Identification of the Protein C/Activated Protein C Binding Sites on the Endothelial Cell Protein C Receptor

IMPLICATIONS FOR A NOVEL MODE OF LIGAND RECOGNITION BY A MAJOR HISTOCOMPATIBILITY COMPLEX CLASS 1-TYPE RECEPTOR*

Patricia C. Y. LiawDagger §, Timothy MatherDagger , Natalia Oganesyan, Gary L. Ferrell, and Charles T. EsmonDagger ||**DaggerDagger

From the Dagger  Cardiovascular Biology Research Program, Oklahoma Medical Research Foundation and the Departments of ** Pathology and || Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center and the  Howard Hughes Medical Institute, Oklahoma City, Oklahoma 73104

Received for publication, November 22, 2000, and in revised form, November 30, 2000



    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The endothelial cell protein C receptor (EPCR) is an endothelial cell-specific transmembrane protein that binds both protein C and activated protein C (APC). EPCR regulates the protein C anticoagulant pathway by binding protein C and augmenting protein C activation by the thrombin-thrombomodulin complex. EPCR is homologous to the MHC class 1/CD1 family, members of which contain two alpha -helices that sit upon an 8-stranded beta -sheet platform. In this study, we identified 10 residues that, when mutated to alanine, result in the loss of protein C/APC binding (Arg-81, Leu-82, Val-83, Glu-86, Arg-87, Phe-146, Tyr-154, Thr-157, Arg-158, and Glu-160). Glutamine substitutions at the four N-linked carbohydrate attachment sites of EPCR have little affect on APC binding, suggesting that the carbohydrate moieties of EPCR are not critical for ligand recognition. We then mapped the epitopes for four anti-human EPCR monoclonal antibodies (mAbs), two of which block EPCR/Fl-APC (APC labeled at the active site with fluorescein) interactions, whereas two do not. These epitopes were localized by generating human-mouse EPCR chimeric proteins, since the mAbs under investigation do not recognize mouse EPCR. We found that 5 of the 10 candidate residues for protein C/APC binding (Arg-81, Leu-82, Val-83, Glu-86, Arg-87) colocalize with the epitope for one of the blocking mAbs. Three-dimensional molecular modeling of EPCR indicates that the 10 protein C/APC binding candidate residues are clustered at the distal end of the two alpha -helical segments. Protein C activation studies on 293 cells that coexpress EPCR variants and thrombomodulin demonstrate that protein C binding to EPCR is necessary for the EPCR-dependent enhancement in protein activation by the thrombin-thrombomodulin complex. These studies indicate that EPCR has exploited the MHC class 1 fold for an alternative and possibly novel mode of ligand recognition. These studies are also the first to identify the protein C/APC binding region of EPCR and may provide useful information about molecular defects in EPCR that could contribute to cardiovascular disease susceptibility.



    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The protein C anticoagulant pathway serves as the major physiologic control of clot formation (reviewed in Refs. 1-3). The pathway is initiated upon binding of thrombin to the endothelial cell surface protein thrombomodulin (TM)1 (4). The thrombin-TM complex activates protein C to activated protein C (APC). APC, in conjunction with its cofactor protein S, degrades factors Va and VIIIa on the phospholipid surface, thereby attenuating the coagulation cascade. Defects in the protein C anticoagulant pathway have been implicated as the underlying risk factors for the development of venous and arterial thrombosis (5-11).

Recent studies demonstrate that human endothelial cells express a transmembrane protein that binds both protein C and APC with high affinity (Kd approx  30 nM) (12). This molecule, named endothelial cell protein C receptor (EPCR), is an endothelial cell-specific, type 1 transmembrane protein that binds to the Gla domain of protein C and APC (13). EPCR enhances the rate of protein C activation by the thrombin-thrombomodulin complex on the endothelial cell surface (14, 15) and when reconstituted into phosphatidylcholine liposomes (16), primarily by decreasing the Km for protein C. A soluble form of EPCR is found in normal human plasma (17) and has been shown to bind to protein C and APC with an affinity similar to that of intact membrane-bound EPCR (15, 17). In contrast to the membrane-bound form, soluble EPCR blocks APC anticoagulant activity (18, 19) by blocking phospholipid interactions (19). Interestingly, soluble EPCR alters the active site of APC, suggesting that the macromolecular specificity of APC may be altered by complex formation with soluble EPCR (19). EPCR also appears to aid in the host response to sepsis since blocking EPCR-protein C interactions in baboons exacerbates the coagulation and inflammatory responses to Escherichia coli (20). Preliminary clinical studies suggest that protein C and APC supplementation is beneficial in sepsis or septic shock (21-23).

