Department of Bioengineering, University of Pennsylvania, Philadelphia, Pennsylvania 19104
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Mechanical ventilation with high
tidal volumes has been shown to contribute to the formation or
worsening of interstitial and alveolar edema. Previously we showed that
application of large biaxial deformations in vitro perturbs the
concentration and distribution of functional tight junction proteins in
alveolar epithelial cells. Using a novel method, we determined that
applied epithelial strain increases paracellular permeability in a
dose- and rate-dependent manner. Primary rat alveolar epithelial cells
were subjected to 12%, 25%, or 37% change in surface area (SA)
cyclic equibiaxial stretch for 1 h. Cells were also stretched
noncyclically at 25%
SA for 1 h. During the experimental
period, a fluorescently tagged ouabain derivative was added to the
apical fluid. Evidence of binding indicated functional failure of the
paracellular transport barrier. The percentage of field area stained
was quantified from microscopic images. There was no significant
evidence of basolateral fluorescent staining at 12%
SA or at 25%
SA applied cyclically or statically. However, cyclic stretch at 37%
SA resulted in significantly more staining than in unstretched cells
(P < 0.0001) or those stretched at either 12%
(P < 0.0001) or 25% cyclic (P < 0.0005) or static (P < 0.05)
SA. These results
suggest that large cyclic tidal volumes may increase paracellular
permeability, potentially resulting in alveolar flooding.
injury; lung; transport
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
PATIENTS SUFFERING FROM acute or chronic lung injury, or who experience poor pulmonary gas exchange, are able to receive respiratory assistance through controlled mechanical ventilation. However, ventilation with excessive regional lung volumes or high pressures can result in the development of ventilator-induced lung injury (VILI) (14). VILI occurs in ~5-15% of ventilated patients and carries an associated mortality rate of 34-60% (14, 20, 25), representing a significant health risk for patients requiring ventilation (20, 25). VILI is commonly marked by the formation or worsening of alveolar air leaks, interstitial and pulmonary edema, and alveolar cell dysfunction, indicative of decreased endothelial and epithelial barrier function (9).
Previously, investigators demonstrated a strong qualitative link between lung inflation volume and pulmonary barrier function in whole lung and animal models. In an open-chest preparation, the number of pulmonary epithelial and endothelial breaks increased at higher lung volumes, regardless of the transmural pressure of the alveolar wall (13). When animals were ventilated with high airway pressures but normal tidal volumes, lungs were uninjured, as determined by wet-to-dry ratio and albumin permeability (8). Conversely, ventilation with high tidal volumes, with or without high airway pressure, resulted in increased wet-to-dry ratios and permeability to albumin in these studies, demonstrating that large tidal volumes impair barrier properties of previously uninjured lungs.
To quantify the extent of barrier dysfunction due to "volutrauma," investigators have measured transport of large and small solutes across the epithelium, often expressing the results as an equivalent pore radius, a measure that increases as the epithelium allows larger molecules to pass more easily. Many of these findings indicate that high static inflation volumes adversely affect the solute and fluid balance that is required for proper pulmonary function (10-12, 15). However, none of these studies examined lungs experiencing cyclic changes in volume, which is the most clinically relevant mode of ventilation. Also, these studies examined lobar or whole lung inflations, which are inherently heterogeneous, and did not examine the effects of lung inflation at the local, or cellular, level.
The cellular structures that provide the majority of resistance to paracellular transport are the gasketlike tight junctions (TJ) of the alveolar epithelium located at the apical end of the intercellular space between adjacent epithelial cells. Intact, they minimize paracellular transport, allowing the cells to regulate transport via transcellular pathways (16). Previously, we demonstrated (5) that application of high magnitudes of cyclic mechanical stretch decreases the concentration of functional TJ proteins in cultured alveolar epithelial cells and alters their distribution around the cell membrane. Given the importance of the TJ in maintaining the alveolar epithelial paracellular transport barrier, we hypothesized that these stretch-induced cytostructural changes are associated with an increase in epithelial paracellular permeability.
