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Blood, Vol. 92 No. 11 (December 1), 1998:
pp. 4188-4197
Intravascular Coagulation Activation in a Murine Model of
Thrombomodulin Deficiency: Effects of Lesion Size, Age, and Hypoxia on
Fibrin Deposition
By
Aileen M. Healy,
Wayne W. Hancock,
Patricia D. Christie,
Helen B. Rayburn, and
Robert D. Rosenberg
From The Pulmonary Center, Boston University School of Medicine; the
Department of Pathology, Harvard Medical School, Boston, MA; and the
Department of Biology, Massachusetts Institute of Technology,
Cambridge, MA.
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ABSTRACT |
We consecutively inactivated both alleles of the thrombomodulin (TM)
gene in murine embryonic stem (ES) cells and generated TM-deficient
(TM /) chimeric mice. Quantitation of an ES-cell
marker and protein C cofactor activity indicates that up to 50% of
pulmonary endothelial cells are ES-cell derived and therefore TM
deficient. Infusions of 125I-fibrinogen into mice show a
significant increase (fourfold, P < .005) in radiolabeled
cross-linked fibrin in TM/ chimeric mouse lung as
compared with wild-type mice. However, only chimeric mice that exhibit
at least a 30% reduction in protein C cofactor activity and are at
least 15 months old display this phenotype. Immunocytochemical
localization of TM in chimeras shows a mosaic pattern of expression in
both large and small blood vessels. Colocalization of cross-linked
fibrin and neo (used to replace TM) reveals that fibrin is deposited in
TM/ regions. However, the fibrin deposits were
largely restricted to pulmonary vessels with a lumenal area greater
than 100 µm2. The hypercoagulable phenotype can be
induced in younger chimeric mice by exposure to hypoxia, which causes a
fivefold increase in  -fibrin levels in lung. Our findings show that
TM chimerism results in spontaneous, intravascular fibrin deposition
that is dependent on age and the magnitude of the TM deficiency.
© 1998 by The American Society of Hematology.
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INTRODUCTION |
THROMBOMODULIN (TM, CD141), the
cell-surface receptor for thrombin that suppresses blood coagulation by
altering thrombin substrate specificity, converts thrombin from a
potent procoagulant (clotting fibrinogen and activating platelets) to
an anticoagulant. The TM-thrombin complex cleaves protein C (PC) to
activated protein C (aPC), which catalyzes the proteolytic degradation
of activated blood clotting factors V and VIII. The physiological role
of the TM/PC pathway has been substantiated by data showing that
thrombin infusions into animals induces generation of aPC, that humans with defects in this pathway experience an acceleration of thrombin generation, and that a suppression of PC activation commonly precedes a
thrombotic episode.1-3 These observations support the
hypothesis that TM plays a critical role in the prevention of thrombus
formation.
TM was initially identified as an endothelial cell
cofactor4; however, subsequent studies documented the
presence of TM on synovial cells, meningeal cells, activated smooth
muscle cells, macrophages, and platelets.5-7 In addition to
its presence on vascular and nonvascular cells of the adult animal, TM
has been localized in the murine embryo as early as gestation day
7.5.8-11 To investigate the role that TM plays in
regulating blood coagulation in adult mice, as well as the function of
TM during murine embryogenesis, we performed gene targeting studies in
murine embryonic stem (ES) cells to generate mice heterozygous or
homozygous for the TM deletion. We previously reported that embryos
homozygous for the TM deletion die in utero by embryonic day
9.5.12 The TM-null phenotype thus precluded further
investigations into the effects of TM homozygous deficiency on
coagulation function in adult mice. As one approach to bypass the
embryonic lethal phenotype of TM-null mice, we generated homozygous
TM-null ES cells to produce TM-deficient
(TM/) chimeric mice.
In the present study, we examined the effects of a TM deficiency in
adult mice. The data show that TM/ chimeric
mice have reduced PC cofactor activity as compared with wild-type mice,
and exhibit an age-dependent increase in urea- and acid-insoluble
intravascular fibrin. Cross-linked fibrin was detected in the pulmonary
vasculature in regions of the vessel wall deficient in TM and fibrin
deposition was largely dependent on the size of the
TM/ region. Furthermore, the heightened
procoagulant phenotype of TM/ chimeric mice
could be reproduced in an age-independent manner by challenging animals
with hypoxia. These results indicate that TM is a necessary component
of the anticoagulant mechanism for the prevention of thrombosis, and
are consistent with the hypothesis that a TM deficiency is a
quantitative trait like hypertension.
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MATERIALS AND METHODS |
Generation of TM/ ES cells and chimeric
mice.
We used a single targeting construct to create TM double-deletion
mutants (TM/). Briefly,
TM+/ ES cells previously generated using the
positive/negative drug selection strategy12,13 were
reselected with the positive selecting agent, geneticin (G418;
GIBCO-BRL, Rockville, MD), as described.14 TM+/ ES cells were grown for 2 weeks in complete
media supplemented with 2.0 mg/mL geneticin and every other day during
drug reselection, washed with phosphate-buffered saline, and refed in
complete media supplemented with geneticin. DNA was isolated from
surviving clones and analyzed by the Southern blot technique and by
polymerase chain reaction to identify the TM-null ES cells. In vitro
differentiation of ES cells was performed as previously
described.15 Embryoid bodies were scored on day 4 of
differentiation for the formation of cystic structures and again on
days 7 and 10 for the presence of blood islands and contracting
myocytes. The TM/ ES cells were injected into
the blastocoel of 3.5-day C57BL/6 embryos, reimplanted into
pseudopregnant recipients, and carried to term. The resulting offspring
were maintained for analysis of a hypercoagulable phenotype.
Preparation of 125I-labeled murine fibrinogen.
