1 Developmental Biology and Cardiothoracic Surgery Program, Childrens Hospital Research Institute of Los Angeles, Los Angeles, California 90027; and 2 Department of Physiology and Cellular Biophysics, Columbia University, New York, New York 10032
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Neovascularization is a key regulatory process in fetal growth and development. Although factors promoting growth and development of the pulmonary vasculature have been investigated, nothing is known regarding the molecular mechanisms that may counteract these stimuli. Endothelial monocyte-activating polypeptide (EMAP) II has recently been identified as an antiangiogenic factor in tumor vascular development. We postulated that EMAP II is a putative negative modulator of lung vascular growth. EMAP II mRNA and protein decrease fivefold (P < 0.01) as the developing lungs in the fetal mouse progress from having poor vascularization (day 14) to having complete vascular development at term (day 18.5). EMAP II protein expression continues to remain low throughout postnatal life and into adulthood, with the exception of a surge that correlates with microvascular maturation. Furthermore, through the use of in situ hybridization and immunohistochemistry, EMAP II is localized throughout the lung, with significant expression in the submyoepithelial area during the early stages of lung development when there is minimal vascular development. In contrast, EMAP II is distributed around the large vessels during the end of vascular development, suggesting that EMAP II modulates the neovascularization process. We speculate that EMAP II is a director of neovascularization in the developing lung.
pulmonary; fetal; vasculogenesis; angiogenesis; antiangiogenesis
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
LUNG MORPHOGENESIS is a complex process that is instructed by master genes as well as by growth factor signals serving as induction and progression factors to modulate branching morphogenesis, cytodifferentiation, and neovascularization. Pulmonary vascular formation is an ongoing process that is highly regulated, involving not only proliferation and differentiation of vascular structures but also regression and stasis of these structures (20). Two distinct and separate processes, vasculogenesis and angiogenesis, are considered responsible for lung vascularization during fetal growth and development (2, 17). First, vasculogenesis is the transdifferentiation and organization of endothelial cells (ECs) into vascular structures. This occurs from randomly distributed mesodermal cells that transdifferentiate into ECs, proliferate, and organize into multicellular structures singularly arranged around a central lumen. Second, angiogenesis is the extension of previously formed vessels into undervascularized regions where differentiated ECs proliferate, sprout from previously formed vessels, and form new vascular structures. Within the mouse embryo (term 18.5 days), formation of the pulmonary circulation occurs predominantly during the canalicular stage (known as the vascular stage) from embryonic days (ED) 16.5 through 17.4 (1, 10, 14, 25, 26). During this period, there is an increase in the number of cells that express von Willebrand factor (vWF). These cells coalesce with other vWF-positive cells to form vascular structures by about ED16 (24). Furthermore, there is an increase in the EC-specific antigen flk-1. This protein is a receptor for vascular endothelial growth factor (VEGF) and is observed in mesenchymal cells surrounding the developing airways at about the same time (15). A recent morphological study by deMello et al. (4) demonstrated that peripheral vasculogenesis and central angiogenesis occur concurrently to form the lung vasculature. Communication between the two networks is rare early in lung development (ED13-14) but increases gradually until a full vascular circuit is established by ED17. Growth factors such as basic fibroblast growth factor (22) and VEGF (11, 16) are known stimulants of migration, proliferation, and transdifferentiation of ECs, leading to the appearance of vascular structures. However, nothing is known of the factors mediating the regression and stasis of the developing pulmonary vasculature.
Endothelial monocyte-activating polypeptide (EMAP) II is an 34-kDa
cytokine-like molecule first identified from murine methylcholanthrene A-induced fibrosarcomas (8). Although initial data
suggested that EMAP II was likely to be a proinflammatory cytokine, in
vivo experiments (9) elucidated only a transient and mild inflammatory response. Recently, in vivo studies (unpublished data)
have shown EMAP II to have striking antiangiogenic properties defined
by inhibition of vessel ingrowth in a Matrigel model and a corneal eye
model. Delivery of EMAP II intraperitoneally to mice
resulted in significant suppression of murine and human tumor growth.
Suppression of tumor growth is likely due to perivascular apoptotic
tissue injury caused by the direct targeting of EMAP II to the
proliferating EC population (21). In addition, we found that EMAP II
induced apoptosis only in ECs in vitro (unpublished data).
