4/low-molecular weight heparin and complexes composed of heparin and murine PF4, protamine, or lysozyme are internalized similarly, suggesting a common endocytic pathway. Uptake of complexes is mediated by macropinocytosis, as shown by inhibition using cytochalasin D and amiloride. Internalized complexes are transported intact to late endosomes, as indicated by co-staining of vesicles with KKO and lysosomal associated membrane protein-2 (LAMP-2). Lastly, we show cellular uptake is accompanied by expression of MHCII and CD83 co-stimulatory molecules.

Conclusions

Taken together, these studies establish a distinct role for heparin in enhancing antigen uptake and activation of the initial steps in the cellular immune response to PF4-containing complexes.

Introduction

Heparin-induced thrombocytopenia (HIT) is a life-threatening thrombotic disorder caused by antibodies to ultra-large complexes (ULCs) of platelet factor 4 (PF4; CXCL4) and heparin. Once antibodies form to PF4/heparin ULCs, it is well-recognized that other negatively charged carbohydrates, such as cell-surface glycosaminoglycans (GAGs) readily substitute for heparin in complex formation. In vitro, HIT antibodies recognize PF4 bound to a variety of cell surfaces, including endothelial cells [1], monocytes [2] and neutrophils [3]. PF4 bound to cellular GAGs serves as an antigenic target for PF4/heparin antibodies and presumably mediates the syndrome of delayed-onset HIT [4–6], a clinical variant of HIT that occurs days to weeks after discontinuation of heparin.

While heparin may be expendable for disease pathogenesis once HIT antibodies form, there is little controversy regarding drug requirement for the initiation of the HIT immune response. With the exception of a few case reports of spontaneous HIT [7, 8], the vast majority of clinically diagnosed HIT occurs in the wake of heparin exposure. Why cell-bound PF4 does not elicit antibody formation, or why HIT does not occur more often as a spontaneous autoimmune disease is not known. Indeed, recent studies indicate that B-cells from mice and healthy adults are capable of producing PF4/heparin specific antibodies in response to inflammatory stimuli independently of heparin exposure [9]. Yet, clinical HIT is infrequently seen in patients with autoimmune disorders [10–12].

To explore the role of heparin in the initiation of the cellular immune response to PF4/heparin, and to understand differences in cellular responses to cell-bound PF4 and PF4/heparin complexes, we studied antigen uptake and processing using labeled PF4, heparin and KKO, a monoclonal antibody specific for PF4/heparin complexes [13]. Using confocal imaging and flow cytometry, we show that heparin markedly augments the uptake of PF4 by monocytes/macrophages and enhances cellular activation leading to expression of immune co-stimulatory molecules. Lastly, we show that uptake of PF4/heparin complexes is mediated through macropinocytosis through pathway shared with the uptake of other complexes formed between heparin and cationic proteins.

Materials and Methods

Reagents

Recombinant human (h) and murine (m) PF4 were purified as previously described [14, 15]. Unfractionated heparin (UFH; Units/mL; Heplock) was purchased from Elkins-Sinn Inc. For stoichiometric calculations involving UFH, we utilized previously published estimates of UFH specific activity at 140 U/mg [14, 16] and mean Mw of 15 kDa [14, 16]. Low molecular weight heparin (LMWH; MW ~4500 Da, 100 mg/mL was purchased from Sanofi-Aventis Pharmaceuticals). Fibronectin from human plasma, E-toxate kit (to detect endotoxin), amiloride, cytochalasin D, protamine, and 4′, 6′-diamidino-2-phenylindole dihydrochloride (DAPI), and lipopolysaccharides (LPS) were purchased from Sigma (St. Louis, MO, USA). Pierce High Capacity Endotoxin Removal Spin Columns were purchased from Thermo Scientific (Rockford, IL). Paraformaldehyde was purchased from Mallinckrodt Chemicals (Raleigh, NC, USA). Ficoll-Paque was purchased from GE Healthcare (Little Chalfont, UK). Human intravenous immunoglobulin (IVIG; Gammunex-C) was procured from Grifols (Los Angeles, CA). Media M199, RPMI 1640, secondary Abs- Streptavidin Alexa Fluor 568, and Alexa Fluor 594, sodium pyruvate, β-mercaptoethanol, minimal essential media (MEM), non-essential amino acids, L-glutamine and heparin-FITC were purchased from Life Technologies (Carlsbad, CA, USA). Fluoromount-G mounting medium was purchased from SouthernBiotech (Birmingham, AL, USA). For confocal studies, HistoBond slides and cover slips were purchased from VWR International (Suwanee, GA). Human GM-CSF and human IL-4 were purchased from PeproTech (Rocky Hill, NJ, USA).

