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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 93 No. 9 (May 1), 1999:
pp. 2951-2958
Direct Evidence of Endothelial Injury in Acute Myocardial
Infarction and Unstable Angina by Demonstration of Circulating
Endothelial Cells
By
Murielle Mutin,
Isabelle Canavy,
Andrew Blann,
Michel Bory,
José Sampol, and
Françoise Dignat-George
From the Laboratoire d'Hématologie et d'Immunologie, U.F.R.
de Pharmacie, Marseille Cedex; Laboratoire d'Hématologie,
Hôpital de la Conception, Marseille Cedex; Service de cardiologie
A, centre hospitalier et universitaire de la Timone, Marseille Cedex,
France; and Haemostasis, Thrombosis and Vascular Biology Unit,
University Department of Medicine, City Hospital, Birmingham, UK.
 |
ABSTRACT |
Circulating endothelial cells (CECs) have been detected in
association with endothelial injury and therefore represent proof of
serious damage to the vascular tree. Our aim was to investigate, using
the technique of immunomagnetic separation, whether the pathological
events in unstable angina (UA) or acute myocardial infarction (AMI)
could cause desquamation of endothelial cells in circulating blood
compared with effort angina (EA) and noncoronary chest pain. A high CEC
count was found in AMI (median, 7.5 cells/mL; interquartile range, 4.1 to 43.5, P < .01 analysis of variance [ANOVA])
and UA (4.5; 0.75 to 13.25 cells/mL, P < .01) within 12 hours
after chest pain as compared with controls (0; 0 to 0 cells/mL) and
stable angina (0; 0 to 0 cells/mL). CEC levels in serial samples peaked
at 15.5 (2.7 to 39) cells/mL 18 to 24 hours after AMI (P < .05 repeated measures ANOVA), but fell steadily after UA.
Regardless of acute coronary events, the isolated cells displayed
morphologic and immunologic features of vascular endothelium. The CECs
were predominantly of macrovascular origin. They did not express the
activation markers intercellular adhesion molecule (ICAM)-1, vascular cell adhesion molecule
(VCAM)-1, and E-selectin, although some were positive for
tissue factor. CECs failed to exhibit characteristics of apoptosis
(TUNEL assay) excluding this event as a possible mechanism
of cell detachment. The presence of CECs provides direct evidence of
endothelial injury in AMI and UA, but not in stable angina, confirming
that these diseases have different etiopathogenic mechanisms.
© 1999 by The American Society of Hematology.
| |
INTRODUCTION |
ENDOTHELIAL INJURY represents a major
initiating step in the pathogenesis of atherosclerosis that can lead to
acute coronary syndromes such as acute myocardial infarction (AMI) and unstable angina (UA).1-3 These acute coronary pathologies
represent high-risk processes with severe life-threatening thrombotic
events. Various experimental models have been developed to improve our understanding of the importance of endothelial integrity and the pathogenic mechanisms of vascular alterations leading to these clinical
situations.4 Fuster et al5 have proposed a
classification of vascular injury that includes stages of increasing
severity. Type I consists of functional alterations of endothelial
cells without substantial morphologic changes. These alterations are followed by lipid accumulation, monocyte and platelet adhesion, and
smooth muscle cell proliferation resulting in plaque formation. Type I
injury can be followed by type II and type III injuries, defined as
endothelial denuding, and intimal injury with or without medial damage.
Finally, the process may be complicated by plaque ulceration, rupture
or erosion, and thrombus formation, which can trigger a heart attack or
sudden death.
Research in clinical settings has been hindered by the inaccessibility
of vascular endothelium in both healthy subjects and patients.
