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Originally published online as doi:10.1189/jlb.0607353 on January 7, 2008

Published online before print January 7, 2008
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(Journal of Leukocyte Biology. 2008;83:902-911.)
© 2008 by Society for Leukocyte Biology

The cytoplasmic domain of tissue factor in macrophages augments cutaneous delayed-type hypersensitivity

Jim Apostolopoulos1, Michael J. Hickey, Laveena Sharma, Piers Davenport, Leon Moussa, Will G. James, Julia L. Gregory, A. Richard Kitching, Ming Li and Peter G. Tipping

Centre for Inflammatory Diseases, Department of Medicine, Monash University, Clayton, Victoria, Australia

1 Correspondence: Department of Medicine, Level 5, Block E, Monash Medical Centre, 246 Clayton Rd., Clayton Victoria, 3168 Australia. E-mail: jim.apostolopoulos{at}med.monash.edu.au


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ABSTRACT
 
In addition to its procoagulant role, tissue factor (TF) has important coagulation-independent roles, including in inflammation. The cytoplasmic domain of TF has been implicated in some of these coagulation-independent roles, particularly cell signaling. To assess the contribution of the cytoplasmic domain of TF to cell-mediated adaptive immunity, the development of cutaneous delayed-type hypersensitivity (DTH) was studied in mice lacking the cytoplasmic domain of TF (TF{delta}CT/{delta}CT mice). DTH responses in sensitized mice were significantly attenuated in TF{delta}CT/{delta}CT mice, and leukocyte-endothelial cell interactions, assessed by intravital microscopy, were impaired significantly. Studies in chimeric mice, created by bone marrow transplantation, showed that the absence of the cytoplasmic domain of TF in leukocytes rather than endothelial cells was responsible for reduced DTH and leukocyte recruitment. DTH responses to OVA could be induced in wild-type mice but not in TF{delta}CT/{delta}CT mice by transfer of activated CD4+ OVA-specific TCR transgenic T cells, demonstrating that the defective DTH response in TF{delta}CT/{delta}CT mice was independent of any defect in T cell activation. Macrophage and neutrophil accumulation and expression of TNF-{alpha} mRNA and phospho-p38-MAPK were reduced significantly in TF{delta}CT/{delta}CT mice, and their macrophages had reduced P-selectin-binding capacity and reduced in vivo emigration in response to MCP-1. These results indicate that leukocyte expression of the cytoplasmic domain of TF contributes to antigen-specific cellular adaptive immune responses via effects on leukocyte recruitment and activation.

Key Words: DTH • coagulation • inflammation • intravital microscopy


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INTRODUCTION
 
Tissue factor (TF) is a single-chain transmembrane glycoprotein that has significant structural homology with type II cytokine receptors [1 ]. In addition to its role in hemostasis and thrombosis, TF has a number of physiological functions in embryogenesis [2 3 4 5 ], angiogenesis [6 ], tumor metastasis [7 ], cell adhesion [8 ], and inflammation [9 , 10 ]. The extracellular domain of TF is responsible for its coagulant activity, and other TF-dependent cellular responses, including Factor VIIa-induced stimulation of IL-1β mRNA expression in keratinocytes [11 ] and phosphorylation of STAT5 [12 ], are also dependent on the extracellular domain of TF. The function of the cytoplasmic domain of TF is less clear. Its potential involvement in cell signaling is suggested by the presence of two serine residues that can be phosphorylated [13 ]. The cytoplasmic domain of TF is required for Factor VIIa-induced cellular responses such as induction of Ca2+ fluxes in myeloid cell lines [14 ], activation of p38 MAPK [15 ], adhesion, migration and spreading of fibroblasts [8 ], smooth muscle migration associated with vascular remodeling following injury [16 ], metastatic tumor growth, and protease-activated receptor 2-dependent angiogenesis [17 ].

Endothelial cells and leukocytes, key cells in inflammatory responses, do not express TF under normal conditions [18 ]. However, in response to inflammatory stimuli including endotoxin [19 , 20 ] and proinflammatory cytokines [21 22 23 ], these cells express high levels of TF. TF expression, in association with macrophage accumulation and fibrin deposition, is a prominent feature of cell-mediated adaptive immune responses such as cutaneous delayed-type hypersensitivity (DTH) [24 ]. Anti-TF mAb treatment significantly reduces local fibrin deposition and skin swelling in DTH [24 ]. Monocyte TF expression correlates closely with T cell-dependent cellular immunity [25 ], and activated T cells and their products are potent inducers of TF expression on monocytes [26 , 27 ].

Several studies suggest that TF signaling can amplify innate inflammatory responses [14 , 28 ]. Proinflammatory functions of the cytoplasmic domain of TF have been demonstrated in a murine model of endotoxemia, in which TF{delta}CT/{delta}CT mice have enhanced survival and reduced serum levels of TNF-{alpha}, IL-1β, and IL-6 [28 ]. In our previous study [29 ], we reported a significant reduction of the severity of antigen-induced arthritis in TF{delta}CT/{delta}CT mice, which was associated with reduced synovial inflammation and proinflammatory cytokine expression. T cell-dependent, cutaneous DTH responses to the disease-inducing antigen were also decreased [29 ].

The aim of the current studies was to further examine the contribution of the cytoplasmic domain of TF to the development of cell-mediated adaptive immune responses by using a model of cutaneous DTH in TF{delta}CT/{delta}CT mice, which lack the 18 carboxyl-terminal amino acids of the cytoplasmic domain of TF but retain normal coagulation function [30 ]. The separate contributions of TF cytoplasmic domain-dependent effects in T cells, macrophages, and endothelial cells were studied. The results demonstrate that the cytoplasmic domain of TF contributes to cutaneous DTH responses by promoting local macrophage recruitment and augmenting TNF-{alpha} expression and p38-MAPK phosphorylation.


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MATERIALS AND METHODS
 
Animals
TF{delta}CT/{delta}CT mice (mixed MF1/129S/v/Swiss background) lacking the 18 carboxyl-terminal amino acids of the cytoplasmic domain of TF were generated by the Cre-lox recombination technique in mice described previously [30 ]. The TF{delta}CT/{delta}CT mice were then backcrossed for nine generations onto a C57/BL6 strain background and along with C57BL/6 [wild-type (WT)] mice, were used in all subsequent experiments, except where indicated. Mice were 8–10 weeks old for all experiments except for the bone marrow transplantation experiments, in which they were 13 weeks old at the time of induction of DTH. All animal experimental protocols were approved by the Monash University Animal Ethics Committee (Australia).

Cutaneous DTH
WT and TF{delta}CT/{delta}CT mice were initially sensitized to a single antigen by a s.c. injection containing 100 µg methylated BSA (mBSA), 100 µg OVA, or 1.6 µg purified protein derivative (PPD) dissolved in CFA (Sigma, Australia), followed by a single booster with the same antigen dissolved in IFA (Sigma) 7 days later. Cutaneous DTH was induced 14 days after the initial immunization by s.c. injection of antigen (10 µg mBSA, 50 µg OVA, or 0.8 µg PPD), dissolved in 20 µl normal saline into the right ear. An irrelevant antigen (casein), at the same concentration, was injected into the opposite (left) ear as a control. Ear thicknesses were measured using a micrometer (Mitutoyo, Kawasaki-shi, Japan) before and 24 h after antigen injection. The antigen-specific DTH response was determined by the difference in the skin-swelling responses between the antigen-challenged and control ear.

