© 1998 American Heart Association, Inc.
Original Contributions |
Expression of Thrombomodulin in Atherosclerotic Lesions and Mitogenic Activity of Recombinant Thrombomodulin in Vascular Smooth Muscle Cells
From the Third Department of Internal Medicine, Faculty of Medicine, Fukui Medical University, Fukui, Japan (G.T., K.O., S.K., E.O., S.T., I.M.); the Department of Pathology, Osaka Medical College, Osaka, Japan (Y.O.); and the Department of Public Health, Showa College of Pharmaceutical Sciences, Tokyo, Japan (H.I.).
Correspondence to Koji Oida, MD, Third Department of Internal Medicine, Fukui Medical University, Matsuoka-cho, Fukui 910-11, Japan. E-mail kojio{at}fmsrsa.fukui-med.ac.jp
 |
Abstract |
|---|
 Top Abstract IntroductionMethodsResultsDiscussionReferences |
|---|
Abstract—Thrombomodulin (TM), a thrombin receptor protein found on the endothelial cell surface, contains 6 tandem epidermal growth factor (EGF)–like structures. Recombinant human TM peptide containing these 6 EGF-like domains (rTME1–6) exhibits mitogenic activity in Swiss 3T3 cells. We examined the localization of TM in atherosclerotic lesions and the effects of rTME1–6 on the growth of cultured rat vascular smooth muscle cells (SMCs). Immunohistochemical analysis demonstrated that TM antigen was localized on monocytes, macrophages, and vascular SMCs. In cultured vascular SMCs, rTME1–6 accelerated [3H]thymidine uptake into DNA in a dose-dependent manner up to 3.4 times the control level. This mitogenic activity was abolished by addition of polyclonal anti-human TM antibody. The rTME1–6–induced mitogenesis was enhanced by EGF. However, a neutralizing monoclonal antibody against the EGF receptor (monoclonal antibody 225) did not inhibit the mitogenic activity of rTME1–6. Calphostin C, a specific protein kinase C inhibitor, and lavendustin-A, an inhibitor of EGF receptor–specific protein tyrosine kinase, inhibited the mitogenic activities of both rTME1–6 and EGF. Finally, rTME1–6 treatment increased the level of phosphorylated mitogen-activated protein kinase in SMCs. Together, these results suggest that TM expression in atherosclerotic lesions may be associated with promotion of atherosclerosis through its mitogenic activity in vascular SMCs.
Key Words: atherosclerosis • thrombomodulin • smooth muscle cells
|
Introduction |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Thrombomodulin (TM) is a high-affinity receptor for thrombin on the endothelial cell surface.1 By forming a complex with thrombin, TM alters the procoagulant activity of thrombin and acts as a cofactor in thrombin-catalyzed activation of protein C.1 Activated protein C inactivates coagulation factors Va and VIIIa, resulting in reduced thrombin formation. Thus, the thrombin-TM pathway is 1 of the major physiological antithrombotic mechanisms in endothelial cells, which makes TM an important regulator involved in maintaining the fluidity of circulating blood. TM consists of 5 distinct domains: the NH2-terminal domain, epidermal growth factor (EGF)–like domain, serine/threonine-rich region, transmembrane domain, and cytoplasmic tail.2 The EGF-like domain consists of 6 tandem EGF-like motifs that are homologous to domain III of the human EGF precursor.3 These EGF-like structures are associated with the cofactor activity of TM during thrombin-dependent protein C activation.4 5 Soluble TM fragments containing the EGF-like domain of native cellular TM6 have been detected in circulating plasma.7 These TM fragments likely are produced by proteolytic cleavage from cellular TM after endothelial cell damage.8
EGF is a mitogen that enhances the proliferation of various types of cells. Its mitogenic activity is mediated by EGF receptors located on the cell surface; these receptors also relay the biological signals of EGF-like growth factors.9 Recently, Hamada et al10 reported that recombinant TM peptide containing the 6 EGF-like domains (rTME1–6) has mitogenic activity in Swiss 3T3 fibroblast cells. However, this mitogenic activity may be mediated by a binding site other than the EGF receptor. To date, however, no studies have investigated whether TM may also have mitogenic activity in cells of the vascular system.
Proliferation of intimal smooth muscle cells (SMCs) is an important contributor to the progression of atherosclerosis.11 Vascular SMC proliferation is controlled by various cytokines and growth factors, depending on the location of the cells in the arterial wall. In the current study, we investigated whether TM binds to vascular cells in atherosclerotic lesions and how rTME1–6 affects the growth of cultured vascular SMCs.
|
Methods |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Materials
Human rTME1–6 was prepared as described previously.10 Human recombinant EGF was purchased from Pepro Teck. Platelet-derived growth factor (PDGF) AB and insulin were purchased from Sigma Chemical Company. Calphostin C12 13 was purchased from Wako. Lavendustin-A13 14 and tyrphostin AG37015 were purchased from Biomol Research Laboratories. A neutralizing monoclonal antibody against the EGF receptor (mAb 225)13 16 was purchased from Research Biochemicals International.
Preparation of Human and Rabbit Aortas
Human aortas were obtained from 6 autopsied cases (3 cases of
lung carcinoma and 1 case each of pancreatic carcinoma, liver
cirrhosis, and diabetes mellitus). Patients included 5 males and 1
female and ranged in age from 52 to 72 years. Rabbit aortas were
obtained from animals (Clea Japan) fed a 1% cholesterol
chow (Oriental Yeast Company Ltd). for 2 months.
