Molecular responses of rat tracheal epithelial cells to
transmembrane pressure
Barbara
Ressler1,
Richard T.
Lee2,
Scott H.
Randell3,
Jeffrey M.
Drazen4, and
Roger D.
Kamm1
1 Department of Mechanical Engineering,
Massachusetts Institute of Technology, Cambridge 02139;
3 Division of Pulmonary and Critical Care
Medicine and 4 Cardiovascular Division,
Brigham and Women's Hospital, Boston, Massachusetts 02115; and
2 Cystic Fibrosis/ Pulmonary Research and
Treatment Center, University of North Carolina School of Medicine,
Chapel Hill, North Carolina 27599
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ABSTRACT |
Smooth muscle constriction in asthma
causes the airway to buckle into a rosette pattern, folding the
epithelium into deep crevasses. The epithelial cells in these folds are
pushed up against each other and thereby experience compressive
stresses. To study the epithelial cell response to compressive stress,
we subjected primary cultures of rat tracheal epithelial cells to
constant elevated pressures on their apical surface (i.e., a
transmembrane pressure) and examined changes in the expression of genes
that are important for extracellular matrix production and maintenance of smooth muscle activation. Northern blot analysis of RNA extracted from cells subjected to transmembrane pressure showed induction of
early growth response-1 (Egr-1), endothelin-1, and transforming growth
factor-
1 in a pressure-dependent and time-dependent manner. Increases in Egr-1 protein were detected by immunohistochemistry. Our
results demonstrate that airway epithelial cells respond rapidly to
compressive stresses. Potential transduction mechanisms of transmembrane pressure were also investigated.
asthma; gene expression; mechanical stress; early growth
response-1; endothelin-1; transforming growth factor-
1
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INTRODUCTION |
IT IS NOW WELL ESTABLISHED that mechanical forces play
a critical role in tissue remodeling and cellular homeostasis. Vascular endothelial cells (32), cardiac myocytes (37), osteoblasts (41), and
glomerular mesangial cells (53) all function in mechanically dynamic
environments; perturbations in mechanical stress are known to have
physiological and pathological implications. Cells in the lung also
experience variable mechanical stress and respond through numerous lung
signal transduction pathways. For example, stretching alveolar
epithelial cells (50) and mechanically perturbing tracheal epithelial
cells (10) result in Ca2+ mobilization. Cyclic stretch of
airway epithelium inhibits prostanoid synthesis (39) and wound repair
(38) in vitro. Cyclic stretch and fluid shear stress on pleural
mesothelial cells has been shown to affect growth factor release (45).
Another important perturbation occurs in the airway when smooth muscle
constricts and the airway wall consequently buckles into a rosette
pattern, causing the epithelium to fold into deep crevasses (51).
Epithelial cells along the folds of these rosettes are pushed up
against each other (Fig. 1) and thus are
subjected to a compressive stress (i.e., pressure or a stress
perpendicular to their surface). Pressures exerted on the epithelium in
an unconstricted airway during normal breathing are seldom > 1 cmH2O (98 Pa) above atmospheric pressure (46) and are much
lower than the compressive stresses exerted by constricting smooth
muscle (>30 cmH2O; 15). We hypothesized that the
compressive stress on airway epithelial cells during
bronchoconstriction could stimulate the cells to modify their
environment in response to this stress much as hemodynamic stresses
induce biological responses in endothelial cells (32).

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Fig. 1.
Schematic of a highly constricted airway. Smooth muscle constriction
causes the airway to buckle into folds. Epithelium within the folds of
the lamina propria is subjected to a normal stress (i.e., a stress
perpendicular to its surface).
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To test this hypothesis, we subjected primary cultures of rat tracheal
epithelial cells to a physiological range of static pressures and
measured changes in gene expression. We examined genes that are known
to be transduced by mechanical forces in vascular endothelial cells and
that also play a role in tissue fibrosis. Early growth response-1
(Egr-1), an immediate-early gene, activates transcription of
platelet-derived growth factor (PDGF; 25), which stimulates fibroblast
proliferation and collagen production (31). Endothelin-1 (ET-1) is a
potent airway contractile agonist and also promotes collagen synthesis
by fibroblasts (4). Transforming growth factor-
1 (TGF-
1) is an
important regulator of lung fibrosis (6) and has been shown to
stimulate expression of fibronectin and collagen genes in fibroblasts
(31). In this study, we demonstrated magnitude- and time-dependent
differential expression of these genes in response to static
transmembrane pressure. Our results suggest that the response of airway
epithelial cells to compressive stresses in constricted airways may
lead to airway fibrosis independent of inflammation in asthma.
