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J. Biol. Chem., Vol. 275, Issue 32, 24304-24312, August 11, 2000
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From the
Received for publication, March 10, 2000, and in revised form, May 2, 2000
Urokinase-type plasminogen activator (uPA) binds
to its receptor (uPAR) with a Kd of about 1 nM. The catalytic activity of the complex is apparent at
uPA concentrations close to Kd. Other functions of
the complex, such as signal transduction, are apparent at much higher
concentrations (35-60 nM). In the present study, we show
that uPA and recombinant soluble uPAR (suPAR), at concentrations that
exceed the Kd and the theoretical saturation levels
(10-80 nM), establish novel interactions that lead to a
further increase in the activity of the single-chain uPA (scuPA)/suPAR
and two-chain uPA (tcuPA)/suPAR complexes. Experiments performed using
dynamic light scattering, gel filtration, and electron microscopy
techniques indicate that suPAR forms dimers and oligomers. The three
techniques provide evidence that the addition of an equimolar
concentration of scuPA leads to the dissociation of these dimers and
oligomers. Biacore data show that suPAR dimers and oligomers bind scuPA
with decreased affinity when compared with monomers. We postulate that
uPAR is present in equilibrium between oligomer/dimer/monomer forms.
The binding of uPA to suPAR dimers and oligomers occurs with lower
affinity than the binding to monomer. These novel interactions regulate
the activity of the resultant complexes and may be involved in uPA/uPAR
mediated signal transduction.
Urokinase-type plasminogen activator
(uPA)1 has long been
implicated in fibrinolysis. uPA uPA binds to its receptor (uPAR) with a Kd of about 1 nM (25, 26), forming a stable complex with an off-rate of
several hours. The resultant complex has been shown to be involved in
two separate biological cascades: (a) plasminogen
activation, which results in proteolytic activity, and (b)
signal transduction, which results in cell adhesion and mitogenesis.
Plasminogen activation by the complex between uPA and its receptor is
apparent at concentrations close to the determined
Kd (27-29), whereas signal transduction is apparent
only at much higher concentrations (35-65 nM) (30, 31).
The lack of agreement between the concentrations required for signal
transduction and the Kd has led to the assumption that the uPA receptor is not involved in uPA-induced signal
transduction (30).
An alternative explanation for the lack of agreement between the known
Kd and the uPA concentration needed to induce signal
transduction is the existence of additional binding epitopes in both
uPA and uPAR, which interact with lower affinity. The possible presence
of more than one kind of interaction is supported by our data (32) and
the data of others (33), which indicate that uPAR contains several
binding epitopes that participate in the binding of uPA to uPAR.
In the present paper we demonstrate the existence of such low affinity
interactions. These novel interactions correlate with a modification of
the activity of the resultant scuPA/suPAR complex and further
activation of tcuPA by suPAR. Furthermore, the low affinity interaction
stems from the capacity of suPAR to form dimers and oligomers that bind
scuPA with relatively lower affinity and dissociate by the addition of
an equimolar concentration scuPA.
Materials
scuPA, the amino-terminal fragment of scuPA (amino acids 1-135;
ATF), low molecular weight scuPA (amino acids 145-411; LMWscuPA), and
low molecular weight tcuPA, scuPA, and suPAR (in part) were all gifts
of Dr. Jack Henkin (Abbott Laboratories, Abbott Park, IL) and were
characterized as described in previous publications (32, 34).
Neutralizing polyclonal anti-uPAR antibodies were a gift of Dr. Douglas
Cines from the University of Pennsylvania. Human thrombin was obtained
from Sigma. tcuPA, Glu-plasminogen, and the plasmin substrate
Spectrozyme PL were purchased from American Diagnostica
(Greenwich, CT). Plasma was obtained from the Hadassah Hospital Blood
Bank (Jerusalem). Blood used to obtain plasma was drawn from
healthy volunteers. 450 ml of blood were collected in bags produced by
Travenol Laboratories (Ashdod, Israel), containing 63 ml of
citrate-phosphate-dextrose (CPD) solution (1.66 g of sodium citrate
(hydrous), 61 g of dextrose, 206 mg of citric acid, and 140 mg of
monobasic sodium phosphate). Plasma was separated by centrifugation at
2500 rpm for 7.5 min and at 4500 rpm for an additional 5 min to remove platelets.