Therefore, we hypothesized that EMAP II is an important regulator of
lung vascularization.
In this study, we report an inverse correlation between EMAP II expression and vascularization within the developing lung. There is a significant decrease in EMAP II mRNA and protein expression in the developing lungs as it transitions from a poorly vascularized structure (ED14) to full fetal lung vascularization at the end of gestation (ED18.5) and through postnatal life into adulthood. This coincides with the neovascularization process. Furthermore, EMAP II is widely distributed, with significant expression in the myoepithelium when the lung is poorly vascularized, suggesting a broad inhibition of vascular development. Localized targeting of EMAP II to perivascular regions of the large arteries occurs later during the vascular stage, suggesting that EMAP II may play a role in maintaining stasis of larger angiogenic vessels while allowing further proliferation and differentiation of the peripheral vasculature. These data suggest a role for EMAP II as a director of neovascularization in the developing lung.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
RNA isolation, RT, and competitive PCR quantification of mRNA. To assess mRNA levels of EMAP II, we obtained fetal lungs from timed-gestation Swiss-Webster mice (Simonsen, Morgan Hill, CA) at ED14-18 of age (term 18.5 days). Specimens were obtained from at least three different litters, and multiple lungs from the same litter were pooled for each specimen. At least three independent RT products were thus evaluated for each data point. Mediastinal structures were removed by microdissection, and total RNA from embryonic lungs was extracted by guanididium thiocyanate after homogenization utilizing a total RNA kit (Qiagen, Santa Clarita, CA). After spectrophotometric quantitation, total RNA was reverse transcribed with oligo(dT) primers as described previously by Zhao and Nishimoto (29). A competitive PCR assay was designed to quantitate subtle developmental differences in gene marker expression. Primers for murine EMAP II were primer I (upstream) 5'-GCATCGCGTCTGGATCTTCGAATT-3' and primer II (downstream) 5'-GTATGTGGCCACACACTCAGCATT-3' (GenBank accession no. U10118). Competitor primers were constructed based on the initial structure of the above designed EMAP II primers. Each composite primer had the target EMAP II gene primer sequence attached to a short stretch of sequence designed to hybridize to the opposite strand of a heterogeneous DNA fragment, thus incorporating itself into the heterogeneous fragment during the PCR amplification and ensuring that the competitor molecules had the same gene-specific primer sequences as the EMAP II cDNA. The heterogeneous DNA was derived from a piece of v-erbB DNA. The competitor PCR product was subsequently 484 bp in size compared with the EMAP II cDNA size of 411 bp, and identities of both EMAP II cDNA and its competitor were confirmed by DNA sequencing.
PCR amplification was carried out in a DNA Robocycler (Stratagene, La Jolla, CA) with a modification of a previously described assay for matrix Gla protein (30) (each time point was repeated 3-5 times). Each sample underwent 35 cycles of denaturation at 94°C for 1 min, annealing at 62°C for 2 min, and extension at 72°C for 2 min after an initial 3-min denaturation at 94°C. The final cycle included a 5-min extension step. Subsequent electrophoresis was performed to separate the target and competitor PCR products that were 411 and 484 bp in size, respectively. The intensity of expression was determined by densitometric analysis of photographed gels (Polaroid 667) with ImageQuant band-analyzing software (Molecular Dynamics, Sunnyvale, CA).
Protein isolation and Western
blotting. Fetuses from timed-gestation Swiss-Webster
mice at ED14-18.5 (term 18.5 days) or postnatal mice through
adulthood were dissected. Afterward, the lungs were isolated and
underwent protein analysis. Briefly, specimens from three different
litters were obtained at each time point, and multiple lungs from the
same litter were pooled for each specimen (for specimens in the fetal
stage). In neonatal mice through adulthood, lungs were isolated from
mice in different litters on three different occasions and subjected to
protein analysis. After the mediastinal structures were removed by
microdissection, the lungs were rinsed in 50 mM Tris, pH 7.4, 0.9 N
NaCl, and 0.01% NaN3. The lungs
were then homogenized by sonification with a Branson sonifier and
stored at 70°C. For Western analysis, homogenates were
cleared by centrifugation at 14,000 g
for 20 min, the protein concentration was determined by the
bicinchoninic acid method, and the samples were normalized by protein content. Equal amounts of protein were then subjected to
electrophoresis on a 12% SDS-PAGE gel, transferred to Immobilon-P, blocked overnight in a casein-based blocking solution (Boehringer Mannheim, Indianapolis, IN), and probed with a rabbit anti-EMAP II
antibody [polyclonal monospecific antibody based on immunoblotting of
plasma and cell extracts; anti-EMAP II IgG blocked the activity of
recombinant EMAP II (rEMAP II) in cell culture assays]. Specific binding was detected with a chemiluminescence substrate (Pierce, Rockford, IL) and XAR-5 film (Eastman Kodak, Rochester, NY).
