Structural and functional characteristics of lung macro - Penn Medicine

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Microvascular Research 67 (2004) 139–151

Structural and functional characteristics of lung macro- and

microvascular endothelial cell phenotypes

Judy King,a Tray Hamil,b Judy Creighton,b Songwei Wu,b Priya Bhat,b

Freda McDonald,a and Troy Stevensb,*

aDepartment of Pathology, Center for Lung Biology, The University of South Alabama College of Medicine, Mobile, AL 36617, USAbDepartment of Pharmacology, Center for Lung Biology, The University of South Alabama College of Medicine, Mobile, AL 36688, USA

Received 11 September 2003

Abstract

Lung macro- and microvascular endothelial cells exhibit unique functional attributes, including signal transduction and barrier properties.

We therefore sought to identify structural and functional features of endothelial cells that discriminate their phenotypes in the fully

differentiated lung. Rat lung macro- (PAEC) and microvascular (PMVEC) endothelial cells each exhibited expression of typical markers.

Screening for reactivity with nine different lectins revealed that Glycine max and Griffonia (Bandeiraea) simplicifolia preferentially bound

microvascular endothelia whereas Helix pomatia preferentially bound macrovascular endothelia. Apposition between the apical

plasmalemma and endoplasmic reticulum was closer in PAECs (8 nm) than in PMVECs (87 nm), implicating this coupling distance in

the larger store operated calcium entry responses observed in macrovascular cells. PMVECs exhibited a faster growth rate than did PAECs

and, once a growth program was initiated by serum, PMVECs sustained growth in the absence of serum. Thus, PAECs and PMVECs differ in

their structure and function, even under similar environmental conditions.

D 2004 Elsevier Inc. All rights reserved.

Keywords: Endoplasmic reticulum; Calcium; Store-operated calcium entry; Proliferation; Lectins

Introduction

Although endothelium lines blood vessels throughout the

circulation, it exhibits highly specialized functions in dif-

ferent vascular sites. In the systemic circulation, permeabil-

ity edema is prominent at post-capillary venules (Thurston

et al., 2000). White blood cell recruitment to sites of

inflammation occurs at high endothelial venules (Cavender,

1990; Colditz, 1985) and, while the blood brain barrier

consists of endothelium with tight cell–cell junctions (Gloor

et al., 2001) that are highly restrictive, both renal glomerular

(Stan et al., 1999) and liver sinusoidal endothelium (Gri-

sham et al., 1975) possess fenestrations that are highly

permeable. It is clear that these distinct endothelial cell

characteristics are at least partly directed by environmental

cues (Stevens et al., 2001).

0026-2862/$ - see front matter D 2004 Elsevier Inc. All rights reserved.

doi:10.1016/j.mvr.2003.11.006

* Corresponding author. Fax: +1-251-460-7452.

E-mail address: [email protected] (T. Stevens).

The embryological origin of endothelial cells may also

contribute to their site-specific function (Stevens et al.,

2001). Studies in the developing lung suggest two distinct

processes form the circulation (deMello and Reid, 2000;

deMello et al., 1997; Hall et al., 2000; Schachtner et al.,

2000; Schwarz et al., 2000), including angiogenesis of

large vessels and vasculogenesis of small vessels. deMello

et al. (1997) used a casting technique to temporally

illustrate vascular tube formation. The earliest formed

vascular structures were observed at embryonic day 14

(E14) in the developing mouse lung. These structures

progressively branched from large vessels at sharp angles,

consistent with angiogenesis, but did not form a contigu-

ous vessel. The parallel growth of blood lakes/islands that

were filled with precursor cells of hematopoietic origin

was observed by transmission electron microscopy until, at

E15, a fusion between angiogenic sprouts and vasculo-

genic blood islands could be resolved using the vascular

casting technique. This issue has also been addressed by

assessing the temporal expression pattern of endothelial

J. King et al. / Microvascular Research 67 (2004) 139–151140

cell markers in developing lung (Schachtner et al., 2000).

Endothelial cells of both large(r) and small vessels express

the VEGF receptor Flk-1 during development, which has

been interpreted to suggest vessels larger than originally

suspected may originate from vasculogenesis. Thus,

while this issue is not completely understood, in the

simplest form, it appears that endothelial cells in large

and small blood vessels are likely to arise from different

progenitors.

Functional studies in in vitro models illustrate that lung

microvascular endothelial cells possess a more restrictive

barrier than their macrovascular counterparts (Chetham et

al., 1999; Kelly et al., 1998; Moore et al., 1998b), and

exhibit unique signaling responses to similar agonists

(Chetham et al., 1999; Kelly et al., 1998; Moore et al.,

1998a; Stevens et al., 1997, 1999, 2001). Distinct site-

specific vascular responses are observed in the intact lung

(Chetham et al., 1999; Khimenko and Taylor, 1999; Qiao

and Bhattacharya, 1991). The lung’s microcirculation is

more restrictive to protein and water flux than is the

macrocirculation (Parker and Yoshikawa, 2002). In con-

trast, macrovascular endothelial cells express more eNOS

(Stevens, unpublished) and generate more nitric oxide (Al-

Mehdi, unpublished) than do microvascular endothelial

cells. Large and small pulmonary vessels appear to exhibit

unique growth or survival properties. Indeed, the lung’s

microcirculation exhibits significantly more plasticity than

previously appreciated (Massaro and Massaro, 1997, 2000,

2001, 2002; Massaro et al., 2000). Emphysema-like lesions

are associated with a decrease in alveolar and capillary

(e.g., microvascular endothelial cell) density, a portion of

which can be rescued by retinoic acid. These findings are

generally compatible with evidence that alveolar cells and

microvascular endothelial cells uniquely regulate one

another’s function, partly dependent upon vascular endo-

thelial cell growth factor (VEGF) signaling to orchestrate

capillary development along the basement membrane of

airway epithelium (Acarregui et al., 1999; Dumont et al.,

1995; Gebb and Shannon, 2000; Lassus et al., 2001;

Shalaby et al., 1997). VEGF stimulates small vessel

formation and microvascular endothelial cell survival.

The VEGF receptor Flk-1 null mice die because blood

islands are disorganized and microvessels do not form

(Shalaby et al., 1995). In the fully developed lung inhibi-

tion of VEGF signaling reduces alveolar septation (as in

emphysema) (Kasahara et al., 2000) and, in combination

with hypoxia, generates microvascular (c100 Am) plexi-

genic lesions (Taraseviciene-Stewart et al., 2001). Thus,

lung endothelial cell origin may be an important determi-

nant of cell phenotype and function. To further determine

the unique attributes of lung macro- and microvascular

endothelial cells, we undertook studies to examine whether

pulmonary artery (PAEC) and microvascular (PMVEC)

endothelial cells isolated from the fully differentiated organ

exhibit distinct structure and function, even under similar

environmental conditions.

