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INTRODUCTION |
Activated protein C
(APC)1 functions as a potent
anticoagulant in vivo by proteolytically inactivating
factors Va and VIIIa, two cofactors that are essential for blood clot
formation (1-3). APC inactivation of factor Va is a complex process
involving at least two proteolytic cleavages. APC cleaves factor Va
rapidly at Arg506 and slowly at Arg306, and
cleavage at Arg306 totally inactivates factor Va (4-7).
Protein S is the cognate cofactor for membrane-bound APC (8), and
protein S stimulates the rate of factor Va cleavage at
Arg306 by 20-fold (9).
APC is a vitamin K-dependent anticoagulant enzyme that has
extensive structural homology to other vitamin K-dependent
enzymes (for reviews see Refs. 1-3). The N-terminal module (amino
acids 1-38) of human APC contains 9
-carboxyglutamic acid (Gla)
residues and is termed the Gla domain. The Gla domain is followed by a domain rich in aromatic residues (also known as the aromatic stack), by
two domains that are homologous to the epidermal growth factor, and
then by a serine protease domain that contains the active site.
As is true for other vitamin K-dependent plasma proteins,
APC binds via the Gla domain to membranes containing negatively charged
phospholipids in the presence of calcium ions (1-3, 10). Light
scattering experiments indicated that two elongated vitamin K-dependent proteins, prothrombin (PT) and factor X,
project radially from the surface when bound to the membrane (11), and
our fluorescence resonance energy transfer (FRET) experiments showed
that the active site of each of the vitamin K-dependent
enzymes is located far (>70 Å) above the membrane surface (12-16),
thereby indicating that they project approximately perpendicularly from
the membrane surface. In the case of membrane-bound APC, its active
site is located an average of 94 Å above the surface (assuming
2 = 2/3; Ref. 12). Our FRET study also revealed that
protein S relocates the active site of membrane-bound APC to a unique position above the membrane surface (84 Å, assuming
2 = 2/3). These FRET results therefore provide a possible structural explanation for the protein S-dependent alteration in the
APC cleavage site on factor Va from Arg506 to
Arg306.
Because of the sequence similarity of the Gla domains in different
vitamin K-dependent proteins, it has been assumed that the
Gla domains of all vitamin K-dependent enzymes must be
structurally and functionally similar (17). Consistent with this view,
exchanging the Gla domain of factor VIIa for that of APC had no affect
on APC plasma anticoagulant activity (18). On the other hand, replacing the Gla domain of factor IXa with that of factor VIIa decreased the
Vmax for factor X activation (19). Furthermore,
the membrane binding affinities of vitamin K-dependent
plasma proteins differ (e.g. Ref. 10). In addition, unique
protein-protein interactions sometimes involve the Gla domain. For
example, factor IXa, factor VIIa, and protein C have been shown to bind
specifically to collagen IV (20), tissue factor (21), and endothelial
protein C receptor (22), respectively, through the Gla domain.
Recently, Smirnov et al. replaced the Gla domain and the
aromatic stack of APC with the corresponding domains of prothrombin to
form the APC-PTGla chimera (23). Exchange of the Gla domains did not
alter the affinity for phosphatidylcholine/phosphatidylserine vesicles
significantly but did increase the rate of factor Va inactivation on
these vesicles. Furthermore, the activity of the chimera was not
increased by protein S. In this study, we test the hypothesis that the
location of the active site in the chimera may be similar to that in
the APC·protein S complex and thereby explain the increased activity
and protein S-independence exhibited by the chimera.
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EXPERIMENTAL PROCEDURES |
Reagents--
Succinimidyl acetylthioacetate, octadecylrhodamine
(OR), and 5-(iodoacetamido)fluorescein were obtained from Molecular
Probes, Eugene, OR. Dioleoylphosphatidylcholine (PC) and
dioleoylphosphatidylserine (PS) were purchased from Avanti Polar
Lipids, Alabaster, AL.
L-3-Phosphatidylcholine-1,2-di[1-14C]oleoyl
([14C]PC) was obtained from Amersham Pharmacia Biotech.
D-Phenylalanyl-L-prolyl-L-arginyl (FPR) chloromethylketone was purchased from Calbiochem. Spectrozyme PCa
and Spectrozyme TH were purchased from American Diagnostica (New York).
Proteins--
Human protein C (24), protein C-PTGla chimera
(25), thrombin (26), protein S (25), and antithrombin III (27) were prepared as before. Protein concentrations were determined using the
following values for molecular weight and
1 cm1% at 280 nm, respectively:
protein C, 62,000 and 14.5 (24); protein S, 75,000 and 9.5 (28);
thrombin, 36,000 and 17.3 (29); and antithrombin III, 56,000 and 6.0 (30).
