From Corvas International, Inc., San Diego,
California 92121 and the ¶ Department of Immunology, Scripps
Research Institute, La Jolla, California 92037
Received for publication, October 5, 2000, and in revised form, December 18, 2000
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Recombinant nematode anticoagulant protein c2
(rNAPc2) is a potent, factor Xa (fXa)-dependent small
protein inhibitor of factor VIIa-tissue factor (fVIIa·TF), which
binds to a site on fXa that is distinct from the catalytic center
(exo-site). In the present study, the role of other fX derivatives in
presenting rNAPc2 to fVIIa·TF is investigated. Catalytically active
and active site blocked fXa, as well as a plasma-derived and an
activation-resistant mutant of zymogen fX bound to rNAPc2 with
comparable affinities (KD = 1-10
nM), and similarly supported the inhibition of fVIIa·TF (Ki* = ~10 pM). The roles
of phospholipid membrane composition in the inhibition of fVIIa·TF by
rNAPc2 were investigated using TF that was either detergent-solubilized
(TFS), or reconstituted into membranes, containing
phosphatidylcholine (TFPC) or a mixture of
phosphatidylcholine and phosphatidylserine (TFPCPS). In the absence of the fX derivative, inhibition of fVIIa·TF was similar for
all three conditions (Ki ~1
µM), whereas the addition of the fX derivative increased
the respective inhibition by 35-, 150-, or 100,000-fold for
TFS, TFPC, and TFPCPS. The removal
of the The blood coagulation response to vascular injury or inflammation
results from a series of amplified reactions, in which several specific
zymogens of serine proteases in plasma are sequentially activated by
limited proteolysis (1). The serine protease factor VIIa
(fVIIa)1 present in the blood
specifically binds to tissue factor (TF), a transmembrane receptor
glycoprotein bound to subendothelial structures or present on the
surface of monocytic or other inflammatory cells, which accumulate at
the site of injury (2). The exposure of TF to circulating blood is the
triggering event that results in the formation of a catalytic complex
(fVIIa·TF) that initiates the amplified cascade of proteolytic events
leading to the formation of the serine protease thrombin (3). The
action of thrombin coupled with the particular rheological environment
found in diseased or damaged vascular beds, result in thrombi with
compositions that vary from platelet-rich, a characteristic of the
arterial vasculature, to fibrin-rich, platelet-poor clots, typical of
the venous vasculature (4).
The pathway leading from the formation of the fVIIa·TF complex to
thrombin proceeds through the serine protease factor Xa (fXa). Factor
Xa is formed by the proteolytic activation of the zymogen factor X (fX)
either by the fVIIa·TF complex or by the catalytic complex composed
of the serine protease factor IXa and its nonenzymatic cofactor factor
VIIIa assembled on an appropriate phospholipid surface (5). Factor Xa
catalyzes the formation of thrombin following assembly into a
macromolecular catalytic complex (prothrombinase) with the nonenzymatic
cofactor factor Va (fVa) that binds to a procoagulant phospholipid
surface, such as activated platelets or inflammatory cells adhered to
the site of vascular damage (6).
The regulation of the blood coagulation involves a variety of
components, most of which act to down-regulate the proteolytic response
initiated following vascular injury. The primary physiological inhibitor of the fVIIa·TF complex, tissue factor pathway inhibitor (TFPI), mediates one of these crucial pathways (7). The efficient inhibition of fVIIa·TF by TFPI requires binding of the inhibitor to
the active site of fXa via the second of its three Kunitz-like inhibitory domains followed by the formation of the final quaternary inhibitory complex with fVIIa·TF, in which the active site of fVIIa
is occupied by the first Kunitz domain of the inhibitor (8). A recent
study suggested that the rate-limiting step governing the inhibition of
fVIIa·TF by TFPI is the binding to fXa, which occurs while fXa is
either bound to or remains in the near vicinity of the fVIIa·TF
complex following zymogen cleavage of fX (9). Therefore, it appears
that the role of fXa in the inhibition of fVIIa·TF by TFPI is that of
an "inhibitory scaffold," upon which is built the final inhibitory
complex between the Kunitz-1 domain of TFPI and the active site of
fVIIa. This proposed mechanism requires the ternary fVIIa·TF·fXa
complex to display a limited half-life or stability. This originates,
in part, from specific protein-protein interactions outside the
catalytic center of fVIIa at exo-sites on fVIIa·TF, to which the
substrate fX or product fXa binds (10, 11).
Previously, we described a potent 84-amino acid, non-Kunitz-like
inhibitor of the fVIIa·TF complex called nematode anticoagulant protein c2 (NAPc2) that was originally isolated from the hematophagous nematode hookworm Ancylostoma caninum (12). An 85-amino acid recombinant form of NAPc2 (rNAPc2) was shown to significantly inhibit
fVIIa·TF-mediated factor IX activation, but only in the presence of
fXa, or fXa that had been irreversibly inhibited with the active-site
inhibitor Glu-Gly-Arg-chloromethylketone (EGR-fXa). The effectiveness
of EGR-fXa as an inhibitory scaffold suggested that rNAPc2 bound to a
region of fXa outside of the catalytic center. The utilization of such
an exo-site by rNAPc2 distinguishes it from TFPI, which has been shown
to require an unoccupied active site in fXa to allow the binding of the
Kunitz 2 domain of the inhibitor (7). Therefore, we propose that,
although rNAPc2 is functionally similar to TFPI with respect to the
requirement of an inhibitory scaffold to mediate its inhibition of
fVIIa·TF, it is mechanistically distinct based on the specific
binding interaction with fXa.
In this report, we characterize the interaction of rNAPc2 with fX
derivatives that support inhibition of fVIIa·TF. We demonstrate that
rNAPc2 can bind with high affinity to zymogen fX, indicating that the
activation status of the inhibitory scaffold is not crucial for the
inhibition of fVIIa·TF by rNAPc2. The phospholipid membrane composition and the Gla-domain of the fX/fXa play critical roles in the
presentation of rNAPc2 that we then show interacts directly with the
active site of fVIIa via a reactive site sequence. Together, these data
support a mechanism of fVIIa·TF inhibition by rNAPc2, which utilizes
an exo-site of either the product of this catalytic complex, fXa, or
more uniquely, the substrate zymogen fX to form the final quaternary
inhibited complex.
Materials
Hepes and Tris buffers, bovine serum albumin (BSA), CHAPS, Tween
20 and all other reagents, not indicated otherwise, were
from Sigma.
