close

Вход

Забыли?

вход по аккаунту

?

625

код для вставкиСкачать
THE ANATOMICAL RECORD 254:61–73 (1999)
Differential Immunolocalization
of VEGF in Rat and Human Adult
Lung, and in Experimental Rat Lung
Fibrosis: Light, Fluorescence,
and Electron Microscopy
HEINZ FEHRENBACH,1* MICHAEL KASPER,2 MICHAEL HAASE,1
DIETER SCHUH,1 AND MARTIN MÜLLER1
1Institute of Pathology, University Clinics ‘‘Carl Gustav Carus,’’
Technical University of Dresden, Dresden, Germany
2Institute of Anatomy, University Clinics ‘‘Carl Gustav Carus,’’
Technical University of Dresden, Dresden, Germany
ABSTRACT
Vascular endothelial growth factor (VEGF) is a cytokine with main
angiogenetic functions in embryonic development and tumor-formation. In
the adult lung, reports of the localization of VEGF were controversial. A
precise cell typing of VEGF-positive pulmonary cells is still lacking. Nothing
is known about a potential role in pulmonary fibrosis. Immunohistochemistry (IH), double immunofluorescence microscopy (DIF), and immunoelectron
microscopy (IEM) were used to study the differential distribution of VEGF in
paraffin-embedded (IH, DIF) and in cryo-substituted, Lowicryl-embedded
(IEM) specimens of normal rat and human lungs and fibrotic rat lungs.
Fibrosis was induced by intratracheal bleomycin treatment. IH and DIF
showed that VEGF was present in surfactant protein (SP) D-positive
alveolar type II pneumocytes, bronchiolar Clara cells, smooth muscle (SM)
cells, and ␣-SM actin-positive myofibroblasts of normal rat and human
lungs. Fibrotic lesions in bleomycin-treated rat lungs were rich in VEGFpositive cells presenting with a heterogeneous phenotype (mainly SP-Dpositive type II pneumocytes, ␣-SM actin-positive myofibroblasts). There
were no signs of angiogenesis. Post-embedding immunogold labeling using
protein A-gold and IgG-gold technique revealed a specific localization of
VEGF to mitochondria, Clara cell secretory granules, and capillary interendothelial cell junctions. The predominant localization of VEGF to bronchiolar and alveolar epithelial and ␣-SM actin-positive cells, and the marked
increase of VEGF-positive type II pneumocytes and myofibroblasts in
fibrotic lung lesions, indicate that in adult lungs VEGF is involved in
processes other than angiogenesis. Anat Rec 254:61–73, 1999.
r 1999 Wiley-Liss, Inc.
Key words: lung; fibrosis; vascular endothelial growth factor; immunocytochemistry
Vascular endothelial growth factor (VEGF), which was
initially described as vascular permeability factor (VPF)
(Senger et al., 1983), is a dimeric glycoprotein that specifically stimulated endothelial cell migration and proliferation, angiogenesis, and increased vascular permeability
(Dvorak et al., 1995; Klagsbrun and D’Amore, 1996).
Moreover, recent studies presented evidence that VEGF
r 1999 WILEY-LISS, INC.
Grant sponsor: Bundesminister für Bildung, Wissenschaft,
Forschung und Technologie; Grant number: 01ZZ5904.
*Correspondence to: H. Fehrenbach, PhD, Institute of Pathology, University Clinics ‘‘Carl Gustav Carus’’, TU Dresden,
Fetscherstra␤e 74, D-01307 Dresden, Germany. Fax: 49 351 458
4328. E-mail: hefeh@rcs.urz.tu-dresden.de
Received 24 March 1998; Accepted 31 July 1998
62
FEHRENBACH ET AL.
may have important, previously unsuspected roles in cell
types other than the endothelium of blood vessels (Brown
et al., 1997; Ergün et al., 1997; Horiuchi and Weller, 1997;
Rooman et al., 1997; Yang and de Bono, 1997). VEGF
protein and transcript have been shown to be present in
various organs of human and rat origin (Berse et al., 1992;
Monacci et al., 1993; Shifren et al., 1994; Tuder et al.,
1995). VEGF was reported to be very abundant in the lung,
with alveolar epithelial cells being the site of highest
immunohistochemical reactivity and mRNA expression
(Maniscalco et al., 1995; Monacci et al., 1993; Shifren et
al., 1994). Evidence has accumulated that VEGF is involved in some pathogenetic and repair processes (Couffinhal et al., 1997; Maniscalco et al., 1995; Shinohara et al.,
1996; Tolnay et al., 1998; Tuder et al., 1995; Voelkel and
Tuder, 1995). However, to our knowledge nothing is known
about a potential role of VEGF in pulmonary fibrosis.
The bleomycin-treated rat lung is an established experimental model of pulmonary fibrosis (Fine and Goldstein,
1997). The structural and ultrastructural alterations associated with fibrogenesis, which include initial endothelial
injury, proliferation of alveolar type II pneumocytes, increased synthesis and deposition of extracellular materials, and subsequent profound remodeling of the pulmonary parenchyma, have been described in detail (Adamson,
1984; Adamson et al., 1986; Adler et al., 1986; Kuhn et al.,
1989). Immunohistochemical studies contributed to the
understanding of the complex processes of remodeling and
repair by a detailed specification of the various cell types
involved as, e.g., the modulation of fibroblasts into myofibroblasts, or the modulation of structural and membrane
proteins of the alveolar epithelium such as changes in the
cytoskeleton, in adhesion molecules and lectin binding
patterns (Kasper and Haroske, 1996; Kasper and Singh,
1995; Vyalov et al., 1993; Woodcock-Mitchell et al., 1984).
In recent years, the focus of interest has been directed
toward understanding the role of cytokines in pulmonary
fibrosis (Wolff and Crystal, 1997).
To gain a better understanding of the biology of VEGF in
the lung, and of its role in pulmonary fibrosis, our study
aimed at the detailed cellular and subcellular localization
of VEGF to the components of both the gas-exchanging
parenchyma and the gas-conducting tree of rat and human
lungs. Double immunofluorescence staining was used in
conjunction with a number of established cell markers
(Kapanci et al., 1992; Kasper et al., 1995a; Kasper and
Singh, 1995) to characterize the nature of VEGF-immunoreactive cells. And postembedding immunogold labeling
was performed to study the subcellular localization of
VEGF by means of electron microscopy.
