Ca2+ entry is essential for cell strain-induced lamellar body fusion in isolated rat type II pneumocytes

Manfred Frick,1 Cristina Bertocchi,1 Paul Jennings,1 Thomas Haller,1 Norbert Mair,1 Wolfgang Singer,2 Walter Pfaller,1 Monika Ritsch-Marte,2 and Paul Dietl1

Departments of 1Physiology and 2Medical Physics, University of Innsbruck, A-6020 Innsbruck, Austria

Submitted 15 September 2003 ; accepted in final form 21 September 2003


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Using a new equibiaxial strain device, we investigated strain-induced Ca2+ signals and their relation to lamellar body (LB) exocytosis in single rat alveolar type II (AT II) cells. The strain device allows observation of single cells while inducing strain to the entire substratum. AT II cells tolerated high strain amplitudes up to 45% increase in cell surface area ({Delta}CSA) without release of lactate dehydrogenase or ATP. Strain exceeding a threshold of ~8% {Delta}CSA resulted in a transient rise of the cytoplasmic Ca2+ concentration in some cells. Higher strain levels increased the fraction of Ca2+-responding cells. The occurrence of strain-induced Ca2+ signals depended on cell-cell contacts, because lone cells (i.e., cells without cell-cell contacts) did not exhibit Ca2+ signals. Above threshold, the amplitude of the Ca2+ signal as well as the number of stimulated LB fusions correlated well with the amplitude of strain. Furthermore, stimulated LB fusions occurred only in cells exhibiting a Ca2+ signal; 50 µM Gd3+ in the bath affected neither Ca2+ signals nor fusions. Intracellular Ca2+ release was triggered at higher strain amplitudes and inhibited by thapsigargin. Removal of bath Ca2+ completely inhibited Ca2+ signals and fusions. We conclude that strain of AT II cells stimulates a Ca2+ entry pathway that is highly sensitive to strain and a prerequisite for subsequent Ca2+ release. Both mechanisms result in a graded response of fusions to strain. Our data also allow us to introduce the term "effective strain" as the physiologically relevant portion of the strain amplitude.

surfactant secretion; alveolar type II cells; mechanical strain; stretch


THE MAIN FUNCTION of alveolar type II (AT II) cells is the synthesis and secretion of surfactant, a lipid-rich, lipoprotein-like material that is essential for respiration by reducing the surface tension of the air-liquid interface in the alveoli (reviewed in Refs. 4, 10, 27). Surfactant is stored in vesicles termed lamellar bodies (LBs) and secreted into the alveolar lumen by exocytosis (8, 13, 18).

A number of hormones, pharmacological agents, and physicochemical factors have been reported to stimulate or regulate surfactant secretion in AT II cells (reviewed in Refs. 3, 14, 19, 28, 39). Among them, strain of AT II cells, which occurs as a result of a deep breath (such as a yawn or a sigh or during exercise), is probably the most potent stimulus for surfactant secretion in vivo (7, 22, 23, 3638). Moreover, Wirtz and Dobbs (37) demonstrated that a single stretch of isolated AT II cells in vitro is sufficient to cause a transient rise in cytosolic Ca2+ concentration ([Ca2+]c) followed by sustained surfactant secretion. This is consistent with many additional lines of evidence that Ca2+ is the major second messenger for LB exocytosis (5, 8, 11, 24).

After fusion of a secretory vesicle with the plasma membrane, vesicle contents are released through the exocytotic fusion pore before being dispersed in the extracellular space. Using the surfactant-staining properties of the lipophilic dye FM 1-43 (13, 18), we previously demonstrated (12, 30) that this release process is very slow in AT II cells and that surfactant does not readily disperse in the extracellular space. One way that a single strain of AT II cells stimulates the secretion of surfactant is via expansion of fusion pores, accelerating the process of surfactant release (12, 30). It was the aim of this study to examine LB fusion responses to strain on the level of single cells under continuous observation before, during, and after strain and to relate these effects to changes of [Ca2+]c. To meet this objective, we designed a new equibiaxial strain system that permits continuous observation of single cells while inducing equibiaxial strain of variable strength and frequency. A major implication of our experiments is that Ca2+ entry, in addition to previous concepts considering intracellular Ca2+ release as the major strain-related event (37), plays an essential role in Ca2+ signaling and LB fusions.


    METHODS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cell culture. AT II cells were isolated from male Sprague-Dawley rats (180–200 g) according to the procedure of Dobbs et al. (6). Isolated cells were plated at low density (~150 cells/mm2) on a 120-µm-thick silicone membrane (Silastic, gloss/gloss finish; Specialty Manufacturing, Saginaw, MI). Membranes were thin and optically clear enough to ensure detailed images of single cells without appreciable distortion compared with glass. Cells were incubated in DMEM supplemented with 24 mM NaHCO3 in a humidified 5% CO2 atmosphere at 37°C for 28–48 h.

Description of strain device. The strain device was designed to enable a continuous observation of single cells while inducing equibiaxial strain to the entire substratum. The principle of this device is similar to previously described methods (15, 16, 29, 31, 32), although improved in several aspects to allow continuous imaging of the same cells throughout the experiment. Any cell of interest can be placed in the area of observation, irrespective of its location on the substratum, and remain there during strain application. Strain application was combined with real-time fluorescence microscopy. The strain device is schematically illustrated in Fig. 1.



