Yonago Acta medica 1997;40:21–30

Expression of Vascular Cell Adhesion Molecule-1 on Human Pulmonary
Artery Endothelial Cells and Human Umbilical Vein Endothelial Cells
α, Interleukin-1β
β and LipopolysacStimulated by Tumor Necrosis Factor-αcharideHiroyuki Sano, Katsuyuki Tomita and Yutaka Hitsuda
Third Department of Internal Medicine, Faculty of Medicine, Tottori University
It has been recently suggested that there is a diversity of endothelial cells (ECs) fromdifferent organs in response to cytokines and lipopolysaccharide (LPS). We examined
whether there was a differential response among human pulmonary artery ECs(HPAECs) and human umbilical vein ECs (HUVECs) with regard to the expression ofα (TNF-αα ),vascular cell adhesion molecule-1 (VCAM-1) by tumor necrosis factor-αβ (IL-1ββ) and LPS stimulation. VCAM-1 expression on these ECs stimulatinterleukin-1β
ed by the stimulators was measured by cellular enzyme-linked immunosorbent assay(ELISA) and flow cytometric analysis. For confirmation of our use of cells as ECs, theexpression of Factor VIII and CD36 were also measured by flow cytometric analysis.
Factor VIII was expressed on unstimulated HPAECs and HUVECs at positive cell,
43.1% and 28.9%, respectively, but not CD36 on these ECs entirely. VCAM-1 was not
expressed on unstimulated HPAECs and HUVECs. Maximal expression of VCAM-1
α , IL-1ββ and LPS at doses of 10
on these ECs was induced 6 h after stimulation by TNF-α
ng/mL, 1 ng/mL and 1 µg/mL, respectively. There was a significant difference in the
maximal expression between these ECs (P < 0.05; Mann-Whitney U test). Furthermore,
VCAM-1 expression by any stimulation was invariably greater on HPAECs than on
HUVECs throughout the course of the experiment (P < 0.01; two-way ANOVA). Therefore, we suggested that ECs from different organs might have diversity with regard to
α , IL-1ββ and LPS stimulation.
VCAM-1 expression by TNF-α
Key words: cellular ELISA; endothelial cells; VCAM-1

Endothelial cells (ECs) play key roles in thecreation of the barrier to endotoxin and interaction with inflammatory cells during migration into the specific organs. Susceptibility ofcultured ECs to endotoxin-induced injury varies
among species and among sites of origin of thecells. The pulmonary artery cell line (Harlan etal., 1983a) and kidney microvascular cell line
(Raghu et al., 1986) are quite susceptible to theeffects of endotoxin whereas human umbilicalvein ECs (HUVECs) (Harlan et al., 1983b; Raghu

et al., 1986) and human omentum (Smedly et al.,1986) are particularly resistant.
The interaction between ECs and inflammatory cells is now considered to form an important part of the inflammatory process throughsome adhesion molecules. ECs are stimulated
by proinflammatory cytokines, e.g., tumor
necrosis factor-α (TNF-α) and interleukin-1β(IL-1β), as well as endotoxin, to increase expression of cell surface adhesion molecules, leading todramatically altered interactions with leuko-

Abbreviations: ANOVA, analysis of variance; EC, endothelial cell; ELAM-1, endothelial-leukocyte adhesion
molecule-1; ELISA, enzyme-linked immunosorbent assay; FACS, fluorescence activated cell scan; FITC,
fluorescein isothiocyanate; HPAEC, human pulmonary artery EC; HUVEC, human umbilical vein EC; ICAM-1,
intercellular adhesion molecule-1; IL-1β, interleukin-1β; LPS, lipopolysaccharide; mAb, monoclonal antibody;
MFI, mean fluorescence intensity; OD, optic density; TNF-α, tumor necrosis factor-α; VCAM-1, vascular celladhesion molecule-1