Activation of Protein C-PTGla--
Human protein C-PTGla (3 mg)
was activated in buffer A (50 mM HEPES (pH 7.5), 150 mM NaCl) plus 5 mM EDTA by incubation with 150 µg of human thrombin at 37 °C in 10.5 ml of final volume. Complete
activation of protein C-PTGla was determined by measuring chromogenic
activity as a function of time until the activity plateaued (24). The
chromogenic activity of the chimera has previously been shown to be
equivalent to native APC (23).
Active Site-directed Labeling of Human APC-PTGla--
APC-PTGla
was generated and isolated as described previously (23). Fl-FPR-labeled
APC-PTGla and human APC were then generated essentially as described
for bovine APC (12).
Factor Va Leiden Isolation and Inactivation--
Factor V Leiden
was purified from a patient homozygous for this dimorphism by a minor
modification of the published procedure (23) in which the barium
precipitation step was omitted and cryoprecipitate was removed before
direct application to the monoclonal antibody column. The purified
factor V Leiden was then activated with thrombin as described for
normal factor V (23).
Factor Va inactivation with APC or the APC-PTGla chimera was studied as
a function of enzyme concentration as described (23). Factor Va
activity was monitored with purified factor Xa, prothrombin, and PC/PS
vesicles. Residual factor Va activity was then determined by reference
to a standard curve of prothrombin activation rate versus
factor Va concentration (23).
Phospholipid Vesicles--
PC/PS (the molar ratio of PC to PS
was 4:1) and 100% PC vesicles were prepared by sonication and
centrifugation as described previously (13). Samples containing OR were
prepared in the same way except that the desired amount of OR (in ethyl
acetate) was added to the phospholipid before lyophilization and
sonication (13). The concentrations of phospholipid and OR in a
purified vesicle sample were determined as before (12), as was
, the OR acceptor density at the vesicle surface.
Gel Filtration Chromatography--
Fl-FPR-APC-PTGla binding to
vesicles was evaluated by incubating 15 nM Fl-FPR-APC-PTGla
with 600 µM PC/PS containing a trace amount of
L-3-phosphatidylcholine-1,2-di[1-14C]oleoyl
in buffer A plus 2 mM CaCl2 for 15 min at room
temperature and then analyzing the sample by gel filtration over a
Superdex 200 FPLC column (Amersham Pharmacia Biotech). The phospholipid vesicles and any bound Fl-FPR-APC-PTGla eluted in the void volume, whereas unbound protein eluted later. The phospholipid concentration was quantified by liquid scintillation counting, whereas
Fl-FPR-APC-PTGla elution was detected by measuring fluorescein emission
intensity. The calcium dependence of Fl-FPR-APC-PTGla binding to
vesicles was examined by performing a parallel incubation and gel
filtration in buffer A plus 5 mM EDTA.
Spectral Measurements--
All spectral measurements, including
determinations of Q, JDA, and
R0 were performed as before (12).
FRET Measurements--
FRET experiments were performed as before
(12), except that the D (donor-containing) and DA (containing donor and
acceptor) microcells initially received 15 nM
Fl-FPR-APC-PTGla (the donor), whereas microcells B (blank) and A
(acceptor-containing) received 15 nM unmodified APC-PTGla.
The initial net emission intensity (Fo) was obtained
by the subtraction of the signal of B from the signals of DA, A, and D.
Samples D and B were then titrated with phospholipid vesicles lacking
the OR acceptor, whereas samples DA and A were titrated with an
equivalent amount of phospholipid vesicles containing OR. The emission
intensity of a sample was measured 5 min after each addition of
phospholipid, a time that was found to be sufficient to reach
equilibrium (i.e. a stable signal). The net intensity of D,
DA, or A (FD, FDA, and
FA, respectively) was obtained by subtracting
the signal of the background B and then correcting for dilution. The
blank signal never exceeded 0.5% of the fluorescent signal of the D or
DA samples. To compensate for any signal in the DA sample caused by
direct excitation of the acceptor, the net dilution-corrected emission
intensity of the A sample was subtracted from that of the DA sample.
The intensity of DA was then normalized by comparison with its own
initial intensity as shown below, as was that of D. Making the
reasonable assumption that the absorbance of the donor dye in the
active site is not altered by the presence of the OR at the membrane
surface, the ratio of the donor quantum yields in the D and DA samples
is given by
|
(Eq. 1)
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where F is the net dilution-corrected emission
intensity of a sample at some point in the titration, and the subscript
o is used to denote the initial intensity of the sample.