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METHODS |
Cell culture.
DMEM-Ham's F-12 medium (DMEM-F-12) and penicillin-streptomycin were
obtained from GIBCO BRL (Life Technologies, Grand Island, NY).
Transferrin and epidermal growth factor (EGF) were from Collaborative Research (Bedford, MA). Bovine pituitaries were purchased from Pel
Freeze (Rogers, AR). All other culture medium additives were obtained
from Sigma (St. Louis, MO). Cells were grown on Transwell-COL tissue
culture inserts (0.4-µm pore size, 24-mm diameter; Corning Costar,
Cambridge, MA). The base of the culture insert consisted of a porous
membrane made of polytetrafluoroethylene coated with type I and III collagen.
Rat tracheal epithelial (RTE) cells were cultured following the
procedure of Kaartinen et al. (22) with slight modification. Cells were
grown in complete medium (CM), which consisted of DMEM-F-12 supplemented with penicillin-streptomycin (100 U/ml and 100 µg/ml, respectively), gentamicin (50 µg/ml), HEPES (30 mM), insulin (10 µg/ml), hydrocortisone (0.1 µg/ml), cholera toxin (0.1 µg/ml), transferrin (5 µg/ml), phosphoethanolamine (10 µM), ethanolamine (80 µM), EGF (25 ng/ml), retinoic acid (5 × 10
8 M), BSA (0.5 mg/ml), and bovine pituitary
extract (1%).
Male Fischer 344 rats, 300-500 g body wt, were killed by
CO2 asphyxiation. Tracheae were sterilely excised, filled
with a 1% protease solution (Sigma) in DMEM-F-12, and
kept overnight at 4°C. Cells were flushed out of the tracheae with
DMEM-F-12 containing 5% heat-inactivated FBS and collected by
centrifugation (500 g, 10 min, 4°C). Cells were resuspended
in a high-protein medium (CM with 3 mg/ml BSA), counted, and seeded on
the top surface of the culture inserts at a density of ~5 × 104 cells/cm2. The lower chamber of the culture
wells contained high-protein medium supplemented with 10% FBS.
Cultures were incubated in a humid environment at 35°C with 3%
CO2 in air.
After 24 h, the medium was changed to CM without serum. The medium was
changed every other day until day 5, after which it was changed
daily. The culture conditions promoted growth of epithelial cells with
a cobblestone morphology; there was no apparent growth of cells with a
fibroblast morphology. After the cells were confluent (day 7),
the medium on the apical surface was removed, and CM was added to the
bottom compartment only to produce an air-liquid interface at the cell
surface. Another 7 days elapsed during which the cells differentiated
into a pseudostratified, mucus-secreting culture with cells of columnar
and basal morphology. Experiments and interventions were conducted on
day 14.
Experimental procedure.
Approximately 12 h before the start of the experiment, the medium was
changed to minimal medium (DMEM-F-12 plus antibiotics only), and the
culture wells were prepared for pressure application in the incubator
(Fig. 2). Stoppers fitted with connectors
were tightly placed in the culture wells. Control culture wells were handled in the same manner and inserted in the apparatus but were not
pressurized. To apply pressure, a large tank filled with incubator air
was pressurized to the desired level with a sphygmomanometer bulb. At
the start of the experiment, the valve from the tank to the cells was
opened, and pressure over the cells (P1) was raised. This
apparatus applied pressure only to the apical surface of the cells; the
basal surface of the culture membrane and the culture medium were
exposed to atmospheric pressure (P2) only. Thus the cells
were subjected to a transmembrane pressure (P1 > P2).

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Fig. 2.
Schematic of a culture well subjected to transmembrane pressure. Air in
the sealed chamber above the cells was pressurized (P1) to
apply a normal stress to the surface of cells and basal surface of
membrane, and culture medium remained exposed to atmospheric pressure
(P2). Thus cells experience a transmembrane pressure
(P1 > P2).
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The pressures (P1) we selected to study were 2, 5, 10, and
20 cmH2O, which were in the range of pressures known to be
generated by airway smooth muscle (15). Samples were collected after 0, 1, and 6 h. Cells harvested at the zero time point were not inserted in
the apparatus and were minimally handled to examine the effects of
manipulation on cell stimulation. At the appropriate time, cells were
lysed for RNA extraction and medium samples were collected. Cell
viability was evaluated by the amount of lactate dehydrogenase (LDH;
Sigma) in the medium. Experimental LDH levels were normalized to LDH
levels of completely lysed cells.
