Adsorption of Fibrinogen to Droplets of Liquid
Hydrophobic Phases
Functionality of the Bound Protein and Biological Implications
Gregory S. Retzinger, Ashley P. DeAnglis, Samantha J. Patuto
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Abstract—Fibrinogen adsorbs spontaneously from aqueous media containing that protein to droplets of liquid hydrophobic
phases dispersed in those same media. Examples of such phases include mineral oils, straight-chain hydrocarbons, and
various plant- and animal-derived oils. Lecithin preexisting on the surface of oil droplets reduces significantly the
amount of fibrinogen that can otherwise bind to them. When bound, fibrinogen remains active in the classic sense of
fibrin gelation. As a consequence, oil droplets coated with fibrinogen can participate in a host of biologically important
adhesive processes in which the protein would be expected to participate. Certain polyanions, eg, heparin, pentosan
polysulfate, dextran sulfate, and suramin, bind to adsorbed fibrin(ogen) and prevent thrombin-dependent adhesion of
fibrinogen-coated surfaces. Thus, these polyanions can be used to prevent adhesion between fibrin(ogen)-coated oil
droplets and other fibrin(ogen)-coated surfaces. Potential practical applications and biological implications of these
phenomena are presented and discussed. (Arterioscler Thromb Vasc Biol. 1998;18:1948-1957.)
Key Words: fibrinogen n atherosclerosis n oils n polyanions n drug delivery
W
e have explored the operation and biological processivity of fibrin(ogen) at interfaces1,2 because we believe such exploration will yield insight relevant to a mechanistic understanding of a host of fibrin(ogen)-dependent
physiological and pathophysiological processes.3–7 Earlier, we
showed that fibrinogen adsorbs avidly from aqueous media
containing the protein to microscopic hydrophobic beads
dispersed in those media.1 Phospholipids or other amphiphiles
preexisting on hydrophobic beads prevent the adsorption of
fibrinogen and other proteins by rendering the surface of the
beads hydrophilic.8,9 When bound to beads, fibrinogen remains functional in the classic sense of fibrin gelation.1 As a
consequence, fibrinogen-coated beads aggregate when stirred
in the presence of thrombin, and they can either be incorporated into developing fibrin clots or adhere to other fibrincoated surfaces they contact.1,2 Importantly, heparin binds to
adsorbed fibrin(ogen) and prevents adhesion between fibrincoated surfaces.10
During the course of our investigations, we “discovered”
that fibrinogen, like many other proteins, adsorbs in stable
fashion from aqueous media to droplets of liquid hydrophobic
phases. As is the case when fibrinogen is bound to solid
polymeric beads, that protein is functional on the surface of
oil droplets. Thus, like fibrinogen-coated beads, fibrinogencoated oil droplets can be incorporated into developing fibrin
clots, and they can adhere to other fibrin-coated surfaces that
they contact.
In this report, we characterize the binding of fibrinogen to
microscopic droplets of several liquid hydrophobic phases.
We demonstrate that the bound protein can mediate adhesion
of these droplets to other fibrin-coated surfaces, and this
adhesion can be prevented by certain polyanions. The relevance of these findings both to the formulation of vehicles for
targeted delivery of drugs and to our understanding of
pathophysiological processes involving deposition of hydrophobic phases, eg, atherosclerosis, is discussed.
Methods
Reagents and Chemicals
Human fibrinogen FIB3 was from Enzyme Research Laboratories.
Before use, the buffer composition of the commercial fibrinogen was
changed by gel permeation chromatography using Sephadex G-25
(Pharmacia) as the matrix material and either 0.02 mol/L Tris-HCl,
pH 7.40, or 0.01 mol/L Na2HPO4 and NaH2PO4, pH 7.40, containing
0.145 mol/L NaCl (PBS) as the eluent. This fibrinogen was then
divided into aliquots and stored frozen at 220°C until use. Before
use, a frozen aliquot of fibrinogen solution was thawed to room
temperature, diluted in buffer as appropriate, and then heated to 37°C
to dissolve any residual cold-precipitable fibrinogen. Fatty acid–free
BSA was from ICN. For some experiments, fibrinogen was uniformly labeled using Na125I from Amersham and Iodo-Gen from
Pierce. Egg yolk L-a-lecithin was from Avanti Polar Lipids. Drakeol
6-VR and Drakeol 32, white mineral oils containing paraffin and
naphthene hydrocarbons ranging from '18 to 36 carbon atoms,
were from Penreco. [1-14C]Dodecane of specific activity 151.7
MBq/mmol was from Sigma Chemical Co. Dodecane, olive oil,
safflower oil, soybean oil, triolein, cholesteryl oleate, squalene,
squalane, ristocetin sulfate, human thrombin (.3000 NIH U/mg),
hirudin (3000 U/mg), the sodium salt of pentosan polysulfate [Mr(ave)
'3800], and the acetate salt of the tetrapeptide Gly-Pro-Arg-Pro
Received April 16, 1998; revision accepted June 5, 1998.
From the Department of Pathology and Laboratory Medicine, University of Cincinnati Medical Center, Cincinnati, Ohio.
Correspondence to Gregory S. Retzinger, MD, PhD, Department of Pathology and Laboratory Medicine, University of Cincinnati Medical Center,
Cincinnati OH 45267-0529.
© 1998 American Heart Association, Inc.