Circulating endothelial cells (CECs) may provide useful material for
the study of vascular injury. They have been detected in the blood in
diverse conditions such as coronary angioplasty, sickle cell disease,
thrombotic thrombocytopenic purpura, and infection with Rickettsia
conorii and cytomegalovirus.6-15 We therefore
hypothesized that AMI and UA could present such a stress to the
vascular system so that endothelial cells would be lost from the intima
and thus appear in the blood. Because preliminary studies suggested
that acute coronary syndromes could be associated with increased levels
of cells presenting endothelial morphologic features,16,17
we combined specific capture and immunological characterization to
investigate CECs in AMI and UA. Consequently, using a previously
developed immunomagnetic separation assay based on S-Endo 1 monoclonal
antibody (MoAb) directed against the endothelial antigen
CD146,10-12,18,19 we studied the number, origin, and surface phenotype of CECs. We also determined whether or not these CECs
were apoptotic and would be simply shed from the intima. We controlled
our AMI and UA specimens by obtaining blood from subjects with effort
angina (EA) and from subjects with noncoronary chest pain.
| |
MATERIALS AND METHODS |
Patient selection and diagnosis.
Subjects were recruited from among those admitted to the coronary care
unit at CHU Timone (Marseille, France). Local ethical committee
approval was obtained according to the Declaration of Helsinki, and
written informed consent was obtained from each subject. They were
categorized into four groups: the target groups AMI and UA and the two
control groups of EA and patients presenting with noncoronary chest
pain. Characteristics and risk factors of patients are listed in
Table 1. All of the patients selected for
the study did not undergo catheterization and percutaneous angioplasty
before and during the time of the study. For each patient included,
standard coronary angiography was performed after the completion of the
study, 8 days after admission for AMI, 5 days after for UA and
controls, and 24 hours after for EA. Coronary artery aspect was
analyzed according to the classification of Ambrose et
al.20 Diagnosis and coronary angiography characteristics of
patients are summarized in Tables 2 and
3.
Diagnosis of AMI was made within 24 hours after the onset of symptoms
according to established criteria of the World Health Organization. At
least two of the three following criteria were present: typical
sustained chest pain, electrocardiogram (ECG) Q wave, and raised peak
creatinine kinase (CPK). All patients showed raised CPK (peak median,
639 IU/mL; interquartile range [IQR], 307 to 988; 12 to
18 hours after admission) and coronary stenoses.
Diagnosis of UA was made within 12 hours after the onset of symptoms
with electrocardiographic changes on ST segment during angina pectoris
crisis. All patients presented creatinine kinase levels within normal
range and subsequently showed coronary stenoses.
Diagnosis of EA was performed by exercise testing with a bicycle
ergometer according to the protocol (30 W each for 3 minutes).21 Systemic arterial pressure and ECG were
continuously supervised. Exercise was stopped when angina pectoris or
ST segment depression appeared.
Diagnosis of control group was made within 12 hours after noncoronary
chest pain. Creatinine kinase was normal. Coronary angiography showed
no stenosis.
All patients from AMI, UA, and control groups received the same medical
treatment: unfragmented heparin (choay: 5,000 IU as a
bolus and 1,000 IU/h during 8 days adaptable in function of activated
partial thromboplastin time [aPTT]), aspirin (160 mg/d), and  blockers (atenolol: 100 mg/d intravenous during 48 hours then orally
during the hospitalization). Only four patients with AMI underwent an
intravenous thrombolysis within 1 to 6 hours delay (tissue plasminogen
activator [actilyse]), 15 mg as a bolus, 0.75 mg/kg over 30 minutes,
and 0.50 mg/kg over 60 minutes because the standard clinical procedure
followed in our hospital is angioplasty for AMI patients hospitalized
within the 6 hours after the onset of symptoms. We have excluded such
patients because in a previous report we have demonstrated that
coronary angioplasty induces the release of endothelial cells in the
circulation.11 Hence, AMI patients included in the present
study were those hospitalized later than 6 hours and who followed the
conventional pharmacologic treatment (unfragmented heparin, aspirin,
and blockers) and the four patients who underwent thrombolysis.
Blood collection.
For CEC quantitation, the first 2 mL of blood drawn were discarded to
avoid contamination by endothelial cells from the punctured vessel
wall. A total of 5 mL of blood was then collected into EDTA. For the
AMI, UA, and control groups, a first blood sample was collected on
arrival at hospital, within 24 hours of the onset of chest pain. Then,
three or four other samples were drawn at intervals, such as every 6 hours, for up to 42 hours. For the EA group, one sample was collected
before undertaking the exercise tolerance test, just after the chest
pain, and again 4 hours later.