21

H. Sano et al.

cytes, e.g., granulocytes and monocytes. In
these interactions, endothelial-leukocyte adhesion molecule-1 (ELAM-1), intercellular adhesion molecule-1 (ICAM-1) and vascular cell adhesion molecule-1 (VCAM-1) are known to play animportant role, as they are presented by the ECs
and interact with corresponding ligands on theleukocyte membranes (Carlos and Harlan, 1994).
VCAM-1 is an especially important adhesion
molecule that is induced on ECs by cytokineand lipopolysaccharide (LPS)-stimulation, andcan mediate the binding of lymphocytes,
eosinophils or monocytes to the ECs in inflammatory lung diseases, e.g., bronchial asthma(Ohkawara and Yamauchi, 1993) and adultrespiratory distress syndrome (Grau et al., 1996).
Until now, the regulation of these proteins,
including ELAM-1, ICAM-1 and VCAM-1, hasbeen most often studied on ECs derived fromthe umbilical vein (Deisher et al., 1993; Gille etal., 1996; Grau et al., 1992; Swerlick et al.,
1992b). It is suspected that the effect of VCAM-1
expression by stimulation of ECs are diverseand complicated, and vary between species andin cellular origin. Swerlick and colleagues(1992a) have already reported that there was asignificant difference in VCAM-1 expression
between human dermal ECs and HUVECs.
We considered that it was necessary to examine the regulation of expression of VCAM-1
on human pulmonary artery ECs (HPAECs) inairway inflammation models. The aim of thisstudy was to compare the reaction profiles of
VCAM-1 expression on HPAECs and on
HUVECs obtained from the same passages after
defined periods of exposure to cytokines andLPS.

CO2 by using EC growth medium consisting ofmodified Hu-Media EB (Kurabo) supplemented with 10 ng/mL recombinant endothelial
growth factor, 5 ng/mL recombinant fibroblast
growth factor, 1 µg/mL hydrocortisone, 50 µg/
mL gentamicin, 50 ng/mL amphotericin B
sulfate and 2% fetal bovine serum in culture
dishes (Falcon 3002, Becton Dickinson
Labware, Lincoln Park, NJ). Experiments werecarried out using cells at the 5th passage, andHUVECs or HPAECs were used for experiments when they had formed confluent monolayers in culture dishes. The confluent ECs
were removed from culture dishes with 0.125%
EDTA (Kurabo) in phosphate buffered saline(PBS) (Research Institute for Medical Disease,
Osaka), centrifuged and resuspended in EC
growth medium at a concentration of 2 × 104
cells/mL. The ECs were then cultured overnight on 0.1% gelatin (Sigma, St. Luois, MO)coated 96-well microtiter plates (Nunc,
Roskilde, Denmark). The ECs were stimulatedby TNF-α, IL-1β and LPS at the desired final
concentration. After stimulation for the required length of time, the wells were washed withwarm EC growth medium. The cells wereconfirmed to be ECs by morphologic criteria andflow cytometric analysis.

Monoclonal antibodies and stimulator
The monoclonal antibody (mAb) directed against
VCAM-1 (CD106; I.G 11B1) and the vonWillebrand factor (Factor VIII) [fluorescein
isothiocyanate (FITC)-conjugated] werepurchased from Serotec Inc. (Bicester, United
Kingdom). The anti-CD36 mAb (IgG1, FITCconjugated) was purchased from Immunotech
Co. (Marseille, France). The biotinylate goatanti-mouse Ig [(Fab)2 fragment] was purchasedfrom Jackson ImmunoResearch Lab. (Avondale,
PA). The phycoerythrin-conjugated antibody waspurchased from Organon Teknika (Durham,
NC). Recombinant human TNF-α and IL-1β
were purchased from Genzyme Diagnostics(Boston, MA). LPS (E. coli, 055, B5) was
purchased from Difco Laboratories (Detroit,
MI).

Materials and Methods

Endothelial cell monolayers
HPAECs and HUVECs were purchased from
Kurabo Co. Ltd. (Osaka, Japan). HPAECs
came from a single donor and were received
cryopreserved (5 × 105 cells) in the 3rd passage.
HUVECs were received in the 1st passage. The
cells were cultured separately at 37°C in 5%
22

Expression of VCAM-1 on HPAECs

Fig. 1. Micrographs of May-Grünwald Giemsa staining of human pulmonary artery endothelial cells (HPAECs)(A) and human umbilical vein endothelial cells (HUVECs) (B) of the 5th passage cells. A and B, × 200.

lysine monohydrochloride and 2.1 mg/mL solution m-periodate for 10 min. This procedure
has been shown to preserve the antigenicity ofmany protein and carbohydrate determinations(Van Ewijk et al., 1980; Van Duinen et al.,Cellular enzyme-linked immunosorbent
assay
The EC monolayers were fixed by incubating witha solution of 2% paraformaldehyde, 0.075% L23