Arterioscler Thromb Vasc Biol. is available at http://www.atvbaha.org
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Retzinger et al
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(GPRP) were also from Sigma. The sodium salts of porcine mucosa
heparins of average molecular weights 14 250, 3700, and 5100 were
from Calbiochem. The sodium salt of unfractionated,
pharmaceutical-grade porcine mucosa heparin, 5000 U/mL, was
from Elkins-Sinn. Sodium suramin was the generous gift of the Drug
Service of the Centers for Disease Control and Prevention, Atlanta,
Ga. The sodium salt of dextran sulfate of Mr(ave) '8000 was from
Sigma, and the sodium salt of dextran sulfate of Mr(ave) '40 000
(range, 37 000 to 43 000) was from Chemical Dynamics. Before its
use, water was first deionized, and then distilled in an all-glass
apparatus. All organic solvents were of a grade suitable for highperformance liquid chromatography. All other chemicals were of the
highest quality available commercially.
Fresh human plasma and fresh serum were prepared from the
blood of healthy donors. For plasma, blood was drawn directly into
evacuated, siliconized glass tubes (Becton Dickinson) containing
sodium citrate, yielding a final concentration of the anticoagulant of
0.013 mol/L. For serum, blood was drawn directly into evacuated
glass tubes (Becton Dickinson) where it was left undisturbed for 2
hours while it clotted at room temperature. Cells were removed from
anticoagulated blood and from clotted blood by centrifugation at
1500g for 15 minutes, and the corresponding plasma or serum was
then aspirated and, unless specified otherwise, used immediately
thereafter. As necessary, the fibrinogen concentration of citrated
plasma samples was determined using the method of Clauss.11
Inactivation of proteinases in citrated plasma was accomplished by
heating the plasma sample at 60°C for 30 minutes. With heating, a
fibrin(ogen)-rich coagulum developed in the plasma. After the
sample was cooled to room temperature, this coagulum was removed. Lyophilized fibrinogen was then added to the medium,
yielding a fibrinogen concentration of 2.931026 mol/L.
Emulsification of Liquid Hydrophobic Phases
High-pressure extrusion was used to prepare emulsions. For this
purpose, either 140 or 220 mL of a liquid hydrophobic phase was
first added to a clean, 12375-mm glass tube. To a tube containing
140 mL of oil was then added 3.5 mL of PBS; to a tube containing
220 mL of oil was then added 5.0 mL of PBS. After brief agitation,
an oil-water mixture was passed 5 times under high pressure (15 000
psi) through the aperture of an automated homogenizer
(EmulsiFlex-20,000-B3).
Isolation of Droplets of Liquid
Hydrophobic Phases
By centrifugation, droplets were separated from the aqueous medium
in which they were prepared. After the droplet-free medium was
aspirated, droplets were washed 3 times with 2.0 mL of fresh buffer
each time. For all droplets other than those containing lecithin, the
relative centrifugal force and duration of the initial centrifugation
were 1500g and 20 minutes, respectively. Subsequently, washes of
lecithin-free droplets were performed by centrifugation at 1500g for
5.0 minutes. To isolate droplets that had been emulsified in the
presence of lecithin, the entire emulsion was first transferred to a
clean, round-bottom, glass tube. The emulsion was then centrifuged
at 9400g for 60 minutes. After the droplet-free medium was
removed, droplets were washed 3 times with 2.0 mL of fresh buffer
each time. Each of these washes involved centrifugation at 9400g for
20 minutes. After their isolation and wash, droplets were redispersed
as necessary to an apparent absorbance of 1.0 at 500 nm with a
cuvette of 1.0-cm-path length. The medium for this purpose was 0.02
mol/L Tris-HCl, pH 7.40, containing 1.0 mg/mL BSA.1
Visualization of Droplets of Liquid
Hydrophobic Phases
Light microscopy was used to visualize both monodisperse droplets
and aggregates of droplets. For this purpose, a small volume of an
emulsion was placed onto a microscope slide and then overlaid with
a coverslip. Permanent records of microscopic views of droplets
were obtained by photomicroscopy.
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Sizing of Droplets of Liquid Hydrophobic Phases
For 1 set of experiments, the size of droplets of various oils was
determined using a laser diffraction particle-size analyzer (LS 230,
Coulter). Refractive indices of 1.47 (olive oil) and 1.33 (water) were
used when fitting the light-scattering data to the instrument’s
preprogrammed sizing algorithm.
Fibrinogen Binding Studies
We assessed the binding of 125I-fibrinogen from buffer to emulsified
droplets of several liquid hydrophobic phases. For this purpose, 1.0
mL of PBS containing 2.80 mg of 125I-fibrinogen was added to 4.0
mL of PBS containing 176 mL of emulsified oil droplets. After
centrifugation of the dispersion, the radioactivity associated with the
droplet-free reaction medium was measured. When droplets contained lecithin, the medium was cleared by centrifugation at 9400g
for 1.0 to 6.0 hours. When droplets did not contain lecithin, the
medium was cleared by centrifugation at 1500g for 20 minutes. The
difference between the total radioactivity of a sample and that
remaining in the medium after separation of the droplets yielded the
radioactivity, hence fibrinogen, bound to the oil droplets.
