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Blood, Vol. 91 No. 6 (March 15), 1998:
pp. 1987-1998
Tissue Factor Regulates Plasminogen Binding and Activation
By
Zhiqiang Fan,
Peter J. Larson,
John Bognacki,
P.N. Raghunath,
John
E. Tomaszewski,
Alice Kuo,
Gabriela Canziani,
Irwin Chaiken,
Douglas B. Cines, and
Abd Al-Roof Higazi
From the Departments of Pathology and Laboratory Medicine and
Medicine and Pediatrics, University of Pennsylvania, Philadelphia, PA;
the Division of Hematology, Children's Hospital of Philadelphia,
Philadelphia, PA; and American Diagnostica, Inc, Greenwich, CT.
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ABSTRACT |
Tissue factor (TF) has been implicated in several important biologic
processes, including fibrin formation, atherogenesis, angiogenesis, and
tumor cell migration. In that plasminogen activators have been
implicated in the same processes, the potential for interactions
between TF and the plasminogen activator system was examined.
Plasminogen was found to bind directly to the extracellular domain of
TF apoprotein (amino acids 1-219) as determined by optical biosensor
interaction analysis. A fragment of plasminogen containing kringles 1 through 3 also bound to TF apoprotein, whereas isolated kringle 4 and
miniplasminogen did not. Expression of TF on the surface of a stably
transfected Chinese hamster ovary (CHO) cell line stimulated
plasminogen binding to the cells by 70% more than to control cells.
Plasminogen bound to a site on the TF apoprotein that appears to be
distinct from the binding site for factors VII and VIIa as judged by a
combination of biosensor and cell assays. TF enhanced two-chain
urokinase (tcuPA) activation of Glu-plasminogen, but not of
miniplasminogen, in a dose-dependent, saturable manner (half maximal
stimulation at 59 pmol/L). TF apoprotein induced an effect similar to
that of relipidated TF, but a relatively higher concentration of the
apoprotein was required (half maximal stimulation at 3.8 nmol/L). The
stimulatory effect of TF on plasminogen activation was confirmed when
plasmin formation was examined directly on sodium dodecyl
sulfate-polyacrylamide gel electrophoresis. In accord with this, TF
inhibited fibrinolysis by approximately 74% at a concentration of 14 nmol/L and almost totally inhibited the binding of equimolar
concentrations of plasminogen to human umbilical vein endothelial cells
and human trophoblasts. Further, CHO cells expressing TF inhibited
uPA-mediated fibrinolysis relative to a wild-type control. TF
apoprotein and plasminogen were found to colocalize in atherosclerotic
plaque. These data suggest that plasminogen localization and activation
may be modulated at extravascular sites through a high-affinity
interaction between kringles 1 through 3 of plasminogen and the
extracellular domain of TF.
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INTRODUCTION |
TISSUE FACTOR (TF) plays a critical role
in the initiation of coagulation.1,2 TF apoprotein is a 263 amino acid integral membrane glycoprotein. TF is not expressed by cells
that interface with the circulation under physiologic
conditions,3 but its expression can be induced on monocytes
and on subsets of endothelial cells4 such as those adjacent
to atherosclerotic plaques5 as part of the response to
inflammation or injury.6,7 TF functions in the form of a
complex with membrane phospholipids to promote the activation of
factors VII, X, and IX. The presence of both components are required
under physiologic conditions for optimal procoagulant
activity.8-11
Clot formation and lysis must be tightly coordinated to secure
hemostasis while maintaining vascular patency. Fibrinolysis is
dependent on the formation of plasmin through the cleavage of
plasminogen by tissue-type plasminogen activator (tPA) and urokinase-type plasminogen activator (uPA) in plasma, on cell surfaces,
and in some extracellular compartments. Plasmin regulates the
activities of coagulation factors such as FV,12 whereas FVa
and FXa have recently been reported to regulate plasminogen activation.13 These observations suggest that potentially
important links may exist between the initiation of coagulation by TF
and the dissolution of fibrin clots by plasmin.
Another potential link comes from recent studies in which TF has been
implicated in angiogenesis,14 vasculogenesis,15 tumor cell invasion,16,17 smooth muscle cell
migration,18 and atherogenesis,5,19-21
processes in which plasminogen activation has been implicated as
well.22-25 The mechanism by which TF participates in these
processes is unknown. The observation that exogenous FVIIa or FXa can
bind to TF in atherosclerotic plaques,5 for example,
indicates that the epitopes for these ligands are not totally occupied
and suggests that binding sites for other ligands may be available as
well. Furthermore, the finding that FVIIa bound to TF in these plaques
expresses enzymatic activity suggests that factors other than tissue
factor protease inhibitor (TFPI) may participate in its regulation at
such extravascular sites. Taken together, these interactions lead us to
ask whether plasminogen binds to TF and whether TF may modulate the
expression of plasminogen activator activity.
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MATERIALS AND METHODS |
Materials.
Glu-plasminogen was purified from human plasma by affinity
chromatography on lysine Sepharose as described.26
Miniplasminogen, high molecular weight two-chain urokinase (tcuPA), and
the plasmin chromogenic substrate Spectrazyme PL
(H-D-norleucyl-hexahydrotyrosyl-lysine-p-nitroanilide diacetate salt)
were provided by American Diagnostica Inc (Greenwich, CT). Murine
monoclonal antibodies to human TF (no. 4509) and plasminogen (no. 3644)
(antiplasminogen) were purchased from American Diagnostica Inc.
Single-chain urokinase (scuPA) was the kind gift of Dr J. Henkin
(Abbott Laboratories, Abbott Park, IL). TF apoproteins in 4 mmol/L
CHAPS were from American Diagnostica as well as generously provided by
Dr L.V.M. Rao (Tyler, TX). Relipidated TF apoproteins were from
American Diagnostica, Baxter Diagnostics, Inc (Deerfield, IL), and Dr
Rao. Factors VII and VIIa were from American Diagnostica and were
purified from plasma (see below). Recombinant soluble extracellular
domain of TF (amino acids 1-219) was the generous gift of Wolfram Ruf
(Scripps Research Institute, La Jolla, CA).
Cells.