Relative cell number

Relative cell number

H. Sano et al.

1984). Fixation was stopped by
300
300
aspiration of a fixative, washing
the cells with 100 mM glycine,
200
200
0.1% bovine serum albumin (BSA)(Sigma) in Hank’s balanced salt
100
100
solution (Bio Whittaker,Walkersville,MD) (blocking solution). The00assay plates were stored at 4°C
10
10
10
10
10
10
10
10
10
10
with 200 µL/well blocking soluFluorescence intensity
Fluorescence intensity
tion. An enzyme-linked immuno- Fig. 2. Flow cytometric analysis of von Willebrand factor on
sorbent assay (ELISA) was used, HPAECs (A) and HUVECs (B). HPAECs and HUVECs wereas previously described (Cui et al., stained by anti-von Willebrand factor monoclonal antibody and were1983), to measure the expression analyzed by a flow cytometry as described in Materials andof VCAM-1 on ECs. The ELISA Methods. Von Willebrand factor was expressed 30 to 40% on
HPAECs and HUVECs.
was performed at room temperature after 3 times washing with
0.1% BSA in PBS. The ECs were incubated for experiment but were always less than 0.01 at
1 h with anti-VCAM-1 antibody (CD106). OD.
After 3 times washing with PBS, biotinylated
goat anti-mouse Ig was added. After 3 times
Immunofluorescence staining and flow
washing with PBS, the enzyme substrate 0.5
cytometrymg/mL o-phenylenediamine and 0.03% hydrogen peroxide in citrate-phosphate buffer The expression of surface antigens on HPAECs(Sumitomo Bakelight Co., Tokyo, Japan) was and HUVECs was detected by the binding ofthen added. Color development was stopped monoclonal antibodies to VCAM-1 (CD 106),with 2 N sulfuric acid, and the optic density CD36 and von Willebrand factor (Factor VIII)(OD) of each well was read at 492 nm in a for 90 min on ice. After washing the cells 3
Micro plate reader (Tosoh Ltd., Tokyo). Test and times with PBS containing 0.1% sodium azide
control measurements were performed 5 times (Wako, Osaka), the expression of CD36 andin each experiment, and the degree of specific Factor VIII was evaluated by indirect fluoresmAb binding was calculated by subtracting the cence after incubation with either FITC- ormean negative control value from each test phycoerythrin-conjugated antibodies (Organon
value. The results were expressed as mean ± Teknika) for 30 min on ice in the dark. VCAMSD for each set of measurements. Negative 1 expression was evaluated by direct immunocontrol values differed from experiment to fluorescence staining with FITC-conjugatedprimary antibody. Flow cytometry
300
300
was performed with 10,000 cells per
sample on a fluorescence activated cell
200
200
scan (FACS) analyzer (Becton
Dickinson, Mountain View, CA). The
100
100
ECs were analyzed by selective gatingbased on forward and side scatter para00meters. Results were expressed as
10
10
10
10
10
10
10
10
10
10
mean fluorescence intensity (MFI),
Fluorescence intensity
Fluorescence intensity which reflects the cells surface density
Fig. 3. Expression of CD36 on HPAECs (A) and HUVECs (B). of the respective marker, and were
Unstimulated HPAECs and HUVECs were stained by anti-CD36
monoclonal antibody and were analyzed by flow cytometry as compared with those obtained withdescribed in Materials and Methods. CD36 was not expressed unlabelled cell and isotype controls
12

3

Relative cell number

Relative cell number

0

0

1

2

3

4

0

1

2

on HPAECs and HUVECs.

24

3

4

4

0

1

2

3

4

Expression of VCAM-1 on HPAECs

Results(Becton Dickinson) with the use of the Lysis II
program (Becton Dickinson).

Morphology of HPAECs and HUVECs
Statistical analysis

All ECs started from small clumps of cells andspreaded into monolayers within 2 weeks.
HPAECs and HUVECs grew slowly at first andthe cells divided only at the edge of each clump.
After 7 to 10 days in culture, however, cell division was more rapid and became confluent
within a matter of days instead of weeks. Unstimulated HPAECs and HUVECs were stainedby May-Grünwald Giemsa methods. HPAECs(Fig. 1A) showed some heterogeneity in shapein that at least 10% of the cells were moreelongated than HUVECs (Fig. 1B), while bothECs appeared to be of the typical cobblestonemorphology in culture.

Data were presented as mean ± SD. The statistical analysis of difference between HPAECs
and HUVECs was performed by two-way
ANOVA and the nonparametric Mann-Whitney
U unpaired test. P values less than 0.05 wereconsidered to be significant. These statisticalanalyses were performed with the Stat View
4.11 statistics package (Abacus Concepts,
Berkeley, CA).

