|
|||||||||
|
|
|
|
|
|
|
|
|
|
|
Blood, 15 June 2007, Vol. 109, No. 12, pp. 5251-5259. Prepublished online as a Blood First Edition Paper on March 1, 2007; DOI 10.1182/blood-2006-10-051334.
HEMOSTASIS, THROMBOSIS, AND VASCULAR BIOLOGY Regulation of macrophage procoagulant responses by the tissue factor cytoplasmic domain in endotoxemia1 The Rockefeller University, New York, NY; 2 Department of Immunology, Scripps Research Institute, La Jolla, CA; 3 Department of Biochemistry, University of Illinois, Champaign-Urbana, IL
Tissue factor (TF) is the primary initiator of coagulation, and the TF pathway mediates signaling through protease-activated receptors (PARs). In sepsis, TF is up-regulated as part of the proinflammatory response in lipopolysaccharide (LPS)–stimulated monocytes leading to systemic coagulation activation. Here we demonstrate that TF cytoplasmic domain–deleted (TF  CT) mice show enhanced and prolonged systemic coagulation activation relative to wild-type upon LPS challenge. However, TFCT mice resolve inflammation earlier and are protected from lethality independent of changes in coagulation. Macrophages from LPS-challenged TFCT mice or LPS-stimulated, in vitro–differentiated bone marrow–derived macrophages show increased TF mRNA and functional activity relative to wild-type, identifying up-regulation of macrophage TF expression as a possible cause for the increase in coagulation of TFCT mice. Increased TF expression of TFCT macrophages does not require PAR2 and is specific for toll-like receptor, but not interferon  receptor, signaling. The presence of the TF cytoplasmic domain suppresses ERK1/2 phosphorylation that is reversed by p38 inhibition leading to enhanced TF expression specifically in wild-type but not TFCT mice. The present study demonstrates a new role of the TF cytoplasmic domain in an autoregulatory pathway that controls LPS-induced TF expression in macrophages and procoagulant responses in endotoxemia.
Disseminated intravascular coagulation driven by tissue factor (TF) is associated with systemic inflammatory disorders, including viral hemorrhagic fevers and severe sepsis (for reviews, see Riewald and Ruf1 and Ruf2). Inhibition of the TF pathway by potent TF-directed inhibitors reduced inflammatory cytokine levels and increased survival in lethal models of bacterial sepsis in primates3–5 and attenuated acute lung injury.6 Although coagulation blockade is likely to be beneficial in sepsis by reducing microthrombosis, subsequent hypoxic organ dysfunction, and inflammation, blocking thrombin generation per se has poor efficacy in prolonging survival in primate models.7,8 Therefore, it remains of interest to understand how signaling of the TF pathway makes contributions to sepsis mortality. Genetic reduction of TF or VIIa levels in mice increased survival in endotoxemia, but protease activated receptor 1 (PAR1) or PAR2 deficiency alone or in combination did not reduce lethality.9–11 Whether the TF pathway contributes to severe systemic inflammation through signaling thus remains incompletely understood.