The following fluorescent antibodies and reagents used for confocal microscopy and flow cytometry, were purchased from eBioscience (San Diego, CA, USA): Anti-human biotin-conjugated CD14, CD1a-PE, biotin conjugated lysozomal associated membrane protein-2 (LAMP-2), anti-human CD1a PE/FITC, anti-human CD14 FITC/PE/AF647, anti-human HLA-DR (MHCII) PE, anti-human CD83 PE, IC fixation buffer and permeabilization buffer. A murine monoclonal antibody (KKO) specific for hPF4/heparin complexes was generated by our laboratory [13] and conjugated with Alexa Fluor 647 (AF647; Molecular Probes) to stain hPF4/heparin complexes.

Cell culture studies

Blood was collected from medication-free healthy donors under a protocol approved by Duke’s Institutional Review Board (Protocol #: Pro 00012901) and consent obtained in accordance with the Declaration of Helsinki. Peripheral blood derived monocytes (PBMCs) were isolated from citrated blood using density separation [17]. For all reagents used in cell-culture assays, final endotoxin levels of antigens and antibodies were below 1 EU/ml (0.1 ng/ml), as measured using a Sigma E-toxate kit. Where necessary, endotoxin was removed using endotoxin removal columns (Pierce).

Generation of human immature dendritic cells (hiDCs)

Dendritic cells were generated from human monocytes isolated from PBMCs by either plastic adherence method or elutriation. Purified monocytes were cultured for 7 days at 37°C, 5% CO₂ at 1×10⁶ cells/ml in RPMI 1640 supplemented with L-glutamine, 10% FBS, gentamicin, sodium pyruvate, β-mercaptoethanol, and non-essential amino acids along with GMCSF and IL-4. Half of the culture media was exchanged every 2 days with fresh media containing twice the concentration of the cytokines mentioned above. After 7 days, cells were harvested and purity was assessed by flow cytometry after staining with anti-CD1a and anti CD14 antibodies.

Confocal microscopy

For confocal microscopy experiments, approximately 1 X 10⁶ cells were allowed to adhere to fibronectin-coated glass coverslips for 2–3 hours. Thereafter, cells were washed with wash buffer containing phosphate buffered saline (PBS, Invitrogen/Life Technologies), 0.5% human serum albumin, 4 mg/mL IVIG and 0.1% Tween-20 and incubated with pre-formed hPF4 ± heparin in M199 media, for varying times at 37°C. To inhibit actin polymerization, cells were incubated with 5 μM cytochalasin D for 1 hour at 37°C along with labeled complexes and then incubated for 24 hours. To inhibit macropinocytosis, cells were pre-incubated with 0.1 mM amiloride in media for 15 minutes prior to addition of labeled complexes. After incubation with antigen, cells were washed and fixed with 4% paraformaldehyde and counterstained with one or more of the following labeled-antibodies: CD14 (monocyte marker), CD1a (dendritic cell marker), KKO (anti-hPF4/heparin monoclonal antibody, mAb), and DAPI (nuclear stain). Coverslips were mounted on slides with Fluoromount-G mounting medium, and examined and photographed with a Leica SP5 inverted confocal microscope using a 100X oil immersion lens (Leica Microsystems, version-LAS AF 2.6; Wetzlar, Germany), with digital zoom (2-10X) and accompanying Leica LAS AF 2.6 software (Buffalo Grove, IL).