Optic density (492 nm)

**

A

2.5

NS

2.0

1.5

C

1.0

C

C

0.5

C

**

Both ECs were trypsinized and fluorescently
labeled with the antibody to Factor VIII. Figure
2 shows that Factor VIII was expressed on unstimulated HPAECs as well as unstimulated
HUVECs at positive cells, 43.1% and 28.9%,
respectively.

0
0

3

6

12

24

Incubation time (h)2.5Optic density (492 nm)

Factor VIII expression on HPAECs and
HUVECsB
2.0

Expression of CD36 on HPAECs and
HUVECs

1.5

mAb-recognizing CD36 failed to identify theexpression of CD36 on HPAECs and HUVECs
by flow cytometric analysis (Fig. 3). Furthermore, the stimulation on HPAECs and
HUVECs by TNF-α (10 ng/mL, 6 h), IL-1β (1ng/mL, 6 h) or LPS (1 µg/mL, 6 h) did not resultin the induction of cell surface CD36.

1.0

0.5

0

0

100 pg/mL 1 ng/mL 10 ng/mL 100 ng/mL

TNF-α concentration
Fig. 4. VCAM-1 expression on HPAECs and
HUVECs measured by cellular ELISA after TNF-α
stimulation. HPAECs (o) and HUVECs (∆) in thetime-dependent manner were stimulated by TNF-α (10ng/mL) (A). HPAECs (hatched columns) and
HUVECs (open columns) in the dose-dependent
manner were stimulated by TNF-α for 6 h (B). Values
are mean ± SD; n = 5. Significantly different (**P <0.01) by analysis with two-way ANOVA and thenonparametric Mann-Whitney U test between
HPAECs and HUVECs in the time-dependent manner.
NS, not significant.

Cellular ELISA analysis of VCAM-1 expression on HPAECs and HUVECs stimulated by TNF-α
VCAM-1 was not expressed on unstimulated
HPAECs nor unstimulated HUVECs, but it was
induced in a time- and dose-dependent manneron HPAECs and HUVECs stimulated by TNFα (Figs. 4A and B). VCAM-1 expression was25

H. Sano et al.

2.5

A
**

2.0

1.5

1.0

C

0.5

C

C
C

**

Optic density (492 nm)

Optic density (492 nm)

2.5

0

1.5

3

6

12

**

1.0

0.5

0
0

A

2.0

24

0

C
3

Incubation time (h)

C

6

12

C

**

24

Incubation time (h)
2.5

2.5

B

Optic density (492 nm)

Optic density (492 nm)

C

2.0

1.5

1.0

0.5

B

@

2.0

1.5

1.0

0.5

0

0

0

0

10 pg/mL 100 pg/mL 1 ng/mL 10 ng/mL

10 ng/mL 100 ng/mL 1 µg/mL 10 µg/mL

LPS concentration

IL-1β concentration

Fig. 6. VCAM-1 expression on HPAECs and
HUVECs measured by cellular ELISA after LPS
stimulation. HPAECs (o) and HUVECs (∆) in thetime-dependent manner were stimulated by LPS (1µg/mL) (A). HPAECs (hatched columns) and
HUVECs (open columns) in the dose-dependent
manner were stimulated by LPS for 6 h (B). Values
are mean ± SD; n = 5. Significantly different (**P <0.01) by analysis with two-way ANOVA and thenonparametric Mann-Whitney U test between
HPAECs and HUVECs in the time-dependent manner.

Fig. 5. VCAM-1 expression on HPAECs and
HUVECs measured by cellular ELISA after IL-1β
stimulation. HPAECs (o) and HUVECs (∆) in thetime-dependent manner were stimulated by IL-1β (1ng/mL) (A). HPAECs (hatched columns) and
HUVECs (open columns) in the dose-dependent
manner were stimulated by IL-1β for 6 h (B). Values
are mean ± SD; n = 5. Significantly different (**P <0.01) by analysis with two-way ANOVA and thenonparametric Mann-Whitney U test between
HPAECs and HUVECs in the time-dependent manner.

greater on HPAECs than on HUVECs in a timedependent manner (P = 0.009; two-way ANOVA)(Fig. 4A). After stimulation by TNF-α with 10
ng/mL, VCAM-1 expression on both ECs
reached a plateau at 6 h. VCAM-1 expression
on HPAECs and HUVECs significantly diminished from the peak level after 24 h (P = 0.009;
Mann-Whitney U test) of TNF-α stimulation,
but not after 12 h (P = 0.251; Mann-Whitney Utest). Maximal expression of VCAM-1 wasalso greater on HPAECs stimulated by TNF-α
than on HUVECs (P = 0.020; Mann-Whitney Utest) (Figs. 4B and 7).

