Deformation-induced lipid trafficking in alveolar epithelial cells

Nicholas E. Vlahakis1, Mark A. Schroeder1, Richard E. Pagano1,2, and Rolf D. Hubmayr1,3

1 Thoracic Diseases Research Unit, Division of Pulmonary and Critical Care Medicine, Department of Internal Medicine, and Departments of 2 Biochemistry and Molecular Biology and 3 Physiology and Biophysics, Mayo Clinic and Foundation, Rochester, Minnesota 55905


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Mechanical ventilation with a high tidal volume results in lung injury that is characterized by blebbing and breaks both between and through alveolar epithelial cells. We developed an in vitro model to simulate ventilator-induced deformation of the alveolar basement membrane and to investigate, in a direct manner, epithelial cell responses to deforming forces. Taking advantage of the novel fluorescent properties of BODIPY lipids and the fluorescent dye FM1-43, we have shown that mechanical deformation of alveolar epithelial cells results in lipid transport to the plasma membrane. Deformation-induced lipid trafficking (DILT) was a vesicular process, rapid in onset, and was associated with a large increase in cell surface area. DILT could be demonstrated in all cells; however, only a small percentage of cells developed plasma membrane breaks that were reversible and nonlethal. Therefore, DILT was not only involved in site-directed wound repair but might also have served as a cytoprotective mechanism against plasma membrane stress failure. This study suggests that DILT is a regulatory mechanism for membrane trafficking in alveolar epithelia and provides a novel biological framework within which to consider alveolar deformation injury and repair.

mechanical ventilation; BODIPY lipids; plasma membrane; lung injury; deforming stress


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

MECHANICAL VENTILATION is the only proven treatment for patients with acute respiratory distress syndrome (28). In these patients, dependent alveolar lung units are collapsed or edema filled, whereas nondependent units are air filled (13). High tidal volume ventilation has been used as a means to recruit dependent alveolar units and thereby improve patient oxygenation. However, in aerated lung units, this leads to alveolar overdistension and, in turn, high lung tissue strains. In animal models, it is readily apparent that high tidal volume mechanical ventilation directly results in lung injury, edema, and inflammation, referred to as ventilator-induced lung injury (VILI) (10, 17, 18). Morphological analysis of the cells that compose the alveolus reveals significant epithelial and endothelial cell injury characterized by blebbing, intercellular and intracellular gap formation, and the resultant denudation of the alveolar basement membrane (11, 12). The aim of the present study was to investigate the effect of basement membrane deformation on alveolar epithelial cell injury. More specifically, we sought to investigate the biological mechanisms used by the alveolar epithelial cell to ensure the integrity of its plasma membrane (PM).

The response of cells to mechanical forces can be viewed very simply by considering the cell as a structural network (cytoskeleton) enveloped by a lipid bilayer (PM). It is the cytoskeleton that determines cell shape and modulates its structure to coordinate shape changes, whereas the PM responds to these cytoskeletal changes to maintain an intact and functional cellular barrier. The cytoskeleton is integrally associated with the extracellular matrix or basement membrane via focal adhesion complexes such that deformations of the basement membrane are transmitted to the overlying cell through these connections. As lung volume increases from functional residual capacity, the alveolar septum unfolds, and at volumes closer to total lung capacity, the septum is placed under stress (3, 14). Thus, with alveolar overdistension, there is a resultant deformation of the basement membrane and a concurrent strain applied to the adherent epithelial cell. Once deformed, the epithelial cell must increase its surface area. This increase in surface area leads to PM stress and, inevitably, stress failure unless the PM is able to "grow." As a means to adapt to this stress, the cell might "unfold" the ruffles of its PM or the lipid bilayer itself might stretch laterally. Although the PM possesses a high shear and bending capacity, it can accommodate lateral stretch of only 3-4%, and thus the protective contribution of the elasticity of the membrane is limited (24). We hypothesized that in response to basement membrane deformation, alveolar epithelial cells translocate lipid molecules to the PM as a further adaptive mechanism to facilitate membrane growth and ultimately prevent membrane rupture.

To study this hypothesis, we used an in vitro cell deformation system to reduce the complex in vivo situation and at the same time simulate alveolar basement membrane strain. With the use of laser confocal microscopy (LCFM), the alveolar epithelial cell surface area was found to increase as a result of substratum deformation. In association with this morphological change, we report that alveolar epithelial cell deformation induced lipid trafficking to the PM in a stretch amplitude-dependent fashion. Surprisingly, cell deformation did not result in a large number of PM breaks, suggesting that deformation-induced lipid trafficking (DILT) might be an important mechanism not only for accommodation of increases in cell surface area but also in the maintenance of cell integrity.


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. Human A549 cells (American Type Culture Collection, Manassas, VA), passages 78-88, were grown on six-well culture plates (BioFlex, Flexcell International, McKeesport, PA). The base of each culture plate consisted of a flexible collagen I-impregnated silicoelastic membrane with a surface area of 9.6 cm2. Ham's F-12K containing L-glutamine (2 mM), FCS (10%), and penicillin-streptomycin-amphotericin B (100 U/ml, 100 µg/ml, and 25 µg/ml, respectively; Sigma, St. Louis, MO) was used as a growth medium. A549 cells were passaged and seeded at a density of 20,800 cells/cm2 (200,000 cells/well) 48 h before each experiment, resulting in an 85-90% confluent monolayer of cells.