Antibodies.
For immunocapture of CECs from whole blood, S-Endo 1 (Biocytex, Marseille, France), a MoAb raised against human umbilical vein endothelial cells (HUVEC) in our laboratory,11 was
used. This MoAb was selected because of its strong reactivity for
endothelial cells from all vascular beds and its negative reaction with
hematopoietic cells, mesothelial cells, or fibroblasts. This antibody
also reacts moderately with smooth muscle cells.19 For the
immunological characterization of the cells, we used the following
antibodies: a rabbit polyclonal antibody against human von Willebrand
factor (vWF, a kind gift from Y. Sultan, Laboratoire d'Hematologie,
Hopital de la Miletrie, Poitiers, France),22 a
murine MoAb against smooth muscle  -actin (IgG2a, clone 1A4; Sigma,
St Quentin-Fallavier, France); MoAbs against the adhesion molecules
intercellular adhesion molecule-1 (ICAM-1) (IgG1, clone F431C2/B7;
Biocytex, Marseille, France), vascular cell adhesion molecule-1
(VCAM-1) (IgG1, clone 1G11; Immunotech, Marseille, France), and
E-Selectin (IgG1, clone 1.2B6; Immunotech). Fluorescein isothiocyanate
(FITC) conjugated MoAb against tissue factor (TF) (IgG1, clone TF9;
Ortho Diagnostic System, Roissy, France) and CD36 (IgG1, clone FA6-152; Immunotech).
The antibodies used as controls were rabbit antiserum for the rabbit
polyclonal antibody (Sigma) and isotype-matched antibodies with
irrelevant binding specificities for the murine MoAbs (IgG1, clone
MOPC-21 and IgG2a, clone UPC-10, Sigma, FITC-conjugated IgG1, clone
679.1MC7, Immunotech).
Goat anti-rabbit and goat anti-mouse antibodies conjugated to
tetramethylrhodamine isothiocyanate (TRITC) (Immunotech) and to FITC
(Silenus, Eurobio, Les Ullis, France), respectively, were used to show
the unconjugated antibodies.
Preparation of immunomagnetic beads.
Monodispersed magnetizable beads (Dynabeads-M-450) were obtained from
Dynal A.S. (Oslo, Norway). They are 4.5-µm diameter polystyrene beads
with rat anti-mouse IgG1 covalently bound to the surface. They were
coated with S-Endo 1 MoAb as a second layer as previously
described.11
Immunomagnetic separation and counting of CECs.
Separation and quantitation of CECs were performed as previously
described.11 A total of 1 mL of whole blood diluted 1:4 in
phosphate-buffered saline (PBS)-0.1% bovine serum albumin-0.1% sodium
azide were mixed with 20 µL of S-Endo 1-coated beads and submitted to
gentle agitation for 30 minutes on a sample mixer (Robbins Scientific,
Bio Techgen, Les Ullis, France). Magnetic beads and rosetted cells were
separated from blood using the MPC6 concentrator (Dynal).
After two washes, the rosetted cells were resuspended in acridin orange
(10 µg/mL in PBS) (Sigma) for cell counting. Analysis was performed
under an optical fluorescence microscope ( exc = 490 nm)
using a Nageotte hemocytometer (CML, Nemours, France) for
quantitation. The criteria retained for the identification of CECs were first, rosettes bearing more than 10 beads
and a cell size in the range of 20 to 50 µm and second, cells with
less than 10 beads, but with a well-preserved and recognizable morphology (clear nucleus in a well delimited cytoplasm and a size,
which might correspond to endothelial cells). For aggregates, the
number of cells was deduced from the number of nuclei or from the
number of spherical rosetted features discriminated in the aggregate.
When cell count was higher than 5 cells/mL, the rest of the blood was
processed for immunomagnetic separation, and rosetted endothelial cells
were either cytocentrifuged onto a glass slide at 200 rpm with low
acceleration (Cytospin, Shandon, UK) or deposited on the slide, dried
in air for further characterization including May-Grünwald-Giemsa
staining, immunologic labeling, and TUNEL assay. They were kept at
 80°C until used.