Flow cytometry of PF4/heparin uptake and cellular activation

Freshly prepared PBMCs were incubated in M199 medium alone or with complexes formed by PF4 (25 μg/mL) and varying heparin concentration. Cells with or without antigen were incubated at 37°C, 5% CO₂ or LPS as a positive control (final concentration 1 μg/ml). After defined time intervals of antigen incubation, cells were washed twice with FACS buffer (PBS with 2 % FBS). Preliminary studies showed that cell surface bound PF4/heparin complexes can be removed by washing cells with excess heparin (data not shown). Therefore, to detect only internalized complexes, cells were incubated for 5 minutes in buffer containing excess heparin (100U/ml) followed by two additional washes. To block Fc receptors, cells were incubated with IVIG at 4°C for 10 minutes and incubated with fluorescently labeled antibodies or isotype antibodies for 30 minutes at 4°C for surface staining. To identify intracellular complexes, cells were incubated with antigen, washed in excess heparin and treated with fixation buffer at room temperature (RT) for 20 minutes. Cells were washed three times with permeabilization buffer, incubated in the same buffer with fluorescently labeled KKO or isotype antibody for 30 minutes at 4°C. Cells were then subjected to two additional washes and, resuspended in FACS buffer. Cells were analyzed using a BD FACS Canto Flow cytometer (BD Biosciences). Analyses were performed using FCS express software (De Novo Software, Los Angeles, CA). Data is reported as mean fluorescence intensity (MFI) of labeled antibody staining.

Statistical Analysis

Enumeration of PF4/heparin ultra-large complexes are expressed as the mean ± standard deviation (SD). Analysis of variance for categorical variables (ANOVA) for 3 or more groups was done using One Way ANOVA analysis with Tukey’s multiple comparison test to determine statistical significance. Statistical analyses were performed using GraphPad Prism (Graph Pad Software Version 5.02, La Jolla, CA).

Results

Monocytes internalize labeled and unlabeled PF4/heparin particles in a time-dependent manner

PF4 and heparin interact through electrostatic forces to form high molecular weight ULCs that are stable at room temperature [16, 18]. To confirm formation and visualization of PF4/heparin complexes using fluorescently labeled heparin, we incubated hPF4 (1–25 μg/ml) and heparin-FITC (0.0125–0.25 U/mL) at various concentrations to yield fixed PF4:heparin molar ratios or PHRs (PHR: 5–7). Consistent with previous studies [17], higher concentrations of PF4 and heparin generated complexes of increasing size (Supplemental Figure 1).

We next asked if PF4/heparin complexes were internalized by monocytes or were retained on the cell-surface when followed over an extended period of time. For these studies, we incubated PF4/heparin-FITC labeled complexes (hereafter, PF4/heparin concentrations will be represented as PF4 in μg/ml and heparin in U/ml; 25/0.25 at PHR=7) with PBMCs for 0–24 hours. In confocal studies, e.g. as shown in Figure 1 (See Supplemental Figure A for accompanying low magnification images), monocytes can be distinguished from lymphocytes by membrane CD14 staining (red stain), which is absent from lymphocytes (depicted “L” in left panel of Figure 1A). PF4/heparin-FITC complexes were evident along the monocyte cell-surface by 2 hours. Internalization of PF4/heparin-FITC particles was notable by 6 hours and maximal by 24 hours. Three-dimensional rendering of 24 hour confocal images confirmed intracellular location of PF4/heparin complexes (Figure 1B). Quantification of percentage of cells with intracellular fluorescent complexes is shown in Figure 1C. Similar kinetics of PF4/heparin uptake was noted using flow cytometry, as shown in Figures 1D & 1E. By flow, maximal MFI of intracellular staining of PF4/heparin complexes by KKO was seen at 24 hours (Figure 1E). Incubation with lower concentrations of PF4 and heparin showed visible internalization of complexes formed at PF4/heparin concentrations as low as 10/0.1 and 5/0.05, but not at 2.5/0.025 (Supplemental Figure 2)

Figure 1. Time dependent uptake of labeled-PF4/heparin complexes & 3D visualization of particle intake.