Fibrinogen was isolated from the platelet-poor plasma of C57BL/6 mice
by two successive precipitations with 25% saturated ammonium
sulfate.16 Murine fibrinogen was radiolabeled with Na125I (Amersham, Arlington Heights, IL) to a
specific activity of approximately 1.0 µCi NaI/µg of fibrinogen
using Iodogen beads (Pierce, Rockford, IL). For
radiolabeling, a 200 µg/250 µL solution (2.2 µmol/L) of
fibrinogen in 0.15 mol/L NaCl, 0.02 mol/L Tris-HCl, pH 7.4, was
incubated with two Iodogen beads and 1.0 mCi Na125I (17.4 mCi 125I/µg of iodine) for 2.5 minutes at room
temperature. Radiolabeled fibrinogen was separated from free NaI by
chromatography on a 0.5 × 12.5-cm G-75 Sephadex column (Sigma,
St Louis, MO). Autoradiography of the
125I-fibrinogen following sodium dodecyl sulfate (SDS)
polyacrylamide gel electrophoresis under reducing or nonreducing
conditions showed that murine fibrinogen was the major radiolabeled
protein, with no significant contaminating proteins.
Quantitation of 125I-labeled fibrin deposition in mice.
TM wild-type mice and TM/ chimeric mice
received 0.5 µCi of 125I-labeled fibrinogen by injection
into the lateral tail vein on days 1, 3, and 5, and the organs were
collected for analysis on day 7 (the half-life of circulating
125I-fibrinogen is approximately 9 to 12 hours).
Forty-eight hours after the final injection, animals received 0.2 mL of
an anticoagulant solution containing 2.4 mg/mL heparin (80 United
States Pharmacopeia units), 10 mg/mL
 -amino-n-caproic acid, and 0.05 µg/mL (42 U/mL) aprotinin (Sigma)
in 0.15 mol/L NaCl. Mice were lightly anesthetized with ether and
killed by cardiac puncture, at which time 1.0 mL of whole blood was
collected for isolation of platelet-poor plasma. Circulating
125I-fibrinogen (cpm/µL) was determined for an aliquot of
plasma, and organs were dissected and tissue fragments prepared for
several analyses. First, organs or organ fragments were snap frozen in liquid nitrogen for later analysis of PC activity and glycerol phosphate isoenzymes. Second, organs were embedded in OCT
compound (Miles Laboratories, Eckhardt, IN) and frozen in
an isopentene bath cooled with dry ice for analysis by
immunohistochemistry. Third, tissue samples were weighed, homogenized
in extraction buffer (0.01 mol/L Na-phosphate, pH 7.5; 10 U/mL heparin;
2 mg/mL EDTA; 10 U/mL aprotinin; and 0.1 mmol/L phenylmethylsulfonyl
fluoride [PMSF]) and prepared for extraction of
125I-fibrinogen and fibrin as described.17
Homogenates were extracted at 4°C for 18 hours, centrifuged at
1,000g for 20 minutes, the supernatants removed, and the
pellets washed once in 0.5 mL of extraction buffer. The supernatant was
again removed and pooled with the previous supernatant. Similarly, both
pellets were combined, resuspended in 1.5 mL of freshly prepared 3 mol/L urea, extracted at 37°C for 2 hours with rotation, and then
centrifuged at 1,000g for 20 minutes. The urea-soluble cpm in
the supernatants and the urea-insoluble cpm in the pellets were
quantified in a gamma counter (LKB 1272 clinigamma, Uppsala, Sweden).
Determination of glycerol phosphate isoenzyme and TM cofactor
activity in TM/ chimeric mice.
The contribution of TM/ ES cells to chimeric
mice was assessed using three criteria; the first was an estimation of
the agouti coat color. The TM-null ES cells were derived from the 129 mouse, which expresses the agouti or brown coat color, and the
recipient embryo is derived from the C57BL/6 mouse, which expresses a
black coat color. The resulting chimeric offspring show a combination of agouti and black coat coloring; the more agouti coloring, the greater the ES-cell contribution to the mouse. The second criteria was
the differential electrophoretic mobility of glycerol phosphate isoenzymes, which is an indication of cell contribution to whole organs.18 The isoforms of glycerol phosphate were
electrophoretically separated on cellulose acetate, detected by
staining for enzyme activity and the relative contribution of each
isoenzyme and therefore each genotype, quantified by densitometry.
Third, the contribution of TM-null ES cells to endothelial cells was
determined using the TM-dependent activation of PC by thrombin.
Sections of mouse lung that had been frozen in liquid nitrogen
immediately after dissection were pulverized with a mortar and pestle
while still frozen. Tissue fragments were extracted in 0.3 mL of lysis
buffer (2% Triton X-100, Sigma; 0.01 mol/L Tris-HCl, pH
8.1; 10 mg/mL pepstatin; 10 mg/mL leupeptin; 0.1 mmol/L PMSF; and
0.02% NaN3) at 4°C for 30 minutes with rotation.
Extracts were then clarified by centrifugation at 16,000g at
4°C for 15 minutes. The protein concentration of the supernatants
was determined by Bradford assay. For cofactor determination, 0.05 µmol/L human PC, and 0.05 µmol/L human thrombin (Enzyme Research
Labs, South Bend, IN) plus 0.5 µg of lung extract was incubated in
0.15 mol/L NaCl; 0.02 mol/L Tris-HCl, pH 7.4; 2.5 mmol/L
CaCl2; and 5 mg/mL bovine serum albumin at 37°C for 15 minutes. Human antithrombin III (Enzyme Research Labs) and hirudin
(Sigma) were then added to stop the reaction and aPC was measured using
the colorimetric reagent, S2236 (Pharmacia, Piscataway,
NJ).
Histology and immunocytochemistry.