Quantitative analysis was accomplished with a Molecular Dynamics
personal densitometer.
Immunohistochemistry and histology. Developing lungs on ED14 and ED18 and postnatally through adulthood were fixed with 4% paraformaldehyde (PFA) and then progressively dehydrated before being embedded in paraffin and sectioned. For immunolocalization of EMAP II (polyclonal monospecific antibody based on immunoblotting of plasma and cell extracts; anti-EMAP II IgG blocked the activity of rEMAP II in cell culture assays) and platelet endothelial cell adhesion molecule (PECAM)-1 antigens (28), we employed rabbit anti-rEMAP II IgG (5 µg/ml) and rat anti-murine PECAM-1 antibody (4 µg/ml). Tissues were deparaffinized and underwent peroxide quenching. After the sections were blocked, they were exposed to anti-EMAP II IgG or rat anti-mouse PECAM-1 overnight at 4°C with a histostain kit from Zymed (San Francisco, CA). Sections were then incubated with secondary biotinylated antibody according to the manufacturer's protocol. A brief incubation with the streptavidin-horseradish peroxidase conjugate system (Zymed) was followed by development with the chromogen substrate 3-amino-9-ethylcarbazole. The preabsorbed immune control was accomplished by incubating the primary antibody with a 50-fold amount of EMAP II protein. Coimmunolocalization was accomplished with a Histostain-DS Kit from Zymed. Briefly, sections were prepared as described above, and then they were blocked and exposed first to anti-mouse PECAM-1 overnight at 4°C. After a brief incubation with a streptavidin-alkaline phosphatase, the second antibody was revealed with 5-bromo-1-chloro-3-indoyl phosphate/nitro blue tetrazolium. Sections underwent blocking and then exposure to the second primary antibody, anti-EMAP II, overnight at 4°C. After a brief incubation with the streptavidin-horseradish peroxidase conjugate system, the second primary antibody was developed with 3-amino-9-ethylcarbazole.
In situ hybridization. Developing lungs (ED14 and 2 days postnatally) were obtained for use for in situ hybridization. The digoxigenin (Dig) RNA probe antisense and sense (control) were made with the Dig RNA labeling kit (SP6/T7) from Boehringer Mannheim. Lungs were fixed in 4% PFA in diethyl pyrocarbonate (DEPC)-treated water, dehydrated with ethanol, and then embedded in paraffin. Using DEPC-treated equipment and solutions, we sectioned the paraffin-embedded specimens, and they were rehydrated and incubated in a prewarmed 5 µg/ml proteinase K solution. Slides were then reimmersed in 4% PFA, treated with 0.25% acetic anhydride, and dehydrated. Sections were exposed to a hybridization solution containing 50% formamide, 10% dextran sulfate, 1 mg/ml of tRNA, 1× Denhardt's solution, 4× saline-sodium citrate (SSC), 50 mM Tris, and 5 mM EDTA that contained 150-300 ng/ml of Dig-labeled RNA probe at 50°C overnight. The RNA probe for EMAP II was 456 bp in size and obtained from a region that has minimal homology with other known proteins. The slides were washed at 55°C in 2× SSC-50% formamide, 1× SSC, and 0.1× SSC for 30 min before being incubated with RNase A (20 µg/ml) for 30 min at 37°C. Slides were then rinsed with 2× SSC, and Dig nucleic acid detection was accomplished with the Genius 3 kit from Boehringer Mannheim. Briefly, slides were incubated in 0.1 M maleic acid-0.15 M NaCl (pH 7.5) for 5 min, and then they underwent blocking in a 1% block reagent. After the blocking, the slides were incubated with anti-Dig-alkaline phosphatase conjugate at 4°C overnight. Slides were then rinsed and incubated with a dilute 5-bromo-1-chloro-3-indoyl phosphate/nitro blue tetrazolium solution for 3 h at room temperature. Afterward, slides were counterstained with a 0.02% fast green solution for 2 min, rinsed in water, air-dried, and mounted.