PBMCs were incubated with hPF4/heparin-FITC complexes (25/0.25; green) for 0, 2, 6, 12 and 24 hours at 37°C and evaluated by confocal microscopy or flow cytometry. Results are representative of 3 or more independent experiments. (A) Time dependent uptake of PF4/heparin complexes. Monocyte (M) cell-surface staining with CD14-PE is shown as red, and cell nuclei shown by DAPI-staining as blue. Lymphocytes (L) shown in the first left hand panel lack staining with CD14. Confocal images shown are at 100X magnification with varying optical zoom (2-5X) as indicated by “OZ”. An accompanying low magnification image is shown in Supplemental Figure A. (B) 3D Visualization of the internalized particles. A 3D image was generated by creating iso-surfaces in XYZ plane using Imaris 7.6 software. (C) Graphical representation of particle uptake. Intact cells with CD14-PE staining and cells with internalization of PF4/heparin particles were counted and the percentage of cells containing intracellular vesicles is shown. (D) Staining of PF4/heparin complexes by KKO-AF647. At depicted time points, PBMCs (10,000 events collected) were gated on CD14 positive cells and stained for intracellular PF4/H complexes using KKO-AF647. (E) Graphical representation of flow cytometric uptake shown in Figure 1D.

To confirm that uptake was attributable to hPF4/heparin complexes and not to the FITC label, we incubated PBMCs with heparin-FITC alone, hPF4-AF647/heparin or unlabeled hPF4/heparin counterstained with a labeled anti-hPF4/heparin monoclonal antibody, KKO-AF647 (Supplemental Figure 3). Using alternative fluorescent labels and KKO binding, these studies confirm that hPF4/heparin complexes, are taken up by monocytes, irrespective of fluorescent label, showing intracellular vesicles containing PF4 co-localized with heparin at 24 hours.

Heparin markedly enhances cellular uptake of PF4

PF4 binds to cell-surface GAGs on a variety of cell-surfaces, including monocytes [19]. To determine if hPF4/GAGs form complexes that are also internalized by monocytes, cells were incubated with unlabeled hPF4 (25 μg) with or without increasing concentrations of unlabeled heparin (0.1–2.5 U/mL; PHRs 16 to 0.67) for 24 hours, followed by fixation, cell permeabilization and counterstaining with KKO-AF647 and uptake was visualized by confocal microscopy (Figure 2A and Supplemental Figure B for accompanying low magnification images) or flow cytometry (Figure 2C & 2D). As shown in Figure 2, uptake of PF4 by monocytes showed marked heparin dependence. Irrespective of technique, uptake of unlabeled hPF4 by PBMCs increased at heparin concentrations up to an optimal PF4:heparin ratio (25/0.25), followed by gradual loss of KKO staining with increasing concentrations of heparin. Figure 2B quantifies confocal image data from these experiments, showing the percentage of CD14-labeled monocytes containing fluorescent intracellular vesicles as a function of heparin concentration. Figure 2C shows comparable flow cytometry data of CD14-gated cells after incubation with complexes and intracellular staining with KKO-AF647. Both methods confirm marked heparin-dependent internalization of PF4 containing complexes by monocytes. Because internalization is maximal at a hPF4/heparin of 25/0.25 (PHR=7), all subsequent studies were performed using these concentrations and PHR.

Figure 2. Heparin augments the uptake of PF4/heparin complexes.

Cells were incubated with unlabeled PF4 (25 μg/ml) alone or incubated with unlabeled heparin (0.1, 0.25, 1 and 2.5 U/ml) for 24 hours at 37°C, and counterstained with KKO-AF647 (shown as yellow to distinguish from CD14-PE, pink color), as described in methods and analyzed by confocal imaging or flow cytometry. Confocal images shown are at 100X magnification with varying optical zoom (2-5X) as indicated by “OZ”. An accompanying low magnification image is shown in Supplemental Figure B. Results are representative of 3 or more independent experiments. (A) Heparin-dependent uptake of PF4/heparin complexes by PBMCs. Monocyte cell surface was stained with CD14-PE (pink), and DNA was stained with DAPI (blue). The images are shown with optical zoom (1-10X) of the respective 100X image. Internalized PF4/heparin complexes are shown with KKO-AF647 (yellow color). (B) Quantification of the hPF4/heparin uptake by PBMCs. The proportion of cells with internalized complexes among the total number of CD14⁺cells in a microscopic field is shown on y-axis as a function of heparin concentration (x-axis). (C) Visualization of internalized PF4/heparin complexes by flow cytometry. Representative flow histogram showing the internalized PF4/heparin complexes (KKO A 647 staining) in the CD14 PE gated monocytes. (D) Mean fluorescence intensity (MFI) of intracellular PF4/heparin complexes labeled with KKO. MFI of cells containing complexes formed with various concentrations of heparin or untreated cells are shown.