Proteins
Recombinant human factor VIIa (fVIIa) was obtained from Novo
Nordisc A/S (Gentofte, Denmark). Recombinant, human full-length tissue
factor (TF) was produced using a baculovirus expression system as
described (13). Purified human proteins factor X (fX), factor IX (fIX),
glutamylglycylarginyl chloromethyl ketone (EGR-ck) modified factor Xa
(EGR-fXa), and des-Gla-domain fXa, modified with EGR-ck
(des-Gla-EGR-fXa) were obtained from Hematologic Technologies (Essex
Junction, VT). These proteins were further purified by immunodepletion
of residual fVII. Similarly, residual fX was removed by immunodepletion
from the fIX preparations. The contaminating activities of the trace
fVII and fX in the fIX preparations were individually assessed in a
quantitative assay for the activation of fX by
fVIIa·TFPCPS and estimated at Recombinant tick anticoagulant peptide (rTAP), rNAPc2, and FLAG-rNAPc2
were expressed in the methylotropic yeast Pichia pastoris and purified to homogeneity as previously described (12). FLAG-rNAPc2 contained the 8-amino acid peptidic sequence (DYKDDDDK) at the NH2 terminus of rNAPc2. The molecular mass of the
recombinant proteins was confirmed by electrospray mass spectrometry,
and protein concentrations determined by quantitative amino acid analysis.
Mutagenesis, expression in transfected dihydrofolate
reductase-deficient Chinese hamster ovary cells, and purification of recombinant factor X (rfX) derivatives and mutants has been described elsewhere (10). Briefly, Ser195 (chymotrypsin numbering;
Ref. 16) of the catalytic triad of rfX was mutated to Ala
(rfXS195A) to eliminate proteolytic and amidolytic
activities of the resulting rfXaS195A. Activation of the
recombinant fX derivative was accomplished using Russel's viper venom,
followed by purification by gel permeation chromatography. To generate
a fX analog that was resistant to scissile bond cleavage by fVIIa·TF,
the P1 residue2
Arg15 of the activation peptide of rfXS195A was
also mutated to Gln, yielding the double mutant
rfXR15Q/S195A.
All protein preparations were judged homogeneous (>95%), following
analysis by SDS-PAGE and staining with Coomassie Brilliant Blue (17).
The concentration of human fX and fXa were determined using an
extinction coefficient at A280
(E TF Reconstitution
The synthetic phospholipids L- The p-nitroanilide-containing chromogenic peptidyl substrate
C-2081 fVIIa was synthesized as a trifluoroacetic acid salt, purified
to homogeneity using HPLC, and lyophilized. The resulting molecular
mass was confirmed by electrospray mass spectrometry. The C-2081
substrate was reconstituted in deionized water just prior to use.
Measurement of the Overall Apparent Dissociation Constant
(Ki*) for the Inhibition of fVIIa·TF-mediated
[3H]fIX activation by rNAPc2
The kinetic measurement of fVIIa·TF-mediated release of
tritiated activation peptide from radiolabeled fIX
([3H]fIX) was performed as described. (12). Briefly, a
complex of fVIIa and TFPCPS was formed in 10 mM
Hepes, 150 mM NaCl, 15 µM BSA, 3.0 mM CaCl2, pH 7.4 (designated assay buffer), for
10 min prior to adding increasing amounts of equimolar rNAPc2 and one
of the following fX derivatives: fX, recombinant fX mutants, fXa,
EGR-fXa, and a complex of fXa and rTAP (rTAP-fXa), which was formed 45 min prior to adding to the fVIIa·TF complex. Following a 30-min
incubation, the reaction was initiated by the addition of the
[3H]fIX, and initial velocities were measured over 10 min, as described previously (12). The final concentration of reactants
in a total volume of 420 µl of assay buffer was: fVIIa (50 pM), TF (2.7 nM), PCPS PLV (6.4 µM), equimolar rNAPc2 and fX derivative (0-1
nM), and [3H]fIX (200 nM,
~5 × Km), and, when included, rTAP (10 nM). The fX derivatives (0-1 nM) in the
absence of rNAPc2 had no effect on the velocity
(V0). The ratio of the inhibited reaction
velocity (Vi) to the respective uninhibited
velocity (V0) (without rNAPc2) was determined
for each concentration of fX derivative-rNAPc2 for three to six
separate experiments. These data were fit by reiterative nonlinear
regression to the quadratic Equation 1 for slow, tight-binding
inhibitors (20, 21) to give the overall equilibrium constant
(Ki*), as described previously (12, 22).
-carboxyglutamic acid-containing domain from the fX
derivative did not affect the binding to rNAPc2, but abolished the
effect of factor Xa as a scaffold for the inhibition of fVIIa·TF by
rNAPc2. The overall anticoagulant potency of rNAPc2, therefore, results from a coordinated recognition of an exo-site on fX/fXa and of the
active site of fVIIa, both of which are properly positioned in the
ternary fVIIa·TF·fX(a) complex assembled on an appropriate phospholipid surface.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
0.001% and
0.0003%,
respectively. The functional, clotting-specific activity of the
purified fIX was measured as 305 units/mg in a clotting assay, using
fIX deficient plasma (George King, Overland Park, KS). Each preparation
of a fXa derivative contained at least 85% of the
-species.
Purified human factor Xa (fXa) was prepared from fX as described
previously (14).
-palmitoyloleoyl
phosphatidylcholine (PC) and L-
-stearoyloleoyl
phosphatidylserine (PS) were obtained from Avanti Polar Lipids
(Alabaster, AL). Full-length TF apoprotein was reconstituted into
phosholipid vesicles (PLV), consisting of 75% PC (w/w) and 25% PS
(w/w) (TFPCPS), or 100% PC (TFPC) in the
presence of detergent, as previously described (19), followed by
dialysis into 10 mM Hepes, 150 mM NaCl, pH 6.5. The TFPC was used immediately following dialysis. The
diameter of the resulting vesicles was measured by light scattering
(Fine Particle Technology, Menlo Park, CA), yielding a volume-weighted Gaussian distribution centered at a mean diameter of 83 ± 20 nm (PCPS PLV) and 112 ± 20 nm (PC PLV). The concentration of
phospholipids in each preparation was determined by a colorimetric
assay for inorganic phosphorus, adapted from Ref. 9.
(Eq. 1)
The Inhibitory Effects of rNAPc2 on the Kinetics of Peptidyl Amidolytic Substrate Hydrolysis by fVIIa
Base-line Kinetic Constants for Substrate Hydrolysis in the Assay-- All studies were performed in assay buffer at ambient temperature (23.5 ± 0.7 °C). The kinetics of hydrolysis of the chromogenic substrate C-2081 by fVIIa were measured under a number of experimental conditions, prior to examining the inhibitory effects of rNAPc2 under these conditions.