Cellular ELISA analysis of VCAM-1 expression on HPAECs and HUVECs stimulated by IL-1β
IL-1β also induced the expression of VCAM-1
on both HPAECs and HUVECs in a time- anddose-dependent manner. There was a significant difference in VCAM-1 expression between
both ECs in a time-dependent manner (P = 0.001;
two-way ANOVA) (Figs. 5A and B). After
stimulation by IL-1β with 1 ng/mL, VCAM-1
expression on both ECs reached maximal value at
6 h. VCAM-1 expression on HPAECs and26

Expression of VCAM-1 on HPAECs

Optic density (492 nm)

HPAEC

*

2.5

*

2.0

HUVEC
300

300

A

B

200

200

100

100

1.5

*
1.0

0.5

0

0
100

101

10 2

103

10 4

100

101

102

103

10 4

102

103

10 4

10 2

10 3

104

102

103

104

0

LPS

1 µg/mL

IL-1β

TNF-α

1 ng/mL

10 ng/mL

Fig. 7. Comparative effects of VCAM-1
expression on HPAECs (hatched columns)
and HUVECs (open columns) after 10 ng/
mL TNF-α, or 1 ng/mL IL-1β or 1 µg/mL
LPS stimulation for 6 h. Data are expressed
as mean ± SD; n = 5. Significantly different(*P < 0.05) by analysis with the MannWhitney unpaired U test between HPAECs
and HUVECs.

HUVECs after 12 h of IL-1β stimulation significantly diminished (P =0.009; Mann-Whitney U test) from thepeak level, and thus IL-1β-induced
expression of VCAM-1 expression on
HPAECs and HUVECs was moretransient than TNF-α-induced expression. We also observed that IL-1β was
not as potent an inducer of VCAM-1
expression on HPAECs and HUVECs
such as TNF-α (Fig. 7). Maximal expression of VCAM-1 stimulation was
also greater on HPAECs stimulated by
IL-1β than on HUVECs (P = 0.020;
Mann-Whitney U test) (Figs. 5B and 7).

Cellular ELISA analysis of VCAM1 expression on HPAECs and
HUVECs stimulated by LPS
LPS also enhanced VCAM-1 expression on both HPAECs and HUVECs ina time- and dose-dependent manner(Fig. 6A and B). There was also a significant difference in VCAM-1 expression between both the ECs in a timedependent manner (P < 0.001; two-way
ANOVA) (Fig. 6A). After stimulation
by LPS with 1 µg/mL, VCAM-1 expres-

300

300

C

Relative cell number

Control

D

200

200

100

100

0

0
100

101

10 2

103

10 4

100

101

300

300

E

F

200

200

100

100

0

0
100

101

102

103

300

10 4

10 0

10 1

300

H

G
200

200

100

100

0

0
10 0

10 1

10 2

103

10 4

10 0

10 1

Fluorescence intensity
Fig. 8. Flow cytometric analysis of VCAM-1 expression.
Unstimulated HPAECs (A) and HUVECs (B) did not express
VCAM-1, but HPAECs (C, E, G) and HUVECs (D, F, H) stimulated with TNF-α (C, D), IL-1β (E, F) and LPS (G, H)
induced to express VCAM-1.

sion on both ECs reached maximal value at 6 h. Although
VCAM-1 expression on HPAECs significantly diminished from the peak level after 12 h (P = 0.009; MannWhitney U test) of LPS stimulation, down regulation of
VCAM-1 expression on HUVECs was not seen until
after 24 h. The stimulation by LPS on HPAECs and
HUVECs resulted in low levels of VCAM-1 induction
when examined to compare LPS stimulation with IL-1β
and TNF-α stimulation (Fig. 7). Maximal expression of
VCAM-1 was obviously greater on HPAECs stimulated27

H. Sano et al.

by LPS than on HUVECs (P = 0.020; MannWhitney U test) (Figs. 6B and 7).