Histological assessment of DTH lesions
Skin from DTH lesions was fixed in Bouin’s solution and embedded in paraffin for fibrinogen staining by immunoperoxidase, as described previously [31 ]. Sections (5 µm) were incubated with rabbit anti-mouse fibrinogen antisera (diluted 1 in 10,000; a gift of Dr. Jay Degen, Cincinnati Children’s Hospital Medical Center, Cincinatti, OH, USA), followed by ABC and diaminobenzidine substrate (Sigma). Fibrinogen staining was scored semiquantitatively in a blinded protocol by rating the extent of staining between 0 and 5 (0, no staining, i.e., background; and 5, intense staining, i.e., the most extensive fibrin deposition). P-selectin expression in skin lesions was assessed in 6 µm sections of snap-frozen tissues. Sections were blocked with 10% sheep serum in 5% BSA/PBS and then incubated with polyclonal rabbit anti-human P-selectin (10 µg/ml, BD Biosciences, San Diego, CA, USA) that cross-reacts with murine P-selectin. The sections were then washed and incubated with FITC-conjugated sheep anti-rabbit IgG (Silenus, Hawthorn, Victoria, Australia) at a dilution of 1 in 50 or with HRP-conjugated sheep anti-rabbit IgG (Silenus) for immunohistological staining. P-selectin was scored semiquantitatively from 0 to 3+ (0, no staining, i.e., background; and 3+, intense fluorescence in any field) in a blinded protocol. Twenty fields within lesions from each mouse were scored and averaged. VCAM-1 and ICAM-1 expression was assessed on 5 µm frozen tissue sections fixed with periodate-lysine paraformaldehyde (PLP) solution. Sections were blocked with 10% normal rabbit serum/10% normal rat serum in 5% BSA/PBS for 1 h and then incubated with a rat anti-mouse ICAM-1 mAb [clone YN1/1.74, American Type Culture Collection (ATCC), Manassas, VA, USA], diluted 1:1000, or rat anti-mouse VCAM-1 mAb (clone 6C71, ATCC), diluted 1 in 250. Sections were washed in PBS, and endogenous peroxidase activity was blocked by incubation with methanol/0.3% H2O2 for 1 h at room temperature. Sections were then incubated with biotinylated rabbit anti-rat IgG (Dako, Carpinteria, CA, USA), which was diluted 1:1000, followed by ABC solution (Dako) for 30 min and 3'-diaminobenzidine tetrahydrochloride, 1 min. Sections were counterstained with hematoxylin, and ICAM-1 and VCAM-1 staining was scored (0–3) using a blinded protocol and semiquantitative scoring scale (0, no staining, i.e., background staining; and 3, intense staining). The expression of VCAM-1 and P-selectin was assessed on cultured lung microvascular endothelial cells isolated from WT and TF{delta}CT/{delta}CT mice by FACS analysis, as described previously [32 ].

Macrophages and neutrophils were counted in DTH lesions by fixing ears in PLP, sectioning, and staining with mAb FA/11 and Gr-1 (respectively) using a three-layer immunoperoxidase technique as described previously [33 ]. Positively stained cells in ear sections within lesions were counted using a graticule in a blinded protocol. Results are expressed as the total number of positively stained cells per mm2.

Assessment of leukocyte trafficking by intravital microscopy
Leukocyte trafficking following antigen challenge was assessed in the microcirculation of the mouse cremaster muscle. Animals were sensitized to OVA, as described above, and challenged 14 days later by intrascrotal injection of OVA (50 µg in 250 µl saline). Twenty-four hours later, animals were anesthetized by i.p injection of 10 mg/kg xylazine (Bayer Pharmaceuticals, Pymble, NSW, Australia) and 200 mg/kg ketamine hydrochloride (Caringbah, NSW, Australia), and the cremaster microvasculature was prepared for examination, as described previously [34 ]. The cremaster microcirculation was visualized using an intravital microscope (Axioplan 2 Imaging, Carl Zeiss Australia) and a color video camera (Sony SSC-DC50AP). The images were recorded using a videocassette recorder (Panasonic NV-HS950) as described previously [34 , 35 ]. Three to four postcapillary venules (25–40 µm in diameter) were examined in each experiment. Venular diameter and the number of rolling, adherent, and emigrated leukocytes were determined off-line during video playback analysis. Rolling leukocytes were defined as cells moving at a velocity less than that of erythrocytes within a given vessel. Leukocyte rolling velocity was determined by measuring the time required for a leukocyte to roll along a 100-µm length of venule and was determined for 20 leukocytes per vessel. Leukocytes were considered adherent to the venular endothelium if they remained stationary for 30 s or longer. The number of leukocytes recruited to the extravascular tissue was determined by counting the leukocytes present in the extravascular tissue within the field of view (shown as interstitial leukocyte accumulation). Three vessels per animal were analyzed for initial measurements of leukocyte rolling and other parameters. Leukocyte rolling, adhesion, and emigration in this model were P-selectin-dependent, as demonstrated by i.v. injection of rat anti-murine P-selectin mAb (RB40.34, BD Biosciences) at a dose shown previously to inhibit P-selectin function in vivo [35 , 36 ].

Assessment of ex vivo splenocyte responses to OVA
Spleens were harvested aseptically from WT and TF{delta}CT/{delta}CT mice sensitized to OVA, and single-cell suspensions were prepared in DMEM containing 5% FCS (Commonwealth Serum Laboratories, Parkville, Victoria, Australia). RBCs were lysed by incubation in 0.15 mol/L NH4Cl, 0.01 mol/L NaHCO3, 0.1 mmol/L EDTA, pH 7.3, for 1 min at 37°C. Total cell numbers were determined using a hemocytometer. OVA-sensitized splenocytes (1x105/well) were cultured for 72 h in round-bottom 96-well plates with OVA at concentrations of 0–100 µg/ml. The proliferative responses were determined by the incorporation of 3H-thymidine during the final 4 h of culture. Antigen-independent, proliferative responses were assessed by culture (as above) in the presence of 1 µg/ml PHA (Sigma) or 10 µg/ml anti-CD3 antibody (clone KT3).

In vivo T cell transfer studies using OT-II cells
OVA-specific CD4+ T cells were obtained from spleens of OT-II mice [37 ], which are transgenic for an OVA-specific TCR. Single splenocyte suspensions were prepared, and RBCs were lysed. Cells were washed, resuspended in RPMI media containing 10% FCS and OVA (1 mg/ml, Sigma), and cultured at 37°C so that OVA-specific T cells could proliferate. Activated OVA-specific T cells were collected after 4.5 days in culture, washed to remove residual OVA, and resuspended at 5 x 106 cells/ml in RPMI. Cell purity was determined by flow cytometry by dual expression of TCR V{alpha}2 [anti-mouse V{alpha}2 mAb, clone B20.1, a gift of Dr. William Heath (The Walter and Eliza Hall Institute of Medical Research, Melbourne, Australia)] and CD4 (anti-CD4 mAb, clone GK1.5) and was routinely ~70%. WT and TF{delta}CT/{delta}CT mice were injected via the tail vein with 5 x 106 OT-II cells in 500 µl RPMI and challenged immediately by a subdermal injection in the right ear of 50 µg OVA containing 5 µg LPS in 20 µl normal saline. Subdermal challenge in the left ear with 50 µg of an irrelevant antigen (casein) and 5 µg LPS in 20 µl normal saline was performed as a control. Antigen-specific DTH responses were calculated from the difference in skin ear swelling to OVA and the irrelevant antigen, measured 24 h after subdermal challenge.

Studies in TF{delta}CT/{delta}CT chimeric mice
Chimeric mice were generated by transplanting bone marrow from TFäCT/äCT mice to WT mice and vice versa. Recipient mice received two doses of 550 rads total body {gamma}-radiation, 3 h apart. Donor bone marrow was extracted from femurs of nonirradiated mice by flushing cells with supplement-free DMEM cell culture medium. Single-cell suspensions were then collected and centrifuged at 500 g for 7 min, the supernatant discarded, and the pellet resuspended in 400 µl medium. RBC (1 ml) lysis buffer (0.15 M NH4Cl, 0.01 M NaHCO3, 0.1 M EDTA, pH 7.3) was added for 1 min at 37°C to lyse RBCs. The pellet was washed twice in medium and then resuspended at 2.5 x 107 cells/ml. Recipient mice were injected i.v. with 200 µl cells (5x106 cells) 24 h after irradiation. Eight weeks were allowed for bone marrow engraftment, at which time, using this protocol, >90% of circulating leukocytes are of the donor phenotype [38 ]. Mice were then immunized with OVA and challenged sub-dermally 21 days later to induce DTH (as outlined above).