The binding of fibrinogen from a citrated plasmalike medium to
olive oil droplets was also assessed. We used for these studies
citrated plasma that had been supplemented with various amounts of
125
I-fibrinogen. The fibrinogen concentration of the citrated virgin
plasma was 7.131026 mol/L. Three milliliters of 125I-fibrinogen–
supplemented plasma was added to an equivalent volume of PBS
containing 132 mL of freshly prepared olive oil droplets. After 30
minutes, this dispersion was mixed with an equivalent volume of
aqueous sucrose, 78% (wt/vol), and the droplets were then floated by
centrifugation for 3 hours at 7000g. The resulting “cream” layer was
washed twice with 10 mL of aqueous sucrose and centrifuged for 1.0
hour at 7000g. The radioactivity associated with the washed cream
layer was then measured. Using the known concentration of endogenous fibrinogen in the plasma, the specific activity of the 125Ifibrinogen supplementing that medium, and the radioactivity associated with the oil droplets, we determined the fibrinogen bound to
the olive oil droplets.
The time dependence of the association of 125I-fibrinogen with oil
droplets dispersed in either buffer or heat-treated, citrated plasma
was assessed as follows. Two milliliters of oil droplets that had been
coated with 125I-fibrinogen was dispersed, with continuous stirring,
into 20 mL of 1 of the 2 aqueous phases. After various lengths of
time, 1.0-mL aliquots of the stirred dispersion were removed, and the
oil and aqueous phases of these aliquots were separated by centrifugation. Subsequently, the radioactivities associated with the oil and
aqueous phases were determined.
Aggregation of Fibrin-Coated Oil Droplets and
Dissociation of Aggregates of Fibrin-Coated
Oil Droplets
When stirred in the presence of thrombin, fibrinogen-coated oil
droplets aggregate, a consequence of interparticle fibrin formation.1
Thus, thrombin-inducible aggregation of such droplets is a convenient measure of the functionality of bound fibrinogen. Several
methods were used to monitor both aggregation of fibrin-coated oil
droplets and dissociation of aggregates of fibrin-coated droplets. One
method involved simply applying 200 mL of an emulsion containing
fibrinogen-coated oil droplets to a smooth surface and then mixing
into this emulsion an amount of thrombin, 0.5 NIH U in 20 mL, with
a wooden spatula. With stirring, droplets coated with a dense layer of
fibrinogen aggregate within seconds after adding the enzyme, a
process obvious to the naked eye. With the addition of certain
substances, aggregates of fibrin-coated particles dissociate, yielding
monodisperse particles.1,10 This dissociation was monitored visually.
Permanent records of these phenomena were obtained
photographically.
Another method used to monitor both the aggregation of droplets
and the dissociation of droplet aggregates involved a platelet
aggregometer.1 A typical aggregation assay was performed at room
temperature as follows. A 0.5-mL dispersion of droplets in 0.02
mol/L Tris-HCl, pH 7.40, containing 1.0 mg/mL BSA was added to
a cylindrical, glass, sample cuvette (ID, 6 mm). The apparent
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Fibrinogen and Liquid Hydrophobic Phases
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absorbance of test dispersions was 1.0 at 500 nm when a cuvette of
1.0-cm-path length was used. As reference material, a dispersion of
polystyrene beads of diameter 0.94560.0064 mm (Seradyn) was
used. The apparent absorbance of this reference material was 0.5 at
500 nm when a cuvette of 1.0-cm-path length was used. Once the
baseline signal of the stirred (1000 rpm) test sample was established,
20 mL of an aqueous solution containing 0.5 NIH U thrombin was
added to the reaction cuvette. The relative absorbance of the sample
as a function of time after addition of the enzyme was then recorded.
For the purpose of this report, a value of 1.0 was assigned arbitrarily
to the maximal change of the aggregometry signal that occurred after
the addition of 0.5 NIH U thrombin to a dispersion of fibrinogencoated droplets of mineral oil (Drakeol 32).
Just as particle aggregation can be monitored turbidimetrically
with a platelet aggregometer, so too, can dissociation of aggregates
of particles be monitored with an aggregometer.1,10 Dissociation of
aggregates of fibrin-coated oil droplets was assessed by using the
same volumes, conditions, and instrument parameters described for
aggregation assays, except that fibrinogen-coated droplets in the
sample cuvette first had to be aggregated. For this purpose, 20 mL of
a buffered aqueous solution of thrombin, 0.5 NIH U, was added to
stirred, monodisperse, fibrinogen-coated droplets. Within 15.0 minutes after addition of the enzyme, droplets had aggregated maximally, and the resulting aggregates could be used to assess aggregate
dissociation. Test reagents were added in 20 mL of water to the
reaction cuvette, and the state of aggregation of the droplets was
followed turbidimetrically as a function of time after the reagent was
added.
Studies Involving Solution-Phase Fibrin Clots
Round-bottom, 12375-mm glass tubes were used as reaction vessels
to explore interactions of variously coated droplets of olive oil and
[1-14C]dodecane, 95/5 vol/vol, with solution-phase fibrin clots. Ten
microliters of buffer containing 0.25 NIH U thrombin was added to
200 mL of PBS containing 1.831025 mol/L fibrinogen. Solutionphase clots formed in this fashion12 were then left undisturbed for 60
minutes. For 1 experiment, 2.0 IU hirudin in 10 mL of buffer was
placed on each of several clots. The exposed, uppermost surface of
each clot, 1.13 cm2, was overlaid with 100 mL of buffer containing
10 mL of droplets of a particular olive oil/[1-14C]dodecane emulsion.
The contents of the reaction vessel were then agitated for 6 seconds
with a vortex apparatus. To remove unbound oil droplets, a clot was
“washed’” twice with 1.0 mL of fresh buffer each time and gentle
agitation. After each wash, the unbound oil droplets were decanted.