HPAECs and HUVECs as macrovessel celllines with TNF-α, IL-1β and LPS stimulation.
Our results in a time- and dose-dependent
response of VCAM-1 expression on HUVECs
stimulated by cytokines and LPS are shown tobe consistent with the observation of Carlos andcolleagues (1990). We found that the degree of
VCAM-1 expression was significantly different
at high doses of these stimulators when compared to the degree of expression found between HPAECs and HUVECs by cellular
ELISA and flow cytometry. The reason for thedifference of the expression on these ECs is notclear. However, these phenomena may be explained at least by the different numbers of surface receptors to cytokines and LPS (Akeson etal., 1992; Lou and Grau, 1993), and the differing regulation of the VCAM-1 gene transcription level. Gille and colleagues (1996) reported
that VCAM-1 expression by IL-1 stimulation
was lower on human dermal microvascular ECs
than on HUVECs, because human dermal
microvascular ECs have specific reductions of
VCAM-1 gene transcription in response to IL1. We considered that the ECs from different
organs might be diverse in their responses tocytokines and LPS. Similar results to ours havebeen reported previously. For instance, Hauser
and colleagues (1993) also found different
levels of VCAM-1 expression on iliac arterial
and venous cultured ECs. In binding assays,
cells cultured from the bovine pulmonary arteryare quite susceptible to the endotoxin effect aspolymorphonuclear neutrophil-EC detachments, whereas HUVECs are particularly
resistant to binding assays (Staub et al., 1982;
Harlan et al., 1983a, 1983b; Meyrick et al.,
1989, 1995).
TNF-α was the most potent signal at thepeak level and the persistent expression of
VCAM-1 on both cell lines. This result was consistent with Swerlick and colleagues (1992a)
showing that TNF-α induced more VCAM-1 on
HUVECs and human dermal ECs than any
cytokines. Although the mechanism of thiscytokine to induce VCAM-1 was not known indetails, Libby and colleagues (1995) demonstrated that TNF-α could induce IL-1 mRNA in
human ECs. We considered that this effect ofFlow cytometry analysis of VCAM-1
expression on HPAECs and HUVECs
Flow cytometry was performed with 10,000
cells per sample on a FACS analyzer. Fifthpassage cells of HPAECs and HUVECs were
stained with mAb-recognizing VCAM-1 aftertrypsinization. HPAECs and HUVECs did not
express VCAM-1 at mean fluorescenceintensity (MFI; 24.2 and 16.7, respectively),
when unstimulated. VCAM-1 expression was
significantly greater on HPAECs than on
HUVECs when stimulated by 10 ng/mL TNF-α(MFI; 193.0 and 113.8, respectively), 1 ng/mL
IL-1β (MFI; 82.4 and 49.1, respectively) or 1
µg/mL LPS (MFI; 28.2 and 18.7, respectively)
for 6 h (Fig. 8).

Discussion
We compared HPAECs with HUVECs with regard to VCAM-1 expression induced by TNFα, IL-1β and LPS. VCAM-1 was expressed
greater on HPAECs than on HUVECs stimulated by these cytokines and LPS. Moreover,
TNF-α was the most potent signal for VCAM-1
expression on both cell lines.
Both cultured cell lines used in this study
were identified as ECs by the expression of
Factor VIII on surfaces and by the typicalcobblestone morphology in confluent culture,but some HPAECs were more elongated than
HUVECs in culture. Both of our cell lines of
the 5th passage expressed Factor VIII in 30 to
40% of the positive cells, because the ECs in thegrowth phase hardly express Factor VIII (Jaffe
et al., 1974; Johnson et al., 1980). Moreover,
we have demonstrated that CD36, which wasreported to be expressed on a majority of
microvessels (Tandon et al., 1989; Swerlick etal., 1992b) but not detected on macrovessels(Knowles et al., 1984; Buckley et al., 1985),
was not expressed on HPAECs and HUVECs
by flow cytometric analysis. So, we examinedthe differential expression of VCAM-1 between
28Expression of VCAM-1 on HPAECs

autocrine might be attributed to the persistentexpression of VCAM-1 by TNF-α.
VCAM-1 is one of the many inducible adhesion molecules that play an important role in thebinding of lymphocytes, monocytes and eosinophils to ECs (Weller et al., 1991; Vonderheide
and Springer, 1992; Meerschaert and Furie,1995). Upregulation of the expression of
VCAM-1 on ECs is an essential mechanism forthe selective accumulation of these inflammatory cells in the lung tissue of patients withasthma (Ohkawa and Yamauchi, 1993;Håkansson et al., 1995). Its regulation has been
examined in detail in HUVECs, but it has notbeen completely investigated in HPAECs. As
we demonstrated that VCAM-1 expression was
greater in HPAECs than in HUVECs, ourresults may imply that inflammatory cells moreeasily migrate into lung tissue. However, inairway inflammation, inflammatory cells
themselves take an important part in migrating
into lung tissue as well as ECs, and are upregulated to express the ligands to VCAM-1,
i.e., CD11b/CD18 and very late antigen-4 bystimulators (Elices et al.,1990; Schwartz et al.,
1990; Håkansson et al., 1995). Therefore, when
inflammatory models are studied, it is necessary to perform not only quantity assay by cellular ELISA but also binding assay.
In conclusion, we clearly demonstrated that
VCAM-1 expression was induced greater on
HPAECs than on HUVECs. So, we think that
there may exist heterogeneity of ECs obtained
from various organs in response to cytokines
and LPS. We suggest that HPAECs rather than
HUVECs should be used for these experimentations in lung inflammatory models.