Assessment of macrophage adhesion molecule expression and chemotactic responses to MCP-1
Mice were immunized with OVA, as described above, and peritoneal macrophages were elicited by i.p. injection with 0.5 ml sterile 3.8% thioglycollate. Macrophages were collected 72 h later by peritoneal lavage with 3 ml ice-cold PBS. Lavage samples were centrifuged at 300 g for 5 min, and the cells were resuspended at 0.5 x 106 cells/ml. Cells were incubated overnight in serum-free DMEM media, and their basal expression of membrane-activated complex 1 (Mac-1) and {alpha}-4-integrin was determined 24 h later by flow cytometry. Mac-1 expression was detected using Alexa Fluor 647-conjugated rat anti-mouse CD11b (integrin {alpha}M chain; Mac-1 chain), diluted 1:100, and {alpha}-4-integrin expression was detected using Alexa Fluor 488-conjugated rat anti-mouse CD49d ({alpha}-4-integrin), diluted 1:100. P-selectin-binding capacity of elicited macrophages was assessed as described previously [35 ]. Cells were incubated with a mouse P-selectin human IgG fusion protein (#555294, BD PharMingen, San Diego, CA, USA) at 25 µg/ml for 30 min at 37°C and then washed and labeled with FITC-conjugated sheep anti-human IgG (1:50) in the presence of 10% mouse serum. After further washing, binding of the P-selectin fusion protein to monocytes (identified using forward- and side-scatter) was determined by flow cytometry (MoFlo Cytomation, Fort Collins, CO, USA). Data are expressed as the increase in mean fluorescence above staining of cells incubated in the absence of the P-selectin fusion protein.

In vivo chemotactic responses of TF{delta}CT/{delta}CT and WT macrophages were assessed by intravital microscopy 4 h after intrascrotal injection of 34.5 ng MCP-1 (in 200 µl saline). Leukocyte emigration and adhesion were assessed in postcapillary venules, as described above. Cremaster muscle was excised and fixed with periodate/lysine/paraformaldehyde. Cryostat cut tissue sections were stained (using a three-layer immunoperoxidase technique) with anti-mouse CD68 mAb (FA/11) and anti-mouse mAb GR-1 to identify macrophages and neutrophils, respectively. The total number of stained cells was counted using computerized image analysis (Scion Image), and results are expressed as the number of positively stained cells relative to the area of tissue in the section.

Real-time PCR: TNF-{alpha} in DTH lesions
Ears were homogenized in 1 ml Trizol (Invitrogen, Carlsbad, CA, USA) with a Kinematica homogenizer (Littan-Lucerne, Switzerland). Samples were incubated at room temperature for 30 min, followed by the addition of 0.2 ml chloroform and then mixed for 15 s and incubated for 2–3 min at room temperature. Samples were spun at 12,000 g for 15 min at 4°C, and 0.5 ml isopropanol was added to the supernatant, which was incubated for 10 min at room temperature and spun at 12,000 g for 10 min. The RNA was washed with 1 ml 75% ethanol in RNase/DNase-free water, spun at 7500 g for 5 min, air-dried, resuspended in water, and stored at –70°C. RNA (5 µg), 150 ng hexamers, and 0.2 mM dNTPs were incubated at 65°C for 5 min and cooled on ice for 1 min. RT buffer, 12 mM MgCl2, 0.044 M DTT, and 1 µl RNase-out (Invitrogen) were added and kept at room temperature for 2 min. Superscript III (50 units) was added, and the samples were incubated for 10 min at room temperature. The samples were then incubated for 50 min at 42°C and stored at –70°C. PCR reactions contained 0.5 µM mouse TNF-{alpha} primers (forward: 5'-GCC TCT TCT CAT TCC TGC TT-3'; reverse: 5'-CAC TTG GTG GTT TGC TAC GA-3') and 1 µl SYBR Green master mix (Roche, Indianapolis, IN, USA) adjusted to a total volume of 20 µl with water. A standard curve with TNF-{alpha} cDNA was used, and values were normalized using β-actin as a housekeeping gene. The optimal temperature ramp cycle was 95°C for 5 s, 60°C for 5 s, and 72°C for 15 s for 40 cycles using a Rotor Gene 3000 light cycler (Corbett, Sydney, NSW, Australia).

p38-MAPK and Erk1/Erk2-MAPK levels in cultured splenocytes from OVA-sensitized mice
Splenocytes were isolated from OVA-sensitized WT and TF{delta}CT/{delta}CT mice, as described earlier, and cultured ex vivo for 2 h with 25 µg/ml OVA. Cell lysates were prepared and then analyzed by Western analysis according to the method recommended by the antibody supplier (Cell Signaling Technologies, Danvers, MA, USA). Samples were run on 12% gels and then transferred to polyvinylidene difluoride membranes, which were blocked for 1 h at room temperature in 1x TBS, 0.1% Tween-20 with 5% nonfat dry milk, and were washed in TBS-T three times for 5 min each wash. Membranes were then incubated overnight at 4°C with primary antibodies (rabbit anti-mouse phospho-p44/42 MAPK, rabbit anti-mouse p44/42 MAPK, rabbit anti-mouse phospho-p38 MAPK, and rabbit anti-mouse p38 MAPK) at 1:1000 in 1x TBS, 5% BSA, 0.1% Tween-20, and then washed three times in TBS-T for 5 min each wash. This was followed by incubation in swine anti-rabbit HRP-conjugated antibodies at 1:2000 in 1x TBS, 0.1% Tween-20 with 5% nonfat dry milk, three washes in TBS-T for 5 min each wash, and exposure to autoradiographic film with chemiluminescent reagent. Results are expressed as a ratio of phosphorylated protein to total protein, as determined by densitometry using Multi-Gauge v.3.0 software.

Statistical analysis
Data were assessed by statistical ANOVA followed by Tukey’s multiple comparison tests or when the experimental design involved two groups only, by Student’s t-test. All data are expressed as mean ± SEM and were considered statistically significant when P < 0.05.


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RESULTS
 
The TF cytoplasmic domain augments cutaneous DTH
Cutaneous DTH responses to three different antigens (OVA, mBSA, or PPD) were assessed in WT and TF{delta}CT/{delta}CT mice. TF{delta}CT/{delta}CT mice had significantly reduced, antigen-specific DTH responses to all three antigens compared with WT mice (Fig. 1 ). Similar significant reductions in DTH responses in WT and TF{delta}CT/{delta}CT mice were observed in initial experiments using littermate-matched mice (OVA, 52% reduction, P<0.05; mBSA 64% reduction, P<0.05; n=8 per group) and in experiments using TF{delta}CT/{delta}CT mice backcrossed on a C57BL/6 background (OVA 65% reduction, P<0.05, n=6 per group). DTH lesions of TF{delta}CT/{delta}CT mice contained significantly fewer Gr-1+ neutrophils (Fig. 2A ) and FA/11+ macrophages (Fig. 2B) compared with lesions in WT mice. There was no significant difference in fibrinogen deposition (Fig. 2C) in OVA-induced DTH lesions in WT compared with TF{delta}CT/{delta}CT mice (fibrinogen score, WT 2.25±0.95; TF{delta}CT/{delta}CT 2.50±0.56; n=6 per group).