Atheromatous plaques of human aortas and fatty streaks
of rabbit aortas were fixed with 10% formalin and embedded in paraffin
for histological and immunohistochemical
examination.
Immunohistochemical Analysis of TM Localization
Polyclonal rabbit anti-human TM antibody was prepared as
described previously.7 Sheep anti-rabbit TM
antibody was prepared by immunizing a sheep with rabbit
TM.17 Immunohistochemical staining was performed
as described by Hsu et al.18 Before staining,
paraffin sections of human and rabbit aortas were digested with 0.1%
trypsin and 10 mmol/L PBS for 30 minutes at 37°C. The human and
rabbit aorta sections were then incubated with 10 µg/mL anti-human TM
antibody and 13 µg/mL anti-rabbit TM antibody, respectively, for 2
hours. Rabbit and sheep preimmune sera served as negative controls.
Antibody-treated sections were washed 3 times in PBS buffer and then
incubated for 60 minutes each in biotinylated secondary antibody and
avidin-conjugated peroxidase (Vector Laboratories); sections were
washed between each step. After the final series of washes, peroxidase
was developed using 0.2% diaminobenzidine and 50 mmol/L Tris-HCl,
pH 7.6, supplemented with 0.3% hydrogen peroxide to yield a brown
reaction product. The reaction was stopped by washing with water,
and the tissues were counterstained with hematoxylin.
In human sections, intimal monocytes and macrophages were
detected immunohistochemically using the monoclonal mouse anti-human
antibody HAM56 (Dako).19 SMCs were identified
using a monoclonal mouse anti-human 
-SM actin antibody
(Sigma).20
Isolation and Culture of Rat Arterial SMCs
Arterial medial SMCs obtained from the thoracic
aorta of male Wistar rats (2 to 3 months old) were isolated by the
outgrowth of explants as described previously.21
In brief, specimens were dissected and cut into small pieces. Explants
were grown in Dulbecco's modified Eagle's medium (DMEM) containing
10% heat-inactivated FBS, penicillin (100 U/mL), and
streptomycin (70 µmol/L) at 37°C in a humidified atmosphere of
5% CO2 and 95% air. Cultured SMCs were used for
the following experiments after repeated (5 to 8) passages.
Cell Proliferation
Rat SMCs were subcultured in 6-well plates at a density of
104 cells per well and incubated for 24 hours at
37°C. The culture medium was removed, and adherent cells were exposed
to 1 nmol/L rTME1–6 in DMEM and 1% FBS. The medium was
replaced every 2 days. Cells were harvested every other day using
0.25% trypsin and 0.02% EDTA, and the number of cells was counted
with a hemocytometer.
DNA Synthesis
Rat SMCs were subcultured in 24-well microplates and grown
almost to confluence to avoid bias from contact inhibition. Cells were
then made quiescent by incubation in serum-free DMEM for 48 hours.
Quiescent cells were exposed to several concentrations of
rTME1–6 peptide and various mitogens for different times
at 37°C and labeled with [3H]thymidine (1
µCi/well) for the last 8 hours of incubation. To investigate the
mechanism of cell growth regulation, several kinase
inhibitors (calphostin C, lavendustin-A, and tyrphostin
AG370) were added at concentrations that inhibited their signaling
pathways but did not affect the basal growth rate of SMCs. The culture
medium was removed, and excess [3H]thymidine
was recovered by washing 4 times with ice-cold PBS and precipitating
with trichloroacetic acid. Cells were detached with 1N NaOH and
collected in a tube. The radioactivity of the collected cells was
determined using a liquid scintillation counter (Aloka). The measured
radioactivity was correlated with the protein concentration, which was
measured by using Lowry's method.
Determination of Mitogen-Activated Protein Kinase Activity
in Cell Extracts
Mitogen-activated protein kinase (MAPK) activity in cell
extracts was measured using a p42/p44 MAPK kit (Amersham).
Growth-arrested rat SMCs were stimulated for 10 minutes at 37°C with
rTME1–6 peptide or EGF and then lysed in 10 mmol/L
Tris, 150 mmol/L NaCl, 2 mmol/L EGTA, 2 mmol/L DTT,
1 mmol/L orthovanadate, 1 mmol/L PMSF, and 1.5 µmol/L
aprotinin, pH 7.4. The lysate was cleared by
centrifugation at 25 000g for 20 minutes at
4°C. MAPK activity was measured by using 10 µL of supernatant and
10 µL of substrate buffer containing the
phosphorylation sequence PLS/TP22
and HEPES buffer containing Mg2+, pH 7.4. The
reaction was terminated after 30 minutes by adding 50 µmol/L
[
-32P]ATP (20 µCi/mmol). MAPK catalyzed
the transfer of the -phosphate group of ATP to a peptide highly
selective for p42/p44 MAPK. The phosphorylated peptide
was separated by spotting onto binding paper. After the paper was
washed with 75 mmol/L orthophosphoric acid and water, the extent
of phosphorylation was determined by liquid
scintillation counting.
MAPK Activity Assay by Western Blot Analysis
MAPK activity also was assessed by Western blot analysis
using the PhosphoPlus MAPK antibody kit (New England BioLabs). Rat SMCs
stimulated for 10 minutes with rTME1–6 peptide or EGF were
lysed in 62.5 mmol/L Tris-HCl, pH 6.8, 2% (wt/vol) SDS, 10%
glycerol, 50 mmol/L DTT, and 0.1% (wt/vol) bromphenol blue. The
lysate was sonicated for 15 seconds, heated at 100°C for 5 minutes,
and centrifuged at 8000g for 5 minutes at 4°C. The
supernatant was subjected to SDS–polyacrylamide gel
electrophoresis, and the resolved proteins were electrophoretically
transferred onto nitrocellulose membranes. The blots were probed with a
rabbit anti-phosphotyrosine MAPK antibody to quantify activated
MAPK or with a rabbit
phosphorylation-state–independent MAPK antibody as a
control. Western blot analysis was performed as described
previously using the primary antibodies at 1/1000
dilutions.23 The immunoreactive proteins were
visualized by using enhanced chemiluminescence.