In the response to microbial pathogen challenge or lipopolysaccharide (LPS)–induced toll-like receptor 4 (TLR4) activation, cells of the monocyte/macrophage lineage are the primary cell types that up-regulate TF to trigger intravascular coagulation.12–15 However, the subsequent crosstalk between coagulation activation and inflammation is complex. Thrombin and PAR signaling may amplify the coagulant response by further inducing TF in endothelial cells16–18 or attenuate coagulation and inflammation through the activated protein C (PC)/endothelial cell PC receptor (EPCR) pathway that uses PARs as signaling receptors.19–22 In addition, endotoxin challenge of TF cytoplasmic domain–deleted (TF
Here, we identify a novel role of the TF cytoplasmic domain in regulating the procoagulant response induced by TLR signaling in macrophages. LPS activation of macrophages is mediated by a receptor complex of CD14 and TLR4.27 The signaling pathways by which LPS induces TF mRNA expression in monocytes have been studied in detail,28 but maturation of monocytes into macrophages promotes diverse phenotypic changes, including the expression of TF29 as well as the TF-VIIa signaling receptor PAR2.30 We find in macrophages that LPS-mediated TF up-regulation is controlled by the TF cytoplasmic domain through p38-dependent regulation of the ERK1/2 MAP kinase pathway. However, regulation of TF expression is independent of PAR2 signaling and specific for TLR, but not interferon
Mouse strains
TF Mouse model of severe endotoxemia
All experiments were carried out in accordance with the National Institutes of Health Animal Protection Guidelines and were approved by The Scripps Research Institute Animal Care and Use Committee. Lipopolysaccharide (LPS, E coli O111:B4; Calbiochem, La Jolla, CA) was dissolved in 37°C normal saline by vortexing for 30 seconds before and after 2 minutes of sonication. Mice between 60 and 80 days of age were intraperitoneally injected under 2% isoflurane with 5 to 9 mg/kg LPS. For each batch of endotoxin, an appropriate dose was established that resulted in 75% to 90% lethality. Because of reported10 sex differences in the dose response to endotoxin, we used male mice only in order to reduce variability. However, the phenotype of TF Two basic experimental protocols were used. For evaluation of coagulation activation and cytokine levels, mice were challenged with LPS for 4, 6, 12, or 18 hours. Ten minutes prior to blood collection, mice were anticoagulated by intraperitoneal injection of 1000 IU heparin (Elkins-Sinn, Cherry Hill, NJ). Under 3% isoflurane anesthesia, blood was drawn from the inferior vena cava into a 3.2% solution of sodium citrate and mice were killed to terminate the experiment. To establish the survival benefit of knock-out strains, mice were challenged with LPS and tightly monitored over the next 7 days without further manipulations, including blood drawing which may have a negative impact on the recovery of animals. Coagulation inhibitors were applied by intravenous tail vein injection at the time of LPS administration. TF expression in peritoneal macrophages after LPS stimulation Mouse peritoneal macrophages were harvested 6 hours after intraperitoneal injection of 5 mg/kg LPS. After killing, peritoneal lavage was performed to harvest resident cells by injecting 3 times 3 to 5 mL sterile cold PBS and recovering the wash through a small abdominal incision. Contaminating red blood cells were lysed with hypotonic buffer and cells were counted. Flow cytometry was used to quantify TF expression on macrophages by gating on CD11b-PerCP and F4/80-PE (Pharmingen, San Diego CA) double-positive cells and detection with fluorescein isothiocyanate (FITC)-conjugated antimouse TF antibody. TF mRNA expression in lungs 6 hours after LPS or sham injection into mice was quantified by TaqMan real-time polymerase chain reaction (PCR) on an ABI PRISM detection system (Applied Biosystems, Foster City, CA). Measurements of thrombin-antithrombin complex (TAT), cytokine levels, and blood counts Citrated blood drawn from the vena cava inferior was immediately centrifuged at 2000g for 10 minutes. Plasma was separated, aliquoted, and stored at –80°C for subsequent determination of TAT and cytokine levels. Enzyme-linked immunosorbent assays (ELISAs) were used to determine TAT (Enzygnost; Dade Behring, Marburg, Germany), IL6, IL1ß, and MCP1 (Quantikine; R&D Systems, Minneapolis, MN) levels. Cytokine profiles were determined by Luminex multiplex analysis (Invitrogen, Carlsbad, CA). For blood counts, 300 µL blood was drawn into EDTA-containing microtainer tubes (Becton Dickinson, Franklin Lakes, NJ) from the inferior vena cava 6 hours after LPS injection. White blood cells, hemoglobin, hematocrit, and platelets were measured on an ADVIA 120 Hematology Analyzer (Bayer, New York, NY) and compared to established reference ranges for mice. Statistical analysis Six to 10 mice per genotype were injected in a block randomized manner and experiments were typically repeated 2 to 4 times. All data are presented as means and standard deviations (mean ± SD), and differences were considered significant at P less than .05. Kaplan-Meier curves were plotted and survival differences were analyzed using a log-rank test. Statistical analysis of continuous data was performed