Figure 1
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  		Figure 1. Cutaneous DTH responses to mBSA (A), OVA (B), and PPD (C) in WT and TF{delta}CT/{delta}CT mice (MF1/129S/v/Swiss background and strain controls). There were significant reductions in the antigen-specific DTH responses to all three antigens in mice lacking the cytoplasmic domain of TF. (*, P<0.05, and **, P<0.001, compared with WT mice; n=8 per group).


Figure 2
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  		Figure 2. Neutrophils (Gr-1+) and macrophages (FA/11+) in ear DTH lesions were identified by immunohistochemistry and counted as described in Materials and Methods. Neutrophil (A) and macrophage (B) accumulation was reduced significantly in TF{delta}CT/{delta}CT mice (*, P<0.05, compared with WT, n=6 per group; **, P<0.05, compared with WT, n=6 per group, respectively). No significant difference in fibrinogen deposition (C) in DTH lesions was observed between WT mice compared with TF{delta}CT/{delta}CT mice (n=6 per group).

TF{delta}CT/{delta}CT mice show reduced antigen-stimulated leukocyte rolling, adhesion, and infiltration
To examine the mechanisms underlying the reduced DTH responses, leukocyte-endothelial cell interactions were assessed in OVA-sensitized WT and TF{delta}CT/{delta}CT mice (n=5 per group) following local OVA challenge. Leukocyte rolling, adhesion, and transmigration in postcapillary venules of TF{delta}CT/{delta}CT mice were reduced significantly compared with WT mice, 24 h post-OVA antigen challenge (Fig. 3A 3B 3C , respectively). In addition, leukocyte-rolling velocity was elevated significantly in TF{delta}CT/{delta}CT mice (WT, 11.1±1.4 µm/s, vs. TF{delta}CT/{delta}CT, 29.3±6.9 µm/s, P<0.01), suggesting reduced leukocyte-endothelial cell interactions in TF{delta}CT/{delta}CT mice. These changes could not be attributed to alterations in blood flow, as RBC velocity (WT, 2.2±0.2 mm/s, vs. TF{delta}CT/{delta}CT, 2.9±0.5 mm/s), venular diameter (WT, 35.0±1.4 µm, vs. TF{delta}CT/{delta}CT, 32.0±1.5 µm), and venular shear rate (WT, 365.2±45.3 s–1, vs. TF{delta}CT/{delta}CT, 450±43.0 s–1) did not differ significantly between WT and TF{delta}CT/{delta}CT mice. In addition, the reduced leukocyte-endothelial cell interactions in TF{delta}CT/{delta}CT mice were not a result of reduced expression of endothelial cell adhesion molecules assessed by semiquantitative histological scoring of P-selectin, ICAM-1, and VCAM-1 in DTH lesions (data not shown).


Figure 3
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  		Figure 3. Leukocyte rolling flux (A), adhesion (B), and interstitial leukocyte emigration (C) assessed by intravital microscopy in postcapillary venules 24 h after OVA challenge in sensitized WT and TF{delta}CT/{delta}CT mice (MF1/129S/v/Swiss background and strain controls; *, P<0.05, compared with WT mice; n=5 per group).

The cytoplasmic domain of TF on leukocytes enhances the DTH response in mice
To examine the role of leukocytes and the endothelium in reducing the DTH response in TF{delta}CT/{delta}CT mice, DTH was induced (with OVA) in chimeric mice produced by bone marrow transplantation. Transplanting TF{delta}CT/{delta}CT bone marrow into WT mice significantly reduced (58%) the DTH response compared with transplanting WT bone marrow into WT mice (i.e., sham chimeras; Fig. 4A ). DTH responses in sham chimeras were similar in magnitude to the DTH responses in nontransplanted WT mice (Fig. 1B) . DTH responses in TF{delta}CT/{delta}CT mice transplanted with TF{delta}CT/{delta}CT bone marrow were equivalent to those in nontransplanted TF{delta}CT/{delta}CT mice. Transplantation of WT bone marrow in TF{delta}CT/{delta}CT mice resulted in DTH responses equivalent to those of WT mice and WT sham chimeras (Fig. 4A) . These studies indicate that the absence of the cytoplasmic domain of TF in leukocytes is responsible for the observed effects on the DTH response. Intravital microscopy of the same chimeric animals (Fig. 4B) revealed a parallel relationship between leukocyte emigration and DTH response; i.e., the absence of the cytoplasmic tail of TF in leukocytes but not in endothelial cells was associated with significantly reduced leukocyte emigration. In support of this observation, the expression of VCAM-1 and P-selectin in cultured lung microvascular endothelial cells isolated from WT and TF{delta}CT/{delta}CT mice was similar (data not shown).


Figure 4
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  		Figure 4. Cutaneous DTH responses to OVA in chimeric mice (A). Bone marrow was transplanted from donors to recipients (donor>>recipient as indicated), and cutaneous DTH was induced 8 weeks later, as described. Attenuated DTH responses in TF{delta}CT/{delta}CT mice were associated with their leukocyte phenotype (n=6 per group; *, P<0.02, WT>>WT vs. TF{delta}CT/{delta}CT>>TF{delta}CT/{delta}CT; **, P<0.005, TF{delta}CT/{delta}CT>>WT vs. WT>>WT; and ***, P<0.005, TF{delta}CT/{delta}CT>>WT vs. WT>>TF{delta}CT/{delta}CT). Interstitial leukocyte accumulation (B) was assessed in the same animals by intravital microscopy in postcapillary venules 24 h after antigen (OVA) challenge in WT and TF{delta}CT/{delta}CT mice. Impaired leukocyte emigration was also associated with the leukocyte phenotype (*, P<0.008, n=6 per group, WT>>TF{delta}CT/{delta}CT compared with TF{delta}CT/{delta}CT>>WT mice).

Ex vivo OVA-stimulated splenocyte proliferation is normal in TF{delta}CT/{delta}CT mice
To determine whether a T cell defect or defects in other leukocytes were responsible for reduced DTH responses in TF{delta}CT/{delta}CT mice, ex vivo antigen-stimulated splenocyte proliferation and T cell transfer studies were performed. OVA-stimulated proliferation of splenocytes from sensitized WT and TF{delta}CT/{delta}CT mice was similar over a range of doses, indicating antigen-stimulated T cell proliferation is unaffected in the absence of the cytoplasmic domain of TF (Fig. 5A ). In addition, antigen-independent, proliferative responses to PHA and anti-CD3 antibody in both groups were similar (Fig. 5B) .


Figure 5
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  		Figure 5. Ex vivo proliferative responses of splenocytes from sensitized mice (A) showed a significant, dose-dependent response to OVA (*, P<0.005, **, P<0.0001, and ***, P<0.0005, compared with no OVA stimulation, n=6 per group). However, there were no significant differences in the proliferative responses of WT (solid bars) and TF{delta}CT/{delta}CT (open bars) mice. Similarly, the proliferative responses to 1 µg/ml PHA and 10 µg/ml anti-CD3 mAb were not significantly different in WT (solid bars) and TF{delta}CT/{delta}CT (open bars) mice (B; n=6 per group; *, P < 0.0001 compared with unstimulated cells; **, P<0.0001, compared with unstimulated cells).

Transfer of OVA-sensitized CD4+ T cells does not restore the DTH responses in TF{delta}CT/{delta}CT mice
To determine if defective in vivo T cell responses could explain the reduced DTH response in TF{delta}CT/{delta}CT mice, T cell transfer studies were performed using purified, ex vivo-activated, OVA-responsive CD4+ T cells from OT-II TCR transgenic mice. These cells were transferred into naïve TF{delta}CT/{delta}CT and WT mice, and the mice were challenged immediately with a subdermal injection of OVA or an irrelevant antigen (casein). Transfer of activated OT-II CD4+ T cells (which have normal TF) to TF{delta}CT/{delta}CT mice failed to induce OVA-specific DTH, whereas transfer of the equivalent number of cells to WT mice induced a significant antigen-specific, skin-swelling response (Fig. 6 ). Responses following T cell transfer in WT mice were similar in magnitude to those observed following active sensitization to OVA in WT mice (Fig. 1B) . Thus, restoration of sensitized, functionally active CD4+ T cells does not overcome the defective DTH responses in TF{delta}CT/{delta}CT mice, suggesting that the critical defect is in other leukocyte populations.