MATERIALS AND METHODS
Experimental Animals
Bleomycin-induced pulmonary fibrosis was examined in
female specific pathogen-free Wistar rats (200–300 g body
weight) which received a single intratracheal dose of 7
units of bleomycin sulphate dissolved in 0.9% NaCl (0.25
ml/kg body weight). Animals were sacrificed at 5 days (n ⫽
3), 24 days (n ⫽ 5), 4 weeks (n ⫽ 4), 5 weeks (n ⫽ 4), and 6
weeks (n ⫽ 5) following bleomycin application. The lungs
of two untreated female Wistar rats were used as a control.
Fixation and tissue processing was performed as described
previously (Kasper et al., 1994). Briefly, after sacrifice the
lungs were perfused via the right ventricle with phosphate-
buffered saline (PBS, pH 7.4) prior to fixation of the lower
lobe of the right lung by immersion into 4% formaldehyde
in PBS. Subsequently, routine paraffin embedding was
performed.
Antibodies and Lectins
Two affinity-purified polyconal antibodies, of rabbit and
goat origin, and one monoclonal anti-VEGF antibody were
used (Santa Cruz Biotechnology, Santa Cruz, CA). The
monoclonal antibody was raised against a peptide corresponding to amino acids 1–191 of VEGF of human origin
(with a deletion from amino acids 142–185) and is claimed
to be non-crossreactive with VEGF-B, VEGF-C, or PlGF.
The polyclonal antibodies were raised against a peptide
corresponding to amino acids 1–20 mapping at the amino
terminus of human VEGF. The rabbit antibody was applied at a 1:40 dilution of the original preparation (100 µg
IgG/ml in PBS containing 0.1% sodium azide and 0.2%
gelatin). The goat antibody was diluted to 1:100 in PBS
containing 0.1% acetylated BSA to reduce unspecific background staining. Since the manufacturer offers the control
peptide, preabsorption controls could be performed by
using the anti-VEGF antibodies preincubated with excess
control peptide. A polyclonal rabbit anti-human factor VIII
antibody (DAKO Diagnostika, Hamburg, Germany) was
used to assess vascularization of fibrotic lung lesions by
immunohistochemistry (IH).
For double immunofluorescence labeling, the rabbit
anti-VEGF antibody was used in conjunction with the
following secondary antibodies, lectins, and conjugates: 1)
polyclonal mouse anti-rat alpha-smooth muscle (␣-SM)
actin antibody (Dianova, Hamburg, Germany), used at a
dilution of 1:50; 2) monoclonal mouse antibody E11 (courtesy of Dr. A. Wetterwald, Zürich, Switzerland), which has
been shown by immunoelectron microscopy (IEM) to specifically label alveolar type I pneumocytes in rat lung
(Wetterwald et al., 1996), used undiluted; 3) monoclonal
mouse antibody against surfactant protein D (SP-D) of rat
(Dr. S. Albrecht, Dresden, Germany), which has been
characterized in detail previously (Kasper et al., 1995a),
and was used undiluted; 4) Ulex europaeus agglutinin
(UEA) (Vector Laboratories, Burlinghame, CA, USA), used
at a dilution of 1:10; 5) Lycopersicon esculentum lectin
(LEL) (Vector), used at a dilution of 1:200; 6) Maclura
pomifera agglutinin (MPA) (Vector), used at a dilution of
1:100; 7) anti-rabbit antibody coupled to 4,6-dichlorotriazinyl-aminofluorescein (DTAF) (Dianova), used at a dilution
of 1:40 for fluorescence labeling of VEGF antibody; 8)
Texas red-coupled anti-mouse antibody (Dianova), used at
a dilution of 1:80 for fluorescence labeling of E11 antibody
and antibody against ␣-SM actin; 9) Texas red-coupled
avidin (Vector), used at a dilution of 1:100 for fluorescence
labeling of the lectins UEA, LEL, and MPA.
For indirect IEM, 20 nm gold-labeled goat anti-rabbit
IgG (Dianova) was used to detect primary rabbit antiVEGF antibody. Alternatively, the primary antibody was
detected by means of protein A coupled to 15 nm gold
(kindly provided by Dr. Posthuma, Utrecht).
Fixation and Tissue Processing for IEM
For IEM of rat lung, tissue blocks of 1 mm3 in size were
fixed by immersion in a mixture of 4% paraformaldehyde
and 0.05% glutaraldehyde in 0.1 M cacodylate buffer (pH
7.4) for 60 min at room temperature. After infiltration with
VEGF IN NORMAL AND FIBROTIC ADULT LUNG
2.3 M sucrose in PBS overnight, tissue blocks were frozen
in liquid nitrogen. Subsequently, tissue blocks were transferred to the precooled methanol (⫺80°C) used as the
medium for cryo-substitution. After 2–3 days at ⫺80°C
and two changes of the substitution medium, low temperature embedding in Lowicryl HM20 and UV polymerization
were performed using a CS Auto (Leica, Nussloch, Germany) as described previously (Kasper et al., 1995b).
IH Staining
For VEGF staining, paraffin sections were mounted on
silane-coated slides, dried overnight, dewaxed, and irradiated with microwave in 0.01 M sodium citrate buffer (pH
6.0) at 750 W (twice for 5 min). The staining procedure was
done at room temperature (RT), and comprised the following steps: 1) washing in PBS, 5 min; 2) 0.3% hydrogen
peroxide, 30 min; 3) 50% fetal calf serum in PBS, 30 min; 4)
affinity purified rabbit polyclonal anti-VEGF antibody
(Santa Cruz Biotechnology), 1:40 dilution in PBS or affinity purified goat polyclonal anti-VEGF antibody (Santa
Cruz Biotechnology), 1:100 dilution in PBS containing
0.1% acetylated BSA (Biotrend); 5) washing in PBS, 10
min; 6) peroxidase-coupled goat anti-rabbit IgG (HRP77;
Dr. Grossmann, Dresden), 1:400 dilution in PBS containing 50% fetal calf serum, 60 min, or anti-goat IgG kit
(Vector); 7) washing with PBS, 10 min; 8) diaminobenzidine (DAB), 8 min; 9) washing in PBS, 1 min; 10) counterstaining with hemalaun, 2–3 sec.
For factor VIII staining (C. Lincoln, personal communication), dewaxed paraffin sections were incubated with
polyclonal antibody (DAKO) diluted at 1:500 in PBS for 1
hr (RT), biotinylated anti-rabbit antibody at 1:200 (Vector)
for 30 min, followed by ABC (Vector Elite) at a dilution of
1:100 for 30 min, and finally counterstained with hemalaun.