Methods
Production of suPAR--
cDNA encoding soluble uPAR (amino
acids 1-277), was generated by polymerase chain reaction using
pTracer-uPAR as templates. The fragments were digested with
BglII and Xho and sub-cloned into the expression
vector (pMT/BiP/V5, Invitrogen). Wild-type suPARs was expressed in
Drosophila Schneider S2 cells (DES system, Invitrogen) as
described by the manufacturer and purified from the media using a
polyclonal anti-uPAR antibody affinity column. SuPAR from Abbott
Laboratories (see "Materials") was in use too.
Measurement of Plasminogen Activator Activity--
The
plasminogen activator activity was measured in the presence of
plasminogen and the plasmin substrate Spectrozyme PL (28). Increasing concentrations of scuPA in the absence or presence of 5 nM suPAR were incubated with 20 nM
Glu-plasminogen and 0.5 mM Spectroyme PL in
phosphate-buffered saline (PBS), pH 7.4, for 20 min. The reaction was
monitored at 405 nm. In another set of experiments, 5 nM
scuPA was incubated in the absence or presence of different
concentrations of suPAR. In control experiments, a 10-fold molar excess
of ATF or 100 nM anti-suPAR antibodies were added to suPAR
before its addition to the reaction mixture.
Kinetic analysis of the plasminogen activation was performed as
described ( (28, 35). Briefly, an equimolar concentration of the
scuPA/suPAR complex (5 nM) was incubated in the presence or
absence of an additional 5 nM suPAR and increasing
concentrations of Glu-plasminogen and Spectrozyme PL in PBS, pH
7.4, at 37 °C. The concentration of generated plasmin was calculated
during each time interval from the rate of change at 405 nm using a
standard curve made with known concentrations of plasmin. The kinetic
parameters of plasmin generation were then plotted using a
double-reciprocal plot of the rates of plasmin generation
versus plasminogen concentration.
Fibrinolysis Assessed by Release of Radioactivity--
Purified
fibrinogen was radiolabeled with 125I and resuspended in
plasma to a specific activity of 30,000 cpm/ml. Clots were formed in
16-mm diameter tissue culture wells (Costar, Cambridge, MA) by
adding 0.4 NIH units to 0.4 ml of plasma. Fibrinolysis was
measured as described previously (41). Briefly, radiolabeled fibrin
clots enriched with 100 nM plasminogen were overlaid for 1 h at 37 °C with 0.4 ml of serum containing 25 nM
plasminogen activator (scuPA or tcuPA, in the absence or presence of
increasing concentrations of suPAR), and the release of radiolabeled
soluble fibrin degradation products was measured.
Size Exclusion Chromatography (SEC)--
A TSK3000SW SEC column
(Beckman) was equilibrated with PBS. Samples (0.1 ml total volume) were
injected using a 0.2-ml loop, and the column was developed with PBS at
a flow rate of 1 ml/min. Absorbance was measured at 220 nm, and data
were collected using a Beckman Gold Chromatography system.
Dynamic Light Scattering--
Dynamic light scattering was
performed with a Dynapro-801 molecular sizing instrument (Protein
Solutions, Inc., Charlottesville, VA) equipped with a 20-µl
micro-sampling cell at room temperature (22 °C). suPAR and scuPA
were in a buffer of 20 mM Tris, pH 7, and 10 mM
MES, pH 5.0, respectively, and were concentrated to 2 mg/ml. A complex
of suPAR and scuPA was made at 1:1 ratio. All protein solutions were
filtered through a membrane of 0.02 µl porosity to remove any dust
prior adding to the micro-sampling cell. Fifteen to 20 µl of
solutions was used to measure the light-scattering signal. For each
sample, at least 10 light-scattering measurements were taken, and the
data were processed by Protein Solution's DynaLS and DYNAMICS
software, version 4.0.
Electron Microscopy of uPA and suPAR--
Rotary-shadowed
samples were prepared by spraying a dilute solution of protein (final
concentration about 15 µg/ml) in a volatile buffer (0.05 M ammonium formate, pH 7.4) and 70% glycerol onto freshly
cleaved mica and shadowing with tungsten followed by deposition of a
carbon film in a vacuum evaporator (Denton Vacuum Co., Cherry Hill, NJ)
(36-38). Preparations of scuPA, suPAR, and mixtures of the suPAR/scuPA
were made at a molar ratio of 2.5:1. The specimens were examined in a
Philips 400 electron microscope (FEI Co., Hillsboro, OR) operating at
80 kV. All experiments were repeated several times, and many
micrographs were taken of randomly selected areas to ensure that the
results were reproducible and representative. About 1000 particles were
counted to determine the prevalence of different particles.