Statistical analysis. Statistical analysis was performed with Student's t-test with the program Statview.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Temporal expression of EMAP II mRNA.
To determine the level of expression of EMAP II mRNA in the developing
lung, levels of EMAP II mRNA were quantitated by competitive RT-PCR.
The EMAP II competitive PCR results are illustrated in Fig.
1. The amount of EMAP II was analyzed by
comparison to the known 10-fg competitor. Results of EMAP II expression
are also compared with a -actin control. We found an inverse
correlation between EMAP II expression and vascularization within the
developing lung. As fetal lungs entered the vascular
stage (ED16.5), there was a significant reduction in EMAP
II mRNA. Furthermore, EMAP II mRNA was further reduced up to 83%
(P < 0.01), comparing ED14 (a poorly
vascularized stage) with ED18 (when there is full fetal vascular
development; term 18.5 days; Fig. 1, C
and D). This significant reduction
in EMAP II mRNA was inversely proportional to the increase in vascular formation within the developing lung.
|
Temporal expression of EMAP II
protein. To assess protein levels of EMAP II in the
developing lung, we quantitated the levels of EMAP II by Western blot
analysis. Equal amounts of lung protein extracts underwent Western blot
analysis (performed in triplicate from three different litters or
animals; Fig. 2) and showed a significant
(fivefold) decrease in EMAP II protein from ED14 to ED18
(P < 0.001) that was maintained into
adulthood (Fig. 2C). During the minimal vascular state, ED14-15, EMAP II was highly expressed. In contrast, there was a marked significant decrease by
ED17, well into the vascular stage. This low level of expression, with
one exception, was maintained throughout life. Interestingly, we noted
that on postnatal days 8-16,
there was a significant surge in the expression of EMAP II within the
lungs, consistent with the period of time that murine lungs
undergo microvascular maturation (Fig
2C). Thus EMAP II protein expression
parallels the decrease in EMAP II mRNA.
|
Spatial expression of EMAP II mRNA and
protein. In situ hybridization revealed that at ED14 in
fetal lungs (Fig. 3,
A and B), EMAP II was produced throughout
the lung, with a predominance in the myoepithelium that lines the
bronchioles. The location of EMAP II mRNA expression
correlates with the same area of expression as other regulators of
vessel formation, flt-1 and flk-1 [VEGF receptors localized by in
situ hybridization (5)], that show increased expression over the
same time period in which EMAP II production is significantly
declining. Furthermore, the myoepithelial region is the area where the
foregut splanchnopleuric mesoderm is located, from
which the vascular supply is derived. In contrast, after full fetal vascular development is reached (postnatal
day 2; Fig. 3,
C and
D), EMAP II is localized
predominantly to the perivascular region, consistent with those results
seen with immunohistochemistry. This result is consistent with that
seen throughout the lung from ED18 into adulthood.
|
Immunolocalization of EMAP II in the lungs of a 14-day embryo indicates
that EMAP II (Fig. 4,
A and
B) is widely expressed throughout
the tissue (Fig. 4B, open
arrows) and bronchioles, with marked
amounts being produced in the myoepithelial region (Fig.
4B, closed arrows).
However, although EMAP II does localize to the blood vessels, it is not
specific for only vascular structures (Fig. 4,
C and
D). Coimmunolocalization (Fig.
4E) confirms that EMAP II is
expressed widely throughout the lungs in the bronchioles, myoepithelium
(Fig. 4E, open arrow), and
perivascular regions (confirmed by coimmunolocalization with the use of
PECAM-1-alkaline phosphatase; Fig. 4E,
closed arrows) and diffusely throughout the lung. In
contrast, by ED18, a time when EMAP II production is markedly reduced,
EMAP II immunolocalizes predominantly to the large pulmonary vessels
(Fig. 5, A
and B), with minimal bronchial expression as defined by sequential sections with PECAM-1 antibody for
vessel identification (Fig. 5, C and
D).
Coimmunolocalization of lungs from a mouse on postnatal
day 2 (Fig.