In this article, we test this hypothesis by examining the alveolar
epithelial paracellular permeability response to changes in the
magnitude and mode of stretch. The common measures of epithelial permeability, such as transepithelial electrical resistance and transport of labeled tracers (3, 6, 18), require the cells to be cultured onto a permeable substrate. Furthermore, stretching of
cultured cells usually involves the use of elastic and flexible substrates. Heretofore, the lack of cell culture substrates that are
permeable, elastic, and biocompatible hindered the measurement of
stretch-induced alterations in paracellular permeability in cultured
cells. In this article we present a new method to assess paracellular
permeability useful for alveolar epithelial cells cultured onto an
impermeable substrate. Changes in paracellular permeability of cultured
alveolar epithelial cells were evaluated after application of stretch
at various magnitudes under both cyclic and static stretch modes. At a
given magnitude, stretch mode did not affect paracellular permeability
in these cells. Paracellular permeability increased after 1 h of
cyclic stretch at 37% change in surface area (SA), consistent with
our previously reported structural data. Together, these results
indicate that sustained cycling at physiological magnitudes of alveolar
strain may impair epithelial barrier function, increasing the
likelihood of pulmonary edema formation or enhancement.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Method rationale.
The technique used for the assessment of paracellular permeability
described in this article is made possible by the polarity that
alveolar epithelial cells display and by the extremely low paracellular
permeability that confluent monolayers of type I cells demonstrate, as
indicated by their high transepithelial electrical resistance of
~2,000 · cm2 (6). These cells
possess functional Na+-K+-ATPases only on the
basolateral surfaces of the plasma membrane (26). The
Na+-K+-ATPases are inhibited by the compound
ouabain, which has a high affinity for the extracellular domain of this
transmembrane protein in all cell types. In designing this study, we
hypothesized that paracellular permeability will increase when cells
are stretched above a certain threshold. Thus, if ouabain is present
only in the fluid surrounding the apical surface of a confluent
monolayer of type I-like cells, stretched cells will allow a greater
amount of ouabain transport through the TJ than unstretched cells and will therefore demonstrate a greater extent of ouabain-ATPase binding.
To detect the appearance of ouabain in the basolateral space and to
measure the extent of binding, we used a derivative of ouabain bound to
the fluorescent dye BODIPY-FL that possesses the same specificity of
binding as unconjugated ouabain.
Cell culture protocol. The protocol for lung cell isolation was approved by the University of Pennsylvania Institutional Animal Care and Use Committee and is based on the method of Dobbs and coworkers (7). Alveolar type II cells were isolated from healthy male Sprague-Dawley rats (180-200 g). The rats were anesthetized with pentobarbital sodium (50 mg/kg body wt ip). The trachea was cannulated, the lungs were mechanically ventilated, an abdominal aortotomy was performed to exsanguinate the animal, and excess blood was removed via pulmonary arterial perfusion. The lungs were then excised and instilled with an elastase solution (3 U/ml; Worthington Biochemical, Lakewood, NJ) and minced in the presence of deoxyribonuclease (Sigma, St. Louis, MO) with a tissue chopper (Sorvall, Newtown, CT). The elastase reaction was stopped with fetal bovine serum (Life Technologies, Rockville, MD). Cells were filtered through progressively finer Nitex mesh (Crosswire Cloth, Bellmawr, NJ), and plated on an IgG-coated culture dish (3 mg/5 ml Tris · HCl). After a 1-h incubation at 37°C, gentle panning isolated type II cells from the macrophages and contaminating cells preferentially adhered to the culture dish. Ultimately, cells were spun down and resuspended in minimum essential medium (MEM) with Earle's salts and supplemented with 10% fetal bovine serum, 25 µg/ml gentamicin, and 0.25 µg/ml amphotericin B (Life Technologies). Cell purity of this isolation procedure is >95%, as determined by phosphine 3R staining of adherent cells (17).
All cells were seeded at a density of 1 × 106 cells/cm2 onto fibronectin-coated (10 µg/cm2; Boehringer Mannheim Biochemicals, Indianapolis, IN) flexible Silastic membranes (Specialty Manufacturing, Saginaw, MI) mounted in custom-made wells. The cells were cultured in MEM supplemented as above for 5 days. It was reported previously that rat alveolar type II cells differentiate into type I cells (which cover >90% of the alveolar surface in vivo) after 5 days under similar culture conditions (4). The medium was replaced daily. By the fifth day of culture, the cells had formed a confluent monolayer and displayed a phenotype consistent with that observed for cultured type I cells. We showed previously (19) that >95% of identically cultured cells stain positively for RTI-40, a type I cell-specific surface antigen. After 5 days, the cells were washed with dye-free Dulbecco's modified Eagle's medium (DMEM; Life Technologies) and subjected to the stretch protocol described in Stretch application. Unstretched wells served as controls in all experiments. Four isolations were performed for each stretch magnitude. Two wells per isolation were stretched at each magnitude, resulting in a total of eight wells for each group. The unstretched control group also consisted of two wells per isolation.Stretch application.