Organs (lung, heart, liver, kidney, and spleen) collected from
wild-type and chimeric mice were fixed in formalin or snap frozen and
stored at  70°C. Formalin-fixed tissues were embedded in
paraffin, sectioned, and stained with hematoxylin and eosin (H&E).
Cryostat sections obtained from snap-frozen tissues were used for the
localization of TM and neomycin phosphotransferase (neo, the product of
the bacterial gene used to replace the TM coding region). To identify
intravascular cross-linked fibrin, acid-insoluble fibrin was also
localized in cryosections.
For TM and neo localization, serial sections (4 µ) fixed in
paraformaldehyde-lysine-periodate (PLP) were incubated overnight at
4°C with a rat anti-mouse TM monoclonal antibody (MoAb; gift of Dr
Stephen Kennel, Oak Ridge Labs, Oak Ridge, TN) and a rabbit anti-neo
antibody (5 Prime 3; Prime Inc, Boulder, CO). To localize acid-insoluble fibrin, cryostat sections were acid washed with 2%
acetic acid in neutral-buffered formalin at room temperature for 20 minutes,19 followed by incubation with a rabbit antibody recognizing fibrinogen and fibrin (Dako, Carpinteria, CA). Sections were washed and bound primary antibodies were detected by successive incubations with bridging antibodies, rabbit peroxidase-antiperoxidase complexes (Dako), the substrate, diaminobenzidine, and finally counterstained with hematoxylin.20 Specificity controls
included the use of isotype-matched rat MoAbs or normal rabbit
immunoglobulin in place of specific primary antibodies: endogenous
peroxidase was blocked using methanol containing 0.03% hydrogen
peroxide.20
Morphometry.
Sequential tissue sections (4 µ) stained for H&E, neo, and
cross-linked fibrin were analyzed using the Bio-Quant Imaging System (R&M Biometrics, Nashville, TN). Four to six sequential
sections were cut for each staining condition and a single section from each condition was chosen as suitable to measure blood vessel lumenal
area. Three nonoverlapping fields (20× objective) and two
nonoverlapping fields (60× objective) were selected from each section, and each chosen field from a section corresponded to the same
field of the sequential sections. Lumenal areas were traced from
digitized images of 175,000 µm2 (20× objective) or
20,000 µm2 (60× objective).
Exposure of TM/ chimeric mice to
hypoxia.
TM/ chimeric mice, aged 6 to 11 months, were
exposed to hypoxic conditions by placing animals in a purge box
(Innovative Systems, Inc, Newton, MA) to achieve a final
oxygen content of 8% for 16 to 18 hours. Animals maintained in room
air were used as normoxic controls. After exposure to normoxic or
hypoxic conditions, mice received a lethal, intraperitoneal injection
of anesthesia, followed by an intravenous injection containing 300 to
450 U of heparin via the lateral tail vein. Whole lung was dissected
and snap frozen in liquid nitrogen.
Cross-linked fibrin was extracted from lung homogenates as described
above (see quantitation of 125I-labeled fibrin). The final
tissue pellet is solubilized in SDS-sample buffer containing 5%
2-mercaptoethanol, incubated at 65°C for 90 minutes,
electrophoresed on a 7.5% polyacrylamide gel, and transferred to PVDF
nylon membrane (Millipore, Bedford, MA) for Western blot analysis of -fibrin. -fibrin was detected using an
MoAb against the N-terminus of the B-fibrin chain (MoAb 59D8; gift
of Dr Marshall Runge, Galveston, TX) and enhanced
chemiluminescence (ECL; Amersham).
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RESULTS |
TM gene inactivation in ES cells and generation of
TM/ chimeras.
We produced an ES-cell clone containing a homozygous deletion of the TM
gene. The resulting clone contained the neo gene in place of the TM
coding region, thereby creating a double knockout or homozygous
TM-deletion mutant. The homozygous TM gene deletion is shown by
Southern blot analysis (Fig 1). The results
shown in Fig 1, lane 3 demonstrate that the 10-kb BglII or
wild-type TM gene fragment is absent in TM/
ES cells, and confirms the presence of a single 3.3-kb band, which
results from the introduction of a novel BglII restriction enzyme site contributed by the neo sequences.
TM/ ES cells were analyzed by Southern
blotting using three different probes corresponding to DNA sequences
both internal to and external to the inactivated locus and all three
probes demonstrate the predicted pattern (not shown). In addition,
TM/ ES cells contain a normal karyotype.
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| Fig 1.
Illustration of targeted TM locus and Southern blot of
TM-null ES cells. (A) Diagram of the targeted TM gene indicating the
5 upstream region of the TM promoter (TM 5), 153 bp of
untranslated TM sequences ( ), the neo expression cassette used to
replace the TM coding region (neo,  ), and 3 TM untranslated
sequences (TM 3). BglII restriction enzyme sites are
indicated at B. (B) Southern blot analysis of ES-cell genomic DNA.
Genomic DNA was isolated from wild-type ES cells (lane 1), from
TM+/ ES cells (lane 2) and from TM/
ES cells (lane 3) and hydrolyzed with BglII restriction enzyme.
The hydrolyzed DNA was electrophoresed, blotted, and hybridized with a
32P-labeled cDNA probe against DNA sequences within the TM
3 region. Markers are indicated in kb.