At the end of the phospholipid titration, the membrane-bound
Fl-FPR-APC-PTGla was released from the membrane surface by the addition
of 5 mM EDTA. After donor release from the membrane, the
spectral measurements were repeated to determine what fraction of the
acceptor-dependent reduction in donor emission intensity was due to Fl-FPR-APC-PTGla binding to the membrane. The
QD/QDA value used in
Equation 2 below was calculated by dividing the QD/QDA value before EDTA
addition by the QD/QDA
value after EDTA addition. This normalization procedure corrects for
the contribution of OR inner filter effects and membrane-binding
independent energy transfer to the observed total reduction in donor
emission intensity.
For experiments with protein S, 15 nM Fl-FPR-APC-PTGla was
first titrated with PC/PS(OR) vesicles until the FRET efficiency reached a constant value. Protein S was then titrated into
membrane-bound Fl-FPR-APC-PTGla up to a final concentration of 300 nM. Identical procedures were used while performing control
experiments with human Fl-FPR-APC, except that excess DTT instead of
EDTA was used to release the fluorescein-labeled heavy chain of
Fl-FPR-APC from the membrane surface (12).
Distance of Closest Approach--
When the extent of energy
transfer between randomly and uniformly distributed donor dyes in one
infinite plane and randomly and uniformly distributed acceptor dyes in
a parallel infinite plane is small, the first term in the approximate
series solution of Dewey and Hammes (31) can be used to solve for
L, the distance of closest approach between the donor and
acceptor dyes in Å,
|
(Eq. 2)
|
where
is the density of acceptor chromophores at the
membrane surface (in OR/Å2), and Ro is
the distance between donor and acceptor dyes at which FRET efficiency
is 50% (in Å). This approach is justified here because
L > 1.5 Ro (32). The extent of FRET
to the acceptor dyes at the inner surface of the 50-Å-thick phospholipid bilayer is negligible and has not been included in our calculations.
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RESULTS |
Extent of Labeling--
Human Fl-FPR-APC-PTGla was prepared as
described under "Experimental Procedures." When the fluorescein
concentration was determined as described by Bock (33), the number of
dyes per protein averaged 0.7 in our preparations of both
Fl-FPR-APC-PTGla and wild-type human Fl-FPR-APC, the same yield that we
obtained previously with bovine Fl-FPR-APC (12). For the experiments
described in this paper, the presence of nonfluorescein-labeled
APC-PTGla or APC molecules in the sample does not interfere with our
interpretation of the spectroscopic data.
Spectral Properties of Fluorescein-labeled Proteins--
The
corrected wavelength of maximum emission and the average values of
quantum yield and of steady-state anisotropy were 520 nm, 0.30, and
0.20 for human Fl-FPR-APC and Fl-FPR-APC-PTGla, the same as previously
published for bovine Fl-FPR-APC (12). Thus, there is no significant
difference in probe environment in the active sites of human and bovine
APC. Furthermore, replacing the Gla domain of APC with the Gla domain
of PT did not alter the spectral properties of the fluorescein dye and,
hence, did not detectably alter the conformation of the active site of
APC.
When PC/PS vesicles were added to either human Fl-FPR-APC or
Fl-FPR-APC-PTGla, no significant changes in fluorescein spectral properties were detected. In addition, the fluorescence lifetime of the
fluorescein (3.9 ns) was unaltered when Fl-FPR-APC-PTGla bound to
PC/PS, so the quantum yield of the fluorescein was unaffected by
membrane binding. Thus, the binding of the protein to a membrane surface did not elicit a detectable alteration in the environment of
the fluorescein dye in the active site of human APC.
Active Site to Membrane Surface Energy Transfer--
In our FRET
experiments, the fluorescein dye in the active site of the protein is
the FRET donor, whereas the rhodamine in OR is the FRET acceptor. The
rhodamine dye is positively charged at pH 7.5 and remains in the
aqueous phase, whereas the hydrophobic octadecyl aliphatic chain
partitions into the lipid bilayer, thereby anchoring the rhodamine
moiety at the aqueous-lipid interface.
When human Fl-FPR-APC-PTGla was titrated with PC/PS vesicles, only a
very small decrease in fluorescein emission intensity was detected (see
Fig. 1, -OR). However, when
Fl-FPR-APC-PTGla was titrated with PC/PS vesicles containing OR, the
fluorescein intensity decreased until the phospholipid added was
sufficient to bind all of the Fl-FPR-APC-PTGla (Fig. 1,
+OR). The association of all of the Fl-FPR-APC-PTGla
molecules with vesicles was confirmed by gel filtration (see below).