EPCR has sequence homology to members of the MHC class I/CD1 family of molecules. MHC class I/CD1 molecules are organized into the alpha 1, alpha 2, and alpha 3 domains followed by a transmembrane region and a short cytoplasmic tail. The alpha 3 domain associates noncovalently with beta -2 microglobulin, although in EPCR this domain is absent. The alpha 1 and alpha 2 domains form a ligand binding groove composed of two antiparallel alpha -helices that sit upon an 8-stranded beta -sheet platform. Although most members of the MHC/CD1 family utilize this groove to bind short peptides, it should be noted that there are exceptions. For example, the neonatal Fc receptor, which shares the MHC fold, has a closed groove that is incapable of binding peptides (24). Instead, the ligand binding interface is on the side of the neonatal Fc receptor.

In this study, we set out to identify the residues within EPCR that are involved in protein C/APC binding. Our approach was to combine data from loss-of-function alanine substitution studies with that obtained from gain-of-function epitope-mapping studies. Given that EPCR differs from other members of the MHC class 1/CD1 family in that (a) it lacks the alpha 3 domain and (b) its ligands are 62-kDa proteins rather than short peptides, these studies will also provide insight into how the MHC structural motif may have evolved to serve different modes of ligand recognition.


    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Materials-- Human thrombin (25), human protein C and APC (26), and recombinant Gla-domainless protein C (GDPC) (27) were prepared as described previously. GDPC is a truncated form of protein C lacking residues 1-46 of the N terminus. Oligonucleotides were synthesized by Operon Technologies Inc. Dulbecco's modified Eagle's medium, fetal bovine serum, and G418 were from Life Technologies, Inc. Sulfo-NHS-LC-biotin UltraLink and Immobilized NeutrAvidin Plus resin were from Pierce. Effectene transfection reagent was from Qiagen, Inc. (Valencia, CA). H-D-(gamma -Carbobenzoxy)-L-prolyl-L-arginine-p-nitroanilide diacetate (Spectrozyme PCa) was from American Diagnostica (Greenwich, CT). All other chemicals were of the highest grade commercially available.

DNA Construction and Mutagenesis-- Human (12) and murine EPCR (28) cDNAs were cloned as XhoI/NotI fragments into the multiple cloning site of the eukaryotic expression vector pcDNA3.1(-) (Invitrogen, San Diego, CA). In vitro mutagenesis to generate and select point mutations was performed using the QuickChange site-directed mutagenesis system as described by the supplier (Stratagene, La Jolla, CA). DNA manipulations to generate deletion mutants or human-murine chimeric cDNA molecules were carried out using standard DNA cloning techniques (29). Double-stranded DNA sequencing was used to verify the authenticity of the mutations.

Transient Expression of Wild-type and Variant Forms of EPCR in 293T Cells-- In the pcDNA3.1(-) vector, expression of EPCR cDNA is under the control of the human cytomegalovirus immediate-early promoter. Transient transfection of 293T cells was performed in 6-well dishes in Dulbecco's modified Eagle's medium containing 10% fetal bovine serum using Qiagen-purified pcDNA3.1(-) constructs employing the Effectene transfection reagent as described by the supplier (Qiagen).

Stable Expression of Wild-type and Variant Forms of EPCR in 293 Cells-- 293 cells were transfected as described above. 48 h post-transfection, the medium was changed to Dulbecco's modified Eagle's medium containing 10% fetal bovine serum and 400 µg/ml G418 (Life Technologies, Inc.). After 2 weeks of drug selection in which the medium was changed every 3 days, drug-resistant colonies were isolated, and the levels of cell surface EPCR expression were determined by flow cytometric analysis as described below.

Fluorescent Labeling of APC-- APC labeled at the active site with fluorescein (Fl-APC) was prepared as described by Bock (30). Briefly, 6.5 ml of 0.2 mg/ml APC was incubated with 6-fold molar excess of Nalpha -((acetylthio)acetyl)-FPR chloromethyl ketone (ATA-PPACK) (Molecular Innovations Inc., Royal Oak MI) in 100 mM Hepes, pH 7.5, 100 mM NaCl, 1 mM EDTA. The labeling reaction was allowed to proceed until the APC was 99% inactive as monitored by the loss of enzymatic activity using Spectrozyme PCa (typically 1 h at room temperature). Excess ATA-PPACK was removed by centrifuging the sample in a molecular mass 10-kDa cut-off Centricon 10 filter (Amicon Inc.) for 15 min at 6,000 × g. All subsequent steps were performed in the dark. A 10-fold molar excess of 5-iodoacetamidofluorescein and one-tenth volume of 1 M hydroxylamine (in 1 M Hepes, pH 7.4) was added to the ATA-PPACK-APC and incubated at room temperature for 2 h. Free fluorescein was removed by gel filtration on a PD-10 column (Amersham Pharmacia Biotech), and the labeled APC was stored at -70 °C.

Fluorescent Labeling of Protein C, JRK 1494, JRK 1535, JRK 1500, and JRK 1513 Monoclonal Antibodies-- Human protein C and four monoclonal antibodies against human EPCR (JRK 1535, JRK 1494, JRK 1500, and JRK 1513) were labeled with fluorescein 5-isothiocyanate as described by Goding (31).

