Microvascular Research 67 (2004) 139 – 151
www.elsevier.com/locate/ymvre
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 Stevens b,*
b
a
Department of Pathology, Center for Lung Biology, The University of South Alabama College of Medicine, Mobile, AL 36617, USA
Department 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 different vascular sites. In the systemic circulation, permeability 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 (Grisham 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).
* Corresponding author. Fax: +1-251-460-7452.
E-mail address: [email protected] (T. Stevens).
0026-2862/$ - see front matter D 2004 Elsevier Inc. All rights reserved.
doi:10.1016/j.mvr.2003.11.006
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 contiguous 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 vasculogenic blood islands could be resolved using the vascular
casting technique. This issue has also been addressed by
assessing the temporal expression pattern of endothelial
140
J. King et al. / Microvascular Research 67 (2004) 139–151
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 sitespecific 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 contrast, macrovascular endothelial cells express more eNOS
(Stevens, unpublished) and generate more nitric oxide (AlMehdi, 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 endothelial 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 inhibition of VEGF signaling reduces alveolar septation (as in
emphysema) (Kasahara et al., 2000) and, in combination
with hypoxia, generates microvascular (c100 Am) plexigenic lesions (Taraseviciene-Stewart et al., 2001). Thus,
lung endothelial cell origin may be an important determinant 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 sternotomy 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 immunostaining 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 15ml 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 microvascular 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 Research 67 (2004) 139–151
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-tetramethylindocarbocyanine-labeled low-density lipoprotein (DiI-acetylated 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 microscopes were used to view the slides.
141
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. FITCconjugated 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 35mm 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 Am
polycarbonate membranes (Nunc, Naperville, IL) for transmission electron microscopy, and grown for 4 days to
Table 1
List of lectins, plant or animal sources, nominal specificities, and staining intensity in PAECs and PMVECs. + = weak staining; ++ = moderate staining; +++ =
strong staining. Staining intensity was approximately the same in all lectins screened, except for Glycine max, Griffonia simplicifolia, and Helix pomatia
Lectin
Source
Nominal specificity
PAEC
PMVEC
Arachis hypogea
Caragana arborescens
Lens culinaris
PNA
CAA
LcH
peanut
pea tree
lentil
++
++
+++
++
++
+++
Lycopersicon esculentum
Ricinus communis
Ulex europaeus
Glycine max
LEA
RCA120
UEA-I
SBA
tomato
castor bean
gorse, furze
soybean
lactose > h-D-galactose
GalNAc
a-mannose > a-glucose,
aGlcNAc
GlcNAch(1,4)GlcNAc
lactose > galactose
a-L-fucose GlcNAch(1,4)GlcNAc
terminal a and h-GalNAc > a and h-Gal
+++
+++
++
+
+++
+++
++
+++
Griffonia simplicifolia
GS-I
N/A
a-galactose > a-GalNAc
+
+++
Helix pomatia
HPA
edible snail
a-GalNAc > h-GalNAc
+++
+
Reference
(Alvarez-Fernandez and
Carretero-Albinana, 1990;
Honda et al., 1986; Kawai
et al., 1988; Mazzuca et al.,
1982; Spicer et al., 1983)
(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)
(Palmer and Bale, 1987;
Taatjes et al., 1990;
Yi et al., 2001)
As indicated, Helix pomatia preferentially binds PAECs, while Glycine max and Griffonia simplicifolia preferentially bind PMVECs.
142
J. King et al. / Microvascular Research 67 (2004) 139–151
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 micrographs. 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 parenchyma 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 microscope glass coverslips (Fisher Scientific, Pittsburgh,
PA) and grown to confluence. Cytosolic Ca2+ was estimated with the Ca2+-sensitive fluorophore fura 2/acetoxymethylester (Molecular Probes, Eugene, OR) according
to methods previously described. Calculations of free
[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. Oneway 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;
Fig. 1. Endothelial cell phenotypes in the intact lung can be discriminated by lectin binding. Green fluorescent stain in the lumen of the pulmonary artery
(arrows) represents staining of endothelial cells with Helix pomatia. No staining was observed with Helix pomatia in the peripheral lung (40, fluorescent
microscope). Staining with Glycine max and Griffonia simplicifolia is absent in the pulmonary artery but is present in peripheral lung capillaries (arrows; 67,
confocal microscope).
J. King et al. / Microvascular Research 67 (2004) 139–151
143
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 according
to Sigma recommendations. In blocking studies, cells were incubated for 30 min with the blocking sugar before a 15-min incubation with the lectin.