To explore the effects of elevated hydrostatic pressure on the cells
(i.e., P1 = P2, with both greater than
atmospheric pressure), the culture plates were placed in a Plexiglas
box that was sealed and then pressurized. Cells in the box experienced
elevated pressure on both the apical and culture membrane surfaces.
There was no strain of the culture membrane, and the pressures were
equal on all surfaces of the culture; thus the cells in this experiment were subjected to hydrostatic pressure only.
To determine the amount of membrane strain that occurred as a result of
transmembrane pressure, we obtained a digital image (Pulnix, Sunnyvale,
CA) of the pressurized culture well and measured the maximum deflection
at the membrane center (wmax) with image-analysis software
(NIH Image version 1.60, Bethesda, MD). Average membrane strain
(
) was calculated according to the
analysis of Williams et al. (49):
=
(wmax/a)2, where
a is the culture membrane radius (a = 12 mm
for our Transwells).
To explore the effects of transmembrane pressure without the associated
strain of the culture membrane, we placed a stiff plastic screen in the
bottom of the culture dish and held the culture membrane flush against
the screen during apical pressure application. Cells were thereby
exposed to the medium at atmospheric pressure while strain of the
culture membrane was minimized.
RNA isolation and hybridization.
Total cellular RNA was isolated with commercially available kits
(RNeasy, QIAGEN, Valencia, CA). After the RNA was concentrated and its
purity and concentration were determined, 15 µg of total RNA were
fractionated on 1.2% agarose-6% formaldehyde gels. RNA was
transferred overnight by capillary action onto nylon membranes (Stratagene, La Jolla, CA) and immobilized with ultraviolet radiation. Northern blot hybridization was performed with a random-primer 32P-labeled 3-kb EcoR I cDNA fragment of mouse
Egr-1 (a kind gift from Dr. V. Sutakme, Beth Israel/Deaconess Medical
Center, Boston, MA), a 600-bp EcoR I cDNA fragment of rat ET-1,
a 1-kb Xba I-Hind III cDNA fragment of rat TGF-
1
(ET-1 and TGF-
1 were kind gifts from Dr. M.-A. Lee, Brigham and
Women's Hospital, Boston, MA), and a 1.1-kb cDNA fragment of human
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) from Clontech (Palo
Alto, CA). Membranes were hybridized with ExpressHyb solution
(Clontech) for 1 h at 68°C and then probed simultaneously for
either Egr-1 and GAPDH or ET-1 and GAPDH, washed, and exposed to X-ray
film for 1-3 days at
80°C with intensifying screens.
Membranes probed with ET-1 and GAPDH were boiled in 0.5% SDS for 10 min and then reprobed for TGF-
1 following the same procedure.
Autoradiograms were quantitatively analyzed by scanning densitometry
and image analysis (Optimas, version 5.2, Bothell, WA). Egr-1, ET-1,
and TGF-
1 signals were normalized to the corresponding GAPDH signal
to correct for variations in loading and transfer. GAPDH was chosen
because it has been shown not to be strongly regulated by other
mechanical stimuli such as shear stress (27) or stretch (3).
Immunohistochemistry.
Culture wells subjected to 10 or 20 cmH2O pressure
for 1 or 6 h were fixed in 4% paraformaldehyde in PBS for 10 min,
permeabilized with 0.3% Triton (Sigma) for 5 min, and blocked with
10% goat serum. Cells were then incubated with rabbit anti-human EGR-1 antibody (1:100 dilution; Santa Cruz Biotechnologies, Santa Cruz, CA)
overnight at 4°C. After being washed, cells were incubated with
biotinylated goat anti-rabbit antibody (1:200 dilution; DAKO, Carpenteria, CA) for 1 h at room temperature. Then the cells were rinsed, incubated with streptavidin fluorescein (1:750 dilution; NEN,
Boston, MA), washed, mounted, and examined by fluorescence microscopy.
Staining specificity was determined by incubating the cells with
nonspecific mouse IgG1 (1:100 dilution; Sigma) or with the biotinylated
secondary antibody only.
Statistics.
Data are expressed as means ± SE unless indicated otherwise.
Data were compared with an unpaired Student's t-test or a
one-way ANOVA combined with Scheffé's test. P values < 0.05 were considered significant.
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RESULTS |
Northern analysis: transmembrane pressure experiments.