References
1 Akeson AL, Mosher LB, Woods CW, Schroeder
KK, Bowlin TL. Human aortic endothelial cells
express the type I but not the type II receptor forinterleukin (IL-1). J Cell Physiol 1992;153:583–
588.
2 Buckley PJ, Dickson SA, Walker WS. Human
splenic sinusoidal lining cells express antigens
associated with monocytes, macrophages, endothelial cells and T lymphocytes. J Immunol
1985;134:2310–2315.
3 Carlos TM, Harlan JM. Leukocyte-endothelial
adhesion molecules. Blood 1994;84:2068–2101.
4 Carlos TM, Schwartz BR, Rosa M, Newman B,
Wayner E, Benjamin C, et al. Vascular cell adhesion molecule-1 (VCAM-1) mediates lymphocyte adherence to cytokine-activated cultured
human endothelial cells. Blood 1990;76:965–
970.
5 Cui YC, Tai PC, Gatter KC, Mason DY, Spry CJ.
A vascular endothelial antigen with restricteddistribution in human fetal, adult, and malignant
tissue. Immunology 1983;49:183–189.
6 Deisher TA, Haddix TL, Montgomery KF,
Pohlman TH, Kaushansky K, Harlan JM. The roleof protein kinase C in the induction of VCAM-1
expression on human umbilical vein endothelial
cells. FEBS 1993;331:285–290.
7 Elices Mj, Osborn L, Takada Y, Crouse C,
Luhowskyj S, Hemler M, et al. VCAM-1 on activated endothelium interacts with the leukocyte
integrin VLA-4 at a site distinct from the VLA-4/
fibronectin binding site. Cell 1990;60:577–584.
8 Gille BJ, Swerlick RA, Lawley TJ, Caughman
SW. Differential regulation of vascular celladhesion molecule-1 gene transcription by tumor
necrosis factor α and interleukin-1α in dermal
microvascular endothelial cells. Blood 1996;87:
211–217.
9 Grau GE, Mili N, Lou JN, Morel DR, Ricou B,
Lucas R, et al. Phenotypic and functional analysis of pulmonary microvascular endothelial cells
from patients with acute respiratory distresssyndrome. Lab Invest 1996;74:761–770.
10 Harlan JM, Harker LA, Reidy MA, Gajdusek
CM, Schwartz SM, Striker GE. Lipopolysaccharide-mediated bovine endothelial cell injury in vitro. Lab Invest 1983a;48:269–274.
11 Harlan JM, Harker LA, Striker GE, Weaver LJ.
Effects of lipopolysaccharide on human endothelial cell in culture. Thromb Res 1983b;29:15–26.
12 Hauser IA, Johnson DR, Madri JA. Differential
induction of VCAM-1 on human iliac venous andarterial endothelial cells and its role in adhesion.
J Immunol 1993;151:5172–5185.
13 Håkansson L, Björnsson E, Janson C, Schmekel
B. Increased adhesion to vascular cell adhesion

Acknowledgments: The authors wish to thank Prof.
Takao Sasaki, Third Dept. of Internal Medicine,Prof. Tohru Mori, Second Dept. of Surgery, Prof.
Yasuaki Kawai, Second Dept. of Physiology, Faculty of Medicine, Tottori University, for their helpful
suggestions in this study. We also would like tothank Dr. Takahiko Tanigawa, Dept. of Bacteriology, Faculty of Medicine, Tottori University, for
his technical suggestion.