This OR-dependent decrease in fluorescein intensity results
largely from FRET from the fluorescein dyes in the active site of the
protein to the rhodamine dyes localized at the phospholipid membrane
surface. To facilitate analysis, the data in Fig. 1 were normalized and
expressed in Fig. 2 as the ratio of donor
quantum yields in the presence and absence of acceptor using Equation 1. The OR-dependent decrease in fluorescein intensity
evident in Figs. 1 and 2 shows that the fluorescein dyes are close
enough to the rhodamine dyes for FRET to occur.

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Fig. 1.
OR dependence of fluorescence of
membrane-bound Fl-FPR-APC-PTGla. Samples containing 15 nM Fl-FPR-APC-PTGla were titrated with PC/PS vesicles that
contained ( ) or lacked OR ( ). Fo is the
initial fluorescence intensity of a sample before the addition of any
vesicles; F is the emission intensity of the sample at any
point in the titration. The acceptor density ( ) in this titration
was 4.73 × 10 4 OR/Å2.
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Fig. 2.
Gla domain and lipid dependence of
Fl-FPR-APC-PTGla FRET to OR. The data of Fig. 1 are replotted here
as QDA/QD ( ). At the
end of the titration, the membrane-bound Fl-FPR-APC-PTGla was released
from the membrane by the addition of EDTA to a final concentration of 5 mM ( ). In a parallel experiment, Fl-FPR-APC was titrated
with PC/PS vesicles in the presence or absence of OR dyes ( ). The
membrane-bound Fl-FPR-APC was released from the membrane by the
addition of DTT to a final concentration of 200 mM ( ).
In another experiment, Fl-FPR-APC-PTGla (initially 15 nM)
was titrated with 100% PC vesicles ( ). The was 2.90 × 10 4 OR/Å2 in the PC experiment and 4.73 × 10 4 OR/Å2 in the others.
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For comparison, wild-type human Fl-FPR-APC was titrated in parallel
with the same stock of PC/PS vesicles used for the Fl-FPR-APC-PTGla titrations. When Fl-FPR-APC was titrated with PC/PS vesicles, no
significant change in fluorescein emission intensity was observed. However, when titrated with PC/PS vesicles containing OR (PC/PS(OR)), the Fl-FPR-APC emission decreased because of FRET, as shown by the
reduction in QDA/QD (Fig.
2).
As a control, Fl-FPR-APC-PTGla was also titrated with 100% PC vesicles
with or without OR because the PT Gla domain requires negatively
charged phospholipids to bind to a membrane surface (34). As expected,
no FRET was observed (Fig. 2). The small reduction in
QDA/QD observed with PC
vesicles (Fig. 2) results from an inner filter effect (a reduction in
detected fluorescein emission caused by the absorption of excitation
and emission light by rhodamine), not from FRET, as we have documented
elsewhere (12).
The data in Fig. 2 also show that the
QDA/QD values for
wild-type human Fl-FPR-APC titrations were always higher than those for
Fl-FPR-APC-PTGla at each point in the titration. Because the proteins
were titrated with the same PC/PS(OR) vesicles, the extent of FRET
between the fluorescein dye in the active site of membrane-bound Fl-FPR-APC-PTGla and OR dyes on the membrane surface was greater than
that between membrane-bound Fl-FPR-APC and OR. The increased efficiency
of energy transfer in the chimeric APC-PTGla relative to wild-type APC
shows that the probe in the active site of the membrane-bound chimera
is not in the same position as that in membrane-bound APC. Thus, the
active sites of membrane-bound wild-type APC and APC-PTGla are
positioned at different locations above the membrane, with the active
site of the chimera closer to the surface and/or rotated so that the
relative orientation of the donor and acceptor transition dipoles is
more parallel.
Reversibility of Energy Transfer--
At the low concentrations of
fluorescent-labeled protein and OR used in our experiments, the average
separation between free protein and OR molecules is too large for
detectable FRET to occur. Thus, if PC/PS-bound Fl-FPR-APC-PTGla or
Fl-FPR-APC is released from the membrane surface at the end of the
experiment, no FRET should occur, and the
QDA/QD value should
return to 1.0. Because vitamin K-dependent proteins require
calcium ions to bind to negatively charged phospholipid surfaces, an
excess of EDTA is commonly used to chelate the calcium ions and
dissociate the protein·membrane complex (e.g. Refs.
13-16).