Cultures of human umbilical vein endothelial cells (HUVECs) were
prepared using published methods27 in Medium 199 supplemented with 10% heat-inactivated fetal calf serum (FCS; Life
Technologies, Bethesda, MD), penicillin-streptomycin, 2 mmol/L
glutamine, 12 U/mL heparin sodium (SoloPak, Elk Grove Village,
IL), Hydrocortisone (1 µg/mL; Clonetics Co, San Diego,
CA), bovine brain extract (12 µg/mL; Clonetics Co), and
human recombinant epidermal growth factor (0.01 µg/mL; Clonetics Co).
The cells were passaged using trypsin-EDTA (GIBCO), frozen in liquid
nitrogen, thawed, and passaged two additional times before use.
Cultures of first trimester human trophoblasts were prepared and
characterized as described.28,29
Development of a Chinese hamster ovary (CHO) cell line expressing
TF.
TF cDNA (clone pHTF8) was obtained from American Type Culture
Collection (Rockville, MD). TF cDNA, corresponding to nucleotides 21 to
1050 of the published sequence30 was cloned into the
Sal I site of the vector pUC18 (Life Technologies, Bethesda,
MD). The vector containing this insert was then digested with
Sau3AI to yield an approximately 1-kb fragment beginning at
nucleotides 54 of TF at its amino terminus and terminating within the
multiple cloning site of the vector at its carboxyterminus. The
fragment was isolated using low-melt gel electrophoresis and a Qiaex II gel purification kit (Qiagen, Chatsworth, PA). The DNA fragment was
ligated into BamHI site of vector pcDNA3 (Invitrogen, Carlsbad, CA). The size and orientation of the insert was confirmed using Sma I and the nucleotide sequence was verified by
dideoxynucleotide chain-termination DNA sequence analysis (University
of Pennsylvania DNA Sequencing Facility, Philadelphia,
PA).
The CHO cells (American Type Culture Collection) were cultured at
37°C in 5% CO2 in Iscove's modified Dulbecco's
medium containing 2 mmol/L L-glutamine, 25 mmol/L HEPES, 10% fetal
bovine serum, 100 U/mL penicillin, 100 µg/mL streptomycin, 0.05 mmol/L hypoxanthine, and 8 nmol/L thymidine (all from Life
Technologies). Before transfection, the CHO cells were seeded onto
15-cm2 petri dishes for at least 24 hours until they
reached approximately 70% confluence. The cells were incubated with
lipofectamine (Life Technologies) and the pcDNA3-TF plasmid that
contains a neomycin resistance gene. The remaining steps were as per
the manufacturer's instruction (Life Technologies). Colonies resistant
to G418 (600 µg/mL; Life Technologies) were selected for analysis.
Individual colonies were trypsinized, transferred to 96-well microtiter
wells (Falcon 3072; Falcon, Franklin Lakes, NJ), and grown to
confluence. Colonies were chosen based on their procoagulant activity
and expression of TF (see below).
The colonies were then characterized for their expression of TF antigen
and FVII-dependent coagulant activity. To measure TF antigen
expression, transfected and control cells were cultured in 96-well
plates until they reached confluency. The cells were then washed twice
with phosphate-buffered saline (PBS) containing 1% bovine serum
albumin (BSA) and incubated with series of buffers each for 1 hour at
room temperature with five washes in PBS/BSA between each incubation.
The cells were incubated sequentially with PBS/2% BSA, 1 µg/mL
anti-TF antibody in PBS/1% BSA, biotinylated goat antimouse IgG (H+L;
Southern Biotechnology Associates, Inc, Birmingham, AL) diluted
1/10,000 in PBS/1% BSA, and streptavidin-horseradish peroxidase (Southern Biotechnology Associates, Inc)
diluted 1/1,000 in PBS/1% BSA according to the manufacturer's
instruction. o-phenylenediamine free base (Sigma Chemical Co, St Louis,
MO) was diluted in phosphate-citrate buffer, pH 5.0, and made 0.03%
H2O2 immediately before its addition to the
cells, and the optical density at 490 nm was measured (Molecular Devices, Palo Alto, CA).
To measure procoagulant activity, CHO cells transfected with TF and the
wild-type parental cells were cultured in 24-well dishes until they
reached confluence. The cells were then washed twice in PBS (without
calcium) and 0.1 mL of citrated normal plasma (0.32% final
concentration) and 0.1 mL PBS (without calcium) was added. Lastly, 0.1 mL of 25 mmol/L CaCl2 was added and the time to form a
visible clot was recorded.
Biosensor analysis of plasminogen binding to TF.
Kinetic analysis of the interaction of plasminogen with TF was
performed using BIA2000 and IAsys optical biosensors (see
Myszka31 and the references therein). Soluble and
apoprotein forms of TF were separately coupled to either CM5-research
grade sensor chip flow cells (BIACORE) or carboxymethyl-dextran
cuvettes (IAsys), using standard amine coupling procedures described by
the manufacturers.32 The coating conditions for TF
immobilization were optimized by observing the electrostatic attraction
to the negatively charged CM-dextran surface at different pHs (2.0 to
5.0 in 10 mmol/L NaAc buffer). TF apoprotein at a concentration of 15 µg/mL, pH 2.3, or soluble TF at a concentration of 15 µg/mL, pH
4.0, were used to coat the sensors' surfaces. The net response chosen
for binding detection was 400 arc seconds and 4,000 RU for the IAsys
and the BIACORE, respectively. Coupling on the BIACORE flow cell was
performed with a flow of 5 µL/min. Immobilized ligand was
equilibrated with PBS, pH 7.4, 0.05% Tween-20 in all experiments,
except when Ca2+ was required in the reaction cell. TBS, pH
7.4, 0.05% Tween-20 was used in the presence of Ca2+
cations. Regeneration of the surface after each binding interaction was
achieved using 10 mmol/L EDTA, pH 9.4, or by competing with the
immobilized ligand for plasminogen binding using a lysine analogue
6-amino hexanoic acid (6-AHA). All experiments were performed at
25°C.
Kinetic measurement and analysis.