Flow Cytometric Analysis of Fluorescein-labeled APC, Protein C, JRK 1535, JRK 1494, JRK 1500, and JRK 1513 Binding to Transfected Cells-- Adherent transfected cells were harvested by incubation at room temperature for 5 min in phosphate-buffered saline (137 mM NaCl, 8 mM Na2HPO4·7H20, 2.7 mM KCl, 1.5 mM KH2PO4) containing 0.02% EDTA. Cells were resuspended in Hanks' balanced salt solution (HBSS) containing 1% bovine serum albumin, 25 mM Hepes, pH 7.5, 3 mM CaCl2, 0.6 mM MgCl2, and 0.02% sodium azide (binding buffer). Cells (1 × 105) were incubated at room temperature with 30 nM Fl-APC or Fl-protein C or 5 µg/ml fluorescein-labeled monoclonal anti-EPCR antibodies for 15 min in the dark. After washing, the cells were resuspended in binding buffer. Bound Fl-APC, Fl-protein C, or fluorescein-labeled antibody was detected on the fluorescence-1 channel on a FACSCalibur (Becton Dickinson). The fluorescence intensity of each sample was analyzed twice.

Triton X-100 Lysis of Transfected 293T Cells-- Confluent transfected 293T cells in a 6-well dish were harvested as described above. The cells were lysed in 50 µl of phosphate-buffered saline containing 1% Triton X-100 at room temperature for 10 min. The levels of EPCR in the cell lysates were analyzed by electrophoresis and immunoblotting as described below.

Biotinylation of Cell Surface Proteins and Precipitation of Biotinylated EPCR-- Confluent transfected 293T cells in a 6-well dish were harvested as described above and resuspended in 90 µl of HBSS. The cells were surface-biotinylated with 10 µl of 5 mg/ml sulfo-NHS-LC-biotin (Pierce) at room temperature for 10 min. After pelleting, the cells were lysed in 200 µl of phosphate-buffered saline containing 1% Triton X-100 for 10 min at room temperature. To precipitate the biotinylated EPCR, 20 µl of UltraLink Immobilized NeutrAvidin Plus resin (Pierce) was added to the lysate and mixed for 1 h at room temperature. The resin was washed 3 times with 0.5 M NaCl, 20 mM Tris-HCl, pH 7.5, 0.5% Triton X-100, then washed once with Tris-buffered saline. Cell surface levels of EPCR were analyzed by electrophoresis of the resin and immunoblotting as described below.

Electrophoresis and Immunoblotting-- Electrophoresis was performed according to the method of Laemmli (32) using 4-20% SDS-polyacrylamide gels. Immunoblotting was performed using JRK 1513 anti-EPCR monoclonal antibody.

Flow Cytometric Analysis of Fl-APC Binding to EPCR Mutants-- The affinities of Fl-APC for EPCR mutants expressed on the surface of 293 cells were determined as follows. Briefly, cells were grown to confluency in T-75 flasks, detached with 0.53 mM EDTA, and suspended in 3 ml of complete HBSS buffer (HBSS containing 3 mM CaCl2, 0.6 mM MgCl2, 1% bovine serum albumin, and 0.02% NaN3). The cells were diluted 1:20 in complete HBSS buffer and incubated with increasing concentrations of Fl-APC at 4 °C for 15 min in the dark. Binding was analyzed on a FACSCalibur flow cytometer (Becton Dickinson). Values of Kd were determined by fitting binding isotherms with a hyperbolic equation using the TableCurveTM program (Jandel Scientific, San Rafael, CA). Previous studies demonstrate that this approach yielded values similar to those obtained with direct radioligand measurement (12).

Molecular Modeling of Human EPCR (hEPCR)-- The hEPCR model was constructed by homology modeling following the method of Greer (33) using the graphics program MAIN (34). The template structure was murine MHC class I H-2Kb (35) (Protein Data Bank accession number 1VAB), with which there was 51% amino acid similarity and 22% identity. The insertion loops present in EPCR between residues Ale-31 and Glu-42, Cys-101 and Glu-106, and Arg-156 and Leu-161 in EPCR were not modeled, and a turn was constructed between Arg-10 and Gln-15, where MHC-I has a 10-residue insert.