Reactions were initiated by the addition of uncomplexed fVIIa (10 nM) or fVIIa (2 nM) in complex with 5 nM TF (TFS, TFPC, TFPCPS) to the individual wells of a 96-well plate (Corning), containing C-2081 (0.05-3.0 mM) in a final volume of 125 µl of assay buffer. Where indicated other reagents were added to the following final concentrations: PCPS or PC PLV (6.8 µM), EGR-fXa (20 nM with free fVIIa, or 5 nM with fVIIa·TF). Factor VIIa and TF were incubated for 10 min prior to the addition to the reaction mixture. Initial reaction velocities were measured as a linear increase in the absorbance at 405 nm (A405 nm) over 10 min at 9-s intervals, using a Thermomax kinetic microplate reader. Measurements were made under steady-state conditions, where less than 5% of the substrate was consumed. The Km was derived from the nonlinear regression fit of the averaged velocities of triplicate reactions versus the respective concentration of C-2081, using Enzfitter software (Biosoft, Cambridge, United Kingdom). Kinetic values were averaged from three independent kinetic determinations, generating the following Km values for each experimental condition: uncomplexed fVIIa (2.6 mM), uncomplexed fVIIa + EGR-fXa (1.8 mM), fVIIa·TF (all TF preparations) (range: 262-356 µM), fVIIa·TF (all TF preparations) + EGR-fXa (range: 272-345 µM).
Assessment of the Time Dependence of the Inhibition of fVIIa and fVIIa·TF by rNAPc2-- The effect of incubation time on the inhibition of fVIIa and fVIIa·TF amidolytic activity by rNAPc2 varied depending on the particular reaction condition used. The extent to which the inhibition for the eight tested reaction conditions was either kinetically "fast" or time-independent, or kinetically "slow" or time-dependent determined how the corresponding dissociation constant (Ki or Ki* ) was determined (23). The relative inhibitory potency of rNAPc2 was measured over a range of concentrations for each reaction condition by two kinetic procedures, using identical concentrations of reactants: 1) fVIIa was added to a mixture of rNAPc2 and substrate to initiate reactions (no pre-incubation between enzyme and rNAPc2), and 2) rNAPc2 was first pre-incubated with fVIIa for 30 min (inhibition in the absence of substrate), followed by addition of substrate to initiate reactions. If the apparent potency of rNAPc2, measured by each of these two procedures was equivalent, then the interaction of rNAPc2 was judged time-independent or fast. The Ki was measured in subsequent experiments, as detailed below for time-independent inhibition. In contrast, if the apparent potency of rNAPc2 measured by procedure 2 was significantly greater than that measured for procedure 1, i.e. determined to be time-dependent or slow, Ki* was subsequently measured as detailed below for time-dependent inhibition.
Measurement of the Overall Equilibrium Dissociation Constant (Ki) for the Time-independent Inhibition of fVIIa and fVIIa·TF by rNAPc2-- The potency of rNAPc2 was measured over a range of substrate concentrations in the presence of increasing concentrations of rNAPc2, and when included, of equimolar EGR-fXa or des-Gla EGR-fXa. Reactions were initiated by the addition of either free fVIIa (10 nM), or preformed fVIIa·TF complex (2 nM fVIIa with 5 nM TFS, TFPSPC, or TFPC) to premixed inhibitor and substrate in the wells of microtiter plates. All reactions were performed in triplicate, and contained in 125 µl of assay buffer, containing a range of six to eight inhibitor concentrations [I] and six substrate concentrations [S]. For reactions with free fVIIa, the concentrations were 1-20 µM ([I]) and 0.1-3.0 mM ([S]), and those with fVIIa·TF complexes were 0.5-10 µM ([I]) and 0.1-2.0 mM ([S]). The initial velocities measured over 10 min under steady-state conditions for three separate experiments were fit by reiterative nonlinear regression to Equation 2, describing a time-independent, classical, reversible competitive inhibitor, to derive the Ki value.
![]() |
(Eq. 2) |
Measurement of the Apparent Overall Equilibrium Dissociation Constant (Ki*) for the Time-dependent Inhibition of fVIIa and fVIIa·TF by rNAPc2-- Varying concentrations of rNAPc2 and equimolar EGR-fXa (0.00025-1 µM) were pre-incubated with a complex of fVIIa (2 nM) and TFS, TFPC, or TFPSPC (5 nM) for 30 min. Initial velocities were measured, following the addition of substrate (650 µM). Preliminary experiments showed that EGR-Xa in the absence of rNAPc2 had no effect on V0. Ratios of the inhibited reaction velocity (Vi) to the uninhibited velocity (V0) for each concentration of rNAPc2 were fit to the quadratic Equation 1 for slow, tight-binding inhibitors for three separate experiments to give the apparent dissociation constant (Ki*).
Determination of the Rate Constants for Inhibition and Overall Equilibrium Dissociation Constant (Ki)-- The rate constants for inhibition of fVIIa·TF by rNAPc2 for each reconstituted TF condition were measured as previously described (25). Briefly, the chromogenic substrate C-2081 (800 µM) was added to the wells of a microtiter plate, containing a range of concentrations of rNAPc2 and equimolar EGR-fXa (0-100 nM) in assay buffer. The reactions were initiated by the addition of a preformed complex of fVIIa (2 nM) and TFS, TFPC, or TFPSPC (5 nM). Progress curves generated over 60 min were analyzed using Equation 3 as described by Cha (25) and Williams (26) and detailed in ref. 24, where P is the measured absorbance defined as a function of initial (Vo) and final (Vs) steady state velocities and the apparent first-order rate constant, kobs, which describes the equilibrium from the initial to the final state.
![]() |
(Eq. 3) |
![]() |
(Eq. 4) |
Determination of rNAPc2 Binding to Inhibitory Scaffolds
Surface Plasmon Resonance Analysis-- Binding kinetics for derivatives of fX to immunocaptured FLAG-rNAPc2 were determined by surface plasmon resonance, using a BIAcore 2000 instrument (Pharmacia Biosensor). The mAb to the FLAG epitope (Anti-FLAG M1, Eastman Kodak Co.) was immobilized on the surface of a CM5 sensor chip by amine coupling, according to manufacturer's recommendations. Recombinant FLAG-rNAPc2 (0.1 mg/ml) was injected onto the sensor chip to saturate the immobilized antibody in Hepes-buffered saline containing 1 mM CHAPS, 0.005% surfactant P20, 5 mM CaCl2, pH 7.4. The kinetics of binding of various derivatives of fX were measured, following the injection of each protein at different concentrations (3-300 nM). Between runs, the Ca2+-dependent mAb was regenerated by eluting bound FLAG-rNAPc2 with 0.1 M EDTA. The kinetic binding constants (ka, kd, and KD) were determined by nonlinear regression analysis of the data from three separate experiments using software provided by the manufacturer. The association rate constant (ka) was calculated from multiple sensorgrams, representing at least five different concentrations of ligand for each experiment. The dissociation rate constant (kd) was calculated from the initial dissociation phase of the binding curves, and the equilibrium dissociation constant (KD) equaled the ratio of kd/ka (27).