Binding of the soluble analyte to the surface-immobilized ligand, in
this case plasminogen and TF, respectively, was detected by a change in
the refractive index as a function of time.33 The detector
response is proportional to the surface concentration of bound
ligand.34 The kinetics of binding were initially evaluated assuming the ligand in solution, or analyte, A (Glu-plasminogen) interacts with the immobilized ligand B, in this case TF, to form complex AB. The net rate of complex formation depends on the free concentration of A and B and on the stability of the formed complex.
Kinetics data were evaluated using relationships described
previously.33,35 For a bimolecular interaction,
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(1)
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the
association and dissociation processes are described respectively by
the relationships
![dR/dt = k<SUB>on</SUB>[A]R<SUB>max</SUB> − (k<SUB>on</SUB>[A] + k<SUB>off</SUB>)R,](/content/vol91/issue6/fulltext/1987/img002.gif)
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(2)
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and
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(3)
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Here,
kon and koff are on and off
rate constants, respectively; [A] is the concentration of analyte
injected onto a flow cell (or into the sensor cuvette) containing the
second interactor B immobilized on a sensor surface; R is the relative
response of the optical biosensor at time t and is proportional to the amount of complex formed; Rmax is the maximum response; and
Ro is the response at the start of the dissociation phase.
When association follows equation 2, a plot of dR/dt versus R should be
linear and yield a slope of  ks = kon[A] + koff, so that a replot of ks versus [A] will yield a slope of kon. For
a single exponential process (equation 3), a plot of
ln(Ro/R) versus time from the dissociation part of the
sensorgram should be a straight line with a slope of koff.
Immunohistochemical colocalization of TF and plasminogen in human
atherosclerotic coronary arteries.
To study the colocalization of TF and plasminogen in vascular samples,
a sequential double immunoenzymatic staining procedure for simultaneous
localization of both antigens in the same tissue section was used.
Vascular samples were obtained from human hearts removed at the time of
transplantation. Replicate 2- to 3-mm sections were fixed by immersing
each sample in 10% neutral buffered formalin overnight at 21°C.
All samples were then embedded in paraffin and serial 5-mm sections
were cut on ProbeOn Plus slides and used for immunocytochemistry. For
immunostaining, a modification of the avidin-biotin peroxidase method
was performed using capillary action technology and the Microprobe
System (Fisher Scientific, Pittsburgh, PA). Briefly, sections were
deparaffinized at 55°C for 30 minutes, placed in xylene, and
hydrated in 100% and then 95% ethanol, and endogenous peroxidase
activity was quenched with a 2.2% (vol/vol)
H2O2/methanol solution added for 5 minutes. All subsequent incubations were performed at room temperature for the
specified time. The slides were then washed with 1× automation buffer (Biomeda Corp, Foster City, CA) and 5% normal horse serum was
added for 20 minutes to block nonspecific binding. Primary and
secondary antibodies were diluted in 1× automation buffer containing 5% normal horse serum. Antibodies to TF and plasminogen were used at 5 µg/mL final concentration. Isotype-matched nonimmune mouse Ig was used as an irrelevant control antibody. The slides were
then incubated with primary antibody for 60 minutes, washed in 1×
automation buffer for 10 minutes, incubated with biotinylated antimouse
antiserum (Vector Laboratories, Burlingame, CA) at a 1:300 dilution for
30 minutes, and washed again with 1× automation buffer. An
alkaline phosphatase-streptavidin-biotin complex (Dako, Capiteria,
CA) was used as a reporter system, and the reaction was
visualized with the alkaline-phosphatase substrate (Alkaline phosphatase substrate kit III-SK 5300; Vector Laboratories) that produces a blue reaction product.
The sections were washed with 1× automation buffer, and once
again blocking was performed with 5% normal horse serum for 20 minutes. Antiplasminogen antibody was then applied for 60 minutes. Sections were then incubated with the antimouse biotinylated secondary antibody for 30 minutes. The slides were then incubated for 30 minutes
with the streptavidin-biotin peroxidase system (Dako) at a 1:50
dilution and developed by adding 0.05% (vol/vol)
3,3 -diaminobenzidine solution (Sigma Chemical Co) and 0.03%
H2O2 (vol/vol) for 5 minutes. Stained sections
were dehydrated, a coverslip was added, and the extent of staining was
evaluated. Doubly stained cells showed mixtures of brown (plasminogen)
and blue (TF) tones.
Expression of human FVII in vitro.
The cDNA for normal FVII (gift of Dr Darrell Stafford, University of
North Carolina, Chapel Hill, NC) was inserted into the multiple cloning site of the expression vector pCMV536 as
described.37 Correct insertion was confirmed by sequence
analysis. Human embryonic kidney cells (293; American Type Culture
Collection) were cotransfected with the pCMV5 and pSV2neo plasmids
using calcium phosphate,38 and the transfected cells were
selected with 500 µg/mL G418 (Life Technologies, Grand Island, NY).
Colonies producing FVII as determined by enzyme-linked immunosorbent
assay were grown to confluence.
Purification of recombinant wild-type FVII.
