Procoagulant and proinflammatory effects of red blood cells on lipopolysaccharide-stimulated monocytes

Summary.

Background:

We aimed to evaluate the mechanisms underlying the effects of red blood cells (RBCs) on the reactivity of monocytes to lipopolysaccharide (LPS) stimulation.

Methods:

Measurements of tissue factor (TF) antigen and activity were performed on freshly isolated white blood cells (WBCs)/platelets resuspended in heparinized plasma, as well as cultured monocytic cells.

Results:

In a dose-dependent manner, RBCs significantly enhanced LPS-induced TF activity and antigen levels in blood monocytes; potentiation of TF activity by both human and murine RBCs did not require the presence of neutrophils and/or platelets. We also measured the levels of monocyte chemotactic protein-1 (MCP-1), the key proinflammatory chemokine that binds to duffy antigen receptor for chemokines (DARC) on RBC surface, in plasma and RBC lysates after the incubation of RBCs with WBC/platelets; at the concentrations corresponding to normal blood counts, RBCs exerted a significant influence on the free plasma levels of MCP-1, with about two-thirds of detectable MCP-1 post-LPS stimulation being associated with RBCs. Critically, DARC-deficient murine RBCs failed to enhance LPS-induced TF activity, confirming the mechanistic significance of RBC-DARC.

Conclusions:

Our study reports a novel mechanism by which RBCs promote procoagulant and proinflammatory sequelae of WBC exposure to LPS, likely mediated by RBC-DARC in the microenvironment(s) that bring monocytes and RBCs in close proximity.

Introduction

Monocytes serve as the main cellular source of tissue factor (TF) in circulation [1]. Transcription factor Sp1 controls basal TF gene expression [2], and TF’s biosynthesis in monocytes is influenced by cell–cell interactions: it has been shown that lipopolysaccharide (LPS)-induced TF activity (TFa) in monocytes is enhanced by neutrophils in a platelet-dependent manner [3]. Screening of signaling pathway inhibitors in the LPS-stimulated blood model revealed that platelet-activating factor receptor and thromboxane A2 receptor are central in augmenting TF expression and activity [4].

Compared with other blood cells known to affect the expression of TF in monocytes, red blood cells (RBCs) are understudied. RBCs (erythrocytes), anuclear cytoplasts with few organelles, mainly serve as carriers of oxygen and as means to remove carbon dioxide and are not yet linked to inflammation, with the exception of sickle cell disease and polycythemia vera [5,6]. RBCs have a wide repertoire of surface receptors, including duffy antigen receptor for chemokines (DARC), a promiscuous receptor for chemokines that binds with various affinities (for a review, see [7]). DARC is present on the RBCs of most whites, whereas it is absent on RBCs of > 90% of African natives and ~65% of African Americans. Because RBC-DARC is primarily associated with Plasmodium falciparum infection, it was surprising that subjects lacking DARC on their RBCs exhibited markedly reduced chemokine levels and lower proinflammatory cytokine response post-LPS challenge [8]. These findings suggest that RBC-DARC may play a major role in modulating proinflammatory responses.

DARC acts as a receptor for monocyte chemotactic protein-1 (MCP-1, or CCL2), platelet factor 4 (PF4, or CXCL4), interleukin 8 (IL-8, or CXCL8), and neutrophil-activating peptide 2 (NAP-2, or CXCL7). All these chemokines induce proinflammatory and procoagulant responses. In a human endotoxemia model, Mayr et al. [9] showed that, compared with RBC-DARC⁺ subjects, RBC-DARC⁻ subjects have a markedly reduced TF-triggered activation of coagulation induced by LPS. Thus, we explored the effect of RBCs on the major procoagulant and proinflammatory entities such as TF and MCP-1, in two distinct ex vivo models.