Methods

Isolation and culture of rat lung endothelial cells

Isolation and culture of rat main pulmonary artery

endothelial cells (PAECs)

Main pulmonary arteries were isolated as previously

described (Creighton et al., 2003; Stevens et al., 1999).

Briefly, 300–400 g Sprague–Dawley rats were euthanized

by an intraperitoneal injection of 50 mg of pentobarbital

sodium (Nembutal, Abbott Laboratories, Chicago, IL).

The heart and lungs were excised en bloc after sternot-

omy and the mainstem pulmonary artery and two vessel

generations were isolated and removed. The artery was

inverted and the intimal lining was carefully scraped

using a scalpel. Harvested cells were then placed into

T25 flasks (Corning Inc., Corning, NY) containing

F12 Nutrient Mixture and Dulbecco’s modified eagle

medium (DMEM) mixture (1:1) supplemented with 10%

fetal bovine serum (FBS), 100 U/ml penicillin, and 100 Ag/ml

streptomycin (Gibco BRL, Grand Island, NY) and passed

up to 15 times. The endothelial cell phenotype was

confirmed by acetylated LDL uptake, Factor VIII-Rag

immunocytochemical staining, and the absence of immu-

nostaining with smooth muscle cell a-actin antibodies.

Pulmonary microvascular endothelial cells (PMVECs)

PMVECs were isolated and cultured using a modified

method described by Stevens et al., 1999 (Creighton et

al., 2003). Male Sprague–Dawley rats (300–400 g) were

euthanized by intraperitoneal injection of 50 mg of

pentobarbital sodium (Nembutal, Abbott Laboratories).

After sternotomy, the heart and lungs were removed en

bloc and placed in a DMEM (Dulbecco’s Modified Eagle

Medium, Gibco BRL) bath containing 90 Ag/ml penicillin

and streptromycin. Thin strips were removed from the

lung periphery adjacent to the pleural surface, finely

minced, and transferred with 2–3 ml DMEM to a 15-

ml conical tube containing 3-ml digestion solution. [0.5 g

BSA, 10,000 units type 2 collagenase (Worthington

Biochemical Co, Lakewood, NJ), and cmf-PBS (Gibco

BRL) to make 10 ml total volume]. The digestion

mixture was allowed to incubate at 37jC for 15 min

before pouring through an 80-mesh sieve into a sterile

200-ml beaker. An additional 5 ml of normal medium

[10% FBS (Fetal Bovine Serum, Hyclone, Logan, UT)

with 30 Ag/ml penicillin and streptromycin in DMEM]

was used to wash the sieve. The isolation mixture was

transferred to a 15 ml conical tube and centrifuged at

300 � g for 5 min, the medium aspirated, and the cells

resuspended with 5 ml complete medium [1 part micro-

vascular conditioned medium: three parts incomplete

medium (80% RPMI 1640, 20% FBS, 12.3 units/ml

Heparin (Elkins-Sinn, Cherry Hill, NJ), and 6.7 Ag/ml

Endogro (Vec Technologies, Rensselaer, NY) with 30 Ag/ml penicillin and streptomycin]. Centrifugation/aspiration

J. King et al. / Microvascular Re

was repeated, the cells resuspended in 2–3 ml complete

medium and allowed to incubate at 37jC for 30 min

before being placed drop wise onto 35-mm culture

dishes. After 1 h at 37jC with 5% CO2, 3 ml of

complete medium was added. The dishes were checked

daily for contaminating cells that were removed by

scraping and aspiration. Endothelial cell colonies were

isolated with cloning rings, trypsinized, re-suspended in

100 Al complete medium and placed as a drop in the

center of a T-25 flask. The cells were allowed to attach

(1 h at 37jC with 5% CO2) before the addition of 5 ml

complete medium. Cultures were characterized using

SEM, uptake of 1,1V-dioctadecyl-3, 3,3V, 3V-tetramethylin-

docarbocyanine-labeled low-density lipoprotein (DiI-acet-

ylated LDL), a lectin-binding panel (see below), and were

routinely passaged by scraping.

Histochemical staining

Rat lung slices were deparaffinized by placing them in an

oven at 60jC for 10 min. They were rinsed with xylene twice

for 5 min, rehydrated with sequential alcohol washes from

100% to 30%, and placed in water for 5 min. After a 15-min

incubation in phosphate buffered saline (PBS) with 0.05%

Tween 20, the FITC-labeled lectins (Sigma, St. Louis,

MO) were applied at a 1:1000 dilution, and incubated

at room temperature for 1 h in the dark. The slides

were rinsed with PBS twice for 10 min and mounted

with fluorescent mounting medium (DAKO, Carpinteria,

CA). Epifluorescent and confocal fluorescent micro-

scopes were used to view the slides.

Table 1

List of lectins, plant or animal sources, nominal specificities, and staining intensity

strong staining. Staining intensity was approximately the same in all lectins scree

Lectin Source Nominal specificity

Arachis hypogea PNA peanut lactose > h-D-galactoseCaragana arborescens CAA pea tree GalNAc

Lens culinaris LcH lentil a-mannose > a-glucos

aGlcNAc

Lycopersicon esculentum LEA tomato GlcNAch(1,4)GlcNAcRicinus communis RCA120 castor bean lactose > galactose

Ulex europaeus UEA-I gorse, furze a-L-fucose GlcNAch(1Glycine max SBA soybean terminal a and h-GalN

Griffonia simplicifolia GS-I N/A a-galactose > a-GalNA

Helix pomatia HPA edible snail a-GalNAc > h-GalNA

As indicated, Helix pomatia preferentially binds PAECs, while Glycine max and

Cell sorting

PAECs and PMVECs were counted using a Coulter

counter (Coulter Corporation, Hialeah, FL), and 4–6 � 105

cells were resuspended in 0.5 ml PBS in flow cytometric

tubes. EGTA (1 mM) was added to facilitate single cell

suspensions, and cells were periodically triturated. FITC-

conjugated lectins (Sigma) were added to the tubes at

increasing concentrations (1:500 to 1:10). After a 20-min

incubation in the dark, the cells were analyzed using the FL-1

channel (FITC) of a flow cytometer. Blocking was performed

with sugars at concentrations recommended by Sigma for

each lectin.

Agglutination experiment

PAECs and PMVECs were grown to confluence on 35-

mm dishes. Lectins were diluted 1:1000, added to the 35

mm dishes and incubated for 15 min. The cells were

trypsinized and triturated to assure single cell suspensions,

then resuspended in PBS. Cells were centrifuged and the

cell pellets were resuspended in PBS. A small drop from

each tube was applied to glass microscope slides and viewed

under a microscope.