In the case of clots that had been overlaid with fibrinogen-coated oil
droplets, an attempt was made to dissociate from the clots any bound
droplets by using 1 of several solutions. These solutions included
(1) 100 mL of buffer containing 0.25 IU plasmin, (2) 100 mL of
buffer containing 5.031023 mol/L GPRP, and (3) 100 mL of buffer
containing 100 USP U unfractionated, pharmaceutical-grade heparin.
Both untreated clots and clots that had been treated with 1 of the test
solutions were then washed another 2 times as described above using
1.0 mL of fresh buffer for each wash. After removing from the
surface those oil droplets that could be liberated, the remaining
clot-associated radioactivity was determined. Before this radioactivity was measured, clots were dissolved by the addition of 0.5 mL of
8.0 mol/L urea. The entire volume of solubilized material ('0.7 mL)
was then added to 5.0 mL of scintillation cocktail (Ultima Gold XR,
Packard) for measurement. Each test sample was prepared in
quadruplicate.
Analysis of Data
Concentration-dependent data were paired with the corresponding
concentrations and then fit to an appropriate equation described in
the text. The best values for the parameters of an equation were
determined using the paired data and a nonlinear least-squares
regression method.13
Results
Fibrinogen Binds to and Stabilizes Droplets of
Emulsified Liquid Hydrophobic Phases
Surfactants can be used to stabilize emulsions consisting of 2
partially miscible or immiscible phases such as mineral oil
Figure 1. Fibrinogen stabilizes emulsified droplets of liquid
hydrophobic phases. Left, Mineral oil (Drakeol 32) floats as a
discrete phase on aqueous buffer; right, fibrinogen stabilizes
microscopic droplets of mineral oil dispersed in an aqueous
phase.
and water (Figure 1). By occupying the interface between the
continuous and discontinuous phases, the surfactant lowers
the interfacial tension of the system, thus reducing the drive
toward self-coalescence of the individual phases. As shown
on the right in Figure 1, fibrinogen, an amphiphilic protein
and as such, a surfactant, effectively stabilizes microscopic
droplets of emulsified mineral oil. In the absence of fibrinogen or other added surfactant, the water and oil of the
emulsion separate back into discrete, continuous phases
within 20 minutes. In contrast, in the presence of fibrinogen,
the oil droplets remains monodisperse, eventually forming
cream layers over a time course ranging in oil-dependent
fashion from hours to days. Liquid hydrophobic phases that
we have found can be emulsified in the presence of fibrinogen
with the resulting droplets remaining monodisperse for various extended periods include, in addition to mineral oils,
olive oil, olive oil/cholesteryl oleate mixtures, triolein, triolein/cholesteryl oleate mixtures, soybean oil, safflower oil,
squalene, squalane, and dodecane. The amounts of fibrinogen
that bind to various oil droplets prepared by high-pressure
extrusion and then exposed to the protein according to our
standard protocol are shown in Figure 2.
Lecithin Prevents the Binding of Fibrinogen to
Droplets of Liquid Hydrophobic Phases;
Cholesteryl Oleate Does Not
Earlier, we showed that lecithin preexisting at rather high
packing density, ie, '0.014 molecule/Å2 ('70 Å2/molecule),
on the surface of hydrophobic beads prevents the binding of
Retzinger et al
Figure 2. Fibrinogen binds to microscopic droplets of various
liquid hydrophobic phases. Oil droplets were dispersed in PBS
containing 1.6531026 mol/L fibrinogen. See text for details.
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fibrinogen to those beads.8 In contrast, cholesteryl oleate
coated to a similar nominal packing density does not influence the rather significant amount of fibrinogen that otherwise binds to the surface of the beads. We wondered whether
lecithin or cholesteryl oleate, when included in the formulation of an oil emulsion, would affect the association of
fibrinogen with oil droplets in the same way that they affect
the binding of the protein to hydrophobic beads.
To address this issue, we first prepared emulsions containing olive oil and various quantities of either lecithin or
cholesteryl oleate. Next, we used laser diffraction to size
droplets prepared from olive oil alone, olive oil and 1.0 mol%
lecithin, and olive oil and 2.0 mol% cholesteryl oleate. We
found that all of the measured droplets had virtually the same
size distribution, a mean diameter of '3.561.8 mm. We then
quantified the binding of 125I-fibrinogen to the droplets. From
the data of Figure 2 and the diameter of the droplets, we
determined that the fibrinogen on droplets of virgin olive oil
likely exists as a monomolecular layer.2,14,15 Furthermore, the
packing density of the fibrinogen, 5.831025 molecules/Å2 (ie,
'17 300 Å2/molecule), is consistent with the molecules of
the protein being oriented with their long axes more normal
than tangential to the interface.2,15 We found next that no
amount of cholesteryl oleate used for these experiments (up
to 4 mol% of the lipid of an emulsion) reduced the quantity
of fibrinogen that associated with droplets of otherwise virgin
olive oil (Figure 3). This result is in keeping with both the oil
solubility and the marginal amphiphilicity of cholesterol
ester.8,16 –18 As shown in the same figure, however, as little as
1.0 mol% lecithin reduced to nearly zero the amount of
fibrinogen that otherwise would bind to olive oil droplets.