for 2 group comparison by the Mann-Whitney U test. Culture of bone marrow–derived macrophages (BMMs) BMMs were in vitro–differentiated using endotoxin-free cell culture reagents according to published protocols.33 Briefly, bone marrow cells were obtained by flushing femurs and tibias with RPMI-1640 medium. Following washing with RBC lysis buffer (Sigma, St Louis, MO), cells were plated into Petri dishes containing RPMI-1640 medium supplemented with 20% to 30% L929 cell–conditioned medium, 10% heat-inactivated FBS, 100 units/mL penicillin, 1000 units/mL streptomycin, and 20 mM L-glutamine and maintained in a 8% CO2 atmosphere. Replacing L929 cell–conditioned medium with recombinant MCSF yielded macrophages with similar properties in the TF-dependent signaling. At day 7, the adherent cellular monolayer was used for experiments. Morphologic and fluorescence-activated cell sorter (FACS) analyses showed that 98% to 100% of cells were macrophages. For analysis of expression of activation markers, BMMs were left unstimulated or stimulated with LPS (1 µg/mL) for 18 hours. Cells were harvested, washed in FACS buffer (PBS, 2% FBS, 2 mM EDTA), and resuspended in FACS staining buffer with Fc-block (Pharmingen) for 10 minutes at 4°C. Cells were then incubated for an additional 30 minutes with fluorochrome-conjugated antibodies at 4°C in the dark. After washing in PBS, cells were analyzed by flow cytometry. Antibodies to CD11b, CD80, CD86, and CD69b were from Pharmingen and anti-TLR4 and F4/80 from eBioscience (San Diego, CA). Characterization of BMMs
BMMs (2 x 105 cells/mL) were plated after 7 days of differentiation and allowed to adhere overnight in L929 cell–conditioned medium. Controls and cells stimulated with LPS (1 µg/mL) at 37°C for the indicated times were rinsed twice with cold PBS and lysed in SDS sample buffer for Western blotting using antiphospho or panspecific ERK1/2 and p38 antibodies (Cell Signaling Technologies, Beverly MA). Fc TF assays
BMMs were stimulated either with LPS (1 µg/mL) or IFN
TF cytoplasmic domain–deleted mice display increased coagulation activation but are protected from lethality independent of altered coagulation We addressed the role of the TF cytoplasmic domain in endotoxemia in a homogenous C57BL/6 genetic background. We established an appropriate endotoxin dose (5-8 mg/kg, depending on individual LPS batches) to yield 75% to 90% lethality in male mice. With the preparations of LPS used, this target lethality was typically achieved at a dose approximately 1.6-fold higher than an LD50, indicating a steep dose-response curve. Lethality in this model occurred 1 to 3 days after endotoxin challenge and all mice that lived for 7 days were considered long-term survivors. At this time, surviving mice had returned to normal feeding behavior, appeared healthy, and were active.
In this model, mice challenged for 6 hours develop an overall leukopenia with increased neutrophil and decreased lymphocyte counts, signs of hemoconcentration, and reduced platelet counts, as also described by Camerer et al.10 No differences in leukocyte or platelet counts were detected between TF
In order to monitor the inflammatory and coagulant response under our challenge conditions, we obtained plasma samples from separate cohorts by terminal bleeding after 4, 6, 12, or 18 hours for determination of thrombin-antithrombin complex (TAT) levels and of circulating cytokines IL6, IL1ß, and MCP1. TF CT mice showed significantly increased coagulation activation, measured as TAT levels, until 12 hours after LPS challenge relative to wild-type mice (Figure 1B). The increase in coagulation activation was accompanied by cytokine production that was indistinguishable from wild-type animals at 6 hours, indicating that LPS-induced inflammatory cytokine elaboration was not regulated by the TF cytoplasmic domain during this period (Figure 1C). At 12 hours, inflammation markers of TFCT mice began to deviate from wild-type controls. After 18 hours, TAT levels were similar between TFCT and wild-type mice. However, levels of all inflammatory cytokines were significantly reduced in TFCT mice, indicating that despite similar coagulation activation, TFCT mice can more readily resolve the systemic inflammatory response.
In order to demonstrate that the measured cytokines are representative of the early inflammatory response of TF
Figure 2B documents that the attenuation of coagulation activation by both inhibitors was efficient for at least 6 hours. Consistent with the study by Camerer et al,10 blocking early coagulation activation by different inhibitors had no effect on the long-term survival of wild-type mice in our model (data not shown). Increased coagulation activation in TF CT mice may have induced antiapoptotic or barrier protective pathways19,22,34,35 without changes in inflammatory parameters. In order to address whether early coagulation activation protected TFCT mice, we administered the potent Xa inhibitor NAP5 at the time of LPS challenge and monitored the survival of these animals versus wild-type and TFCT controls (Figure 2D). Attenuation of early coagulation activation had no effect on the survival of TFCT mice, suggesting that these mice are protected by alterations in direct TF-dependent signaling, rather than changes in coagulation activation.