Cell strain device. A portable vacuum stretch device was developed for cell stretching and imaging by confocal microscopy. The BioFlex wells were placed in a Teflon-like (Delrin) rubber-sealed mold with six 1.5-cm-diameter loading posts and then fixed to the confocal microscope movable stage mount for easy imaging maneuverability (Fig. 1). The loading posts were lubricated with Braycote grease 804 (Castrol, Irvine, CA) to minimize friction between the membrane and loading post. The membrane and, in turn, the attached cells were then deformed from below by a portable vacuum device connected to the Delrin mold. Cells could be visualized both in their undeformed and deformed states with LCFM. A single stretch of 7 or 25% was applied to the cells and held on average for 1-2 min to allow for focusing and image acquisition.


View larger version (62K):
[in this window]
[in a new window]
 
Fig. 1.   Cell strain device for laser confocal microscopy (LCFM) imaging of alveolar epithelial cells. Diagram shows relationship to the confocal X,Y stage and water-immersion lens. Photo at top, representative single, 1-µm confocal images of cells demonstrating typically observed changes in cell cross-sectional area with deformation.

Correlation of vacuum pressure to membrane strain amplitude. To determine the strain amplitude applied to the BioFlex membranes by the portable vacuum device, five ink marks in a cross configuration were imprinted on the membrane overlying the loading post. The membrane was imaged with an analog video camera (charge-coupled device TRV82, Sony, New York, NY) in both the undeformed and strained states at vacuum pressures varying from 0 to 70 kPa. With a PC-based digital frame grabber (Imaging Technology, Woburn, MA), specific video frames were digitized. Individual marker positions were identified and saved with a user interactive PC-based software package (Thoracic Diseases Research Unit, Mayo Clinic, Rochester, MN). Percent membrane strain was calculated by tracking the changes in distance between all dot pair combinations and then averaging these results. Thus stretch amplitude refers to the percent change in the length of any line element in the membrane of the well.

Cell imaging. Cells were imaged with an argon ion laser scanning confocal microscope (Bio-Rad MRC 600, Hercules, CA) mounted on an upright microscope (BH2, Olympus, Melville, NY) immediately after membrane lipid labeling. Optical sections (1 µm) were obtained with a ×40 water-immersion objective lens (numerical aperture = 0.75) (Carl Zeiss, Thornwood, NY) and by moving the stage in one direction alone to eliminate backlash in the stepper motor. Fluorescent labels were excited with blue [wavelength (lambda ) = 488 nm] laser light, and emission wavelengths were collected simultaneously by green (lambda  = 520-560 nm) and red (lambda  > 590 nm) filters (Omega Optical, Brattleboro, VT) for BODIPY and by a single long band-pass filter block (lambda  > 515 nm) for 1,1-dioctadecyl-3,3,3'3'-tetramethylindocarbocyanine perchlorate (DiI), FM1-43, and FITC. At each optical section plane, images were digitized at eight-bit resolution, averaged over three scans with Kalman filtering to reduce background noise, and stored in arrays of 768 × 512 pixels.

Cell surface area and fluorescence quantitation. Cell surface area, membrane fluorescence characteristics, and computer-generated three-dimensional images of cells were calculated with an image display and manipulation package (ANALYZE, Mayo Foundation, Rochester, MN) (16). For surface area analysis, the entire cell was scanned; for fluorescence intensity measurements, a single image slice through the middle of the cell was acquired. In both cases, a single cell in both the undeformed and deformed states was scanned and analyzed. For surface area, the scan images were reconstructed into three-dimensional images and median filtered (5 × 5 × 5) to remove broad backgrounds and maintain small cell features (7). Surface area measurements were verified with established computer-based stereological techniques (16). Seed points for regions of interest were utilized and then fit to the cell by hand-drawn pixel intensity limitations.

Labeling of cell membranes with lipid probes. Three different fluorescent lipid labels were used for membrane labeling: DiI, BODIPY-FL C5-lactosylceramide (LacCer), and FM1-43 (Molecular Probes, Eugene, OR). For surface area calculations, cells were washed with 1× PBS and then incubated with 0.1% DiI (in ethanol) at 37°C for 10 min to label the PM. For lipid trafficking experiments, cell PMs were labeled with BODIPY-LacCer by washing with ice-cold 1× HEPES-buffered MEM (HMEM; without glucose) and incubation with 3 nM BODIPY-LacCer-defatted BSA complex (6) for 30 min at 4°C. Cells were then washed again with ice-cold HMEM before LCFM imaging. Finally, the intracellular membranes were labeled with 2 µM FM1-43 for 4 h at 37°C and then washed with 1× PBS before being imaged. In separate experiments, cells were ATP depleted by incubating them at 37°C for 30 min in the presence of 5 mM sodium azide (Sigma) and 50 mM 2-deoxy-D-glucose (Sigma).