29

H. Sano et al.

human endothelium. J Clin Invest 1990;85:
2019–2022.
25 Smedly LA, Tonnesen MG, Sandhaus RA,
Haslett C, Guthrle LA, Johnston RB, et al.
Neutrophil-mediated injury to endothelial cells:
enhancement by endotoxin and essential role ofneutrophil elastase. J Clin Invest 1986;77:1233–
1234.
26 Springer TA. Adhesion receptors of immune
system. Nature 1990; 346:425–434.
27 Staub NE, Schultz EL, Albertine KH. Leukocytes and pulmonary microvascular injury. Ann
NY Acad Sci 1982;38:332–343.
28 Swerlick RA, Lee KH, Li LJ, Sepp NT, Caughman
SW, Lawley TJ. Regulation of vascular cell adhesion molecule 1 on human dermal microvascularendothelial cells. J Immunol 1992a;149:698–705.
29 Swerlick RA, Lee KH, Wick TM, Lawley TJ.
Human dermal microvascular endothelial but not
human umbilical vein endothelial cells express
CD36 in vivo and in vitro. J Immunol 1992b;148:
78–83.
30 Tandon NN, Lipsky RH, BurgessWH, Jamieson
GA. Isolation and characterization of platelet
glycoprotein IV (CD36). J Biol Chem 1989;262:
7570–7575.
31 Van Duinen SG, Mauw BJ, de Graaff Reitsma
CB, Ruiter DJ. Methods in laboratory investigation: immunoeletron microscopic methods fordemonstration of antigens on normal humanmelanocytes and other epidermal cells. Lab
Invest 1984;50:733–741.
32 Van Ewijk W, Coffman RC, Weissman IL.
Immunoelectron microscopy of cell surface antigens: a quantitative analysis of antibody binding
after different fixation protocols. Histochem J
1980;12:349–361.
33 Vonderheide RH, Springer TA. Lymphocyte
adhesion through very late antigen 4: evidence
for a novel binding site in the alternatively spliced domain of vascular cell adhesion molecule 1
and an additional alpha 4 integrin counterreceptor on stimulated endothelium. J Exp Med
1992;175:1433–1442.
34 Weller PF, Rand TH, Goelz SE, Chi RG, Lobb
RR. Human eosinophil adherence to vascularendothelium mediated by binding to vascular celladhesion molecule 1 and endothelial leukocyte
adhesion molecule 1. Proc Natl Acad Sci 1991;
88:7430–7433.

molecule-1 and intercellular adhesion molecule-1
of eosinophils from patients with asthma. J
Allergy Clin Immunol 1995;96:941–950.
14 Jaffe EA, Hoyer LW, Nachman RL. Synthesis of
von Willebrand factor by cultured human endothelial cells. Proc Natl Acad Sci 1974;71:1901–
1909.
15 Johnson AR. Human pulmonary endothelial
cells in culture: activities of cells from arteries
and cells from veins. J Clin Invest 1980;65:841–
850.
16 Knowles DM, Tolidjian B, Marboe C, D'Agati,
Grimes M, Chess L. Monoclonal anti-humanmonocyte antibodies OKM1 and OKM5 possess
distinctive tissue distributions including differential reactivity with vascular endothelium. J
Immunol 1984;132:2170–2173.
17 Libby P, Sukhova G, Lee RT, Galis ZS. Cytokines regulate vascular functions related to stability of the atherosclerotic plaque. J Cardiovasc
Pharmacol 1995;25:9–12.
18 Lou J, Grou GE. TNF in vascular pathology: theimportance of platelet-endothelium interactions.
Res Immunol 1993;144:355–363.
19 Meerschaert J, Furie MB. The adhesion molecules used by monocytes for migration acrossendothelium include CD11a/CD18, CD11b/
CD18 and VLA-4 on monocytes and ICAM-1,
VCAM-1 and other ligands on endothelium. J
Immunol 995;154:4099–4112.
20 Meyrick B, Hoover R, Jones MR, Berry LC,
Brigham KL. In vitro effects of endotoxin on
bovine and sheep lung microvascular and pulmonary artery endothelial cells. J Cell Physiol 1989;
138:165–174.
21 Meyrick B, Berry LC, Christman BW. Response ofcultured human pulmonary artery endothelial cells
to endotoxin. Am J Physiol 1995;268:239–244.
22 Ohkawa Y, Yamauchi K. Expression of adhesion
molecules in bronchial asthma. Nippon Kyobu
Shikkan Gakkai Zasshi 1993;31:125–131 (inJapanese).
23 Raghu G, Striker LJ, Striker GE. Lipopolysaccharide-mediated injury of cultured humanglomerular endothelial cells. Clin Immunol
Immunopathol 1986;38:275–281.
24 Schwartz BR, Wayner EA, Carlos TM, Ochs HD,
Carter WG, Harlan JM. Identification of adherence molecules modulating adhesion of CD11/
CD18-deficient lymphoblastoid cell to cultured(Received December 20, 1996, Accepted January 10, 1997)30