Department of Physiology and Cell Biology, Albany Medical College, Albany, New York 12208
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ABSTRACT |
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Exposure of confluent pulmonary arterial
endothelial monolayers to tumor necrosis factor (TNF)- causes both a
reorganization and/or disruption of fibronectin (Fn) in
the extracellular matrix and an increase in transendothelial protein
permeability. However, the factors initiating this response to TNF-
have not been defined. Because TNF-
can induce proteinase expression
in endothelial cells, we determined whether proteinases cause both the
alteration of the Fn matrix and the permeability increase as is often
speculated. Incubation of calf pulmonary arterial endothelial
monolayers with TNF-
(200 U/ml) for 18 h caused a disruption of the
Fn matrix and an increase in transendothelial protein permeability. A
reduced colocalization of cell-surface
5
1-Fn
integrins with the Fn fibers in focal contacts was also observed.
TNF-
treatment of endothelial monolayers with matrices prelabeled
with 125I-human Fn (hFn) did not
cause the release of Fn fragments or alter the content of Fn antigen in
the medium as analyzed by SDS-PAGE coupled with autoradiography. Both
the content and fragmentation pattern of Fn within the cell layer and
the insoluble Fn matrix also appeared unchanged after TNF-
exposure
as confirmed by Western immunoblot. Fn-substrate zymography revealed
that TNF-
increased the expression of two proteinases within the
conditioned medium in which activity could be blocked by aprotinin but
not by EDTA, 1,10-phenanthroline, leupeptin, or pepstatin. However,
inhibition of the Fn proteolytic activity of these two serine
proteinases did not prevent either the TNF-
-induced disruption of
the Fn matrix or the increase in permeability. Thus the reorganization and/or disruption of the Fn matrix and the temporally associated increase in endothelial permeability caused by TNF-
appear not to be
due to proteolytic degradation of Fn within the extracellular matrix.
In contrast, decreased
5
1-Fn
integrin interaction with Fn fibers in the matrix may be important in
the response to TNF-
exposure.
lung vascular permeability; fibronectin; tumor necrosis factor-; proteinases
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INTRODUCTION |
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PULMONARY ENDOTHELIAL PROTEIN permeability is increased
during inflammatory-induced septic lung injury, an event associated with the release of cytokines such as tumor necrosis factor (TNF)- from activated monocytes and macrophages sequestered in the lung (19,
29, 30). This increase in lung vascular permeability contributes to the
etiology of pulmonary edema and acute respiratory distress syndrome
(ARDS) in septic surgical and trauma patients (1, 23).
Under both in vivo and in vitro conditions, TNF-
has been shown to
decrease the integrity of the endothelial barrier (5, 7, 9, 19, 21),
but the mechanism of this alteration in endothelial barrier
permeability is unclear.
Fibronectin (Fn) in the subendothelial matrix influences cell adhesion
to the substratum, a process that can affect endothelial cell shape and
cytoskeletal conformation (11, 25) and, therefore, the integrity of the
endothelial barrier. Exposure of calf pulmonary arterial endothelial
(CPAE) monolayers to TNF- causes both an increase in protein
permeability and a temporally associated reorganization and/or
disruption of the fibrillar organization pattern of Fn in the
extracellular matrix (ECM) (5, 37). Indeed, the insoluble pool of Fn
normally incorporated within the lung matrix is disturbed by many of
the same agents that can also increase lung vascular permeability in
vivo (6, 17, 20, 33), suggesting a potential functional relationship
between stability of the ECM and integrity of the lung vascular
barrier. The addition of purified soluble human plasma Fn (hFn) to the
medium of TNF-
-treated endothelial monolayers can prevent as well as
reverse the TNF-
-induced increase in protein permeability, and this
protective effect appears to require incorporation of the added soluble
Fn into the ECM (5, 35). Such findings suggest that the increase in
protein permeability caused by TNF-
may be due to disruption of the
Fn matrix, leading to altered endothelial adhesion and barrier
integrity.
Exposure of isolated endothelial cells to TNF- causes the release of
the serine proteinase urokinase-type plasminogen activator (uPA) as
well as of two matrix metalloproteinases, i.e., gelatinase B (MMP-9),
and stromelysin-1 (MMP-3) (8, 14, 16, 32). Accordingly, these
proteinases could potentially mediate the altered integrity of the ECM
and thus destabilize a confluent endothelial cell monolayer. However,
TNF-
exposure can also increase the release of proteinase inhibitors
such as plasminogen activator inhibitor-1 from endothelial cells (31).