Fluorescence spectrophotometry. The FM1-43 fluorescence (FFM1-43) in the supernatant from culture wells of A549 cells that were undeformed or strained by 25% was measured in an SLM 8000C spectrofluorometer (Urbana, IL). Cells were loaded with FM1-43 as described in Labeling of cell membranes with lipid probes and washed three times with PBS. Cells were exposed to no strain or to 25% strain, and the supernatant from the culture wells was collected. Twenty milliliters of supernatant were combined with 180 ml of 1% SDS (Sigma) in a cuvette, and the fluorescence was measured (lambda ex = 488 nm; lambda em = 565 nm).

Determination of PM breaks. To determine whether cellular deformation was associated with breaks in the PM, cells were incubated with FITC-dextran (Dx) (molecular mass = 4, 70, or 150 kDa; Fluka, Milwaukee, WI) for 10 min at 37°C and then stretched using the same parameters as described in Cell strain device. Cells with homogeneous intracellular distribution of FITC-Dx were counted in five low-power (×40) microscope fields in each culture well for each experiment and tabulated as a percent of the total cells counted (between 200 and 300 cells in total). Six separate experiments were performed for each Dx size.

Statistical analysis. All measurements are presented as means ± SD. Statistical comparisons between experimental conditions were made with Student's t-test for paired observations. Significance was assumed at P < 0.05 with respect to a two-tailed probability distribution. For stretch analyses, each cell served as its own static control. Quantification of lipid trafficking was also compared against a time-control cell population that was exposed to the same environmental conditions including laser light exposure time. Analysis of variance was used to determine 1) membrane strain uniformity resulting from applied transmembrane pressures and 2) strain effect on percent injured cells across different molecular mass FITC-Dx molecules.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

Validation of membrane strain amplitude and uniformity. The degree of membrane strain produced by the vacuum pump at increasing pressures and its reproducibility and uniformity were analyzed. As demonstrated in Fig. 2, this strain-pressure relationship was nonlinear and varied on average by 2-3% at any given pressure. Cell deformation experiments were performed at 20 and 40 kPa (150 and 300 mmHg) pressure, resulting in membrane strains of 7 ± 2.3 and 25 ± 2.8%, respectively. The values for percent membrane strain represent the mean results of strain values from each of 6 ink-marked pairs in each experiment (12 in total). Using multiple regression analysis of variance, we determined that membrane strain was uniform when analyzed across all 12 ink-marked pairs (P = 0.7).


View larger version (13K):
[in this window]
[in a new window]
 
Fig. 2.   Relationship between applied transmembrane pressure and resultant membrane strain in the cell strain device. Membrane strain was calculated from changes in distance between pairs of ink marks printed on the deformable membrane. Each data point represents mean ± SD strain value of 12 ink-marked pairs (6 separate pairs from each experiment). Applied pressures of 20 kPa (150 mmHg; arrow at left) and 40 kPa (300 mmHg; arrow at right) were used in all subsequent experiments.

Cell surface area increases as a result of membrane strain. To determine whether cell surface area increased after membrane deformation, the PMs of cells were labeled with DiI and imaged by LCFM in the undeformed and deformed states. Five separate cells were then analyzed after either 7 or 25% strain. As a result of membrane deformation, cells were found to flatten and elongate, with the greatest change in cell cross-sectional area occurring in the basalmost portions of the cells (Fig. 3A). Figure 1 (top) shows a single LCFM image of cells labeled with BODIPY and qualitatively demonstrates typical observed changes in basal cell cross-sectional area after deformation. The application of 7 or 25% membrane strain resulted in an average increase (P < 0.01) in cell surface area of 18 ± 6 or 35 ± 9%, respectively (Fig. 3B). Thus changes in cell surface area were strain amplitude dependent.


View larger version (17K):
[in this window]
[in a new window]
 
Fig. 3.   Deformation-induced changes in cell surface area. A: representative cross-sectional area-height graph of a single cell. Abscissa, 1-µm LCFM image slices taken from the base to the apex of the cell. After deformation, cells flattened and elongated, with the most pronounced increase in cross-sectional area occurring in the basalmost portions of the cell. B: each data point is the change in cell surface area of a single cell from the undeformed to the deformed state. Strain increased cell surface area in an amplitude-dependent fashion with cells exposed to a 7 or 25% strain, increasing their surface area by 18 ± 6 and 35 ± 9%, respectively. P < 0.01.

Cell strain reduces PM lipid label concentration. In these experiments, BODIPY-LacCer was used to label the PM. The concentration of BODIPY label within the PM can be quantitated by the red-to-green fluorescence emission ratio (R/G). With decreasing molar concentrations in lipid membranes, BODIPY lipids exhibit a concentration-dependent spectral shift in fluorescent wavelength emission (from red to green) (21). The R/G of each pixel within the PM and an average value of the entire PM were measured for cells in single-image slices in the undeformed and deformed states. The effect of 25% stretch on the R/G of the PM is illustrated in the representative frequency distribution of all PM pixels in a single cross-sectional image of the cell (Fig. 4A). A leftward shift in the R/G distribution was apparent after 25% strain, representing a decrease in the molar density of labeled lipid in the PM. The photo at the top of Fig. 1 demonstrates this qualitatively. The average decrease in the R/G of the PM of cells stretched by 25% compared with their prestretched state in five separate experiments (15 individual cells) was 59%. Compared with the unstretched time-control cell population, stretch resulted in a 3.2 times greater decrease (P < 0.002) in the average R/G of the PM (Fig. 4B); this decrease suggested trafficking of unlabeled lipid to the PM.