Statistical Analyses
All results are expressed as mean±SD. Statistical significance
was determined with Student's t test and 1-way ANOVA as
appropriate. The Stat View statistical program (Abacus Concepts) was
used. Values of P<0.05 were considered statistically
significant.
|
Results |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Localization of TM in Human and Rabbit Atherosclerotic Lesions
The presence of TM was determined immunohistochemically in atherosclerotic lesions of human aortas obtained from autopsied cases as well as in fatty streaks of rabbit aortas obtained from animals fed a 1% cholesterol diet. In the human aortas, TM was detected on monocytes and macrophages, which were identified using mAb HAM56, and intimal SMCs, which were identified using an antibody against
-SM actin (Figure 1C
). Medial SMCs also expressed TM
(Figure 1D). In the rabbit aortas, TM expression was demonstrated in
foamy macrophages, intima, and endothelial
cells (Figure 1F). In contrast to what was observed in human aortas,
medial SMCs in rabbit aortas expressed little TM. In both human (Figure 1E) and rabbit (Figure 1G) aortas, the specificity of the anti-TM
antibody was demonstrated on endothelial cells at the
vasa vasorum in the adventitia. An absorption test was performed to
evaluate the specificity of the polyclonal antibody for TM; incubation
of the polyclonal antibody with excess TM before
immunostaining led to a complete disappearance of the
staining reaction.
|
Effect of rTME1–6 on SMC Growth and DNA
Synthesis
To investigate the effects of rTME1–6 on SMC
proliferation, the cells were cultured in DMEM and 1% FBS in the
presence or absence of 1 nmol/L rTME1–6 for 14 days.
rTME1–6 significantly increased SMC proliferation 3.8-fold
(Figure 2). We also analyzed the
effects of rTME1–6 on
[3H]thymidine uptake into the DNA of rat SMCs.
After the cells were made quiescent by incubation in serum-free DMEM,
they were incubated with or without 1 nmol/L rTME1–6 for
various periods of time. [3H]Thymidine was
added for the last 8 hours. Maximal
[3H]thymidine incorporation into DNA, which
occurred 28 hours after addition of rTME1–6, was about
3.4-fold higher than that of cells cultured in serum-free medium
(Figure 3A). The dose dependence of
rTME1–6–mediated growth stimulation was analyzed
by culturing the cells in serum-free medium with increasing
rTME1–6 concentrations for 28 hours. These
analyses showed that [3H]thymidine
incorporation reached a plateau at about 3.5 times the control levels
at a concentration of 1 nmol/L rTME1–6 (Figure 3B). In
contrast, boiled rTME1–6 had no effect on either cell
proliferation or [3H]thymidine uptake into the
DNA of rat SMCs.
|
)
or without (
) 1 nmol/L rTME1–6. Medium was replaced
every 2 days. Cells were counted every other day. Data are mean±SD of
triplicate cultures. Student's t test was used to
evaluate differences between the 2 groups at various times.
*P<0.05, **P<0.01.
|
) 1 nmol/L rTME1–6 for various times.
[3H]Thymidine was added 8 hours before the indicated
times. B, Quiescent cells were exposed to various concentrations of
rTME1–6 for 28 hours (
Effect of rTME1–6 and Various Growth Factors on
[3H]Thymidine Uptake Into SMC DNA
rTME1–6, insulin, EGF, and PDGF each increased
[3H]thymidine uptake in a dose-dependent
manner. At the saturating concentration of each growth factor,
rTME1–6 increased [3H]thymidine
incorporation by 3.4-fold; insulin, 3.5-fold; EGF, 4.5-fold; and PDGF,
5.1-fold (Table 1). We then evaluated the
effect of coculturing rTME1–6 with each of these growth
factors on [3H]thymidine uptake.
rTME1–6 stimulated [3H]thymidine
uptake into the DNA of rat SMCs even at an EGF concentration of 1.6
nmol/L, which had a maximal effect on uptake (Table 1). Incubation with
a 100-fold molar excess of polyclonal rabbit anti-human TM antibody
completely inhibited the rTME1–6–induced increase in
[3H]thymidine uptake but had no effect on the
increase in uptake induced by EGF or PDGF (Figure 4).
|
|
Effect of Neutralizing Anti-EGF Receptor Antibody on
rTME1–6–Induced Mitogenic Activity
The mitogenic activity of rTME1–6 in
Swiss 3T3 cells has been shown to be mediated by binding of the peptide
to a specific site different from the EGF receptor. To determine
whether this also was true in rat SMCs, we examined the effect of mAb
225 on rTME1–6–induced mitogenic activity in
rat SMCs (Figure 5). mAb 225 is directed
against the extracellular domain of the human EGF receptor and can
inhibit ligand binding of the EGF family to both human and rat EGF
receptors.13 16 To determine its effects,
quiescent cells were exposed to rTME1–6 (1 nmol/L) or EGF
(1.6 nmol/L) in the absence or presence of mAb 225 (200 nmol/L) for 28
hours at 37°C. The antibody did not significantly inhibit the
rTME1–6–induced increase in
[3H]thymidine uptake.