Protein C Activation on Transfected 293 Cells-- Stably transfected 293 cells in 24-well plates were washed 3 times with phosphate-buffered saline. The cells were preincubated for 5 min at room temperature with 0.5 ml of HBSS containing 25 mM Hepes, pH 7.5, 0.1% bovine serum albumin, 3 mM CaCl2, and 0.6 mM MgCl2 before the addition of 0.2 µM human protein C or 0.2 µM GDPC. Protein C or GDPC activation was initiated by the addition of 10 nM thrombin. In some cases, 0.5 µM anti-EPCR mAb JRK 1494 was added before protein C and preincubated with the cells for 10 min at room temperature. After 30 min at 37 °C, 100 µl of the reactions were stopped by the addition of 20 µl of 1.66 mg/ml antithrombin containing 20 mM EDTA. 50 µl of the supernatant was transferred into a 96-well microplate, and amidolytic activities of APC were determined toward 0.2 mM Spectrozyme PCa substrate in 20 mm Tris-HCl, pH 7.5, 150 mM NaCl. The rates of substrate cleavage were measured in a Vmax microplate reader (Molecular Devices). All determinations were performed in duplicate. Under the conditions employed in this assay, less than 10% of the protein C was activated during the assay, as determined by reference to a standard curve of fresh fully activated protein C versus absorbance units/min.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Expression of EPCR Variants in 293 Cells-- hEPCR, bovine EPCR, and murine EPCR (mEPCR) are single-chain transmembrane glycoproteins containing 221, 222, and 225 amino acids, respectively. The amino acid comparisons of human, bovine, and murine EPCR are shown in Fig. 1. cDNAs encoding human and mouse EPCR were cloned into the eukaryotic expression vector pcDNA3.1(-) and expressed in 293 cells. The apparent molecular mass of wild-type hEPCR, as determined by SDS-polyacrylamide gel electrophoresis and immunoblot analysis (~46 kDa), is approximately twice that of its predicted molecular mass (~24 kDa), consistent with the presence of carbohydrate moieties on EPCR (Fig. 2). Glutamine substitutions at the four N-glycosylation consensus sites (Asn-30, Asn-47, Asn-119, Asn-155) result in EPCR variants with decreased molecular masses (Fig. 2). The electrophoretic mobility of the mutant proteins increases proportionally with the number of N-glycosylation sites ablated. In contrast, hEPCR variant L1, which contains alanine substitutions at five residues (Ser-71, Gln-75, Thr-145, Arg-156, glu-163), exhibits the same electrophoretic mobility as wild-type hEPCR (Fig. 2).



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Fig. 1.   Amino acid sequence comparisons of human, bovine, and murine EPCR. Residues enclosed in green circles are those that, when mutated to alanine, retain APC binding activity. Residues enclosed in blue circles are those that, when mutated to alanine, result in the loss of all mAb epitopes as well as APC binding and, hence, appear to have global effects on EPCR conformation. The 10 residues marked with red asterisks are those that, when mutated to alanine, resulted in loss of APC binding but retain mAb epitopes. The only exception is Arg-81, which resulted in the loss of APC as well as JRK 1494 binding. The seven residues marked with arrows are alanine substitutions that result in the loss of either the JRK 1513 epitope (Leu-37, Thr-38, His-39) or the JRK 1494 epitope (Arg-81) or the JRK 1500 epitope (Arg-129, Glu-130, Arg-131).



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Fig. 2.   Immunoblot analysis of cell surface proteins of 293 cells transfected with hEPCR variants. 293 cells were transiently transfected with cDNAs encoding wild-type hEPCR (WT), hEPCR mutant L1, hEPCR mutant N47Q, hEPCR mutant N119Q, and hEPCR mutant N30Q/N155Q. 48 h post-transfection, the cells were harvested, and cell surface proteins were isolated as described under "Experimental Procedures." The proteins were subjected to electrophoresis in a 4-20% SDS-polyacrylamide gel under nonreducing conditions and transferred to nitrocellulose. The immunoblot was probed with JRK 1513, a monoclonal antibody against hEPCR. Molecular mass standards are on the left as indicated.

Selection of Amino Acid Residues on EPCR for Mutagenesis-- Human, bovine, and murine EPCR all bind saturated and in a Ca2+-dependent manner to Fl-APC (28). Fig. 1 shows the locations of the residues in the extracellular domain of EPCR that we selected for individual or multiple mutations to alanine. Most of the alanine substitutions were directed at conserved residues, but some were also directed at nonconserved regions of EPCR.

Interaction of EPCR Alanine Mutants with Fl-APC and Fluorescein-labeled Anti-hEPCR Monoclonal Antibodies-- The affinities of EPCR variants for Fl-APC were first qualitatively assessed by flow cytometric analysis. The cDNAs of the variants were transiently transfected into 293 cells, and Fl-APC binding was monitored on a FACSCalibur flow cytometer. The analyses were performed in the presence of 30 nM Fl-APC, which is the dissociation constant for the interaction of hEPCR with APC (36). The ligand binding properties of the cell surface EPCR variants were compared with 293 cells transfected with hEPCR (positive control) and pcDNA3.1(-) vector (negative control). Western blotting analysis of whole cell lysates and cell surface proteins was performed in parallel to qualitatively monitor the EPCR antigen levels of the variants.