View larger version (18K):
[in this window]
[in a new window]
 
Fig. 4.   Deformation-induced changes in plasma membrane (PM) BODIPY concentration. A: representative frequency distribution graph of BODIPY-lactosylceramide (LacCer) concentration [red-to-green ratio (R/G)] in the PM of a single cell before and after 25% strain. Each data point is the percentage of PM pixels with a given fluorescent R/G. There was a shift to the left in the R/G of the cell PM after deformation, representing a decrease in the molar density of lipid label within the PM. B: each data point is the mean percentage decrease in PM BODIPY concentration of a single cell compared with itself over time alone (time-control cells) or after 25% strain. *Cells strained by 25% demonstrated a 3.2 times greater decrease in PM label concentration compared with time-control cells (n = 5), P < 0.002.

Substratum deformation induces lipid trafficking to the plasma membrane in a strain amplitude- and energy-dependent fashion. To directly measure whether deformation resulted in trafficking of lipid molecules, we fluorescently labeled intracellular lipid membranes with the styryl lipid label FM1-43 (Fig. 5A). FM1-43 is an amphiphilic molecule that is characterized by an ~50-fold loss in fluorescence when transferred from the membrane bilayer to the aqueous phase (4). Thus the change in whole cell fluorescence intensity after stretch served as a measure of lipid trafficking to the PM. The leftward shift in the frequency distribution of pixel intensity for a representative single-image slice of a cell labeled with FM1-43 after 25% strain is shown in Fig. 5B. In three separate experiments, cells labeled with FM1-43 were deformed by either 7 or 25% compared with unstretched time-control cells (15 separate cells in each group). Deformation resulted in lipid transport to the PM, and a DILT response was determined in which 25% strain resulted in a twofold greater (P < 0.001) trafficking response than that in cells with 7% strain (33 ± 12.3 vs. 16 ± 7.7%) and a sevenfold greater (P < 0.001) trafficking response than that in the time-control cells (33 ± 12.3 vs. 4.9 ± 3.5%; Fig. 5C). After deformation, cells were found to exclude trypan blue and could be recultured with normal morphology and growth characteristics over 48-120 h (data not shown). In two further experiments, cells were ATP depleted by incubation with sodium azide and 2-deoxy-D-glucose. The DILT response was not apparent in these ATP-depleted cells compared with that in the time-control cell population (Fig. 5D), suggesting that DILT is an energy-dependent process.


View larger version (38K):
[in this window]
[in a new window]
 
Fig. 5.   Deformation-induced lipid trafficking (DILT) to the PM. A: representative confocal image of A549 cells labeled with FM1-43 over 4 h at 37°C showing numerous labeled intracellular vesicles in all cells. B: representative frequency distribution graph of the change in FM1-43 fluorescence (FFM1-43) of a single confocal image from a single cell before deformation and during 25% strain. There was a shift to the left in the graph after deformation, representing a decrease in total fluorescent intensity within the cell and thus exocytosis of FM1-43. C: mean percent decrease in FFM1-43 intensity compared with itself over time alone (time control) or after 7 or 25% strain. Bars, means ± SD of all the cells within each group; n = 3 separate experiments. *P < 0.001 vs. time-control cell population. **P < 0.001 vs. 7% substratum strain. D: DILT is an energy-dependent process that is abrogated in the presence of ATP-depleting medium. Each data point represents the mean percent decrease in FFM1-43 intensity in the presence of azide and 2-deoxy-D-glucose compared with itself over time alone (time control) or after 25% strain. There was no significant difference between time-control and strained cells, P = 0.5.

To provide further evidence that FM1-43 was indeed exocytosed as a result of substratum deformation, samples of supernatant from undeformed and 25% strained culture wells were analyzed by fluorescence spectrofluorometry (two separate experiments; six wells each from undeformed and 25% strained cells). The supernatant from deformed culture wells contained on average 13% more (P = 0.007) detectable FFM1-43 compared with supernatant from undeformed culture wells.

Cell stretch induces a small number of PM breaks. After a 10-min incubation at 37°C with FITC-Dx, the distribution of intracellular fluorescence was found to be diffuse and homogeneous (representative of PM breaks) or granular and heterogeneous (representative of noninjurious endocytosis) (19). Cells were stretched by the same parameters as in the previous experiments, and FITC-Dx was found to be homogeneously distributed throughout the cell (Fig. 6A) in only a small percentage (0.6-2.7%) of the cells overlying the middle of the loading post (Fig. 6B). In five separate experiments at 25% strain, 2.8% of cells demonstrated PM injury in the presence of 70-kDa FITC-Dx. This was 4.9 times greater (P < 0.02) than that in cells exposed to a 7% strain (Fig. 6B). This strain amplitude dependence of PM injury was apparent after 25% strain for all Dx sizes (P < 0.003). Seven percent strain did not result in a significantly greater cell injury (P > 0.05) when compared with that in unstrained cells. The degree of PM injury was not dependent (P = 0.2) on the size of Dx studied.