Double Immunofluorescence Staining
Paraffin sections of normal rat lungs (n ⫽ 2) and of
experimental rat lungs fixed at 24 days (n ⫽ 2) and 5 weeks
(n ⫽ 2) after bleomycin treatment, respectively, were
investigated using the polyclonal rabbit anti-VEGF antibody. Dewaxed and microwave-treated sections were
stained at RT as follows: 1) washing in PBS containing 50
mM glycine, 15 min; 2) PBS containing 0.2% gelatine
(PBG) and 0.5% bovine serum albumin (BSA), twice, 10
min each; 3) primary antibody, 1:40 dilution in PBG, 45
min; 4) rinsing with PBG, four times, 2 min each; 5)
washing with PBS, twice, 2 min each; 6) DTAF-coupled
goat anti-rabbit IgG (Dianova), 1:200 dilution in PBS; 7)
rinsing with PBS, six times, 2 min each; 8) secondary
antibody (see above) in PBG or lectin (see above) in PBS,
45 min unless stated otherwise; 9) rinsing in PBS, six
times, 2 min each; 10) Texas red-coupled goat anti-mouse
IgG (Dianova) or Texas red-coupled avidin (Vector); 11)
washing with PBS, six times, 2 min each; 12) mounting in
glycerol-PBS (9:1) containing 2.5% 1,4-Diazobicyclo2.2.2.Octane (DABCO, Janssen, Beerse, Belgium) to reduce fading of fluorescent dyes. Double-stained sections
were examined with an Olympus BH-2 microscope
equipped with a reflected light fluorescence device (Olympus, Tokyo, Japan). Dye specific fluorescence was selected
with standard fluorescence filter sets (Olympus, Tokyo,
Japan) to demonstrate fluorescein and Texas red staining,
respectively.
63
Immunogold Labeling
For ultrastructural localization of VEGF, ultrathin sections were collected on 200-mesh nickel grids coated with
3% collodion. Labeling was performed on a sheet of Parafilm in a humid chamber at RT unless indicated otherwise,
and comprised: 1) washing in PBS containing 50 mM
glycine, 15 min; 2) PBG containing 0.5% BSA, twice, 10
min each; 3) primary antibody, i.e., polyclonal rabbit
anti-VEGF Ab, 1:20 dilution in PBG, overnight at 8°C; 4)
rinsing with PBG, four times, 2 min each; 5) washing with
PBS, twice, 2 min each; 6) 20 nm gold-coupled goat antirabbit IgG (Dianova), 1:50 dilution in PBS; alternatively,
15 nm gold-coupled protein A (Aurion) was used at a
dilution of 1:50 in PBS; 7) rinsing with PBS, six times, 2
min each; 8) counterstaining with 2% uranyl acetate and
Reynolds lead citrate. Labeled ultrathin sections were
examined with a Zeiss EM 900 operated at 80 kV.
Specificity Controls
Three specificity controls were performed with each
secondary detection system. The primary antibody against
VEGF was substituted by 1) primary antibody absorbed to
control peptide (Santa Cruz Biotechnology); 2) an irrelevant polyclonal rabbit IgG at an equivalent concentration; or 3) primary antibody was omitted. The lectin
binding was specifically blocked by the corresponding
sugar (for details, see Kasper and Singh, 1995).
Western Blotting
Frozen lung tissue was pulverized under liquid nitrogen.
After addition of lysis buffer (20 mM HEPES, pH7.9, 400
mM NaCl, 1mM EGTA, 1 mM EDTA, 1% Triton X-100,
0.5% sodium desoxycholate, 0.5% NP-40, 1 mM DTT, 0.5
mM PMSF) the suspension was homogenized for 1 min
using an Ultraturrax (IKA Labortechnik, Staufen, Germany). After an incubation time of 10 min, cellular debris
was pelleted at 12,000g for 5 min. The supernatant was
stored at -80°C. Proteins were fractionated with denaturing polyacrylamide electrophoresis (Laemmli, 1970). Proteins were blotted onto nylon membranes (Pall, Dreieich,
Germany). Polyclonal rabbit anti-VEGF antibody was
used at a dilution of 1:220. A horseradish peroxidaselabeled donkey anti-rabbit secondary antibody (Amersham, Braunschweig, Germany) was used at a dilution of
1:1000. Super Signal Ultra chemiluminescent substrate
(Pierce, via Merck, Dresden, Germany) was applied according to the manufacturer’s instructions. Membranes were
exposed to Hyperfilm-ECL (Amersham) for 5–10 min.
RESULTS
Immunohistochemistry
Irrespective of the antibody used, IH staining of paraffinembedded normal rat and human lung, revealed the
presence of VEGF in cells not only of the gas-exchanging
parenchyma, but also of the gas-conducting bronchioles
(Fig. 1A–D). Bronchiolar staining comprised the nonciliated epithelial Clara cells and smooth muscle cells of the
peribronchiolar arterioles. The bronchiolar tunica muscularis exhibited moderate staining in human lung and only
a trace amount of staining in rat lung. Endothelial cells
were devoid of immunohistochemical reaction product.
The most prominent alveolar staining was seen in type II
pneumocytes (Fig. 1C,D). Alveolar macrophages exhibited
Figure 1.
65
VEGF IN NORMAL AND FIBROTIC ADULT LUNG
TABLE 1. Immunohistochemical labeling pattern of rat pulmonary cells during
the development of bleomycin-induced fibogenesis
Time after
treatment
Cell type
Gas-conducting region
Bronchial epithelium
ciliated
nonciliated
Vascular endothelium
Smooth muscle cells
Gas-exchanging region
Alveolar epithelium
type I cells
type II cells
Alveolar macrophages
Interstitial cells
Capillary endothelium
Control
(n ⫽ 2)
5 days
(n ⫽ 3)
24 days
(n ⫽ 5)
4 weeks
(n ⫽ 4)
5 weeks
(n ⫽ 4)
6 weeks
(n ⫽ 5)
⫺
⫹⫹
⫺
⫹
⫺
⫹⫹
⫺
⫹
⫺
⫹⫹
(⫺)
⫹
⫺
⫹⫹
(⫺)
⫹
⫺
⫹⫹
(⫺)
⫹
⫺
⫹⫹
(⫺)
⫹
(⫺)
⫹⫹
(⫹)
(⫹)
(⫺)
(⫺)
⫹⫹/>
⫹/>
(⫹)
(⫺)
(⫺)
⫹⫹/>
⫹/>
⫹/⫹⫹>
(⫺)
(⫺)
⫹⫹/>
⫹/>
⫹/⫹⫹>
(⫺)
(⫺)
⫹⫹/>
⫹/>
⫹/⫹⫹>
(⫺)
(⫺)
⫹⫹/>
⫹/>
⫹/⫹⫹>
(⫺)
⫺ ⫽ Not stained.