FVII-producing cell lines were expanded in Dulbecco's modified
Eagle's medium (DMEM)/F12 medium contain 10% FCS,
penicillin/streptomycin, L-glutamine, and 6 µg/mL vitamin K. The
cells were cultured in roller bottles in serum-free medium containing
10 µg/mL of a supplement containing insulin, transferrin, and
selenium (Boehringer Mannheim, Indianapolis, IN) for up to 14 days, and
the culture supernatants were harvested at approximately 24-hour
intervals. Clones expressed FVII at a concentration of approximately 1 mg/L/24 hr. Culture supernatants were filtered through cellulose
acetate (Schleicher and Schuel, Keene, NH). Benzamidine (10 mmol/L) and
protease inhibitors (Complete protease inhibitor cocktail; Boehringer
Mannheim) were added (0.5 µL 25× stock/mL conditioned medium),
and the samples were stored at 20°C. Thawed culture medium
was diluted with 1.5 vol of 20 mmol/L Tris, pH 7.2/10 mmol/L
benzamidine/5 mmol/L EDTA, mixed with 10 mL of Q-Sepharose beads
(Pharmacia Biotech, Uppsala, Sweden) equilibrated in 20 mmol/L Tris, pH
7.2/60 mmol/L NaCl/10 mmol/L benzamidine/5 mmol/L EDTA and stirred at
4°C for 20 minutes. Q-Sepharose beads were washed in a scintered
glass funnel with 20 mmol/L Tris, pH 7.2/10 mmol/L benazmidine before
use. Bound protein was eluted with 20 mmol/L Tris (pH 7.2/750 mmol/L
NaCl/10 mmol/L benzamidine). Pooled fractions containing protein were mixed with 4 vol of 20 mmol/L Tris, pH 7.4/10 mmol/L benzamidine and
made 10 mmol/L in CaCl2. The pooled fractions were applied to a calcium-dependent anti-FVII monoclonal antibody-Sepharose column
(gift of Dr T. Jorgensen, NovoNordisk, Copenhagen, Denmark) equilibrated with 20 mmol/L Tris, pH 7.2/100 mmol/L NaCl/10 mmol/L CaCl2/50 mmol/L benzamidine, and a buffer containing 20 mmol/L Tris (pH 7.2/1 mol/L NaCl/10 mmol/L CaCl2/50 mmol/L
benzamidine) was used to elute protein nonspecifically bound to the
column. The column was then re-equilibrated, and recombinant FVII was eluted with 20 mmol/L Tris, pH 7.2/100 mmol/L NaCl/20 mmol/L EDTA/50 mmol/L benzamidine. The pooled material was concentrated using Centriprep-30 concentrators (Amicon, Beverly, MA) and quantified using
Coomasie blue (Pierce, Rockford, IL). Samples were assayed for purity
using an 8% to 25% polyacrylamide gel gradient and visualized by
silver staining (Bio-Rad, Hercules, CA). Analysis of
 -carboxyglutamic acid (Gla) content was performed as
described.39 Recombinant FVII had a Gla content that was
similar to plasma-derived FVII (Enzyme Research Laboratories, South
Bend, IN).
Competition between FVII and plasminogen for TF binding and
activity.
The effect of plasminogen on FVII binding to TF was evaluated by
BIAcore biosensor assay. Increasing concentrations of plasminogen were
injected into the flow cell and association allowed to reach steady
state in each case. After unbound plasminogen was washed out of the
flow cell by buffer alone, FVII was injected at a concentration of 149 nmol/L and its association and dissociation phases were recorded. The
extents of association of FVII after varying plasminogen binding were
compared.
Next, the potential for FVII and plasminogen to compete for activity
was studied in two ways. First, relipidated TF (10 nmol/L) was
incubated with 125I-plasminogen (100 nmol/L) in the absence
or in the presence of FVII or FVIIa (100 nmol/L each) and the specific
binding to HUVECs was measured as described above. Second, TF (10 nmol/L) was added to 0.1 mL of recalcified, citrated pooled normal
plasma in the absence or presence of plasminogen (100 nmol/L) and the
time to form a visible clot was determined.
Measurement of plasminogen activator activity.
Two methods were used. First, plasminogen activator activity was
measured as previously described.40 Briefly, Glu- or
mini-plasminogen (5 nmol/L), tcuPA (5 nmol/L), and Spectrazyme PL (100 to 167 µmol/L) were incubated with varying concentrations of TF in
0.1 mol/L Tris-HCl, pH 8.0, or PBS, pH 7.4 (60 µL final volume), in
96-well Falcon Multiwell tissue culture dishes (Becton Dickinson,
Lincoln Park, NJ), and the optical density at 405 nmol/L was measured at various times. Second, conversion of plasminogen to plasmin was
analyzed using sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE). In these experiments, Glu-plasminogen (3 µmol/L) was incubated with tcuPA (400 nmol/L) in the absence or
presence of various concentrations of TF apoprotein (0 to 450 nmol/L;
American Diagnostica) in 57 mmol/L Tris-HCl, pH 8.0 (final volume, 70 µL), at 37°C for 20 minutes. To stop the reaction, 5 µL of
4× SDS-PAGE sample buffer containing 10% 2-mercaptoethanol was
added to 15 µL of the reaction mix. The samples were boiled for 5 minutes, and the proteins were separated using a 10% SDS 12%
polyacrylamide gel that was then stained with Coomasie blue.
Contribution of TF apoprotein and phospholipids to plasminogen
activation.
Three sets of experiments were performed to explore the contribution of
the phospholipid component of TF to plasminogen activation. First, the
effects of the TF apoprotein and relipidated TF were compared. Second,
purified phospholipids in the same ratio as incorporated into
relipidated TF were studied. Third, human serum albumin (Sigma) was
relipidated in the same manner as TF. To do this, a mixture containing
equimolar ratios of phosphatidyl serine, phosphatidyl choline, and
phosphatidyl ethanolamine (Avanti Polar Lipids Inc, Alabaster, AL)
dissolved in 100% ethanol was dried under nitrogen. The phospholipids
were resuspended in 10 mmol/L Tris HCl, pH 7.5, and added to an
equimolar concentration of albumin overnight at 22°C in the
presence of deoxycholate (0.05% final concentration). Excess detergent
was removed by extensive dialysis against PBS.
Effect of TF on plasminogen binding to cells.
Two sets of experiments were performed. First, Glu-plasminogen was
radiolabeled with Na 125I (Dupont-NEN, Boston, MA) using
Iodo-Beads (Pierce, Rockford, IL). The cells were washed twice with 1%
albumin in Dulbecco's PBS supplemented with 0.9 mmol/L
CaCl2 and 0.5 mmol/L MgCl2 (GIBCO). 125I-labeled plasminogen (100 nmol/L) was added to the
cells alone or in the presence of various concentrations of TF
apoprotein for 1 hour at 15°C in the case of trophoblasts or at
4°C when HUVECs were studied. The cells were washed four times in
the same buffer to remove unbound plasminogen. The bound plasminogen
was then eluted by exposing the cells to glycine buffer, pH 3.0, for 7 to 10 minutes and the radioactivity was measured. Nonspecific binding
was determined by performing the experiment in the presence of 15 mmol/L 6-AHA (Sigma). Specific binding was calculated from the
difference between total and specific binding. In a second set of
experiments, the binding of 125I-Glu-plasminogen to CHO
cells was measured. CHO cells expressing TF and parental wild-type
cells (4 × 104) were incubated with 50 nmol/L
125I-plasminogen in the presence or absence of 100 mmol/L
6-AHA for 2 hours at 37°C. The cells were washed three times in
PBS, the cell-associated radioactivity was measured, and the specific
binding was calculated.