View larger version (44K):
[in this window]
[in a new window]
 
Fig. 6.   Deformation-induced PM breaks. A: representative LCFM image of an epithelial cell strained by 25% demonstrating homogeneous uptake of 70-kDa FITC-dextran (Dx) in a single cell, thus identifying breaks in its PM. Surrounding cells showed only their normal autofluorescence pattern, suggesting no sustained PM breaks. B: percentage of cells with PM breaks with the use of different molecular weight FITC-Dx after no strain, 7% strain, or 25% strain. Only 25% strain (*) resulted in a significantly greater number of cells with PM breaks when compared with no strain (F = 21.19; *P < 0.001) and also resulted in more breaks when compared with 7% strain (**P < 0.05). These results were not dependent on the size of FITC-Dx that was used (F = 1.74; P = 0.18).

We also wished to determine whether the strain-induced breaks were lethal to the cells and whether they were reversible in nature. First, cells were recultured after deformation experiments, and those that had taken up FITC-Dx were found to be adherent to the culture plate and to have normal morphology 36-72 h after reculturing. This suggests that the PM injury was nonlethal. Second, to determine whether the breaks were reversible, cells were incubated with FITC-Dx immediately after the stretch was released (n = 4 cells). The number of cells taking up FITC-Dx in a homogeneous fashion was no different (P > 0.05) when compared with uptake in unstrained cells, suggesting that the induced PM breaks were reversible.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

We report a heretofore undescribed response of alveolar epithelial cells to deformation, namely, increased lipid transport to the PM. This DILT response was rapid in onset (observed within 60-90 s of substratum strain) and vesicular in nature and may represent an important regulatory mechanism for membrane transport in alveolar epithelial cells. Cell deformation induced by substratum strain resulted in significant increases in overall cell surface area. However, in spite of the significant changes in cell shape, the proportion of cells sustaining PM breaks was small. These breaks were nonlethal and reversible in nature, suggesting that at least one important role of DILT is reparative, site-directed lipid transport. However, DILT was observed in all cells, not just those sustaining PM breaks, suggesting that DILT might also represent a cytoprotective response against PM stress failure.

The present study was performed with an in vitro alveolar epithelial cell system and was established to achieve two goals: 1) to simulate in vivo basement membrane strain that occurs during mechanical ventilation-induced alveolar distension and 2) to allow LCFM imaging of cells in the undeformed and deformed states. In achieving these goals, we were able to reduce complex in vivo conditions to a meaningful experimental setup in which direct biological consequences of definable mechanical strains could be accurately measured. We chose to study the effects of a maximal substratum strain value of 25%. The relevance of this strain amplitude in the setting of VILI has been addressed previously (33). Briefly, despite numerous studies (3, 20, 32) using different measurement techniques, in situ strain of alveolar epithelial cells has never been precisely quantified. Most evidence suggests that the alveolus deforms little during quiet breathing. However, there is reason to think that injurious strains are imposed on the lung parenchyma during mechanical ventilation with high tidal volumes (10). Using the parenchymal marker technique, Rodarte and colleagues (25) measured the strains of approx 1-cm3 lung regions in intact dogs and reported values as high as 40% during inspiratory capacity maneuvers (25). Although these estimates cannot be directly extrapolated to basement membrane or alveolar wall strain, they nevertheless place the 25% substratum strain used in our present study in a biologically relevant context.

Type I epithelial cells cover >90% of the alveolar surface and are believed to be exposed to a greater strain than type II cells, which are cuboidal in shape, reside in alveolar corners, and might be sheltered from deforming stresses (32). Type I cells might therefore be a more relevant substrate for deformation-induced lung injury research. Unfortunately, primary type I cell culture systems have been incompletely characterized to date and are not readily available (22). Thus A549 cells were used as a representative alveolar epithelial cell model. This epithelial cell line exhibits intercellular adhesion molecule-1 surface receptor expression (a characteristic of the type I phenotype) (22, 36) and possesses functioning proinflammatory signaling pathways (33). Because A549 cells do not form polarized monolayers and might not communicate with each other through gap junctions, the biological relevance of our findings might be questioned. However, the DILT responses of primary rat type II alveolar epithelial cells grown for 5 days in culture were virtually identical to those of the A549 cells described in Fig. 5 (Vlahakis and Hubmayr, unpublished observations). This finding supports the notion that A549 cells serve as a representative model system for our experimental questions.

By taking advantage of the unique fluorescent characteristics of BODIPY-labeled lipid analogs and of the lipophilic dye FM1-43, we were able to demonstrate that deformation induces lipid transport to the PM. To control for the effects of temperature, time, and light exposure on the membrane lipids of the cell, we compared the strained cell population with cell populations treated in an identical fashion except for deformation (time-control cells). Using BODIPY labeling of the PM, we showed that deformation induced a decrease in the molar concentration of lipid label within the PM. This finding suggested that unlabeled endogenous lipids were transported to the PM, thereby reducing the molar density of the fluorescent lipid in the membrane. Strain-induced unfolding of cell surface ruffles would not have affected the BODIPY fluorescence spectrum because unfolding does not alter the intermolecular distance of BODIPY label within the plane of the lipid bilayer (9). However, the BODIPY data could also be interpreted as a selective loss of fluorescent label from a physically strained PM. Therefore, to specifically demonstrate lipid transport to the PM in association with cell deformation, we used the lipophilic dye FM1-43. FM1-43 is known to label intracellular vesicles, which "recycle" to the PM, and has been used extensively as a means to study exocytosis (4). Epithelial cell deformation resulted in exocytosis of these labeled vesicles, and this process occurred in a strain amplitude-dependent fashion. Thus these results provided direct confirmation that cell deformation induces lipid transport to the PM.