|
Effects of Calphostin C, Lavendustin-A, and Tyrphostin AG370 on
rTME1–6–Induced DNA Synthesis
To investigate the mitogenic signaling pathway
underlying the activity of rTME1–6, we analyzed
the effects of calphostin C (a protein kinase C inhibitor),
lavendustin-A (a protein tyrosine kinase inhibitor specific
for the EGF receptor), and tyrphostin AG370 (a tyrosine kinase
inhibitor specific for the PDGF receptor) on
mitogenic activity in SMCs (Figure 6). Quiescent cells were treated with
rTME1–6, EGF, PDGF, or insulin in the presence or
absence of calphostin C (0.1 µmol/L), lavendustin-A
(1 µmol/L), or tyrphostin AG370 (50 µmol/L) for 28 hours
at 37°C, with [3H]thymidine (1 µCi/well)
added for the last 8 hours. The concentrations of inhibitor
used did not affect basal [3H]thymidine
uptake.
|
Calphostin C treatment reduced the increase in
[3H]thymidine uptake induced by
rTME1–6 or EGF to 60±4% and 55±5%, respectively, of
the control value (Figure 6A
). In contrast, the increase in
[3H]thymidine uptake caused by PDGF and insulin
was not significantly affected. Lavendustin-A also significantly
inhibited the increase in [3H]thymidine uptake
caused by rTME1–6 (61±4% of the control value) and EGF
(68±7% of the control value) but not by PDGF and insulin (Figure 6B).
Finally, tyrphostin AG370 significantly inhibited the
mitogenic activity of PDGF (35±5% of the control value)
but not of rTME1–6 and EGF (Figure 6C).
MAPK Activity in Rat SMCs Treated With
rTME1–6
To examine whether MAPK was involved in
rTME1–6–induced mitogenic activity, MAPK
activity was determined by 2 different methods, an enzyme bioassay and
immunological detection of activated MAPK. In the enzyme
bioassay, MAPK activity is calculated from the extent to which
[32P] is transferred from the -phosphate of
ATP to a peptide that is highly selective for p42/p44 MAPK. MAPK
activity was stimulated by both rTME1–6 (275±22% of the
control value) and EGF (248±14% of the control value). This effect
was concentration dependent (Table 2).
|
For immunological analysis, both total and active MAPK were
visualized by Western blotting. Immunoblots of the lysates
of cells treated with rTME1–6 or EGF and probed with
anti-phosphotyrosine MAPK antibody showed higher levels of
activated phosphotyrosine MAPK than did control cells. However,
the levels of total MAPK in the cell lysates did not differ among
rTME1–6–treated, EGF-treated, and control cells (Figure 7).
|
|
Discussion |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
TM is widely distributed in the endothelium of human arteries and veins, lymphatic vessels, syncytiotrophoblasts,24 synovial lining cells,25 monocytes,26 platelets,27 and neutrophils.28 It acts as a cofactor during thrombin-dependent protein C activation. The anticoagulant activity of TM depends on the local cell-membrane status.29 30 TM serves as a regulator in maintaining the fluidity of circulating blood; accordingly, TM downregulation on damaged endothelial cells potentiates the prothrombotic properties of the vascular wall, promotes coagulation, and contributes to thrombosis.
The current study has demonstrated TM antigen expression in atherosclerotic lesions. TM was constantly expressed in monocytes and macrophages in the atherosclerotic lesions of human and rabbit aortas. In human aortas, TM expression also occurred in intimal and medial SMCs. In rabbit aortas, in contrast, no TM expression was detectable in medial SMCs. These differences in TM expression in the vascular SMCs of humans and rabbits may be attributable to species differences.
TM contains several regions that are homologous to EGF. Mature EGF, which is produced by proteolytic cleavage of the extracellular domain of an integral membrane precursor, is present in various body fluids, such as circulating plasma.31 EGF binds to a specific receptor on the plasma membrane of cells to initiate mitogenic activity. TM possesses EGF-like domains. A recent study found that the rTME1–6 peptide, which includes the EGF-like domains, has mitogenic activity in Swiss 3T3 fibroblasts.10 In the current study, we expanded this observation by demonstrating that rTME1–6 also has mitogenic activity in cultured vascular SMCs. It appears likely that a specific binding site for rTME1–6 that may be structurally homologous to the EGF receptor and that may be required for the mitogenic activity of the EGF-like TM domain exists. rTME1–6 further increased [3H]thymidine incorporation into the DNA of rat SMCs even at the concentration of EGF that showed a maximal effect on [3H]thymidine uptake. On the other hand, a neutralizing monoclonal anti-EGF receptor antibody inhibited the EGF-induced, but not the rTME1–6–induced, synthesis of DNA. These findings suggest that the binding site that mediates the mitogenic activity of rTME1–6 is not the EGF receptor.
These observations suggest that rTME1–6 binds to a site that differs from the EGF-binding domain. To clarify this explanation, we studied a ligand-binding assay using 125I-labeled rTME1–6. 125I–rTME1–6 (32 000 cpm/mg) was prepared by using the lactoperoxidase method. Quiescent rat SMCs were exposed to several concentrations of 125I–rTME1–6 with and without a 100-fold excess of cold rTME1–6 for 3 hours at 37°C. Specific binding of 125I–rTME1–6 to cultured SMCs was calculated as the difference between the radioactivity bound in the absence of cold rTME1–6 and that bound in its presence. Scatchard plot analysis of specific binding indicated a single class of binding site (data not shown).