Antigen specificity of complex uptake

To determine whether internalization of hPF4/heparin complexes requires recognition of human PF4 or UFH specifically, we performed experiments using other positively-charged, heparin-binding proteins {murine PF4 (mPF4), protamine (PRT) and lysozyme (LYS)} and hPF4 complexed with LMWH. In these experiments, mPF4, PRT or LYS was incubated with heparin-FITC at stoichiometric ratios known to form complexes [20]. In other experiments, hPF4-AF488 was incubated with increasing concentrations of LMWH. As shown in Figure 3A (See Supplemental Figure C for low magnification), heparin-bound complexes consisting of mPF4, PRT or LYS were all internalized by monocytes and visualized as intracellular vesicles at 24 hours. When hPF4 was incubated with LMWH, we noted a LMWH-dependent increase in PF4/LMWH complexes, as seen with hPF4/unfractionated heparin (as seen in Figure 2A). These studies demonstrate that the uptake of hPF4/heparin by monocytes is mediated by the formation of complexes rather than through an unidentified hPF4-receptor mediated pathway.

Figure 3. Antigen specificity of complex uptake.

Confocal images shown are at 100X magnification with varying optical zoom (2-5X) as indicated in Figure by “OZ”. An accompanying low magnification image is shown in Supplemental Figure C. Results are representative of three independent experiments. (A) Monocyte uptake of other positively charged protein/heparin complexes. PBMCs were incubated with complexes of other positively charged proteins and heparin-FITC, including mPF4 (25/0.25), PRT (31/0.4), LYS (31.2/0.7) for 24 hours at 37°C. Monocytes are stained with CD14-PE (pink) and DNA stained with DAPI (blue). (B) Monocyte uptake of PF4/LMWH complexes. Cells were incubated with PF4 alone (25 μg/ml) or PF4 and LMWH at varying ratios (0.1, 0.5, 1 and 2.5 μg/ml) at 37°C for 24 hours. Monocytes are stained with CD14-PE (pink) and PF4/LMWH (~4.5 KDa) complexes were detected using monoclonal Ab KKO-AF488 (green). (C) Quantification of the hPF4/LMWH uptake by PBMCs. The proportion of cells with internalized complexes among the total number of CD14⁺cells in a microscopic field is shown on y-axis as a function of LMWH concentration (x-axis). Significant differences (p<0.05) are indicated by star symbol.

Cellular specificity of PF4/heparin complex uptake

To determine if cellular uptake of PF4/heparin is specific to monocytes, we examined uptake by neutrophils and dendritic cells (DCs), other intravascular cells that are capable of ingesting and clearing particulate antigens. As shown in Figure 4A (See Supplemental Figure D for accompanying low magnification images), when hPF4/heparin complexes were incubated with neutrophils, the labeled complexes remained in the extracellular space and there was minimal intracellular uptake (Figure 4A, left panel). Furthermore, when monocytes (Figure 4A, middle panel) or monocytes together with neutrophils were incubated with hPF4/heparin (Figure 4A, right panel), uptake was restricted to the CD14-expressing monocytes. To determine if DCs internalize hPF4/heparin complexes, PBMCs-derived human immature DCs (hiDCs) were incubated with hPF4/heparin-FITC and stained for DC markers CD1a or monocyte CD14. As shown in Figure 4B (See Supplemental Figure D for accompanying low magnification images), internalization of hPF4/heparin complexes was readily evident in cells expressing CD1a and lacking CD14 (characteristic of DCs).

Figure 4. Cellular specificity of PF4/heparin complex uptake.