Materials and methods

Chemicals, reagents, and mice

LPS (Escherichia coli O26:B6) was from Difco Laboratories (Detroit, MI, USA); Lymphoprep and Polymorph-prep were both from Axis-Shield (Oslo, Norway); and phorbol 12-myristate 13-acetate (PMA) was from Sigma-Aldrich (St Louis, MO, USA). The 6- to 8-week-old wild-type (WT) C57BL/6 mice or DARC-null mice on C57BL/6 background (strain 29873; MMRRC-UNC, Chapel Hill, NC, USA) were used to collect RBCs via intracardiac blood draw.

TF mRNA measurements

RNA was isolated from THP-1 cells by using the RNeasy kit (Qiagen, Valencia, CA, USA). cDNA was synthesized by using Transcriptor Reverse Transcriptase (Roche, Indianapolis, IN, USA), and quantitative reverse transcription–polymerase chain reaction was performed by using the previously validated TaqMan probe/primer set for human full-length TF [10]. Total TF antigen in THP-1 cells was quantified by using a highly sensitive ELISA (Zymutest RK042A; Hyphen BioMed, Neuville-sur-Oise, France).

Cell culture maintenance

Murine monocyte/macrophage cell line J774A.1 (ATCC, Manassas, VA, USA) was maintained in DMEM (10–013-CV; Cellgro, Manassas, VA, USA) supplemented with 10% fetal bovine serum; human monocytic cell line THP-1 (ATCC) was maintained in RPMI (10–040-CV; Cellgro) supplemented with 10% fetal bovine serum (35–010-CV; Cellgro) and standard antibiotics, antimycotics, and β-mercaptoethanol (30–004-CI; Cellgro). For CCR5 antagonist studies, THP-1 cells were incubated overnight with CCR5-specific inhibitor Maraviroc (10 μmol L⁻¹, PZ0002; Sigma Aldrich).

Blood collection, cell isolation, and stimulation with LPS

Venous blood was drawn from healthy consenting donors into a plastic syringe with a 19-gauge needle and transferred to polycarbonate tubes containing Fragmin (10 IE mL⁻¹; Pharmacia, Uppsala, Sweden); the study was approved by the Institutional Ethics Committee. Cells were counted with use of an Sysmex K 1000 (TOA Medical Electronics, Kobe, Japan). Blood cell isolation was carried out ≤ 1 h post-blood collection; RBCs, granulocytes, and mononuclear cells were isolated via centrifugation by applying 2 mL of whole blood on top of a two-layer density gradient consisting of 3.0 mL Lympho-prep on top of 3.0 mL Polymorphprep, as described previously [3]. RBCs were collected from the bottom and washed 3 times in 0.15 mol L⁻¹ NaCl. The quantities of RBCs added were from 25 to 400 μL of 10⁷ RBCs μL⁻¹, yielding the concentration range from 0.3 × 10⁹ to 4.4 × 10⁹ mL⁻¹ RBCs in the reactions. After washing, cells were resuspended in plasma containing Fragmin. Test aliquots were prepared by combining mononuclear cells, granulocytes, and platelets from 1 mL blood in 0.5 mL plasma (Fragmin), corrected for volume by 0.15 mol L⁻¹ NaCl if needed and RBCs in the above range. LPS was then added at 5 ng mL⁻¹ (with or without PMA), and incubation was carried out for 2 h at 37 °C in a rotary incubator and terminated by adding 0.1 mL 5.0 mmol L⁻¹ EDTA. Mononuclear cells were isolated, frozen, and thawed immediately before measurements of TFa as described later. In some experiments, incubated samples were centrifuged for the collection of plasma to quantify MCP-1 by using an ELISA (‘Ready-Set-Go’; eBioscience, San Diego, CA, USA); MCP-1 was also measured in RBC lysates.