(⫺) ⫽ Weak staining of single cells cannot be excluded.
(⫹) ⫽ Weak to moderate staining restricted to some cells.
⫹ ⫽ Weak to moderate, generalized staining.
⫹⫹ ⫽ Strong, generalized staining.
> ⫽ Cell proliferation.
/ ⫽ In normal/fibrotic regions, respectively.
only weak if any staining for VEGF in rat lungs (Fig. 1C),
but were regularly stained in the human lung specimens
(Fig. 1D).
In the experimental rat lungs treated with bleomycin,
interstitial fibrotic regions were observed in the alveolar
parenchyma 24 days or more after treatment. Notably,
fibrotic regions exhibited strong immunoreactivity for
VEGF (Fig. 1E,F,G; Table 1). In the early phase, characterized by hyperplasia of the alveolar epithelium, type II
pneumocytes were easily identifiable (Fig. 1E). With progressing fibrosis, however, interstitial fibrotic lesions were
densely populated by VEGF-positive cells, which could not
unequivocally be ascribed to a distinct cell type (Fig. 1F,G).
There were no signs of increased vascularization of the
fibrotic lesions as indicated by factor VIII staining (not
shown).
Immunofluorescence Microscopy
The staining patterns observed by double immunofluorescence labeling experiments are summarized in Table 2.
Fig. 1. Abbreviations for figures: A: alveolar space; AM: alveolar
macrophage; BE: bronchiolar epithelium; C: capillary lumen; E: endothelium; G: secretory granule; L: lamellar body; M: mitochondria; Mu: tunica
muscularis; Nu: nucleus; P1: type I pneumocyte; P2: type II pneumocyte;
PL: pleura; SM: smooth muscle cell; V: vessel. IH staining for VEGF of
normal (A, C) and bleomycin-treated rat lung (E, F) 24 days and (G, H) 6
weeks after treatment, and of normal (B, D) human lung. In the
gas-conducting system (A, B, G), VEGF-staining is seen in Clara cells
(arrowheads) and smooth muscle cells of the tunica muscularis (Mu). In
normal pulmonary parenchyma (C, D), VEGF-staining is prominent in
alveolar type II pneumocytes (P2); note difference in staining intensity
between rat and human alveolar macrophages (AM). In fibrotic rat lung 24
days after bleomycin-treatment (E), metaplasia of alveolar epithelium with
numerous VEGF-positive type II pneumocytes is seen; note focal staining
of thin leaflet of transformed alveolar epithelium (arrowhead) and of
capillary endothelium (arrow). Areas of interstitial fibrotic lesions (F, G)
present with numerous, heterogeneous VEGF-staining cells. Staining is
completely abolished by preincubation of the antibody with control peptide
(H). Scale bar represents 50 µm (A–F), and 100 µm (G, H).
Double staining for VEGF and Ulex europaeus agglutinin
(UEA) confirmed the absence of VEGF from vascular
endothelial cells of normal rat lung specimens (Figs. 2A,
4A,B). Double staining with anti-VEGF antibody and
Lycopersicon esculentum lectin (LEL) confirmed the presence of VEGF-immunoreactivity in surfactant protein D
(SP-D)-positive Clara cells and its absence from ciliated
cells of the bronchiolar epithelium (Figs. 2D, 3A,E). Looking at the parenchyma, double immunofluorescence staining for SP-D and VEGF revealed that far more alveolar
cells stained positive for VEGF than for SP-D, which is
specific for type II pneumocytes (Fig. 2C). Double staining
for VEGF and the type I pneumocyte-specific monoclonal
antibody E11 indicated that VEGF was absent from the
thin type I cell leaflets of the air–blood-barrier (Fig. 2B).
Double staining for VEGF and Maclura pomifera agglutinin (MPA), which also labels the entire population of
alveolar macrophages (Kasper et al., 1994), confirmed that
in the rat VEGF was weakly present in this cell type (not
shown). Colocalization of ␣-SM actin and VEGF was seen
around small vessels (Fig. 3B,F), and at alveolar entrance
rings (Fig. 3C,G), which are known to contain ␣-SM actin
positive pericytes and ring muscles, respectively (Kapanci
et al., 1992). In all cell types, staining for VEGF was largely
cytoplasmatic and frequently of granular appearance.
In the experimental lungs, double immunofluorescence
staining revealed a remarkable heterogeneity of VEGFimmunoreactive cells in fibrotic regions. The population
comprised cells that stained for SP-D (Fig. 2E,F), and for
␣-SM actin (Fig. 3D,H), respectively. Flattened epithelial
cells lining the alveolar wall stained more intensely for
SP-D (Fig. 2E) than those cells within fibrotic lesions (Fig.
2F). While cells staining for the lectins MPA and UEA also
contributed to the VEGF-positive cell population in fibrotic
regions, cells double-labeled for the type I pneumocytespecific antibody E11 did not (data not shown). Double
labeling for VEGF and UEA revealed that, in contrast to
control lungs, individual vascular endothelial cells of
fibrotic rat lungs were VEGF-immunoreactive (Fig. 4C,D).
66
FEHRENBACH ET AL.
TABLE 2. Results of immunofluorescence double labeling of normal rat lung
Cell marker
Cell type
Gas-conducting region
Bronchial epithelium
ciliated
nonciliated
Vascular endothelium
Smooth muscle cells
Gas-exchanging region
Alveolar macrophages
Alveolar epithelium
type I cells
type II cells
Interstitial cells
Myofibroblasts
Pericytes
Capillary endothelium
Antibodies
Lectins
VEGF
E11
SP-D
␣-SMA
LEL
MPA
UEA
⫺
⫹⫹
⫺
⫹
n.e.
n.e.
n.e.
n.e.
⫺
⫹⫹
⫺
⫺
⫺
⫺
⫺
⫹⫹
⫹⫹
⫺
⫺
⫺
⫹
⫹
⫺
⫺
⫹
⫹
⫹⫹
⫺
(⫹)
⫺
(⫹)
⫺
⫺
⫹⫹
⫹
⫺
⫹⫹
⫹
⫺
⫺
⫹⫹
⫺
⫺
⫹⫹
⫺
⫺
⫹⫹
⫹
⫹
(⫹)
(⫹)
⫺
⫺
⫺
⫺
⫺
⫺
⫺
⫹
⫹
⫺
n.e.
n.e.
⫺
n.e.
n.e.
⫺
n.e.
n.e.
⫹
⫺ ⫽ Not labeled.
(⫹) ⫽ Labeling not generalized.
⫹ ⫽ Labeling generalized, but weak.