Immunocapture of rNAPc2 to Immobilized fX Derivatives-- The binding of FLAG-rNAPc2 to immunocaptured fX derivatives was measured by modified enzyme-linked immunoassay detection method. A murine monoclonal antibody (mAb 10)3 produced against human fX, which was shown to recognize all fX derivatives equally well, was used for solid phase immobilization of the various fX-derived inhibitory scaffolds. mAb 10 was bound to microtiter plate wells following an overnight incubation in phosphate-buffered saline (PBS) at 4 °C followed by washing with PBS containing 0.05% Tween 20 (v/v). Blocking with PBS containing 1% BSA (w/v), and 2% mannose (w/v) was followed by washing, and the addition of a saturating amount of one of the fX derivatives (final concentration 50 nM) in Hepes-buffered saline containing 15 µM BSA. Following a 1-h incubation and washing, varying concentrations of FLAG-rNAPc2 (0-100 nM) were added and incubated for 1 h followed by washing and addition of the HRP-anti-FLAG mAb M1 conjugate. Following a 1-h incubation and washing, bound peroxidase activity was visualized using 3,3',5,5'-tetramethylbenzidine dihydrochloride hydrate and 1 N H2SO4. The end point absorbance (A450-650 nm) was read, and the data were analyzed by fitting A450-650 nm versus [fX derivative] using reiterative nonlinear regression to the binding equation Y = Bmax * [L]/(KD + [L]), where [L] represents the concentration of fX derivative, Bmax the maximal binding, and KD the measured dissociation constant. Three separate binding experiments were performed for each fX derivative, and the reported KD value represented the mean of the three resolved KD values from each of those experiments.
Identification of the Cleavage Site in the Reactive Loop of rNAPc2
rNAPc2 was added to both a preformed complex of fVIIa·TFPCPS, or TFPCPS alone (control), and incubated for 0-5 h at 37 °C. The final concentration of reactants in 300 µl of 25 mM Hepes, 150 mM NaCl, 5 mM CaCl2, pH 7.4, was 90 µM rNAPc2, 1.9 µM TFPCPS, 966 µM PCPS PLV, and 0.9 µM rfVIIa. At various time intervals (0-5 h), aliquots were quenched with EDTA and submitted to SDS-PAGE followed by Coomassie Blue staining, which demonstrated that rNAPc2 was completely cleaved by the fVIIa·TF into two distinct bands by 5 h. The control sample ran as a single band, comparable to the starting material.
The remaining sample was reduced and carboxymethylated by adding
dithiothreitol (10 mM) and heating at 100 °C for 2 min
in denaturing buffer (final concentration 0.1 M Tris, 6 M guanidine HCl, pH 8.0) followed by the addition of sodium
iodoacetamide (NaIOAc) in denaturing buffer (1 mM) and an
additional 30-min incubation in the dark at 23 °C. Additional NaIOAc
(40 mM) and dithiothreitol (43 mM) were added
and incubated sequentially, followed by separation using reverse phase
HPLC C18 column chromatography (Vydac). Fractions
containing two distinct absorbance peaks were collected, pooled, and
lyophilized. SDS-PAGE analysis confirmed that each of the two bands
from the original digestion aliquot comigrated with a corresponding
band from one of the eluted pools. The two individual purified pools
were then subjected to NH2-terminal sequence analysis
(Protein and Nucleic Acid Facility, Stanford University Medical Center,
Stanford, CA).
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
rNAPc2-mediated Inhibition of fVIIa·TF by Derivatives of Zymogen
and Activated Factor X--
Previously, the inhibition of fVIIa·TF
by rNAPc2 was shown to be dependent on fXa that was either
catalytically active or irreversibly inactivated with the peptidyl
chloromethylketone EGR-ck (EGR-fXa) (12). To further define the
repertoire of inhibitory scaffolds used by rNAPc2, we investigated
zymogen fX, and a pre-formed complex of fXa with the macromolecular
inhibitor, rTAP (rTAP· fXa), and compared their effects to those of
fXa and EGR-fXa in the activation of [3H]fIX by
fVIIa·TFPCPS. As shown in Fig.
1, increasing amounts of equimolar
mixtures of rNAPc2 and each of the fX derivatives resulted in potent
inhibition of fVIIa·TFPCPS, reflected in a concentration-dependent reduction in the activation of
[3H]fIX. In the absence of added fX derivative, rNAPc2
did not inhibit [3H]fIX activation over the same
concentration range, where complete inhibition was observed with each
derivative.
|
To determine the apparent overall equilibrium dissociation constant (Ki*) for the fX derivatives, the data shown in Fig. 1 were fit to Equation 1, which describes mechanism-independent, slow, tight-binding inhibition (Table I). These data suggested that all of the factor X derivatives tested, including zymogen fX, catalytically active and inhibited forms of fXa, were similarly effective as inhibitory scaffolds for rNAPc2 in this assay, where the measured Ki* values ranged from 4 to 13 pM.
|
We could not rule out the possibility that the result with zymogen fX, shown in Table I, was due to the interaction of rNAPc2 with fXa that could have been formed during the course of the incubation with fVIIa·TFPCPS. To directly address this issue, a recombinant form of wild-type fX (rfX) was compared with the following recombinant mutant fX derivatives, all of which contained the S195A mutation in the catalytic triad, rendering them incapable of forming a catalytically active fXa: rfXS195A, rfXaS195 (activated rfXS195A, prepared using Russel's viper venom), and rfXR15Q/S195A (resistant to proteolytic activation by fVIIa·TFPCPS due to the mutation of the scissile bond Arg15 to Gln (Ref. 10)). As shown in Table II, recombinant fX and the active-site mutant rfXS195A and rfXaS195A were indistinguishable from each other, and were only about 3-fold less potent than the corresponding plasma-derived zymogen and active enzyme counterparts. This slightly increased Ki* for rNAPc2 inhibition obtained with the recombinant forms of fX may be a reflection of subtle differences in either their post-translational modifications or purification. Despite this, the proteolytically resistant form of rfX (rfXR15Q/S195A) was equipotent to the other recombinant derivatives, demonstrating that proteolytic activation of zymogen fX was not required for the formation of a high affinity inhibitory complex with fVIIa·TF.
|
High Affinity Binding of rNAPc2 to Zymogen and Activated Forms of
fX--
The data in Table I and II suggest a possible direct
interaction of rNAPc2 with the various fX-derived scaffolds, each of which resulted in a similarly effective bimolecular complex that then
inhibited fVIIa·TF. Therefore, to quantify the energetic contribution
of the protein-protein interaction between rNAPc2 and the fX
derivatives to inhibitory complex formation, direct binding
measurements were performed with these fX derivatives and rNAPc2 that
contained the amino-terminal FLAG sequence (rFLAG-NAPc2) to facilitate
its immobilization and detection. The addition of the FLAG sequence did
not affect the inhibitory potency of rNAPc2 in the presence of EGR-fXa
in the [3H]fIX activation assay as described above (data
not shown). For surface plasmon resonance (BIAcore) measurements,
immobilized anti-FLAG antibody was used to immunocapture rFLAG-NAPc2
for determination of the binding parameters for the various fX
derivatives (Fig. 2). Human
plasma-derived fXa (Fig. 2A), EGR-fXa, and the recombinant mutant rfXaS195A all bound with high affinity to
rFLAG-NAPc2, with KD values ranging from 0.6 to
1.0 nM (Table III).