Immunofluorescence procedure.
Rosetted endothelial cells were rehydrated in PBS, fixed in
paraformaldehyde (PFA; Fluka Chemie, Buchs, Switzerland)
3% and permeabilized with 0.1% Triton X100 (Prolabo,
Vaulx-en-Velin, France) or 0.5% NP-40. Nonspecific sites were coated
with PBS-0.5% milk powder, which was used for antibody dilutions.
Slides were then proceeded for double immunostaining.
They were incubated with anti-vWF antibody together with one of the
following unconjugated MoAbs (antismooth muscle-specific -actin,
ICAM-1, VCAM-1, and E-selectin) previously described, or
FITC-conjugated MoAbs against TF and CD36, during 3 hours. They were
then washed and incubated with the second step reagents:
TRITC-goat anti-rabbit for vWF antibody, FITC-goat
anti-mouse for the unconjugated MoAbs, or PBS-milk for FITC-conjugated
first MoAbs during 2 hours. The slides were then washed with PBS
incubated 5 minutes with 1 µg/mL propidium iodide (Sigma), washed,
and mounted in Moeviol. Cell preparations were analyzed with a Leica
confocal microscope (Lyon, France).
Detection of apoptotic cells.
Apoptotic cells were identified by the detection of DNA strand breaks
using the technique of in situ terminal deoxynucleotidyl transferase-mediated deoxyuridine triphosphate (dUTP)
nick end labeling (TUNEL)23 using a commercial kit: the TdT
in situ apoptosis detection kit-fluorescein (Genzyme Diagnostics,
Cambridge, UK). Biotinylated nucleotides were incorporated into the
apoptically generated DNA ends using TdT. The covalently bound
biotinylated nucleotides were detected using a streptavidin-fluorescein
conjugate. Nuclei were counterstained with a solution of 1 µg/mL
propidium iodide during 5 minutes. Cell preparations were analyzed with a Leica confocal microscope.
Statistical analysis.
The sign test was used to show that the distribution of the CEC data
was nonnormal and is therefore presented as median and interquartile
range (IQR). Data from the four groups of subjects (AMI, UA, EA, and
controls) was analyzed by the Kruskal-Wallis test with three degrees of
freedom. It was then log transformed to allow Tukey's posthoc test to
be applied. Serial data were analyzed by repeated measures (Friedman's
two-way) analysis of variance on Minitab 10 extra.
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RESULTS |
Cross-sectional estimation of CECs.
The levels of CECs were determined on blood samples obtained within 12 hours of the development of symptoms (Fig
1). Median CEC count was 7.5 cells/mL (IQR, 1.5 to 43.5 cells/mL) in
the AMI group and 4.5 cells/mL (IQR, 0.75 to 13.25) in UA. In both the
EA (before test exercise) and control groups, no CECs were detected
(median, both 0; IQR, both 0 to 0). Log transformation of these data
followed by Tukey's test showed levels to be higher in AMI than in
both EA and controls (both P < .01). Levels in UA were higher
than in both EA and controls (both P < .05). Levels between
AMI and UA and between EA and controls were not significantly different. Our data does not support the hypothesis that pharmacologic treatment and the release of endothelial cells are linked because the
four groups received the same medication, but only AMI and UA patients
presented CECs (Table 4). It is noteworthy
that among the four patients treated by thrombolysis, two presented
high levels of CECs (100 and 550 cells/mL), while the remainder
presented low levels (0 and 7 cells/mL within the first 12 hours).
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| Fig 1.
Quantitation of CECs. Levels of CECs in patients with AMI
(n = 21) and UA (n = 32) compared to subjects with EA (n
= 13) and to subjects with noncoronary chest pain (C, n = 14)
within 12 hours of the development of symptoms. Bar represents median
value.
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Serial estimation of CECs.
Five serial blood samples were obtained from 26 patients suffering from
an AMI, four samples from 20 patients in UA, and three samples from 13 patients undergoing the treadmill exercise test (Table 4). Data were
analyzed by Friedman's two-way repeated measures of analysis of
variance (ANOVA). For the AMI, there was a significant peak in CECs at
18 to 24 hours (P = .009 overall repeated measures ANOVA,
P < .05 peak v <12 hours sample and v 30 to
42 hours sample). There was no change in levels of CECs in UA
(P = .451) or in EA, despite the immediate peak postexercise due to exercise testing (P = .417).