Transmission electron microscopy

PAECs and PMVECs were seeded (PMVEC density

2.7 � 105; PAEC density 6.7 � 105) onto 0.4 Ampolycarbonate membranes (Nunc, Naperville, IL) for trans-

mission electron microscopy, and grown for 4 days to

search 67 (2004) 139–151 141

in PAECs and PMVECs. + = weak staining; ++ = moderate staining; +++ =

ned, except for Glycine max, Griffonia simplicifolia, and Helix pomatia

PAEC PMVEC Reference

++ ++

++ ++

e, +++ +++

+++ +++

+++ +++

,4)GlcNAc ++ ++

Ac > a and h-Gal + +++ (Alvarez-Fernandez and

Carretero-Albinana, 1990;

Honda et al., 1986; Kawai

et al., 1988; Mazzuca et al.,

1982; Spicer et al., 1983)

c + +++ (Bankston et al., 1991;

Del Vecchio et al., 1992;

Gumkowski et al., 1987;

Magee et al., 1994;

Schnitzer et al., 1994;

Tsokos et al., 2002)

c +++ + (Palmer and Bale, 1987;

Taatjes et al., 1990;

Yi et al., 2001)

Griffonia simplicifolia preferentially bind PMVECs.


confluence. Cultures were fixed in 3% glutaraldehyde in

cacodylate buffer, rinsed in cacodylate buffer, and post-fixed

for 30 min with 1% osmium tetroxide. The cells were

dehydrated using a graded alcohol series. Portions of the

filters were embedded in PolyBed 812 Resin (Polysciences

Inc., Warrington, PA). Thick sections (1 Am) were cut with

glass knives and stained with 1% toluidine blue. Thin

sections (80 nm) were cut with a diamond knife and then

stained with uranyl acetate and Reynold’s lead citrate.

Cultures were examined and photographed using a Philips

CM 100 transmission electron microscope (FEI Company,

Hillsboro, OR). Measurements were made from the micro-

graphs. Measurements of endothelial cell length were made

only if the nucleus of the cell was in the section.

Portions of the pulmonary artery and the lung paren-

chyma were fixed in 3% glutaraldehyde in cacodylate

buffer by immersion in fixative or vascular perfusion.

The specimens were rinsed in cacodylate buffer, post-fixed

for 1 h with 1% osmium tetroxide, and then prepared as

described above. Measurements were made from the

micrographs. Measurements of endothelial cell length were

made only if the nucleus of the cell was in the section.

Cytosolic calcium

Endothelial cells were seeded onto 25-mm circle mi-

croscope glass coverslips (Fisher Scientific, Pittsburgh,

PA) and grown to confluence. Cytosolic Ca2+ was esti-

mated with the Ca2+-sensitive fluorophore fura 2/acetox-

ymethylester (Molecular Probes, Eugene, OR) according

to methods previously described. Calculations of free

Fig. 1. Endothelial cell phenotypes in the intact lung can be discriminated by le

(arrows) represents staining of endothelial cells with Helix pomatia. No staining

microscope). Staining with Glycine max and Griffonia simplicifolia is absent in the

confocal microscope).

[Ca2+]i are routinely made using modifications of the

formula described by Grynkiewicz et al. (1985).

Cell growth

Endothelial cells were seeded at 1 � 105 cells per well

in six well plates at n = 3. Cells were seeded in normal

media containing DMEM, 10% FBS (or as otherwise

noted), and 1 � pen/strep. Every 24 h for 6 days after

the seeding date, cells were photographed, resuspended

using trypsin, and counted using a Coulter counter.

Data analysis

Numerical data are reported as mean F SEM. One-

way ANOVA was used to evaluate differences between

experimental groups, with a Student Newman–Keuls

post hoc test as appropriate. Significance was considered

P < 0.05.

Results

Lectin binding to lung macro- and microvascular

endothelial cells

Lectin binding has previously been utilized as an effective

method of discriminating between macro- and microvascular

endothelial cells (Abdi et al., 1995; Del Vecchio et al., 1992;

Fischer et al., 2000; Gumkowski et al., 1987; Lotan et al.,

1994; Magee et al., 1994; Norgard-Sumnicht et al., 1995;

ctin binding. Green fluorescent stain in the lumen of the pulmonary artery

was observed with Helix pomatia in the peripheral lung (40�, fluorescent

pulmonary artery but is present in peripheral lung capillaries (arrows; 67�,

Fig. 2. Endothelial cell phenotypes in vitro can be discriminated by lectin binding. Increased staining intensity is demonstrated by a right shift in the

fluorescence intensity of sorted cells. Helix pomatia exhibits a more intense fluorescence in PAECs than in PMVECs. Glycine max and Griffonia simplicifolia

exhibit a more intense fluorescence in PMVECs when compared to PAECs. Inset pictures show control cells without lectin treatment (yellow), and cells treated

with a-GalNAc to block Helix pomatia, h-GalNAc to block Glycine max, and a-galactose to block Griffonia simplicifolia (blue). Sugars were added accordingto Sigma recommendations. In blocking studies, cells were incubated for 30 min with the blocking sugar before a 15-min incubation with the lectin.

J. King et al. / Microvascular Research 67 (2004) 139–151 143

Schnitzer et al., 1994). Nine different lectins were therefore

screened for binding to the rat pulmonary artery and micro-

vascular endothelial cell surface (Table 1). Of the nine lectins

Fig. 3. Lectin-induced agglutination discriminates endothelial cell phenotypes in vi

Griffonia simplicifolia selectively agglutinate PMVECs. Cells were trypsinized an

presence of lectins. Arrows indicate cell clumps. Pictures are representative of fiv

examined, six did not distinguish between macro- and

microvascular cell types, while three demonstrated a prefer-

ential binding pattern. In vivo, FITC-labeled Helix pomatia

tro. Helix pomatia selectively agglutinates PAECs whereas Glycine max and

d triturated in to single cell suspensions, then allowed to agglutinate in the

e separate experiments.


staining was observed selectively in macrovascular endothe-

lia. The green fluorescent stain can be seen lining the lumen

of the pulmonary artery (Fig. 1), but is absent in the

peripheral lung. FITC-labeled Glycine max and Griffonia

simplicifolia stained microvascular cells preferentially.

Green fluorescent stain can be seen lining the lumen of

capillaries in peripheral lung but is absent in the pulmonary

artery (Fig. 1).

Fluorescence-activated cell sorting (FACS) was used to

confirm that cells isolated and cultured in vitro retained their

in vivo phenotype. H. pomatia staining was prominent in

macrovascular cells, while G. max and G. simplicifolia

staining was prominent in microvascular cells (Fig. 2).