To investigate the signaling pathway that underlies the mitogenic activity of rTME1–6, we analyzed the effects of 3 kinase inhibitors. These include calphostin C, the protein kinase C inhibitor; lavendustin-A, which inhibits a protein tyrosine kinase specific for the EGF receptor; and tyrphostin AG370, a tyrosine kinase inhibitor specific for the PDGF receptor.
Protein kinase C–mediated signaling is 1 of the mitogenic pathways affecting vascular SMCs. Leszczynski et al12 reported that rat vascular SMCs cease to proliferate and undergo apoptosis when they are exposed to calphostin C. Protein kinase C also may be involved in EGF-dependent mitogenic activity by activating phospholipase C and adenyl cyclase.32 33 In the current study, treatment with calphostin C significantly reduced the [3H]thymidine uptake induced by rTME1–6 and EGF, suggesting that the mitogenic activity of rTME1–6, as well as the mitogenic activity of EGF, involves protein kinase C activation.
Antiproliferative compounds called tyrphostins34 specifically inhibit the EGF receptor tyrosine kinase. These compounds also inhibit EGF receptor autophosphorylation and EGF-dependent tyrosine phosphorylation of endogenous substrates, such as phospholipase CII.35 Moreover, the intracellular effects of tyrphostins are quantitatively correlated with the inhibition of EGF-dependent cell proliferation.35 One tyrphostin, lavendustin-A, significantly reduces the mitogenic activity of EGF.13 14 Similarly, in the current study, lavendustin-A significantly reduced the uptake of [3H]thymidine by SMCs in response to both rTME1–6 and EGF.
Tyrphostin AG370 is a highly potent blocker of PDGF-induced mitogenesis. Bryckaert et al15 reported that tyrphostin AG370 inhibits PDGF-induced mitogenesis and phosphorylation without affecting receptor-mediated PDGF binding and internalization. We demonstrated in this study that in SMCs, tyrphostin AG370 significantly inhibited PDGF-induced DNA synthesis but had no effect on rTME1–6– or EGF-induced DNA synthesis. Together, these results suggest that protein kinase C signaling and lavendustin-A–sensitive tyrosine kinase signaling are involved in the rTME1–6–induced mitogenic activity on SMCs. Thus, rTME1–6 appears to act through a mechanism similar to that of EGF.
Phosphorylation of cellular protein is important in regulating cell growth and the responses to extracellular stimuli. It is well known that the response to mitogens binding to receptors with intrinsic tyrosine kinase activity involves the activation of MAPKs. In addition, MAPKs are activated by stimuli, such as cytokines; hormones that bind to G protein–coupled receptors; and physical forces, such as fluid shear stress and stretch.36 Consequently, MAPK appears to serve as a common signaling mechanism for a variety of extracellular stimuli involved in the regulation of cell growth and function. Kusuhara et al37 showed that oxidized LDL stimulates MAPK activity in cultured vascular SMCs and macrophages. Accordingly, MAPK may play a significant role in the pathways by which oxidized LDL contributes to altered cellular function associated with atherogenesis. Although the effects of rTME1–6 on the MAPK activity of macrophages have not been examined, rTME1–6–mediated stimulation of MAPK in vascular SMCs may contribute to atherogenesis via such a mechanism.
On endothelial cells, TM may serve as an anticoagulant.1 Oxidized LDL reduces TM transcription in cultured human endothelial cells. The resulting decrease in anticoagulant activity on the cell surface may contribute to the development of atherosclerosis.38 In monocytic THP-1 cells, however, oxidized LDL has the opposite effect and upregulates TM expression.39 TM expression in circulating monocytes may play a role in preventing clotting and inflammation,40 and the oxidized LDL–induced upregulation of TM in monocytes may compensate for TM downregulation in endothelial cells. In the current study, we demonstrated that rTME1–6 also has a mitogenic effect on vascular SMCs, which suggests a role for TM in atherogenesis. Thus, it appears that TM expression may have different biological effects in different cell types. The physiological roles of TM expression in vascular SMCs and macrophages in atherosclerotic lesions remain to be elucidated.
Soluble TM fragments containing the EGF-like domain of native
TM6 are present in circulating
plasma.7 In the current study, we demonstrated
that rTME1–6 concentrations of 1 nmol/L have a
mitogenic effect on vascular SMCs. The plasma levels of
soluble TM antigen are 
0.1 nmol/L to 0.4 nmol/L in healthy adults
but can exceed 1 nmol/L in patients with disorders such as diabetes
mellitus,41 disseminated intravascular
coagulation,42 and systemic lupus
erythematosus.43 To
contribute to the proliferation of vascular SMCs, the soluble TM
fragments in the circulating plasma must pass through the layer of
endothelial cells lining the blood vessels. We
hypothesize that the TM expressed by intimal SMCs and
macrophages exerts its mitogenic activity on
neighboring SMCs only after its release from dead cells. Alternatively,
native TM on the cell surface may have a mitogenic activity
comparable to that of the membrane-anchored heparin-binding EGF-like
growth factor (pro HB-EGF), which exhibits mitogenic
activity in vascular SMCs similar to mature
HB-EGF.44 HB-EGF, a member of the EGF family, is
expressed in macrophages, endothelial cells,
platelets, T cells, and vascular SMCs. Its expression is
particularly high in atherosclerotic lesions compared with normal
vessels.45 Several other growth factors that are
synthesized from membrane-anchored proteins are also biologically
active in their transmembrane forms. For example, the precursor of
transforming growth factor- stimulates EGF receptor
phosphorylation and
mitogenesis.46 This type of biological activity
has been termed "juxtacrine stimulation." Consequently,
membrane-anchored TM may act on intimal SMCs via a juxtacrine
mechanism.