Figure 3 shows representative Northern
blots of RNA collected from RTE cells subjected to 20 cmH2O
pressure for 0, 1, and 6 h. Egr-1 hybridization signals were easily
apparent after 1 h of stimulation and returned to baseline control
levels after 6 h of steady pressure stimulation. ET-1 hybridization
signals were elevated after both 1 and 6 h of stimulation. TGF-
1
hybridization signals were elevated after only 6 h of stimulation.

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Fig. 3.
Expression of early growth response-1 (Egr-1), endothelin-1 (ET-1),
transforming growth factor- 1 (TGF- 1), and
glyceraldehyde-3-phosphate dehydrogenase (GAPDH) after 1-6 h of 20 cmH2O pressure stimulation. Egr-1 mRNA was upregulated
after 1 h of pressure stimulation. ET-1 mRNA was elevated with 1 and 6 h of pressure. TGF- 1 mRNA was clearly present after 6 h of pressure
application. +, With; , without.
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A summary of densitometric data from the Northern analysis is given in
Fig. 4. After 1 h of steady pressure
stimulation, Egr-1 mRNA levels were elevated above control levels: 18.5 ± 4.7-fold (n = 9 experiments; P < 0.0001) with 20 cmH2O pressure, 6.8 ± 1.2-fold (n = 8 experiments; P < 0.0001) with 10 cmH2O pressure, and 3.4 ± 0.6-fold (n = 6 experiments; P < 0.001)
with 5 cmH2O pressure. In all experiments, Egr-1 expression
returned to baseline levels after 6 h of stimulation. Pressures of 2 cmH2O did not significantly stimulate Egr-1 expression.
ET-1 mRNA levels were elevated above control only in the 20 cmH2O experiments in which ET-1 levels were 4.1 ± 0.7-fold (n = 5 experiments; P < 0.01) over control
level after 1 h and 4.7 ± 1.0-fold (n = 5 experiments; P < 0.01) over control level after 6 h. TGF-
1 mRNA
expression was increased over control level by a factor of 4.6 ± 0.7 (n = 7 experiments; P < 0.0001) and 2.2 ± 0.2 (n = 5 experiments; P < 0.0001) after 6 h of 20 and
10 cmH2O pressure stimulation, respectively; 5 and 2 cmH2O pressure failed to significantly induce TGF-
1
expression. Expression of GADPH mRNA was highly consistent with
ethidium bromide staining of the 18S bands and did not vary with the
magnitude of pressure applied (data not shown).

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Fig. 4.
Densitometric analysis of Egr-1, ET-1, and TGF- 1 at 20 (A),
10 (B), 5 (C), and 2 (D) cmH2O.
Values were normalized to control levels. Note different scales of
y-axes. Significantly greater than control level: **P < 0.0001; P < 0.001; *P < 0.01.
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Northern analysis: hydrostatic pressure and no-strain experiments.
To study the effects of hydrostatic pressure on gene expression, we
subjected RTE cells to 10 cmH2O transmembrane pressure or
hydrostatic pressure for 0, 1, or 6 h. Hydrostatic pressure did not
induce Egr-1 mRNA expression but transmembrane pressure did at the 1-h
time point (Fig. 5A). Transmembrane
pressure, but not hydrostatic pressure, also increased TGF-
1 mRNA
expression approximately twofold after 6 h (Fig. 5B).

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Fig. 5.
A: Egr-1 mRNA expression after 1 h of 10 cmH2O
transmembrane pressure or hydrostatic pressure application. B:
TGF- 1 mRNA expression after 6 h of 10 cmH2O
transmembrane pressure or hydrostatic pressure application.
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The average strain of the culture membrane
(
) was calculated to be 2.3% at 20 cmH2O transmembrane pressure, 1.5% at 10 cmH2O, 1.0% at 5 cmH2O, and 0.5% at 2 cmH2O. Because the membrane was fixed at the edges, the
strain pattern was anisotropic and nonuniform (49). To reduce membrane
strain, a rigid mesh support with 800-µm-diameter pores was inserted;
membrane strain with the support in place was calculated to be less
than one-tenth the average strain of the unsupported membrane (8).
Cells were subjected to 10 cmH2O transmembrane pressure
with and without the mesh supports. Thus the cells were subjected to
1.5% average membrane strain without support or <0.15% strain in
the mesh pores with support. Representative Northern blots for Egr-1
mRNA in cells exposed to 10 cmH2O transmembrane pressure for 1 h with and without membrane strain are shown in Fig.