(A) Studies of neutrophil uptake. Peripheral blood derived neutrophils, monocytes and mixture of neutrophils and monocytes were incubated with PF4/heparin-FITC (green) particles at ratio 25/0.25 for 24 hours at 37°C. Left-most panel shows only isolated neutrophils with PF4/heparin complexes; middle panel shows monocyte populations with PF4/heparin complexes and right-most panel shows combined monocytes and neutrophils. Neutrophils are stained with CD66-APC (red) (shown with white arrows) and monocytes are stained with CD14-PE (pink), and cellular DNA with DAPI (blue). (B) Studies of hiDCs uptake. PBMC derived hiDCs were incubated with PF4/heparin-FITC complexes (green) and stained with CD1a-PE (pink) and DNA is counterstained with DAPI (blue). The complexes were also co-stained with KKO AF647 (red border around green complexes). Images shown are the optical zoom (2-5X) of the respective 100X image. An accompanying low magnification image is shown in Supplemental Figure D. Confocal images shown are at 100X magnification with varying optical zoom (1-10X) as indicated by “OZ”. Results are representative of three independent experiments.

Mechanism of cellular uptake of PF4/heparin

Cellular uptake of antigen is known to be size-dependent [21]. For soluble antigens (<500 nm in size), uptake occurs via diverse endocytic mechanisms involving coat proteins [22]. For large or particulate antigens (>500 nm), uptake generally occurs via macropinocytosis or receptor-mediated phagocytosis. These two mechanisms are active cellular processes that require cytoskeletal rearrangement and organization of intracellular vesicles [23]. To examine the mechanism of cellular uptake of hPF4/heparin, a series of studies were undertaken to examine the role of temperature, cytoskeleton and endocytosis of hPF4/heparin-bearing vesicles. As shown in Figure 5A (See Supplemental Figure E for accompanying low magnification images), uptake of PF4/H was temperature-dependent. There was virtually no internalization at 4°C as compared to 37°C. To examine requirements for cytoskeletal rearrangement, PBMCs were pre-incubated with cytochalasin D, an actin polymerization inhibitor that blocks cytoskeletal reorganization. As shown in Figure 5B, monocytes treated with cytochalasin D did not internalize hPF4/heparin complexes. Because internalization of hPF4/heparin, PRT/heparin and LYS/heparin complexes was not protein specific, we asked if internalization occurred via macropinocytosis, a ligand-independent pathway for uptake of large macromolecules. Macropinocytosis, unlike phagocytosis, is highly sensitive to amiloride, a Na⁺/H⁺ exchanger [24]. For these studies, we pre-treated PBMCs with amiloride or vehicle followed by hPF4/heparin complexes. As shown in Figure 5C, we noted marked attenuation of cellular internalization of hPF4/heparin in the presence of amiloride as compared to vehicle treated cells. Finally, to determine if intracellular complexes are processed through an endosomal pathway, we co-stained hPF4/heparin containing vesicles with LAMP 2, a marker of late endosomal maturation. As shown in Figure 5D, hPF4/heparin-containing vesicles stained positive for LAMP- 2, indicating that by 24 hours the complexes had accumulated in a late endosomal compartment.

Figure 5. Mechanisms of cellular uptake of PF4/heparin complexes.

Mechanism of cellular uptake was investigated as described in methods. For all studies, monocytes stained with CD14-PE (pink) and DNA stained with DAPI (blue). Confocal images shown are at 100X magnification with varying optical zoom (2-5X) as indicated by “OZ”. An accompanying low magnification image is shown in Supplemental Figure E. Results are representative of three independent experiments. (A) Effects of temperature. Cells were incubated at 4°C or 37°C. Complexes were co-stained with KKO AF647 (red). (B) Requirements for cytoskeletal rearrangement. Cells were treated with or without Cytochalasin D (5 μM) and incubated with PF4/heparin-FITC (green). Complexes were co-stained with KKO AF647 (red) (C) Effects of amiloride. Cells were incubated with amiloride (0.1 mM) for 15 minutes followed by incubation with PF4/heparin-FITC complexes (green). (D) Co-localization of LAMP-2 with intracellular PF4/heparin complexes. Cells were stained with LAMP-2-Biotin/Streptavidin AF 594 (colored as yellow). Complexes were stained with heparin-FITC (green). The co-localization of LAMP-2 (yellow) and PF4/heparin-FITC (green) complexes is shown in the additional image.