Morphologic aspects.
After immunomagnetic separation and cell observation in the
hemocytometer, the same morphologic features were noticed between cells
harvested from AMI and from UA. In both cases, different cytological
patterns were observed after staining with the fluorescent probe
acridin orange or with May-Grünwald-Giemsa: a majority of cells
were 20 to 50 µm in diameter with a large nucleolated nucleus and a
less fluorescent cytoplasm surrounded by beads; they were found alone
or clumped together (Fig 2A and B). We
sometimes observed spindle-shaped cells presenting elongated nuclear
configuration (Fig 2C) and sheets of CECs remaining largely intact with
well-preserved nuclei and well-delineated cytoplasm (Fig 2D).
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| Fig 2.
Cytological analysis of CECs isolated with magnetic beads
coated with S-Endo 1 antibody. Each panel shows CECs isolated from
donors with AMI or UA. Panels A, B, and C show CECs stained with
acridin orange presenting a round shape, clear nuclei and nucleoli
(arrows) (A and B) or a spindle shape (C). Panel D shows a
cell sheet stained with May-GrünwaldGiemsa. Numerous beads
(focus or out of focus) are evident in all panels (arrowheads); they
have a diameter of 4.5 µm.
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Immunofluorescence staining.
To confirm the endothelial origin of the circulating cells and exclude
contamination by smooth muscle cells, double immunological stainings
for vWF and smooth muscle-specific -actin were performed on the
recovered cells. On all cell types examined, vWF was detected as
granular structures uniformly distributed in cytoplasm
(Fig 3A through C), whereas antismooth
muscle-specific -actin labeling was negative. Cell staining was
compared with control slides of mixed human umbilical vein endothelial
cells (HUVEC) (vWF positive) and smooth muscle cells
(-actin positive) (Fig 3D).
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| Fig 3.
Immunofluorescence analysis of CECs. (A, B, and C)
CECs isolated from donors with AMI or UA. Cells of (D) are mixed
endothelial and smooth muscle cells of human umbilical vein for
control. All are stained for both intracellular vWF (red) and
smooth muscle-specific  actin (green). Round- and spindle-shaped
cells (A and B), cell sheets (C), and umbilical vein endothelial cells
(D) show the same granular pattern of vWF expression. The presence of
smooth muscle-specific actin is detected only in control smooth
muscle cells (D). Nuclei are counterstained red with propidium iodide.
The green halo around beads comes from anti-mouse FITC-secondary
antibody bound on the S-endo 1-coated beads.
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Information regarding the origin of CECs was obtained by staining with
CD36, a marker that is present predominantly in microvascular endothelium.24-26 By double-staining for CD36 and vWF, we
showed that the cells isolated in samples from patients with AMI (n = 6) or with UA (n = 6) did not express CD36 and therefore were predominantly of macrovascular origin. Negative and positive controls were provided by lymphocytes and granulocytes (CD36 negative) and
monocytes and platelets (CD36 positive) on blood smears. To assess
whether endothelial cells circulate in an activated state, we analyzed
on the same AMI and UA samples, the dual expression of vWF with
molecules that appear on activated endothelial cells, namely ICAM-1,
VCAM-1, E-selectin, and tissue factor. Only tissue factor was expressed
by 25% of the CECs examined. Cell staining was compared with negative
and positive control slides of resting and TNF (10 ng/mL) activated HUVEC.
Detection of DNA degradation.
To assess if apoptosis accounted for cell desquamation, we investigated
nuclear DNA fragmentation of CECs from four patients with AMI and four
patients with UA displaying the highest levels of CECs. In each sample
examined, less than 10% of cells was found to be positive. The
majority, including the endothelial sheets, were devoid of green
staining showing the lack of DNA breaks, but showed a uniform nuclear
staining with propidium iodide
(Fig 4).
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| Fig 4.