Indeed, from the volume of oil used for the experiment (176
mL); the mean diameter of the spherical droplets; and the
weight average molecular weights of lecithin (760) and
triolein (885, “olive oil”), one calculates that the point of
intersection of the 2 extrapolated linear regions of the figure
corresponds to a nominal molecular area for each lecithin
molecule of '43 Å.2 This nominal molecular area is in good
agreement with that of lecithin maximally packed at the
air-water interface, '75 Å.2,19 The difference between these 2
areas may be more apparent than real because, in the case of
oil droplets, the phospholipid partitions between the bulk and
surface phases of the oil.17 Taken together, these data are
December 1998
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Figure 3. Effect of lipid amphiphiles on the binding of fibrinogen
to droplets of olive oil. When included in the oil phase, lecithin
(F) reduces the binding of fibrinogen to droplets of otherwise
virgin olive oil; cholesteryl oleate (E) does not. The straight-line
regions of data that relate to lecithin represent an empiric fit of
the data to a titration curve. The arrow indicates lecithin content
at the intersection of the extrapolated straight-line fits. All data
points represent mean6SD of duplicate determinations.
eminently consistent with the proposal that fibrinogen is
excluded from the surface of the droplets when that surface is
rendered hydrophilic as a consequence of occupancy by
“tightly packed” phospholipid.8
Fibrinogen Bound to Droplets of Liquid
Hydrophobic Phases Is Functional
Fibrinogen-coated droplets of liquid hydrophobic phases can
be isolated and then redispersed in fresh aqueous medium
containing either no protein or proteins other than fibrinogen,
eg, BSA (Figure 4A). The fibrinogen bound to oil droplets is
functional, as demonstrated by the macroscopic aggregation
of droplets when they are stirred in the presence of thrombin
(Figure 4B). As expected, thrombin-induced aggregation of
fibrinogen-coated droplets is inhibited by hirudin, a rapid and
potent inhibitor of this enzyme (Figure 4C). Photomicrographs of monodisperse, fibrinogen-coated mineral oil droplets and aggregates of fibrin-coated mineral oil droplets are
shown in Figure 5.
Droplet aggregation and related phenomena can be monitored conveniently using either a platelet aggregometer or
another photometric device. As shown in Figure 6, the
apparent absorbance of a stirred dispersion of fibrinogencoated oil droplets decreases rapidly after the addition of
thrombin to the dispersion. This decrease in absorbance
corresponds to the aggregation of droplets, a consequence of
interparticle fibrin dimerization.1 By inhibiting thrombin,
hirudin prevents aggregation. Thus, just as fibrin(ogen) binds
to and remains operational on solid, microscopic, hydrophobic beads,1 so too, does fibrinogen bind to and remain
operational on droplets of liquid hydrophobic phases.
We assessed next whether various measures would liberate
fibrin-coated oil droplets from the surface of a solution-phase
fibrin clot.12 For this purpose, we first prepared olive oil
droplets containing 0.5 mol% [1-14C]dodecane. These radiolabeled droplets were coated with fibrinogen, washed, and
concentrated. An aliquot of the concentrated, fibrinogencoated droplets was then overlaid onto the exposed surface of
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Fibrinogen and Liquid Hydrophobic Phases
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Figure 6. Aggregometry tracing of fibrinogen-coated droplets of
mineral oil (Drakeol 32). A, In the presence of thrombin, the
apparent absorbance of a stirred dispersion of fibrinogencoated oil droplets decreases with time. B, Hirudin inhibits the
aggregation of fibrinogen-coated droplets that otherwise occurs
after addition of thrombin. The arrow indicates addition of
thrombin. See Reference 1 and text for details.
Figure 4. Macroscopic assessment of fibrinogen-coated droplets of mineral oil (Drakeol 32). A, In the absence of thrombin,
fibrinogen-coated droplets are monodisperse and form a confluent layer. B, With the addition of thrombin, fibrinogen-coated
droplets aggregate. C, Hirudin inhibits the aggregation of
fibrinogen-coated droplets that otherwise occurs in the presence of thrombin.
a uniform, thrombin-containing, fibrin clot, and the remaining
radioactivity associated with the clot after 1 of several
treatments was determined (Figure 7). As expected, hirudin
coadministered with fibrinogen-coated oil droplets reduced
significantly the association of droplets with the clot, indicat-
Figure 5. Microscopic assessment of fibrinogen-coated droplets of mineral oil (Drakeol 32). The droplets are of diameter
'3.561.8 mm. Left, Fibrinogen-coated droplets in the absence
of thrombin; right, fibrinogen-coated droplets in the presence of
thrombin.
ing that adherence of the droplets to the clot is likely a
consequence of noncovalent interactions between the fibrin
of the clot and the fibrin generated on the droplets. Plasmin,
GPRP, and unfractionated, pharmaceutical-grade heparin
each effectively dislodge droplets bound to the surface of
clots. The mechanism by which plasmin liberates droplets
undoubtedly involves digestion of fibrin existing between the
droplets and the clot surface, because plasminogen does not
free fibrin-coated particles from the surface of a solutionphase clot.12 Heparin10 and GPRP,20,21 on the other hand, likely
interfere with noncovalent interactions between fibrin on the
droplets and the fibrin at the clot surface. We conclude from
these experiments that the functionality of fibrin(ogen) bound
to oil droplets is similar in all measured respects to that of
fibrin(ogen) in solution.
Ristocetin Flocculates Fibrinogen-Coated
Oil Droplets
Ristocetin dimers flocculate fibrinogen, a consequence of
complexation of the bifunctional dimers with certain b-turns
Figure 7. Effect of various agents on the adherence of droplets
of olive oil:[1-14C]dodecane, 95/5 vol/vol, to a solution-phase
fibrin clot. Results are expressed as mean6SEM (n54) percentage of control radioactivity associated with clots in the absence
of any treatment. See Reference 12 and text for details.