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The prolonged growth of ES cells in culture diminishes the pluripotency
of the stem cells and thus the ability to contribute uniformly to
cellular lineages during embryonic differentiation.18 We
tested TM/ ES-cell pluripotency by in vitro
and in vivo differentiation.15,21 ES-cell differentiation
under nonadherent culture conditions results in the formation of cystic
structures called embryoid bodies, which display specific developmental
markers. Hemoglobin-containing blood islands appear after 7 days of
suspension culture and clusters of contracting cells are present after
10 days in culture. These structures represent the differentiation of
hematopoietic and endothelial cells as well as cardiomyocytes. ES cells
were grown in suspension culture for 4, 7, or 11 days and scored for
the presence of cystic structures on day 4, acetylated-low density lipoprotein (LDL) uptake on day 7, and contracting cells
on day 11.22 Between 39 and 100 embryoid bodies were
counted for each condition in duplicate. There were no significant
differences in the number of cystic structures, embryoid bodies
staining positive for acetylated-LDL uptake, or contracting muscle
among all three genotypes. These results suggest that
TM/ ES cells differentiate in vitro at the
same frequency as controls. The presence of all three of these
characteristic differentiation markers in TM/
ES cells suggests that the prolonged culture conditions required to
delete the second TM allele did not adversely affect ES-cell differentiation in vitro. TM/ ES-cell
pluripotency was shown in vivo by generating ES-cell-derived chimeric
mice.
The resulting chimeric offspring (129 ES cells on a C57BL/6 background)
were initially categorized by the degree of agouti coat coloring, which
ranged from 20% to 80%. The agouti locus and the TM gene are both
located on chromosome 2 in the mouse; therefore, the agouti coat
coloring is an approximation of TM chimerism.23 For
analysis of the TM-deficiency, the animals were separated into two
groups: high-percentage chimeras (50% to 80% agouti coat coloring) or
low-percentage chimeras (20% to 50% agouti coat coloring). All
TM/ chimeras appear healthy and normal in
size and weight. Routine histological analyses of
TM/ chimeras from 2 months to 2 years of age
show no genotype-specific pathologies in any of the tissues examined or
within the lumen of blood vessels where TM is widely expressed.
Age-related increase in urea-insoluble fibrin in
TM/ chimeric mice.
To investigate the effects of a TM deficiency on the coagulation
system, we examined TM/ chimeric mice for the
spontaneous deposition of 125I-labeled fibrin in heart,
lung, liver, spleen, and kidney. Whole organ homogenates were extracted
in urea, which results in two fractions. The first fraction is composed
of urea-soluble cpm that contain 125I-fibrinogen, soluble
fibrin, and fibrin degradation products, and the second fraction is
composed of urea-insoluble cpm that contain cross-linked
125I-fibrin and fibrin degradation products (A.M.H. and
R.D.R., unpublished observations, November
1994).17 The high-percentage chimeras have up
to a fivefold increase in urea-insoluble cpm in lung (P < .005) and a twofold increase in urea-insoluble cpm in liver (P < .01) as compared with age-matched, wild-type mice and
low-percentage chimeras (Fig 2,
top). No difference is observed in urea-insoluble cpm in heart, kidney,
or spleen (results not shown). This increase in urea-insoluble cpm is
age dependent; animals attaining 15 to 20 months are affected, whereas
animals 6 to 11 months remain unaffected (Fig 2, bottom). These results
suggest that an age-related, prothrombotic phenotype develops
spontaneously in the lungs and liver of TM/
chimeric mice.
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| Fig 2.
Quantitation of 125I-labeled cross-linked
fibrin in TM/ chimeric and wild-type mice.
125I-labeled fibrinogen was intravenously infused into mice
and urea-insoluble extracts were prepared. 125I-labeled
urea-insoluble counts were quantitated and the cpm from five or six
animals from each genotype was averaged. Cross-linked fibrin from lung
and liver of 15- to 20-month-old mice (top) and 6- to 11-month-old mice
(bottom) is shown. ( ), TM+/+; ( ), high-percentage
chimeras; ( ), low-percentage chimeras.
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Contribution of TM/ ES cells to chimeric
mice.
The marked increase of urea-insoluble 125I-fibrin in the
TM/ chimeric mouse lung led us to analyze the
contribution of TM/ ES cells to lung tissue.
We used the differential expression of the glycerol phosphate
isoenzymes (GPI): the ES cell is homozygous for the "type a"
isoenzyme whereas the C57BL/6 host is homozygous for the "type b"
isoenzyme.18 The percentage of GPIa detected in whole lung
lysates is shown in Table 1. The GPIa
average value for the high-percentage chimeras is 33% (range, 26% to
40%) and the GPIa average value for the low-percentage chimeras is
20% (range, 18% to 23%). These results indicate the percentage of chimeric lung that is ES-cell derived. However, these values represent the ES-cell contribution to all cell types in the lung, not just the
endothelium.
To assess the TM/ ES-cell contribution to the
endothelium, TM cofactor activity was measured in these same lung
extracts. The average TM cofactor activity for the high-percentage
chimeras is 14 ng aPC/minute compared with 20 ng aPC/minute in
wild-type mice. This represents a 31% reduction from wild-type levels.
The average TM cofactor activity for the low-percentage chimeras is 17 ng aPC/minute or a 17% reduction from wild-type levels. These values
correspond closely to the values obtained with the GPI assay for
TM/ ES-cell contribution to lung and indicate
that, on average, from 17% to 31% of the endothelium in
TM/ chimeric mice do not express TM.
TM expression in chimeric lung.
TM is expressed at the lumenal surface of virtually all vascular
endothelium.6,24,25 This is shown in the murine lung using
immunocytochemistry to examine tissue sections from wild-type animals,
where we found all pulmonary endothelial cells stain positive for TM
(Fig 3a). TM immunolocalization
provides a means to distinguish TM-expressing and nonexpressing
endothelium, thereby identifying those endothelial cells presumably
derived from TM-null ES cells. When we examined the pulmonary
endothelium of TM/ chimeric mice, we found TM
localized in a mosaic pattern. Cross sections through single vessels
showed an intact endothelium with examples of either all endothelial
cells staining positive for TM (Fig 3b), endothelial cells staining
positive adjacent to endothelial cells negative for TM (Fig 3c), or
endothelial surfaces entirely negative for TM staining (Fig
3d). A similar TM expression pattern was observed in the
vasculature of liver and kidney in TM/
chimeric mice regardless of age (not shown). The mosaic expression pattern in TM/ chimeric mice is not
restricted to a subset of vessels; mosaicism is observed in large- and
small-caliber vessels in both the arterial and venous vasculature.