To investigate the correlation of TM expression with proliferation of SMCs, we performed double immunohistochemical staining of proliferating cell nuclear antigen (PCNA) and TM on atherosclerotic lesions under several conditions. Double immunofluorescent staining of the atheromatous plaques showed only TM-positive vascular SMCs. There was no distribution of PCNA-positive cells in the intima. A few PCNA-positive spindle cells, which are negative for TM, were found in the adventitia but not in the intima. It is well known that there are very few PCNA-positive cells within human atherosclerotic lesions. On the other hand, it has almost been established that intimal SMCs have mitogenic activity. We consider it very difficult to detect PCNA-positive SMCs immunohistochemically, mostly because of the rapid cell cycle of the SMCs, which makes detection of PCNA-positive cells difficult.
Chemotaxis of medial SMCs to the intima and proliferation of intimal SMCs are important in atherogenesis.11 If TM shares biological functions with other members of the EGF family, it may contribute to SMC differentiation and chemotaxis. However, additional research on TM is required to elucidate its role in atherosclerosis.
|
Acknowledgments |
|---|
We thank Naoyo Yamaguchi for her excellent technical assistance.
Received July 29, 1997; accepted May 18, 1998.
|
References |
|---|
|
TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
1. Esmon CT. The role of protein C and thrombomodulin in the regulation of blood coagulation. J Biol Chem. 1989;264:4743–4746.
2. Suzuki K, Kusumoto S, Deyashiki Y, Nishioka J, Maruyama I, Zushi M, Kawahara S, Honda G, Yamamoto S, Horiguchi S. Structure and expression of human thrombomodulin, a thrombin receptor on endothelium acting as a cofactor for protein C activation. EMBO J. 1987;6:1891–1897.[Medline] [Order article via Infotrieve]
3. Wen D, Dittman WA, Ye RD, Deaven LL, Majerus PW, Sadler JE. Human thrombomodulin: complete cDNA sequence and chromosome localization of the gene. Biochemistry. 1987;26:4350–4357.[Medline] [Order article via Infotrieve]
4.
Zushi M, Gomi K, Yamamoto S, Maruyama I, Hayashi T,
Suzuki K. The last three consecutive epidermal growth factor–like
structures of human thrombomodulin comprise the minimum functional
domain for protein C–activating cofactor activity and anticoagulant
activity. J Biol Chem. 1989;264:10351–10353.
5.
Tsiang M, Lents SR, Sadler JE. Functional domains of
membrane-bound human thrombomodulin: EGF-like domains four to six and
the serine/threonine-rich domain are required for cofactor activity.
J Biol Chem. 1992;267:6164–6170.
6. Jackson DE, Tetaz TJ, Salem HH, Mitchell CA. Purification and characterization of two forms of soluble thrombomodulin from human urine. Eur J Biochem. 1994;221:1079–1087.[Medline] [Order article via Infotrieve]
7. Ishii H, Nakano M, Tsubouchi J, Ishikawa T, Uchiyama H, Hiraishi S, Tahara C, Miyajima Y, Kazama M. Establishment of enzyme immunoassay of human thrombomodulin in plasma and urine using monoclonal antibodies. Thromb Haemost. 1990;63:157–162.[Medline] [Order article via Infotrieve]
8. Ishii H, Uchiyama H, Kazama M. Soluble thrombomodulin antigen in conditioned medium is increased by damage of endothelial cells. Thromb Haemost. 1991;65:618–623.[Medline] [Order article via Infotrieve]
9. Cohen S. Epidermal growth factor. In Vitro Cell Dev Biol. 1987;23:239–246.[Medline] [Order article via Infotrieve]
10.
Hamada H, Ishii H, Sakyo K, Horie S, Nishiki K, Kazama
M. The epidermal growth factor–like domain of recombinant human
thrombomodulin exhibits mitogenic activity for Swiss 3T3
cells. Blood. 1995;86:225–233.
11. Ross R. The pathogenesis of atherosclerosis: a perspective for the 1990s. Nature. 1993;362:801–809.[Medline] [Order article via Infotrieve]
12. Leszczynski D, Zhao Y, Luokkamaki M, Foegh ML. Apoptosis of vascular smooth muscle cells: protein kinase C and oncoprotein Bcl-2 are involved in regulation of apoptosis in non-transformed rat vascular smooth muscle cells. Am J Pathol. 1994;145:1265–1270.[Abstract]
13.
Kornyei JL, Li X, Lei ZM, Rao CV. Analysis of
epidermal growth factor action in human myometrial smooth muscle cells.
J Endocrinol. 1995;146:261–270.
14.
Hsu CY, Persons PE, Spada AP, Bednar RA, Levitzki A,
Zilberstein A. Kinetic analysis of the inhibition of the
epidermal growth factor receptor tyrosine kinase by lavendustin-A and
its analogue. J Biol Chem. 1991;266:21105–21112.
15. Bryckaert MC, Eldor A, Fontenay M, Gazit A, Osherov N, Gilon C, Levitzki A, Tobelem G. Inhibition of platelet-derived growth factor–induced mitogenesis and tyrosine kinase activity in cultured bone marrow fibroblasts by tyrphostins. Exp Cell Res. 1992;199:255–261.[Medline] [Order article via Infotrieve]
16.