From this perspective, the proteolytic activity of a proteinase
secreted by endothelial cells may, in theory, be regulated or
attenuated by endogenous inhibitors also secreted by the same cells.
Thus although the addition of TNF-
to the culture medium of CPAE
monolayers does cause the release of proteinases, their speculated role
in both the TNF-
-induced disruption of Fn in the ECM and the
increase in protein permeability has yet to be validated.
The present study was designed to determine whether the reorganization
and/or disruption of the Fn matrix as well as the temporally associated
increase in transendothelial protein permeability (5, 37, 38) observed
in CPAE monolayers after exposure to TNF- was dependent on
proteolysis of Fn in the subendothelial ECM. Our study, which is the
first to analyze the content, molecular weight, and deposition pattern
of Fn within the subendothelial matrix of endothelial monolayers after
TNF-
exposure, suggests that both the reorganization of Fn in the
ECM and the increase in monolayer protein permeability are not
dependent on the proteolytic degradation of matrix localized Fn. In
contrast, these TNF-
-induced changes in endothelial permeability and
ECM organization may be associated with a reduced colocalization of
5
1-Fn
integrins with fine Fn fibers within the ECM.
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METHODS |
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Endothelial Cell Monolayer
CPAE cells (American Type Culture Collection CCL-209, Rockville, MD) were grown in MEM (GIBCO BRL, Grand Island, NY) supplemented with 20% fetal bovine serum (FBS; Hyclone), penicillin (100 U/ml), and streptomycin (100 µg/ml). Experiments were performed between passages 17 and 23. When the CPAE monolayers were treated with TNF-TNF-, Leukocyte Elastase, and Proteinase Inhibitors
Antibodies
The following antibodies were purchased: rabbit anti-bovine Fn (bFn; Calbiochem, La Jolla, CA), FITC-conjugated goat F(ab')2 fragment to hFn (Cappel, Durham, NC), rabbit anti-bovine collagen IV (Biodesign International, Kennebunk, ME), and mouse monoclonal antibody to humanImmunofluorescence
CPAE cells were seeded (280,000 cells/well) on glass coverslips in 12-well tissue culture dishes and grown to confluence (3 days). The cell monolayers were then treated with medium alone or medium containing TNF-Immunofluorescence Protocols
Detection of bFn. After the TNF-Simultaneous detection of bFn and hFn.
After the TNF- treatment interval was over, the monolayers were
prepared for analysis by dual-label immunofluorescence. The primary
antibody used to detect bFn was a rabbit anti-bFn (1:100), and the
primary antibody used to detect hFn was a FITC-conjugated goat
F(ab')2 fragment to hFn
(1:50). The secondary antibody, RITC-conjugated goat anti-rabbit IgG
(1:50), was then added to visualize bFn. To ensure specificity of the
antibodies and avoid cross-reactivity, the antibodies to either bFn or
hFn were subjected to affinity chromatography against the opposite
species antigen with either hFn- or bFn-Sepharose columns,
respectively.
Simultaneous detection of hFn and bovine collagen
IV. Similar to the tracer experiments, we seeded CPAE
cells in the presence of medium containing FBS (deficient in bFn) plus
exogenous hFn. Soluble plasma hFn added to the culture medium before
confluence will incorporate into the ECM and colocalize with the
endogenous bFn. Moreover, after treatment with TNF-, both the
endogenous bFn and the added hFn will show similar reorganization
and/or disruption. The monolayers were treated with TNF-
for 18 h
and then prepared for dual-label immunofluorescence. The hFn was
detected with an FITC-conjugated goat anti-hFn (1:100), and collagen IV was detected with a rabbit anti-bovine collagen IV IgG and an RITC-conjugated goat anti-rabbit IgG (1:100). Such a dual-label approach to detect collagen IV and bFn in the matrix of the same monolayer avoided a potential antibody cross-reactivity with rabbit anti-bFn and rabbit anti-bovine collagen IV.
Simultaneous detection of hFn and
5
1-Fn
integrins.