Surface Plasmon Resonance--
Binding of scuPA to suPAR was
measured using a BIA 3000 optical Biosensor (Biacore, AB,
Sweden) (39). This method detects binding interactions in real time by
measuring changes in the refractive index at a biospecific
surface and enables association and dissociation rate constants to be
calculated. For these studies, recombinant suPAR and recombinant scuPA
were coupled to CM5 research grade sensor chip flow cells (Biacore) via
standard amine coupling procedures (40) using
N-hydroxysuccinimide/methyl-N'-[3-(dimethylamino) propyl] carbodiimine hydrochloride (Pierce) at a level of 1000 relative units each. Sensor surfaces were coated with ligands (10 µg/ml) in 10 mM NaAc buffer, pH 5.0. Following
immobilization, unreacted groups were blocked with 1 M
ethanolamine, pH 8.5. A third flow cell, similarly activated and
blocked without immobilization of protein, served as a control surface.
The binding buffer was PBS, pH 7.4, 0.005% Tween 20. Binding of scuPA
was measured at 25 °C at a flow rate of 30 µl/min for 6.7 min followed by 3 min of dissociation. The bulk shift due to changes in
the refractive index was measured using the control surface and was
subtracted from the binding signal at each condition to correct for
nonspecific signals. Surfaces were regenerated with a single 30-s pulse
of 1 M NaCl, pH 3.4, followed by an injection of binding
buffer for 1 min to remove this high salt solution. All injections were
performed in a random fashion using the RANDOM command in the automated method. The response at equilibruim (Req) was calculated as the average
response over the last 10 s of association.
Data were analyzed by both linear (Scatchard) and nonlinear regression.
Linear transformation (43) was performed using Excel 97 software,
fitting the equation Req/C = Our previous data, as well as those of other laboratories, show
that suPAR stimulates the activity of scuPA at concentrations equal or
close to the Kd (27-29). In the initial
experiments, we examined the effect of increasing the concentrations of
the ligands on the activity of the resultant complex. ScuPA (5 nM) was incubated with increasing concentrations of suPAR.
As expected, the addition of an equimolar concentration of suPAR
stimulated the plasminogen activation activity of scuPA (Fig.
1). An additional increase in the
concentration of suPAR resulted in a further increase in scuPA
activity. The stimulatory effect of suPAR was
dose-dependent and saturable. SuPAR alone had no
plasminogen activation activity. The presentation of the data as a
reciprocal plot (Fig. 1B) shows that half-maximal
stimulation was achieved at a suPAR concentration of 30 nM.
To prove that the apparent saturation resulted from a limited
scuPA/suPAR interaction and was not the result of other limiting factors, such as the amount of plasminogen, or of the chromogenic plasmin substrate, we added additional amounts of scuPA in the plateau
region of the curve (scuPA/suPAR ratio 5/80). Fig.
2 shows that by increasing the
concentration of scuPA from 5 to 10 nM at a fixed
concentration of suPAR (80 nM, which gave a maximal response at 5 nM suPAR), a further increase of scuPA
activity was obtained. In addition, Fig. 2 shows that the stimulatory
effect of suPAR on scuPA activity could be abolished by the addition of
ATF. This observation supports the conclusion that the observed plateau
was the result of a limited interaction with suPAR. The inhibitory
effect of ATF indicates that the additional interactions depend on the
occurrence of primary binding between scuPA and suPAR through the ATF.
To further support the role of suPAR in the stimulatory effect, we
added polyclonal anti-uPAR antibodies. Fig. 2 shows that the anti-uPAR
antibodies had the same inhibitory effect as ATF, whereas irrelevant
IgG has no effect on the activity of the complex (not shown).
Novel Interactions between Urokinase and Its Receptor*
,
,,,
, and§§§
Department of Clinical Biochemistry, Hebrew
University-Hadassah Medical Centers, Jerusalem, Israel IL-91120,
the Departments of § Pathology and Laboratory Medicine and
¶ Cell and Developmental Biology, University of Pennsylvania,
Philadelphia, Pennsylvania 19104, the Center for Thrombosis and
Haemostasis Research, Beth Israel Deconess Medical Center and
Harvard Medical School, Boston, Massachusetts 02215, and the
** Department of Biology, Angstrom Pharmaceuticals Inc,
San Diego, California 92121
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
/ mice show a tendency toward
spontaneous thrombosis (1-3) and are more prone to form thrombi when
exposed to endotoxin (1) or hypoxia (4) or when the uPA gene is disrupted in otherwise healthy tissue-type plasminogen
activator/ mice (1). Down-regulation of uPA expression in
wild-type mice also correlates with fibrin deposition in response to
endotoxin (5). Additional biologic functions that have been attributed to uPA and its receptor include cell adhesion (6-17) and
differentiation (18-24).