TFa quantification

First, blood monocyte–associated TFa in reconstituted whole blood samples was quantified in a two-stage amidolytic assay based on the ability of human TF to accelerate the activation of human factor X (FX) by FVIIa, followed by the FXa conversion of prothrombin to thrombin in the presence of human FVa [11]; thrombin was quantified using Th-1 substrate, and color generated was determined at 405 nm by using a ThermoMax micro-plate reader (Molecular Devices, Silicon Valley, CA, USA). Crude rabbit brain extract was used as a standard for TFa, with undiluted activity assigned at 1 U mL⁻¹. Second, monocytic cell–associated TFa was also tested in a two-stage assay whereby the rate of FX-to-FXa conversion by the TF–FVIIa complex is assessed directly without the prothrombinase stage; we previously described this assay and showed its utility in measuring low-level TFa in platelet-free plasma [12]. In brief, fresh whole blood was collected from healthy donors with sodium citrate anticoagulant and centrifuged at 2500 × g for 15 min. Plasma and buffy coat were removed, and the remaining RBCs were washed 3× with PBS. RBCs were incubated with THP-1 or J7741.A cells, after which RBCs were lysed using RBC lysis buffer (154 mmol L⁻¹ NH₄Cl, 0.127 mmol L⁻¹ EDTA, 11.9 mmol L⁻¹ NaHCO₃), and THP-1/J7741.A cells were washed until no visible remnants of hemoglobin were observed. Cells were resus-pended in a solution buffered by HEPES and combined with 10 nmol L⁻¹ human FVIIa, 150 nmol L⁻¹ human FX (both from Enzyme Research Laboratories, South Bend, IN, USA), and 5 mmol L⁻¹ CaCl₂ in the wells of a 96-well plate; after 15 min of incubation, samples were transferred to another 96-well plate containing EDTABicine buffer, which terminates the production of FXa [12]. Chromogenic substrate Pefachrome Xa (Enzyme Research Laboratories) was then added to the wells, and the increase in optical density at 405 nm was captured every 5 min over a 1-h period on a kinetic plate reader (Molecular Devices) prewarmed to 37 °C. Optical densities at 405 nm rose linearly with time, and 1-h readouts/end point data were used for statistical analyses. Human FVIIa and FX supplied by Enzyme Research Laboratories were previously shown to be suitable in assessing human and murine TFa with high specificity [13]. All experiments were run in triplicate with ‘no LPS/+RBC’ controls to exclude hemoglobin autofluorescence; lipi-dated recombinant human TF (RT-0300; Haematologic Technologies, Inc., Essex Junction, VT, USA) was used to generate a standard curve.

Model of endotoxemia

C57BL/6 WT and DARC-null mice were injected intraperitoneally with LPS (5 mg/kg, Sigma Aldrich E. coli O111:B4) or saline, and after 5 h blood was collected from inferior vena cava into 3.5% sodium citrate–coated syringes. Whole blood was centrifuged (4000 × g for 15 min), plasma was separated, and RBCs were lysed with RBC lysis buffer. Leukocytes were dual-labeled with a goat anti-mouse TF polyclonal antibody (AF3178; R&D Systems, Minneapolis, MN, USA)/Alexa Fluor 594–conjugated rat anti-mouse antibody (Life Technologies, Grand Island, NY, USA) and anti-mouse CD11b-FITC (60001; Stemcell Technologies, Vancouver, BC, Canada) and analyzed by flow cytometry on an LSR-Fortessa (BD Biosciences, San Jose, CA, USA). Forward scatter and side scatter profiles were used to gate cells for size and exclude cell debris from data analysis.

Statistics

Statistical analysis was performed with SigmaPlot 12 (Systat, Chicago, IL, USA); all results are reported as mean ± SD. For two-group comparisons of parametric data, we used a two-tailed Student’s t-test, while non-parametric data was analyzed with Mann–Whitney rank sum. Multiple groups were assessed using one-way ANOVA on ranks, one-way ANOVA, or two-way ANOVA, when appropriate. For correlation analyses, Pearson’s coefficients were derived, and P ≤ 0.05 was considered significant.