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| Fig 3.
TM immunolocalization in wild-type (Wt) and
TM/ chimeric (Ch) lung. PLP-fixed cryosections of
mouse lung were stained with an anti-TM rat MoAb and counterstained
with hematoxylin. TM is localized to the intimal surface of wild-type
mice (a) and some TM/ chimeric vessels (b). TM
mosaicism is shown in cross sections through blood vessels containing
patchy TM staining (c); arrows indicate endothelial nuclei confirming
the presence of an intact intima. Also observed are blood vessels
completely negative for TM localization (d); arrow indicates adjacent
blood vessel immunoreactive with anti-TM antibodies. Original
magnification × 630.
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Immunolocalization of cross-linked fibrin and neo in chimeric mouse
lung.
To map the intravascular fibrin deposits that were initially identified
by quantitating urea-insoluble 125I-labeled fibrin (Fig 2),
we immunolocalized acid-insoluble fibrin in murine lung.19
Only focal and low-level fibrin deposition was detected in lung
sections from wild-type mice (Fig 4a),
which was in contrast to the intense fibrin labeling in corresponding sections from TM/ chimeric mice (Fig 4b). In
general, the most remarkable fibrin localization occurred in small
muscular arteries, although fibrin deposition was also evident in large
and small veins. The localization of acid-insoluble fibrin in the 6- to
11-month-old TM/ chimeric mice was comparable
with that observed for wild-type mice (not shown).
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| Fig 4.
Localization of acid-washed fibrin/fibrinogen in the
murine pulmonary vasculature. PLP-fixed cryosections of mouse lung were
acid washed to solubilize fibrinogen and fibrin degradation products
before immunolocalization with an anti-fibrin/fibrinogen IgG and
counterstaining with hematoxylin. Wild-type (TM+/+)
mice between 15 and 20 months of age display little or no
immunoreactivity at the intimal surface (representative image shown in
a), whereas age-matched TM/ chimeric (Ch TM) mice
display a strong cross-linked fibrin immunolocalization at the lumenal
surface of blood vessels (b). Endothelial cell nuclei are indicated by
arrowheads. Original magnification × 400.
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We next determined whether fibrin is deposited on those regions of the
intimal surface deficient in TM. As stated, the neo gene is expressed
in endothelial cells derived from TM/ ES
cells. Therefore, endothelial cells immunoreactive with anti-neo antibodies lack TM. Colocalization studies in serial sections showed
that pulmonary blood vessels of TM/ chimeric
mice which displayed labeling for acid-insoluble fibrin (Fig 5, arrows) were also immunoreactive
for neo (Fig 5, arrows), whereas sites lacking fibrin deposition were
correspondingly deficient in neo expression (Fig 5, asterisk).
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| Fig 5.
Colocalization of cross-linked fibrin and neo in
TM/ chimeric mouse lung. PLP-fixed cryosections of
mouse lung were stained with anti-neo antibody and acid-washed serial
sections stained with the anti-fibrin/fibrinogen antibody before
counterstaining with hematoxylin. Shown is a representative image from
a high-percentage chimeric mouse in the 15- to 20-month age group.
Low-power views of serial cross sections show that neo (a) is present
in the same blood vessels that demonstrate immunoreactivity for
cross-linked fibrin (XL-Fibrin, b). Higher power magnification, c and
d, of the same images as shown in a and b. Arrows indicate blood
vessels. Original magnification in panels a and b, × 50; panels c and
d, × 125.
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Although we could colocalize neo and fibrin in sequential sections of a
single vessel, we observed that fibrin localization was not present in
all neo-positive vessels and was almost completely absent from the
microvasculature. To determine whether we could further classify the
types of vessels showing fibrin deposition, we measured the lumenal
areas of all blood vessels present within an H&E-stained lung section
as compared with the lumenal areas of blood vessels that stained
positive for neo and fibrin. The morphometric analysis of sequential
lung sections showed that neo-positive (ie,
TM/) vessels displayed a similar frequency
distribution as the H&E-stained sections
(Table 2): 88% of total blood vessels and
71% of neo-positive blood vessels have a lumenal area less than 100 µm2. This is in contrast to the frequency distribution of
the cross-linked, fibrin-stained vessels, because only 15% of
fibrin-stained vessels have a lumenal area less than 100 µm2. Although the possibility exists that the absence of
cross-linked fibrin in the microvasculature results from a fixation
artifact, we found that the localization of acid-insoluble fibrin in
another TM/ mouse, the TMPro/Pro
mouse, showed the presence of cross-linked fibrin deposited throughout the pulmonary vasculature in both microvessels and in larger caliber vessels.26 Collectively, the results of the
immunocytochemical and morphometric analyses suggest that fibrin
deposition is a single-vessel effect controlled not only by the extent
of the TM deficiency but also by vessel size and type.
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Table 2.
Comparison of Total Blood Vessels With Neo-Stained
and Cross-Linked, Fibrin-Stained Pulmonary Blood Vessels
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Fibrin deposition in response to hypoxia.
The results of the 125I-labeled fibrin analysis in the 6- to 11-month-old TM/ chimeric mice (Fig 2)
suggest that an increased thrombotic potential exists in these animals
but is manifested only in response to a second stimulus such as
increased age or environmental factors. To test this hypothesis, we
measured the endogenous -fibrin levels in 6- to 11-month-old
TM/ chimeric mice either exposed to hypoxia
(8% O2) or maintained at normoxia (21% O2).