5E) confirms the perivascular
location of EMAP II [3-amino-9-ethylcarbazole-labeled EMAP II
(Fig. 5E, closed arrows) and
PECAM-1-alkaline phosphatase (Fig. 5E,
open arrow)]. Interestingly, it appears that
EMAP II is localizing to the smooth muscle cell (Fig.
5E, closed arrows), consistent with other modulators of vessel growth such as FGF. With one
exception (postnatal days
8-16), subsequent sections obtained from
postnatal day 1 through adulthood are
consistent with those results seen on ED18. During
days 8-16 in
postnatal murine lung (Fig. 6,
A and
B), coinciding with a surge of EMAP II protein (Fig. 2C), EMAP II
distribution is widespread in both large and capillary
vessels. Of note, EMAP II is distributed in a perivascular,
subendothelial pattern in larger vessels (6B, arrows). This is
confirmed through the use of PECAM-1 immunolocalization on sequential
sections (Fig. 6, C and
D), where PECAM-1 is localized to
endothelial cell junctions (Fig. 6D, arrows). This period of time is consistent with microvascular maturation of the lung within the
mouse. EMAP II immunolocalization specificity was confirmed through the
use of a preabsorbed control (data not shown). Thus, when EMAP II is
highly expressed in the early stages of lung development before
significant vascularization of the lung, its distribution is throughout
the lung. This is reflected in the spatial expression of EMAP II mRNA
and protein, with its localization in the myoepithelial region
(considered to be the foregut splanchnopleuric mesoderm responsible for
vascular formation), suggesting that it may be functioning as a
regulator of vascular development. However, during the later stages
when there is rapid neovascularization of the lungs, EMAP II expression
is limited to the large vessels. This suggests that the primary
function of EMAP II later in gestation is to stabilize or cause stasis
of vessel angiogenesis, whereas its decreased expression in the rest of
the lung may facilitate the neovascularization process.
|
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Neovascularization is critical to the growth and development of the fetal lung. Two distinct processes, vasculogenesis (the transdifferentiation and organization of cells into vasculature) and angiogenesis (the proliferation and sprouting of vessels from previously formed vessels) organize the appearance of vascular structures within the mesenchyme of embryonic lungs (2, 17). The development of a mature pulmonary circulation is dependent on the angiogenic ingrowth of central pulmonary vessels from the heart and linkage of these vessels to the distal capillary networks formed by vasculogenesis (3). These two separate embryonic pulmonary vascular systems connect before birth, forming a vascular circuit composed of the central pulmonary artery, branch arteries, capillaries, and veins (19). These two systems arise concurrently and gradually form a uniform vascular system by ED17 (4).
Regulation of new blood vessel growth is controlled by vasoactive agents, growth factors (13, 16, 22, 23), matrix proteins, and tissue interactions (7, 12, 18, 27). Effectors of blood vessel growth display a marked heterogeneity in the developmental responses of the large and small vessels. The initial characterization of neovascularization within the lung was limited by vessel markers that identified vessel development (vascular formation noted on ED16) such as vWF (24). However, Vecchi et al. (28) noted that PECAM-1 antibody was a reliable marker for the identification of murine embryological ECs; it is expressed within the murine lung vasculature as early as the pseudoglandular stage. The presence of an active developing lung vasculature within the pseudoglandular stage was confirmed by the morphological study of deMello et al. (4). Furthermore, positive modulators of neovascularization such as VEGF have been noted to have an impact on the behavior of primordial ECs engaged in vasculogenesis and are important in the vascular patterning and regulation of vessel size (5).
Lung development, including neovascularization, is classically divided into five chronological stages in the mouse and is based on the growth and differentiation of specific pulmonary epithelial structures: 1) the embryonic stage is from ED9.5 to ED11.5 in the mouse, 2) airway construction occurs in the pseudoglandular stage from ED11.5 to ED16.5, 3) vessels develop in the canalicular stage from ED16.5 to ED17.4 (also known as the vascular stage), 4) the number of terminal sacs and vascularization increase and type I and II epithelial cells differentiate from ED17.4 to postnatal day 5 in the terminal sac stage, and 5) during the alveolar stage, from postnatal days 5-14, there is development and maturation of alveolar ducts and alveoli (1, 10, 14, 25, 26). Hence factors that enhance lung neovascular formation in the developing lung would be expected to have their greatest influence during the canalicular and terminal stages, whereas negative modulators of neovascularization would be expected to exert the greatest influence up to the canalicular stage.