6. Cells subjected to 10 cmH2O
transmembrane pressure without membrane strain for 1 h exhibited a 4.8 ± 0.5-fold (n = 5 experiments) increase in Egr-1 mRNA. Cells
under the same pressure but with membrane strain had a 7.0 ± 1.1-fold
(n = 9 experiments) increase in Egr-1 mRNA after 1 h. The
difference in levels of Egr-1 mRNA between the strained and supported
samples was not significant (P = 0.18). TGF-
1 mRNA was also
elevated in both strained (1.7-fold) and unstrained (2.1-fold) cells
after 6 h of pressure stimulation, but too few experiments were
performed to determine significance (data not shown).

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Fig. 6.
Effect of membrane strain on Egr-1 mRNA expression. Experiment was 10 cmH2O pressure applied for 1 h with and without a mesh
support to remove membrane strain.
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Immunohistochemistry.
Increased levels of Egr-1 protein were detected in cells subjected to 6 h of 20 (Fig. 7A) and 10 cmH2O (data not shown) pressure stimulation. There was no
detectable immunofluorescence above background in cells subjected to 1 h of pressure stimulation (data not shown) or in control cells (Fig.
7B). Specificity of the staining was confirmed by the absence
of staining with exposure to nonspecific mouse IgG1 (Fig. 7C)
and to the biotinylated secondary antibody (data not shown).

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Fig. 7.
Immunofluorescence of Egr-1. Cells exposed to 20 cmH2O for
6 h (A) showed increased Egr-1 protein immunostaining.
B: control cells with no pressure. C: cells stained
with irrelevant mouse IgG antibody.
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Cell death.
At 6 h and 20 cmH2O pressure, LDH levels indicated 16 ± 5% (SD) cell death (n = 47 experiments). At all other time
points and pressures, <2% of the cells died.
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DISCUSSION |
Implications for airway remodeling and asthma.
Cells in mechanically dynamic environments such as endothelial cells
(32), osteoblasts (41), and fibroblasts (14) often cause stress
adaptation in the surrounding tissue if the mechanical environment
deviates from normal. We present evidence that mechanical forces
equivalent to those resulting from bronchoconstriction can transduce
signals in airway epithelial cells. In response to compressive
stresses, which we believe mimic those exerted on epithelial cells in
the folds of a highly constricted airway, rat tracheal epithelial cells
differentially upregulated mRNA encoding expression of Egr-1, ET-1, and
TGF-
1 in a magnitude- and time-dependent manner. In the case of
Egr-1, we also demonstrated by immunohistochemistry that elevated mRNA
levels are accompanied by elevated Egr-1 protein levels.
The compressive stresses applied were estimated to be in the
physiological range of those generated by smooth muscle. For example,
canine airway smooth muscle has been shown to generate intraluminal
pressures > 30 cmH2O in small bronchi with ACh
stimulation (15). The magnitude of compressive stress acting on the
epithelium of a constricted airway (such as that depicted in Ref. 51)
can be estimated by modeling the airway as a thick-walled cylinder under external pressure. A simple force balance shows that the mean
circumferential wall stress (
) is related
to the effective external pressure due to smooth muscle constriction
(Psm) through the following expression:
(R/t + 1)Psm, where R is the inside radius and
t is the wall thickness, both of the constricted airway (43).
Using the estimated value of R/t
0.5 (51) and substituting Psm = 30 cmH2O gives
45 cmH2O. A thick-walled cylinder is a crude model of a buckled airway and does not provide any
details of the stress pattern on the epithelium. Extension of previous
finite-element models of the airway (48) leads to a more accurate
estimation of the type and magnitude of stresses experienced by the
epithelium during smooth muscle constriction that are consistent with
this rough estimate (33).
After rapid smooth muscle constriction, the airway tissue may initially
be regarded as incompressible and the epithelial cells within the folds
would experience no transcellular pressure difference. However, if
constriction continues, fluid within the high-pressure regions of the
airway wall may shift to regions of lower fluid pressure within the
wall or in the luminal space. As this occurs, stresses within the
constricted airway would be increasingly borne by the matrix, and the
epithelial cells within the folds would eventually experience a
transcellular fluid pressure gradient such as that imposed in our in
vitro system. Employing poroelastic computational models of the airway
wall would be helpful in determining the time-varying fluid stresses
within a constricted airway.