Peritoneal TF
Circulating monocytes are critical in the initial inflammatory response to endotoxin. Because monocytes express TF only after LPS induction, we reasoned that the TF cytoplasmic domain cannot play a regulatory role in monocytes. Consistently, all early inflammatory markers were found to be unchanged and TF
Maturation of monocytes into macrophages results in TF expression, but macrophages remain sensitive to LPS stimulation by further up-regulation of TF.29 To test whether macrophages potentially contribute to systemic coagulation activation, we designed an in vivo experiment where resident macrophages were stimulated with LPS for 6 hours and TF expression was evaluated on CD11b and F4/80 double-positive cells isolated from the peritoneal cavity. LPS stimulation increased TF expression on peritoneal macrophages from basal levels by approximately 2-fold in wild-type mice. TF
TF
In order to distinguish between direct effects of the TF cytoplasmic domain in macrophages versus indirect effects of TF cytoplasmic domain signaling originating from other cell types in vivo, we further characterized TF induction in bone marrow–derived macrophages. Bone marrow from wild-type, TF
TF CT BMMs show increased LPS-induced TF expression
TF expression was analyzed after 6 hours of LPS stimulation by measuring TF clotting activity and VIIa-dependent Xa generation, as well as by determining TF mRNA levels by quantitative real-time PCR. Baseline TF activity was similar in all genotypes, but BMMs from TF
The TF cytoplasmic domain suppresses TLR4-mediated ERK1/2 activation dependent on p38 activation
Mitogen-activated protein kinase (MAPK) pathways downstream of TLR4 signaling are required for TF induction.39 We analyzed ERK1/2 and p38 phosphorylation in LPS-stimulated BMMs from wild-type and TF
Pharmacological inhibition of ERK1/2 activation or PI3 kinase by U0126 or LY292004, respectively, prevented TF induction in BMMs of both wild-type and TF CT mice (Figure 6B). The PI3 kinase inhibitor LY292004 also blocked LPS-induced ERK1/2 phosphorylation (Figure 6A). Thus, TF induction is ERK1/2 dependent and PI3 kinase is upstream of ERK1/2 signaling. Of interest, blocking p38 with SB203580 increased ERK1/2 phosphorylation (Figure 6A) in wild-type mice, but had no effect on ERK1/2 phosphorylation in TFCT mice, indicating that the TF cytoplasmic domain may act as a scaffold to promote p38-dependent suppression of ERK1/2 phosphorylation.
Macrophage differentiation was carried out in conditioned medium from L929 cells that may contain factors that were responsible for the observed phenotypic changes of TF CT macrophages. To exclude this possibility, we generated BMMs using recombinant macrophage colony-stimulating factor. Figure 6B shown that the p38-dependent, differential ERK1/2 phosphorylation between wild-type and TFCT macrophages was independent of an unidentified factor in the L929 cell–conditioned medium. Inhibition of p38 also increased TF induction in wild-type BMMs, but TF expression in TFCT BMMs was similar in the presence or absence of p38 inhibitor (Figure 6C). Taken together, these experiments identify p38-dependent regulation of ERK1/2 as the underlying mechanism by which the TF cytoplasmic domain specifically controls the expression levels of TF in LPS-stimulated macrophages.
These experiments provide new insight into functions of the TF cytoplasmic domain in the procoagulant response during endotoxemia in vivo. TLR4 engagement activates various members of the mitogen-activated protein kinase (MAPK) family, including ERK1/2, c-Jun amino terminal kinases (JNKs), and p38 in monocytes leading to TLR4-mediated induction of inflammatory mediators and TF. TF gene induction in monocytes has been extensively studied and shown to involve both NF-  B and MAP kinase pathways.28 Macrophages, unlike endothelial cells and monocytes, up-regulate TF during maturation and constitutively express TF.29 Although the TF cytoplasmic domain per se does not regulate the basal TF expression, it imposes regulatory control over TLR4-mediated, but not interferon -mediated, TF induction in macrophages. ERK1/2 phosphorylation induced by TLR4 was found to be entirely dependent on PI3 kinase activation in macrophages. In monocytes and endothelial cells, TF expression in response to endotoxin or angiogenic growth factors is known to be negatively regulated by PI3 kinase signaling.40,41 Our data do not exclude that PI3 kinase activation also regulates TF induction in macrophages. However, the requirement for PI3 kinase to activate ERK1/2 precludes further experiments to address this question, because in macrophages TF up-regulation is entirely dependent on ERK1/2 similar to other cell types. The presented results identify a regulatory pathway in which the TF cytoplasmic domain suppresses ERK1/2 phosphorylation through a p38 pathway. Negative regulation of ERK1/2 activation by p38 has also been documented for growth factor,42 G-protein–coupled receptor,43,44 cytokine receptor,45 and innate immune46 signaling.