Fan Z, Mendelsohn J, Masui H, Kumar R. Regulation of
epidermal growth factor receptor in NIH3T3/HER14 cells by antireceptor
monoclonal antibodies. J Biol Chem. 1993;268:21073–21079.
17. Uchiyama H, Ohtani H, Hiraishi S, Horie S, Ishii H, Kazama M. Changes in plasma thrombomodulin antigen in rabbit developing endotoxin-induced disseminated intravascular coagulation and the effect of heparin. Thromb Res. 1992;65:593–604.[Medline] [Order article via Infotrieve]
18. Hsu SM, Raine L, Fanger H. The use of avidin-biotin-peroxidase complex (ABC) in immunoperoxidase technique: a comparison between ABC and unlabeled antibody (PAP) procedures. J Histochem Cytochem. 1981;29:577–580.[Abstract]
19. Grown A, Tsukada T, Ross R. Human atherosclerosis II: immunocytochemical analysis of the cellular composition of human atherosclerotic lesions. Am J Pathol. 1986;125:191–207.[Abstract]
20.
Skalli O, Ropraz P, Trzeciak A, Benzonana G, Gillesen
D, Gabbiani G. A monoclonal antibody against -smooth muscle
actin: a new probe for smooth muscle differentiation. J Cell
Biol. 1986;103:2787–2796.
21. Takahashi S, Oida K, Fujiwara R, Maeda H, Hayashi S, Takai H, Tamai T, Nakai T, Miyabo S. Effect of cilostazole, a cyclic AMP phosphodiesterase inhibitor, on the proliferation of rat aortic smooth muscle cells in culture. J Cardiovasc Pharmacol. 1992;20:900–906.[Medline] [Order article via Infotrieve]
22.
Clark LI, Sanghera JS, Pelech SL. Definition of a
consensus sequence for peptide substrate recognition by p44mpk, the
meiosis-activated myelin basic protein kinase. J
Biol Chem. 1991;266:15180–15184.
23.
Oida K, Takai H, Maeda H, Takahashi S, Shimada A,
Suzuki J, Tamai T, Nakai T, Miyabo S. Apolipoprotein (a) is present
in urine and its excretion is decreased in patients with renal failure.
Clin Chem. 1992;38:2244–2248.
24.
Maruyama I, Bell CE, Majerus PW. Thrombomodulin is
found on endothelium of arteries, veins, capillaries,
and lymphatics, and on syncytiotrophoblast of human placenta.
J Cell Biol. 1985;101:363–371.
25. Boffa MC, Burke B, Haudenschild CC. Preservation of thrombomodulin antigen on vascular and extravascular surfaces. J Histochem Cytochem. 1987;35:1267–1276.[Abstract]
26.
McCachren SS, Diggs J, Weinberg JB, Dittman WA.
Thrombomodulin expression by human blood monocytes and by human
synovial tissue lining macrophages. Blood. 1991;78:3128–3132.
27.
Suzuki K, Nishioka J, Hayashi T, Kosaka Y.
Functionally active thrombomodulin is present in human
platelets. J Biochem. 1988;104:628–632.
28.
Conway EM, Nowakowski B, Steiner MM. Human neutrophils
synthesize thrombomodulin that does not promote thrombin-dependent
protein C activation. Blood. 1992;80:1254–1263.
29.
Galvin JB, Kurosawa S, Moore KL, Esmon CT, Esmon NL.
Reconstitution of rabbit thrombomodulin into phospholipid vesicles.
J Biol Chem. 1987;262:2199–2205.
30. Horie S, Ishii H, Hara H, Kazama M. Enhancement of thrombin-thrombomodulin–catalyzed protein C activation by phosphatidylethanolamine containing unsaturated fatty acids: possible physiological significance of phosphatidylethanolamine in anticoagulant activity of thrombomodulin. Biochem J. 1994;301:683–691.
31. Jon T, Itoh M, Katsumi K, Yokoyama Y, Takeuchi T, Kato T, Wada W, Tanaka R. Physiological concentrations of human epidermal growth factor in biological fluids: use of a sensitive enzyme immunoassay. Clin Chim Acta. 1986;158:81–90.[Medline] [Order article via Infotrieve]
32.
Nishibe S, Wahl MI, Hernandez-Sotomayor SMT, Tonks NK,
Rhee SG, Carpenter G. Increase of catalytic activity of phospholipase
C-1 by tyrosine phosphorylation.
Science. 1990;250:1253–1255.
33.
Carpenter G, Cohen S. Epidermal growth factor.
J Biol Chem. 1990;265:7709–7712.
34.
Yaish P, Gazit A, Gilon C, Levitzki A. Blocking of
EGF-dependent cell proliferation by EGF receptor kinase
inhibitors. Science. 1988;242:933–935.
35.
Lyall RM, Zilberstein A, Gazit A, Gilon C, Levitzki A,
Schlessinger J. Tyrphostins inhibit epidermal growth factor
(EGF)-receptor tyrosine kinase activity in living cells and
EGF-stimulated cell proliferation. J Biol Chem. 1989;264:14503–14509.
36. Pelech SL, Sanghera JS. MAP kinases: charting the regulatory pathways. Science. 1993;257:1355–1356.
37.
Kusuhara M, Alan C, Angelina C, Bradford CB. Oxidized
LDL stimulates mitogen-activated protein kinase in smooth
muscle cells and macrophages. Arterioscler Thromb Vasc
Biol. 1997;17:141–148.
38.
Ishii H, Kizaki K, Horie S, Kazama M. Oxidized low
density lipoprotein reduces thrombomodulin transcription in cultured
human endothelial cells through degradation of the
lipoprotein in lysosomes. J Biol Chem. 1996;271:8458–8465.