Results

RBCs enhance LPS-induced TFa and TF protein levels in blood monocytes

In earlier studies of TFa levels in LPS-stimulated blood of healthy individuals, we reported a connection between family history of myocardial infarction and hyperreactivity of blood monocytes in response to low-dose LPS [14,15]. We have now analyzed RBC counts and monocyte TFa in healthy subjects (n = 260) and determined that there exists a weak yet highly significant positive correlation between RBC counts and monocyte TFa (r = 0.199, P = 0.001, Fig. 1A), raising the possibility that RBCs may mechanistically contribute to LPS-induced prothrombotic state in systemic inflammation. LPS-induced TFa in blood monocytes varies between individuals [15]; thus, we collected blood from several individuals to ensure the subject-unbiased nature of the changes in blood monocyte TFa. As shown in Fig. 1B, RBCs enhanced LPS-induced TFa in a dose-dependent manner with a significant increase beginning at as low as 0.3 × 10⁹ mL⁻¹ RBCs, plateauing at 2.2–4.4 × 10⁹ mL⁻¹ RBCs. To examine whether this increase in TFa is solely due to RBCs supplying PS-positive surfaces for prothrombinase complexes [16], we also quantified blood monocyte–associated TF protein by ELISA (IMUBIND TF; American Diagnostica, Greenwich, CT, USA). As shown in Fig. 1C, the trend of TF protein increase reached significance at 1.1 × 10⁹ mL⁻¹ RBCs per sample. Because in the absence of RBCs PMA is known to (i) amplify LPS-induced TFa in blood monocytes and (ii) activate platelets and neutrophils – the two cell types that influence TFa in monocytes [3] – we checked whether PMA has an impact on the effect of RBCs on LPS-induced TFa; the addition of 5 ng mL⁻¹ PMA in combination with LPS did not affect the potentiation of TFa by RBCs (data not shown).

Fig. 1.

(A) Correlation between red blood cell (RBC) counts and tissue factor (TF) activity (TFa) in blood monocytes of healthy subjects (n = 260). (B) RBCs enhance lipopolysaccharide (LPS)-induced TFa in blood monocytes. Isolated RBCs were added at various concentrations to mononuclear cells recombined with neutrophils and platelets in plasma anticoagulated with Fragmin and incubated with lipopolysaccharide (LPS) (5 ng mL⁻¹) for 2 h at 37 °C in a rotary shaker. After incubation, mononuclear cells were isolated, washed, frozen, and tested for TFa after thawing. The baseline TFa value was 49 ± 9.6 mU/1 million cells; n = 3. (C) RBCs increase LPS-induced TF protein levels in monocytes (test system is the same as in B); the baseline TF antigen value was 307 ± 134 pg mL⁻¹; n = 3. *P < 0.05.

Neutrophils and/or platelets are not required for RBC-induced potentiation of TFa

We next examined whether the effect of RBCs on monocyte-associated TFa observed in the whole blood system can also be observed in a system without plasma, neutrophils, and/or platelets and, if so, whether superpotentiation of F3 expression is responsible for this phenomenon. A time course of TF mRNA expression has yielded a classic acute-response pattern whereby the maximal induction of TF mRNA by LPS was reached by 2 h and was largely gone at 4 h. RBCs significantly increased TF mRNA levels at 2 h post-LPS exposure (Fig 2A). As could be expected from the mRNA data, TF protein levels peaked at 4 h with RBCs significantly superinducing TF protein levels at 2 and 4 h (Fig. 2B); for all subsequent studies, 4-h incubation was used. The TFa results we obtained using our FXa generation assay (Fig. 2C) were qualitatively analogous to those shown in Fig 1; moreover, in the absence of LPS, RBCs did not affect TFa in monocytic cells (Fig. 2A–C), and preincubation of cell lysates with inhibitory anti-TF antibody 7G11 [17] completely abolished FXa generation (Fig. 2C), confirming that our readout was highly specific for TF.

Fig. 2.

Effects of lipopolysaccharide (LPS) and red blood cell (RBC) stimulation on tissue factor (TF) mRNA (A), total TF antigen (B), and TF activity (TFa) (C) levels in human monocytic cells (THP-1). Cells were stimulated with LPS for 4 h at 37 °C with or without human RBCs with the final RBC concentration-per-well equal to 4.4 billion mL⁻¹. THP-1 cell lysates were assessed for TFa (C) using a two-step FXa generation assay. n = 3 (A–C); *P < 0.05.