Substratum deformation was also found to increase epithelial cell surface area. This increase in surface area provides further evidence for DILT and suggests that the cell might grow PM to accommodate increases in surface area. Based on lipid tether force-displacement data, it has been argued that fibroblasts and neuronal cells maintain a PM "reservoir" consisting of membrane ruffles that unfold to accommodate increases in cell surface area (24). Unfortunately, the confocal microscope is unable to resolve subtle PM invaginations and ruffles that are readily apparent on electron micrographs of A549 cells (images not shown). As a result, our reported surface area values are an "apparent" measure of the true cell surface areas, and their changes with deformation do not reflect the relative contributions of PM unfolding and lipid transport to the PM. Nonetheless, these findings do support the concept of a membrane reservoir and suggest that alveolar epithelial cells use intracellular lipid vesicles as a result of basement membrane deformation.

In contrast to DILT, which occurred in all deformed cells, we found that substratum deformation resulted in only a small percentage of cells with PM breaks, even at the greatest strain (25%). This was surprising in light of the large increases in cell surface area that resulted from this deformation. We believe that this finding lends further support to the adaptive role of DILT in the accommodation of cell surface area. In addition, the reversible and nonlethal nature of these breaks further speaks to the cytoprotective role of DILT and provides evidence for the first time that rapid transport of vesicles to the site of PM injury is a deformation-induced repair process in alveolar epithelial cells. The varying molecular size of FITC-Dx and its homogeneous pattern of cellular uptake after cell stress has been used by investigators as a marker of PM breaks or "porosity" (19). It is important to note, however, that a homogeneous pattern of FITC-Dx uptake depends on the nature of the induced PM breaks. If the breaks are small in dimension, large in number, and open for a short period of time, the cell might still take up low molecular mass molecules, although not in a homogeneous fashion. This raises an important issue with regard to the validity of techniques that use the cellular permeability of fluorescent and low molecular mass (<1 kDa) molecules (such as ethidium homodimer and propidium iodide) to determine cell death (31). Our results suggest that in experiments where cells have been physically stressed, these molecules might indeed be markers of PM disruptions rather than cell death.

Although the percentage of cells with breaks was found to be small, the potential biological relevance for the lung might still be great. Even injury of a single cell can be a sufficient trigger for the initiation of tissue inflammation. Gap junction and local paracrine communication between epithelial cells would potentially allow PM injury of a single cell to be magnified and lung inflammation to be potentiated (27). Both alveolar overdistension in vivo (30) and alveolar epithelial cell deformation in vitro result in the production of numerous proteins including interleukin-8, one of the key inflammatory molecules associated with VILI (23, 33). As these protein molecules are "chaperoned" by associated lipid vesicles to the cell surface, PM surface area might increase as a result. Thus DILT might simply be a secondary phenomenon of protein expression or secretion by the cell. This point is underscored when specifically considering the alveolar type II cell in which surfactant exocytotic events occur regularly. Indeed, studies (2, 35) have shown that cell deformation stimulates type II rat alveolar epithelial cells to release surfactant. Therefore, the findings in our system that recycling vesicles are transported to the PM as a result of deformation speak to a more unique underlying relevance for DILT in the alveolar epithelium other than that of surfactant secretion alone in type II cells.

Our data using FFM1-43 support the notion that lipid transport to the PM results from vesicular transport. The rapidity of this response suggests that the cell might either store vesicles in the immediate sub-PM region or that vesicles early in other trafficking pathways might preferentially be targeted to the PM. Studies in which cells were deformed in the presence of ATP-depleting media demonstrated that lipid trafficking was abrogated, suggesting that DILT is an energy-dependent process. The upstream transducer(s) of DILT remains to be defined, in particular, the role of the interaction of extracellular matrix (alveolar basement membrane) and integrins (of the alveolar epithelial cell). It is likely that the cytoskeleton (5, 34) and focal adhesion complexes (8) are essential for transduction of these mechanical forces. Sheetz and Dai (26) also found, in fibroblasts, that PM tension might play an important role in transducing cell morphology, motility, and membrane trafficking within the cell. In addition, they elegantly showed that cells maintain a PM reservoir that serves to buffer mechanical stress-induced changes in PM tension. This reservoir increases with cytoskeletal disruption or increases in PM tension (24). Although we did not measure PM tension, our results do support the concept of a PM reservoir in alveolar epithelial cells and provide direct evidence that, in part, intracellular vesicles make up this reservoir and might be utilized in the setting of mechanical deformation.