Wells were mounted onto a custom-built cell-stretching device capable
of applying equibiaxial strain to the samples at a precise user-defined
magnitude and frequency, as previously described (22).
Cells were stretched cyclically at 15 cycles/min for 1 h at 12%,
25%, or 37% SA. These changes in surface area approximately correspond to strains experienced by the alveolar epithelium in vivo at
70%, 90%, and 100% total lung capacity (TLC), respectively (23). An additional set of wells was stretched statically
(0 cycles/min) for 1 h at 25%
SA. As a positive control, a
separate population of unstretched cells was scraped once with a rubber policeman. During the 1-h stretch period, or for 1 h after the scrape, the apical surface of the stretched and unstretched cells was
bathed in DMEM containing 2 µM BODIPY-ouabain (Molecular Probes, Eugene, OR). Nonspecific binding was evaluated by adding 2 µM unconjugated BODIPY (Molecular Probes) to a separate population of two
wells stretched at 37%
SA. Cells were stretched in darkness to
prevent photobleaching of the dye. The temperature of the stretch device was maintained at 20°C to minimize endocytosis of the dye but
not impair cellular homeostasis.
Microscopic examination. Wells were mounted onto a Nikon TE-300 inverted epifluorescence microscope and examined at ×20 magnification. Two fields from each well were selected randomly, and two images of each field with a phase objective or a green (BODIPY-ouabain fluorescence) emission filter were captured and stored with Metamorph imaging software (Universal Imaging, West Chester, PA) and a microscope-mounted Hamamatsu camera and controller. Identical image acquisition times were used for all images of each type acquired.
After all images were captured, we wanted to distinguish whether the observed green fluorescence was due to endocytosis of BODIPY-ouabain into the cell or whether BODIPY-ouabain had in fact bound to the basolateral surface of the plasma membrane. Anti-BODIPY antibody (Molecular Probes) was added in a 1:40 dilution to one of the two wells from each group in every isolation. All wells were then kept overnight at 4°C to inhibit nonspecific endocytosis (21, 24), and images were obtained again as above on the following day. Anti-BODIPY, when bound with BODIPY-ouabain, quenches the fluorescent signal of the BODIPY-ouabain bound to Na+-K+-ATPase pumps. Given the large size of the anti-BODIPY molecule (radius ~15-20 Å) and the low incubation temperature, it was expected that anti-BODIPY would not be internalized by the cells. To rule out the possibility that loss of BODIPY-ouabain fluorescence overnight could be attributed to dye photobleaching and not antibody quenching, the other well from each group was incubated overnight without the antibody for comparison. If both wells remained brightly fluorescent, the BODIPY-ouabain had been internalized. If both became dark, the BODIPY-ouabain had been photobleached. However, if wells treated with anti-BODIPY demonstrated reduced fluorescence compared with those without antibody treatment, we would conclude that the BODIPY-ouabain was bound in the extracellular domain.Additional experimentation.
The influence of specific endocytotic pathways on cell staining was
determined by treating one unstretched well and one well stretched at
37% SA from each of three isolations with 5 µM phenylarsine oxide
(PAO; Sigma), an inhibitor of receptor-mediated endocytosis. These
wells were treated during the entire 1-h experimental period with PAO,
BODIPY-ouabain, and ethidium homodimer-1 and examined as described in
Microscopic examination. One untreated, unstretched well and
one untreated well stretched at 37%
SA from each isolation served
as untreated controls. Treatment of alveolar epithelial cells with PAO
using a similar concentration and duration has been shown to decrease
endocytotic activity without adversely affecting cellular viability
(2). In a limited manner, cells were also treated with a
higher concentration of PAO (10 µM), but these results were no
different than those at 5 µM (data not shown). Significantly more
BODIPY-ouabain staining in the stretched, PAO-treated group compared
with the stretched, untreated group would indicate that
receptor-mediated endocytosis was involved in stretch-induced
BODIPY-ouabain binding.