The transiently transfected cells were also screened for the ability to bind fluorescein isothiocyanate-labeled anti-human EPCR monoclonal antibodies. Our laboratory has raised a panel of anti-human EPCR monoclonal antibodies, four of which are used in this study. These antibodies recognize human EPCR but not its mouse counterpart. As shown in Fig. 3, JRK 1494 and JRK 1535 mAbs block hEPCR/Fl-APC interactions, whereas JRK 1500 and JRK 1513 mAbs do not. As expected, protein C also blocks hEPCR/Fl-APC interactions, consistent with previous studies demonstrating that EPCR binds to the Gla domain of both protein C and APC (13). The effect of these four mAbs on the interaction between hEPCR and Fl-protein C is identical to that observed on the interaction between hEPCR and Fl-APC (data not shown).



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Fig. 3.   Flow cytometric analysis of Fl-APC binding to hEPCR in the presence of protein C and anti-hEPCR mAbs. 293 cells stably expressing hEPCR were pre-incubated at room temperature with 500 nM protein C, JRK 1494, JRK 1535, JRK 1500, or JRK 1513 in the presence of 3 mM CaCl2 and 0.6 mM MgCl2. After 15 min, 30 nM Fl-APC was added to the cells and incubated at room temperature for an additional 15 min. Binding of Fl-APC to the cells was analyzed by flow cytometry.

The results of the flow cytometric analyses and Western blotting analyses are summarized in Fig. 1. Residues enclosed in green circles are those that, when mutated to alanine, do not affect the binding of Fl-APC to the transfected cells. These residues include the four N-linked carbohydrate attachment sites at Asn-30, Asn-47, Asn-119, and Asn-115, suggesting that the carbohydrate moieties of EPCR are not critical for protein C/APC interactions. The 10 residues marked with red asterisks (Arg-81, Leu-82, Val-83, Glu-86, Arg-87, Phe-146, Tyr-154, Thr-157, Arg-158, and Glu-160) are those that, when mutated to alanine, result in cell surface expression but no detectable Fl-APC binding, suggesting that these residues are involved in EPCR-APC interactions. As expected, these 10 mutants do not bind to Fl-protein C. Residues enclosed in blue circles are those that, when mutated to alanine, result in the loss of detectable intracellular and cell surface expression, suggesting that these residues are critical for secondary structure integrity of EPCR. Fig. 1 also shows residues that, when mutated to alanine, result in the loss of binding to fluorescein isothiocyanate-labeled anti-EPCR monoclonal antibodies (denoted by arrows).

Determination of the Affinities of EPCR Variants for Fl-APC-- As mentioned above, the initial flow cytometric characterization of the variants was performed in the presence of 30 nM Fl-APC. Thus, to determine qualitatively the affinities of EPCR variants for Fl-APC, stably transfected 293 cells were generated. The affinities of 11 EPCR variants for Fl-APC were determined by monitoring the changes in cell fluorescence during Fl-APC titration. These variants were chosen to represent a wide range of ligand binding properties. Binding was analyzed by flow cytometry, and Kd values were determined by fitting binding isotherms to a hyperbolic equation. To correct for nonspecific binding, 293 cells transfected with pcDNA3.1(-) vector alone were utilized.

As shown in Table I, hEPCR, mEPCR, and clone 64-3 (a hEPCR-mEPCR chimera) all have similar binding affinities to Fl-APC (Kd = 31 ± 28, 46 ± 15, and 51 ± 22 nM, respectively). Alanine substitutions that result in the removal of the epitope for JRK 1500 do not influence the binding of hEPCR to Fl-APC (Kd = 55 ± 11 nM). Alanine substitution of 5 residues in the groove of hEPCR (clone L1) or removal of two N-linked carbohydrate attachment sites (clone A/D sugar) result in a modest decrease in binding affinity to Fl-APC (Kd = 161 ± 52 and 87 ± 4 nM, respectively). In contrast, the mutations E86A, R87A, F146A, Y154A, and R158A, all of which result in variants that do not bind Fl-APC as demonstrated in the initial qualitative screens (Fig. 1), decreased the affinity of hEPCR for Fl-APC greater than 30-fold (Kd values of >= 1000 nM).


                              
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Table I
Dissociation constants for the interaction of Fl-APC with EPCR variants expressed in 293 cells
The affinities of Fl-APC for EPCR variants were measured by flow cytometry as described under "Experimental Procedures." The experiments were performed in Hepes-buffered saline containing 3 mM CaCl2 and 0.6 mM MgCl2. The values correspond to the mean and the S.E. of at least two determinations.