⫹⫹ ⫽ Strong generalized labeling.
n.e. ⫽ Not specifically examined.
Immunoelectron Microscopy
Indirect immunogold labeling of freeze-substituted parenchymal tissues embedded in Lowicryl HM20 with polyclonal rabbit anti-VEGF antibody revealed a predominant
staining of mitochondria of type II pneumocytes, both in
rat (Fig. 5A–C) and human lung (Fig. 5D). Identical
labeling patterns were seen using gold-coupled IgG (Fig.
5A,D) or protein A-gold (Fig. 5C) to detect the primary
antibody. Few gold granules were observed over the
nucleus, and even less over endoplasmic reticulum, cytosol, and the surrounding extracellular matrix. Staining
was absent from the surfactant storing lamellar bodies.
However, gold particles were not restricted to alveolar type
II pneumocytes, but were observed to be present over
mitochondria of vascular smooth muscle cells (not shown),
bronchiolar Clara cells (Fig. 6A), and of virtually every cell
type of the alveolar septum. The Clara cells were characterized by additional specific labeling of their secretory
granules (Fig. 6A), and the capillary endothelium was
unique in that VEGF-labeling additionally localized to
intercellular junctions (Fig. 6B). In control sections, which
were concurrently performed with every labeling sequence, only slight unspecific labeling was seen (Fig. 5B).
Western Blotting
Based on Western blots of total lung protein extracts,
which were performed using the polyclonal rabbit antibody against VEGF, we could show that a single protein
band was present at about 22 to 26 kD (Fig. 7). This
corresponds to Western blot data reported from extracts of
human testis and seminiferous tubules, which were characterized by a protein band at 24 kD (Ergün et al., 1997). A
second protein band at 49 kD indicative of a dimeric form
of VEGF in human testis, was not observed in the extracts
of rat lung tissue.
DISCUSSION
The present study investigated the differential distribution of VEGF by means of cell targeting using double
immunofluorescence in normal and fibrotic rat and normal
human lung. The intracellular localization of VEGF was
examined by IEM. A remarkable heterogeneity of the
VEGF-immunoreactive cell population was observed by
double immunofluorescence microscopy both in normal
and fibrotic lungs.
VEGF in Normal Adult Lungs
A number of studies have shown that VEGF is abundantly present in alveolar type II pneumocytes of normal
adult lungs (Christou et al., 1998; Maniscalco et al., 1995,
1997; Monacci et al., 1993; Shifren et al., 1994; Tuder et al.,
1995). In our study, VEGF-immunoreactivity was observed not to be restricted to this alveolar cell type, but to
be present in bronchiolar Clara cells, smooth muscle cells,
and alveolar myofibroblasts as well, while ciliated bronchiolar epithelial cells and vascular endothelial cells did not
stain. These observations extend previous reports of the
presence of VEGF in pulmonary cell types other than
alveolar type II pneumocytes. The results are in line with
the studies of Monacci et al. (1993), who, from in situ
hybridization experiments, reported that VEGF mRNA
was present in virtually every pulmonary alveolar cell of
adult rat lung. Shifren et al. (1994) in their study of fetal
and adult human tissues observed VEGF-immunoreactivity in pulmonary epithelial cells, myocytes (including
smooth muscle cells of vessels), but not in vascular endothelial cells. Tuder et al. (1995) reported VEGF mRNA to be
present in type II pneumocytes and alveolar macrophages
in control rats, while hypoxia induced additional expression in bronchiolar epithelial cells and in vascular smooth
muscle cells. In contrast, Maniscalco et al. (1995), studying
rabbit tissues, reported that VEGF mRNA and peptide
were enriched in type II pneumocytes, but absent from
alveolar macrophages, fibroblasts, and endothelial cells.
Further, these authors reported only little or no VEGFexpression in large vessel endothelial cells, airway cells,
smooth muscle cells, and type I pneumocytes. We have
shown that VEGF is present in ␣-SM actin-positive pericytes and vascular smooth muscle cells of normal adult
Fig. 2. Double immunofluorescence staining of normal (A–D) and
fibrotic (E, F) rat lung. Green (fluorescein staining) indicates VEGFimmunoreactivity; red (Texas Red staining) indicates UEA (A), E11 (B),
and SP-D (C, D) labeling; orange to yellow indicates colocalization of both
markers. (A) While only smooth muscle cells stain for VEGF (green), with
some cells also staining for UEA (yellow), the endothelial cells are only
positive for UEA (arrowheads). (B) VEGF is seen in type II pneumocyte
(arrowheads), but is absent from E11-labeled type I pneumocytes (red).
(C) Type II pneumocytes staining for SP-D are always VEGF-positive as
indicated by yellow to orange fluorescence, while numerous other cells of
the alveolar septum only stained for VEGF (green). (D) Clara cells stained
positively for both VEGF and SP-D. (E, F) After 24 days of bleomycintreatment, flattened alveolar epithelial cells stained strongly for SP-D and
VEGF (E), while only weak staining for SP-D was seen in cells within
fibrotic lesions (F). Scale bar ⫽ 100 µm (A, C), and 20 µm in (B, D–F).
68
FEHRENBACH ET AL.
Figure 3.
VEGF IN NORMAL AND FIBROTIC ADULT LUNG
69
Fig. 4. Double immunofluorescence staining of arterial vessel wall of
normal (A, B) and bleomycin-treated (C, D) rat lung to demonstrate
immunoreactivity for VEGF (A, C) and UEA (B, D), respectively. While in
control lung tissue UEA-positive endothelium (B) is devoid of VEGF-
immunoreactivity (A), individual UEA-positive vascular endothelial cells
(D) were seen in fibrotic lungs to be immunoreactive for VEGF (arrowheads in C). Arterial smooth muscle cells (SM) were immunoreactive for
VEGF in both normal and fibrotic lungs. Scale bar ⫽ 20 µm.
lungs, which is in line with in situ hybridization studies
performed to localize VEGF mRNA (Shifren et al., 1994;
Tuder et al., 1995). Although the precise role of VEGF in
these cell types still remains to be determined, one may
follow Senger et al. (1993), who proposed that vascular
smooth muscle-VEGF might be involved in the maintenance of normal vascular function.