Data analysis. With Metamorph, a fluorescence intensity threshold was applied to each fluorescence image to examine the signal above background fluorescence levels. Background fluorescence levels were determined for each isolation by selecting a large region from each unstretched fluorescence image and calculating the maximum pixel intensity in each of these regions. This intensity was used as the threshold for all stretched and unstretched images from that isolation. Occasional regions that contained non-cell-specific autofluorescence were not included. For each image, the pixels possessing intensities above the threshold and below saturation were counted and expressed as a percentage of total field area. Average intensity of these pixels was also calculated for each field of stretched cells. Stained area and intensity measurements from unstretched and stretched wells were compared with the Tukey test for significant differences across multiple groups (28). Results are expressed as means ± SE.
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Extent of BODIPY-ouabain stained area.
Both stretched and unstretched cells displayed at least a small degree
of BODIPY-ouabain staining (Fig. 1).
Cells stretched cyclically for 1 h at 12% SA (2.4 ± 0.6% of field area stained) or 25%
SA (4.2 ± 0.9%) did not
experience significantly greater BODIPY-ouabain staining than
unstretched, untreated cells (2.1 ± 0.3%). In contrast, cyclic
stretch at 37%
SA increased the area of BODIPY-ouabain staining to
16.8 ± 4.7% of the field. This increase was significant compared
with unstretched cells and with all other stretched populations (Fig.
2). BODIPY-ouabain staining was not
preferentially colocalized with ethidium homodimer-1 binding, indicating that plasma membrane rupture was not a main factor by which
BODIPY-ouabain was able to bind to cells (Fig.
3). Previously, we showed
(5) that this stretch regimen alters distribution and
expression of the functional TJ protein occludin in similarly cultured
cells. Together, these results indicate that stretch magnitude is a
determinant of the extent of paracellular barrier dysfunction in
cultured alveolar epithelial cells. To determine whether this increase
in fluorescence could be due to an increase in visible cellular surface
area, we measured the surface areas of individual cells before stretch
application and compared them to the areas of the same cells measured
after 1 h of 37%
SA cyclic stretch. Paired analysis of the
measured areas did not reveal a significant change in cell surface area
(data not shown).
|
|
|
|
|
|
Intensity of BODIPY-ouabain staining.
Average intensity of suprathreshold pixels in stretched cell fields did
not differ significantly from those in unstretched control wells at
12% (108.0 ± 4.3% of average intensity of controls), 25%
cyclic SA (103.9 ± 6.6%), and 25% static
SA (121.0 ± 5.5%; Fig. 7). However, at 37%
SA, intensity was significantly higher than in controls (130.7 ± 5.9%; P < 0.01). This result suggests that this
magnitude of stretch increases the amount as well as the extent of
BODIPY-ouabain binding to the basolateral surface.
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Our findings support the hypothesis that moderate static and
cyclic inflation volumes do not adversely affect paracellular barrier
function. The BODIPY-ouabain staining in unstretched, untreated cells
was low, averaging 2.1% of the microscopic field areas examined.
Cyclic stretch for 1 h at 12% or 25% SA did not result in an
increase in fluorescence. One hour of static stretch at 25%
SA did
not produce a significantly different staining intensity compared with
control levels or with staining after cyclic stretch at 25%
SA.
Thus we conclude that paracellular permeability did not increase in
cells stretched at these magnitudes, which correspond roughly to
inflations to lung volumes averaging 70% and 90% TLC
(23). In contrast, 1 h of cyclic inflations to 100%
TLC (23) (37%
SA) resulted in significantly
increased paracellular transport (16.8% of the field stained),
compared with all other experimental groups.