39. Oida K, Tohda G, Ishii H, Horie S, Kohno M, Okada E, Suzuki J, Nakai T, Miyamori I. Effect of oxidized low density lipoprotein on thrombomodulin expression by THP-1 cells. Thromb Haemost. 1997;78:1228–1233.[Medline] [Order article via Infotrieve]
40. McCachren SS, Diggs J, Weinberg JB, Dittman WA. Thrombomodulin expression by human blood monocytes and by human synovial tissue lining macrophages. Blood. 1991;78:3128–3132.
41. Oida K, Takai H, Maeda H, Takahashi S, Tamai T, Nakai T, Miyabo S, Ishii H. Plasma thrombomodulin concentration in diabetes mellitus. Diabetes Res Clin Pract. 1990;10:193–196.[Medline] [Order article via Infotrieve]
42.
Takano S, Kimura S, Ohdama S, Aoki N. Plasma
thrombomodulin in health and diseases. Blood. 1990;76:2024–2029.
43. Kawakami M, Kitani A, Hara M, Harigai M, Suzuki K, Kawaguchi Y, Ishii H, Kazama M, Kawagoe M, Nakamura H. Plasma thrombomodulin and a 2-plasmin inhibitor-plasmin complex are elevated in active systemic lupus erythematosus. J Rheumatol. 1992;19:1704–1709.[Medline] [Order article via Infotrieve]
44.
Higashiyama S, Abraham JA, Klagsbrun M. Heparin-binding
EGF-like growth factor stimulation of smooth muscle cell migration:
dependence on interactions with cell surface heparin sulfate.
J Cell Biol. 1993;122:933–940.
45.
Higashiyama S, Abraham JA, Miller J, Fiddes JC,
Klagsbrun M. A heparin-binding growth factor secreted by
macrophage-like cells that is related to EGF.
Science. 1991;251:936–939.
46.
Anklesaria P, Teixido J, Laiho M, Pierce JH,
Greenberger JS, Massague J. Cell-cell adhesion mediated by binding
of membrane-anchored transforming growth factor to epidermal
growth factor receptors promotes cell proliferation. Proc Natl
Acad Sci U S A. 1990;87:3289–3293.
This article has been cited by other articles:
 |
|
 |
|
  I.-C. Lo, T.-M. Lin, L.-H. Chou, S.-L. Liu, L.-W. Wu, G.-Y. Shi, H.-L. Wu, and M. J. Jiang Ets-1 mediates platelet-derived growth factor-BB-induced thrombomodulin expression in human vascular smooth muscle cells Cardiovasc Res, March 1, 2009; 81(4): 771 - 779. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 K. Rabausch, E. Bretschneider, M. Sarbia, J. Meyer-Kirchrath, P. Censarek, R. Pape, J. W. Fischer, K. Schror, and A.-A. Weber Regulation of Thrombomodulin Expression in Human Vascular Smooth Muscle Cells by COX-2-Derived Prostaglandins Circ. Res., January 7, 2005; 96(1): e1 - e6. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. V. de Wouwer, D. Collen, and E. M. Conway Thrombomodulin-Protein C-EPCR System: Integrated to Regulate Coagulation and Inflammation Arterioscler. Thromb. Vasc. Biol., August 1, 2004; 24(8): 1374 - 1383. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 C. E. Hafer-Macko, F. M. Ivey, K. A. Gyure, J. D. Sorkin, and R. F. Macko Thrombomodulin Deficiency in Human Diabetic Nerve Microvasculature Diabetes, June 1, 2002; 51(6): 1957 - 1963. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Y. Kim, P. L. Walinsky, F. D. Kolodgie, C. Bian, J. L. Sperry, C. B. Deming, E. A. Peck, J. G. Shake, G. B. Ang, R. H. Sohn, et al. Early Loss of Thrombomodulin Expression Impairs Vein Graft Thromboresistance: Implications for Vein Graft Failure Circ. Res., February 8, 2002; 90(2): 205 - 212. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
H. Weiler, V. Lindner, B. Kerlin, B. H. Isermann, S. B. Hendrickson, B. C. Cooley, D. A. Meh, M. W. Mosesson, N. W. Shworak, M. J. Post, et al. Characterization of a Mouse Model for Thrombomodulin Deficiency Arterioscler. Thromb. Vasc. Biol., September 1, 2001; 21(9): 1531 - 1537. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. E. Freedman Thrombin, Thrombomodulin, and Extracellular Signal-Regulated Kinases Regulating Cellular Proliferation Circ. Res., April 13, 2001; 88(7): 651 - 653. [Full Text] [PDF]
|
|
|
|
|
|
L. Le Flem, L. Mennen, M.-L. Aubry, M. Aiach, P.-Y. Scarabin, J. Emmerich, and M. Alhenc-Gelas Thrombomodulin Promoter Mutations, Venous Thrombosis, and Varicose Veins Arterioscler. Thromb. Vasc. Biol., March 1, 2001; 21(3): 445 - 451. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Y. Kim, P. L. Walinsky, F. D. Kolodgie, C. Bian, J. L. Sperry, C. B. Deming, E. A. Peck, J. G. Shake, G. B. Ang, R. H. Sohn, et al. Early Loss of Thrombomodulin Expression Impairs Vein Graft Thromboresistance: Implications for Vein Graft Failure Circ. Res., February 8, 2002; 90(2): 205 - 212. [Abstract] [Full Text] [PDF]
|
|
| |||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||