An important assumption with regard to the endothelial monolayer on a
Fn-rich ECM is that cell-surface integrins, especially
5
1-Fn
integrins, located primarily on the basal surface of such adherent
cells, are bound or colocalized with Arg-Gly-Asp (RGD) sites in Fn
fibers in the matrix. Such colocalization should take place within
focal contact regions (3). Whether there would be reduced
colocalization after TNF-
exposure is not known. In the present
study, the cells were pulsed before confluence with hFn, the rapid
covalent incorporation of which into the ECM allowed for the
simultaneous staining of Fn within the matrix and cell-surface
5
1-integrins.
The monolayers were again treated with TNF-
for 18 h and then
prepared for dual-label immunofluorescence. On the same monolayers, hFn
was detected with an FITC-conjugated goat anti-hFn (1:100), and
cell-surface
5
1-integrins
were detected with both the mouse monoclonal antibody to human
5
1-integrin (1:500) and a secondary antibody that was an RITC-conjugated goat anti-mouse IgG (1:100).
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RESULTS |
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The data in Fig. 2, obtained with the use
of monoclonal antibodies to the
5
1-complex,
document the existence of
5
1-Fn integrins on the pulmonary endothelial cells in culture in focal adhesion structures (Fig. 2A) as
well as the presence of a fibrillar Fn subendothelial matrix (Fig.
2B). An 18-h exposure of CPAE
monolayers to TNF-
(200 U/ml) caused both a reorganization of the
fibrillar Fn matrix (Fig. 2E) and a
marked reduction in the immunofluorescent detection (red staining),
indicative of
5
1-integrins
within focal contact-like structures on many of the cells (Fig.
2D). Figure 2,
C and
F, are the computer-generated
overlapping images of both fields, confirming extensive colocalization
of the
5
1-integrins with the fine Fn fibers in the matrix (yellow staining) of normal monolayers (Fig. 2C) but reduced
association after TNF-
(Fig. 2F).
Such fluorescent images, which were observed repeatedly in separate
studies, are consistent with the possibility that TNF-
may disturb
the interaction of endothelial cell-surface
5
1-Fn integrins with Fn fibers in the matrix.
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To determine whether the rearrangement of Fn in the matrix of CPAE
monolayers was specific to Fn, we examined the organization of another
major matrix protein found in the ECM of pulmonary endothelial cell
matrices, i.e., collagen IV (Fig. 3).
Confluent endothelial monolayers were treated with medium alone or
medium containing TNF- (200 U/ml) for 18 h, after which time the
cell layers were fixed, permeabilized, and processed for dual-label immunofluorescence to examine both Fn and collagen IV in the ECM in the
same cells. As shown in Fig. 3, after treatment with TNF-
, there was
a dramatic reorganization and/or disruption of Fn in the matrix,
whereas collagen IV appeared to be organized in large bands both before
and after TNF-
treatment.
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To determine whether the reorganization of matrix Fn after TNF- was
due to proteolysis of the matrix-localized Fn, we used CPAE monolayers
with subendothelial matrices preloaded with small tracer doses
(1-2 µg) of soluble
125I-hFn (Fig.
4). To initially verify the model, we first
documented that the radiolabeled soluble hFn added to the medium as a
tracer would be incorporated into the ECM and that this newly
incorporated hFn would also be rearranged after TNF-
exposure
similar to endogenous bFn. Accordingly, dual-label immunofluorescence
was utilized to examine the organization of both endogenous bFn and
preloaded hFn after TNF-
exposure. As shown in Fig. 4, the soluble
hFn tracer added to the culture medium of the CPAE monolayer did
incorporate into the ECM (Fig. 4B)
where it colocalized in a fibrillar pattern with endogenous bFn (Fig.
4A). Moreover, as predicted, both
endogenous bFn (Fig. 4A) and the
newly added hFn already incorporated into the ECM (Fig.
4B) became rearranged in the
disrupted pattern after exposure to TNF-
(Fig. 4,
C and
D), validating that the hFn tracer behaved in a manner similar to endogenous bFn.
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It should be noted that in the above experiment (Fig. 4),
only tracer (1-2 µg) doses of
125I-hFn were used, not the
typical 300- to 600-µg hFn dose used to attenuate or normalize the
protein permeability after TNF- exposure (5, 37, 38). If the larger
treatment hFn dose is used, then the deposition pattern of the added
plasma Fn within the ECM appears, in part, dependent on when hFn is
added to the culture medium in relationship to the TNF-
challenge.