LPS-exposed murine monocytes respond to murine RBC in a DARC-dependent manner

Next, we examined whether our findings are unique to human monocytes and/or RBCs. As shown in Fig. 3A, murine RBCs significantly potentiated TFa in cultured murine monocytes/macrophages at a concentration as low as 1.1 × 10⁹ mL⁻¹; again, this effect of RBCs was contingent on LPS stimulation. To examine whether the observed effect of RBCs is dependent on the presence of RBC-DARC, we compared the effects of WT RBCs and DARC-null RBCs on LPS-elicited TFa. As shown in Fig. 3B, DARC-null RBCs failed to potentiate murine monocyte TFa triggered by LPS, suggesting that RBCs likely exert their effect via DARC-bound chemokines, of which MCP-1 is a substantial constituent [7]. We then examined the contribution of DARC to the overall TF response of blood monocytes under severe inflammatory conditions: in a model of LPS-induced endotoxemia, DARC-null mice exhibited a highly significant, ~30% decrease in the induction of TF-positive monocytic cells compared with WT mice (Fig. 3C).

Fig. 3.

(A) Effect of lipopolysaccharide (LPS) and red blood cell (RBC) stimulation on tissue factor (TF) activity (TFa) in murine monocytes/macrophages (J774A.1). Cells were stimulated with LPS for 4 h at 37 °C with or without murine RBCs with the final RBC concentration per well equal to 4.4 billion mL⁻¹; J774A.1 lysates were assessed for TFa using a two-step FXa generation assay; n = 3. (B) Duffy antigen receptor for chemokines (DARC)-null RBCs have no effect on murine monocyte TFa induced by LPS stimulation; experimental conditions are the same as in A; n = 3. (C) Blood monocytes (CD11b⁺) in DARC-null mice exhibit a significantly dampened response in TF upregulation (CD11b⁺ TF⁺) on intraperitoneal LPS injection (endotoxemia model, see Materials and methods for experimental details); n = 3–6 mice per cohort. *P < 0.05; **P < 0.005; ***P < 0.001.

RBCs modulate the distribution of LPS-induced MCP-1

MCP-1 binds to RBC-DARC [7] and, because MCP-1 is able to raise TFa and protein levels in THP-1 cells via Ca²⁺ mobilization and protein kinase C activation[18], we hypothesized that MCP-1 bound to RBCDARC may account for the increase in blood monocyte–associated TF antigen/TFa as RBCs come in contact and interact with blood monocytes. MCP-1 was measured in plasma after centrifugation of the LPS-stimulated white blood cells (WBCs) and platelets, as well as in lysates of isolated RBCs. As shown in Fig 4A, MCP-1 levels in the RBC lysates rose substantially and correlated with the amount of added RBCs to the incubation mixture of WBCs/platelets; at higher RBC concentrations, the majority of detectable MCP-1 redistributed from plasma to RBCs, reaching about two-thirds of total MCP-1 in the system at 4.4 × 10⁹ mL⁻¹ RBCs. We then examined whether CCR5, a major receptor for MCP-1 expressed on monocytes, plays a significant role in the observed effects of RBCs: as shown in Fig. 4B, preincubation of THP-1 cells with a specific CCR5 inhibitor completely eliminated TFa potentiation by RBCs.

Fig. 4.

(A) Total amount of monocyte chemotactic protein-1 (MCP-1) in plasma and red blood cell (RBC) lysates after lipopolysaccharide (LPS) stimulation of isolated mononuclear cells combined with neutrophils and platelets and various concentrations of RBCs. Details of the test system is the same as in Fig. 1; n = 3. (B), Inhibition of CCR5 eliminates RBC-mediated increase of TFa expressed by LPS-stimulated THP-1 cells; MVC, Maraviroc; 7G11, inhibitory anti-TF antibody (ref. 17); n = 3. *P < 0.05; i.t., in trend (0.1 > P > 0.05). TF, tissue factor; TFa, TF activity.