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
Req/Kd + Rmax/Kd, where
Req is the response at equilibrium, C is the concentration of analyte in solution, Kd is
the equilibrium dissociation constant, and Rmax
is the maximal specific binding to the surface. Nonlinear regression
was performed using GraphPad PRISM 2.0 fitting the binding isotherm
directly (Req = C·Rmax/(C + Kd).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (9K):
[in a new window]
Fig. 1.
The effect of increasing concentrations of
suPAR on scuPA-mediated plasminogen activation. A, the
activity of 5 nM scuPA in the presence of increasing
concentrations of suPAR ( 
). suPAR alone had no effect on plasminogen
activation(
). B, double-reciprocal plot of the data
presented above.
View larger version (14K):
[in a new window]
Fig. 2.
The effect of suPAR, suPAR + ATF, or
anti-suPAR antibodies on the activity of the scuPA/suPAR complex.
Plasminogen activation by 5 nM scuPA alone (column
1), 5 nM scuPA with 80 nM suPAR
(column 2), 10 nM scuPA with 80 nM
suPAR (column 3), 10 nM scuPA/80 nM
suPAR, and 400 nM ATF (column 4), and 10 nM scuPA/80 nM suPAR and 200 nM
anti-suPAR antibodies (column 5). The activity was
determined as described in the legend for Fig. 1.
Fig. 3 shows that at a scuPA/suPAR ratio
of 1:2, the Vmax, but not the
Km, was greater than with equimolar concentrations. The increase in the Vmax could also be the
result of contamination of suPAR by uPA, but this possibility could be
excluded because suPAR alone had no activity (Fig. 1) and ATF inhibited
the activity. Another possibility is the presence in suPAR of a
protease that cleaves scuPA to tcuPA. To exclude this possibility, we
incubated scuPA with suPAR for 24 h. Under these conditions, scuPA
continued to migrate as a single band when analyzed by reducing
SDS-PAGE as described elsewhere (28) (data not shown).
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These results suggest that suPAR establishes novel interactions with
scuPA that are apparent only at higher concentrations of the ligands
and that participation of these binding epitopes induces additional
conformational changes in the complex, leading to further activation.
To support these conclusions, we used a clot lysis assay. Fig.
4A shows that the
scuPA-mediated fibrinolysis of human plasma-derived clots was
stimulated in a dose-dependent and saturable manner by
suPAR concentrations that exceeded those of scuPA and the
Kd.
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In previous publications, we showed that suPAR had almost no effect on tcuPA activity when both ligands were present in equimolar concentrations (28, 41). The next question to be addressed was, therefore, whether interactions established at higher concentrations of suPAR affected tcuPA activity.
Fig. 4B shows that tcuPA-mediated fibrinolytic activity on human plasma clots was stimulated by suPAR. Marginal stimulation (10%) was observed at a 1:1 concentration ratio, and a further increase of suPAR concentration induced a dose-dependent and saturable stimulation. Maximal stimulation (more than 5-fold) was obtained at tcuPA/suPAR ratios of 1:6. To exclude the possibility that the stimulation was the result of suPAR activity on contaminating scuPA, tcuPA was analyzed by SDS-PAGE. Overloaded gels did not reveal the presence of scuPA in the preparation (not shown).
To determine whether the effect of suPAR was due to binding to tcuPA or to some other mechanism, we examined the effect of ATF on the stimulation by suPAR. Fig. 4B shows that the effect of suPAR on tcuPA could be completely inhibited by ATF and that suPAR had no effect on the activity of LMWuPA. Anti uPAR antibodies abolished the effect of suPAR on tcuPA (not shown). In addition, ATF inhibited the stimulatory effect of suPAR on scuPA-mediated fibrinolysis, and suPAR had no effect on LMWscuPA activity (Fig. 4A).