In general, VEGF-staining has been reported to be
cytoplasmatic (Couffinhal et al., 1997; Ergün et al., 1997;
Shifren et al., 1994). However, Park et al. (1993) presented
evidence that there may be differences in the localization
of the different isoforms. By means of a preembedding
immunogold technique and electron microscopy, immuno-
reactivity for VEGF189/206 has been localized to extracellular matrix-like material in transfected human embryonic
kidney CEN4 cells expressing the respective isoform (Park
et al., 1993). Using postembedding immunogold and protein A-gold techniques, we could demonstrate by electron
microscopy that VEGF-immunoreactivity was enriched in
mitochondria. Specificity of the labeling was confirmed by
preabsorption of the primary antibody to the peptide used
for immunization, which always abolished mitochondrial
staining. Labeling was most prominent in type II pneumocytes and bronchiolar Clara cells, but was observed in
virtually every cell type. This again corresponds to in situ
hybridization studies showing the widespread presence of
VEGF mRNA in rat lung (Monacci et al., 1993). While the
strong immunoreactivity of the secretory granules of Clara
cells may indicate that VEGF is released along this
secretory pathway, the significance of the high level of
VEGF-immunoreactivity in mitochondria remains unclear. By analogy, another growth factor, TGF-␤1, has been
localized to mitochondria of rat and mouse cardiac myocytes and rat hepatocytes (Heine et al., 1991). As with
VEGF, the functional role of TGF-␤1 in mitochondria is
still unknown. One may speculate that mitochondria
represent a potential intracellular site of phosphorylation.
In agreement with previous studies (Maniscalco et al.,
1995; Shifren et al., 1994; Tuder et al., 1995), vascular
Fig. 3. Double immunofluorescence staining of normal (A–C, E–G)
and bleomycin-treated (D, H) rat lung. (A–D) VEGF immunofluorescence
labeling with corresponding staining for (E) LEL, and (F–H) ␣-SM actin.
(A, E) Intense fluorescence of VEGF-positive granules is confined to
apical portion of Clara cells (arrowheads). (B, F) VEGF-positive staining is
seen to colocalize with ␣-SM actin-positive pericytes at capillary-venular
junction (arrowheads); asterisks indicate two VEGF-positive type II
pneumocytes. (C, G) VEGF-positive, ␣-SM actin-positive myofibroblasts
at an alveolar entrance ring. (D, H) Fibrotic lesions contain VEGF-positive,
␣-SM actin-positive cells (black arrowheads), VEGF-positive, ␣-SM actinnegative cells (arrow) as well as VEGF-negative, ␣-SM actin-positive cells
(white arrowhead). Scale bar ⫽ 10 µm (A–C, E–G), and 50 µm (D, H).
70
FEHRENBACH ET AL.
Fig. 5. Indirect immunogold labeling of cryo-substituted, Lowicrylembedded rat (A, B, C) and human (D) lung. Mitochondria (M) of alveolar
type II pneumocytes are specifically stained by the affinity-purified, polyclonal
rabbit anti-VEGF antibody, while lamellar bodies (L) are devoid of labeling
irrespective of using gold-conjugated anti-rabbit IgG (A, D) or proteinA-gold (C)
as secondary detection system. Staining of mitochondria and interendothelial
cell junctions (arrowhead) is not seen with the primary VEGF-antibody being
preabsorbed to control peptide (B). Scale bars ⫽ 1 µm.
endothelial cells of normal rat lungs did not stain for
VEGF by IH. However, IEM revealed discrete immunogold
labeling for VEGF of interendothelial cell junctions of
alveolar capillaries in normal rat lungs. This specific
localization of VEGF to a structure actively involved in the
regulation of capillary permeability may indicate a poten-
VEGF IN NORMAL AND FIBROTIC ADULT LUNG
71
Fig. 6. Indirect IgG-gold labeling of cryo-substituted, Lowicryl embedded normal rat lung. Beside mitochondrial
staining, specific labeling was seen (A) in secretory granules (G) of bronchiolar Clara cells, and (B) to be associated
with interendothelial junctions (arrowheads) of capillary endothelium (E). Scale bars ⫽ 1 µm.
a dose- and time-dependent increase in permeability (Wang
et al., 1996). In the frog mesentery, exposure to 1 nM
VEGF rapidly and transiently increased microvessel hydraulic conductivity within 30 sec and returned to control
within 2 min (Bates and Curry, 1996).
VEGF in Pulmonary Fibrosis
Fig. 7. Western blot of protein extract of total lung tissue. A single
protein band at about 24 kD was detected by means of the rabbit
polyclonal antibody against VEGF used in most of the immunostaining
experiments.
tial role of VEGF in the regulation of normal capillary
function. While the involvement of VEGF in the induction
of hyperpermeability of tumor-associated neovessels is
widely accepted (Dvorak et al., 1995; Feng et al., 1996; Qu
et al., 1995; Senger et al., 1993), evidence supporting a role
for VEGF in the physiological regulation of vascular
permeability has been presented only recently. In cultured
bovine brain microvessel endothelial cells, VEGF induced
In fibrotic regions of bleomycin-treated rat lungs, VEGFimmunoreactive cells were abundantly present. By double
immunofluorescence microscopy, alveolar type II pneumocytes as identified by their staining for surfactant protein
D (SP-D), and myofibroblasts as identified by ␣-SM actinstaining, were prominent contributors to the VEGFimmunoreactive population of cells in rat lung fibrotic
regions. Notably, SP-D staining of cells within fibrotic
regions was less intense compared with cells facing the
alveolar air space. This may correspond to the finding that
type II pneumocytes containing abundant VEGF mRNA
have little if any SP-C mRNA, and vice versa (Maniscalco
et al., 1995). Early morphometric studies have shown that
at 28 days after bleomycin-instillation, the largest increase in any cell population was in contractile interstitial
cells, which have subsequently been shown to be predominantly ␣-SM actin-positive myofibroblasts (Adler et al.,
1986, 1989; Vyalov et al., 1993). Proliferation of alveolar
myofibroblasts was observed as early as 24 hr to 3 days
after intratracheal bleomycin administration, and preceded the formation of collagen fibers (Vyalov et al., 1993).
Knowing that hypoxia is a potent pathological stimulus for
the upregulation of VEGF (Namiki et al., 1995; Stavri et
al., 1995; Tuder et al., 1995), the increase in VEGF-positive
cells in fibrotic lung regions may be a response to an
impaired oxygen supply of the thickened fibrotic septa.