However, as documented in the case of bovine Fl-FPR-APC (12), we
observed that the EDTA-stimulated dissociation of the human Fl-FPR-APC·PC/PS complex was too slow and incomplete to allow us to
use this approach for examining the reversibility of Fl-FPR-APC-to-OR FRET (data not shown). We therefore used an excess of DTT to reduce the
disulfide bond between the two chains of Fl-FPR-APC and thereby release
the fluorescein-labeled heavy chain from the vesicle surface. Upon
addition of excess DTT, the value of
QDA/QD increased to a
value close to 1.0 (0.94-0.99 depending on the acceptor density). We
have shown earlier that this small residual OR-dependent
decrease in donor intensity (Fig. 2, open triangle) that
cannot be reversed by DTT (or EDTA for the chimera; see below) is
caused by an inner filter effect (12). Thus, only the changes in
QDA/QD resulting from
membrane binding were used to calculate the distance of closest approach between the fluorescein and rhodamine dyes (i.e.
the DTT- or EDTA-reversible
QDA/QD).
Interestingly, in contrast to wild-type APC, when excess EDTA was added
to PC/PS-bound Fl-FPR-APC-PTGla, the chelation of the calcium ions
resulted in an immediate increase in fluorescein emission intensity in
the DA sample cuvette such that the value of
QDA/QD returned to a
value close to 1.0 (Fig. 2, open circle). This spectral
change results from the dissociation of Fl-FPR-APC-PTGla from the
membrane surface. The Fl-FPR-APC-PTGla release was confirmed by gel
filtration chromatography (see below). Thus, the replacement of the Gla
domain of wild-type human APC with that of the Gla domain of PT yields
a chimeric protein with the membrane-binding properties that
correspond, as expected, to PT, not APC.
Phospholipid Dependence of Fl-FPR-APC-PTGla to OR Energy
Transfer--
Because prothrombin does not bind to membrane surfaces
that lack acidic phospholipids (10), one would not expect to see FRET
if Fl-FPR-APC-PTGla was titrated with PC vesicles containing OR. As
shown in Fig. 2 (open squares), only a very small decrease in fluorescein intensity was observed when Fl-FPR-APC-PTGla was titrated with PC(OR). This OR-dependent decrease in donor
intensity was due to an inner filter effect rather than to
membrane-binding-dependent FRET, as evidenced by the fact
that the magnitude of this decrease was nearly equivalent to that
observed when Fl-FPR-APC-PTGla and Fl-FPR-APC were dissociated from the
membranes with excess EDTA and DTT, respectively (Fig. 2).
Association of Fl-FPR-APC-PTGla with Membranes Detected by Gel
Filtration--
The magnitude of FRET can be determined accurately
only if the fraction of membrane-bound Fl-FPR-APC-PTGla molecules is
known. To address this issue, Fl-FPR-APC-PTGla was incubated with a
large excess of PC/PS vesicles in the presence of Ca2+, and
the distribution of free and membrane-bound chimera was then determined
using gel filtration. Fl-FPR-APC-PTGla bound to PC/PS vesicles will
elute in the excluded volume, whereas unbound Fl-FPR-APC-PTGla will
elute in the included volume. More than 98% of the Fl-FPR-APC-PTGla
fluorescence co-eluted with the radioactive vesicles (Fig.
3A), thereby demonstrating
that essentially all of the Fl-FPR-APC-PTGla molecules can bind to the
PC/PS vesicles and participate in FRET. In contrast, no fluorescence
was detected co-eluting with the radioactive vesicle peak when the
incubation lacked Ca2+ (Fig. 3B).

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Fig. 3.
Fl-FPR-APC-PTGla binding to PC/PS
vesicles. Fl-FPR-APC-PTGla (15 nM) was incubated with
600 µM PC/PS vesicles in buffer A containing either 2 mM CaCl2 (A) or 5 mM
EDTA (B) and then chromatographed over a Superdex 200 gel
filtration column. Each fraction was examined for Fl-FPR-APC-PTGla
content by fluorescein emission intensity ( ) and for phospholipid
content
(L-3-phosphatidylcholine-1,2-di[1-14C]oleoyl)
by scintillation counting ( ).
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Distance of Closest Approach: Active Site to Membrane
Surface--
Ro, the distance at which the
efficiency of FRET is 50% efficient, was determined as before (12),
assuming that the refractive index of the medium between the donor and
acceptor is 1.4 and that the transition dipoles of the donor and
acceptor dyes are oriented randomly during the excited state lifetime
of the donor (i.e.