EMAP II, originally identified as an antiangiogenic protein, has been shown to be a potent inhibitor of vessel growth and differentiation in vivo and in vitro (21; unpublished data). Thus EMAP II may play a role in lung vascularization. Consistent with the ability of EMAP II to modulate vessel growth and coinciding with the pseudoglandular stage, levels of EMAP II mRNA (Fig. 1) and protein (Fig. 2) are significantly elevated. Furthermore, the wide distribution of EMAP II throughout the lung in the pseudoglandular stage (Fig. 4, A and B), within the mesenchymal cell population, bronchioles, and myoepithelial region, suggests that EMAP II exerts an antiangiogenic as well as an antivasculogenic effect during early lung development.
It has been shown that lung formation is derived from a composite of endodermal and mesodermal tissues. The vasculature of the lung is derived from the foregut splanchnopleuric mesoderm, which surrounds the epithelium as it grows out from the mediastinum into the pleural space, whereas the endoderm of the lung bud gives rise to the mucosal lining of the bronchi and epithelial cells of the alveoli. During the early stages of lung development, expression of EMAP II is increased within the mesenchyme surrounding the bronchioles, an area consistent with the mesoderm. Therefore, the spatial-temporal expression of EMAP II is consistent with its potential role as a negative modulator of vessel growth because vessels arise in the foregut splanchnopleuric mesoderm at later stages of lung development. Furthermore, EMAP II expression within the bronchioles may function as a traffic director for primordial ECs by preventing the development of capillaries within the bronchioles. We have shown that EMAP II has a direct suppressive effect on the dividing and growing ECs (21) and speculate that this suppressive effector is especially strong during fetal growth and development. As a negative modulator of vascular growth, EMAP II appears to be important early in lung development when cell proliferation, neovascularization, and cell dedifferentiation are the main emphasis. Consistent with this observation, Gebb and Shannon (6) recently showed that this same region, the area of the foregut splanchnopleuric mesoderm (myoepithelial layer), contains the VEGF receptors that increase significantly during the vascular stage within the rat, consistent with the time when EMAP II expression is undergoing marked reduction.
In contrast to high expression of EMAP II in the early stages of lung development, during the later vascular stage, there is a significant decline in EMAP II mRNA (Fig. 1) and protein (Fig. 2). This suggests that during rapid neovascularization, EMAP II is no longer present to exert a general negative regulatory effect on vascular formation. The level of EMAP II remains low until postnatal day 8 when there is a marked surge in its expression. This coincides with microvascular maturation within the murine lung. At this stage, EMAP II is found in capillaries within the air spaces and in the musculature surrounding large vessels, indicating that EMAP II is holding further vessel development in check. Furthermore, it is intriguing that by ED18 and later, EMAP II appears to be produced within the basement membrane of the blood vessel (Figs. 5 and 6). This is interesting because it is the same location where other positive regulators of vessel growth, such as FGF and VEGF, are made. Therefore, this further supports the modulatory role of EMAP II in vessel growth and stability.
Hence we have shown that during early embryological development when there is minimal vascular development, EMAP II is highly expressed in the developing lung. It is distributed throughout the mesenchyme and strongly concentrated within the myoepithelial layer. EMAP II localization is coincident with the region of the foregut splanchnopleuric mesoderm where the pulmonary vasculature develops. Within this region, we see an initial marked expression of EMAP II that decreases in correlation with the rise in vasculature. At the point of microvascular maturation, EMAP II expression increases and is associated with capillaries and large vessels. The localization of EMAP II to the large vessels during the vascular stage and through adulthood suggests that EMAP II may further influence vascular growth by stabilizing existing vascular structures; however, further investigation is needed.
Therefore, the striking elevation of EMAP II during the early minimal vascular stage of lung development and its rapid decline during the vascularization stage invite the speculation that EMAP II may be a director of endothelial dedifferentiation (vasculogenesis) and the angiogenic process of embryonic endothelial lung development.
![]() |
ACKNOWLEDGEMENTS |
---|
This research was supported by American Heart Association Grant-in-Aid 1131-GI1 (to M. Schwarz) and National Heart, Lung, and Blood Institute Grants HL-44060, HL-49772, and HL-60231 (to D. Warburton).