It is widely accepted that uPA binds to suPAR at a 1:1 ratio. uPA binds to its receptor with high affinity and a Kd of 1 nM. The observation that suPAR exerts a stimulatory effect at concentrations that exceed the 1:1 ratio and the theoretical saturating concentrations suggests the existence of another kind of interaction of suPAR with uPA. This additional interaction proceeds with relatively lower affinity and leads to additional functional modification of the uPA/uPAR complex. This hypothesis is supported by the observation that the activity of the resultant complex is higher and by data from the literature that demonstrate that uPA-induced signal transduction is observed only at concentrations that are much higher than would be predicted from the Kd of the bimolecular complex formation (30, 42).
In an attempt to verify the existence of previously unknown
interactions between uPA and uPAR, we used several different
approaches. First, we used the size exclusion technique. Fig.
5A shows that suPAR exhibited
a complex equilibrium when analyzed by SEC-HPLC. SuPAR migrated in two
peaks at 4.7 and 8.1 min retention time (RT). In
contrast, scuPA migrated as a single peak of 10.9 RT (not shown). Fig. 5B shows
that on SDS-PAGE, scuPA and suPAR injected into the HPLC run as single
bands.
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In an attempt to examine the effect of scuPA on the equilibrium between the different forms of suPAR, we incubated scuPA with suPAR at several molar ratios. Fig. 5B shows the results of the addition of scuPA at a 1:2 ratio (suPAR excess). Under these conditions, no monomeric scuPA was present, and the second suPAR peak migrated at 8.4 rather than 8.1 min. SDS-PAGE analysis of the protein present in this peak demonstrated that it represents a scuPA-suPAR complex (Fig. 6B). Fig. 5C presents the results of the addition of scuPA at a 1:1 ratio. Under these conditions, scuPA shifted both peaks of suPAR to a peak of RT = 8.4 min, which represents the scuPA/suPAR complex (Fig. 6B). These data suggest that the higher molecular mass form of suPAR is able to interact with scuPA, albeit at a lower affinity.
It should be noted that the monomeric scuPA-suPAR complex elutes with a RT (8.4 min) comparable with the 8.1-min peak of suPAR. This fact suggest that the suPAR that eluted with RT = 8.1 min is a dimer.
The size exclusion technique shows the status of the isolated ligands
and complexes but does not provide information as to the situation
under equilibrium conditions. To examine the status of the reactants
under equilibrium conditions, we used dynamic light scattering. The
dynamic light scattering measures translational diffusion coefficients
(DT) of proteins through monitoring the scattered light of protein in solution. Based on
DT, the hydrodynamic radius
(Rh) of the protein can be calculated using the Stokes-Einstein equation: Rh = kbT/6
DT,
with Boltzmann's constant (kb), solvent viscosity
(), and temperature Kelvin (T). The molecular
weight of the protein is then estimated by fitting the measured
Rh to a standard curve derived from 25 globular
proteins. Proteins that are either nonspherical in shape or partially
unfolded will show apparent molecular weights that are higher than
their theoretical molecule weights.
Fig. 7A shows the
regularization histogram of scuPA, and illustrates that scuPA is
primarily a single species in aqueous solution. Using a monomodal
cumulatant analysis, scuPA was found to have an averaged hydrodynamic
radius (Rh) of 3.77 nm with 16% polydispersity (0.6 nm), corresponding to an apparent molecular mass of 73 kDa,
based on the standard size/weight relationship for globular
proteins.
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The matrix-assisted laser desorption ionization (MALDI) mass spectrum of scuPA and SDS-PAGE of scuPA gave a molecular mass of 48,140 Da and ~47 kDa, respectively (data not shown), and both are close to its theoretical molecular mass, 46,511 Da, derived from protein sequence. The difference of apparent molecular weight from mass when compared with theoretical calculation is believed to be due to the contribution of the carbohydrate present in the protein.
In contrast to the monodispersity of scuPA, the regularization histogram of suPAR (Fig. 7B) reveals some large aggregates with an Rh ~40 nm along with small size aggregates (Rh ~4 nm). Monomodal cumulatant analysis gives an Rh of 5.32 nm and a 45% polydispersity (2.4 nm), corresponding to a molecular mass of 167 kDa. This large polydispersity also clearly suggests the existence of aggregates. A bimodal cumulatant analysis, assuming the existence of two species of suPAR in the aqueous solution, suggests that 98% of suPAR has an Rh of 4.2 nm, corresponding to an apparent molecular mass of 97.7 kDa, and the remaining 2% has an Rh of 19.3 nm, corresponding to an apparent molecular mass of 5109 kDa.