Controls (e.g., cells without FITC-labeled lectin) did not

fluoresce and specific sugars in competitive binding studies

prevented staining (inset, Fig. 2). To confirm specificity of

these lectins for their respective endothelial cell type,

agglutination studies were performed in which lectin-treated

endothelial cells were trypsinized and dispersed. As is seen

in Fig. 3, H. pomatia selectively agglutinated PAECs in the

presence of trypsin whereas G. max and G. simplicifolia

selectively agglutinated PMVECs in the presence of trypsin.

These findings suggest that rat lung endothelial cells possess

similar surface sugars that can be distinguished from lectin

binding, both in vivo and in vitro.

Lung endothelial cell morphology

Few studies have documented the morphological char-

acteristics of rat lung PAECs and PMVECs, particularly

under identical culture conditions. We therefore examined

ultrastructural characteristics of these cell types in vitro

and in situ. Both PAECs and PMVECs in culture exhibited

round to oval nuclei, few mitochondria, rough endoplasmic

reticulum, junctions between cells, and surface projections

(Fig. 4). Vesicles consistent with caveolae (50–80 nm)

were present in both cell types, and vesicles consistent

with clathrin-coated pits were observed in PAECs (100–

150 nm). Groups of filaments were observed along the

basal membrane of both cell types, however, they were

more prominent in the PAECs (data not shown). By

transmission electron microscopy PAECs measured 8.8–

38.3 Am in diameter and 2.5–7.1 Am in maximum height.

By transmission electron microscopy PMVECs measured

9.0–37.5 Am in diameter and 2.9–7.1 Am in maximum

height.

Similar characteristics were observed in situ (Fig. 4).

Native PAECs exhibited numerous vesicles and occasional

Weibel-Palade bodies along with some mitochondria and

Fig. 4. Endothelial cell morphology in vitro. Ultrastructural assessment of

PAECs [panel A] and PMVECs [panel B] in culture-demonstrated typical

appearance of mitochondria (M), rough endoplasmic reticulum (RER), and

nucleus (N). Caveolae- or clathrin-coated pits (C) were observed.

Organelles were similarly observed in perfusion fixed lung pulmonary

artery [panel C] and capillary [panel D]. F denotes filter; L denotes lumen.

Fig. 5. RER-plasmalemma coupling distinguishes endothelial cell phenotypes. (A) Typical cytosolic calcium response to activation of store operated calcium entry

using thapsigargin (1 AM) demonstrates a lower response in PMVECs than in PAECs. (B) Transmission electron micrograph reveals that RER can be observed

immediately adjacent to the apical cell membrane in PAECs. (C) In addition, RER can be observed immediately adjacent to a vesicle consistent in size with a

clathrin-coated pit (e.g.,c100 nm). RER were also observed nearby caveolae-like structures (50–80 nm) in PAECs. RER do not similarly approach the apical

plasma membrane (D) or vesicles (E) in PMVECs. (F) Summary data reveal the RER-apical membrane distance is approximately 100 nm in PAECs (n = 44

specimens) and 250 nm in PMVECs (n = 50 specimens). *Denotes significantly different from PAEC.


Fig. 7. PMVECs possess a greater proliferative index that do PAECs. (A)

Serum-stimulated (10%) cell growth was observed over a 6-day period.

After a 2-day lag phase both PAECs and PMVECs exhibited log phase

growth, although PMVECs grew faster than did PAECs. (B) Serum-

restriction (0.1%) inhibited the growth of both cell types. (C) However,

serum stimulation during the lag phase was sufficient to initiate PMVEC

growth, even when cells were serum-deprived during the log phase. Such

treatment inhibited the growth of PAECs. *Denotes different from PAECs.

(D) Phase contrast images illustrate that PAECs and PMVECs grow at

different rates. Each cell type was seeded in 6-well plates at 105 cells/well,

and grown in the presence of 10% serum. PMVECs reached confluence on

day 4, whereas PAECs reached confluence on day 6.

Fig. 6. Cell junctions differ between PAECs (A) and PMVECs (B) in vitro.

PMVECs occasionally exhibited cytoplasmic processes (CP) not found in

PAECs. RER was found nearby cell borders, although always closer to the

membrane in PAECs than in PMVECs. ‘‘A’’ denotes apical cell side. ‘‘F’’

denotes filter. ‘‘N’’ denotes nucleus.


RER. The RERwas as close as 58 nm from the apical surface.

Projections extend from the apical surface of the native

pulmonary artery endothelial cells, with more numerous

projections at the junctions between cells. Many of the

cellular projections are thin. In situ PAECs measured 11.0–

29.7 Am in diameter and 2.1–7.2 Am in maximum height.

Maximal cell height was in the area of the nucleus, where the

cell extended into the lumen. Nuclei were primarily oval

shaped. The peripheral parts of the cell were thinner, mea-

suring as little as 62.5 nm in height when perfused fixed

vessels were examined. In situ capillaries of the lung paren-

chyma contained two to three endothelial cells in a vascular

cross-section. The cell’s periphery was very thin, measuring

only 15.8 nm in some areas; organelles were absent in this

thin periphery. Numerous vesicles were present throughout

the cells, with the exception of the thinnest regions. Rare

profiles of RER were present, located primarily in the thicker

portions of the cell. RER was seen as close as 89 nm from the

apical surface. A few scattered mitochondria were present.

Apical (luminal) surfaces of the endothelial cells were often

flattened, although scattered projections were observed. The

capillary endothelial cells measured 10.3–26.9 Am in diam-

eter and 2.6–7.2 Am in maximum height. Because of the

Fig. 7 (continued).


small caliber of the capillaries, the nucleus caused a distinct

bulging into the lumen.

RER-membrane coupling: relevance to calcium signaling

We have previously observed that the thapsigargin-in-

duced store operated calcium entry response is lower in

PMVECs than it is in PAECs (Kelly et al., 1998; Moore

et al., 1998b; Stevens et al., 1997, 1999). Store-operated

calcium entry pathways are activated by depletion of cal-

cium in the endoplasmic reticulum (Putney, 1986). At

present, the signal(s) linking calcium store depletion to

activation of store-operated calcium entry is unclear, al-

though three separate models have been developed to


address putative coupling mechanism(s) (for review, see

(Parekh and Penner, 1997; Putney, 2001; Putney and

Ribeiro, 2000)). Both conformational coupling and secretion

coupling models implicate a ‘‘physical’’ relationship be-

tween the endoplasmic reticulum and plasmalemma in

activation of store-operated calcium entry channels. We

therefore examined the morphological distribution of endo-

plasmic reticulum in PAECs and PMVECs, to evaluate

whether physical coupling between the endoplasmic reticu-

lum and plasma membrane could provide a plausible expla-

nation for the decreased store operated calcium entry

response in PMVECs. RER was observed close to the

apical, lateral (junctions between cells), and basal cell

membranes in both cell types, although the pattern of

distribution differed significantly in PAECs and PMVECs.