Figure 6
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  		Figure 6. DTH responses of naïve WT (n=5) and TF{delta}CT/{delta}CT (n=7) mice following transfer of activated, OVA-specific OT-II TCR transgenic CD4+ T cells and OVA challenge in the right ear. Casein was used as an irrelevant antigen in the contralateral ear. Transfer of OVA-activated T cells (expressing full-length TF) enabled normal DTH responses to OVA in WT mice but not TF{delta}CT/{delta}CT mice (*, P<0.001, compared with casein challenge in WT mice, n=6 per group; **, P<0.001, compared with OVA challenge in WT mice, n=6 per group).

Macrophages from TF{delta}CT/{delta}CT mice have reduced capacity to bind P-selectin and reduced migration in response to MCP-1
Elicited peritoneal macrophages were analyzed to examine the role of specific adhesion molecules in the reduced leukocyte-endothelial cells interactions in TF{delta}CT/{delta}CT mice. Macrophages were deactivated by overnight culture in the absence of serum. Their capacity to bind a mouse P-selectin-human IgG fusion protein was then assessed in a functional assay for P-selectin glycoprotein ligand 1 (PSGL-1) [39 ]. P-selectin binding was reduced significantly in macrophages from TF{delta}CT/{delta}CT mice compared with WT mice (Fig. 7A ), indicating a functional defect in their ability to interact with P-selection and induce leukocyte rolling. In contrast, no difference was observed in macrophage expression of Mac-1 and the {alpha}4-integrin (Fig. 7B and 7C , respectively).


Figure 7
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  		Figure 7. P-selectin-binding capacity (A), expression of Mac-1 (B), and {alpha}-4-integrin (C) on elicited macrophages isolated from TF{delta}CT/{delta}CT and WT mice, as determined by flow cytometric analysis. There was a significant reduction in P-selectin binding in TF{delta}CT/{delta}CT mice, indicating reduced PSGL-1 function (*, P<0.05, compared with WT mice, n=6 per group).

To explore the possibility that the reduced leukocyte recruitment in TF{delta}CT/{delta}CT mice was a result of impairment in the migratory ability of leukocytes in these mice, in vivo chemotactic responses to MCP-1 (CCL2) were examined in cremasteric, postcapillary venules by intravital microscopy. MCP-1-induced leukocyte emigration (Fig. 8A ) was reduced significantly by 50% in TF{delta}CT/{delta}CT mice compared with WT mice, with a significant reduction of macrophages (60%) but no significant reduction in neutrophils (Fig. 8B) . In WT mice, ~75% of leukocytes recruited in response to MCP-1 were CD68+ macrophages, and the remaining cells were Gr-1+ neutrophils (Fig. 8B) . These findings provide direct in vivo evidence that absence of the TF cytoplasmic domain is associated with impaired macrophage migratory responses.


Figure 8
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  		Figure 8. Reduced interstitial leukocyte emigration in response to administration of MCP-1 (A) was demonstrated in postcapillary venules by intravital microscopy in TF{delta}CT/{delta}CT mice (*, P<0.001, compared with WT mice; WT, n=4; and TF{delta}CT/{delta}CT, n=6). Emigrating leukocytes were identified in cremaster muscle sections (B) by staining with anti-CD68 (macrophages) and anti-Gr-1 (neutrophils). The reduced leukocyte emigration was a result of reduced CD68+ macrophage numbers (*, P<0.001, compared with WT mice; WT, n=4; and TF{delta}CT/{delta}CT, n=6). There was no significant difference in Gr-1+ neutrophil numbers.

Deletion of the cytoplasmic domain of TF suppresses the expression of TNF-{alpha} and the phosphorylation of p38-MAPK
Phosphorylation of MAPKs in splenocytes following ex vivo antigen stimulation was quantitated by Western blotting to examine effects of the absence of the cytoplasmic domain of TF on intracellular signaling pathways activated during immune activation. A significant (50%) reduction in phosphorylation of p38-MAPK (Fig. 9A ) and increased phosphorylation of Erk1- and Erk2-MAPK (Fig. 9B) were demonstrated in TF{delta}CT/{delta}CT mice compared with WT mice. A similar increase in phosphorylated Erk1 and Erk2-MAPK has been reported in TF{delta}CT/{delta}CT mice in a model of endotoxemia, where the authors concluded that the TF cytoplasmic domain suppresses Erk1/2 phosphorylation [40 ]. Also, reduced phosphorylation of p38 MAPK in cell lines expressing TF lacking the cytoplasmic domain has been associated with impaired migratory behavior in vitro [15 ].


Figure 9
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  		Figure 9. Quantitative Western blot analysis of ex vivo OVA-stimulated splenocytes from sensitized WT and TF{delta}CT/{delta}CT mice shows a reduction in the phosphorylation of p38-MAPK (A) relative to total p38 in TF{delta}CT/{delta}CT mice (*, P<0.005, compared with WT, n=5 per group). In contrast, there is an increase in the phosphorylation of Erk1- and Erk2-MAPK (B) relative to total Erk1 and Erk2 in TF{delta}CT/{delta}CT mice (*, P<0.005, compared with WT, n=5 per group; **, P<0.001, compared with WT, n=5 per group). TNF-{alpha} mRNA levels in DTH lesions measured by real-time PCR and expressed as a ratio relative to β-actin (C) were significantly lower in TF{delta}CT/{delta}CT mice compared with WT mice (*, P<0.05, compared with WT, n=6 per group).

In conjunction with reduced macrophage (60%) and neutrophil (50%) accumulation, TNF-{alpha} mRNA levels were significantly lower (~80%) in DTH lesions of TF{delta}CT/{delta}CT mice (Fig. 9C) . The greater reduction of TNF-{alpha} production compared with the reduction in leukocyte accumulation would be consistent with less TNF-{alpha} production per recruited leukocyte (suggestive of reduced leukocyte activation as well as recruitment) and/or reduced TNF-{alpha} production by other cells at the site of the inflammatory response. Previously, elicited macrophages from TF{delta}CT/{delta}CT mice have also been shown to express significantly less TNF-{alpha} than WT mice [29 ].


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DISCUSSION
 
Prominent accumulation of macrophages expressing high levels of TF [24 ] and extravascular fibrin deposition [41 ] contributing to skin swelling [42 ] indicate a key role for the procoagulant function of TF in DTH responses. However, studies of "DTH-like" inflammatory renal injury showing that inhibition of TF provides significant protection, even in the absence of fibrinogen [43 ], suggest additional coagulation-independent roles for TF in adaptive immunity.

In this study, the development of cutaneous DTH was assessed in mice lacking the cytoplasmic domain of TF. In these mice, the extracellular and transmembrane domains of TF are expressed, and coagulation function is normal [29 ]. The results demonstrate an important contribution of the cytoplasmic domain of TF to the development of cutaneous DTH responses. TF{delta}CT/{delta}CT mice had significant reductions in DTH (30–60%) to three standard antigens (OVA, mBSA, and PPD), reduced macrophage recruitment in DTH lesions, and significant defects in leukocyte-endothelial interactions in the effector phase of DTH responses. Similar amounts of fibrin deposition in these DTH lesions indicate that the reduced response is not a result of impaired procoagulant function.