Upon combination of suPAR with scuPA at a 1:1 ratio, the light-scattering pattern appears to be monodispersed with an Rh of ~4.1 nm and 19% polydispersity, corresponding to a molecular mass of 91 kDa (Fig. 7C). The lack of a signal at Rh = 3.77 demonstrates the absence of uncomplexed scuPA. These data are consistent with the dissociation of suPAR dimers for the preferential formation of 1:1 scuPA:suPAR complexes.
The scuPA/suPAR complex is smaller in size (4.1 nm) than the suPAR dimer (4.2 nm). This finding is in agreement with the SEC data showing that the scuPA/suPAR complex elutes with a longer RT as compared with suPAR dimers (RT = 8.4 versus 8.1 min). The combined data of SEC and dynamic light scattering suggest a more compact and smaller hydrodynamic radius in the scuPA-suPAR complex.
We also used electron microscopy to determine the nature of the
complexes formed between scuPA and suPAR. Rotary shadowed preparations
of suPAR were examined by electron microscopy. The protein was
initially at a concentration of 20 µM and was diluted about 200× into 0.05 M ammonium formate at pH 7.4 and 70%
glycerol just before spraying onto mica, prior to further preparation
for microscopy. Electron microscopic examination revealed two different particles (Fig. 8A). The
smaller of the two was a globular particle with a diameter of about 10 nm or about 7-8 nm after subtraction of the thickness of the coating
of metal. The larger particle consisted of two of the smaller nodules
side-by-side to form dimers. Under the buffer and protein concentration
conditions of this experiment, 41% of the particles or 55% of the
protein were in the form of dimers and 2% of the particles or 5% of
the protein were trimers, whereas the remainder were monomers. It
should be noted, however, that these numbers cannot be compared
directly with the SEC or light-scattering results because of the
different buffer and protein concentrations.
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Rotary-shadowed preparations of scuPA were also examined by electron microscopy (Fig. 8B). These molecules appeared to be globular particles just slightly larger in size than the suPAR particles, but they were entirely monomeric. The difference in size is consistent with the difference in molecular mass; scuPA has 411 amino acids, whereas suPAR has 282 amino acid residues. Some particles appeared to be oblong with an axial ratio of about 0.85, while others were nearly the same in both dimensions. These two types of images may represent different views of the same structure.
Electron microscopy (EM) was also carried out on rotary-shadowed preparations of scuPA and suPAR incubated together for 30 min. In images of a mixture of suPAR and scuPA, there were also monomeric and dimeric structures. The small difference in size of scuPA and suPAR did not allow us to unequivocally identify monomers as one or the other in these mixtures (Fig. 8C).
Optical biosensors were employed to further examine the data obtained by size exclusion, which suggested that some forms of suPAR bind scuPA less efficiently than others. The data obtained by size exclusion, dynamic light scattering, and EM show that suPAR, but not scuPA, dimerizes/aggregates. Our aim, therefore, was to compare the behavior of both molecules in solution and to see how the postulated dimerization/aggregation of suPAR affects its affinity toward scuPA. For this purpose, either suPAR or scuPA were immobilized to CM5 carboxymethydextran biosensor surfaces, and the corresponding ligand was added at various concentrations. A wide range of concentrations was used to examine concentrations that induce scuPA activity (Figs. 1 and 4A) as well as signal transduction. To avoid relying on kinetic data (which are more likely to be affected by immobilization chemistry), analysis of equilibrium binding was performed to determine ligand affinity.
Nonlinear regression of the single-site bimolecular binding
isotherms was performed for both the binding of scuPA to suPAR (Fig. 9, A and C)
and suPAR to scuPA (Fig. 9, B and D). For scuPA in solution, a Kd of 4.2 ± 0.6 nM
was obtained (R2 = 0.98) whereas a
Kd of 65 ± 8 nM
(R2 = 0.99) was calculated when suPAR was
in solution. As seen by the values of R2, the
fit for this model was quite good. Interestingly, for linear regression
of Scatchard-transformed data, this was not the case, specifically when
suPAR was in solution. As can be seen from the plots, the data for
suPAR in solution is curvilinear, suggesting a more complex binding
interaction, whereas a linear plot is obtained with scuPA in solution.