Measurements taken from transmission electron micro-

graphs revealed that the RER was as close as 8 nm to the

apical cell membrane in PAECs and as close as 87 nm to the

apical cell membrane in PMVECs (Fig. 5). Summary data of

membrane associated organelles indicated that, on average,

RER is nearly 2.5-fold closer to the plasmalemma in PAECs

than in PMVECs. Together, these findings suggest that the

proximity of RER to the plasmalemma may contribute to the

differential calcium signaling responses seen in these cell

types.

PMVECs form a more restrictive macromolecular barrier

than do PAECs (Chetham et al., 1999; Kelly et al., 1998).

Since macromolecular flux occurs at least partly through

intercellular junctions, we examined sites of cell–cell con-

tact in PMVECs and PAECs (Fig. 6). Electron dense

structures were observed at cell–cell borders. In both cell

types, RER could be resolved near cell–cell borders nearby

electron dense structures that contribute to cell–cell adhe-

sion, although the RER was closer to cell–cell borders in

PAECs than in PMVECs. RER was seen as close as 13 nm

from the cell membrane between PAECs and as close as 77

nm from the cell membrane between PMVECs. RER was

observed as close as 14 nm from the basal cell membrane in

PAECs and 76 nm from the basal cell membrane in

PMVECs. Store-operated calcium entry channels have not

presently been resolved within cell junctions. However,

these findings suggest that activation of store-operated

calcium entry may provide a calcium source nearby sites

of cell adhesion.

Endothelial cell growth

To further characterize lung endothelial cells, we evalu-

ated the growth rates of PAECs and PMVECs. Trypsinized

cells were triturated to single cell suspensions and re-seeded

in the presence of 10% serum. Both PAECs and PMVECs

exhibited a characteristic 2-day lag phase followed by log

phase growth (Fig. 7). PMVECs grew faster than PAECs.

Growth of both cell types was inhibited when cells were

incubated with 0.1% serum for 5 days. However, when cells

were incubated with 10% serum during lag phase growth

and then switched to 0.1% serum, PMVECs grew almost

normally whereas PAEC growth was significantly inhibited.

Together, these findings suggest that PMVECs possess a

unique pro-proliferative phenotype that is not present in

PAECs.

Discussion

Our present studies were founded on the hypothesis

that PAEC and PMVEC phenotypes are distinct, in part

due to their epigenetic origin. If this hypothesis is true,

then the cells should retain distinct functions in vitro

when their environments are similar. We approached this

hypothesis using structure–function analyses, evaluating

morphological characteristics of the cells along with

functional endpoints.

Lectins are protein agglutinins isolated from various

plant and animal sources that have proven useful for

distinguishing between cell phenotypes(Abdi et al., 1995;

Del Vecchio et al., 1992; Fischer et al., 2000; Gumkowski et

al., 1987; Lotan et al., 1994; Magee et al., 1994; Norgard-

Sumnicht et al., 1995; Schnitzer et al., 1994; Symon and

Wardlaw, 1996). Lectins interact with cell surface carbohy-

drates and therefore cell-specific lectin binding provides

important information regarding glycocalyx characteristics.

As in earlier studies, G. simplicifolia in particular selectively

interacted with PMVECs in vivo and in vitro. This lectin

exhibits affinity for a-galactose, indicating the PMVEC

glycocalyx is enriched with an a-galactose containing

carbohydrate. In contrast to prior studies, which primarily

observed H. pomatia binding to alveolar cells, we found H.

pomatia interacted with PAECs with preference over

PMVECs in vivo and in vitro. H. pomatia exhibits affinity

for a- and h-N-acetyl-galactosamine, indicating the rat

PAEC glycocalyx is enriched with an a- and h-N-acetyl-galactosamine carbohydrate. Since the glycocalyx contrib-

utes to cell–cell recognition, evidence for a differential

‘‘structure’’ of the PAEC and PMVEC glycocalyx suggests

these cells function distinctly in response to inflammatory

stimuli. Indeed, selectins bind homing receptors on the

endothelial cell glycocalyx (Symon and Wardlaw, 1996),

and bacteria interact with the endothelial cell surface

through adhesions that bind the glycocalyx (Hoppe et al.,

1997). The contribution of such distinct surface carbohy-

drate structures to site-specific inflammatory responses will

be important to resolve.

PAECs and PMVECs possessed significant morpholog-

ical distinctions, both in the intact circulation and in

culture. Association between the apical plasmalemma and

endoplasmic reticulum is closer in PAECs than in

PMVECs, implicating this membrane to organelle coupling

in calcium-mediated signal transduction. The principal

mode of calcium entry in endothelial cells is through so-

called store operated calcium entry pathways (Moore et al.,

1998b; Nilius and Droogmans, 2001), where calcium store


depletion in the endoplasmic reticulum activates calcium

entry across the plasmalemma. Neither the mechanism of

membrane channel activation nor the molecular identity of

membrane channels is well understood. However, certain

mammalian transient receptor proteins (TRPC) contribute

subunits to store-operated calcium entry channels (Birn-

baumer et al., 1996; Freichel et al., 1999; Hofmann et al.,

2000). Endothelial cells express these channels (Brough et

al., 2001; Moore et al., 1998a; Wu et al., 2001) and, in

recent studies, TRPC1 (Rosado and Sage, 2000) and

TRPC3 (Birnbaumer et al., 2000; Boulay et al., 1999;

Kiselyov et al., 1999) have been immunoprecipitated with

inositol 1,4,5-trisphosphate receptors that reside the endo-

plasmic reticulum. These biochemical studies implicate

direct coupling between the plasmalemma and endoplasmic

reticulum in mechanism(s) underlying channel activation.

Our present finding that the plasmalemma and endoplasmic

reticulum are immediately adjacent in PAECs lends further

support for the necessity of direct coupling between the

membrane and organelle in channel activation. Indeed,

PAECs possess more prominent store operated calcium

entry pathways than do PMVECs (Kelly et al., 1998;

Stevens et al., 1997, 1999).

Direct activation of store-operated calcium entry using

thapsigargin induces reorganization of f-actin, myosin light

chain phosphorylation and rapid intercellular gap formation

in PAECs (Moore et al., 1998a, 2000; Norwood et al.,

2000), whereas PMVECs are resistant to this calcium-

mediated gap formation (Kelly et al., 1998). In our present

studies, close coupling was observed between the endoplas-

mic reticulum and plasma membrane between PAECs,

raising the possibility that store-operated calcium entry

channels are functionally localized to sites of cell–cell

adhesion. Impetus for this possibility comes from skeletal

muscle, where coupling between the transverse tubule and

sarcoplasmic reticulum is essential for calcium-mediated

contraction (Isenberg et al., 1996). At present, putative

store-operated calcium entry channels are known to be

enriched in caveolae (Lockwich et al., 2000) and have not

been localized to membrane borders between cells. In

PMVECs, endoplasmic reticulum was not closely associated

with membrane borders between cells, particularly in the

intact circulation. Reduced endoplasmic reticulum–mem-

brane coupling may contribute to the enhanced barrier

function of PMVECs.