2 = 2/3). The spectral
overlap integral (JDA) for fluorescein in Fl-FPR-APC-PTGla and OR totaled 3.62 × 1015
M
1 cm
1nm4, and this
yielded an Ro of 50.3 Å, the same
Ro value obtained for both human and bovine
Fl-FPR-APC-to-OR FRET (12).
Because the magnitude of the observed FRET depends upon the density of
OR at the membrane surface (
), the extent of energy transfer was
determined over a range of
values. The results from eight
independent experiments at five different
values with human
Fl-FPR-APC-PTGla and from five independent experiments with human
Fl-FPR-APC are tabulated in Table I.
L, the distance of closest approach between the plane of
donor dyes in the active sites of membrane-bound enzymes and the plane
of OR acceptor dyes at the membrane surface, averaged 88.7 Å for human
Fl-FPR-APC-PTGla, whereas the average L value of the human
Fl-FPR-APC was 94.3 Å. The uncertainty noted in Table I reflects the
random experimental error in our experiments. The absolute values of
L reported in Table I are also uncertain because the
relative orientation of the donor and acceptor transition dipoles
cannot be determined experimentally, and we have assumed
2 = 2/3. As discussed previously, the uncertainty in
Ro in our experiments because of this assumption is
approximately ± 10% (Ref. 12 and references therein). However,
for the purposes of this study, the absolute value of L is
not important, whereas the relative efficiencies of FRET for the
chimera and wild-type proteins are very important. There was no
dependence of L on the method used to release
Fl-FPR-APC-PTGla from the membrane, because the same average
L value was obtained for titrations reversed by the addition
of excess EDTA or DTT.
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Table I
Active site location above the membrane surface detected by FRET to OR
Ro was 50.3 Å in all experiments. Since
2 was assumed to be 2/3 in all experiments, any difference
in FRET due to the addition of protein S is assumed to be due to
translational motion, i.e. a difference in L
rather than a rotational motion that would alter Ro.
See "Results".
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The average L value obtained for human Fl-FPR-APC-PTGla,
assuming a
2 value of 2/3, was 5.6 Å shorter than
either human or bovine Fl-FPR-APC. Because the spectral properties of
the fluorescein dye in Fl-FPR-APC-PTGla are the same as those in both
human and bovine Fl-FPR-APC, the difference in FRET efficiency between
the chimera and the wild-type proteins must arise from a difference in
the heights of the active sites of the membrane-bound proteins above
the membrane surface and/or from a difference in fluorescein
orientation (
2). A difference in
2 could
arise either from different orientations of the protease domains of the
chimeric and wild-type APC relative to the membrane surface or from
different rotational freedoms of the donor dyes in the active sites of
the two proteins. Because fluorescein anisotropy was the same for the
wild-type APC species and the chimera, we conclude that the active-site
probe has the same rotational freedom in each enzyme.
When the FRET results obtained with human Fl-FPR-APC-PTGla are compared
with those obtained with human Fl-FPR-APC, the S.D. values for the
average L values appear to overlap (Table I). However a
rigorous statistical analysis of these data using the Tukey HSD method
to compare the means (35) reveals that the average L values
for these membrane-bound proteins are different at the 98% confidence
level (p < 0.02). (The critical criterion is not
whether the limit on individual means overlap but rather whether the
limit on the differences between the means includes zero. Stated
another way, the S.E. of the differences is more important than the
differences in the S.E. values.) Therefore the difference between the
locations of the active sites of APC and APC-PTGla above the membrane
is statistically significant.
Protein S Dependence of FRET--
The active site of
membrane-bound bovine APC moves upon binding to bovine protein S (12).
Because the effect of protein S on APC has been shown to be
species-specific (36), we wanted to determine whether the protein
S-dependent alteration in the topography of membrane-bound
APC observed with the bovine proteins also occurs in the human system.
Thus, we have here examined the effect of human protein S on the
location of the active sites of membrane-bound human Fl-FPR-APC and
human Fl-FPR-APC-PTGla. Samples of human Fl-FPR-APC were first titrated
with sufficient PC/PS vesicles (±OR) to bind all of the Fl-FPR-APC and
an excess of protein S cofactor. When human protein S was titrated into these samples, little change in fluorescein emission intensity was
observed in the D sample lacking OR (data not shown). However, when
human protein S was titrated into the human Fl-FPR-APC·PC/PS sample
containing PC/PS·OR (the DA cuvette), the donor intensity decreased
until all of the Fl-FPR-APC was bound to protein S. This protein
S-dependent change in human Fl-FPR-APC emission is expressed in Fig. 4 as the relative
QDA/QD; specifically the
ratio (QDA/QD)+protein
S/(QDA/QD)-protein
S. This spectral change was saturatable with respect to protein
S concentration, suggesting that it reflects APC·protein S complex
formation. The protein S-dependent change in FRET
efficiency seen in Fig. 4 can occur either because of translational
(closer to the membrane) and/or rotational (more parallel alignment of
donor-acceptor transition dipoles) movement of the active site of
membrane-bound Fl-FPR-APC relative to the membrane surface. Assuming
that this movement is solely translational, the average height of the
fluorescein in this membrane-bound APC·protein S complex would be 84 Å above the membrane surface. Thus, the human protein S relocates the active site of human APC to the same extent observed with the bovine
proteins.