In the absence of smooth muscle constriction, the airway is unbuckled
and the epithelium only experiences those stresses associated with
airway wall strain and variations in air pressure. Wall strain is
difficult to estimate, but it has been reported that the circumference of the supporting basement membrane changes little during changes in
airway caliber (19). Variations in air pressure in the airways amount
to only ~1 cmH2O with quiet breathing (47). Thus
compressive stresses acting on the epithelium in an unconstricted
airway during normal breathing are likely to be well below those in a
constricted airway and also below those that resulted in signal
transduction in our experiments. During positive-pressure ventilation,
however, airway pressures reach levels in excess of 35 cmH2O. This has led others to suggest that mechanical
ventilation may lead to trauma and remodeling of the lung parenchyma
(5, 13), although the cause may be high alveolar distension (13), high
pulmonary capillary pressures (5), or high oscillatory airway pressures.
We chose not to study pressures higher than 20 cmH2O to
avoid rupturing the culture membrane and because of the decrease in cell viability observed at 20 cmH2O. The cause of this cell
death is unknown. Vital staining of the culture wells after 6 h of 20 cmH2O transmembrane pressure showed predominantly live
cells, with small clusters of dead cells scattered throughout the
culture well (data not shown). The late onset of cell death in the
high-pressure experiments suggests an apoptotic rather than a necrotic
pathway, but this was not determined in these experiments. There is
intriguing evidence of a link between cell viability and cytoskeletal
integrity. Disruption of cytoskeletal components has been shown to lead
to apoptosis either by liberation of enzymes or by loss of mechanical tension at points of cell contact (see Ref. 21 for review). If the
stress of transmembrane pressure application disrupted the cytoskeleton
of the RTE cells, then the resulting cell death may have been a natural
consequence of this disruption. Because mRNA upregulation was observed
at pressures where cell death was undetectable and at the early time
point in the high-pressure experiments, gene transcription is not
believed to depend on cell death.
Bronchoconstriction in asthma may occur rapidly after exposure to
antigen or may occur several hours after antigen exposure and last
several hours to several days (46). Thus epithelial cells in a
constricted airway may be exposed to mechanical stresses for periods
much longer than 6 h, the maximum exposure time in our experiments. We
limited our study to a time course consistent with the duration of
smooth muscle constriction during an acute asthma attack. Our results
show that epithelial cells respond rapidly to mechanical stresses and
suggest that repeated bouts of bronchoconstriction, even for relatively
short times, may indeed result in accumulation of extracellular matrix
in the airway wall.
RTE cells were used as a model system to study the cellular response to
forces in constricting bronchi. The epithelial cell phenotypes vary
along the length of the bronchial tree, so it is not known whether rat
bronchial epithelial cells would respond to compressive stresses in the
same manner as tracheal epithelial cells. Further experiments would be
required to determine regional differences in the response of
epithelial cells to mechanical stress.
This study focused on genes thought to influence the asthmatic airway.
Egr-1 is an immediate-early gene and zinc finger transcription factor.
It has been shown to play a role in cell proliferation (17), has been
detected at sites of vascular injury (24), and is upregulated by
increased cardiac ventricular loads (pressures) in cat heart muscle
(36). Egr-1 also activates transcription of PDGF (24, 25), which
stimulates fibroblast proliferation and collagen production (31). Other
genes such as TGF-
1 and Egr-1 itself also have nucleotide
recognition sites for Egr-1. Egr-1 is known to be sensitive to
mechanical stimulation (29); our experiments showed that Egr-1 was
particularly sensitive to the magnitude of normal stress and was
upregulated with as little as 5 cmH2O of pressure. The lack
of upregulation at 2 cmH2O indicates that the manipulations
required to apply the steady pressure did not initiate
mechanotransduction. The time course for transcription was similar to
that in endothelial cells subjected to shear stress, with early (30 min
to 1 h) enhanced transcription and a decrease to baseline by 6 h (40).
ET-1 is a potent airway contractile agonist that also promotes collagen
synthesis by fibroblasts (4). Its transcription appears to be highly
dependent on the type and magnitude of stress. Physiological levels of
fluid shear stress on vascular endothelial cells cause rapid
downregulation of ET-1 mRNA in a dose-dependent manner (27). Stretch,
on the other hand, has been shown to either upregulate ET-1 mRNA
expression (44) or have no effect (27) in endothelial cell cultures.
Our experiments showed a different pattern of ET-1 mRNA expression,
with rapid (1 h) and sustained (6 h) elevation when 20 cmH2O was used as the stimulus. Interpretation of this
finding is complicated, however, by the cell death observed in
experiments conducted at 20 cmH2O.