Retzinger et al
December 1998
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Figure 8. Ristocetin dimers flocculate fibrinogen-coated droplets of olive oil. See text for details.
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of the protein.22 As shown in Figure 8, dimeric ristocetin also
flocculates fibrinogen-coated oil droplets. Ristocetin dimers
do not, however, flocculate droplets coated with an irrelevant
protein, ie, BSA. Thus, as assessed using ristocetin as a
structural probe, fibrinogen retains important solution-phase
features when it is bound to oil droplets.
Fibrinogen Is Stable on Droplets of Liquid
Hydrophobic Phases
Fibrinogen dissociates rather slowly from oil droplets dispersed in buffer: only '4% of the radioactivity bound
initially to saturation on droplets of either mineral oil
(Drakeol 32) or olive oil is liberated from the droplets when
they are incubated in buffer for 48 hours. In the case of olive
oil, this holds true even when the aqueous phase is heattreated, fibrinogen-supplemented plasma: 95% of the radioactivity remains associated with the droplets after 48 hours.
Such results are consistent with the notion that the binding of
fibrinogen to hydrophobic surfaces is essentially irreversible,
a phenomenon due at least in part to interfacial “denaturation” of the protein.1,8,14,15 The protein is not so denatured as
to lose function, however. As shown in Figure 9, the
thrombin-inducible aggregation of fibrinogen-coated olive oil
droplets changes little, if any, after incubation for 24 hours in
citrated plasma.
Fibrinogen Adsorbs From Plasma to Droplets of
Liquid Hydrophobic Phases and Is Functional
Fibrinogen adsorbs rapidly and in relatively large quantity
from blood to solid, hydrophobic surfaces in contact with that
medium.4,23 For this reason, we assessed next whether fibrinogen would adsorb from citrated plasma to oil droplets. Such
an assessment seemed apropos, because biologically relevant
lipid particles in vivo, ie, lipoproteins, are bathed continuously in a fibrinogen-rich medium, and fibrinogen appears to
contribute to the initiation, development, and growth of
atherosclerotic plaques.3,24,25
As shown in Figure 10, olive oil droplets exposed to fresh,
citrated plasma but not droplets exposed to fresh serum
aggregate in the presence of thrombin, indicating that fibrinogen adsorbs from plasma to the droplets and is functional.
As shown in Figure 11, the rate and extent of aggregation of
plasma-exposed oil droplets depend in turn on the concentra-
Figure 9. Aggregometry tracing of fibrinogen-coated droplets of
olive oil before (A) and after (B) incubating them for 24 hours in
heat-treated, fibrinogen-supplemented (2.931026 mol/L),
citrated normal plasma. The arrow indicates addition of
thrombin.
tion of fibrinogen in the plasma. Using 125I-fibrinogen as a
tracer, we found that the quantity of fibrinogen adsorbed to
droplets incubated in plasma containing 7.131026 mol/L (241
mg%) fibrinogen was 0.40 mg/mL oil, and the quantity of
fibrinogen bound to droplets from the plasma containing
13.231026 mol/L (450 mg%) fibrinogen was 0.87 mg/mL oil.
Thus, while plasma proteins, in addition to fibrinogen, must
contribute to the final layer of protein adsorbed from plasma
to oil droplets, the fibrinogen that does adsorb is functional,
and its interfacial concentration increases with increasing
plasma concentration of the protein.
Heparins and Other Polyanions Prevent the
Mutual Adhesion of Fibrin-Coated Surfaces
Earlier, we demonstrated that heparins and related polyanions, in the absence of any cofactor, bind with high affinity to
fibrin(ogen) adsorbed to solid, polymeric beads.10 As consequences of this binding, the mutual adhesion of fibrinogencoated beads that otherwise occurs in the presence of thrombin can be prevented, and preexisting aggregates of fibrincoated beads can be dissociated. Discovery of these
phenomena led us to suggest that the binding of heparin to
adsorbed fibrin(ogen) might serve some general, cofactorindependent, “antiadhesive” function.10
Believing,10 as do others,26 –29 that the direct interaction of
heparin with fibrin(ogen) contributes in vivo to the anticoag-
1954
Fibrinogen and Liquid Hydrophobic Phases
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Figure 12. Dose-dependent inhibition of thrombin-inducible
aggregation of fibrinogen-coated droplets of mineral oil (Drakeol
32). F, Heparin, Mr(ave)514 250, Kd50.7060.10 nmol/L; Œ, heparin, Mr(ave)55100, Kd516.2166.55 nmol/L; E, pentosan polysulfate, Mr53800, Kd517.2560.92 nmol/L; D, dextran sulfate,
Mr(ave)58000, Kd523.7863.47 nmol/L; M, heparin, Mr(ave)53700,
Kd524.4261.18 nmol/L; f, suramin, Mr51429,
Kd510 04061668 nmol/L; and l, dextran sulfate,
Mr(ave)540 000. See Reference 10 and text for details.