The data obtained from pairing TM/ chimeric
mice by agouti coat coloring and challenging one animal from each pair
with hypoxia are shown in Fig 6. These data
show that exposure of TM/ chimeric mice to
hypoxia results in a fourfold increase in cross-linked -fibrin in
lung over paired controls (P < .005). Wild-type mice exposed
to hypoxia show a similar fourfold increase in cross-linked -fibrin
in lung compared with normoxic control mice. However, the ambient
levels of -fibrin are sixfold higher in the
TM/ chimeras as compared with wild-type mice
(ie, 0.11 µg fibrin/.1 g tissue v 0.50 µg fibrin/.1 g
tissue in wild-type mice under normoxic versus hypoxic conditions,
respectively, and 0.62 µg fibrin/.1 g tissue v 2.64 µg
fibrin/.1 g tissue in TM/ chimeras).
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| Fig 6.
Hypoxia induces fibrin deposition in
TM/ chimeric mice. (A) Western blot of -fibrin
from 6- to 11-month-old TM chimeras. Mice were paired according to
agouti coat coloring and one animal from each pair was maintained under
normoxic conditions (lanes 1, 2, and 3) while the other was exposed to
hypoxic conditions (lanes 4, 5, and 6). Western blot analysis of
urea-insoluble tissue samples was conducted using an MoAb against the
amino-terminus of B-fibrin. (B) Western blot of purified -fibrin
showing the standard curve containing: lane 1, 250 ng; lane 2, 150 ng;
lane 3, 100 ng; lane 4, 50 ng; and lane 5, 20 ng. (C) Quantitation of
cross-linked fibrin from Western blot shown in A. Values are expressed
as µg -fibrin per 100 mg of lung tissue.
|
|
| |
DISCUSSION |
Mutations in the TM/PC anticoagulant pathway are quite common in
humans. Alterations in PC function are often associated with an
increased risk of thrombosis, although many affected individuals remain
free of thrombotic complications.27 Conversely, if a population of patients displaying thrombotic complications are identified with heterozygous deficiencies in the TM/PC pathway, the
occurrence of a thrombotic phenotype in these patients is 10 to 25 times higher than in the general population. These findings suggest
that an inherited predisposition to thrombosis may result from an
additional mutation in the anticoagulant pathways. It has long been
postulated that a variant TM gene could be the second genetic defect
precipitating a thrombotic episode. This hypothesis has gone largely
untested because the endothelium is not readily sampled to test for
such defects.28 Therefore, we generated a murine model for
TM deficiency to directly test how alterations in TM function
contribute to the pathogenesis of thrombosis.
We replaced both alleles of the TM gene with the neo gene in ES cells
and used these homozygous mutant ES cells to generate chimeric mice
showing distinct TM deficiencies. We analyzed
TM/ chimeric mice for the presence of a
thrombotic propensity by quantitating urea-insoluble
125I-fibrin extracted from organs of mice infused with
125I-fibrinogen and by the immunolocalization of
acid-insoluble fibrin in these same organs. The amount of insoluble or
cross-linked fibrin corresponds to ES-cell contribution to whole lung,
as measured by the isoenzymes of glycerol phosphate and the
thrombin-dependent activation of PC. We found that the chimeric mice
containing the highest contribution from the TM-null ES cells (ie,
high-percentage chimeras containing up to a 40% relative contribution
from ES cells, shown in Table 1) are those animals expressing both the lowest TM levels and a significant increase in
125I-labeled, urea-insoluble fibrin. This thrombotic
phenotype is age dependent: high-percentage chimeras between 15 and 20 months show a significant increase in urea-insoluble fibrin in lung and liver. The high-percentage chimeras between 6 and 11 months have not
yet developed this phenotype, whereas wild-type mice and low-percentage TM chimeras do not show increased urea-insoluble
125I-fibrin, regardless of age. These results are
consistent with spontaneous intravascular fibrin formation in
TM/ mice and suggest that over a prolonged
time period a TM deficiency alters the hemostatic mechanism in favor of
the procoagulant response.
The results of our investigation reveal several factors that contribute
to the development of a prothrombotic phenotype and provide insight
into the physical characteristics of vascular lesion formation. Our
most surprising finding is that the actual physical size of the
TM/ area or procoagulant foci limits fibrin
formation and deposition. With few exceptions, it is those
TM/ vessels showing a lumenal area in excess
of 100 µm2 that also show cross-linked fibrin deposition.
Moreover, these focal gaps in anticoagulant function are autonomous of
hemostatic function in general because animals with the highest TM
deficiency show no manifestation of thrombotic disease other than these
localized, intravascular fibrin deposits. Using chimeric technology, we
can now test whether this single-vessel, quantitative trait is also generated from other mutations in the coagulation mechanism.
Our results indicate that coagulation activation and thrombin
generation occur normally in mice and that TM functions to counter the
procoagulant effects of thrombin. As stated, the increased urea-insoluble 125I-fibrin observed in
TM/ chimeric mice results from focal gaps in
TM expression. However, the morphometric analysis of sequential lung
sections localizing cross-linked fibrin as compared with neo shows that
fibrin deposits are disproportionately absent from the
TM/ microvessels. Although the
mechanism remains unclear, one possibility is that the TM pathway (ie,
either TM and/or aPC) functioning on wild-type endothelium may
be sufficient to neutralize thrombin generated at the lumenal surface
of neighboring TM/ cells and this ability is
lost when the distance between TM-expressing and nonexpressing cells
becomes too great. Whether fibrin deposition is the direct result of
decreased TM or the presumed decrease in aPC generation remains
unknown, in part because of the difficulty in quantifying local aPC.