Effect of TF on uPA-mediated fibrinolysis.
Two sets of experiments were performed to assess the effect of TF on
uPA-mediated fibrinolysis. First, fibrinogen containing plasminogen
(Calbiochem, San Diego, CA) was reconstituted in PBS to a concentration
of 3 mg/mL. Thrombin (Sigma) at 0.025 NIH U/mL was added to 30 mL of
the fibrinogen mixture and the fibrin clot was decanted onto a plastic
lid. After 60 minutes at 22°C, 10 µL of PBS containing scuPA or
tcuPA (1 µmol/L final concentration) was added alone or in the
presence of TF (14 nmol/L). After 2 hours, the clot was washed four
times with PBS, trypan blue was added, and the samples were incubated
overnight at 22°C. The clot was then washed, photographed, and
scanned for total reflectance using a Hoefer GS 300 Scanning
Densitometer (Hoefer Scientific Instruments, San Francisco, CA). Each
image was assigned a value for total reflectance in arbitrary units.
The relative amount of reflectance was quantified by integrating the
peak reflectance and normalizing each value to the largest area
measured. Second, the effect of CHO cells expressing TF and wild-type
on the lysis of fibrin clots was compared. CHO cells were used both as
the source of uPA and TF in these experiments. The CHO cells (4 × 105) were detached with EDTA, washed in PBS, and added for
3 hours at 37°C to a fibrin clot formed in the presence of
plasminogen (500 µmol/L) or buffer. The wells were photographed and
the size of the zones of lysis was determined using a scanning
densitometer as described above.
| |
RESULTS |
The purpose of this study was to determine whether plasminogen binds to
TF and whether this binding modulates plasminogen activation.
Binding of plasminogen to immobilized TF.
To determine whether plasminogen interacts directly with TF, we
examined the capacity of plasminogen to bind to a soluble, immobilized
fragment of recombinant TF apoprotein comprising the extracellular
domain of the protein (amino acids 1-219) using optical biosensor
interaction analysis. As shown in Fig 1A,
plasminogen was found to bind to this fragment in a dose-dependent
manner. Linearization of both the association and dissociation phases according to equations 2 and 3, respectively (see the Materials and
Methods), showed that these processes did not fit strictly to a single
bimolecular process. In general, nonlinear dR/dt plots such as shown in
Fig 1B could reflect multiple binding sites with different affinities,
cooperativity or more complex models. To make approximations of binding
constants in the current experiments, the dR/dt plots were divided into
fast and slow components (components 1 and 2, respectively), in a
manner used previously (see, for example, Casanovas and
Springer41). Linear fits of components 1 and 2 to equation
2 yielded ks plots (Fig 1C) that led to kon values of 4 × 104 and 2 × 103 mol/L 1 s1,
respectively. Fitting the early dissociation data to equation 3 (Fig
1D) led to a calculated koff of 3.8 × 103 s1. Hence, from these linear
fits, apparent equilibrium Kd values were obtained of 100 nmol/L and 2 µmol/L for components 1 and 2, respectively.
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| Fig 1.
Kinetic interaction analysis of Glu-plasminogen binding
to TF as determined with a BIAcore optical biosensor. (A) Sensorgrams for interaction of denoted concentrations of Glu-plasminogen to sensor-immobilized TF extracellular domain. Association phase: 3-minute
injections of Glu-plasminogen at 30 µL/min. Dissociation phase:
4-minute injections of buffer alone at 100 µL/min. (B) dR/dt plots of
association data according to equation 1. Linear fits are shown to two
different components of the data, the early component (cpt. 1) and the
late component (cpt. 2). (C) Concentration dependence of ks values
determined from linear fits of cpts. 1 and 2 to equation 1. (D)
Dissociation phase data from 1 µmol/L plasminogen sensorgram plotted
according to equation 2.
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|
We next examined whether plasminogen had an effect on FVII and FVIIa
binding to TF apoprotein. FVIIa was found to bind to immobilized TF
extracellular domain by biosensor analysis (data not shown), consistent
with previously published results.42,43 We then tested
whether plasminogen prebinding to the immobilized TF affected
subsequent binding of FVII. As shown in Fig
2, preinjection of plasminogen at increasing concentrations did not
significantly reduce the extent of subsequent FVII binding, as judged
by the maximum response signal after FVII injection, corrected for the signal due to bound plasminogen. A similar finding was observed when
the order of injection was reversed, ie, FVII followed by plasminogen
(data not shown). Hence, plasminogen and FVII do not appear to block
each other's binding to TF completely and likely bind at independent
sites.
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| Fig 2.
FVII interaction with TF-plasminogen complex as
determined by BIA2000 analysis. Glu-plasminogen was injected, at
concentrations of 0.125 µmol/L (sensorgram 3), 0.25 µmol/L
(sensorgram 2), and 1.0 µmol/L (sensorgram 1), onto
sensor-immobilized soluble TF and the sensor surface then washed as in
Fig 1. Then, FVII was injected in buffer containing 4 mmol/L
CaCl2 for 3 minutes at 30 µL/min, followed by a wash in
buffer alone containing 4 mmol/L CaCl2. Sensorgram 4 shows
binding of FVII to immobilized TF after preinjection and wash with
buffers containing no Glu-plasminogen. Sensorgrams labeled 1 through 4 for FVII are continuations of the same-numbered sensorgrams for
Glu-plasminogen. (Inset) Bmax values for FVII bound after prebinding of
Glu-plasminogen at the different concentrations. Bmax values are the
difference between the maximum observed response for FVII association
and the response due to bound Glu-plasminogen just before FVII was
injected.
|
|
Role of the kringles in the binding of plasminogen to TF.