Figure 10. Macroscopic assessment of fibrinogen-coated droplets of olive oil. A, Droplets of olive oil coated from an aqueous
solution containing fibrinogen alone as protein; B, droplets of
olive oil coated with fibrinogen alone and then exposed to
thrombin; C, droplets of olive oil coated with fibrinogen alone
and then exposed simultaneously to hirudin and thrombin; D,
olive oil droplets isolated and washed after exposure to citrated
plasma (fibrinogen57.131026 mol/L); E, same as D but after
addition of thrombin; F, same as D but after simultaneous addition of hirudin and thrombin; G, droplets of olive oil isolated and
washed after exposure to normal serum; and H, same as G but
after addition of thrombin.
ulant activity of the mucopolysaccharide, we tested whether
heparins and other polyanions prevent thrombin-inducible
aggregation of fibrinogen-coated droplets of mineral oil. For
these studies, we used unfractionated heparin, low-molecularweight heparins, pentosan polysulfate, suramin, and dextran
sulfates. As shown in Figure 12, all of the materials tested
reduced in a dose-dependent fashion the maximal rate of
aggregation of droplets. For all but dextran sulfate of Mr(ave)
'40 000, the reduction in rate as a function of polyanion
concentration obeys well the relationship Vobs5Vmax/(11
P/Kd), where Vmax is the maximal rate of aggregation in the
absence of polyanion, Vobs is the observed rate of aggregation
in the presence of polyanion, P is the molarity of the
polyanion, and Kd is the equilibrium dissociation constant of
the complex. The polyanions fall into 4 distinct groups.
Unfractionated heparin [Mr(ave) '14 250] is by far the most
potent inhibitor of the series, followed by, in decreasing order
of potency, the group consisting of the sulfated polysaccharides of low molecular weight [ie, Mr(ave)’s between 4000 and
8000], suramin [Mr51429], and high-molecular-weight dextran sulfate [Mr(ave) '40 000]. For the heparins and dextran
sulfates, these results are the same as those obtained using
fibrin(ogen)-coated, solid, polymeric beads.10 The apparent
dependence on the length of the dextran sulfate polymer
compared with that of heparin and the relative impotency of
suramin with respect to unfractionated heparin probably
indicate some fundamental structural requirement for complex formation.
Discussion
Figure 11. Aggregometry tracing of droplets of olive oil isolated
and washed after exposure to either citrated plasma or serum.
A, Plasma fibrinogen concentration513.231026 mol/L; B,
plasma fibrinogen concentration57.131026 mol/L; C, same as B
but hirudin was added before the addition of thrombin; and D,
serum. The arrow indicates addition of thrombin.
We have demonstrated that fibrinogen adsorbs spontaneously
from aqueous media containing that protein to droplets of
liquid hydrophobic phases dispersed in those same media.
Lecithin preexisting on the surface of oil droplets reduces
significantly the amount of fibrinogen that can otherwise bind
to the droplets. When bound, fibrinogen is stable and remains
active in the classic sense of fibrin gelation. As a consequence, oil droplets coated with fibrinogen can participate in
a host of biologically important adhesive processes in which
the protein would otherwise be expected to participate.
Certain polyanions appear to bind to adsorbed fibrin(ogen)
and prevent thrombin-dependent adhesion of fibrin-coated
Retzinger et al
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surfaces. What follows is a discussion of potential practical
applications and theoretical implications of these findings.
Because so many pathological lesions, eg, tumors,
granulomas, bacterial abscesses, atherosclerotic plaques—
indeed, virtually all sites of inflammation— have a thrombotic component,30 –32 it would seem worthwhile generally
to develop vehicles to deliver therapeutic drugs or imaging
agents to sites of fibrin deposition. Fibrinogen-coated oil
droplets might provide such a vehicle, especially for the
delivery of water-insoluble molecules. Although the use of
oil droplets for the delivery of hydrophobic molecules is
not new, the site-specific targeting of such droplets has
been problematic.33 Functional fibrinogen adsorbed “irreversibly” to drug-loaded oil droplets might serve to focus
those droplets to pathological sites that express thrombin
activity, thereby locally increasing drug concentration
while globally limiting nonspecific effects of therapy.
Fibrinogen-coated oil droplets might also be formulated
as adjuvants for vaccines. Earlier, we proposed that fibrinogen adsorbs from plasma to the surface of mineral oil
droplets of Freund’s and related adjuvants and acts synergistically with amphiphilic lipids expressed on the surface
of those droplets to elicit acute inflammation, granuloma
formation, immune adjuvancy, and hemorrhage.5,34 Because at that time it was our intention to expose the surface
of oil droplets as the site of biological activity of adjuvant
lipids, we substituted solid, microscopic, hydrophobic
beads for the oil droplets of conventional adjuvant emulsions.35 Using the bead system, we demonstrated conclusively that fibrinogen binds avidly and with high affinity to
beads coated with adjuvant lipids and that the bound
protein acts synergistically with the lipid microenvironment to produce the biological effects traditionally associated with adjuvant oil emulsions.5,34 Since then, many
adjuvant preparations that include oil and 1 or another
amphiphilic lipid have been formulated for successful
vaccine use.36 We suggest that precoating adjuvant oil
droplets with fibrinogen might optimize the immuneenhancing potential of the droplets, perhaps by either
facilitating or promoting the interactions of the particles
with macrophages.