Discussion

We demonstrate that RBCs, in a dose-dependent manner, amplify the LPS-induced TFa in human monocytes in (i) a cell model based on recombining WBCs and platelets in Fragmin-anticoagulated plasma and (ii) monocytic cells not exposed to blood cell types other than RBCs (Figs. 1–2). This effect is also seen when murine cells are used (Fig. 3), indicating that this phenomenon should be taken into consideration when interpreting results obtained in murine models of thrombosis tied to acute and/or chronic inflammation, especially those with a known or suspected bacterial component.

We previously showed that neutrophils, in a platelet-dependent interaction, enhance LPS-induced TFa in monocytes, particularly when PMA was added with LPS as stimuli [3]. However, this interaction appeared only to be associated with TF, as there was no effect on cytokine production. Here, we report that RBCs play a major role in subcompartmentalization of MCP-1 produced by blood monocytes in response to LPS, which likely accounts for the observed increase in monocyte TF protein when RBCs are present at the concentration corresponding to that of normal RBC counts (Figs. 1C, 4A). This suggests a specific interaction between the RBCs and monocytes, and one may conclude that the mechanism of this interaction differs from the neutrophil–platelet interaction with monocytes, because the absence of platelets and neutrophils had no effect on the RBC-induced upregulation of the monocyte TFa (Figs. 2C, 3A). Our results obtained with DARC-null RBCs (Fig. 3B), while completely novel, are in agreement with previous reports on RBC-DARC and its effect on TF-triggered coagulation. After infusion of low-dose LPS in healthy RBC-DARC⁺ and RBCDARC⁻ subjects, RBC-DARC⁻ subjects exhibited a markedly reduced procoagulant response [9]. Accordingly, LPS increased microparticle-associated TFa less potently in RBC-DARC⁺ subjects compared with RBC-DARC⁺ subjects; prothrombotic state was also more pronounced in RBC-DARC⁺ subjects, as were the MCP-1 levels, and it was suggested that, in the local chemokine milieu, MCP-1 may influence the levels of microparticle-associated TF [9,19]. Interestingly, Mayr et al. showed that infusion of MCP-1 into healthy volunteers markedly enhanced circulating monocytes yet it did not induce coagulation [20], suggesting that biologically significant levels of LPS in blood is a required constituent for the effect of RBCs. In that light, we note that circulating human monocytes can be chronically activated by bacterial endotoxins due to low-grade, proinflammatory conditions associated with a compromised blood–tissue barrier, such as in periodontitis, which is known to exacerbate atherosclerosis [21].

Platelet factor 4 (PF4), which is released quite instantly in blood incubated with LPS, plays an important role in TF expression in LPS-stimulated monocytes [11]. PF4 upregulates LPS-induced TFa in a granulocyte-dependent reaction where P-selectin plays a major role. However, we failed to see any contribution by neutrophils and platelets in our cell-based systems. It is possible that traces of platelets in mononuclear cell preparations may have provided PF4 to at least partially account for the RBC effect described in the present study. In this context, we note that PF4 and DARC are both required for platelet-mediated killing of P. falciparum [22]; however, molecular mediators involved in this phenomenon are unknown. We also demonstrate that, at low concentrations, RBCs do not significantly impact LPS-induced MCP-1 in the plasma, but at normal hematocrit levels, the majority of MCP-1 protein produced by blood monocytes in response to LPS is associated with the RBCs. Thus, it is reasonable to propose that systemically, LPS-stimulated monocytes may affect the circulating levels of MCP-1 and/or other proinflammatory markers produced by other blood and vascular cells, in an RBC-DARC–dependent manner. Further mechanistic studies will address whether a physical interaction of RBCs and monocytes is required for the RBC-DARC/monocyte-CCR5–dependent potentiation of LPS-induced TF expression on blood monocytes reported here.

In conclusion, our results support the notion that RBCs likely mediate major proinflammatory and procoagulant events in circulation, and further studies examining this phenomenon are thus highly warranted.