If soluble hFn is added at
18 h (with the TNF-
), then much of
the added hFn also is reorganized in a disrupted pattern by 12-18
h, like endogenous bFn (5), because it is rapidly incorporated into the
ECM, even within 2-3 h. Under these circumstances, it appears difficult to visualize by immunofluorescence the smaller portion of hFn
incorporated within the ECM in a fibrillar pattern because of the
intense background of a disrupted Fn matrix. However, we observed that
if hFn is added 12 h after the addition of TNF-
, when endogenous bFn
is already reorganized or disrupted, then 6 h later when the
permeability is lowered, the added hFn can be readily visualized (37)
in a fibrillar deposition in the ECM.
CPAE monolayers with a prelabeled Fn matrix were then exposed to
TNF- (200 U/ml) for 18 h, and both radiolabeled Fn released into the
medium and the presence of either intact or fragmented Fn in the medium
were analyzed with SDS-PAGE and autoradiography. The amount of
125I-Fn in the medium of both
control and TNF-
-treated monolayers was not significantly different
(Fig.
5A).
Also, analysis of medium samples diluted with 12% TCA to allow for
separation of 125I-bound Fn from
free 125I indicated similar
amounts of free 125I in the
control and TNF-
-treated monolayers (control, 12.0 ± 0.46%;
TNF-
, 8.6 ± 0.38%). In addition, SDS-PAGE and autoradiographic analysis (Fig. 5B) showed that Fn
present in the medium from both control and TNF-
-treated monolayers
was not different. Thus exposure of the monolayer to TNF-
did not
cause any significant release of Fn fragments into the culture medium.
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We next analyzed the relative amount of
125I-Fn associated with either the
cell layer (pool I) or
incorporated in the subendothelial matrix (pool
II) after TNF- exposure (18 h). To our surprise, after an 18-h exposure to TNF-
(200 U/ml), there was no significant decrease in either the amount of
125I-Fn bound to the cell layer
(pool I) or the
125I-Fn incorporated into the ECM
(pool II; Fig.
6A).
Also, SDS-PAGE and autoradiographic analysis showed that TNF-
exposure caused no Fn fragments either in the cell layer or within the
ECM (Fig. 6B). To verify that our
autoradiographic technique was sensitive enough to detect Fn fragments
if they existed, parallel experiments were done after leukocyte
elastase, a proteinase known to degrade Fn, was added to the
endothelial monolayers. Leukocyte elastase (2 × 10
4 U/ml) added for 18 h
caused both the release of Fn fragments and a loss of Fn from the cell
layer-matrix as expected (Curtis and Saba, unpublished
data). These findings indicated that TNF-
exposure
caused a reorganization of Fn in the matrix without an actual loss of
Fn from the matrix or any obvious proteolysis of Fn incorporated within
the ECM.
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SDS-PAGE and immunoblotting were also used to examine the endogenous
bFn after treatment with TNF-. Confluent monolayers were treated
with either medium alone (control) or medium containing 200 U/ml of
TNF-
for 18 h. Then, either the medium (Fig.
7A) or
the resulting cell layer and matrix (Fig.
7B) were analyzed by SDS-PAGE and
immunoblotting with an antibody specific to bFn. As predicted,
high-molecular-weight Fn multimers located in the stacker portion of
the gel, as previously shown (Fig. 6), could not be transferred and
therefore visualized during immunoblotting. TNF-
did not cause any
change in the amount of Fn detected within the medium or cell
layer-matrix (Fig. 7). Thus exposure of the CPAE monolayer to 200 U/ml
of TNF-
for 18 h does not appear to cause any major release of Fn
fragments from the matrix into the culture medium or significant
proteolysis of Fn already incorporated in the ECM.
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We then employed sensitive Fn-substrate zymography to determine whether
Fn-degrading proteinases were increased in both the medium and the
associated cell layer-ECM after TNF-. Confluent CPAE cells were
first treated with either medium alone (control) or medium containing
200 U/ml of TNF-
, and after 18 h, an equal volume of medium or cell
layer-matrix was analyzed by Fn-substrate zymography. As shown in Fig.
8A,
TNF-
treatment caused an increased expression in proteinase bands in
the medium, with two distinct molecular weights. These two proteinase
bands were also present in the cell layer-matrix, but their activity
was not changed with TNF-
(Fig.
