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Blood, 1 June 2001, Vol. 97, No. 11, pp. 3658-3661
BRIEF REPORT
Resting and activated endothelial cells are increased in the
peripheral blood of cancer patients
Patrizia Mancuso,
Alessandra Burlini,
Giancarlo Pruneri,
Aron Goldhirsch,
Giovanni Martinelli, and
Francesco Bertolini
From the Divisions of Hematology-Oncology,
Pathology-Laboratory Medicine, and Medical Oncology, European Institute
of Oncology, Milan, Italy.
 |
Abstract |
Circulating endothelial cells (CECs) were enumerated in 20 healthy
controls and 76 newly diagnosed cancer patients by means of 4-color
flow cytometry. In breast cancer (n = 46) and lymphoma (n = 30)
patients, both resting and activated CECs were increased by 5-fold
(P < .0008 vs control). CECs significantly correlated with plasma levels of vascular cell adhesion molecule-1 and vascular endothelial growth factor. Resting and activated CECs were similar to
healthy controls in 7 lymphoma patients achieving complete remission
after chemotherapy, and activated CECs were found to decrease in 13 breast cancer patients evaluated before and 24 hours after quadrantectomy.
(Blood. 2001;97:3658-3661)
© 2001 by The American Society of Hematology.
| |
Introduction |
Circulating endothelial cells (CECs) have been
found to be increased in the peripheral blood (PB) of patients affected
by sickle cell anemia,1 cytomegalovirus,2
rickettsial3 infection, myocardial infarction, and
endotoxinemia.4,5 Moreover, increased CECs have
been reported in patients bearing intravascular
instrumentation.6 Considering that the generation of
new blood vessels (angiogenesis) and co-option of preexisting vessels
are crucial steps in cancer progression,7,8 we developed a
novel 4-color flow cytometry procedure to measure CECs and circulating
endothelial cell progenitors (CEPs) in cancer patients. Studies were
performed in whole blood with commercially available monoclonal antibodies.
| |
Study design |
PB was collected in ethylenediaminetetraacetic acid (EDTA) tubes
through 21G needles in 76 newly diagnosed cancer patients (30 with
lymphoma and 46 with breast cancer [BC]) and 20 controls. Among
lymphoma patients, 28 had B-cell low-grade (n = 16, including 8 patients with chronic lymphocytic lymphoma) or high-grade
(n = 12) non-Hodgkin lymphoma and 2 had Hodgkin disease. Five
lymphoma patients (1 mantle cell, 1 marginal zone, 3 chonic lymphocytic lymphoma) had leukemic disease. Among 46 BC patients, all with infiltrating duct carcinoma, 9 were in stage N0, 10 had axillary lymph node metastases, and 27 had distant metastatic diseases. Patients
bearing intravascular instrumentation were excluded from the study.
A panel of monoclonal antibodies, including anti-CD45 to exclude
hematopoietic cells, anti-CD31, -CD34, -CD36, -CD105, -CD106, -CD133,
and -P1H12,1,9 and appropriate analysis gates (Figure 1) were used to enumerate resting and
activated CECs and CEPs. Monoclonal antibodies (Table
1) were conjugated with fluorescein isothiocyanate (FITC), R-phycoerythrin (PE), peridinin chlorophyll protein (PerCP), or allophycocyanin (APC), and cell suspensions were
evaluated by a FACSCalibur equipped with a second red-diode laser
(Becton Dickinson, San Jose, CA). Absolute cell numbers were calculated
by reference fluorescent beads, and "lyse-no-wash" procedures were
used to increase sensitivity and reproducibility.10 After
acquisition of at least 100 000 cells per PB sample, analyses were
considered as informative when adequate numbers of events (> 100,
typically 3-400) were collected in the CEC enumeration gates.
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| Figure 1.
Four-color flow cytometry evaluation of circulating endothelial
cells and endothelial progenitors.
(A) Representative panel (left) shows the analysis gate used to exclude
platelets, death cells, and debris and the reference beads used to
obtain absolute cell count. The other panel (right) shows the gate used
to exclude hematopoietic cells expressing the CD45 antigen. (B,C)
Middle and bottom panels indicate negative controls and the expression
of antigens used to evaluate resting (CD31 and CD34), activated (CD105
and CD106), and progenitor (CD133) endothelial cells. (B) Middle panels
show the resting phenotype of a representative healthy control; (C)
bottom panels show the more activated phenotype of a representative
newly diagnosed BC patient. R4 indicates CD45-hematopoietic progenitors
depicted by high CD34 expression.
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View this table:
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|
Table 1.
Target antigens, related cluster designations, antibody
clones, and conjugation of monoclonal antibodies used in the
study
|
|
Resting CECs were defined as negative for hematopoietic marker CD45;
positive for endothelial markers P1H12, CD31, and CD34; negative for
activation markers CD105 and CD106; and negative for the progenitor
marker CD133. Activated CECs were defined as CD45 ,
P1H12+, CD31+, CD34+, CD105
or CD106+, and CD133. According to
Rafii,11 CEPs were depicted by expression of CD133.