Stretching mesangial cell cultures caused induction of TGF-
1 mRNA,
but long periods of stretch (36-48 h) were required (34, 53). In
the present experiments, TGF-
1 mRNA levels were clearly increased
after only 6 h of pressure stimulation. TGF-
1 mRNA was elevated more
than four times the control level with high pressure (20 cmH2O) and was doubled with 10 cmH2O pressure.
Thus our results are consistent with those of Berg et al. (5), who reported that TGF-
1 mRNA expression levels in rabbit lung parenchyma were increased by a factor of four after only 4 h of high airway pressure ventilation. Our results also raise the possibility that airway epithelial cells were the source of the increased TGF-
1 mRNA
levels noted by Berg et al. Because of the relatively short duration of
our experiments, we did not investigate whether transmembrane pressure
increased production of ET-1 and TGF-
1 proteins because there would
likely be a delay of several hours between expression of message and
production of protein. Further experiments would be necessary to
determine whether transmembrane pressure also leads to production,
secretion, and activation of these proteins.
Mechanisms of gene induction.
Although it is unclear exactly how transmembrane pressure stimulates
gene expression in epithelial cell cultures, we are able to eliminate
certain potential causes. In our experiments, we observed average
culture membrane strains as high as 2.3%. Although cyclic strains as
low as 1% are adequate to produce changes in gene expression (52), no
studies have shown any effect of such small static strains. Our own
experiments demonstrated elevated levels of Egr-1 mRNA even when
membrane strain was reduced 10-fold. Therefore, membrane strain is not
responsible for our findings.
Experiments were also performed to eliminate the effects of hydrostatic
pressure as a mechanism for gene induction in our cultures. Although
hydrostatic pressures in the range of 10-170 cmH2O
have been shown to enhance cell proliferation (23), DNA synthesis (16),
or release of cellular protein (1), there are no examples of
hydrostatic pressure affecting gene expression levels. Our experiments
did not show any changes in mRNA expression with 10 cmH2O
hydrostatic pressure. It is unclear how hydrostatic pressure would
stimulate the cells because this would not lead to cell deformation. It
is possible that changes in dissolved gas partial pressures induced the
changes seen in the experiments by Acevedo et al. (1) with elevated
hydrostatic pressure. However, in other studies in which pressure was
elevated with the use of an inert gas and medium pH,
PO2, and PCO2 were
carefully monitored, similar responses were seen (16, 23). In our
experiments, pressurization would cause a small (<1%) increase in
the gas partial pressures above the cells, but pressure in the culture
medium was not elevated. If changes in the gas partial pressure had
induced gene expression, then we would have expected to also observe
upregulation in the hydrostatic pressure experiments. Because we did
not, we conclude that the slight changes in gas partial pressures were not responsible for the changes in gene expression that occurred with
transmembrane pressure application.
We can only speculate on how the cell is deformed to transduce the
signal, assuming that deformation, leading to protein conformational changes, is ultimately responsible. Raising the pressure above the
cells might cause liquid to be expelled from the cells, raising intracellular osmotic pressure and inducing cell volume regulation. Using osmotic water permeabilities of 40-50 µm/s for distal
airway epithelium (12) in the analysis of Fischbarg et al. (11) and assuming no concentration gradients to retard water flux (for maximum
cell volume change) give a change in cell layer height of
5-7 × 10
4 µm/s with 20 cmH2O pressure application. These changes could cause a
transient reduction in cell layer thickness before volume regulation
and might therefore cause sufficient deformation to transduce a
biological signal.
Transmembrane pressure could also force liquid through the
intercellular gap, producing a shear stress on the cells. In an experiment similar to ours, Dull et al. (9) measured solute and albumin
fluxes across endothelial cell monolayers by pressurizing the medium on
the apical surface of the cells. Dull et al. also grew their cells on
Transwell inserts, and the pressures they used ranged from 5 to 10 cmH2O. Volume fluxes of medium without albumin ranged from
3 × 10
5 to 1.4 × 10
4
ml/s, which would be enough to force 1 ml of liquid across the cells in
2-9 h. The amount of surface liquid initially covering our cell
cultures would be inadequate to produce such high volume fluxes, and
the surface of our cells did not appear dry after 6 h of pressure application.