PF4/heparin internalization is accompanied by cellular activation

We then asked if the internalization of hPF4/heparin was followed by cellular activation and upregulation of cellular activation molecules (MHCII, CD86 and CD83). To do so, we performed flow cytometric studies of monocytic cells bearing hPF4/heparin complexes. Gating by CD14⁺ cells revealed increased expression of MHCII (Figure 6A) and CD83 (Figure 6B), but not CD86 expression (data not shown) at 24 hours. Taken together, these studies confirm that cellular uptake of PF4/heparin is accompanied by cellular activation and expression of co-stimulatory molecules likely involved in the initiation of the immune response.

Figure 6. Uptake of PF4/heparin complexes trigger cellular activation.

(A) Cellular expression of MHCII. Cells incubated with varying PF4/heparin complexes were stained for flow cytometry as described in materials and methods section. Mean fluorescence intensity of MHC II expression on CD14 FITC positive cells was determined and expressed relative to the mean fluorescence intensity (MFI) of the control untreated cells. (B) Cellular expression of CD83. Cells incubated with varying PF4/heparin complexes were gated by FITC and CD14-PE-expressing cells, co-stained with CD83-APC and analyzed for MFI (mean ± SD). The mean ± SD from three independent experiments is shown.

Discussion

PF4 is an abundant platelet protein (1.36 + 0.24 ng/fL × 10⁻⁶) [25] released at sites of platelet activation. PF4 binds to cellular GAGs with high affinity and forms antigenic complexes that are recognized by PF4/heparin antibodies [1, 2]. Yet, in the host, the PF4/GAG complex differs markedly from the PF4/heparin complex in its ability to elicit an immune response.

Several theories have been advanced to explain why the PF4 self-antigen is immunogenic in some settings but not others. Greinacher and colleagues suggest that immunogenicity to PF4 may be primed by prior bacterial infection [26–28]. In this bacterial immunization model, infections associated with platelet activation and PF4 release generate circulating PF4/bacterial complexes that activate the innate immune system and prime the immune system to respond to subsequent heparin exposure. Wang and colleagues suggest that PF4 immunogenicity results from a breach in peripheral tolerance mechanisms. These investigators find auto-reactive B-cells with anti-PF4 specificity in healthy mice and humans [9]. Their studies suggest that inflammation, surgery and/or infection may breach self-tolerance and lead to proliferation of auto-reactive B-cells and PF4/heparin antibody production.

Our studies, described here, offer additional insights as to why an immune response to PF4 occurs more commonly in the context of heparin exposure, and rarely in its absence. Our data indicate that heparin plays an essential role in rendering PF4 immunogenic by virtue of its ability to aggregate this positively charged protein to form a particulate antigen. Our previous studies have shown that heparin forms ULCs with PF4 and other positively charged proteins in a heparin-dependent manner [16, 20], and that complexes formed at distinct stoichiometric ratios of PF4:heparin are far more immunogenic in mice than PF4 itself. In continuation of these studies, we now show that this aggregating property of heparin or LMWH promotes uptake of positively charged proteins by antigen-presenting cells. In these studies, although PF4 forms antigenic complexes with cellular GAGs [2, 19], we show that heparin augments cellular internalization perhaps because it is more efficient at aggregating PF4 complexes. Using KKO, we note modest uptake of complexes when cells are incubated with PF4 alone, but marked increase in KKO binding when cells are pre-incubated with PF4 and heparin or LMWH (Figures 2 & 3B). While UFH and LMWH showed differences in complex formation and monocyte uptake, it should be noted that our studies were not designed to compare the immunogenicity of these two compounds, as the PHRs of circulating UFH or LMWH are not known. In other studies shown in Figure 3A, we show that cellular uptake is protein-independent, as complexes composed of heparin and other positively charged proteins (mPF4, protamine and lysozyme) are similarly ingested.