DTH responses require initial T cell sensitization to antigen presented in secondary lymphoid organs and a subsequent secondary response to local antigen challenge. Recruitment of leukocytes to the site of antigen challenge in DTH responses requires local recognition of antigen by memory T cells, which direct recruitment of additional effector cells, including T cells, macrophages, and neutrophils, to mediate the local inflammatory response. Circulating leukocytes undergo a sequence of rolling, followed by firm adhesion to activated endothelial cells in the postcapillary venules prior to endothelial transmigration into an inflammatory site [44 ]. These leukocyte-endothelial interactions in postcapillary venules of the cremaster muscle were examined by intravital microscopy following local antigen challenge in sensitized TF{delta}CT/{delta}CT mice, and significantly impaired leukocyte rolling and adhesion to the endothelium and reduced leukocyte accumulation were demonstrated. These results suggest an important role for the cytoplasmic domain of TF in antigen-stimulated, leukocyte-endothelial cell interactions and in leukocyte recruitment during cell-mediated adaptive immune responses. Similar impairment of leukocyte-endothelial cell interactions in response to an innate inflammatory stimulus was observed following endotoxin-challenge in TF{delta}CT/{delta}CT mice [28 ].

To determine whether leukocytes or endothelial cells were responsible for attenuated DTH responses in TF{delta}CT/{delta}CT mice, we used bone marrow transplantation to reconstitute WT mice with TF{delta}CT/{delta}CT bone marrow (and vice versa). TF{delta}CT/{delta}CT mice transplanted with WT bone marrow had normal DTH responses, equivalent to that seen in nontransplanted WT mice and WT mice transplanted with WT bone marrow (sham transplants). However, WT mice transplanted with TF{delta}CT/{delta}CT bone marrow had significantly reduced DTH responses (equivalent to nontransplanted and "sham"-transplanted TF{delta}CT/{delta}CT mice), indicating that the expression of the cytoplasmic domain of TF in leukocytes rather than in endothelial cells is required for development of the cutaneous DTH response. The absence of a demonstrable difference in adhesion molecule expression by endothelial cells in TF{delta}CT/{delta}CT mice is consistent with the hypothesis that reduction in leukocyte/endothelial interactions is not a result of an endothelial cell adhesion molecule defect.

To explore which leukocyte subpopulation was responsible for impaired DTH, we examined T cell responses in TF{delta}CT/{delta}CT mice, ex vivo and in vivo. CD4+ T cells are central to the development of DTH via their ability to recognize antigens, develop antigen-specific memory, and coordinate the cellular response upon peripheral antigen challenge. To determine whether deficient T cell activation occurred in the absence of the cytoplasmic domain of TF, ex vivo antigen-stimulated proliferation of splenocytes from mice sensitized in vivo to OVA was assessed. Splenocyte proliferation to OVA was unaffected in mice lacking the cytoplasmic domain of TF, indicating no apparent defect in primary antigen presentation, recognition, or activation of their T cells. To determine if the defect in DTH responses could be overcome in TF{delta}CT/{delta}CT mice by providing WT-activated CD4+ T cells, passive transfer studies were performed using OT-II CD4+ T cells (expressing intact TF), which were activated by OVA ex vivo. Transfer of these activated T cells resulted in normal DTH responses following cutaneous OVA challenge in naïve WT mice but failed to facilitate DTH responses in TF{delta}CT/{delta}CT mice, demonstrating that provision of competent, activated T cells does not overcome their impaired DTH response. Together with the normal antigen-stimulated proliferation responses, these findings indicate that a defect in T cell function is not primarily responsible for the attenuation of DTH in the absence of the cytoplasmic domain of TF.

As T cell function appeared intact in TF{delta}CT/{delta}CT mice, the function of other effector leukocytes involved in DTH responses, particularly macrophages, was studied. These experiments focused on the multistep process required for leukocyte recruitment into DTH lesions. First, molecules associated with leukocyte rolling were examined. The observation of reduced rolling in postcapillary venules of TF{delta}CT/{delta}CT mice undergoing antigen-specific DTH responses suggested altered expression and/or function of molecules, which mediate rolling. Previous studies of DTH responses have implicated the P-selectin/PSGL-1 pathway [33 , 45 , 46 ]. This was confirmed in the present study by demonstrating that P-selectin inhibition effectively eliminated rolling in wild-type and TF{delta}CT/{delta}CT mice (data not shown). Given the absence of any difference in endothelial P-selectin expression, it was hypothesized that the defect may be related to an altered leukocyte P-selectin-binding function. Using isolated peritoneal macrophages, we demonstrated that the absence of the TF cytoplasmic domain is associated with reduced P-selectin-binding activity. This suggests that the TF cytoplasmic domain contributes to macrophage expression of functional P-selectin ligands, such as PSGL-1. Furthermore, this provides a mechanism to explain the reduction in rolling observed in TF{delta}CT/{delta}CT mice.

The subsequent step of leukocyte adhesion was also significantly reduced in TF{delta}CT/{delta}CT mice. This may have reflected the reduction in leukocyte rolling, although in some inflammatory models, leukocyte rolling must be reduced to a greater degree than that observed in the present study to result in significant impairment of leukocyte adhesion [35 , 47 ]. An alternative explanation could be that the expression and/or function of adhesion ligand/receptor molecules associated with leukocyte adhesion were altered in TF{delta}CT/{delta}CT mice. Two candidate pathways for this effect are the Mac-1/ICAM-1 interaction and the {alpha}4-integrin/VCAM-1 pathway. No clear differences in endothelial expression of ICAM-1 and VCAM-1 were detected, supporting the finding from the chimeric mouse experiments that the defect was independent of the endothelial phenotype. Macrophage expression of Mac-1 and the {alpha}4-integrin was also comparable between wild-type and TF{delta}CT/{delta}CT mice, indicating that leukocyte integrin expression was not markedly reduced in TF{delta}CT/{delta}CT mice, although functional alterations in these adhesion molecules cannot be excluded.

Effector responses in DTH require leukocyte-endothelial cell transmigration, and the TF cytoplasmic domain has been shown to promote migration of various cell types in vitro. The presence of the cytoplasmic domain of TF enhances migration of a human bladder carcinoma cell line (J82) by activation of p38 and Rac1 [15 ] and promotes adhesion, spreading, and migration in this cell line by binding to ABP-280 [8 ]. The absence of the cytoplasmic domain of TF impairs migration of mouse aortic smooth muscle cells [16 ] and impairs chemotaxis of transfected porcine aortic endothelial cells [48 ]. To assess the migratory responses of macrophages lacking the cytoplasmic domain of TF independently of other complex leukocyte and endothelial activation signals involved with antigen-stimulated DTH responses, we investigated their ability to migrate in vivo in response to the macrophage-specific chemokine MCP-1. TF{delta}CT/{delta}CT mice displayed reduced monocyte migration, despite achieving comparable levels of leukocyte adhesion within the microvasculature. These findings provide strong evidence that the presence of the cytoplasmic domain of TF in macrophages promotes their ability to transmigrate across the endothelium and accumulate in cutaneous DTH responses.

TNF-{alpha} plays an important role in neutrophil recruitment and p38-MAPK activation during inflammation [49 ]. p38-MAPK is activated by TNF-{alpha} and regulates a wide range of inflammatory responses (including leukocyte chemotaxis and emigration) in many different cells [49 ]. Inhibition of p38-MAPK can reduce TNF-{alpha}-induced inflammatory responses and MCP-1-mediated chemotaxis [50 ]. Peritoneal macrophages from TF{delta}CT/{delta}CT mice with arthritis have previously been shown to have reduced TNF mRNA expression [29 ]. Impaired leukocyte TNF-{alpha} production in DTH lesions of TF{delta}CT/{delta}CT mice could provide a mechanistic explanation for impaired p38-MAPK activation and reduced leukocyte chemotaxis into DTH lesions of TF{delta}CT/{delta}CT mice. Impaired p38-MAPK signaling may also blunt expression of other proinflammatory genes, which contribute to the local DTH response.

In summary, these studies provide in vivo evidence for a role of the cytoplasmic domain of TF in macrophages in leukocyte-endothelial interactions, inflammatory cell recruitment and activation, and tissue injury in adaptive immune responses, which initiate cutaneous DTH.