8B). They were likely serine
proteinases because their degradative activity was completely inhibited
by aprotinin (4 µg/ml) but not by EDTA (20 mM), 1,10-phenanthroline
(0.1 mM), leupeptin (8 µg/ml), or pepstatin (0.4-1.2 µg/ml),
which have inhibitory activities toward metalloproteinases (EDTA and
1,10 phenanthroline), trypsin-like and cysteine proteinases
(leupeptin), and aspartic proteinases (pepstatin; Fig.
9).
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Because increased protease activity obviously existed, we then
determined whether the TNF--induced rearrangement of the Fn matrix
could be blocked by the serine proteinase inhibitor aprotinin. To our
surprise, the TNF-
-induced Fn matrix reorganization was identical
(Fig. 10) in the absence and presence of
aprotinin (4 µg/ml), further supporting the conclusion that the
reorganization of the Fn matrix was likely not dependent on Fn
proteolysis. Actually, a dose-response study was performed with
aprotinin at concentrations as high as 12 µg/ml, but, even at this
higher dose, aprotinin had no ability to block the TNF-
-induced
matrix reorganization (micrographs not shown). We also observed that
the TNF-
-induced increase in endothelial protein permeability as
measured by 125I-albumin clearance
was also not blocked by the serine proteinase inhibitor aprotinin (Fig.
11). Indeed, the typical 300-400%
increase in protein permeability in response to TNF-
was seen (Fig.
11) in both the absence and presence of aprotinin (4 µg/ml),
suggesting that the permeability increase was also being mediated by a
process independent of the proteolysis of Fn incorporated with
the ECM.
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DISCUSSION |
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The inflammatory cytokine TNF- has been shown to cause an increase
in the protein permeability of lung endothelial monolayers and a
dramatic disruption of the fibrillar Fn in the ECM (5, 37). Also, the
addition of soluble RGD peptides [but not Arg-Gly-Glu (RGE)
peptides] or antibodies to
5
1-integrins
to the culture medium of endothelial monolayers can increase their
protein permeability and rearrange the Fn in the ECM similar to the
pattern observed with TNF-
(5), suggesting that the TNF-
-induced
decrease in endothelial mono-layer integrity and rearrangement of the
Fn-rich matrix may be mediated by a change in the ability of the
cells to adhere to Fn in the matrix. The effect of these agents on the protein permeability of lung endothelial monolayers could be initiated by proteinase modification of the matrix Fn because TNF-
causes the
release of both uPA and the metalloproteinases gelatinase B and
stromelysin-1 from endothelial cells (8, 14, 16, 32).
RGD peptides themselves as well as
anti-5
1-integrins
have the ability to potentially stimulate protease secretion from the endothelial cells. Werb et al. (36) demonstrated that the adhesion of
fibroblasts to immobilized Fn peptides or to immobilized antibody against the
5
1-integrin
induced both collagenase and stromelysin gene expression, whereas
adhesion of fibroblasts to intact Fn did not induce this response. They
suggested that intact Fn may signal the cell differently than either an
antibody to the
5
1-receptor or RGD peptides. In the present study, we could not detect evidence of
Fn proteolysis after TNF-
treatment, suggesting that the
TNF-
-induced increase in endothelial protein permeability may be due
to a change in the ability of cell-surface integrins to bind to matrix
Fn. This alternate mechanism is supported by our observed change in the
surface distribution of the
5
1-integrin
after TNF-
exposure (Fig. 2), indicative of reduced interaction or
actual dissociation of many of the cell-surface
5
1-Fn
integrins with Fn fibers located in the subendothelial matrix.
In blood vessels, the ECM underlying the endothelium supports cell
attachment, spreading, migration, and proliferation. After injury, it
may be the composition and organization of the ECM that is directing
the responses of the endothelial cells (11). In response to binding an
immobilized ligand, such as Fn within the ECM, many integrins will
cluster and become immobilized within focal adhesion complexes due to
binding interactions with actin-associated cytoskeletal proteins (2).