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Fig. 4.
Protein S dependence of Fl-FPR-APC and
Fl-FPR-APC-PTGla energy transfer to OR. Human Fl-FPR-APC
(initially 15 nM) was initially titrated with 50 µM PC/PS in the presence or absence of OR dyes ( = 3.38 × 10 4 OR/Å2) to a
QDA/QD value of 0.87. At
this point in the titration, human protein S was titrated into the
membrane-bound Fl-FPR-APC, and
QDA/QD decreased further
to 0.82. In the above figure, the values of
QDA/QD at different
protein S concentrations are expressed relative to the
QDA/QD of 0.87 obtained
in the absence of protein S ( ). Fl-FPR-APC-PTGla (initially 15 nM) was titrated with 50 µM concentrations of
the same PC/PS vesicles in the absence or presence of OR, and
QDA/QD decreased to 0.84, at which point human protein S was titrated into the membrane-bound
chimera sample. The values of
QDA/QD at different
protein S concentrations are expressed in the above figure relative to
the QDA/QD of 0.84 obtained in the absence of protein S ( ). In a third titration with
the same vesicles, bovine Fl-FPR-APC (initially 15 nm) was titrated
with 50 µM PC/PS vesicles in the presence or absence of
OR to yield QDA/QD value
of 0.87. Human protein S was then titrated into the sample as
indicated, and the relative
QDA/QD values shown are
calculated relative to the
QDA/QD value of 0.87 obtained in the absence of protein S ( ).
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Although human and bovine protein S appear to elicit the same
topographical change upon binding to their cognate membrane-bound enzymes, these changes are species-specific. When human protein S was
titrated into a sample of bovine Fl-FPR-APC, no change in FRET
efficiency was observed (Fig. 4). Thus, the absence of a cofactor-dependent structural change correlates with the
inability of human protein S to stimulate bovine APC function (36,
37).
Strikingly, when the human Fl-FPR-APC-PTGla·PC/PS(±OR) complex was
titrated with human protein S, no protein S-dependent
increase in the efficiency of energy transfer was observed, even when
high concentrations of protein S were added to the complex (Fig. 4). Because human protein S does not elicit any fluorescein spectral changes upon association with human Fl-FPR-APC and because
protein S does not stimulate APC-PTGla activity (23), we cannot
ascertain whether or not protein S is binding to Fl-FPR-APC-PTGla in
these experiments. However, it is clear that protein S does not elicit the same change in the topographies of APC-PTGla·PC/PS and wild-type APC·PC/PS. Given the absence of any protein S-dependent
stimulation of APC-PTGla activity (23), it is interesting that
protein S also does not have any influence on the location of the
active site of membrane-bound Fl-FPR-APC-PTGla.
Inactivation of Factor Va Leiden--
Because the active sites of
the chimera and of APC in the presence of protein S have similar
locations, it raised the possibility that the chimera and the
APC·protein S complex might be functionally similar. If the protein
S-dependent enhancement of factor Va cleavage at
Arg306 by APC (9) is caused by the movement of the APC
active site, then the chimera might cleave factor Va at
Arg306 at a rate similar to that of APC·protein S. In
factor Va Leiden, the most common source of APC resistance,
Arg506 is mutated to Gln, leaving only Arg306
available for APC inactivation of factor Va (9, 38). We therefore
compared the dose-response curves of factor Va Leiden inactivation by
APC in the presence and absence of protein S to that of the chimera in
the absence of protein S (Fig. 5).
Consistent with the above proposal, the dose-response curve for the
inactivation of factor Va by the chimera is very similar to that of APC
plus protein S, and both curves are shifted to the left relative to APC
in the absence of protein S. Consistent with earlier findings of the
Rosing group (9), protein S had a much greater effect on factor Va
Leiden inactivation than we or others had observed on normal factor Va
(23).

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Fig. 5.