Detection of apoptotic cells with TUNEL assay.
(A and B) CECs isolated from donors with AMI or UA. (C) Control
apoptotic HL60 cells. Nuclei of CECs are counterstained red with
propidium iodide. A large majority of CECs including endothelial sheets
are negative (A). Only rare cells show DNA breaks (B). The control
apoptotic HL60 cells are positive (C).
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DISCUSSION |
To our knowledge, this is the first combined specific capture and
immunologic demonstration of CECs in AMI and UA. Because CECs were not
detected in the controls or in the EA group, their presence in blood is
indicative of endothelial cell detachment in acute coronary syndromes
with probable atheroma plaque rupture. Therefore, our data supplements
preliminary reports that suggested the presence of cells or cell
"carcasses" with endothelial morphological features in these two
pathologies without demonstrating their endothelial nature and their
viability.16,17
In certain other vascular diseases, CECs have been reported to be an
"ex vivo" indicator of vascular injury. The level of CECs varies
according to the extent of the endothelial lesion. A high endothelial
cell count is found in widespread vascular damage associated with
rickettsial vasculitis (2 to 1,600 cells/mL),10 sickle cell
crisis (46 to 430 cells/mL),6-8 or cytomegalovirus infection (0 to 50 endothelial cells per 2 × 105
ficoll-separated mononuclear cells).14,15 However, the
number of CECs found in localized vessel damage, such as after coronary angioplasty, is just over the normal (mean values, approximately 5 to
10 cells/mL).11
In the present study, a large range of CEC counts was found in AMI as
well as in UA, suggesting a wide variation of vessel injury associated
with these events. Notably, some CECs were still observed in the blood
in some patients up to 42 hours after the onset of chest pain. This may
reflect the half-life of these cells, as already reported in patients
undergoing coronary angioplasty (at least 24 hours),11 or
it may reflect protracted vascular irritability or delayed endothelial
cell detachment from the subendothelium of the vessel wall. No cells
were found at rest in EA; this result was not significantly modified
after exercise testing, suggesting that ischemia associated with stable
angina does not induce significant endothelial cell desquamation. In
addition, we can exclude the possibility that the level of CECs
reflects drug treatment because controls (who suffered from a
noncoronary chest pain) received the same medication as patients
(including heparin therapy), but had no CECs. Furthermore, unpublished
data of a group of patients not suffering an acute coronary disease and
treated with various doses of unfractionated heparin (0.1 to 0.5 IU/mL)
did not show any CECs.
The cytological heterogeneity of cells isolated both in AMI and UA is
noteworthy. The majority of CECs presented classical endothelial
morphologic features with rounded shapes. These cells showed the
typical granular cytoplasmic distribution of vWF and the absence of
smooth muscle-specific actin consistent with their endothelial
nature. The same typical granular staining was found on spindle-shaped
cells and endothelial sheets. Such circulating endothelial sheets were
previously described in a dog model of myocardial infarction and were
shown to come from endocardial endothelium.27 Because no
specific immunologic marker for this type of endothelium is available,
it was not possible to show that the CECs are indeed from
cardiovascular origin. These cells tend to be predominantly of
macrovascular origin as defined by the negativity of CD36 staining. We
cannot be completely confident that CD36 CECs are
not microvascular, as some CD36 endothelial cells
have been identified in dermal microvessels.26 The majority
of the isolated cells did not present a proadhesive phenotype, as
evidenced by the absence of expression of three adhesion molecules
ICAM-1, VCAM-1, and E-selectin. This observation can be linked to the
recent demonstration that endothelial cells differentially express cell
adhesion molecules depending on the size of blood vessels, with the
most prominent expression of E-selectin and ICAM in
microvessels.28 Alternatively, it is also possible that CEC
activation state does not accurately reflect the phenotype of the
endothelium remaining attached in situ. Compared with data published by
Solovey et al,7 all we can conclude is that the CECs
recovered in acute coronary diseases and in sickle cell anemia are
different in terms of origin and activated phenotype. A small proportion of CECs expressed tissue factor, suggesting that circulating endothelium may have a procoagulant phenotype, although this hypothesis clearly demands confirmation. Recently, a circulating population of
endothelial putative progenitor cells that expressed vascular endothelial growth factor receptor were identified from
peripheral blood of healthy subjects by CD34 magnetic
beads.29 These cells should differ from those recovered in
this study, as the CECs isolated with S-Endo 1 beads are not detectable
in healthy individuals.