The plasminogen molecule contains 5 kringle domains, each of which
contains a lysine binding site. These kringles mediate the binding of
plasminogen to fibrin, to other proteins such as Lp(a), and to cells.
Binding of plasminogen to these sites is prevented by the lysine
analogue, 6-AHA.44-48 Therefore, we next examined the
effect of 6-AHA on the binding of plasminogen to TF. As shown in
Fig 3A, plasminogen bound directly to
full-length TF apoprotein immobilized on a biosensor surface, in this
case measured using the IAsys biosensor. Only a portion of bound
plasminogen was eluted when the sensor surface was washed with buffer
alone. The remaining bound plasminogen was completely washed off by
injecting buffer containing 20 mmol/L 6-AHA. In accord with this
finding, a plasminogen fragment containing kringles 1 through 3 bound
to immobilized TF apoprotein (Fig 3B), whereas isolated kringle 4 and
miniplasminogen (which lacks kringles 1 through 4) did not (data not
shown).
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| Fig 3.
Comparison of binding of Glu-plasminogen (A) and kringles
1 through 3 (B) to immobilized TF apoprotein as observed with an IAsys
optical biosensor. Association and dissociation (wash) buffers were as
for the binding of Glu-plasminogen with immobilized soluble TF (Fig 1,
first part of Fig 2). Both proteins were injected into the sensor
cuvette at 500 nmol/L and the association phase was observed.
Dissociation was observed in dissociation buffer after 5 quick (5 seconds) rinses of the sensor cuvette with dissociation buffer. In the
case of the plasminogen experiment, 20 mmol/L 6-AHA was added to the
rinsed cuvette after buffer-induced dissociation.
|
|
Relationship of TF to cellular binding sites for plasminogen.
Experiments were performed to determine whether the binding of TF to
plasminogen affects its binding to cells, a process that is known to be
mediated through its kringles. To do this, we measured the binding of
125I-plasminogen to HUVECs in the presence and absence of
TF. TF inhibits the binding of plasminogen to HUVECs in a
dose-dependent manner (Fig 4A). Specific
binding of plasminogen was almost totally inhibited by equimolar
concentrations of TF, an effect that was achieved only at much higher
concentrations of 6-AHA (15 mmol/L). The same results were obtained
using TF from three different sources and using another type of cell,
ie, cultured human trophoblasts (Fig 4B). As shown above (Fig 2), the
binding of FVII to TF apoprotein is unaffected by the binding of
plasminogen. Consistent with this result, preincubation of TF (100 nmol/L) with either FVII or FVIIa (100 nmol/L) had no effect on its
capacity to inhibit the binding of plasminogen (100 nmol/L) to cells or
to alter FVII-dependent procoagulant activity (not shown). Thus, these
data are consistent with the experiments cited above using soluble
proteins that suggest that plasminogen binds to TF through its kringles
at a site on the apoprotein distinct from the binding site for FVII and
FVIIa.
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| Fig 4.
TF inhibits the binding of Glu-plasminogen to cells.
125I-Glu-plasminogen (100 nmol/L) was added to HUVECs (A)
or trophoblasts (B) in the presence of the indicated concentrations of
TF or 15 mmol/L 6-AHA and the cell-associated radioactivity was
measured. The results are expressed relative to the binding of
radiolabeled plasminogen in the absence of TF. The mean ± SD of three
separate experiments performed in triplicate is shown.
|
|
Effect of TF on plasminogen activator activity.
Plasminogen undergoes a marked conformational change as a result of
binding of certain ligands to its kringles.49 This change in conformation alters the susceptibility of the protein to activation by plasminogen activators.50-52 In view of this, we next
asked whether the binding between TF and the kringles of plasminogen affected plasminogen activation by uPA. To examine this possibility, we
incubated tcuPA (5 nmol/L), Glu-plasminogen (5 nmol/L), and a plasmin
chromogenic substrate (100 µmol/L) in the presence and absence of 1.1 nmol/L TF. The addition of TF stimulated the formation of plasmin
(Fig 5). Plasmin formation was stimulated
by TF in a dose-dependent and saturable manner
(Fig 6). Half-maximal stimulation was
observed at a TF concentration of 59 pmol/L. Virtually identical results were obtained using each of three sources of TF (data not
shown).
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| Fig 5.
Effect of TF on uPA-mediated activation of
Glu-plasminogen. Glu-plasminogen (5 nmol/L) was incubated with tcuPA (5 nmol/L) and the plasmin chromogenic substrate (100 µmol/L) in the
absence or in the presence of TF (1.1 nmol/L). The mean ± SD of three separate experiments performed in triplicate is shown.
|
|
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| Fig 6.
Concentration-dependent effect of TF on uPA-mediated
activation of Glu-plasminogen. Glu-plasminogen (5 nmol/L) was incubated with tcuPA (5 nmol/L), the plasmin chromogenic substrate (100 µmol/L), and the indicated concentrations of relipidated TF for 15 minutes under the conditions described in Fig 4 and the absorbance at
405 nm was measured. The data are expressed relative to the absorbance
measured in the absence of TF. The mean ± SD of three separate
experiments performed in triplicate is shown.
|
|
We next examined the contribution of TF apoprotein and its phospholipid
component to uPA-mediated plasminogen activation. The results shown in
Fig 7 indicate that TF apoprotein
stimulated uPA-mediated plasmin generation. Half maximal stimulation
was achieved at a concentration of 3.8 nmol/L. Thus, a higher
concentration of the apoprotein was needed to stimulate plasmin
formation to the same extent as relipidated protein. The results shown
in Fig 7 also indicate that high concentrations of TF apoprotein
attenuated the stimulatory effect on plasmin formation. The fact that
higher concentrations of the apoprotein were required for plasminogen activation compared with relipidated TF suggest either that the phospholipids themselves express activity or that the phospholipids facilitate the activity of the apoprotein. To distinguish between these
two possibilities, we measured the plasminogen activator activity of a
mixture of purified phospholipids present in the same ratio as
relipidated TF and the activity of human serum albumin reconstituted
with phospholipids in the same manner as TF. In contrast to the
stimulatory effect of the TF apoprotein, neither the isolated
phospholipid components of TF nor relipidated human serum albumin had
any effect on plasminogen activation.