Plasma-derived zymogen fX (Fig. 2B) and the
activation-resistant recombinant mutant rfXR15Q/S195A bound
to rFLAG-NAPc2 with association rates that were ~5-10-fold lower
than those measured for the various fXa species, while the dissociation
rates of fX versus fXa were <2-fold different. A possible
explanation for the slower association kinetics of the zymogen forms
with rNAPc2 could stem from the known partial disorder of the zymogen
serine protease domain prior to its activation, and formation of the
stabilizing Ile16-Asp194 salt bridge, (28, 29)
requiring an induced fit during the association phase of the
immunocaptured FLAG-rNAPc2 with the zymogen fX.
|
|
Also shown in Table III are the results from an alternative ELISA-based assay that measured the interaction between rFLAG-NAPc2 and several derivatives of fX, which were immobilized using a fX monoclonal antibody as opposed to the surface plasmon resonance measurements which were made with immobilized FLAG-rNAPc2. In this assay, rFLAG-NAPc2 bound to fXa species, including fXa that lacks the Gla-domain, with KD values of 1.5-2.8 nM, which are comparable to those measured using the BIAcore method (Table III). However, it appeared that rFLAG-NAPc2 bound to immunocaptured, plasma-derived fX with a higher affinity, as reflected in the roughly 4-fold reduction in the KD compared with the BIAcore measurement. The time scale of the individual incubation steps in the ELISA format minimize effects of different association rates and often confer sensitivity to dissociation differences. Thus, the similarity in affinity of rNAPc2 for zymogen fX compared with those for the fXa species, measured by ELISA can be considered a reflection of the similar dissociation rates, also evident from the BIAcore measurements.
Nevertheless, both assays demonstrate highly stable complexes between of rNAPc2 and either the zymogen or the activated forms of fX. This finding provides a rational for the similar potency of all fX species in stabilizing the quaternary, inhibited complex, as demonstrated in Tables I and II.
Phospholipid Membrane Requirement for the Inhibition of fVIIa·TF by rNAPc2-- The activation of the macromolecular substrate fX by fVIIa·TF has been shown to be critically dependent on the Gla-domain-mediated phospholipid binding of fX and specific protein-protein interaction with the fVIIa·TF complex (30, 31). The influence of the supporting phospholipid surface on the ability of the fX derivatives to deliver rNAPc2 to the fVIIa·TF complex was assessed using a chromogenic assay that measured the rNAPc2-dependent inhibition of fVIIa·TF amidolytic activity. In contrast to the [3H]fIX activation assay, the amidolytic activity of fVIIa·TF is independent of a phospholipid surface, allowing for the comparison of detergent-solubilized TF (TFS) with TF reconstituted with the neutral phospholipid phosphatidylcholine (TFPC) or a mixture of phosphatidylcholine and the anionic phospholipid phosphatidylserine (TFPCPS). The inhibition of fVIIa by rNAPc2 in the absence of TF and phospholipid membrane was relatively poor in the presence or absence of the inhibitory scaffold EGR-fXa (Ki 6.8-8.8 µM; Table IV). In contrast, the presence of TF increased the inhibition of fVIIa by rNAPc2 ~7-fold (Ki = ~1 µM) and was independent on the phospholipid composition. The effect of TF on the inhibition of fVIIa by rNAPc2 is likely related to the conformational stabilization and allosteric enhancement of catalytic efficiency of the protease domain of fVIIa by TF, as evidenced by the similar magnitude of improvement (~7-fold) in the measured Km for hydrolysis of the chromogenic substrate under these experimental conditions (data shown under "Experimental Procedures"). These data demonstrate that the phospholipid surface did not directly affect the inhibition of fVIIa·TF by rNAPc2 in the absence of an appropriate inhibitory scaffold.
|
The addition of EGR-fXa had a profound effect on the inhibition of fVIIa·TF amidolytic activity, depending on the preparation of TF used. In the absence of a phospholipid membrane (fVIIa·TFS), EGR-fXa increased inhibitory potency, as reflected in ~30-fold decrease in Ki* to 28.5 nM (Table IV). The inhibition of fVIIa·TFPC by rNAPc2 was improved by ~5-fold (Ki* of 6.5 nM), compared with that with TFS, suggesting that a neutral membrane surface could facilitate quaternary complex formation. The effect of EGR-fXa was greatest, when fVII-TFPCPS was used, resulting in an ~100,000-fold decrease in Ki* (9.6 pM) compared with fVII-TFPCPS without EGR-fXa (Ki = 0.96 µM) clearly implicating the procoagulant, anionic PS component as responsible for optimizing quaternary complex formation.
The anionic Gla-domain of fX is known to play a crucial role in the interaction of this coagulation protease both with a pro-coagulant membrane surface (32), and directly with the fVIIa·TF complex (15). To further demonstrate the importance of this domain for the activity of these inhibitory scaffolds, we used fX derivatives lacking the Gla-domain. The removal of the first 45 amino acids of the Gla-domain from fXa by chymotrypsin digestion resulted in a single NH2 terminus and chromogenic substrate hydrolysis that was comparable to full-length fXa (data not shown). In addition, the binding of des-Gla-fXa to rFLAG-NAPc2 was indistinguishable from full-length fXa, demonstrating that this domain was not crucial for rNAPc2 binding (Table III). Active site-blocked des-Gla-EGR-fXa was a completely ineffective inhibitory scaffold for the inhibition of fVIIa·TF amidolytic activity by rNAPc2 (Table IV). Inhibition of fVIIa·TF in the presence of des-Gla-EGR-fXa was indistinguishable from the inhibition of fVIIa·TF without an added inhibitory scaffold under any of the TF reconstitution conditions. This indicates that the primary role of the Gla-domain of the inhibitory scaffold is to mediate a productive interaction with fVIIa·TF on the membrane surface.
Certain assumptions were made regarding the kinetic mechanism governing
the inhibition of fVIIa amidolytic activity by rNAPc2 under the various
conditions described in Table IV. These assumptions were based on the
experimental observation that the inhibition by rNAPc2 of fVIIa was
either kinetically fast (time-independent) or slow
(time-dependent) relative to the rate of substrate
hydrolysis. The experimental approach took these differences into
account, and the measured Ki or
Ki* values are listed accordingly in Table IV
for the respective fast or slow kinetic behavior. For most conditions,
the inhibition of fVIIa·TF by rNAPc2 could be best described as fast
(time-independent), reversible, competitive inhibition as shown in Fig.
3A for the inhibition of
fVIIa·TFPCPS by rNAPc2. Only upon the addition of EGR-fXa
could the kinetics of fVIIa·TF inhibition be best described as slow
(time-dependent) and competitive, as shown in Fig.