Our data indicate that the TF cytoplasmic domain may act as a scaffold to orchestrate a crosstalk between p38 and ERK1/2 signaling. Ott et al have shown that ligation of TF can stimulate the small GTPase rac and p38 in a TF cytoplasmic domain–dependent manner in cancer cells.25 The observed migratory response in their experimental system may reflect cytosolic sequestration of ERK1/2 following simultaneous activation of p38 and ERK1/2. Consistently, TF Maturation of monocytes into macrophages induces diverse phenotypic changes, including the up-regulation of PAR2.30 Deletion of PAR2 showed that the TF cytoplasmic domain influences TLR4 signaling independent of PAR2, providing evidence for an autonomous signaling function of TF to suppress in a p38-dependent manner ERK1/2 activation and TF up-regulation. Negative regulatory roles of the TF cytoplasmic domain were also demonstrated in angiogenesis. Because diabetic neoangiogenesis showed colocalization of phosphorylated TF with PAR2,26 PAR2 signaling may serve as an upstream signal to turn off the regulatory control of TF through phosphorylation. Our data indicate that one immediate downstream effect of such loss of regulatory control in cells exposed to LPS is the up-regulation of TF, which conversely could amplify TF-dependent PAR2 signaling responses.
Loss of the TF cytoplasmic domain leads to an enhanced coagulopathy in a model of severe endotoxemia. Peritoneal macrophages from LPS-stimulated mice showed enhanced TF expression independent of PAR2 expression, validating in vivo findings with in vitro–differentiated, bone marrow–derived macrophages. Although coagulation activation was exacerbated in TF
Robust coagulation activation improves sepsis outcome in heterozygous factor V Leiden carriers,21 but not thrombomodulin mutant mice,48 implicating the protein C pathway. Although enhanced activation of the protein C pathway and resulting barrier and cytoprotective signaling may have contributed to the improved survival of TF
The presented in vitro and in vivo data identify macrophages as a critical cell type in which the TF cytoplasmic domain plays a regulatory role in TLR4-dependent TF induction. Whether macrophages are the sole cell type that causes the enhanced circulating TAT level in this model remains to be addressed. In particular, in late-stage endotoxemia, endothelial cells and other myeloid inflammatory cells in the spleen express TF,49 and these cells may contribute to the sustained coagulation activation at later stages when the cytokine profiles indicate that activation of the innate immune system is attenuated in TF
Contribution: J.A. designed and performed experiments, analyzed and interpreted data, and wrote the paper; F.N. performed research and analyzed data; T.K., Y.K.L., and G.B. performed research; J.H.M. provided vital reagents; W.R. supervised the project and wrote the paper. J.A. and F.N. contributed equally to this study. Conflict-of-interest disclosure: The authors declare no competing financial interests. Correspondence: Wolfram Ruf, Department of Immunology, SP258, The Scripps Research Institute, 10550 North Torrey Pines Road, La Jolla, CA 92037; e-mail: ruf{at}scripps.edu.
This study was supported by NIH grants HL77753 and HL78614 (W.R.), and stipend NI 540/2-1 from the Deutsche Forschungsgemeinschaft (F.N.).
We thank Cindi Biazak, Pablito Tejada, Jennifer Royce, and Rachael Deleers for their excellent technical assistance, and Drs Peter Carmeliet and Patricia Andrade-Gordon for providing TF
Submitted October 12, 2006; accepted February 27, 2007.
Prepublished online as Blood First Edition Paper, March 1, 2007
DOI: 10.1182/blood-2006-10-051334
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Top
Abstract
Introduction
, after 2 hours of stimulation (
3).