One known cause of such deviation from linearity is dimerization of
ligand in solution, which lowers the effective concentration of monomer
(43); this not only results in a curvilinear Scatchard plot, but also
in an apparent increase in Kd. These data give
further support to the formation of suPAR dimmers in solution.
Furthermore, these data demonstrate directly that this dimerization
decreases the effective affinity of scuPA for its receptor.
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DISCUSSION |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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Our data show that scuPA and suPAR are able to establish a novel interaction that appears at concentrations that are higher than the accepted Kd and the predicted saturation. These interactions result in increased activity of the complex with half-maximal activity at 30 nM. These conclusions are based on several observations, which follow.
The increase of suPAR concentrations to levels that exceed the theoretical saturation of a simple 1:1 complex induces a further stimulation of scuPA/suPAR activity. The stimulatory effect of suPAR cannot be attributed to contaminants or artifacts for several reasons. 1) Because suPAR by itself has no plasminogen activation activity, contamination by PA cannot be a factor. 2) scuPA was not cleaved by suPAR to tcuPA, and tcuPA activity was stimulated by suPAR. These observations exclude the involvement of contaminants with protease activity that could induce scuPA cleavage to tcuPA. 3) The abolition of the suPAR-induced activation by neutralizing anti-uPAR antibodies or by ATF.
Three approaches were used to verify previously unknown interactions between suPAR/suPAR molecules and between scuPA and suPAR. The three approaches used show that suPAR is in equilibrium between different forms (oligomers/dimers/monomers). It is clear that the addition of equimolar concentrations of scuPA leads to the dissociation of all forms of suPAR dimerization/oligomerizaion. The observation that the monomeric scuPA-suPAR complex elutes with a longer RT as compared with any form of suPAR monomer alone (RT = 8.4 versus 8.1 min) indicates that there is no detectable suPAR monomer. This conclusion is supported by the observation that scuPA runs at an RT of 10.9 min.
Dynamic light scattering measures the translational diffusion coefficient of the protein by monitoring the decay of an autocorrelation function with time. The hydrodynamic radius (Rh) of the protein can then be calculated using the Stokes-Einstein equation. The molecular mass of the protein is estimated by fitting the measured Rh to a standard curve derived from 25 globular proteins. Proteins with major axial components, or that are spheroidal or partially unfolded, will show an apparent molecular mass that are higher than their sequence molecular mass. This is the case for scuPA, where dynamic light scattering gives a molecular mass of 73 kDa, which is much higher than the molecular mass (46.5 kDa) determined from the sequence. This excludes the possibility for a scuPA dimer and indicates that scuPA might have an elongated shape.
This conclusion is consistent with the data published by Mangle et al. (44) using small angle neutron scattering. It is also consistent with the modular structure of scuPA, which consists of an EGF domain, a kringle domain, and a serine protease domain. Futhermore, this conclusion is supported by the data obtained using SEC or EM techniques.
At high concentrations, suPAR exists mostly (98%) as a species with a molecular mass of 97.7 kDa, consistent with the formation of dimers. It is difficult to exclude the possibility of the formation of suPAR trimers, but it can be concluded that under the experimental conditions, suPAR does not exist as a monomer. The ability of suPAR to form dimers was confirmed using SEC and EM.
In contrast to the dimerization of suPAR alone, the scuPA-suPAR complex at equimolar concentrations is clearly a single species (Fig. 7C) with an Rh of 4.1 nm and an apparent molecular mass of 91 kDa. It is interesting to note that the scuPA-suPAR complex has a size (4.1 nm) similar to that of the suPAR dimer (4.2 nm). This finding is in agreement with the SEC data, which shows that both suPAR and the scuPA-suPAR complex elute with similar retention times in a size exclusion column. The combined data of SEC and dynamic light scattering suggest a more compact and smaller hydrodynamic radius in the scuPA-suPAR complex.
Biophysical techniques (light scattering and SEC) and electron
microscopy are complementary, as demonstrated, for example, by
recent studies of the formation of oligomers and conformational changes
of the integrin 
IIb
3 (45). The former
methods give quantitative results under aqueous conditions but can be
somewhat difficult to interpret. On the other hand, electron microscopy provides a physical picture that may be used to interpret the biophysical measurements, although any quantification of the images must be done very cautiously because of artifacts arising from the
conditions and methods of preparation.