Alternatively, the other natural anticoagulant pathways (ie, the
heparin-antithrombin III mechanism, the vasoactive substances
prostacyclin and nitric oxide, and the fibrinolytic mechanism) may
compensate for the loss of TM function, thereby preventing fibrin
deposits in small TM/ regions.
A similar compensatory effect is observed when urea-insoluble
125I-fibrin levels are compared between the high- and
low-percentage chimeras. The increased urea-insoluble
125I-fibrin observed only in the high-percentage chimeras
suggests that a threshold exists for TM anticoagulant function.
Chimeric mice showing reductions from wild-type TM levels of
approximately 24% or more display an increased and localized
cross-linked fibrin, whereas mice showing reductions less than 24%
remain unaffected, even though these levels are decreased from
wild-type TM levels. This observation further supports the supposition
that either the TM/PC pathway functioning in wild-type endothelial
cells or the other natural anticoagulant pathways complement
TM/ regions. The phenotype observed in
TM/ chimeric mice may be analogous to the
results obtained with intravenous infusions into mice of neutralizing
anti-TM antibody, which blocks cell-surface-associated TM activity in
vivo and enhances the likelihood of a lethal thrombotic event after
injection of thrombin into the tail vein.29 Collectively,
these results suggest that thrombosis is a quantitative trait, ie, the
development of a thrombotic phenotype is correlated with the magnitude
of the reduction in TM functional levels.
A second contributing factor to the prothrombotic phenotype is
organ-specific variations in endothelial cell hemostatic function. The
evidence emerging from studies in transgenic and knockout mice suggests
that the coagulation mechanism is controlled by the local
microenvironment present in different organs and
tissues.30,31 Hemodynamic forces, cell-cell interactions,
soluble factors, and tissue-specific gene expression all contribute to
locally regulated hemostasis.
Indeed, comparison of the TM/ chimeric mouse
with another TM-mutant mouse, the TMPro/Pro mouse, shows
distinct qualitative and quantitative differences in their respective
hemostatic function. In our previous investigation26 we
characterized the thrombotic phenotype of the TMPro/Pro
mice, which carry a single targeted mutation in the TM coding region
that corresponds to a Glu to Pro substitution at position 387 in the
human gene. This point mutation results in expression of the intact TM
receptor in vivo, but PC activation is reduced by 95%. Therefore, the
TM/ chimeric mice display a more subtle
mutation (PC activation reduced by 31%) and the intensity of the
thrombotic phenotype is diminished as compared with the
TMPro/Pro mice. For example, ambient fibrin levels in the
lungs of TMPro/Pro mice are increased 30-fold over
wild-type mice, whereas fibrin levels in the lungs of the
TM/ chimeric mice are increased only 5-fold
over wild-type mice, as determined by quantitative Western blotting.
Also, TMPro/Pro mice exposed to hypoxia (8% O2
for 16 hours) show a 10-fold increase in fibrin over normoxic
TMPro/Pro mice as compared with only a 4-fold increase
observed in the TM/ chimeric mice (Fig 6). A
second difference was noted when comparing the results of the
immunocytochemical analysis of lung sections from the
TMPro/Pro mice, which revealed fibrin deposition in
all segments of the vascular system (ie, microvessels, arteries, and
veins), with the TM/ chimeric mice, which
revealed preferential fibrin deposition in larger vessels. TM mosaicism
most likely contributes to this vascular bed-specific fibrin deposition
observed only in the TM/ chimeric mice.
Finally, organ-specific differences in hemostatic function result from
the distinct TM mutations. The TM/ chimeric
mice, like the TMPro/Pro mice, display a dramatic increase
in fibrin deposition in lung, but the TMPro/Pro mice also
display a marked fibrin deposition in the spleen and heart, whereas the
TM/ chimeric mice show an increased fibrin
deposition in liver. Worth noting are the two different methods used to
quantify cross-linked fibrin present in the vital organs of mice.
Infusions of 125I-labeled fibrinogen and measurements of
cross-linked radiolabeled fibrin, used to examine
TM/ chimeric mice, reflect both fibrin
deposition and dissolution over the 1-week time course. The
quantitative Western blot for fibrin -chains measures steady-state
fibrin deposition. However, we do detect significant increased fibrin
in the lungs of TM/ chimeric mice using both
techniques, suggesting that the organ-specific differences in TM
function are not solely caused by methodology. A second possibility
contributing to organ-specific differences in fibrin deposition between
the two TM/ mice is the age at which animals
were analyzed. The TM/ chimeric mice were
examined both at 6 to 11 months and at 16 to 20 months, whereas the
TMPro/Pro mice were all less than 1 year old. However,
knockout mice for the fibrinolytic proteins, urokinase, and tissue
plasminogen activator also display organ-specific differences in fibrin
deposition as measured by the quantitative Western blot
technique,26 indicating that organ-specific variations in
endothelial cell hemostatic function exist and are not restricted to
the TM/PC pathway.
The major similarity between the TM/ chimeric
and TMPro/Pro mice is the increased fibrin deposition found
in lungs. This may reflect the preferential expression of TM in normal
mouse lung as compared with other highly vascularized
organs24 and suggests that in the lung the anticoagulant
system may be more susceptible to alterations in the hemostatic
mechanism and therefore more susceptible to thrombosis.