![]() |
FOOTNOTES |
---|
M. Schwarz completed this work during the tenure of a Clinician Scientist Award from the American Heart Association (93004140).
Address for reprint requests: M. Schwarz, Childrens Hospital of Los Angeles, Cardiothoracic Intensive Care Medicine, 4650 Sunset Blvd., MS #66, Los Angeles, CA 90027.
Received 7 November 1997; accepted in final form 29 October 1998.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Burri, P. H.,
J. Dbaly,
and
E. R. Weibel.
The postnatal growth of the rat lung. I. Morphometry.
Anat. Rec.
178:
711-730,
1974[Medline].
2.
Coffin, J. D.,
J. Harrison,
S. Schwartz,
and
R. Heimark.
Angioblast differentiation and morphogenesis of the vascular endothelium in the mouse embryo.
Dev. Biol.
148:
51-62,
1991[Medline].
3.
DeMello, D.,
and
L. Reid.
Pre and postnatal development of the pulmonary circulation.
In: Basic Mechanisms of Pediatric Respiratory Disease: Cellular and Integrative, edited by V. Chernik,
and B. Mellins. Philadelphia, PA: Decker, 1991, p. 36-54.
4.
DeMello, D.,
D. Sawyer,
N. Galvin,
and
L. Reid.
Early fetal development of lung vasculature.
Am. J. Respir. Cell Mol. Biol.
16:
568-581,
1997[Abstract].
5.
Drake, C.,
and
C. Little.
Exogenous vascular endothelial growth factor induces malformed and hyperfused vessels during embryonic neovascularization.
Proc. Natl. Acad. Sci. USA
92:
7657-7661,
1995[Abstract].
6.
Gebb, S.,
and
J. Shannon.
Tissue interactions mediate early events in pulmonary vasculogenesis (Abstract).
Am. J. Respir. Crit. Care Med.
157:
A208,
1998.
7.
Ignotz, R. A.,
and
J. Massague.
Transforming growth factor-beta stimulates the expression of fibronectin and collagen and their incorporation into the extracellular matrix.
J. Biol. Chem.
261:
4337-4345,
1986
8.
Kao, J.,
J. Ryan,
J. Brett,
J. Chen,
H. Chen,
Y. G. Fan,
G. Godman,
P. C. Familletti,
F. Wang,
Y. C. Pan,
D. Stern,
and
M. Clauss.
Endothelial monocyte-activating polypeptide (EMAP) II. A novel tumor-derived polypeptide that activates host response mechanisms.
J. Biol. Chem.
267:
20239-20247,
1992
9.
Kao, J.,
K. Houck,
Y. Fan,
I. Haehnel,
S. K. Libutti,
M. L. Kayton,
T. Grikscheit,
J. Chabot,
R. Nowygrod,
S. Greenberg,
W. Kuang,
D. W. Leung,
J. R. Hayward,
W. Kissel,
M. Heath,
J. Brett,
and
D. M. Stern.
Characterization of a novel tumor-derived cytokine. Endothelial monocyte-activating polypeptide II.
J. Biol. Chem.
269:
25106-25119,
1994
10.
Kauffman, S. L.
Histogenesis of the papillary Clara cell adenoma.
Am. J. Pathol.
103:
174-180,
1981[Abstract].
11.
Kim, J.,
B. Li,
J. Winer,
M. Armanini,
N. Gillett,
H. Phillips,
and
N. Ferrara.
Inhibition of VEGF-induced angiogenesis suppresses tumor growth in vivo.
Nature
362:
841-844,
1993[Medline].
12.
Lane, T. F.,
M. L. Iruela-Arispe,
R. S. Johnson,
and
E. H. Sage.
SPARC is a source of copper-binding peptides that stimulate angiogenesis.
J. Cell Biol.
125:
929-943,
1994[Abstract].
13.
Maciag, T.,
T. Mehlman,
R. Friesel,
and
A. Schrieber.
Heparin binds endothelial cell growth factor, the principal cell mitogen in bovine brain.
Science
225:
932-935,
1984[Medline].
14.
Meyrick, B.,
and
L. Reid.
Ultrastructure of alveolar lining and its development.
In: Development of the Lungs, edited by W. Hoden. New York: Dekker, 1977, p. 135-214.
15.