Mutagenesis to Map Monoclonal Antibody Epitopes on hEPCR-- A potential weakness of alanine substitution mutagenesis is that lack of ligand recognition may be a consequence of destabilizations in tertiary structure or loop conformations of the native molecule rather than in the removal of ligand binding residues. We thus aimed to merge the above loss-of-function data with data obtained from gain-of-function studies. Since our anti-hEPCR mAbs do not recognize mEPCR, we generated human/mouse chimeric proteins to delineate the mAb epitopes. The overall goal is to see if the epitopes for the blocking mAbs colocalize with the 10 residues that, when mutated to alanine, lose Fl-APC binding ability. If so, this would increase our confidence in the definitive assignment of a role for Arg-81, Leu-82, Val-83, Glu-86, Arg-87, Phe-146, Tyr-154, Thr-157, Arg-158, and Glu-160 in protein C/APC binding. The cDNAs of five human-mouse chimeras were transfected into 293 cells, and ligand binding was monitored by flow cytometry. Fig. 4A shows the results of this epitope-mapping strategy. The epitope for JRK 1494, a blocking mAb, is localized to hEPCR (Trp-26-Val-116). Consistent with this map, mutation of Arg-81 to alanine abolishes Fl-JRK 1494 and Fl-APC binding but does not affect the binding of the other three mAbs (Fig. 1). Residues Val-25 to Leu-52 contain the epitope for JRK 1513, a nonblocking mAb. Mutations of Leu-37, Thr-38, and His-39 to alanine abolishes Fl-JRK 1513 binding but does not affect the binding of Fl-APC nor the other three mAbs (Fig. 1). The C-terminal half of hEPCR (Phe-113 to Cys-222) contains the epitopes for both JRK 1500 (nonblocking) and JRK 1535 (blocking). The epitope for JRK 1500 likely included residues Arg-127, Glu-129, and Arg-130 since clone 64-3 (R127A, E129A, R130A) does not bind Fl-JRK 1500 but does bind to Fl-APC as well as to the other three mAbs (Fig. 1).



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Fig. 4.   Gain-of-function mutagenesis to map mAb epitopes on hEPCR. A, schematic representation of human-mouse EPCR chimeras. The amino acid residues at the junction between the human and mouse cDNAs are shown. The cDNAs of hEPCR, mEPCR, and the six human-mouse EPCR chimeras were transfected into 293 cells, and binding to Fl-APC and fluorescein isothiocyanate-labeled mAbs was monitored by flow cytometry as described under "Experimental Procedures." The symbols + and - designate binding and lack of binding to the ligands, respectively. B, summary of the results of the loss-of-function alanine mutagenesis studies and the gain-of-function epitope-mapping studies. The 10 candidate residues for protein C/APC binding are shown in the top schematic diagram. The epitopes for the four mAbs are shown below.

Fig. 4B summarizes the results of the loss-of-function and gain-of-function studies. Five of the 10 candidate residues for protein C/APC binding (Arg-81, Leu-82, Val-83, Glu-86, Arg-87) colocalize with the epitope for the blocking mAb JRK 1494. In contrast, the epitopes for JRK 1513 and JRK 1500, both of which are nonblocking mAbs, do not colocalize with any of the 10 candidate residues for APC/protein C binding. Western blot analysis revealed that hEPCR is immunoreactive only with JRK 1513 mAb, suggesting that JRK 1494, JRK 1500, and JRK 1535 mAbs recognize conformation-dependent epitopes (data not shown).

Fig. 5 shows a three-dimensional ribbon model of the extracellular domain of human EPCR built using mouse CD1 as a structural template. The domain consists of an eight-stranded antiparallel beta -pleated sheet with two antiparallel alpha -helices (helix 1= residues 58 to 83, helix 2= residues137 to 179). In the left panel, areas shown in green are residues that can be mutated to alanine without loss of Fl-APC binding. Regions in blue, when mutated, result in loss of cell surface expression and, hence, are likely mutations that influence protein folding. Regions in red are the amino acid side chains of 7 of the 10 protein C-APC binding candidate residues. Thr-157, Arg-158, and Glu-159 are not shown since they reside in a region of EPCR that was not modeled due to lack of homology to mouse CD1. The right panel shows the epitopes for JRK 1494 (red), JRK 1535 (red), JRK 1513 (white), and JRK 1500 (pink). Based on our three-dimensional molecular model, the candidate residues for protein C/APC binding are located at the distal end of the two alpha -helical segments that form the putative ligand binding groove.



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Fig. 5.   Molecular model of hEPCR based on the murine MHC class 1 H-2Kb structure. Left panel, regions in red are the amino acid side chains of 7 of the 10 protein C/APC binding candidate residues. Thr-157, Arg-158, and Glu-159 are not shown, since they reside in a region of EPCR that was not modeled due to lack of homology to mouse MHC class 1 H-2Kb. Areas shown in green, many of which are in the groove region, could be mutated to alanine without loss of APC binding. Regions in blue, when mutated, result in loss of cell surface expression and, hence, are likely mutations that influence protein folding. Right panel, the epitopes for the nonblocking antibodies are shown in white and pink (JRK 1513 and JRK 1500, respectively) and the epitopes for the blocking antibodies are shown in red (JRK 1494 and JRK 1535). Black line, with competitor; gray area, without competitor.