3B for the inhibition of fVIIa·TFPCPS by
rNAPc2 in the presence of EGR-fXa.
|
Analysis of representative progress curves for inhibition of amidolytic
substrate hydrolysis by fVIIa·TFS,
fVIIa·TFPC, and fVIIa·TFPCPS using Equation 3 yielded values for kobs as described under
"Experimental Procedures." The linear relationship between the
measured kobs and the concentration of
rNAPc2·EGR-fXa complex shown in Fig. 4
was consistent with a slow, competitive interaction between
rNAPc2·EGR-fXa and fVIIa·TF for all three conditions. The relative
rates of association (k1) and dissociation
(k1) derived from the data shown in Fig. 4,
were used to calculate the respective Ki values of
9.2 pM, 2.0 nM, and 8.4 nM for
fVIIa·TFPCPS, fVIIa·TFPC, and
fVIIa·TFS, which were in agreement with the
Ki* values in Table IV, determined using a
mechanism-independent approach (Table I). The calculated
k1 and k
1 values gave further insight into differences between the membrane surfaces in
fXa-dependent inhibition of fVIIa·TF by rNAPc2. By
assuming a first order dissociation, the half-life
(t1/2) of the inhibited complex was found to be
dramatically stabilized by PS (fVIIa·TFPCPS)
(t1/2 = 4.3 ± 1 h), as compared with
fVIIa·TFS (t1/2 = 15 ± 4 min) or
fVIIa·TFPC (t1/2 = 13 ± 4 min).
Although the negatively charged PS headgroups facilitated optimal
fXa-dependent association (~10-fold more rapidly than on
the PC membrane), the most striking effect of PS appeared to be the
stabilization of the inhibited complex.
|
Confirmation of the Putative Reactive Loop of rNAPc2--
The
inhibition data described above suggest that rNAPc2 either interacts
directly with the active site of fVIIa, or sterically prevents access
of the substrate to the active site. Previously, we proposed the
putative reactive loop region of rNAPc2
(42LVR45V), based on the homology to the
Ascaris family of serine protease inhibitors and to another member of
the NAP family (NAP-5, formerly AcAP5) (12). To confirm a direct
interaction between rNAPc2 and fVIIa, fVIIa·TFPCPS was
incubated with excess rNAPc2 for 5 h at 37 °C under reaction
conditions that allowed for the slow cleavage of the inhibitor.
Amino-terminal sequence analysis of two resulting peptides revealed the
sequences: NH2-1KATMQ6(C-Me), which
corresponded to the native amino terminus of rNAPc2, and
NH2-45V(C-Me)HQD50C-Me, which
contained the predicted P1' residue2 at
Val45. These data confirmed that the predicted reactive
loop of rNAPc2 directly interacts with the active site of fVIIa·TF
presumably in canonical fashion as has been described for other small
protein inhibitors of serine proteases (33, 34). It should be noted that these experiments were conducted under conditions, which favored
inhibitor cleavage, and therefore were not analogous to the conditions
used for determining the inhibitory constants described above. There is
no evidence that rNAPc2 is proteolytically cleaved under the conditions
used for determining the inhibition constants, which is similar to
other small protein serine protease inhibitors (33-35).
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
This study characterizes key interactions that allow for tight (Ki* = 10 pM) complex formation between the small protein inhibitor rNAPc2 and fVIIa·TF in the presence of an inhibitory scaffold derived from fX. Active site occupancy of fXa with small peptidyl (EGR-ck) or larger small protein (rTAP) inhibitors did not appreciably influence the affinity of rNAPc2 for fXa, strongly suggesting that this inhibitor is binding to a high affinity accessory binding site or exo-site. Since binding of rTAP to fXa does not influence the function of fXa as an inhibitory scaffold, it is clear that the binding site for rNAPc2 is distinct from the proposed accessory site used by rTAP (36). The high affinity binding of rNAPc2 to fX suggests that a binding site for rNAPc2 is also exposed in the zymogen, although ongoing biochemical and x-ray crystallographic structural studies are necessary to conclusively demonstrate whether the binding site on fXa and fX are identical. However, the high affinity binding of rNAPc2 to zymogen fX is a unique characteristic of this inhibitor that should influence its biological properties. Most notably, the biochemical data predict that rNAPc2 should form a stable complex with physiologic plasma concentrations of fX zymogen. This was recently confirmed by the finding that rNAPc2 quantitatively binds to circulating fX, when administered to humans and other species such as the rat, dog, and cynomolgus monkey (37).
The formation of this stable complex of rNAPc2 with physiological
concentrations of fX strongly support a sequential inhibitory mechanism
for rNAPc2 in vivo, as schematically depicted in Fig. 5. In step 1 of this mechanism, rNAPc2
binds to zymogen fX or activated fXa, either in solution or bound to an
appropriate anionic phospholipid surface. Although contributions of a
membrane surface to this initial interaction cannot be entirely ruled
out, rNAPc2 can form a sufficiently stable and physiologically relevant
complex with fX in the absence phospholipid. In step 2, the rNAPc2·fX inhibitory scaffold docks to fVIIa·TF. This docking is highly dependent on the phospholipid composition of the membrane, in which TF
resides. There is a strong requirement for a pro-coagulant surface
containing the anionic phospholipid PS for maximal inhibition, as
evidenced by the differential rates of association of the
rNAPc2·EGR-fXa complex with the fVIIa·TF between TFPCPS
and TFPC.
|
The interaction of the rNAPc2·fX(a) complex with membrane-associated fVIIa·TF appears to be predominately mediated by the Gla-domain of the inhibitory scaffold. However, even in the absence of a phospholipid membrane, deletion of the Gla-domain in fXa resulted in an inactive scaffold, indicating that the Gla-domain is also playing a role in docking to a binding interface formed by fVIIa and TF. A similar role of the Gla-domain in binding to the fVIIa·TF complex has been previously suggested for the substrate fX, and the fXa·TFPI complex (9, 15). Thus, the inhibitor appears to utilize a docking mode similar to that of the substrate (fX) for delivery to the active site of fVIIa, which is the last step of the reaction mechanism proposed in Fig. 5. Solid evidence of a fVIIa·TF cleavage site in the reactive loop of rNAPc2 confirms docking of the inhibitor to the active site of fVIIa. Even though the reactive loop region of rNAPc2 was shown to be cleaved by fVIIa·TF under certain reaction conditions, it is likely that there is an equilibrium reached between the cleaved and un-cleaved inhibitor (rNAPc2·fX(a)) and the enzyme (fVIIa·TF), resulting in a stable inhibitory complex, which similarly occurs with other small protein inhibitors of serine proteases (33, 34, 38).