Schnitzer et al., 1994). Nine different lectins were therefore
screened for binding to the rat pulmonary artery and microvascular endothelial cell surface (Table 1). Of the nine lectins
examined, six did not distinguish between macro- and
microvascular cell types, while three demonstrated a preferential binding pattern. In vivo, FITC-labeled Helix pomatia
Fig. 3. Lectin-induced agglutination discriminates endothelial cell phenotypes in vitro. Helix pomatia selectively agglutinates PAECs whereas Glycine max and
Griffonia simplicifolia selectively agglutinate PMVECs. Cells were trypsinized and triturated in to single cell suspensions, then allowed to agglutinate in the
presence of lectins. Arrows indicate cell clumps. Pictures are representative of five separate experiments.
144
J. King et al. / Microvascular Research 67 (2004) 139–151
staining was observed selectively in macrovascular endothelia. 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 characteristics 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.
J. King et al. / Microvascular Research 67 (2004) 139–151
145
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.
146
J. King et al. / Microvascular Research 67 (2004) 139–151
flattened, although scattered projections were observed. The
capillary endothelial cells measured 10.3– 26.9 Am in diameter and 2.6– 7.2 Am in maximum height. Because of the
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 RER was 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, measuring as little as 62.5 nm in height when perfused fixed
vessels were examined. In situ capillaries of the lung parenchyma 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
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) Serumrestriction (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.
J. King et al. / Microvascular Research 67 (2004) 139–151
147
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-induced 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 calcium in the endoplasmic reticulum (Putney, 1986). At
present, the signal(s) linking calcium store depletion to
activation of store-operated calcium entry is unclear, although three separate models have been developed to
148
J. King et al. / Microvascular Research 67 (2004) 139–151
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 between the endoplasmic reticulum and plasmalemma in
activation of store-operated calcium entry channels. We
therefore examined the morphological distribution of endoplasmic reticulum in PAECs and PMVECs, to evaluate
whether physical coupling between the endoplasmic reticulum and plasma membrane could provide a plausible explanation 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 micrographs 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 contact 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 adhesion, 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 evaluated 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; NorgardSumnicht et al., 1995; Schnitzer et al., 1994; Symon and
Wardlaw, 1996). Lectins interact with cell surface carbohydrates 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-acetylgalactosamine carbohydrate. Since the glycocalyx contributes 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 carbohydrate structures to site-specific inflammatory responses will
be important to resolve.
PAECs and PMVECs possessed significant morphological 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 socalled store operated calcium entry pathways (Moore et al.,
1998b; Nilius and Droogmans, 2001), where calcium store
J. King et al. / Microvascular Research 67 (2004) 139–151
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 (Birnbaumer 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 endoplasmic 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 calciummediated gap formation (Kelly et al., 1998). In our present
studies, close coupling was observed between the endoplasmic 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 –membrane 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 findings 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.
149
In summary, PAECs and PMVECs differ structurally
and functionally—even when their environments are similar. These results support the idea that a cell’s embryological (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 bronchogenic 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 lectins. 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. Recent 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. Modulation of Ca(2+) entry by polypeptides of the inositol 1,4, 5- trisphosphate 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+-selective, 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-operated 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.
150
J. King et al. / Microvascular Research 67 (2004) 139–151
Del Vecchio, P.J., Siflinger-Birnboim, A., Belloni, P.N., Holleran, L.A.,
Lum, H., Malik, A.B., 1992. Culture and characterization of pulmonary 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., Breitman, M.L., 1995. Vascularization of the mouse embryo: a study of flk1, 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 endothelial 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. Scanning 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 microscopy. 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 permeability 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-terminal 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., Ambudkar, 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 elastase-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 pulmonary endothelial cells expressing Trp1. Am. J. Physiol. 275,
L574 – L582.
Moore, T.M., Chetham, P.M., Kelly, J.J., Stevens, T., 1998b. Signal transduction 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 bovine 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 pomatia lectin-binding sites in mouse lung elastic fibers. Histochemistry 88,
91 – 95.
Parekh, A.B., Penner, R., 1997. Store depletion and calcium influx. Physiol. 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.
J. King et al. / Microvascular Research 67 (2004) 139–151
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-trisphosphate 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. Biochem. 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., Breitman, 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 properties 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,
151
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., Garcia, J.G., Hebbel, R.P., Tuder, R.M., Garfinkel, S., 2001. NHLBI workshop 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. Histochem. 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 deathdependent pulmonary endothelial cell proliferation and severe pulmonary hypertension. FASEB J. 15, 427 – 438.
Thurston, G., Baluk, P., McDonald, D.M., 2000. Determinants of endothelial cell phenotype in venules. Microcirculation 7, 67 – 80.
Tsokos, M., Anders, S., Paulsen, F., 2002. Lectin binding patterns of alveolar epithelium and subepithelial seromucous glands of the bronchi in
sepsis and controls—An approach to characterize the non-specific immunological 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.