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| Fig 7.
Concentration-dependent effect of TF on uPA-mediated
activation of Glu-plasminogen. Glu-plasminogen (5 nmol/L) was incubated with tcuPA (5 nmol/L), the plasmin chromogenic substrate (100 µmol/L), and the indicated concentrations of TF apoprotein for 15 minutes under the conditions described in Fig 4 and the absorbance at
405 nm was measured. The data are expressed relative to the absorbance
measured in the absence of TF. The mean ± SD of three separate
experiments performed in triplicate is shown.
|
|
Acceleration of plasmin formation by uPA in the presence of TF was also
evident using gel analysis. In these experiments, Glu-plasminogen (3 µmol/L) was incubated with tcuPA (400 nmol/L) in the presence of
various concentrations of TF for 20 minutes, and the reaction mixture
was analyzed using SDS-PAGE under reducing conditions
(Fig 8). TF apoprotein stimulated the
conversion of plasminogen to plasmin in a dose-dependent manner,
consistent with its effect on the generation of plasmin activity.
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| Fig 8.
TF stimulates uPA-mediated conversion of plasminogen to
plasmin. Glu-plasminogen (3 µmol/L) was incubated with tcuPA (400 nmol/L) and with various concentrations of TF apoprotein for 20 minutes, and the reaction mixture was analyzed using SDS-PAGE under
reducing conditions. Lane 1 contains plasminogen alone. Lanes 2 through
4 contain plasminogen, tcuPA, and increasing concentrations of TF (lane
2, 40 nmol/L TF; lane 3, 100 nmol/L TF; and lane 4, 300 nmol/L TF).
|
|
To determine whether TF stimulated uPA-mediated plasminogen activation
through a mechanism similar to lysine analogues, such as
6-AHA,53-55 the activation of miniplasminogen, which lacks
the kringles 1 through 4 region that contains the high-affinity
Lys-binding sites,47 was analyzed. TF had no effect on the
activation of miniplasminogen by tcuPA under the identical conditions
in which stimulation of full-length Glu-plasminogen was observed (data not shown).
The data cited above suggest that plasminogen binds to TF through its
kringles, thereby promoting its activation by uPA. Binding of other
ligands, such as 6-AHA and oleic acid, to the kringles of plasminogen
has been shown to inhibit plasmin(ogen)-mediated fibrinolysis, even
though these same reagents stimulate its activity on small
substrates.50,56 The results shown in
Fig 9 indicate that TF behaves in a similar
way. Addition of TF (14 nmol/L) to plasminogen inhibited fibrinolysis
by 74%.
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| Fig 9.
Effect of TF on uPA- and plasminogen-mediated
fibrinolysis. scuPA (1 µmol/L) was added to a fibrin clot in PBS (A1
and B1) or PBS containing 14 nmol/L TF (A2 and B2) or TF without tcuPA (C1 and C2) for 2 hours. The clot was incubated with trypan blue, washed, photographed, and scanned, and its size was measured by densitinometric analysis as described in the Materials and Methods. The
fibrin clot measured 0.75 and 0.84 cm2 in the absence of TF
and 0.18 and 0.22 cm2 in the presence of TF. The data shown
are representative of three experiments so performed. TF caused a
similar increase in clot size when tcuPA (1 µmol/L) was substituted
for scuPA.
|
|
To determine whether these results using soluble TF can be generalized
to the cell surface, we studied plasminogen binding and activation
using a stably transfected CHO cell line that expresses full length
human TF (CHO-TF). Specific binding of plasminogen to CHO-TF was
stimulated 70% relative to the parental wild-type cells
(Fig 10). Furthermore, the addition of
CHO-TF to plasma clots inhibited fibrinolysis relative to wild-type
cells (Fig 11), similar to the effect of
soluble TF shown above.
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| Fig 10.
Binding of TF to cell-associated TF. CHO cells
expressing TF and wild-type cells (4 × 104) was incubated
with 50 nmol/L 125I-plasminogen in the
presence or absence of 100 mmol/L 6-AHA for 2 hours at 37°C. The
cells were washed three times in PBS, the cell-associated radioactivity
was measured, and the specific binding was calculated. The mean ± SEM
of three experiments is shown.
|
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| Fig 11.
Inhibition of fibrinolysis by CHO-TF. CHO cells (4 × 105) expressing TF (A) and wild-type cells (B) were
detached with EDTA, washed in PBS, and added for 3 hours at 37°C to
a fibrin clot formed in the presence of plasminogen (0.5 µmol/L;
lanes 3 and 4) or buffer (lanes 1 and 2). The wells were photographed
and the size of the zone lysis evident in the center of each well was
measured by scanning densitometry. The picture shown is representative of three experiments performed under identical conditions.
|
|
Histochemical colocalization of TF and plasminogen in vivo.
The data shown above indicate that an interaction occurs between TF
apoprotein and plasminogen in vitro. To examine whether such
interactions occur in vivo, we examined the relationship between the
distribution of TF and plasminogen in human atherosclerotic coronary
arteries using a dual-labeling approach
(Fig 12). TF apoprotein was distributed
in several locations within the atherosclerotic vessel wall, as
described previously by others.5,19-21 The most intense
staining was found in the adventitia, but TF apoprotein was also
readily identified in the medial layer, in acellular plaque, and in
large vessel endothelium adjacent to the plaque. Plasminogen was found
in large vessel and microvascular endothelium, in medial and intimal
smooth muscle cells, and in the acellular plaque. Colocalization of TF
apoprotein and plasminogen was seen in luminal endothelial cells
overlying the plaque (Fig 12A), within the acellular portions of the
plaques (Fig 12B and C), and within medial smooth cells (Fig 12D) and
smooth muscle cells surrounding small vessels in the adventitia (Fig
12E). No staining was seen when an isotype control Ig was used (Fig
12F).
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| Fig 12.
Immunohistochemical colocalization of TF apoprotein and
plasminogen in human atherosclerotic coronary artery. TF apoprotein stained blue; plasminogen stained brown (see the Materials and Methods). A large coronary artery is shown at an original magnification ×200. (A) Endothelial cells overlying plaque. (B) Acellular plaque with cholesterol cleft. (C) Acellular plaque matrix. (C) Adventitia. (E) Media of small artery within coronary artery. (F)
Negative Ig control.
|
|
| |
DISCUSSION |
The results of this study show that plasminogen binds with high
affinity to an epitope(s) on the extracellular domain of TF apoprotein.