Given that mechanical stimuli can induce endocytosis in other kinds of epithelia (1), we performed specific studies to rule out this pathway as a significant mechanism responsible for the observed increase in fluorescence after stretch. First, PAO treatment did not affect dye binding in stretched cells (Fig. 5), ruling out the contribution of receptor-mediated endocytosis to the process of BODIPY-ouabain binding. Second, staining cells after stretch instead of during stretch did not affect the extent of BODIPY-ouabain binding, ruling out the contribution of specific stretch-induced endocytosis. Third, stretch studies with BODIPY-ouabain administered at 37°C were followed by treatment with anti-BODIPY at 4°C. This temperature has been reported to block endocytosis in similar alveolar epithelial cells (21, 24), restricting the quenching action of anti-BODIPY to externally bound BODIPY-ouabain only. The treatment quenched the fluorescent staining from 16.8% to 1.2% of the field area (Fig. 4). Thus we conclude that most of the BODIPY-ouabain is bound extracellularly, ruling out nonspecific endocytotic pathways. Fourth, our TX experiments, which demonstrated significantly higher fluorescence after cell permeabilization compared with stretch-induced fluorescence levels (Fig. 6), disprove the hypothesis that stretch-induced increases in plasma membrane permeability, rather than TJ permeability, are primarily responsible for the findings presented in Fig. 2. Fifth, we show that stretch did not increase the amount of cell staining with unconjugated BODIPY (Fig. 2), confirming that the stretch-induced increases in fluorescence were specific to increased numbers of BODIPY-ouabain-bound sites rather than nonspecific stretch-induced increases in intracellular dye concentration. Finally, BODIPY-ouabain fluorescence was demonstrated in unstretched and intact cells near a scraped monolayer region (Fig. 1), suggesting that any type of TJ disruption, not merely mechanical stretch, can result in increased BODIPY-ouabain staining.
Our data also reveal additional information regarding BODIPY-ouabain
binding in these cells. The fact that anti-BODIPY is apparently able to
diffuse through the TJ and quench BODIPY-ouabain fluorescence provides
evidence that barrier injury in this model is irreversible over the
24-h experimental period. The absence of a strong correlation between
regions of strong BODIPY-ouabain and ethidium homodimer-1 binding
indicate that plasma membrane rupture does not by itself produce
cellular BODIPY-ouabain binding. Also, because treatment with
unconjugated BODIPY did not produce a significant difference in
staining between cells stretched at 37% SA and unstretched cells,
we can conclude that the binding of BODIPY-ouabain to the cell exterior
was specific to the Na+-K+-ATPase sites. The
average intensity of the stained regions also increased at 37%
SA,
indicating that a higher amount of BODIPY-ouabain was bound in these
regions than in the stained regions of monolayers stretched at lower magnitudes.
Our cyclic stretch data from the current study indicate the possibility
of a stretch magnitude threshold between 25% and 37% SA above
which barrier integrity suffers. This stretch magnitude corresponds to
the average strain experienced by the alveolar epithelium between 90%
and 100% TLC and is the same threshold magnitude previously discovered
for TJ structural disruption in cells cultured under identical
conditions (5). Both studies provide evidence that high,
yet physiologically relevant, epithelial strains can cause alveolar
dysfunction and injury in an in vitro preparation.
Similar threshold-dependent, stretch-induced TJ injury at high lung inflation volumes has been observed in animal preparations using tracers of similar size to BODIPY-ouabain (radius ~20 Å). Egan and coworkers examined the effects of static inflation on whole sheep and rabbit lungs, noting that inflation below 100% TLC did not produce an increase in paracellular permeability to tracers of 14- to 34-Å radius (11, 12). Inflation volumes above 100% TLC removed all resistance to paracellular transport of these molecules, indicative of a complete loss of epithelial barrier function (11, 12). These injury thresholds determined in whole lung studies agree with our structural (5) and functional (current study) findings in vitro in a primary cell culture preparation.
Additional permeability assessment has been performed in animals with tracers of smaller size (radius <10 Å), which provide more sensitive detection of impaired barrier properties. Egan et al. (10, 12) observed that static inflation of dog and sheep lungs at 75-80% TLC produced an increase in paracellular permeability to tracers of small size. In contrast, Kim and Crandall (15) noted that static inflation of bullfrog lungs within the physiological range did not result in increased paracellular transport to these tracers and that only supraphysiological inflation caused increased solute and fluid transport across the alveolar barrier in these animals. Thus there is some disagreement in the literature over the extent of injury measured with small solutes at static inflation volumes below 100% TLC, but the results of Kim and Crandall support the current findings of a decrease in TJ integrity at or above 100% TLC.
In humans, repeated inflation of the whole lung to 80-85% TLC can occur in well-trained athletes during vigorous exercise with no apparent loss of pulmonary epithelial barrier function (27). These lung inflation volumes correspond to an alveolar epithelial change in surface area of <25% (23), a stretch magnitude that was found to be below the injury threshold in this study. Thus these in vivo observations are consistent with the results of the in vitro experiments presented here.