Although some of the monoclonal antibodies used in our CEC panel
(namely, CD31, CD34, CD105, and CD133) react with defined hematopoietic
cell populations in addition to CECs, the P1H12 monoclonal antibody is
highly specific as an endothelial marker. Unlike other commonly used
endothelial markers, P1H12 specifically localizes to endothelial cells
of all vessels including cancerous tissues and does not react with
hematopoietic or epithelial cells.9 Regarding activated
CEC markers, high levels of CD105 (endoglin), a receptor for
transforming growth factor- , and vascular cell adhesion molecule-1
(VCAM-1) are currently considered hallmarks of activated endothelial
cells either in vitro and in tissues undergoing angiogenesis in
vivo.12,13
Sensitivity and specificity of our procedure were evaluated by serial
dilution of human umbilical cord endothelial cells (expressing an
"activated CEC" phenotype) in the U-937 cell line. The detection limit of our procedure was 0.1 cell/µL, and specificity was more than
90%. Anti-CD34 antibodies conjugated with APC, when compared with PE,
have decreased mean fluorescence intensity.10 Although most CD34+ hematopoietic progenitors were excluded from our
analysis gate by CD45 expression,10 APC-conjugated
anti-CD34 was useful to discriminate between the very tiny population
of CD45-hematopoietic progenitors, which were
CD34+++(bright), and CD34+ CECs (Figure 1).
Microvessel density (MVD) was evaluated by anti-CD34 staining in
paraffin-embedded tumor samples as we described in detail elsewhere.14 In brief, sections were immunostained with
CD34-reactive Qbend/10 (Signet Laboratories, Dedham, MA). The
substitution of the primary antibody with nonimmune mouse serum was
used as negative control. For MVD enumeration, at least 10 fields were
evaluated at ×250 magnification (0.78 mm3). Any
brown-staining endothelial cell (or cluster) that was clearly separated
from adjacent microvessels was considered a single, countable
microvessel, and vessel lumens were not a prerequisite to define a
structure as a microvessel. In duplicate independent readings, MVD
intrareader variability was found to be 14% ± 10% (r = 0.879).
Circulating vascular endothelial growth factor (VEGF) and VCAM-1
were measured in the plasma of patients and controls by commercial enzyme-linked immunosorbent assay kits (R&D Systems, Minneapolis, MN,
and Biosource, Camarillo, CA, respectively) as we previously described.15 It has been suggested in the past that VEGF
measurements in EDTA-plasma samples may in part reflect VEGF release
from activated platelets. As described by Wynendaele et
al,16 however, EDTA may influence platelet shape, but no
statistically significant difference in VEGF levels is observed in
plasma samples collected with EDTA versus plasma samples collected with
4 different anticoagulants (sodium citrate, theophylline, adenosine,
and dipyridamole) to obtain maximal platelet stabilization.
| |
Results and discussion |
In healthy controls (n = 20), mean values of resting and
activated CECs were 7.9/µL (95% confidence interval [CI],
4.7-11.1) and 1.2/µL (95% CI, 0.1-2.3), respectively. Seven female
controls were reevaluated during the menstrual period associated with
physiologic active angiogenesis. Mean activated CECs were found to
increase from 2.4/µL (95% CI, < 0.1-5.5) to 4.4/µL (95% CI,
< 0.1-10.5). However, this trend did not reach statistical
significance (P = .15 by Wilcoxon matched-pairs test).
In 76 newly diagnosed patients, mean resting and activated CECs
were 39.1/µL (95% CI, 16.8-61.4) and 6.8/µL (95% CI, 5.0-8.6), ie, increased by 5-fold (P < .0008 vs control by
ANOVA). CEC distribution was normal in controls and skewed in
patients. Three of 5 lymphoma patients in leukemic phase contributed to
most of the skewing observed in CEC distribution among patients; and
lymphoma patients, compared with BC patients, had higher mean resting
(78.0/µL [95% CI, 22.8-133.5] vs 15.8/µL [95% CI, 12.0-19.7],
P = 0.0078 by ANOVA) and activated (8.7/µL [95% CI,
5.1-12.3] vs 5.6/µL [95% CI, 3.7-7.4], P = 0.18)
CECs. Differences between BC patients with early or metastatic diseases
were not significant. In controls and patients, the number of CECs did
not significantly increase with age. Studies are ongoing to fully
understand the clinical features associated with the particularly
elevated CEC values observed in some patients, and repeated CEC
measurements in patients and controls indicated a low
longitudinal CEC variation. Evaluation of CD36 expression showed that
in patients and controls at least half of the CECs were microvascular
in origin.1 In patients and controls, the count of resting
and activated CECs did not correlate with the count of white
cells, red cells, or platelets.