PMVECs grew at a faster rate than did PAECs but,

remarkably, they exhibited a unique serum-stimulated

growth program. Indeed, whereas 0.1% serum growth

arrested PMVECs, incubation of PMVECs in 10% serum

during lag phase growth was sufficient to sustain rapid

proliferation in 0.1% serum during the log phase. Similar

serum exposure did not sustain PAEC growth. These find-

ings suggest PMVECs exhibit a unique growth program,

wherein activation of the paradigm can be sustained by

autocrine factors. Factors that mediate the autocrine growth

capacity of PMVECs will be essential to resolve.

In summary, PAECs and PMVECs differ structurally

and functionally—even when their environments are sim-

ilar. These results support the idea that a cell’s embryo-

logical (e.g., epigenetic) origin may impact its function

even in the fully differentiated organ (Stevens et al., 2001).

Appreciation for both the epigenetic and environmental

determinants of cell phenotype reveal important insight

into how site-specific function can be achieved. In future

studies, it will be important to consider the interplay

between cell origin and environmental cues in regulating

cell behavior.

Acknowledgments

We thank Dr. Ray Hester for his participation in this

work. Supported by HL66299 and HL60024 (T. Stevens).

References

Abdi, K., Kobzik, L., Li, X., Mentzer, S.J., 1995. Expression of membrane

glycoconjugates on sheep lung endothelium. Lab. Invest. 72, 445–452.

Acarregui, M.J., Penisten, S.T., Goss, K.L., Ramirez, K., Snyder, J.M.,

1999. Vascular endothelial growth factor gene expression in human

fetal lung in vitro. Am. J. Respir. Cell Mol. Biol. 20, 14–23.

Alvarez-Fernandez, E., Carretero-Albinana, L., 1990. Lectin histochemistry

of normal bronchopulmonary tissues and common forms of broncho-

genic carcinoma. Arch. Pathol. Lab. Med. 114, 475–481.

Bankston, P.W., Porter, G.A., Milici, A.J., Palade, G.E., 1991. Differential

and specific labeling of epithelial and vascular endothelial cells of the

rat lung by Lycopersicon esculentum and Griffonia simplicifolia I lec-

tins. Eur. J. Cell. Biol. 54, 187–195.

Birnbaumer, L., Zhu, X., Jiang, M., Boulay, G., Peyton, M., Vannier, B.,

Brown, D., Platano, D., Sadeghi, H., Stefani, E., Birnbaumer, M.,

1996. On the molecular basis and regulation of cellular capacitative

calcium entry: roles for Trp proteins. Proc. Natl. Acad. Sci. U. S. A.

93, 15195–15202.

Birnbaumer, L., Boulay, G., Brown, D., Jiang, M., Dietrich, A., Mikoshiba,

K., Zhu, X., Qin, N., 2000. Mechanism of capacitative Ca2+ entry

(CCE): interaction between IP3 receptor and TRP links the internal

calcium storage compartment to plasma membrane CCE channels. Re-

cent Prog. Horm. Res. 55, 127–161.

Boulay, G., Brown, D.M., Qin, N., Jiang, M., Dietrich, A., Zhu, M.X.,

Chen, Z., Birnbaumer, M., Mikoshiba, K., Birnbaumer, L., 1999. Mod-

ulation of Ca(2+) entry by polypeptides of the inositol 1,4, 5- trisphos-

phate receptor (IP3R) that bind transient receptor potential (TRP):

evidence for roles of TRP and IP3R in store depletion-activated

Ca(2+) entry. Proc. Natl. Acad. Sci. U. S. A. 96, 14955–14960.

Brough, G.H., Wu, S., Cioffi, D., Moore, T.M., Li, M., Dean, N., Stevens,

T., 2001. Contribution of endogenously expressed Trp1 to a Ca2+-se-

lective, store-operated Ca2+ entry pathway. FASEB J. 15, 1727–1738.

Cavender, D.E., 1990. Organ-specific and non-organ-specific lymphocyte

receptors for vascular endothelium. J. Invest. Dermatol. 94, 41S–48S.

Chetham, P.M., Babal, P., Bridges, J.P., Moore, T.M., Stevens, T., 1999.

Segmental regulation of pulmonary vascular permeability by store-op-

erated Ca2+ entry. Am. J. Physiol. 276, L41–L50.

Colditz, I.G., 1985. Margination and emigration of leucocytes. Surv. Synth.

Pathol. Res. 4, 44–68.

Creighton, J., Masada, N., Cooper, D.M.F., Stevens, T., 2003. Coordinate

regulation of membrane cAMP by calcium inhibited adenylyl cyclase

(type 6) and phosphodiesterase (type 4) activities. Am. J. Physiol. 284,

L100–L107.


Del Vecchio, P.J., Siflinger-Birnboim, A., Belloni, P.N., Holleran, L.A.,

Lum, H., Malik, A.B., 1992. Culture and characterization of pulmo-

nary microvascular endothelial cells. In Vitro Cell. Dev. Biol. 28A,

711–715.

deMello, D.E., Reid, L.M., 2000. Embryonic and early fetal development

of human lung vasculature and its functional implications. Pediatr. Dev.

Pathol. 3, 439–449.

deMello, D.E., Sawyer, D., Galvin, N., Reid, L.M., 1997. Early fetal

development of lung vasculature. Am. J. Respir. Cell Mol. Biol. 16,

568–581.

Dumont, D.J., Fong, G.H., Puri, M.C., Gradwohl, G., Alitalo, K., Breit-

man, M.L., 1995. Vascularization of the mouse embryo: a study of flk-

1, tek, tie, and vascular endothelial growth factor expression during

development. Dev. Dyn. 203, 80–92.

Fischer, E., Wagner, M., Bertsch, T., 2000. Cepaea hortensis agglutinin-I,

specific for oglycosidically linked sialic acids, selectively labels endo-

thelial cells of distinct vascular beds. Histochem. J. 32, 105–109.

Freichel, M., Schweig, U., Stauffenberger, S., Freise, D., Schorb, W.,

Flockerzi, V., 1999. Storeoperated cation channels in the heart

and cells of the cardiovascular system. Cell. Physiol. Biochem. 9,

270–283.

Gebb, S.A., Shannon, J.M., 2000. Tissue interactions mediate early events

in pulmonary vasculogenesis. Dev. Dyn. 217, 159–169.