Biomedical materials scientists have long appreciated
that fibrinogen, of all the plasma proteins, adsorbs rapidly
and preferentially to hydrophobic, polymeric materials in
contact with blood.4,23 As a consequence of this adsorption,
blood clots are nucleated on the surface of the material
often leading, in the case of circulatory prosthetics, to the
development of an occlusive thrombus. Thus, 1 aim of
biomedical materials researchers is formulation of polymers that do not bind fibrinogen. (Importantly, a successful means that prevents the adsorption of fibrinogen and
other proteins to a surface involves coating that surface
with phospholipids.37) Another aim of biomedical materials researchers is identification of the region(s) of fibrinogen that actually contacts hydrophobic, polymeric surfaces.15,38 Attempts at this second aim have met with limited
success, both because the binding of fibrinogen to hydrophobic surfaces is essentially irreversible and because the
protein and its remnants are difficult to elute from hydrophobic surfaces.14 We propose that droplets of liquid
hydrophobic phases may help determine the identity of the
December 1998
1955
“surface recognition site(s)” of fibrinogen. Because droplets of the oils used here are readily soluble in organic
phases, there is no need to elute the protein or an adherent
remnant of the protein from the droplets. It is enough to
simply dissolve the underlying hydrophobic “scaffold,”
leaving as residual only proteinaceous material.
Perhaps 1 of the more important concepts to derive from
this study is that heparin and related polyanions prevent
adhesion between fibrin-coated surfaces, modeled here by
droplets of liquid hydrophobic phases. The sensitivity and
specificity of the phenomenon to unfractionated heparin
support the notion that this antiadhesive property exists by
design,10 begging further investigation of the phenomenon.
As concerns practical application, we suspect that these
polyanions might be used advantageously to prevent or
even reverse a host of deleterious, fibrin(ogen)-mediated,
adhesive events, particularly those occurring at sites of
inflammation.6,7,30 –32,39,40
For quite some time, biomedical scientists studying
atherosclerosis have focused on the role(s) of lipids in that
disorder. More recently, investigators have begun to seriously consider mechanistic roles for fibrinogen in atherogenesis,41– 43 in part because fibrin(ogen) is a ubiquitous
and significant component of advanced lesions of atherosclerosis, ie, plaques.3,24,25,40,44 – 46 We ourselves have proposed that the affinity of fibrinogen for extracellular
deposits of atheromatous lipids contributes to the morbid
and mortal thrombotic consequences of atherosclerosis8:
because clots are nucleated by adsorbed fibrinogen,
fibrinogen-coated lipid surfaces should be predisposed to
thrombosis. However, fibrinogen is a significant component of even the earliest detectable precursor of the
atherosclerotic plaque, the fatty streak.42 What role, if any,
could fibrinogen play in the earliest stages of plaque
development?
Blood, a fibrinogen-rich medium, is replete with lipidladen lipoproteins that percolate through the walls of blood
vessels and into the tissues. Popular theories hold that
lipoproteins, particularly LDLs, are somehow retained
within arterial walls, and this retention accounts for the
accumulation there of lipids that will eventually constitute
the plaque.47,48 Given the results of our studies, those of
others,46,49 and current opinion regarding plaque initiation,
development, and growth, it is reasonable to propose that
the localization and accumulation in vivo of lipoproteins—
or, for that matter, any lipid particle—will be mediated at
least in part by fibrinogen adsorbed to those particles. This
adsorption in turn will be dictated by the expression of
hydrophobic domains on the surface of the particles. Thus,
localization and accumulation of lipid particles within the
vasculature need not involve any specific interaction
between some particle-resident amphiphile, eg, apolipoprotein(a),50 and fibrin(ogen): for fibrinogen-coated particles to accumulate, there need only be focal production of
thrombin, such as occurs at any site of inflammation.32
Many approaches could be used to assess the validity of
this proposal. One approach might involve measuring
directly the binding of fibrinogen to various lipoproteins
and, if fibrinogen binds to them, its functionality once
bound (G.S.R. et al, unpublished data, 1997). Another
approach might involve correlating plasma fibrinogen
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Fibrinogen and Liquid Hydrophobic Phases
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concentration with some lipid-dependent aspect of plaque
growth. Still another approach might involve use of
anticoagulants as a means to prevent or limit plaque
development. All of these biological approaches have
merit, and all would likely yield valuable mechanistic
insight relevant to both atherosclerosis and its therapy.
However, physicochemical approaches of the sort presented here should also provide valuable information pertinent
to a mechanistic understanding of atherosclerosis, because
the adhesive potential conferred to a lipid particle by
adsorbed fibrinogen makes the presence of that protein on
the particle singularly important.
Which factors should be expected to influence the
“nonspecific” binding of fibrinogen to hydrophobic lipid
particles? We have demonstrated several, including the
solution-phase concentration of fibrinogen, the solutionphase concentration of species that compete with fibrinogen for the surface of particles, and amphiphiles preexisting on the surface of particles. The concentration of
particles, a measure of lipid “load,” will also influence the
equilibrium distribution of the protein, as will measures
that affect the affinity and/or capacity of individual particles for the protein. Given all of these factors, it is
reasonable to presume that the disposition of lipid particles
that percolate through the walls of the vasculature will
depend on the dynamic equilibrium that must normally
exist between bound and free forms of fibrinogen.
Acknowledgments
This work was supported by funds to G.S.R. from the Department of
Pathology and Laboratory Medicine, University of Cincinnati, and
by a Focused Giving Award to G.S.R. from Johnson & Johnson.
G.S.R. thanks Ruth Mary Retzinger for inspiration.
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Adsorption of Fibrinogen to Droplets of Liquid Hydrophobic Phases: Functionality of the
Bound Protein and Biological Implications
Gregory S. Retzinger, Ashley P. DeAnglis and Samantha J. Patuto
Arterioscler Thromb Vasc Biol. 1998;18:1948-1957
doi: 10.1161/01.ATV.18.12.1948
Arteriosclerosis, Thrombosis, and Vascular Biology is published by the American Heart Association, 7272
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