The structural link among ECM, integrins, and the actin cytoskeleton
stabilizes cell adhesion and also provides the mechanical basis by
which integrins can contribute to endothelial shape and associated
endothelial barrier integrity. Changes in the ability of the cells to
bind to the ECM results in cytoskeletal reorganization and global
changes in cell shape (13, 25). Some insight into the mechanism by
which changes in adherence to the ECM promotes coordinated changes in
cell shape is apparent from the findings of Sims et al. (25). They
showed that changes in cell shape can be caused by cytoskeletal tension
generated by an actomyosin filament sliding mechanism, which is
physically resisted or opposed by the binding of cell-surface integrins
to immobilized adhesion sites within the ECM. These studies suggest that the ability of the integrins, such as
5
1-Fn
integrin, to bind to attachment sites with the ECM may be what controls
cell shape by physically resisting the resident or ongoing cytoskeletal tension.
Fn exists in two general forms: soluble Fn in the plasma or lymph fluid
and insoluble Fn (plasma or cellular) found in the ECM (22, 23). Plasma
Fn can incorporate into the tissue ECM and codistribute with locally
produced cellular Fn (22, 23). In the lung, Fn has been localized to
the subendothelial matrix, under epithelial cells, and in the
interstitial matrix (28). The release of Fn from the lung ECM has been
associated with lung vascular injury (6, 17, 20, 33). For example,
Peters et al. (17) documented the release of intact cellular Fn from the matrix of isolated perfused rabbit lungs after oxidant- or leukocyte-induced vascular injury. Release of intact extra domain (ED)-rich cellular Fn from the perfused rabbit lung after
oxidant-induced vascular injury is not prevented by prior inhibition of
protein synthesis in the lung (33). Moreover, postoperative
gram-negative bacterial infusion in sheep, which elicits an increase in
lung protein permeability, also causes the rapid release of
matrix-localized ED1-containing Fn
from the lung interstitial ECM into the postnodal lymph (20). In
essentially all of these in vivo experiments, very few Fn fragments
could be detected. This suggests that the loss of lung barrier
integrity in vivo coupled with the large water flux across the
endothelial barrier may have facilitated the washout of intact Fn from
the disrupted interstitial matrix by a process unrelated to Fn
proteolysis, or, because soluble Fn can be rapidly assembled
and/or incorporated into the ECM (5, 12, 18), perhaps there is
a rapid matrix Fn disassembly process activated by TNF-.
The exposure of lung endothelial monolayers to TNF- for 18 h did not
change the content or molecular weight of the Fn recovered from the
conditioned medium or the cell layer-ECM. It should be noted that
Stolpen et al. (27), using human umbilical vein endothelial cells,
showed a loss of Fn from the matrix by immunofluorescent techniques,
but this loss of stainable Fn was not apparent until 72-96 h after
TNF-
treatment, a time course much greater than that used in the
present study. Partridge and colleagues (15, 16) used SDS-PAGE analysis
(under reducing conditions) and reported loss of a 220-kDa band (Fn
monomer) after treatment of microvessel endothelial cells with TNF-
,
but loss of the Fn monomer did not correspond with any increase in Fn
fragments. Additionally, because the high-molecular-weight multimers of
matrix Fn, which are known to be retarded within the stacking portion
of the gel, were not shown, the loss of Fn monomers after TNF-
may
simply be due to an increase in high-molecular-weight Fn multimers
resistant to reducing agents.
Niedbala and Picarella (14) showed that TNF- treatment of human
umbilical vein endothelial cells (low passage) caused proteolysis of a
[3H]glucosamine-labeled
ECM as measured by the release of radioactivity into the medium over
time. This TNF-
-mediated ECM degradation was plasminogen dependent
and could be inhibited by an anticatalytic uPA monoclonal antibody.
Plasmin can degrade multiple ECM components, so the relationship of
this finding to any potential proteolysis of matrix Fn can only be
speculated. The present study, which is the first extensive analysis of
the content, molecular weight, and organization of Fn in the
subendothelial matrix after treatment with the cytokine TNF-
,
suggests that TNF-
alters the organization of Fn in the matrix and
increases the protein permeability of the endothelial cell monolayers
independent of Fn proteolysis.
Proteinase activity against Fn was readily detected in both the medium
and cell layer-matrix, and such proteolytic activity was increased in
the medium after TNF- treatment. The proteinase activity detected in
the Fn zymogram was likely plasmin because the bands detected had a
molecular mass of ~90 kDa, and their Fn-degrading
activity was inhibited by aprotinin. However, because of the numerous
inhibitors in the serum, it is unlikely that any additional plasmin
released into the culture medium would have had any significant
degradative activity against the Fn found in the subendothelial matrix.