Gloor, S.M., Wachtel, M., Bolliger, M.F., Ishihara, H., Landmann, R., Frei,

K., 2001. Molecular and cellular permeability control at the blood–

brain barrier. Brain Res. Brain Res. Rev. 36, 258–264.

Grisham, J.W., Nopanitaya, W., Compagno, J., Nagel, A.E., 1975. Scan-

ning electron microscopy of normal rat liver: the surface structure of its

cells and tissue components. Am. J. Anat. 144, 295–321.

Grynkiewicz, G., Poenie, M., Tsien, R.Y., 1985. A new generation of Ca2+

indicators with greatly improved fluorescence properties. J. Biol. Chem.

260, 3440–3450.

Gumkowski, F., Kaminska, G., Kaminski, M., Morrissey, L.W., Auerbach,

R., 1987. Heterogeneity of mouse vascular endothelium. In vitro studies

of lymphatic, large blood vessel and microvascular endothelial cells.

Blood Vessels 24, 11–23.

Hall, S.M., Hislop, A.A., Pierce, C.M., Haworth, S.G., 2000. Prenatal

origins of human intrapulmonary arteries: formation and smooth muscle

maturation. Am. J. Respir. Cell Mol. Biol. 23, 194–203.

Hofmann, T., Schaefer, M., Schultz, G., Gudermann, T., 2000. Transient

receptor potential channels as molecular substrates of receptor-mediated

cation entry. J. Mol. Med. 78, 14–25.

Honda, T., Ono, K., Katsuyama, T., Nakayama, J., Akamatsu, T., 1986.

Mucosubstance histochemistry of the normal mucosa and epithelial

neoplasms of the lung. Acta Pathol. Jpn. 36, 665–680.

Hoppe, H.C., de Wet, B.J., Cywes, C., Daffe, M., Ehlers, M.R., 1997.

Identification of phosphatidylinositol mannoside as a mycobacterial

adhesin mediating both direct and opsonic binding to nonphagocytic

mammalian cells. Infect. Immun. 65, 3896–3905.

Isenberg, G., Etter, E.F., Wendt-Gallitelli, M.F., Schiefer, A., Carrington,

W.A., Tuft, R.A., Fay, F.S., 1996. Intrasarcomere [Ca2+] gradients in

ventricular myocytes revealed by high speed digital imaging micros-

copy. Proc. Natl. Acad. Sci. U. S. A. 93, 5413–5418.

Kasahara, Y., Tuder, R.M., Taraseviciene-Stewart, L., Le Cras, T.D.,

Abman, S., Hirth, P.K., Waltenberger, J., Voelkel, N.F., 2000. Inhibition

of VEGF receptors causes lung cell apoptosis and emphysema. J. Clin.

Invest. 106, 1311–1319.

Kawai, T., Greenberg, S.D., Titus, J.L., 1988. Lectin histochemistry

of normal lung and pulmonary adenocarcinoma. Mod. Path. 1,

485–492.

Kelly, J.J., Moore, T.M., Babal, P., Diwan, A.H., Stevens, T., Thompson,

W.J., 1998. Pulmonary microvascular and macrovascular endothelial

cells: differential regulation of Ca2+ and permeability. Am. J. Physiol.

274, L810–L819.

Khimenko, P.L., Taylor, A.E., 1999. Segmental microvascular permeabil-

ity in ischemia– reperfusion injury in rat lung. Am. J. Physiol. 276,

L958–L960.

Kiselyov, K., Mignery, G.A., Zhu, M.X., Muallem, S., 1999. The N-termi-

nal domain of the IP3 receptor gates store-operated hTrp3 channels.

Mol. Cell 4, 423–429.

Lassus, P., Turanlahti, M., Heikkila, P., Andersson, L.C., Nupponen, I.,

Sarnesto, A., Andersson, S., 2001. Pulmonary vascular endothelial

growth factor and Flt-1 in fetuses, in acute and chronic lung disease,

and in persistent pulmonary hypertension of the newborn. Am. J.

Respir. Crit. Care Med. 164, 1981–1987.

Lockwich, T.P., Liu, X., Singh, B.B., Jadlowiec, J., Weiland, S., Ambud-

kar, I.S., 2000. Assembly of Trp1 in a signaling complex associated

with caveolin-scaffolding lipid raft domains. J. Biol. Chem. 275,

11934–11942.

Lotan, R., Belloni, P.N., Tressler, R.J., Lotan, D., Xu, X.C., Nicolson, G.L.,

1994. Expression of galectins on microvessel endothelial cells and their

involvement in tumour cell adhesion. Glycoconj. J. 11, 462–468.

Magee, J.C., Stone, A.E., Oldham, K.T., Guice, K.S., 1994. Isolation,

culture, and characterization of rat lung microvascular endothelial cells.

Am. J. Physiol. 267, L433–L441.

Massaro, G.D., Massaro, D., 1997. Retinoic acid treatment abrogates elas-

tase-induced pulmonary emphysema in rats. Nat. Med. 3, 675–677.

Massaro, G.D., Massaro, D., 2000. Retinoic acid treatment partially

rescues failed septation in rats and in mice. Am. J. Physiol. 278,

L955–L960.

Massaro, D., Massaro, G.D., 2001. Pulmonary alveolus formation: critical

period, retinoid regulation and plasticity. Novartis Found. Symp. 234,

229–236.

Massaro, D., Massaro, G.D., 2002. Invited Review: Pulmonary alveoli:

formation, the ‘‘call for oxygen,’’ and other regulators. Am. J. Physiol.

282, L345–L358.

Massaro, G.D., Massaro, D., Chan, W.Y., Clerch, L.B., Ghyselinck, N.,

Chambon, P., Chandraratna, R.A., 2000. Retinoic acid receptor-beta: an

endogenous inhibitor of the perinatal formation of pulmonary alveoli.

Physiol. Genomics 4, 51–57.

Mazzuca, M., Lhermitte, M., Lafitte, J.J., Roussel, P., 1982. Use of lectins

for detection of glycoconjugates in the glandular cells of the human

bronchial mucosa. J. Histochem. Cytochem. 30, 956–966.

Moore, T.M., Brough, G.H., Babal, P., Kelly, J.J., Li, M., Stevens, T.,

1998a. Store-operated calcium entry promotes shape change in pul-

monary endothelial cells expressing Trp1. Am. J. Physiol. 275,

L574–L582.

Moore, T.M., Chetham, P.M., Kelly, J.J., Stevens, T., 1998b. Signal trans-

duction and regulation of lung endothelial cell permeability. Interaction

between calcium and cAMP. Am. J. Physiol. 275, L203–L222.