To assess the potential mechanism of cell detachment, we investigated
whether apoptosis was involved. However, 90% of the CECs showed a
remarkably good cytoplasmic and nuclear morphologic preservation
without the usual morphologic features of apoptosis. Because one of the
biochemical hallmarks of apoptosis is fragmentation of chromatin, we
analyzed these cells with a TUNEL assay. In the majority, including the
endothelial sheets, the CECs showed a lack of DNA breakage indicating
that they were not in apoptosis when they detached from the vessel
wall. Moreover, it has been reported that in advanced atheromatous
lesions or in coronary restenosis, apoptosis concerns mostly smooth
muscle cells and monocytes/macrophages.30,31 In the present
study, less than 10% of CECs analyzed showed DNA signs of apoptosis.
It is possible that the apoptotic changes in these cells occurred after
detachment when they were already in the blood, which may be a function
of the lag time between detachment and blood collection.
Apart from apoptosis, several other possibilities can be considered for
the mechanisms responsible for endothelial cell detachment. It might be
due to (1) mechanical dislodgment of cells during plaque rupture (as
supported by angiographic data); (2) proteolysis of subendothelial
matrix proteins triggered by u-PA or t-PA-mediated plasminogen
activation32,33; (3) prolonged ischemia of the heart muscle
can lead to the detachment of the most sensitive endothelial cells such
as endocardial endothelial cells; and (4) oxidative burst occurring
after blood reperfusion.34-36 Another possibility is that
these cells represent cells released from the microcirculation during
ischemia. The fact that CECs are CD36 and probably
not of microvascular origin does not favor this hypothesis.
In conclusion, the measurement of CECs represents a direct evidence of
injury of the endothelial lining among patients with acute coronary
disease. This information can be obtained by a simple, noninvasive test
that may be useful for endothelial exploration and may yield new
insights in the pathophysiology of endothelial injury. Further study
will determine if this endothelial marker can be used as a diagnostic
index, especially in UA. Furthermore, the fact that CECs are not
apoptotic suggests potential use as autologous vectors for diagnosis of
endothelial abnormalities and for vascular therapy.
| |
ACKNOWLEDGMENT |
The authors are grateful to Drs Victor Gurewich and Valentin Fuster for
suggestions and comments on the manuscript and to Biocytex and
Immunotech companies for providing antibodies. We thank Dr Serge Yvorra
for his contribution in the early phase of this study. We also thank
Robert Pistoresi and Michel Dehri for confocal microscope analyses,
Annie Bottari and Andrée Boyer for secretarial work.
| |
FOOTNOTES |
Submitted June 15, 1998; accepted December 18, 1998.
Supported in part by Hoechst Laboratories.
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 Françoise Dignat-George,
PhD, Laboratoire d'Hématologie et d'Immunologie,
U.F.R. de Pharmacie, 27 Bd Jean Moulin, 13385 Marseille Cedex 5, France; e-mail: hematim{at}pharmacie.univ-mrs.fr.
| |
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Monocytes coexpress endothelial and macrophagocytic lineage markers and form cord-like structures in Matrigel(R) under angiogenic conditions
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M. MUTUNGA, B. FULTON, R. BULLOCK, A. BATCHELOR, A. GASCOIGNE, J. I. GILLESPIE, and S. V. BAUDOUIN
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Am. J. Respir. Crit. Care Med.,
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[Abstract]
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E. J. Topol and J. S. Yadav
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F. Dignat-George, A. Blann, and J. Sampol
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T. Stefanec
Circulating Apoptotic Endothelial Cells
Blood,
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J. Yu, S. Tian, L. Metheny-Barlow, L.-J. Chew, A. J. Hayes, H. Pan, G.-L. Yu, and L.-Y. Li
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TOP
ABSTRACT
INTRODUCTION