The isolation of suPAR dimers and oligomers by SEC and EM indicates that the suPAR/suPAR interaction is a stable one. On the other hand, the clean and complete disruption of the suPAR dimer and oligomer by scuPA shown here suggests that the interaction of self-association of suPAR is relatively weak, compared with the interaction between suPAR and scuPA, which has a dissociation constant in the nanomolar range.
The SEC shows that suPAR in oligomers binds scuPA with lower affinity as compared with when it is present in dimers. The optical biosensor experiments are consistent with this observation, demonstrating a lower apparent affinity of suPAR to scuPA when suPAR is in solution at relatively high concentrations and not only monomers, but also higher order oligomers, are present. In addition, the deviation from linearity of Scatchard plots for the condition when suPAR is free in solution contrasted to when scuPA is free in solution is expected from this interaction model.
SEC, dynamic light scattering, and electron microscopy techniques indicate for the first time that suPAR can spontaneously form dimers and oligomers. The three techniques provide evidence that the addition of scuPA leads to the dissociation of these dimers and oligomers. Thus, the binding of scuPA to suPAR leads to changes in the properties of suPAR by interfering with the dimer/oligomers structure. This may lead to the exposure of binding epitopes in suPAR that will permit its interactions with other ligands, including vitronectin (7-9) or thrombospondin (8). By permitting novel interactions of uPAR, such a process may be involved in signal transduction.
Based on the data presented, we postulate that suPAR is present in equilibrium between monomeric, dimeric, and oligomeric forms. The binding of uPA to suPAR at equimolar concentrations induces the dissociation of the suPAR dimers. We may therefore find high moleculat weight entities when suPAR is in excess, although we have not been able to demonstrate the existence of such entities to date; this may be because of the fragile nature of such heterogenic complexes. The fragility of a suPAR/suPAR/scuPA complex may stem from the fact that although suPAR can establish interactions with scuPA or with another molecule of suPAR, it prefers the interaction with scuPA. Thus, when scuPA is present in equimolar concentration, all of the scuPA is bound to suPAR, whereas if suPAR is in excess, the free molecules of suPAR can still interact with suPAR that is bound to scuPA. The observation that in the presence of a molar excess of suPAR, the activity of uPA is greater than its activity at equimolar concentration, along with the capacity of suPAR to form dimers and oligomers, strongly supports the possibility that such an entity does exist. More efforts will be invested to demonstrate the existence of these entities.
Furthermore, the exact mechanism by which these complexes are assembled
and disassembled and whether these complexes have any biological
relevance need to be elucidated in the future. Indeed, the mechanism by
which uPAR binds uPA is not fully understood. Our data (32) and those
of others (33) indicate that uPAR contains several binding epitopes
that participate in the binding of uPA. These binding epitopes are
distributed on the three homologous domains that form the uPA receptor.
The existence of these different epitopes on uPAR may help to provide
an explanation for the formation of these novel complexes.
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ACKNOWLEDGEMENTS |
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We thank Dr. Gabriella Canziani and Dr. Irwin Chaiken of the University of Pennsylvania Biosensor Core for technical assistance with the surface plasmon resonance experiments.
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FOOTNOTES |
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* This work was supported in part by Grants HL60169 (to A. A. H.), HL58107 (to A. A. H.), and HL30954 (to J. W. W.) from the National Institutes of Health.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.
Present address: Attenuon, LLC, 10130 Sorento Valley Rd.,
Suite B, San Diego, CA 92121.
§§ To whom correspondence should be addressed: Dept. of Pathology and Laboratory Medicine, 513A Stellar-Chance, 422 Curie Blvd., Philadelphia, PA 19104. Fax: 215-573-2012; E-mail: abd@md2.huji.ac.il.
Published, JBC Papers in Press, May 4, 2000, DOI 10.1074/jbc.M002024200
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ABBREVIATIONS |
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The abbreviations used are: uPA, urokinase-type plasminogen activator; uPAR, uPA receptor; scuPA, single-chain uPA; tcuPA, two-chain uPA; ATF, amino-terminal fragment of urokinase; LMW, low molecular weight; suPAR, recombinant soluble uPA; MES, 4-morpholineethanesulfonic acid; HPLC, high pressure liquid chromatography; EM, electron microscopy; PAGE, polyacrylamide gel electrophoresis; PBS, phosphate-buffered saline; SEC, size exclusion chromatography.
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REFERENCES |
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TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
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, shows the activity of 25 nM LMWscuPA
(A) or LMWtcuPA (B) in the presence of increasing
concentrations of suPAR.