VEGF was shown to induce several rapid cellular responses in cultured human endothelial cells as, e.g., a
72
FEHRENBACH ET AL.
twofold increase in the release of Von Willebrand factor
(vWF) (Brock et al., 1991). In radiation-induced pulmonary fibrosis, the number of vWF-positive cells per parenchymal unit decreased during the first 2 months after
irradiation and showed a marked increase thereafter
(Kasper et al., 1996). The fact that we could not observe a
similar increase in vascularization in our bleomycin model
may be explained by the shorter period of 6 weeks of
observation in this study. Recently, VEGF and one of its
receptors (flt-1) have been demonstrated to be elevated in
activated macrophages in pulmonary sarcoidosis (Tolnay
et al., 1998). Further, VEGF has been shown to act as a
chemoattractant of mast cells (Gruber et al., 1995), which
are well known to immigrate into fibrotic lung lesions
(Goto et al., 1984). Thus, the elevated level of VEGFpositive type II pneumocytes and myofibroblasts may
induce immigration and aggregation of activated macrophages and mast cells in fibrotic lung lesions.
We conclude that the predominant localization of VEGFimmunoreactivity to bronchiolar and alveolar epithelial
cells, and to ␣-SM actin-positive cells in adult rat and
human lungs, and the marked increase of VEGF-positive
cells in the absence of an induction of vascularization in
fibrotic lesions of bleomycin-treated rat lungs, indicate
that VEGF exerts its action in processes other than
angiogenesis in the adult lung.
ACKNOWLEDGMENTS
The expert technical assistance of I. Peterson, B. Rudolph, S. Petzold, S. Langer, and H. Seidel is acknowledged
with thanks. Professor Dr. K. Kayser (Thoraxclinic, University of Heidelberg, Germany) kindly provided the human
material used in this study. We thank Professor Dr. K.-W.
Wenzel and Dr. R. Koslowski (Institute of Physiological
Chemistry, TU Dresden, Germany) who provided the
bleomycin rat lung model.
REFERENCES
Adamson IYR. 1984. Drug-induced pulmonary fibrosis. Environ Health
Perspect 55:25–36.
Adamson IYR, Orr FW, Young L. 1986. Effects of injury and repair of
the pulmonary endothelium on lung metastasis after bleomycin. J
Pathol 150:279–287.
Adler KB, Callahan LM, Evans JN. 1986. Cellular alterations in the
alveolar wall in bleomycin-induced pulmonary fibrosis in rats. An
ultrastructural morphometric study. Am Rev Respir Dis 133:1043–
1048.
Adler KB, Low RB, Leslie KO, Mitchell J, Evans JN. 1989. Contractile
cells in normal and fibrotic lung. Lab Invest 60:473–485.
Bates DO, Curry FE. 1996. Vascular endothelial growth factor increases hydraulic conductivity of isolated perfused microvessels. Am
J Physiol 271:H2520–H2528.
Berse B, Brown LF, Van de Water L, Dvorak HF, Senger DR. 1992.
Vascular permeability factor (vascular endothelial growth factor)
gene is expressed differentially in normal tissues, macrophages, and
tumors. Mol Biol Cell 3:211–220.
Brock TA, Dvorak HF, Senger DR. 1991. Tumor-secreted vascular
permeability factor increases cytosolic Ca2⫹ and von Willebrand
factor release in human endothelial cells. Am J Pathol 138:213–221.
Brown LF, Detmar M, Tognazzi K, Abu-Jawdeh G, Iruela-Arispe ML.
1997. Uterine smooth muscle cells express functional receptors (flt-1
and KDR) for vascular permeability factor/vascular endothelial
growth factor. Lab Invest 76:245–255.
Christou H, Yoshida A, Arthur V, Morita T, Kourembanas S. 1998.
Increased vascular endothelial growth factor production in the
lungs of rats with hypoxia-induced pulmonary hypertension. Am J
Respir Cell Mol Biol 18:768–776.
Couffinhal T, Kearney M, Witzenbichler B, Chen D, Murohara T,
Losordo DW, Symes J, Isner JM. 1997. Vascular endothelial growth
factor/vascular permeability factor (VEGF/VPF) in normal and
artheriosclerotic human arteries. Am J Pathol 150:1673–1685.
Dvorak HF, Brown LF, Detmar M, Dvorak AM. 1995. Vascular
permeability factor/vascular endothelial growth factor, microvascular hyperpermeability, and angiogenesis. Am J Pathol 146:1029–
1039.
Ergün S, Kilic N, Fiedler W, Mukhopadhyay AK. 1997. Vascular
endothelial growth factor and its receptors in normal human
testicular tissue. Mol Cell Endocrinol 131:9–20.
Feng D, Nagy JA, Hipp J, Dvorak HF, Dvorak AM. 1996. Vesiculovacuolar organelles and the regulation of venule permeability to
macromolecules by vascular permeability factor, histamine, and
serotonin. J Exp Med 183:1981–1986.
Fine A, Goldstein RH. 1997. Animal models of pulmonary fibrosis. In:
The Lung. Scientific Foundations. Crystal RG, West JB, Barnes PJ,
Weibel ER, editors. Philadelphia, New York: Lippincott/Raven Publishers. p 2525–2536.
Goto T, Befus D, Low R, Bienenstock J. 1984. Mast cell heterogeneity
and hyperplasia in bleomycin-induced pulmonary fibrosis of rats.
Am Rev Respir Dis 130:797–802.
Gruber BL, Marchese MJ, Kew R. 1995. Angiogenic factors stimulate
mast-cell migration. Blood 86:2488–2493.
Heine UI, Burmester JK, Flanders KC, Danielpour D, Munoz EF,
Roberts AB, Sporn MB. 1991. Localization of transforming growth
factor-beta 1 in mitochondria of murine heart and liver. Cell Regul
2:467–477.
Horiuchi T, Weller PF. 1997. Expression of vascular endothelial
growth factor by human eosinophils: Upregulation by granulocyte
macrophage colony-stimulating factor and interleukin-5. Am J
Respir Cell Mol Biol 17:70–77.
Kapanci Y, Ribaux C, Chaponnier C, Gabbiani G. 1992. Cytoskeletal
features of alveolar myofibroblasts and pericytes in normal human
and rat lung. J Histochem Cytochem 40:1955–1963.
Kasper M, Haroske G. 1996. Alterations in the alveolar epithelium
after injury leading to pulmonary fibrosis. Histol Histopathol 11:463–
483.
Kasper M, Singh G. 1995. Epithelial lung cell marker: Current tools
for cell typing. Histol Histopathol 10:155–169.
Kasper M, Sakai K, Koslowski R, Wenzel KW, Haroske G, Schuh D,
Müller M. 1994. Localization of surfactant protein A (SP-A) in
alveolar macrophage subpopulations of normal and fibrotic rat lung.
Histochemistry 102:345–352.
Kasper M, Albrecht S, Grossmann H, Grosser M, Schuh D, Müller M.
1995a. Monoclonal antibodies to surfactant protein D: Evaluation of
immunoreactivity in normal rat lung and in a radiation-induced
fibrosis model. Exp Lung Res 21:577–588.