Influence of EPCR Variants on Protein C Activation Rates-- Previous studies demonstrated that binding of protein C to EPCR enhances the rate of protein C activation by the thrombin-TM complex (14-16). In this study, we coexpressed EPCR variants and TM in 293 cells to confirm the requirement for protein C binding in the EPCR-dependent enhancement in protein C activation. The following cell lines were used in these studies: (a) 293 cells, (b) 293 cells stably transfected with TM cDNA, (c) 293 cells stably transfected with both TM and hEPCR, (d) 293 cells stably transfected with both TM and E86A hEPCR, and (e) 293 cells transfected with both TM and A/D sugar hEPCR. Protein C activation was performed on the cell surface in the absence or presence of JRK 1494, a mAb that blocks protein C binding to EPCR. We also determined the activation rates of Gla-domainless protein C, which is a derivative of protein C that cannot bind to EPCR but in solution is activated by the thrombin-TM complex at the same rate as full-length protein C (37). As shown in Fig. 6, the levels of protein C and GDPC activation on 293 cells is low, and protein C activation is not influenced by pre-incubation with JRK 1494. In 293 cells expressing either TM alone or TM and EPCR variants, the activation rate of GDPC is ~7-fold higher than that observed in 293 cells. This rate is relatively constant between the four TM-expressing cell lines, as is to be expected since the cell lines express similar levels of TM as measured by 125I-radiolabeled CTM 1009 anti-human TM Fab fragments (Fig. 6, bottom of graph). Interestingly, the addition of JRK 1494 to 293 cells expressing TM alone inhibited protein C activation rates approximately 2-fold. A likely explanation for this observation is that transfection of 293 cells with TM cDNA also increases the levels of cell surface EPCR. Indeed, cell surface EPCR antigen is higher in TM-expressing cells compared with 293 cells as measured by 125I-radiolabeled JRK 1535 anti-human EPCR Fab fragments (Fig. 6, bottom of graph) and by flow cytometry using fluorescently-labeled JRK 1500 and JRK 1535 mAbs (data not shown).



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Fig. 6.   Protein C activation of 293 cells expressing EPCR variants and TM. Protein C activation was performed on confluent 293 cells and on confluent 293 cells expressing either TM or TM and EPCR variants as described under "Experimental Procedures." Gray bars, activation rate of protein C; black bars, activation rate of protein C in the presence of 500 nM inhibitory anti-hEPCR mAb JRK 1494; white bars, activation rate of GDPC. The bars represent the mean, whereas the lines above the bars reflect the S.E. of at least two determinations. The number of EPCR and TM molecules expressed per cell surface are indicated on the figure as are the EPCR:TM ratios of each cell line. WT, wild type.

In the presence of hEPCR and A/D sugar hEPCR, the protein C activation rate by the thombin-TM complex is increased by 12.1- and 10.5-fold, respectively (Fig. 6). Preincubation with JRK 1494 blocked protein C activation rates to near that observed with 293 cells transfected with TM cDNA alone. In contrast, even at an EPCR:TM ratio of 8:1, E86A hEPCR does not augment protein C activation by the thrombin-TM complex on the cell surface. Taken together, these studies confirm that binding of protein C to EPCR is necessary for the EPCR-dependent enhancement in protein C activation by the thrombin-TM complex.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Although EPCR has been implicated as an important regulatory protein of coagulation and inflammation, until now no information was available related to the critical contact sites between EPCR and protein C/APC. In this study, we have identified 10 amino acids (Arg-81, Leu-82, Val-83, Glu-86, Arg-87, Phe-146, Tyr-154, Thr-157, Arg-158, and Glu-160) that, when individually mutated to alanine residues, result in EPCR variants that lose protein C/APC binding ability. We believe that these alanine substitutions reflect the removal of ligand binding residues rather than the disruption of structural integrity of EPCR for the following reasons. First, using human-mouse chimeric EPCR constructs, half of the point mutations responsible for loss of protein C/APC binding were mapped to the epitope responsible for binding one of the inhibitory antibodies (Fig. 4). Second, each of the 10 EPCR variants was screened for the ability to bind four anti-EPCR mAbs, three of which recognize conformation-dependent epitopes (Fig. 1). All of these EPCR variants retain the ability to bind the anti-EPCR mAbs, suggesting that the mutations have not perturbed the three-dimensional conformation of EPCR. The only exception is hEPCR R81A, which does not bind to JRK 1494, suggesting that alanine substitution of Arg-81 removes both protein C/APC and JRK 1494 binding. Third, molecular modeling of EPCR indicates that the 10 candidate residues are clustered in the distal end of the two alpha -helical segments of EPCR.

This study has also identified several residues in EPCR that, when mutated, result in the loss of intracellular and cell surface expression (Fig. 1). Given the importance of the protein C pathway and the roles that EPCR plays in this pathway, it follows that mutations in EPCR that impair protein C/APC binding or impair EPCR expression would likely increase thrombotic risk. This assumption appears to be supported by preliminary clinical studies by Merati et al. (38), in which an EPCR loss-of-function mutation has been identified that is more prevalent in patients with deep vein thrombosis compared with controls.