The main advantage provided by this in vitro experimental system is that the local physical environment of a monolayer of cultured cells can be controlled more readily than that of a whole lung. Furthermore, the relative contributions of the cellular and extracellular components of the monolayer and of the apical milieu can be investigated more effectively than in whole lungs. However, although our model of alveolar epithelial injury possesses type I-like pneumocytes that cover the vast majority of the alveolar epithelium, it is not fully representative of conditions in the in vivo alveolus. It does not contain macrophages or fibroblasts, and therefore the composition of the extracellular matrix and of the interstitial and luminal milieu may not be identical to that of the viable whole lung. Additionally, after 5 days in culture, the majority of type II cells differentiate to type I phenotype (19), resulting in a lack of surfactant production that may alter the surface tension milieu of the in vitro alveolar epithelium. Once the basic mechanisms of stretch-induced alveolar epithelial dysfunction are elucidated with new methods such as that presented here, a more representative in vitro model of the epithelium may be constructed to determine how these other constituents of the in vivo alveolus affect the susceptibility of alveolar epithelial cells to mechanical injury.
Using a novel technique for the assessment of paracellular permeability, we have demonstrated that the paracellular permeability of stretched alveolar epithelial cells in culture is dependent on the magnitude of the applied stretch. This study represents the first time that paracellular permeability has been assessed in cultured and mechanically stretched epithelial cells, and this method should prove useful to other transport physiologists, especially those interested in cellular injury. Although this study provides evidence that alveolar epithelial barrier integrity is altered at high magnitudes of applied strain, for a more complete description there is a need to supplement these findings with data from tracers of varying size and charge to determine critical cell deformations required to disrupt the epithelial barrier to transport of small charged solutes, large macromolecules, and other types of physiologically relevant compounds. After identifying the chemical pathways by which excessive stretch is transduced into an injurious cellular response, we may develop injury interventions to reduce the severity of stretch-induced alveolar epithelial injury and minimize the incidence and mortality of ventilator-induced lung injury.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. James F. Sanzo of the University of Pennsylvania for assistance with microscopy and Dr. Kenneth R. Spring of the National Institutes of Health for helpful discussions.
![]() |
FOOTNOTES |
---|
This work was supported by National Heart, Lung, and Blood Institute Grant HL-57204, National Science Foundation Grant BES-9702088, and the Whitaker Foundation. K. J. Cavanaugh was supported by a Whitaker Graduate Fellowship.
Address for reprint requests and other correspondence: S. S. Margulies, Dept. of Bioengineering, Univ. of Pennsylvania, 3320 Smith Walk, Philadelphia, PA 19104 (E-mail: margulie{at}seas.upenn.edu).
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. Section 1734 solely to indicate this fact.
August 14, 2002;10.1152/ajpcell.00341.2002
Received 23 July 2002; accepted in final form 6 August 2002.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Apodaca, G.
Modulation of membrane traffic by mechanical stimuli.
Am J Physiol Renal Physiol
282:
F179-F190,
2002
2.
Bates, SR,
Tao JQ,
Schaller S,
Fisher AB,
and
Shuman H.
Lamellar body membrane turnover is stimulated by secretagogues.
Am J Physiol Lung Cell Mol Physiol
278:
L443-L452,
2000
3.
Berg, MM,
Kim KJ,
Lubman RL,
and
Crandall ED.
Hydrophilic solute transport across rat alveolar epithelium.
J Appl Physiol
66:
2320-2327,
1989
4.
Borok, Z,
Danto SI,
Zabski SM,
and
Crandall ED.
Defined medium for primary culture de novo of adult rat alveolar epithelial cells.
In Vitro Cell Dev Biol Anim
30A:
99-104,
1994.
5.
Cavanaugh, KJ,
Oswari J,
and
Margulies SS.
Role of stretch on tight junction structure in alveolar epithelial cells.
Am J Respir Cell Mol Biol
25:
584-591,
2001
6.
Cheek, JM,
Kim KJ,
and
Crandall ED.
Tight monolayers of rat alveolar epithelial cells: bioelectric properties and active sodium transport.
Am J Physiol Cell Physiol
256:
C688-C693,
1989
7.
Dobbs, LG,
Gonzalez R,
and
Williams MC.
An improved method for isolating type II cells in high yield and purity.
Am Rev Respir Dis
134:
141-145,
1986[ISI][Medline].
8.
Dreyfuss, D,
and
Saumon G.
Role of tidal volume, FRC, and end-inspiratory volume in the development of pulmonary edema following mechanical ventilation.