Plasminogen binding to TF likely occurs through its kringles, because a
fragment that contains kringles 1 through 3 also binds to TF, whereas
isolated kringle 4 and miniplasminogen do not. Plasminogen and
FVII/FVIIa appear to bind to different sites on TF. Preincubation of
biosensor-immobilized TF extracellular domain with plasminogen did not
decrease the on-rate of FVIIa binding and preincubation of TF with FVII
or FVIIa had no effect on its capacity to inhibit the binding of
plasminogen to cells. Furthermore, preincubation of TF with plasminogen
had no effect on TF-induced procoagulant activity.
The results of the biosensor experiments, in particular the definitive
response transitions in the sensorgrams showing direct mass buildup at
the sensor surface upon plasminogen addition to the bulk phase and
subsequent mass reduction when plasminogen is removed, demonstrate
convincingly that plasminogen binds directly to the soluble domain of
TF in vitro. However, the calculated rate constants obtained from the
kinetics for this interaction must be considered to be estimates in
that they were calculated assuming the existence of two sites of
interaction with differing affinity (data fit in Fig 1B). This latter
assumption is not yet proven, and the sensorgram data would also be
consistent with more complicated models, such as cooperativity.
Calculation of affinities using such alternative models would yield
different values for kon, although it is likely that the
fastest rates so calculated would not be severely different from the
faster rate determined here for component 1 (Fig 1B and C).
Interestingly, TF affects plasminogen activation and its binding to
cells at concentrations substantially below the estimated kd (see
below). This suggests that the biological (membrane-associated) and
biosensor-immobilized conformations of TF may differ, that full
occupancy is not required for biologic activity of TF, or that there
exists a much higher affinity interaction not detectable in the time
scale of the sensor method. A better understanding of the structural
requirements for the interaction between plasminogen and TF will be
needed to distinguish among these possibilities.
TF also stimulates the activation of plasminogen by uPA measured using
a small plasmin substrate. Stimulation of urokinase-mediated plasminogen activation by TF was dose-dependent. Plasminogen activation by uPA was stimulated to the same extent by TF apoprotein and re-relipidated protein, although higher concentrations of the former
were required. Neither isolated phospholipids nor human serum albumin
relipidated in an identical manner as TF-accelerated plasminogen
activation, suggesting that the apoprotein is the active moiety in the
complex. The contribution of the lipid to plasminogen activation may be
similar to its presumed role in accelerating the activation of factor X
by the TF-VIIa complex and TF-VII by FXa,11 in this case by
providing an additional surface to which plasminogen or uPA may bind.
TF apoprotein and Glu-plasminogen appear to form a 1:1 molar complex
under the experimental conditions in that equimolar concentrations of
TF almost totally inhibited the specific binding of plasminogen to
HUVECs (Fig 5). Furthermore, 3.8 nmol/L TF apoprotein stimulated the
activation of 5 nmol/L plasminogen by approximately 50% (Fig 6).
However, the observation that the activation of 5 nmol/L plasminogen increased until the TF concentration reached 1 nmol/L suggests that a
more complex interaction has occurred. This was especially evident at
high concentrations of purified apoprotein (Fig 6). These data are
consistent with one of the possible explanations of the biosensor
binding isotherms mentioned above (Fig 1), suggesting from the dR/dt
plots the possible existence of a second, lower-affinity binding site
for binding of plasminogen to TF. Binding of TF to plasminogen through this second site appears to impede further stimulation. Less likely, the apoprotein may be degraded by
plasmin.57
Although TF promoted the plasminogen activator activity of uPA on
plasminogen when a small chromogenic substrate was used, TF inhibited
uPA-mediated fibrinolysis. The capacity of TF to stimulate uPA and
plasminogen-mediated cleavage of a small plasmin substrate while
inhibiting fibrinolysis is similar to the effects of
6-AHA50,56 and oleic acid.58 The difference
between the two situations likely results from binding of TF to the
lysine binding sites of plasmin(ogen) that are involved in
fibrinolysis. In accord with this interpretation, the effect of TF on
plasminogen activation against the chromogenic plasmin substrate was
not detected using a plasminogen fragment missing kringles 1 through 4, which are known to contain the high-affinity lysine binding
sites.47 Furthermore, TF inhibited the binding of
plasminogen to endothelial cells, a process mediated through its lysine
binding sites,22,59,60 in accord with the results of
optical biosensor studies showing that binding is mediated through
kringles 1 through 3. Thus, it is likely that TF occupies the lysine
binding sites because of the high affinity of this interaction, and
therein prevents plasmin(ogen) from binding to fibrin.
The results of this study raise the possibility that TF may contribute
a high-affinity cellular binding site for plasminogen at sites of
vessel injury or inflammation. For example, TF is abundant in
atherosclerotic plaques,5,19,21,61 whereas the concentration of plasminogen is very low compared with its
concentration in plasma.62 We observed that plasminogen and
TF colocalized in specific cellular and acellular portions of the
plaque. Thus, TF may regulate the availability of plasminogen in such
extracellular compartments because of its high affinity relative to
other plasminogen binding sites described to date. In this manner, the
capacity of TF to regulate plasminogen activation on cell surfaces may contribute to its reported effects on
angiogenesis,14 tumor invasiveness,16,17 and atherosclerosis.19
| |
FOOTNOTES |
Submitted March 19, 1997;
accepted October 28, 1997.
Supported in part through National Institutes of Health Grants No.
HL40387, HL50970, and HL49839 (D.B.C.) and Grant No. 960105000 from the
American Heart Association (A.A.H.).
Address reprint requests to Douglas Cines, MD, Room 513A,
Stellar-Chance Laboratories, University of Pennsylvania, 420 Curie Blvd, Philadelphia, PA 19104.
The publication costs of this article were defrayed in part by page
charge payment. This article must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
| |
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Abstract
Introduction
Abstract