Comparison of the TM/ chimeric mice with mice
heterozygous for the TM mutation (TM+/) is also
instructive. TM+/ mice have a 50% reduction in
functional TM levels and protein C cofactor activity12 as
compared with the 31% reduction in cofactor activity in
TM/ chimeras. However,
TM+/ mice show a uniform reduction in TM throughout
the vasculature in contrast to the mosaic pattern found in
TM/ chimeras. We do not detect an increase in
fibrin deposition/dissolution (as determined by
125I-fibrinogen infusions) in TM+/ mice
regardless of age (results not shown); this is in contrast to the
4-fold increase in fibrin deposition/dissolution in the TM chimeric
mice (Fig 2). The TM+/ mice have a 3-fold increase
in steady-state fibrin deposition26 as compared with the
5-fold increase over wild-type levels observed in
TM/ chimeras. Therefore, the
TM+/ mice have slightly lower levels of cofactor
activity, but display a less pronounced phenotype. These results
suggest that a focal TM deficiency is more likely to perturb hemostasis
than a comparable but uniform TM deficiency. However, when these two
mutant mice are exposed to hypoxia, it is the TM+/
mice that display a more striking phenotype. The
TM+/ mice show a 10-fold increase in steady-state
fibrin levels26 as compared with a 4-fold increase in TM
chimeras (Fig 6). These results suggest that exposure to a systemic
thrombogenic challenge causes a more marked response in animals with a
greater overall reduction in TM cofactor activity.
The third key element of the prothrombotic phenotype is age. The data
support the existing hypothesis that age-related alterations in the
coagulation mechanism predispose individuals harboring inherited,
silent mutations in the anticoagulant mechanism to thrombosis. Indeed,
the onset of thrombotic disease in humans has long been associated with
advancing age. In addition to increased circulating levels of
coagulation factors, the results of studies indicate significant
increases in coagulation enzyme activation and fibrinolytic enzyme
activity are age related.32,33 This documentation of an
age-dependent increase in coagulation system activity has led
investigators to propose that an increase in circulating clotting
factor activity may be a marker for predicting a "prethrombotic
state". However, the results of a recent study have shown that in a
group of healthy centenarians (25 individuals between the ages of 100 and 102 years), both coagulation system activation and fibrinolytic
activity were markedly increased over younger healthy controls, yet
these individuals were free of detectable thrombosis.34
These results suggest that increased coagulation system activity
associated with advanced age may be necessary but is not sufficient to
induce a thrombotic state.
Age-dependent increases in coagulation system activity are not
restricted to humans. Age-specific increases in the clotting factor,
factor IX, have also been documented in C57BL/6 mice. Factor IX mRNA
and cofactor activity are significantly upregulated in 28-month-old
mice as compared with 2-month-old mice.35 Thus, the
combined effects of increased coagulation system activity in aging
animals and the prolonged imbalance in the TM anticoagulant pathway are
sufficient to induce the thrombotic state.
Finally, oxygen deprivation induces a thrombotic phenotype, but the
mechanisms by which hypoxia-induced pulmonary thrombosis occurs are
unknown. Results of several recent studies in mice have defined events
leading up to coagulation activation in response to hypoxia that may
effect TM anticoagulant function. Within the first 2 to 4 hours after
exposure to hypoxia and before the onset of fibrin deposition an
increase in endothelial cell synthesized interleukin-6 is
noted.36 Increased cytokine levels are closely followed by
the appearance of pulmonary mononuclear phagocytes expressing increased
tissue factor and within 8 hours fibrin formation is
detected.37 Although the precise role the anticoagulant
pathway plays in contributing to these early events has not been
directly addressed, we noted that TM protein levels in lung remain
unchanged after mice were exposed to hypoxia for 24 hours.26 In the present study, we found that oxygen
deprivation induces a thrombotic phenotype in both wild-type and in
TM/ mice as expected. However, the ambient
levels of steady-state fibrin are increased fivefold to sixfold in the
6- to 11-month-old TM/ mice as compared with
wild-type mice, and fibrin formation in response to hypoxia is also
increased fivefold to sixfold in TM/ chimeras
compared with wild-type mice. These data suggest that the TM pathway
mediates the intensity of the thrombotic event and lends further
support to the hypothesis that a threshold for anticoagulant function
exists and is required for controlling hemostasis.
Extensive studies have shown the critical role that both heritable and
acquired traits play in promoting intravascular thrombosis. Our data
confirm and extend these previous findings and demonstrate that the
extent of the TM deficiencies, the physical dimensions of the
TM/ regions, and the stresses associated with
advancing age or environmental factors (ie, oxygen deprivation)
accelerate the onset and severity of a thrombotic event. In conclusion,
our findings support the novel concept that thrombosis resulting from a
TM deficiency is a single-vessel, quantitative trait.
| |
ACKNOWLEDGMENT |
We thank Dr D.L. Rosene for the use of the BioQuant Imaging System and
Dr E. Edelman for the use of an imaging work station.
| |
FOOTNOTES |
Submitted June 17, 1998;
accepted August 4, 1998.
Supported by National Institutes of Health Grants No. RO1 HL53396, PO1
41484 (R.D.R.), and RO1 HL60579 (W.W.H.). A.M.H. was the recipient of
an American Heart Association Fellowship Award from the Massachusetts
Affiliate.
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.
Address reprint requests to Aileen M. Healy, PhD, The Pulmonary Center,
Boston University School of Medicine, 80 E Concord St, Boston, MA
02118; e-mail: ahealy{at}bupula.bu.edu.
| |
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Expression of Thrombomodulin and Consequences of Thrombomodulin Deficiency during Healing of Cutaneous Wounds
Am. J. Pathol.,
November 1, 1999;
155(5):
1569 - 1575.
[Abstract]
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S.-F. Yan, N. Mackman, W. Kisiel, D. M. Stern, and D. J. Pinsky
Hypoxia/Hypoxemia-Induced Activation of the Procoagulant Pathways and the Pathogenesis of Ischemia-Associated Thrombosis
Arterioscler Thromb Vasc Biol,
September 1, 1999;
19(9):
2029 - 2035.
[Abstract]
[Full Text]
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Abstract
Introduction