The requirement for an inhibitory scaffold composed of a fX derivative is similar to TFPI, which, like rNAPc2, forms a stable complex with fVIIa·TF assembled on a phospholipid surface. Even though the level of potency of fVIIa·TF inhibition is quite similar between TFPI (Ki* = 8 pM (Ref. 9)) and rNAPc2 (Ki* = 10 pM), there are several significant differences between the two inhibitors that distinguish their respective mechanisms for achieving inhibition of fVIIa·TF. First and foremost is the difference between the two inhibitors with respect to the inhibitory scaffold requirement. Catalytically active fXa is exclusively required by TFPI for the inhibition of fVIIa·TF due to the requirement for an accessible catalytic center by which the second Kunitz domain of TFPI can bind to fXa (8). As described above, there is no such requirement for catalytically active fXa for rNAPc2-mediated inhibition of fVIIa·TF, as evidenced by the roughly equivalent extent of inhibition using inhibited forms of fXa (EGR-fXa and rTAP·fXa) or the zymogen fX. Both rNAPc2 and TFPI share the same requirements of an anionic phospholipid surface as well as an intact Gla-domain on the fX(a) inhibitory scaffold for maximal inhibition of fVIIa·TF. It is likely that both the protein-lipid and to some extent the protein-protein interactions of the Gla-domain with the membrane-associated fVIIa·TF complex are the principal factors responsible for the observed specificity and potency of fVIIa·TF inhibition by rNAPc2. This is also true for the TFPI·fXa complex, based on the similar degree of inhibition observed for native TFPI and a hybrid protein composed of Kunitz domain I and the light chain of fXa (39). The function of the fX(a) derivative for both rNAPc2 and TFPI appears to be the same: to properly present a canonical loop region of each inhibitor to the active site of fVIIa.
Overall, it is remarkable that such structurally unique proteins like
TFPI and rNAPc2 derived from such disparate species (vertebrates and
nematode parasites, respectively) have evolved such a similar strategy
of utilizing an inhibitory scaffold derived from fX to block blood
coagulation by inhibiting the fVIIa·TF complex. A possible
explanation for this evolutionary convergence may lie in the fact that
the ternary fVIIa·TF·fX(a) complex has a sufficient biological
half-life on cells or procoagulant surfaces to serve as a relevant
target for an inhibitory mechanism that simultaneously exploits
recognition determinants in both the docked fX(a) and the active site
of fVIIa. To our knowledge, the binding of rNAPc2 to fX provides the
first description of a substrate-mediated delivery of an inhibitor to
an enzyme complex. The binding of rNAPc2 to zymogen fX may offer a
uniquely effective strategy for inhibiting the initiation of blood
coagulation mediated by fVIIa·TF in vivo. This is based on
the expectation that circulating rNAPc2·fX complexes would rapidly
inhibit fVIIa·TF prior to the initiation of coagulation, obviating
the need for any fXa generation as would be required by TFPI. The
relative difference in affinity of the rNAPc2·fX complex
(Ki = ~10 pM) versus fX
(Km=~100-200 nM (Refs. 9 and 11)) for
fVIIa·TFPCPS would favor the formation of the quaternary
complex (rNAPc2·fX·fVIIa·TF) even under conditions where fX is
not saturated with rNAPc2. Therefore, in most cases of clinical
thrombosis where the amount of fVIIa·TF generated at the site of
vascular damage would be limited by the exposure of tissue factor,
antithrombotic efficacy would be predicted at concentrations of rNAPc2
far below the plasma concentration of fX. Indeed, clinical efficacy of
rNAPc2 has been demonstrated at a dose that yielded a plasma
concentration of approximately one tenth of the circulating fX
concentration (40). Therefore, the unique properties of rNAPc2 not only
offer future opportunities in gaining a better understanding of the
mechanistic details of fVIIa·TF-mediated initiation of blood
coagulation, but also a possible new approach to clinical anticoagulant therapy.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. W. Rote for excellent and invaluable assistance in the preparation and review of this manuscript, and Dr. S. Krishnaswamy for critical comments and review of the manuscript.
![]() |
FOOTNOTES |
---|
* 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.
§ To whom correspondence should be addressed: Corvas International, Inc., 3030 Science Park Rd., San Diego, CA 92121. Tel.: 858-455-9800; Fax: 858-455-7895; E-mail: peter_bergum@corvas.com.
Present address: Alcon Laboratories, Forth Worth, TX 76134.
Published, JBC Papers in Press, January 3, 2001, DOI 10.1074/jbc.M009116200
2 The residue nomenclature of Schecter and Berger is used (41).
3 T. S. Edgington, unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are:
fVIIa, factor VIIa;
EGR-fXa, L-glutamyl-L-glycyl-L-arginyl
chloromethyl ketone-modified factor Xa;
Gla-domain, -carboxyglutamic
acid-containing domain;
des-Gla-EGR-fXa, L-glutamyl-L-glycyl-L-arginyl
chloromethyl ketone-modified factor Xa lacking the
-carboxyglutamic
acid-containing domain;
PC, L-
-palmitoyloleoyl
phosphatidylcholine;
PS, L-
-stearoyloleoyl
phosphatidylserine;
PLV, phospholipid vesicle;
PCPS, 75% (w/w)
phosphatidylcholine, 25% (w/w) phosphatidylserine vesicles;
fX, factor
X;
fXa, factor Xa;
rfX, recombinant human factor X;
rfXS195A, recombinant human factor X containing the
site-directed mutation of S195A;
rfXR15Q/S195A, recombinant
human factor X containing site-directed mutations S195A and
R15Q;
fVIIa·TF, complex of factor VIIa and tissue factor;
rNAPc2, recombinant nematode anticoagulant protein c2;
FLAG-rNAPc2, rNAPc2 with
an additional 8-amino acid sequence (DYKDDDDK) at the NH2
terminus;
rTAP, recombinant tick anticoagulant peptide;
TF, human
full-length tissue factor;
TFs, detergent-solubilized
tissue factor;
TFPC, tissue factor reconstituted into 100%
(w/w) phosphatidylcholine vesicles;
TFPCPS, tissue factor
reconstituted into 75% (w/w) phosphatidylcholine, 25% (w/w)
phosphatidylserine vesicles;
TFPI, human tissue factor pathway
inhibitor;
HPLC, high performance liquid chromatography;
CHAPS, 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulfonic acid;
BSA, bovine serum albumin;
PBS, phosphate-buffered saline;
mAb, monoclonal
antibody;
PAGE, polyacrylamide gel electrophoresis;
ELISA, enzyme-linked immunosorbent assay.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Mann, K. G. (1999) Thromb. Haemostasis 82, 165-174[Medline] [Order article via Infotrieve] |
2. | Martin, D. M. A., Wiiger, T., and Prydz, H. (1998) Thromb. Res. 90, 1-25[CrossRef][Medline] [Order article via Infotrieve] |
3. | Rapaport, S. I., and Rao, V. M. (1992) Arterioscler. Thromb. 12, 1111-1121[Medline] [Order article via Infotrieve] |
4. | Hirsh, J., and Weitz, J. I. (1999) Semin. Hematol. 36, 118-132[Medline] [Order article via Infotrieve] |
5. | Jenny, N. S., and Mann, K. G. (1998) Thrombosis and Hemorrhage , pp. 3-27, Williams and Wilkins, Baltimore, MD |
6. | Mann, K. G., Krishnaswamy, S., and Lawson, J. H. (1992) Semin. Hematol. 29, 213-226[Medline] [Order article via Infotrieve] |
7. | Broze, G. J., Jr. (1995) Annu. Rev. Med. 46, 103-112[CrossRef][Medline] [Order article via Infotrieve] |
8. | Girard, T. J., Warren, L. A., Novotny, W. F., Likert, K. M., Brown, S. G., Miletich, J. P., and Broze, G. J., Jr. (1989) Nature 338, 518-520[CrossRef][Medline] [Order article via Infotrieve] |
9. |
Baugh, R. J.,
Broze, G. J., Jr.,
and Krishnaswamy, S.