Kasper M, Huber O, Grossmann H, Rudolph B, Tränkner C, Müller M.
Immunocytochemical distribution of E-cadherin in normal and
injured lung tissue of the rat. 1995b. Histochem Cell Biol 104:383–
390.
Kasper M, Schöbl R, Haroske G, Fischer R, Neubert F, Dimmer V,
Müller M. 1996. Distribution of von Willebrand factor in capillary
endothelial cells of rat lungs with pulmonary fibrosis. Exp Toxicol
Pathol 48:283–288.
Klagsbrun M, D’Amore PA. 1996. Vascular endothelial growth factor
and its receptors. Cytokine Growth Factor Rev 7:259–270.
Kuhn C, Boldt J, King TE Jr, Crouch E, Vartio T, McDonald JA. 1989.
An immunohistochemical study of architectural remodeling and
connective tissue synthesis in pulmonary fibrosis. Am Rev Respir
Dis 140:1693–1703.
Laemmli UK. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680–685.
Maniscalco WM, Watkins RH, Finkelstein JN, Campbell MH. 1995.
Vascular endothelial growth factor mRNA increases in alveolar
epithelial cells during recovery from oxygen injury. Am J Respir Cell
Mol Biol 13:377–386.
Maniscalco WM, Watkins RH, D’Angio CT, Ryan RM. 1997. Hyperoxic
injury decreases alveolar epithelial cell expression of vascular
endothelial growth factor (VEGF) in neonatal rabbit lung. Am J
Respir Cell Mol Biol 16:557–567.
VEGF IN NORMAL AND FIBROTIC ADULT LUNG
Monacci WT, Merrill MJ, Oldfield EH. 1993. Expression of vascular
permeability factor/vascular endothelial growth factor in normal rat
tissues. Am J Physiol 264:C995–C1002.
Namiki A, Brogi E, Kearney M, Kim EA, Wu T, Couffinhal T,
Varticovski L, Isner JM. 1995. Hypoxia induces vascular endothelial
growth factor in cultured human endothelial cells. J Biol Chem
270:31189–31195.
Park JE, Keller GA, Ferrara N. 1993. The vascular endothelial growth
factor (VEGF) isoforms: Differential deposition into the subepithelial extracellular matrix and bioactivity of extracellular matrixbound VEGF. Mol Biol Cell 4:1317–1326.
Qu H, Nagy JA, Senger DR, Dvorak HF, Dvorak AM. 1995. Ultrastructural localization of vascular permeability factor/vascular endothelial growth factor (VPF/VEGF) to the abluminal plasma membrane
and vesiculovacuolar organelles of tumor microvascular endothelium. J Histochem Cytochem 43:381–389.
Rooman I, Schuit F, Bouwens L. 1997. Effect of vascular endothelial
growth factor on growth and differentiation of pancreatic ductal
epithelium. Lab Invest 76:225–232.
Senger DR, Galli SJ, Dvorak AM, Perruzzi CA, Harvey VS, Dvorak
HF. 1983. Tumor cells secrete a vascular permeability factor that
promotes accumulation of ascites fluid. Science 219:983–985.
Senger DR, Van De Water L, Brown LF, Nagy JA, Yeo KT, Yeo TK,
Berse B, Jackman RW, Dvorak AM, Dvorak HF. 1993. Vascular
permeability factor (VPF, VEGF) in tumor biology. Cancer Metastasis Rev. 12:303–324.
Shifren JL, Doldi N, Ferrara N, Mesiano S, Jaffe RB. 1994. In the
human fetus, vascular endothelial growth factor is expressed in
epithelial cells and myocytes, but not vascular endothelium: Implications for mode of action. J Clin Endocrinol Metab 79:316–322.
Shinohara K, Shinohara T, Mochizuki N, Mochizuki Y, Sawa H, Kohya
T, Fujita M, Fujioka Y, Kitabatake A, Nagashima K. 1996. Expression of vascular endothelial growth factor in human myocardial
infarction. Heart Vessels 11:113–122.
Stavri GT, Zachary IC, Baskerville PA, Martin JF, Erusalimsky JD.
73
1995. Basic fibroblast growth factor upregulates the expression of
vascular endothelial growth factor in vascular smooth muscle cells.
Synergistic interaction with hypoxia. Circulation 92:11–14.
Tolnay E, Kuhnen C, Voss B, Wiethege T, Müller KM. 1998. Expression
and localization of vascular endothelial growth factor and its
receptor flt in pulmonary sarcoidosis. Virchows Arch 432:61–65.
Tuder RM, Flook BE, Voelkel NF. 1995. Increased gene expression for
VEGF and the VEGF receptors KDR/Flk and Flt in lungs exposed to
acute or to chronic hypoxia. Modulation of gene expression by nitric
oxide. J Clin Invest 95:1798–1807.
Voelkel NF, Tuder RM. 1995. Cellular and molecular mechanisms in
the pathogenesis of severe pulmonary hypertension. Eur Respir J.
8:2129–2138.
Vyalov SL, Gabbiani G, Kapanci Y. 1993. Rat alveolar myofibroblasts
acquire alpha-smooth muscle actin expression during bleomycininduced pulmonary fibrosis. Am J Pathol 143:1754–1765.
Wang W, Merrill MJ, Borchardt RT. 1996. Vascular endothelial growth
factor affects permeability of brain microvessel endothelial cells in
vitro. Am J Physiol 271:C1973–C1980.
Wetterwald A, Hoffstetter W, Cecchini MG, Lanske B, Wagner C,
Fleisch H, Atkinson M. 1996. Characterization and cloning of the
E11 antigen, a marker expressed by rat osteoblasts and osteocytes.
Bone 18:125–132.
Wolff G, Crystal RG. 1997. Biology of pulmonary fibrosis. In: The
Lung. Scientific Foundations. Crystal RG, West JB, Barnes PJ,
Weibel ER, editors. Philadelphia, New York: Lippincott/Raven Publishers. p 2509–2524.
Woodcock-Mitchell J, Adler KB, Low RB. 1984. Immunohistochemical
identification of cell types in normal and in bleomycin-induced
fibrotic rat lung. Cellular origins of interstitial cells. Am Rev Respir
Dis 130:910–916.
Yang W, de Bono DP. 1997. A new role for vascular endothelial growth
factor and fibroblast growth factors: Increasing endothelial resistance to oxidative stress. Febs Lett 403:139–142.
Документ
Категория
Без категории
Просмотров
4
Размер файла
1 310 Кб
Теги
625
1/--страниц
Пожаловаться на содержимое документа