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ACKNOWLEDGEMENTS
 
These studies were supported by a Program grant from the National Health and Medical Research Council of Australia (NH&MRC) and a NH&MRC Fellowship grant (P. G. T.).

Received June 6, 2007; revised October 29, 2007; accepted November 30, 2007.


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REFERENCES
 
    1
  1. Camerer, E., Kolsto, A. B., Prydz, H. (1996) Cell biology of tissue factor, the principal initiator of blood coagulation Thromb. Res. 81,1-41[CrossRef][Medline]
    2. 2
  2. Carmeliet, P., Mackman, N., Moons, L., Luther, T., Gressens, P., Van Vlaenderen, I., Demunck, H., Kasper, M., Breier, G., Evrard, P., et al (1996) Role of tissue factor in embryonic blood vessel development Nature 383,73-75[CrossRef][Medline]
    3. 3
  3. Bugge, T. H., Xiao, Q., Kombrinck, K. W., Flick, M. J., Holmback, K., Danton, M. J., Colbert, M. C., Witte, D. P., Fujikawa, K., Davie, E. W., Degen, J. L. (1996) Fatal embryonic bleeding events in mice lacking tissue factor, the cell-associated initiator of blood coagulation Proc. Natl. Acad. Sci. USA 93,6258-6263[Abstract/Free Full Text]
    4. 4
  4. Parry, G. C., Mackman, N. (2000) Mouse embryogenesis requires the tissue factor extracellular domain but not the cytoplasmic domain J. Clin. Invest. 105,1547-1554[Medline]
    5. 5
  5. Luther, T., Flossel, C., Mackman, N., Bierhaus, A., Kasper, M., Albrecht, S., Sage, E. H., Iruela-Arispe, L., Grossmann, H., Strohlein, A., et al (1996) Tissue factor expression during human and mouse development Am. J. Pathol. 149,101-113[Abstract]
    6. 6
  6. Abe, K., Shoji, M., Chen, J., Bierhaus, A., Danave, I., Micko, C., Casper, K., Dillehay, D. L., Nawroth, P. P., Rickles, F. R. (1999) Regulation of vascular endothelial growth factor production and angiogenesis by the cytoplasmic tail of tissue factor Proc. Natl. Acad. Sci. USA 96,8663-8668[Abstract/Free Full Text]
    7. 7
  7. Mueller, B. M., Ruf, W. (1998) Requirement for binding of catalytically active factor VIIa in tissue factor-dependent experimental metastasis J. Clin. Invest. 101,1372-1378[Medline]
    8. 8
  8. Ott, I., Fischer, E. G., Miyagi, Y., Mueller, B. M., Ruf, W. (1998) A role for tissue factor in cell adhesion and migration mediated by interaction with actin-binding protein 280 J. Cell Biol. 140,1241-1253[Abstract/Free Full Text]
    9. 9
  9. Bokarewa, M. I., Morrissey, J. H., Tarkowski, A. (2002) Tissue factor as a proinflammatory agent Arthritis Res. 4,190-195[CrossRef][Medline]
    10. 10
  10. Luther, T., Mackman, N. (2001) Tissue factor in the heart Multiple roles in hemostasis, thrombosis, and inflammation. Trends Cardiovasc. Med. 11,307-312
    11. 11
  11. Camerer, E., Gjernes, E., Wiiger, M., Pringle, S., Prydz, H. (2000) Binding of factor VIIa to tissue factor on keratinocytes induces gene expression J. Biol. Chem. 275,6580-6585[Abstract/Free Full Text]
    12. 12
  12. Versteeg, H. H., Spek, C. A., Slofstra, S. H., Diks, S. H., Richel, D. J., Peppelenbosch, M. P. (2004) FVIIa:TF induces cell survival via G12/G13-dependent Jak/STAT activation and BclXL production Circ. Res. 94,1032-1040[Abstract/Free Full Text]
    13. 13
  13. Mody, R. S., Carson, S. D. (1997) Tissue factor cytoplasmic domain peptide is multiply phosphorylated in vitro Biochemistry 36,7869-7875[CrossRef][Medline]
    14. 14
  14. Cunningham, M. A., Romas, P., Hutchinson, P., Holdsworth, S. R., Tipping, P. G. (1999) Tissue factor and factor VIIa receptor/ligand interactions induce proinflammatory effects in macrophages Blood 94,3413-3420[Abstract/Free Full Text]
    15. 15
  15. Ott, I., Weigand, B., Michl, R., Seitz, I., Sabbari-Erfani, N., Neumann, F. J., Schomig, A. (2005) Tissue factor cytoplasmic domain stimulates migration by activation of the GTPase Rac1 and the mitogen-activated protein kinase p38 Circulation 111,349-355[Abstract/Free Full Text]
    16. 16
  16. Ott, I., Michaelis, C., Schuermann, M., Steppich, B., Seitz, I., Dewerchin, M., Zohlnhofer, D., Wessely, R., Rudelius, M., Schomig, A., Carmeliet, P. (2005) Vascular remodeling in mice lacking the cytoplasmic domain of tissue factor Circ. Res. 97,293-298[Abstract/Free Full Text]
    17. 17
  17. Belting, M., Dorrell, M. I., Sandgren, S., Aguilar, E., Ahamed, J., Dorfleutner, A., Carmeliet, P., Mueller, B. M., Friedlander, M., Ruf, W. (2004) Regulation of angiogenesis by tissue factor cytoplasmic domain signaling Nat. Med. 10,502-509[CrossRef][Medline]
    18. 18
  18. Drake, T. A., Morrissey, J. H., Edgington, T. S. (1989) Selective cellular expression of tissue factor in human tissues Implications for disorders of hemostasis and thrombosis. Am. J. Pathol. 134,1087-1097
    19. 19
  19. Gregory, S. A., Morrissey, J. H., Edgington, T. S. (1989) Regulation of tissue factor gene expression in the monocyte procoagulant response to endotoxin Mol. Cell. Biol. 9,2752-2755[Abstract/Free Full Text]
    20. 20
  20. Colucci, M., Balconi, G., Lorenzet, R., Pietra, A., Locati, D., Donati, M. B., Semeraro, N. (1983) Cultured human endothelial cells generate tissue factor in response to endotoxin J. Clin. Invest. 71,1893-1896[Medline]
    21. 21
  21. Schwager, I., Jungi, T. W. (1994) Effect of human recombinant cytokines on the induction of macrophage procoagulant activity Blood 83,152-160[Abstract/Free Full Text]
    22. 22
  22. Bevilacqua, M. P., Pober, J. S., Majeau, G. R., Fiers, W., Cotran, R. S., Gimbrone, M. A., Jr (1986) Recombinant tumor necrosis factor induces procoagulant activity in cultured human vascular endothelium: characterization and comparison with the actions of interleukin 1 Proc. Natl. Acad. Sci. USA 83,4533-4537[Abstract/Free Full Text]
    23. 23
  23. Bevilacqua, M. P., Pober, J. S., Wheeler, M. E., Cotran, R. S., Gimbrone, M. A., Jr (1985) Interleukin-1 activation of vascular endothelium Effects on procoagulant activity and leukocyte adhesion. Am. J. Pathol. 121,394-403
    24. 24
  24. Imamura, T., Iyama, K., Takeya, M., Kambara, T., Nakamura, S. (1993) Role of macrophage tissue factor in the development of the delayed hypersensitivity reaction in monkey skin Cell. Immunol. 152,614-622[CrossRef][Medline]
    25. 25
  25. Geczy, C. L., Meyer, P. A. (1982) Leukocyte procoagulant activity in man: an in vitro correlate of delayed-type hypersensitivity J. Immunol. 128,331-336[Abstract]
    26. 26