If fluid flux did occur, we may estimate the magnitude of shear stress
(
w) by a force balance equating the pressure gradient (
P) acting over the cell layer and the shear stress in the
intercellular space:
P · h
2
wL, where L is the thickness of the
cell layer and h is the average intercellular gap size. The
maximum
P (20 cmH2O) applied across a 20-µm-thick cell
layer with an h of 200 nm (estimated from the transmission
electron micrographs) may produce a shear stress of ~0.1
cmH2O (or 100 dyn/cm2) on the cells. Shear
stresses as low as 10-25 dyn/cm2 have been shown to
induce gene expression in endothelial cell cultures (28, 32), so this
mechanism of stimulation cannot be ruled out.
Pressure applied to the apical surface might also cause the cells to be
deformed into the pores of the culture membrane. One study using hollow
fiber membranes for separation of stem cells from bone marrow showed
that long extensions of the cell membrane and cytoplasm extruded into
the pores of the filtration membrane with 40 mmHg positive-pressure
application (30). We attempted to visualize cell deformations at the
membrane surface by transmission electron microscopy, but this proved
inconclusive because the Transwell membrane was destroyed during
processing (data not shown). No obvious cell extensions in the
transmission electron or light micrographs were seen, however.
Histological processing may have removed the appearance of the
extensions, or the deformations into the pores were too small to detect
by our methods. Deformations in the epithelial cell membranes are
visible in micrographs of highly constricted airways (51). If
deformation of the cell membrane is the stimulus for our response, then
it is likely that this mechanism is present in both the in vitro and in
vivo situations.
In highly constricted airways, there are likely to be several
mechanical forces acting on the cells within the wall. Compressive stresses on the epithelial cells within the folds appear to deform the
cell shape, squeezing the cells out of the fold toward the luminal
space. Fluid movement within the wall and out of the wall may produce
extracellular fluid shear stress and transcellular fluid pressure
gradients, leading to cellular volume change and deformation.
Therefore, potential transduction mechanisms in these transmembrane
pressure experiments (such as deformation of the cell shape, fluid
shear stress, and cellular fluid loss) are also likely to be present in
constricted airways in vivo.
Concluding observations.
The structure of the airway wall changes markedly in patients with
asthma (20). Beneath the epithelial basement membrane is a fibrotic
layer that is significantly thickened (35). It is generally believed
that the increased amount of extracellular matrix material in the
subepithelial collagen layer is produced by myofibroblasts (7) in
response to inflammation (18, 35). Our results suggest that another
potential mechanism may be at work, that the response of airway
epithelial cells to mechanical stresses during prolonged smooth muscle
constriction leads to elaboration of transcription factors and
cytokines that could lead to airway wall remodeling. The compressive
stresses studied were similar to those stresses in a constricted,
buckled airway and are not present during normal lung function. The
genes we selected to study have been shown to play a role in tissue
remodeling. Egr-1 activates transcription of important remodeling genes
such as PDGF and TGF-
1 (24). ET-1 is a potent bronchoconstrictor, stimulates collagen secretion from fibroblasts (4), and is present in
bronchial epithelium of symptomatic asthmatic patients (2). TGF-
1
plays a key role in lung fibrosis (6) and is expressed in asthmatic
airways (26, 42). Thus production of these profibrotic mediators by the
epithelium in response to mechanical stress could stimulate the
fibroblasts to remodel the airway wall.
In conclusion, airway epithelial cell cultures stimulated by static
transmembrane pressure upregulate expression of mRNAs for Egr-1, ET-1,
and TGF-
1 in a magnitude- and time-dependent manner. Egr-1 protein
is also upregulated by mechanical stress. It is unclear how
transmembrane pressures stimulate our cell cultures, but we can
eliminate the effects of hydrostatic pressure and membrane strain as
possible mechanisms. The gene induction response may be similar to that
of airway epithelium in vivo when severe prolonged smooth muscle
constriction causes the airway wall to buckle and the epithelium folds
into deep crevasses.
 |
ACKNOWLEDGEMENTS |
The authors thank Melody A. Swartz for assistance with immunohistochemistry.
 |
FOOTNOTES |
This work was supported by the Whitaker Foundation; National Heart,
Lung, and Blood Institute Grants HL-33009 and HL-54759; and the Freeman Foundation.
The costs of publication of this
article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: R. D. Kamm,
Massachusetts Institute of Technology, Dept. of Mechanical Engineering,
Rm. 3-260, 77 Massachusetts Ave., Cambridge, MA 02139 (E-mail:
rdkamm{at}mit.edu).
Received 13 July 1999; accepted in final form 18 January 2000.
 |
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