Millauer, B.,
S. Wizigmann-Voos,
H. Schnurch,
R. Martinez,
N. P. H. Moller,
W. Risau,
and
A. Ullrich.
High affinity VEGF binding and development expression suggest Flk-1 as a major regulator of vasculogenesis and angiogenesis.
Cell
72:
835-846,
1993[Medline].
16.
Plate, K.,
G. Breier,
H. Weich,
and
W. Risau.
Vascular endothelial growth factor is a potential tumor angiogenesis factor in human gliomas in vivo.
Nature
359:
845-848,
1992[Medline].
17.
Poole, T. J.,
and
J. D. Coffin.
Vasculogenesis and angiogenesis: two distinct morphogenetic mechanisms establish embryonic vascular pattern.
J. Exp. Zool.
251:
224-231,
1989[Medline].
18.
Risau, W.,
and
V. Lemmon.
Changes in the vascular extracellular matrix during embryonic vasculogenesis and angiogenesis.
Dev. Biol.
125:
441-450,
1988[Medline].
19.
Roman, J.
Cell-cell and cell-matrix interactions in development of the lung vasculature.
In: Lung Growth and Development, edited by J. McDonald. New York: Dekker, 1997, p. 365-399.
20.
Scavo, L.,
R. Ertsey,
C. Chapin,
L. Allen,
and
J. Kitterman.
Apoptosis in the development of rat and human fetal lungs.
Am. J. Respir. Cell Mol. Biol.
18:
21-31,
1998
21.
Schwarz, M.,
J. Brett,
J. Li,
J. Hayward,
R. Schwar,
J. Kao,
O. Chappey,
J. Wautier,
J. Chabot,
P. Lo Gerfo,
and
D. Stern.
Endothelial monocyte-activating polypeptide (EMAP) II, a novel antiangiogenic protein, suppresses tumor growth and induces apoptosis in endothelial cells (Abstract).
Circ. Supp
l.92:
34,
1995.
22.
Shing, Y.,
J. Folkman,
R. Sullivan,
C. Butterfield,
J. Murray,
and
M. Klagsbrun.
Heparin affinity: purification of a tumor-derived capillary endothelial cell growth factor.
Science
225:
932-935,
1984[Medline].
23.
Stenmark, K.,
and
M. Weiser.
Vascular development and function.
In: Pediatrics and Perinatology, The Scientific Basis (2nd ed.). London: Arnold, 1996, p. 683-693.
24.
Sundell, C. L.,
and
J. Roman.
Control of lung vasculogenesis by extracellular matrix composition (Abstract).
Mol. Biol. Cell
5:
179a,
1994.
25.
Ten Have-Opbroek, A. A.
The development of the lung in mammals: an analysis of concepts and findings.
Am. J. Anat.
162:
201-219,
1981[Medline].
26.
Thurlbeck, W.
Pre- and postnatal organ development.
In: Basic Mechanisms of Pediatric Respiratory Disease: Cellular and Integrative, edited by V. Chernik,
and B. Mellins. Philadelphia, PA: Decker, 1991, p. 23-35.
27.
Tonnesen, M. G.,
D. Jenkins, Jr.,
S. L. Siegal,
L. A. Lee,
J. C. Huff,
and
R. A. F. Clark.
Expression of fibronectin, laminin, and factor VIII-related antigen during development of the human cutaneous microvasculature.
J. Invest. Dermatol.
85:
564-568,
1985[Abstract].
28.
Vecchi, A.,
C. Garlanda,
M. Lampugnani,
M. Resnati,
C. Matteucci,
A. Stoppacciaro,
H. Schnurch,
W. Risau,
L. Rucco,
A. Mantovani,
and
E. Dejana.
Monoclonal antibodies specific for endothelial cells of mouse blood vessels. Their application in the identification of adult and embryonic endothelium.
Eur. J. Cell Biol.
63:
247-254,
1994[Medline].
29.
Zhao, J.,
and
S. K. Nishimoto.
An RNA-competitive polymerase chain reaction method for human matrix gamma-carboxyglutamic acid protein mRNA measurement.
Anal. Biochem.
228:
162-164,
1995[Medline].
30.
Zhao, J.,
and
D. Warburton.
Matrix Gla protein gene expression is induced by transforming growth factor- in embryonic lung culture.
Am. J. Physiol.
273 (Lung Cell. Mol. Physiol. 17):
L282-L287,
1997