VEGF is produced by most tumor cells7,8 and is
involved in CEP mobilization.17 Mean VEGF was 83 pg/mL
(95% CI, 27-140) and 192 pg/mL (95% CI, 103-281) in controls and
patients, respectively (P = .03 by ANOVA). Correlation
between MVD and VEGF and between MVD and CECs did not reach statistical
significance. On the other hand, a positive correlation
(r = 0.419, P = .009 by multiple regression)
was found between CECs per microliter and plasma VEGF. As shown in
Figure 2, a normal distribution of
resting CECs, activated CECs, and plasma VEGF was observed in controls,
whereas a switch to increased VEGF, increased CECs, and activated CEC
phenotype was observed in cancer patients. Circulating VCAM-1, a
glycoprotein produced by angiogenic endothelial cells, was
significantly increased in patients (mean 1496 ng/mL
[95% CI, 1161-1831]) compared with controls (838 ng/mL
[95% CI, 737-938], P = .003). VCAM-1 strongly correlated with CECs per microliter (r = 0.582,
P < .0001) but not with MVD, and lymphoma patients had
significantly higher VCAM-1 levels compared with those with BC
(P = .002).
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| Figure 2.
Three-dimensional surface plots showing plasma VEGF and
resting and activated CECs in healthy subjects and cancer patients.
A normal distribution was prevalent in controls, whereas a switch to
increased VEGF, increased CECs, and an activated CEC phenotype was
observed in cancer patients.
|
|
CECs were found to be similar to control values in 7 lymphoma patients
who achieved complete remission after chemotherapy. In these patients,
mean resting and activated CECs were 12.8/µL (95% CI, 4.0-21.6) and
0.8/µL (95% CI, 0.4-1.2), respectively (P = .0001 and
.63 vs newly diagnosed patients and healthy controls, respectively).
Furthermore, CECs were found to decrease in 13 BC patients evaluated
before and 24 hours after quadrantectomy (Figure
3). In these BC patients, activated CECs
significantly decreased from 9.0/µL (95% CI, 2.4-15.5) to 2.0/µL
(95% CI, 0.9-3.1, P = .0107 by Wilcoxon matched-pairs
test), whereas mean resting CECs decreased from 25.3/µL (95% CI,
16.3-34.4) to 16.4/µL (95% CI, 10.0-22.8), and statistical
significance was borderline (P = .0546).
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| Figure 3.
CECs in 13 BC patients before and after quadrantectomy.
A decrease was observed in both activated and resting CECs.
P values were calculated by the Wilcoxon matched-pairs
test.
|
|
CEPs were below 0.5/µL in all controls and newly diagnosed patients
evaluated. Higher CEP counts were found in 4 of 11 patients evaluated
while recovering from high-dose chemotherapy-induced aplasia and in 2 of 7 healthy controls evaluated during the menstrual period. In these 6 cases, CEP values ranged from 1.1/µL to 9/µL.
Increased CECs have been described so far only in conditions that have
in common the presence of vascular injury.1-6 Our finding
that resting and activated CECs are increased in newly diagnosed cancer
patients and decline after cure underlines the crucial role of
angiogenesis in both solid tumors and hematopoietic malignancies.
Moreover, resting and activated CECs appear to be novel and promising
surrogate angiogenesis markers. Interestingly, our data about the
increase of activated (CD105 or CD106+) CECs in
cancer patients offer a rationale for recent reports about the
predictive potential of soluble CD105 and CD106 in the PB of BC
patients.18,19
The increase of resting and activated CECs in cancer patients may be
due to different biological reasons: CECs may derive from newly formed
tumor vessels or, alternatively, represent ingress of proliferating
endothelial cells from neighboring normal tissue or even from distant
uninvolved vessels activated by tumor's derived cytokines. To better
distinguish between these hypotheses, we are evaluating CEC kinetic and
cell cycle in animal models of human cancer. In parallel, the
measurement of CEC viability is currently under investigation to
ascertain whether a given drug therapy has antiangiogenic properties.
| |
Note added in proof |
Chang et al20 have recently provided evidence indicating
that tumor blood vessels may be mosaics in which both endothelial and
tumor cells form the luminal surface. These data provide a possible
explanation for the finding of high CEC numbers in cancer patients.
| |
Footnotes |
Submitted September 25, 2000; accepted February 2, 2001.
The publication costs of this
article were defrayed in part by
page charge payment. Therefore,
and solely to indicate this fact,
this article is hereby marked
"advertisement"
in accordance with 18 U.S.C.
section 1734.
Reprints: Francesco Bertolini, Hematology-Oncology, European
Institute of Oncology, via Ripamonti 435, 20141 Milan, Italy;
e-mail: francesco.bertolini{at}ieo.it.
| |
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Top
Abstract
Introduction