(1998)
J. Biol. Chem.
273,
4378-4386 |
10. |
Shobe, J.,
Dickinson, C. D.,
Edgington, T. S.,
and Ruf, W.
(1999)
J. Biol. Chem.
274,
24171-24175 |
11. |
Baugh, R. J.,
Dickinson, C. D.,
Ruf, W.,
and Krishnaswamy, S.
(2000)
J. Biol. Chem.
275,
28826-28833 |
12. |
Stanssens, P.,
Bergum, P. W.,
Gansemans, Y.,
Jespers, L.,
Laroche, Y.,
Huang, S.,
Maki, S. L.,
Messens, J.,
Lauwereys, M.,
Cappello, M.,
Hotez, P. J.,
Lasters, I.,
and Vlasuk, G. P.
(1996)
Proc. Natl. Acad. Sci. U. S. A.
93,
2149-2154 |
13. | Ruf, W. (1994) Biochemistry 33, 11631-11636[Medline] [Order article via Infotrieve] |
14. | Bock, P. E., Craig, P. A., Olson, S. T., and Singh, P. (1989) Arch. Biochem. Biophys. 273, 375-388[Medline] [Order article via Infotrieve] |
15. | Ruf, W., Shobe, J., Rao, S. M., Dickinson, C. D., Olson, A., and Edgington, T. S. (1999) Biochemistry 38, 1957-1966[CrossRef][Medline] [Order article via Infotrieve] |
16. | Bode, W., Mayr, I., Baumann, U., Huber, R., and Hofsteenge, J. (1989) EMBO J. 8, 3417-3475 |
17. | Laemmli, U. K. (1970) Nature 227, 680-685[Medline] [Order article via Infotrieve] |
18. | Di Scipio, R. G., Hermodson, M. A., and Davie, E. W. (1977) Biochemistry 16, 5253-5260[Medline] [Order article via Infotrieve] |
19. | Ruf, W., Miles, D. J., Rehemtulla, A., and Edgington, T. S. (1993) Methods Enzymol. 222, 209-224[Medline] [Order article via Infotrieve] |
20. | Morrison, J. F. (1969) Biochim. Biophys. Acta 185, 269-286[Medline] [Order article via Infotrieve] |
21. | Morrison, J. F. (1982) Trends Biochem. Sci. 7, 102-105[CrossRef] |
22. | Krishnaswamy, S., Vlasuk, G. P., and Bergum, P. W. (1994) Biochemistry 33, 7897-7907[Medline] [Order article via Infotrieve] |
23. | Morrison, J. F., and Walsh, C. T. (1988) Adv. Enzymol. Relat. Areas Mol. Biol. 61, 201-301[Medline] [Order article via Infotrieve] |
24. | Håkansson, K., Tulinsky, A., Ableman, M. M., Miller, T. A., Vlasuk, G. P., Bergum, P. W., Lim-Wilby, M. S. L., and Brunck, T. K. (1995) Biorg. Med. Chem. 3, 1009-1017[CrossRef][Medline] [Order article via Infotrieve] |
25. | Cha, S. (1975) Biochem. Pharmacol. 24, 2177-2185[CrossRef][Medline] [Order article via Infotrieve] |
26. | Williams, J. W., and Morrison, J. F. (1979) Methods Enzymol. 63, 437-467[Medline] [Order article via Infotrieve] |
27. |
Kelly, C. R.,
Dickinson, C. D.,
and Ruf, W.
(1997)
J. Biol. Chem.
272,
17467-17472 |
28. | Huber, R., and Bode, W. (1978) Acc. Chem. Res. 11, 114-122 |
29. | Bode, W., Schwager, P., and Huber, R. (1978) J. Mol. Biol. 118, 99-112[Medline] [Order article via Infotrieve] |
30. |
Krishnaswamy, S.,
Field, K. A.,
Edgington, T. S.,
Morrissey, J. H.,
and Mann, K. G.
(1992)
J. Biol. Chem.
267,
26110-26120 |
31. |
Ruf, W.,
Miles, D. J.,
Rehemtulla, A.,
and Edgington, T. S.
(1992)
J. Biol. Chem.
267,
6375-6381 |
32. | Jackson, C. M. (1984) Prog. Hemost. Thromb. 7, 55-109[Medline] [Order article via Infotrieve] |
33. | Laskowski, M., Jr., and Kato, I. (1980) Annu. Rev. Biochem. 49, 593-626[CrossRef][Medline] [Order article via Infotrieve] |
34. | Bode, W., and Huber, R. (1991) Biomed. Biochim. Acta 50, 437-446[Medline] [Order article via Infotrieve] |
35. | Dunwiddie, C. T., Vlasuk, G. P., and Nutt, E. M. (1992) Arch. Biochem. Biophys. 294, 647-653[Medline] [Order article via Infotrieve] |
36. | Wei, A., Alexander, R. S., Duke, J., Ross, H., Rosenfeld, S. A., and Chang, C. H. (1998) J. Mol. Biol. 283, 147-154[CrossRef][Medline] [Order article via Infotrieve] |
37. | Vlasuk, G. P., Maki, S., Cruikshank, A., Rote, W. E., and Bergum, P. W. (1998) Blood 92 Suppl. I, 1488 |
38. | Ardelt, W., and Laskowski, M., Jr. (1991) J. Mol. Biol. 220, 1041-1053[Medline] [Order article via Infotrieve] |
39. | Girard, T. J., Macphail, L. A., Likert, K. M., Novotny, W. F., Miletich, J. P., and Broze, G. J., Jr. (1990) Science 248, 1421-1424[Medline] [Order article via Infotrieve] |
40. | Lee, A., Agnelli, G., Büller, H., Ginsberg, J., Heit, J., Rote, W., Vlasuk, G., and Gent, M. (2000) Blood 96, 491a |
41. | Schecter, I., and Berger, A. (1967) Biochem. Biophys. Res. Commun. 27, 157-162[Medline] [Order article via Infotrieve] |