Inactivation of factor Va Leiden by APC and
APC-PtGla. Factor Va Leiden (0.2 nM) was inactivated
by APC ( , ) or the chimera ( ) at the concentrations indicated
for 30 min at room temperature. When present ( ), the protein S
concentration was 70 nM. The reaction mixtures contained 10 µg/ml PC/PS vesicles. Remaining factor Va activity was determined by
prothrombinase assays containing 1 nM factor Xa, 1.4 M prothrombin, 10 µg/ml PC/PS.
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DISCUSSION |
Our previous FRET studies have revealed that a feature shared by
most coagulation enzyme·cofactor complexes is an alteration in the
topography of the membrane-bound enzyme upon binding to the cofactor.
For example, tissue factor was found to alter the location of the
active site of membrane-bound factor VIIa relative to the membrane
surface (15), and this position was dictated by the cofactor, not by
the membrane binding Gla domain of factor VIIa (39). Also, factor Va
binding to factor Xa on a membrane surface causes translocation and/or
rotation of the active site of the enzyme relative to the membrane
surface (13). In addition, the cofactor may alter the conformation of
the zymogen and/or the location of the scissile bond in the substrate
above the membrane surface, as is the case with factor Va and the
prothrombin activation intermediate, meizothrombin (16). The alignment
of enzyme active sites with the scissile bond in the substrates cannot
account for all cofactor effects, because most cofactors can stimulate zymogen activation by their cognate enzyme in the absence of membrane surfaces (40). Protein S differs from other cofactors in that it cannot
enhance factor Va inactivation by APC in the absence of phospholipid
(41). This observation makes the protein S system ideally suited to
examine the importance of aligning the active site of the enzyme with
the scissile bond of the substrate, particularly because we recently
observed that protein S moves the active site of APC as much as 10 Å closer to the membrane surface (12).
An approach to testing the functional significance of the latter
protein S-dependent change in membrane topography is to
develop an APC homologue in which the location of the active site above the membrane surface in the absence of protein S is similar to that of
the APC·protein S complex. The FRET measurements reported here
indicate that the topography of the active site of the chimera and the
APC·protein S complex are similar. That this change in topography has
functional consequences is borne out by the observation that factor Va
Leiden is inactivated on PC/PS vesicles at comparable rates by the
APC·protein S complex and by the chimera in the absence of protein S. This is true despite the fact that the chimera and APC bind PS/PC
vesicles with comparable affinity (23). This suggests that protein S
functions, at least in part, by aligning the APC active site with the
Arg306 cleavage site in factor Va.
The fact that the PT Gla domain can be substituted for the APC Gla
domain without reducing the rate of factor Va inactivation argues
strongly that the Gla domain is not directly involved in substrate
recognition by APC. It is extremely unlikely that the prothrombin Gla
domain improves the activity of the chimera in the absence of protein S
over wild-type APC because of direct interactions between the
prothrombin Gla domain and factor Va. For example, in the absence of
membranes, the heavy chain of factor Va binds intact prothrombin and
prethrombin 1, a derivative of prothrombin that lacks the Gla domain,
with the same affinity (42). The observation that APC-PTGla inactivates
membrane-bound factor Va Leiden at the same rate as the APC·protein S
complex demonstrates that replacing the Gla domain of APC with that of prothrombin is functionally equivalent to binding protein S to APC.
The nature of the FRET-detected difference in active-site locations
between membrane-bound APC and APC-PTGla cannot be determined unambiguously. Assuming that the observed increase in FRET efficiency for Fl-FPR-APC-PTGla relative to Fl-FPR-APC was due solely to translational movement, the average height of the active site above the
membrane was about 6 Å less with APC-PTGla than with APC (Table I). If
the exchange of the Gla domains also caused the active site to rotate
relative to the planar bilayer surface, then the actual change in
height caused by the domain swap could be more or less than 6 Å.
Although the FRET measurements cannot tell us the exact magnitude of
the structural change, they do show that exchanging the Gla domains
caused the active site of the enzyme to move significantly relative to
the membrane surface, which, as discussed above, probably accounts for
the selective enhancement of cleavage at Arg306 in factor Va.
Although the position of the active site above the membrane surface has
been postulated to be functionally important (12, 13, 15, 16, 43), the
data reported here constitute the first direct demonstration that
modulation of active site location can alter enzyme activity and that
the cofactor can be rendered irrelevant by the appropriate
repositioning of the active site. Thus, we conclude that
protein-dependent modulation of topography (specifically,
structure relative to the membrane surface) is an effective means to
regulate the activity of membrane-bound enzymes involved in hemostasis
and likely in other membrane-dependent processes.