In conclusion, this study provides novel in vitro evidence that basement membrane deformation induces a rapid, strain amplitude-dependent transport of lipids to the PM of alveolar epithelial cells. This DILT response serves, in part, to repair PM breaks in order to maintain PM integrity and cell viability. At the same time, DILT can be viewed as a cytoprotective mechanism against PM stress failure and represents an attractive target for pharmacotherapy of VILI. The recently concluded Acute Respiratory Distress Syndrome Network Trial (1) conclusively showed that smaller ventilator-delivered tidal volumes improve patient survival. In a number of animal models, mechanical ventilation with high tidal volumes has been shown to raise vascular permeability and promote lung inflammation. PM stress failure of alveolar lining cells is a plausible mechanotransduction event underlying these cardinal manifestations of VILI. The inability of epithelial and endothelial cells to accommodate large basement membrane strains can lead to cell injury that manifests as gap formation within and across the alveolar-capillary barrier. The consequent leakage of serum into the alveolar space impairs surfactant function and alters alveolar micromechanics. The accumulation of serum proteins can trigger phagocytosis and proinflammatory signaling. Another potential inflammatory trigger is the calcium influx through transient endothelial and epithelial PM wounds (29). Wounding-induced cell activation has been extensively studied in other systems. It is one of many possible mechanisms through which cell deformation can initiate nuclear factor-kappa B activation, proinflammatory gene transcription, and cytokine release (15, 19). Thus DILT represents a newly described regulatory mechanism for membrane traffic in alveolar epithelium and provides a novel biological framework within which to consider alveolar deformation injury and repair.


    ACKNOWLEDGEMENTS

We thank Dr. G. C. Sieck and Dr. Y. S. Prakash for help with the confocal microscopy, Jon J. Camp for assistance with image analysis, and L. L. Oeltjenbruns for preparing the manuscript.


    FOOTNOTES

This work was supported by National Institutes of Health Grants R01-HL-63178 and GM-22942, a Glaxo Wellcome Pulmonary Fellowship Research grant, and institutional support from the Clinician Investigator Program and the Thoracic Diseases Research Unit (Mayo Clinic).

Address for reprint requests and other correspondence: N. E. Vlahakis, Rm. 4-411 Alfred Bldg., Mayo Clinic, 200 First St. SW, Rochester, MN 55905 (E-mail: vlahakis.nicholas{at}mayo.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.

Received 27 July 2000; accepted in final form 30 November 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Acute Respiratory Distress Syndrome Network. Ventilation with lower tidal volumes compared with traditional tidal volumes for acute lung injury and the acute respiratory distress syndrome. N Engl J Med 342: 1301-1308, 2000[Abstract/Free Full Text].

2.   Ashino, Y, Ying X, Dobbs LG, and Bhattacharya J. [Ca2+]i oscillations regulate type II cell exocytosis in the pulmonary alveolus. Am J Physiol Lung Cell Mol Physiol 279: L5-L13, 2000[Abstract/Free Full Text].

3.   Bachofen, H, Schurch S, Urbinelli M, and Weibel ER. Relations among alveolar surface tension, surface area, volume, and recoil pressure. J Appl Physiol 62: 1878-1887, 1987[Abstract/Free Full Text].

4.   Betz, WJ, Mao F, and Bewick GS. Activity-dependent fluorescent staining and destaining of living verterbrate motor nerve terminals. J Neurosci 12: 363-375, 1992[Abstract].

5.   Bi, G-Q, Morris RL, Liao G, Alderton JM, Scholey JM, and Steinhardt RA. Kinesin- and myosin-driven steps of vesicle recruitment for Ca2+-regulated exocytosis. J Cell Biol 138: 999-1008, 1997[Abstract/Free Full Text].

6.   Chen, C-S, Martin O, and Pagano RE. Changes in the spectral properties of a plasma membrane lipid analog during the first seconds of endocytosis in living cells. Biophys J 72: 37-50, 1997[Abstract].

7.   Chen, H, Swedlow JR, Grote M, Sedat JW, and Agard DA. The collection, processing, and display of digital three-dimensional images of biological specimens. In: Handbook of Biological Confocal Microscopy (2nd ed.), edited by Pawley JB.. New York: Plenum, 1995, p. 201-203.

8.   Chicurel, ME, Singer RH, Meyer CJ, and Ingber DE. Integrin binding and mechanical tension induce movement of mRNA and ribosomes to focal adhesions. Nature 392: 730-733, 1998[ISI][Medline].

9.   Dahim, M, Mizuno NK, and Brockman HL. A free volume model for BODIPY fluorescence in monolayers (Abstract). Biophys J 78: 180A, 2000[ISI].

10.   Dreyfuss, D, and Saumon G. Ventilator-induced lung injury: lessons from experimental studies. Am J Respir Crit Care Med 157: 294-323, 1998[Free Full Text].

11.   Dreyfuss, D, Soler P, Basset G, and Saumon G. High inflation pressure pulmonary edema: respective effects of high airway pressure, high tidal volume and positive end-expiratory pressure. Am Rev Respir Dis 137: 1159-1164, 1988[ISI][Medline].

12.   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[Abstract/Free Full Text].

13.   Gattinoni, L, D'Andrea L, Pelosi P, Vitale G, Pesenti A, and Fumagalli R. Regional effects and mechanism of positive end-expiratory pressure in early adult respiratory distress syndrome. JAMA 269: 2122-2127, 1993[Abstract].

14.   Gil, J, Bachofen H, Gehr P, and Weibel ER. Alveolar volume-surface area relation in air- and saline-filled lungs fixed by vascular perfusion. J Appl Physiol 47: 990-1001, 1979[Abstract/Free Full Text].