Am Rev Respir Dis
148:
1194-1203,
1993[ISI][Medline].
9.
Dreyfuss, D,
and
Saumon G.
Ventilator-induced lung injury: lessons from experimental studies.
Am J Respir Crit Care Med
157:
294-323,
1998
10.
Egan, EA.
Response of alveolar epithelial solute permeability to changes in lung inflation.
J Appl Physiol
49:
1032-1036,
1980
11.
Egan, EA.
Lung inflation, lung solute permeability, and alveolar edema.
J Appl Physiol
53:
121-125,
1982
12.
Egan, EA,
Nelson RM,
and
Olver RE.
Lung inflation and alveolar permeability to non-electrolytes in the adult sheep in vivo.
J Physiol
260:
409-424,
1976[Abstract].
13.
Fu, Z,
Costello ML,
Tsukimoto K,
Prediletto R,
Elliott AR,
Mathieu-Costello O,
and
West JB.
High lung volume increases stress failure in pulmonary capillaries.
J Appl Physiol
73:
123-133,
1992
14.
Haake, R,
Schlichtig R,
Ulstad DR,
and
Henschen RR.
Barotrauma. Pathophysiology, risk factors, and prevention.
Chest
91:
608-613,
1987[ISI][Medline].
15.
Kim, KJ,
and
Crandall ED.
Effects of lung inflation on alveolar epithelial solute and water transport properties.
J Appl Physiol
52:
1498-1505,
1982
16.
Lubman, RL,
Kim KJ,
and
Crandall ED.
Alveolar epithelial barrier properties.
In: The Lung: Scientific Foundations (2nd ed.), edited by Crystal RG,
and West JB.. Philadelphia: Lippincott-Raven, 1997, p. 585-602.
17.
Mason, RJ.
Transepithelial transport by pulmonary alveolar type II cells in primary culture.
Proc Natl Acad Sci USA
79:
6033-6037,
1982[Abstract].
18.
Matsukawa, Y,
Lee VH,
Crandall ED,
and
Kim KJ.
Size-dependent dextran transport across rat alveolar epithelial cell monolayers.
J Pharm Sci
86:
305-309,
1997[ISI][Medline].
19.
Oswari, J,
Matthay MA,
and
Margulies SS.
Keratinocyte growth factor reduces alveolar epithelial susceptibility to in vitro mechanical deformation.
Am J Physiol Lung Cell Mol Physiol
281:
L1068-L1077,
2001
20.
Parker, JC,
Hernandez LA,
and
Peevy KJ.
Mechanisms of ventilator-induced lung injury.
Crit Care Med
21:
131-143,
1993[ISI][Medline].
21.
Stroetz, RW,
Vlahakis NE,
Walters BJ,
Schroeder MA,
and
Hubmayr RD.
Validation of a new live cell strain system: characterization of plasma membrane stress failure.
J Appl Physiol
90:
2361-2370,
2001
22.
Tschumperlin, DJ,
and
Margulies SS.
Equibiaxial deformation-induced injury of alveolar epithelial cells in vitro.
Am J Physiol Lung Cell Mol Physiol
275:
L1173-L1183,
1998
23.
Tschumperlin, DJ,
and
Margulies SS.
Alveolar epithelial surface area-volume relationship in isolated rat lungs.
J Appl Physiol
86:
2026-2033,
1999
24.
Vlahakis, NE,
Schreoder MA,
Pagano RE,
and
Hubmayr RD.
Deformation-induced lipid trafficking in alveolar epithelial cells is both temperature and cholesterol dependent (Abstract).
FASEB J
15:
A494,
2001.
25.
Ware, LB,
and
Matthay MA.
The acute respiratory distress syndrome.
N Engl J Med
342:
1334-1349,
2000
26.
Waters, CM,
Ridge KM,
Sunio G,
Venetsanou K,
and
Sznajder JI.
Mechanical stretching of alveolar epithelial cells increases Na+-K+-ATPase activity.
J Appl Physiol
87:
715-721,
1999
27.
Whipp, BJ,
and
Pardy RL.
Breathing during exercise.
In: Handbook of Physiology. The Respiratory System. Mechanics of Breathing. Bethesda, MD: Am Physiol Soc, 1986, sect. 1, vol. III, pt. 2, p. 605-629.
28.
Zar, J.
Biostatistical Analysis. Upper Saddle River, NJ: Prentice Hall, 1999.