Moore, T.M., Norwood, N.R., Creighton, J.R., Babal, P., Brough, G.H.,

Shasby, D.M., Stevens, T., 2000. Receptor-dependent activation of

store-operated calcium entry increases endothelial cell permeability.

Am. J. Physiol. 279, L691–L698.

Nilius, B., Droogmans, G., 2001. Ion channels and their functional role in

vascular endothelium. Physiol. Rev. 81, 1415–1459.

Norgard-Sumnicht, K.E., Roux, L., Toomre, D.K., Manzi, A., Freeze, H.H.,

Varki, A., 1995. Unusual anionic N-linked oligosaccharides from bo-

vine lung. J. Biol. Chem. 270, 27634–27645.

Norwood, N., Moore, T.M., Dean, D.A., Bhattacharjee, R., Li, M., Stevens,

T., 2000. Store-operated calcium entry and increased endothelial cell

permeability. Am. J. Physiol. 279, L815–L824.

Palmer, K.C., Bale, L.A., 1987. Ultrastructural localization of Helix poma-

tia lectin-binding sites in mouse lung elastic fibers. Histochemistry 88,

91–95.

Parekh, A.B., Penner, R., 1997. Store depletion and calcium influx. Phys-

iol. Rev. 77, 901–930.

Parker, J.C., Yoshikawa, S., 2002. Vascular segmental permeabilities at

high peak inflation pressure in isolated rat lungs. Am. J. Physiol.

283, L1203–L1209.

Putney Jr., J.W., 1986. A model for receptor-regulated calcium entry. Cell

Calcium 7, 1–12.

Putne Jr., J.W., 2001. Cell biology. Channelling calcium. Nature 410,

648–649.


Putney Jr., J.W., Ribeiro, C.M., 2000. Signaling pathways between the

plasma membrane and endoplasmic reticulum calcium stores. Cell.

Mol. Life Sci. 57, 1272–1286.

Qiao, R.L., Bhattacharya, J., 1991. Segmental barrier properties of the

pulmonary microvascular bed. J. Appl. Physiol. 71, 2152–2159.

Rosado, J.A., Sage, S.O., 2000. Coupling between inositol 1,4,5-trisphos-

phate receptors and human transient receptor potential channel

1 when intracellular Ca2+ stores are depleted. Biochem. J. 350

(Pt. 3), 631–635.

Schachtner, S.K., Wang, Y., Scott Baldwin, H., 2000. Qualitative and

quantitative analysis of embryonic pulmonary vessel formation. Am.

J. Respir. Cell Mol. Biol. 22, 157–165.

Schnitzer, J.E., Siflinger-Birnboim, A., Del Vecchio, P.J., Malik, A.B.,

1994. Segmental differentiation of permeability, protein glycosylation,

and morphology of cultured bovine lung vascular endothelium. Bio-

chem. Biophys. Res. Commun. 199, 11–19.

Schwarz, M.A., Zhang, F., Lane, J.E., Schachtner, S., Jin, Y., Deutsch, G.,

Starnes, V., Pitt, B.R., 2000. Angiogenesis and morphogenesis of

murine fetal distal lung in an allograft model. Am. J. Physiol. 278,

L1000–L1007.

Shalaby, F., Rossant, J., Yamaguchi, T.P., Gertsenstein, M., Wu, X.F., Breit-

man, M.L., Schuh, A.C., 1995. Failure of blood-island formation and

vasculogenesis in Flk-1-deficient mice. Nature 376, 62–66.

Shalaby, F., Ho, J., Stanford, W.L., Fischer, K.D., Schuh, A.C., Schwartz,

L., Bernstein, A., Rossant, J., 1997. A requirement for Flk1 in primitive

and definitive hematopoiesis and vasculogenesis. Cell 89, 981–990.

Spicer, S.S., Schulte, B.A., Thomopoulos, G.N., 1983. Histochemical prop-

erties of the respiratory tract epithelium in different species. Am. Rev.

Respir. Dis. 128, S20–S26.

Stan, R.V., Kubitza, M., Palade, G.E., 1999. PV-1 is a component of the

fenestral and stomatal diaphragms in fenestrated endothelia. Proc. Natl.

Acad. Sci. U. S. A. 96, 13203–13207.

Stevens, T., Fouty, B., Hepler, L., Richardson, D., Brough, G., McMurtry,

I.F., Rodman, D.M., 1997. Cytosolic Ca2+ and adenylyl cyclase

responses in phenotypically distinct pulmonary endothelial cells.

Am. J. Physiol. 272, L51–L59.

Stevens, T., Creighton, J., Thompson, W.J., 1999. Control of cAMP in lung

endothelial cell phenotypes. Implications for control of barrier function.

Am. J. Physiol. 277, L119–L126.

Stevens, T., Rosenberg, R., Aird, W., Quertermous, T., Johnson, F.L., Gar-

cia, J.G., Hebbel, R.P., Tuder, R.M., Garfinkel, S., 2001. NHLBI work-

shop report: endothelial cell phenotypes in heart, lung, and blood

diseases. Am. J. Physiol. 281, C1422–C1433.

Symon, F.A., Wardlaw, A.J., 1996. Selectins and their counter receptors: a

bitter sweet attraction. Thorax 51, 1155–1157.

Taatjes, D.J., Barcomb, L.A., Leslie, K.O., Low, R.B., 1990. Lectin binding

patterns to terminal sugars of rat lung alveolar epithelial cells. J. His-

tochem. Cytochem. 38, 233–244.

Taraseviciene-Stewart, L., Kasahara, Y., Alger, L., Hirth, P., Mc Mahon, G.,

Waltenberger, J., Voelkel, N.F., Tuder, R.M., 2001. Inhibition of the

VEGF receptor 2 combined with chronic hypoxia causes cell death-

dependent pulmonary endothelial cell proliferation and severe pulmo-

nary hypertension. FASEB J. 15, 427–438.

Thurston, G., Baluk, P., McDonald, D.M., 2000. Determinants of endothe-

lial cell phenotype in venules. Microcirculation 7, 67–80.

Tsokos, M., Anders, S., Paulsen, F., 2002. Lectin binding patterns of alveo-

lar epithelium and subepithelial seromucous glands of the bronchi in

sepsis and controls—An approach to characterize the non-specific im-

munological response of the human lung to sepsis. Virchows Arch. 440,

181–186.

Wu, S., Sangerman, J., Li, M., Brough, G.H., Goodman, S.R., Stevens, T.,

2001. Essential control of an endothelial cell ISOC by the spectrin

membrane skeleton. J. Cell Biol. 154, 1225–1233.

Yi, S.M., Harson, R.E., Zabner, J., Welsh, M.J., 2001. Lectin binding and

endocytosis at the apical surface of human airway epithelia. Gene Ther.

8, 1826–1832.

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