It is possible that the aprotinin we added did not have access to
cell-associated plasmin, but we added a large concentration of
aprotinin over a long time period. Because aprotinin is 6.5 kDa, we did
not have reason to believe that it would be limited in gaining access
to the basolateral cell surface even in control conditions. In the
TNF-
treatment conditions, the cells lose their cell-cell contacts
so aprotinin would have full access. Using the same confluent
endothelial monolayers, we have already shown that the addition of
antibodies to
5
1-integrins can increase endothelial protein permeability in 18 h. The fact that
the antibody to
5
1-integrin,
which is very large compared with aprotinin, has access to the
basolateral surface is proof to us that aprotinin (6.5 kDa) can get to
these sites, especially after TNF-
. We also measured plasmin
activity after either TNF-
alone or TNF-
coupled with aprotinin.
We observed that aprotinin decreased the plasmin activity exhibited
after TNF-
(Curtis and Saba, unpublished data),
consistent with previous findings (26) that aprotinin inhibits both
fluid-phase and cell-surface plasmin activity. However, aprotinin was
unable to block the TNF-
effect on the monolayer. Collectively, such
data suggest that an aprotinin-sensitive proteinase did not cause the
TNF-
-induced Fn matrix disruption or the increase in protein
permeability that we observed.
Disruption of the subendothelial Fn matrix after TNF- exposure as
reflected by the condensation of the Fn matrix into thicker bands or
aggregates of fibers was repeatedly observed. TNF-
treatment of
endothelial cells did not appear to cause any major reorganization of
collagen IV, suggesting that Fn matrix reorganization and/or disruption
was specific. Although this response appears to be independent of Fn
proteolysis, it could potentially be explained by an altered mechanical
balance between the ECM and the cytoskeleton in accordance with the
concept of Wang and colleagues (34, 35).
Although disruption of the Fn matrix after TNF- exposure appears not
to be due to a proteolytic attack of Fn already incorporated within the
ECM, the functional relationship of the matrix Fn disruption to the
observed increase in transendothelial protein permeability remains to
be clarified. Both RGD peptides and antibodies to
5
1-integrin can each increase the protein permeability of CPAE monolayers as well
as cause a disruption of the Fn matrix identical to those seen after
TNF-
exposure (5, 37, 38). Moreover, the soluble plasma Fn added to
the culture medium of lung endothelial monolayers can incorporate into
the ECM and attenuate the increase in protein permeability observed
with all three agents. This suggests that the changes caused by RGD
peptides,
5
1-integrin
antibodies, or TNF-
may be mediated by a similar mechanism. Because
the ECM can indirectly modulate endothelial barrier function by
influencing endothelial cell shape, cell spreading, and cell-matrix
adhesion due, in part, to the binding of integrins to matrix molecules, it would appear that reduced interaction of
5
1-Fn
integrins with Fn fibers in the matrix may be the basis for the
response we observed. The previous findings of Curtis et al.
(5) of enhanced cell spreading and a decrease in
endothelial protein permeability after matrix incorporation of added
soluble Fn coupled with the present data indicating that proteolysis is
not mediating the response support this conclusion.
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ACKNOWLEDGEMENTS |
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The secretarial assistance of Debbie Moran is much appreciated.
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FOOTNOTES |
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This study was supported primarily by National Institute of General Medical Sciences Grant GM-21447 and in part by National Cancer Institute Grant CA-69612.
T. M. Curtis was a predoctoral trainee in the Department of Physiology and Cell Biology (Albany Medical College, Albany, NY), was supported by National Heart, Lung, and Blood Institute Training Grant T32-HL-07194, and is currently a postdoctoral fellow at the University of Virginia (Charlottesville). R. F. Rotundo was a postdoctoral fellow in the Department of Physiology and Cell Biology during these studies, was supported by National Heart, Lung and Blood Institute Training Grant T32-HL-07529, and is currently a Research Associate at Albany Medical College.
Address for reprint requests: T. M. Saba, Dept. of Physiology and Cell Biology (A-134), Albany Medical College, 47 New Scotland Ave., Albany, NY 12208.
Received 21 November 1997; accepted in final form 7 April 1998.
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