We have also provided evidence of glycosylation at all four N-glycosylation consensus sites of hEPCR (Asn-30, Asn-47, Asn-119, and Asn-155). The glycosylation contributes to nearly half of the apparent molecular mass of the molecule. Human EPCR variants containing mutations in the carbohydrate attachment sites exhibited decreased molecular masses compared with wild-type EPCR (Fig. 2). These findings are consistent with previous studies in our laboratory showing that treatment of hEPCR with endoglycosidase F/peptide-N-glycosidase reduces the apparent molecular mass from 46 to 28.5 kDa on SDS-polyacrylamide gel electrophoresis (36). The fact that glutamine substitutions at the glycosylation sites do not affect Fl-APC nor Fl-protein C binding significantly (Fig. 1) suggests that the carbohydrate moieties of EPCR are not critical for APC/protein C recognition. Instead, the N-glycosylation of hEPCR may contribute to the protection of EPCR from proteolytic degradation or may serve as specific recognition domains for other, as yet unidentified, ligands.

Our three-dimensional molecular model of hEPCR was constructed by homology modeling using the murine MHC class 1 H-2Kb as the structural template (35). The model suggests that hEPCR is folded into a ligand binding groove composed of two anti-parallel alpha -helices that sit upon an eight-stranded beta -sheet platform (Fig. 5), a fold that is characteristic of members of the MHC class1/CD1 family of receptors. The overall structure of our hEPCR model is similar to that developed by Villoutreix et al. (39) using the x-ray structure of mouse CD1 as template.

MHC class 1 molecules bind to octamer and nonamer peptides (35, 40), whereas CD1 molecules recognize lipids and glycolipids (reviewed in Ref. 41). In the case of MHC class 1 receptors, peptide ligands are tethered by hydrogen bonds between backbone atoms of the peptide and side chain residues in the alpha -helices that line the groove (42). These interactions are further supplemented by contacts of polymorphic MHC groove side chains with a few "anchor" side chains in the peptide (40). The anchor residues of the peptide occupy depressions within the MHC groove. In contrast, the interaction between CD1 molecules and lipids involves extensive hydrophobic interactions in deeply buried depressions within the CD1 groove (43).

Unlike conventional MHC class 1/CD1 proteins, EPCR lacks the alpha 3 domain, and its ligands are large proteins rather than short peptides or lipids. The results of this study indicate that the APC/protein C binding region of EPCR is located in the distal end of the two alpha -helical segments that form the putative binding groove. To our knowledge, this is the first report of such a binding motif for a member of the MHC class 1/CD1 family of molecules. There are other examples of ligand binding versatility of the MHC class1/CD1 fold. For example, the neonatal Fc receptor has a proline residue in the alpha 2 helix that produces a kink in the alpha 2 helix, resulting in a closed groove (24). Although the peptide binding groove is lost, the neonatal Fc receptor has evolved to recognize the Fc portion of immunoglobulins on the side of the Fc receptor. In the case of MIC-A, a stress-inducible antigen restricted to gut epithelium, the peptide binding groove is absent due to disordering in one of the groove-defining helices (44). Potential receptor interaction surfaces are on the "underside" of the beta -sheet platform.

The present study is the first to identify the protein C/APC binding region of EPCR. This information may provide a framework to help guide interpretation of future genetic screening studies. This work also suggests that EPCR has exploited the MHC class 1 fold for an alternative and possibly novel mode of ligand recognition.


    ACKNOWLEDGEMENTS

We thank Dr. Naomi L. Esmon for critical reading of the manuscript and for many helpful discussions. We are grateful to Nici Barnard in the preparation of this manuscript.


    FOOTNOTES

* This work was supported in part by NHLBI, National Institutes of Health Grant P01 HL54804.The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§ Recipient of a Research Fellowship from the Heart and Stroke Foundation of Canada.

Dagger Dagger An investigator with the Howard Hughes Medical Institute. To whom correspondence should be addressed: Howard Hughes Medical Institute, Oklahoma Medical Research Foundation, 825 NE 13th St., Oklahoma City, OK 73104. Tel.: 405-271-7571; Fax: 405-271-3137; E-mail: Charles-Esmon@omrf.ouhsc.edu.

Published, JBC Papers in Press, November 30, 2000, DOI 10.1074/jbc.M010572200


    ABBREVIATIONS

The abbreviations used are: TM, thrombomodulin; APC, activated protein C; EPCR, endothelial cell protein C receptor; hEPCR and mEPCR, human and murine EPCR, respectively; HBSS, Hanks' balanced salt solution; mAb, monoclonal antibody; GDPC, Gla-domainless protein C; Fl-APC, APC labeled at the active site with fluorescein; ATA-PPACK, Nalpha -((acetylthio)acetyl)-FPR chloromethyl ketone; MHC, major histocompatibility complex.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES


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