It is generally recognized that particulate antigens stimulate the immune response with far greater potency than soluble antigens [29], whether these are of infectious origin, occur as drug contaminants [30], or found in vaccine formulations [31]. Based on our findings of complex size (Supplemental Figure 1), cellular uptake (Figures 1 & 2) and heightened expression of MHCII and CD83 expression (Figure 6), we speculate that PF4/heparin complexes exert their immunogenicity as particulate antigen. Although the mechanisms of immunogenicity of particulate antigen are not well-understood and likely vary by antigen [30], one unifying biological feature of such antigens is their ability, relative to soluble antigen, to enhance endocytosis by antigen presenting cells (APC) [32–34]. For particulate antigen, size and surface charge of particles [35] are considered important biophysical features for cellular uptake. Whereas some studies have shown the importance of positive charge [35, 36], others have not [37]. In keeping with our previous findings [14], we note that charge plays a more prominent role than particle size in the immunogenicity of PF4/heparin, as complexes formed with greater amounts of PF4 relative to heparin (PHR 7>2), were more likely to be endocytosed. Consistent with published studies of the effects of cellular uptake, we confirm that increased endocytosis is associated with evidence of cellular activation leading to CD83 expression[35].

Based on our findings of ligand-independence and susceptibility to amiloride [21], it appears likely heparin-bound complexes enter into monocytes and DCs via macropinocytosis. Macropinocytosis is a non-saturable, constitutive mechanism by which macrophages and dendritic cells sample large volumes of extracellular fluid for foreign antigen [38]. After internalization, macropinosomes migrate centrally and fuse with endo-lysosomal compartments, where the fate of antigen cargo is determined [38, 39]. In these lysosomal compartments, membrane permeable antigens readily diffuse out, but non-permeable antigens are subject to varying extents of proteolytic degradation based on chemical and physical composition [39].

It has been the long-standing belief that complete lysosomal degradation promotes antigen presentation by generating short peptides essential for MHC class I or class II binding and transport to the cell-surface. However, recent studies indicate that antigens with limited susceptibility to proteolysis may be more immunogenic in vivo [40]. Using two model proteins and their isoforms, Delamarre and colleagues showed that proteins that were more resistant to proteolysis in DC lysosomes were more immunogenic in vivo [40]. Because unprocessed antigen taken up by macropinocytosis can be regurgitated into the extracellular environment, it is speculated that antigen resistance to proteolytic digestion increases immunogenicity by enabling antigen persistence in lymph and splenic microenvironments [41]. While additional studies are needed to confirm that our finding of intact PF4/heparin complexes in lysosomes (Figure 5) is due to relative proteolytic resistance, it is reasonable to speculate that structural preservation of complexes could lead to localized and persistent activation of specialized B-cell populations, such as marginal zone B-cells [9, 41].

At this time, our studies do not address the requirements for MHCII-restricted antigen presentation or other cellular contributions (T-cells and/or B-cells) in the immune pathogenesis of HIT. Nonetheless, our studies reconcile several important clinical observations that remain unexplained. Our model explains, in part, why heparin exposure is necessary to initiate disease and might not lead to clinical recurrence in settings where pathogenic PF4/heparin ratios are not attained during drug re-exposure. Although PF4 levels may be elevated in various disease states and may bind to cellular GAGs or bacteria, our studies show that heparin enhances the immunogenicity of PF4 by rendering it a particulate antigen. Our studies also help to explain why HIT occurs commonly in settings such as cardiopulmonary bypass [42, 43], where high levels of circulating PF4 and heparin result in larger (Supplemental Figure 1) and more abundant complexes. Finally, our model provides an explanation for the in vivo immunogenicity of other heparin-binding proteins, such as protamine [20, 44–46] or lysozyme [20] Thus, our in vitro and in vivo studies predict that any positively–charged proteins or PF4 complexed with other anionic molecules (e.g. RNA, DNA) capable of forming complexes are likely to be taken up by monocytes and have the potential to become immunogenic in vivo.

In summary, we show that heparin essentially facilitates uptake of PF4/heparin complexes by APCs through formation of particulate antigen/ULCs. Enhanced cellular uptake of complexes promotes cellular activation and likely contributes to in vivo immunogenicity of PF4/heparin complexes. Future therapeutic strategies targeting ULC formation [15] might be useful not only for the treatment of HIT and its clinical complications, but also in the prevention of HIT and its immune response.