  26. Gregory, S. A., Kornbluth, R. S., Helin, H., Remold, H. G., Edgington, T. S. (1986) Monocyte procoagulant inducing factor: a lymphokine involved in the T cell-instructed monocyte procoagulant response to antigen J. Immunol. 137,3231-3239[Abstract]
    27. 27
  27. Geczy, C. L. (1984) Induction of macrophage procoagulant by products of activated lymphocytes Haemostasis 14,400-411[Medline]
    28. 28
  28. Sharma, L., Melis, E., Hickey, M. J., Clyne, C. D., Erlich, J., Khachigian, L. M., Davenport, P., Morand, E., Carmeliet, P., Tipping, P. G. (2004) The cytoplasmic domain of tissue factor contributes to leukocyte recruitment and death in endotoxemia Am. J. Pathol. 165,331-340[Abstract/Free Full Text]
    29. 29
  29. Yang, Y. H., Hall, P., Milenkovski, G., Sharma, L., Hutchinson, P., Melis, E., Carmeliet, P., Tipping, P., Morand, E. (2004) Reduction in arthritis severity and modulation of immune function in tissue factor cytoplasmic domain mutant mice Am. J. Pathol. 164,109-117[Abstract/Free Full Text]
    30. 30
  30. Melis, E., Moons, L., De Mol, M., Herbert, J. M., Mackman, N., Collen, D., Carmeliet, P., Dewerchin, M. (2001) Targeted deletion of the cytosolic domain of tissue factor in mice does not affect development Biochem. Biophys. Res. Commun. 286,580-586[CrossRef][Medline]
    31. 31
  31. Drew, A. F., Tucker, H. L., Liu, H., Witte, D. P., Degen, J. L., Tipping, P. G. (2001) Crescentic glomerulonephritis is diminished in fibrinogen-deficient mice Am. J. Physiol. Renal Physiol. 281,F1157-F1163[Abstract/Free Full Text]
    32. 32
  32. Gregory, J. L., Morand, E. F., McKeown, S. J., Ralph, J. A., Hall, P., Yang, Y. H., McColl, S. R., Hickey, M. J. (2006) Macrophage migration inhibitory factor induces macrophage recruitment via CC chemokine ligand 2 J. Immunol. 177,8072-8079[Abstract/Free Full Text]
    33. 33
  33. Tipping, P. G., Huang, X. R., Berndt, M. C., Holdsworth, S. R. (1996) P-selectin directs T lymphocyte-mediated injury in delayed-type hypersensitivity responses: studies in glomerulonephritis and cutaneous delayed-type hypersensitivity Eur. J. Immunol. 26,454-460[Medline]
    34. 34
  34. Hickey, M. J., Sharkey, K. A., Sihota, E. G., Reinhardt, P. H., MacMicking, J. D., Nathan, C., Kubes, P. (1997) Inducible nitric oxide synthase (iNOS)-deficient mice have enhanced leukocyte-endothelium interactions in endotoxemia FASEB J. 11,955-964[Abstract]
    35. 35
  35. Hickey, M. J., Bullard, D. C., Issekutz, A., James, W. G. (2002) Leukocyte-endothelial cell interactions are enhanced in dermal postcapillary venules of MRL/faslpr (lupus-prone) mice: roles of P- and E-selectin J. Immunol. 168,4728-4736[Abstract/Free Full Text]
    36. 36
  36. Norman, M. U., Van De Velde, N. C., Timoshanko, J. R., Issekutz, A., Hickey, M. J. (2003) Overlapping roles of endothelial selectins and vascular cell adhesion molecule-1 in immune complex-induced leukocyte recruitment in the cremasteric microvasculature Am. J. Pathol. 163,1491-1503[Abstract/Free Full Text]
    37. 37
  37. Barnden, M. J., Allison, J., Heath, W. R., Carbone, F. R. (1998) Defective TCR expression in transgenic mice constructed using cDNA-based {alpha}- and β-chain genes under the control of heterologous regulatory elements Immunol. Cell Biol. 76,34-40[CrossRef][Medline]
    38. 38
  38. Timoshanko, J. R., Holdsworth, S. R., Kitching, A. R., Tipping, P. G. (2002) IFN-{gamma} production by intrinsic renal cells and bone marrow-derived cells is required for full expression of crescentic glomerulonephritis in mice J. Immunol. 168,4135-4141[Abstract/Free Full Text]
    39. 39
  39. Li, F., Wilkins, P. P., Crawley, S., Weinstein, J., Cummings, R. D., McEver, R. P. (1996) Post-translational modifications of recombinant P-selectin glycoprotein ligand-1 required for binding to P- and E-selectin J. Biol. Chem. 271,3255-3264[Abstract/Free Full Text]
    40. 40
  40. Ahamed, J., Niessen, F., Kurokawa, T., Lee, Y. K., Bhattacharjee, G., Morrissey, J. H., Ruf, W. (2007) Regulation of macrophage procoagulant responses by the tissue factor cytoplasmic domain in endotoxemia Blood 109,5251-5259[Abstract/Free Full Text]
    41. 41
  41. Colvin, R. B., Johnson, R. A., Mihm, M. C., Jr, Dvorak, H. F. (1973) Role of the clotting system in cell-mediated hypersensitivity. I. Fibrin deposition in delayed skin reactions in man J. Exp. Med. 138,686-698[Abstract]
    42. 42
  42. Colvin, R. B., Mosesson, M. W., Dvorak, H. F. (1979) Delayed-type hypersensitivity skin reactions in congenital afibrinogenemia lack fibrin deposition and induration J. Clin. Invest. 63,1302-1306[Medline]
    43. 43
  43. Cunningham, M. A., Kitching, A. R., Tipping, P. G., Holdsworth, S. R. (2004) Fibrin independent proinflammatory effects of tissue factor in experimental crescentic glomerulonephritis Kidney Int. 66,647-654[CrossRef][Medline]
    44. 44
  44. Springer, T. A. (1995) Traffic signals on endothelium for lymphocyte recirculation and leukocyte emigration Annu. Rev. Physiol. 57,827-872[CrossRef][Medline]
    45. 45
  45. Kanwar, S., Bullard, D. C., Hickey, M. J., Smith, C. W., Beaudet, A. L., Wolitzky, B. A., Kubes, P. (1997) The association between {alpha}4-integrin, P-selectin, and E-selectin in an allergic model of inflammation J. Exp. Med. 185,1077-1087[Abstract/Free Full Text]
    46. 46
  46. McCafferty, D. M., Kanwar, S., Granger, D. N., Kubes, P. (2000) E/P-selectin-deficient mice: an optimal mutation for abrogating antigen but not tumor necrosis factor-{alpha}-induced immune responses Eur. J. Immunol. 30,2362-2371[CrossRef][Medline]
    47. 47
  47. Kubes, P., Jutila, M., Payne, D. (1995) Therapeutic potential of inhibiting leukocyte rolling in ischemia/reperfusion J. Clin. Invest. 95,2510-2519[Medline]
    48. 48
  48. Siegbahn, A., Johnell, M., Sorensen, B. B., Petersen, L. C., Heldin, C. H. (2005) Regulation of chemotaxis by the cytoplasmic domain of tissue factor Thromb. Haemost. 93,27-34[Medline]
    49. 49
  49. Nwariaku, F. E., Chang, J., Zhu, X., Liu, Z., Duffy, S. L., Halaihel, N. H., Terada, L., Turnage, R. H. (2002) The role of p38 MAP kinase in tumor necrosis factor-induced redistribution of vascular endothelial cadherin and increased endothelial permeability Shock 18,82-85[Medline]
    50. 50
  50. Lokuta, M. A., Huttenlocher, A. (2005) TNF-{alpha} promotes a stop signal that inhibits neutrophil polarization and migration via a p38 MAPK pathway J. Leukoc. Biol. 78,210-219[Abstract/Free Full Text]




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