15.   Grembowicz, KP, Sprague D, and McNeil PL. Temporary disruption of the plasma membrane is required for c-fos expression in response to mechanical stress. Mol Biol Cell 10: 1247-1257, 1999[Abstract/Free Full Text].

16.   Hanson, DP, Robb RA, Aharon S, Augustine KE, Cameron BM, Camp JJ, Karwoski RA, Larson AG, Stacy MC, and Workman EL. New software toolkits for comprehensive visualization and analysis of three-dimensional multimodal biomedical images. JAMA 10: 229-230, 1997.

17.   International Consensus Conference in Intensive Care Medicine. Ventilator-associated lung injury in ARDS. Am J Respir Crit Care Med 160: 2118-2124, 1999[Free Full Text].

18.   Lecuona, E, Saldías F, Comellas A, Ridge K, Guerrero C, and Sznajder JI. Ventilator-associated lung injury decreases lung ability to clear edema in rats. Am J Respir Crit Care Med 159: 603-609, 1999[Abstract/Free Full Text].

19.   McNeil, PL, and Steinhardt RA. Loss, restoration, and maintenance of plasma membrane integrity. J Cell Biol 137: 1-4, 1997[Free Full Text].

20.   Mercer, RR, Laco JM, and Crapo JD. Three-dimensional reconstruction of alveoli in the rat lung for pressure-volume relationships. J Appl Physiol 62: 1480-1487, 1987[Abstract/Free Full Text].

21.   Pagano, RE, Martin OC, Kang HC, and Haugland RP. A novel fluorescent ceramide analog for studying membrane traffic in animal cells: accumulation at the Golgi apparatus results in altered spectral properties of the sphingolipid precursor. J Cell Biol 113: 1267-1279, 1991[Abstract].

22.   Paine, R, III, and Simon RH. Expanding the frontiers of lung biology through the creative use of alveolar epithelial cells in culture. Am J Physiol Lung Cell Mol Physiol 270: L484-L486, 1996[Free Full Text].

23.   Pugin, J, Dunn I, Jolliet P, Tassaux D, Magnenat J-L, Nicod LP, and Chevrolet J-C. Activation of human macrophages by mechanical ventilation in vitro. Am J Physiol Lung Cell Mol Physiol 275: L1040-L1050, 1998[Abstract/Free Full Text].

24.   Raucher, D, and Sheetz MP. Characteristics of a membrane reservoir buffering membrane tension. Biophys J 77: 1992-2002, 1999[Abstract/Free Full Text].

25.   Rodarte, JF, Hubmayr RD, Stamenovic D, and Walters BJ. Regional lung strain in dogs during deflation from total lung capacity. J Appl Physiol 58: 164-172, 1985[Abstract/Free Full Text].

26.   Sheetz, MP, and Dai J. Modulation of membrane dynamics and cell motility by membrane tension. Trends Cell Biol 6: 85-89, 1996[ISI].

27.   Simon, RH, and Paine R, III. Participation of alveolar epithelial cells in lung inflammation. J Lab Clin Med 126: 108-118, 1995[ISI][Medline].

28.   Slutsky, AS. Mechanical ventilation. American College of Chest Physicians' Consensus Conference. Chest 104: 1833-1859, 1993[ISI][Medline].

29.   Terasaki, M, Miyake K, and McNeil PL. Large plasma membrane disruptions are rapidly resealed by Ca2+-dependent vesicle-vesicle fusion events. J Cell Biol 139: 63-74, 1997[Abstract/Free Full Text].

30.   Tremblay, L, Valenza F, Ribeiro SP, Li J, and Slutsky AS. Injurious ventilatory strategies increase cytokines and c-fos mRNA expression in an isolated rat lung model. J Clin Invest 99: 944-952, 1997[Abstract/Free Full Text].

31.   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[Abstract/Free Full Text].

32.   Tschumperlin, DJ, and Margulies SS. Alveolar epithelial surface area-volume relationship in isolated rat lungs. J Appl Physiol 86: 2026-2033, 1999[Abstract/Free Full Text].

33.   Vlahakis, NE, Schroeder MA, Limper AH, and Hubmayr RD. Stretch induces cytokine release by alveolar epithelial cells in vitro. Am J Physiol Lung Cell Mol Physiol 277: L167-L173, 1999[Abstract/Free Full Text].

34.   Wang, N, Butler JP, and Ingber DE. Mechanotransduction across the cell surface and through the cytoskeleton. Science 260: 1124-1127, 1993[ISI][Medline].

35.   Wirtz, HR, and Dobbs LG. Calcium mobilization and exocytosis after one mechanical stretch of lung epithelial cells. Science 250: 1266-1269, 1990[ISI][Medline].

36.   Yu, ML, and Limper AH. Pneumocystis carinii induces ICAM-1 expression in lung epithelial cells through a TNF-alpha -mediated mechanism. Am J Physiol Lung Cell Mol Physiol 273: L1103-L1111, 1997[Abstract/Free Full Text].


Am J Physiol Lung Cell Mol Physiol 280(5):L938-L946
1040-0605/01 $5.00 Copyright © 2001 the American Physiological Society