View larger version (44K):
[in this window]
[in a new window]
 
Fig. 1. Cell strain device. Three-dimensional construction (top) and schematic drawing (bottom) of the strain device (colors correspond to 3-dimensional drawing). Cells are grown on an elastic, optically clear, inert silicone membrane (memb), which is fixed in a clamping device (clamp), and placed on the stage of an inverted microscope (st). The actual strain device consists of two parts, aligned around the optical path of the microscope (dotted line). The lower part (supp) consists of 2 bearing rings separated by an indention groove (i.g.) and serves as support for the membrane. The upper part consists of 2 cylinders that can be moved independently in the z-direction. The outer cylinder (o.c.) is lowered to clamp the membrane tightly between the cylinder and the outer bearing ring. Thus the cells are fixed in the central portion of the clamped membrane and aligned to the optical path of the objective (obj). Thereafter, lowering the inner cylinder (i.c.) (into the indention groove) by twisting the screw (scr) results in actual strain of the clamped membrane. This is done either manually or by a computer-controlled step motor. Green arrows indicate twisting of screw and resulting movement of the inner cylinder. Red arrow indicates cell strain due to membrane deformation.

 

Cells are grown on an elastic, optically clear, inert silicone membrane that is fixed in a custom-made clamping device and placed on the stage of an inverted microscope (Axiovert 135 TV; Zeiss). The actual strain device consists of two parts, which are aligned to the optical path of the microscope. The lower part consists of two bearing rings separated by an indention groove. Both rings serve as support for the silicone membrane and are lubricated to reduce friction. The upper part consists of two cylinders that can be moved independently in the z-direction. For strain application, the cells of interest are placed in the visual field of the microscope. The outer cylinder is then lowered until the silicone membrane is tightly clamped between the cylinder and the outer bearing ring. After the cells are fixed in the center portion of the clamped membrane, lowering the inner cylinder into the indention groove results in actual straining of the clamped membrane. This is done either manually or by a computer-controlled step motor.

Neither lateral shifting of the cells, because they are positioned in the center of the strain field, nor out-of-focus problems occur, because cells remain in the same plane during application of strain. As a result, cells can be observed by real-time imaging at high spatial and temporal resolution at various strain levels and/or frequencies.

Validation of membrane deformation during strain. To test for uniformity of membrane strain, we followed the method described by Tschumperlin and Margulies (32). Briefly, membranes were marked with an approximate center dot and three dots along four radial spokes (Fig. 2C). After insertion into the strain device, the positions of the individual dots at five different strain levels (due to displacement of the inner cylinder) were captured by a charge-coupled device camera (Fig. 2C). Distances between dot locations were analyzed with image analysis software (T.I.L.L. Photonics), and the data were used to calculate circumferential ({epsilon}{theta}) and radial ({epsilon}r) strains ({epsilon}{theta} = u/r, {epsilon}r = {partial}u/{partial}r, where r is the radial position in the relaxed state and u is the radial displacement in the strained state; see Fig. 2). If {epsilon}{theta} = {epsilon}r, the strain field is uniform (equibiaxial), indicating equal strains in all directions. Therefore, we compared the {epsilon}r and {epsilon}{theta} values at different strain levels. The dependence of average {epsilon}{theta} amplitude on r and average {epsilon}r amplitude on {theta} (radial direction) were measured at every indention of the inner cylinder and calculated over five membranes.



View larger version (43K):
[in this window]
[in a new window]
 
Fig. 2. A: circumferential and radial membrane strains. To demonstrate the uniformity of membrane deformation, circumferential strains and radial strains of 5 membranes (means ± SD) were compared at identical indention depth (5.2 mm). Left: circumferential strain at 3 radial positions (corresponding to marks on the membrane in C). Results indicate that strain was independent of the radial position. Right: radial strain in 4 directions, indicating that strain was independent of orientation. The circumferential and radial strains do not differ; therefore, the strain field is equal in all directions (equibiaxial). B: increase in membrane surface area ({Delta}MSA) as a function of ring indention. {Delta}MSA was computed at different ring indentions by using 5 membranes (means ± SD) to characterize the range of operation. C: photograph of membrane in relaxed (left) and strained (right) configurations. Marks seen on the membrane were used for the qualitative assessment of strain (r, radius; u, radial displacement).

 

Membrane deformations were computed according to Tschumperlin and Margulies (32). The stretch ratio ({lambda}) in an equibiaxial strain field is defined as {epsilon}{theta} = {epsilon}r = {lambda} - 1 and can be computed according to {lambda} = (r + u)/r. We calculated the average {lambda} for every radial position at every strain level by averaging the {lambda} values of the same radial position across all four spokes. The membrane surface area (MSA) represents the area of the upper membrane surface on which the cells were grown. Changes in membrane surface area (%{Delta}MSA) were determined by the method described in detail by Tschumperlin and Margulies (32) and were calculated by % {Delta}MSA = ({lambda}2 - 1) x 100. The cell surface area (CSA) is defined as the area within the margin of a single cell; its measurement is described in Determination of cellular strain.

Determination of cellular strain. Cells loaded with fura 2-AM were washed and placed in the strain device. With a two-dimensional imaging system (T.I.L.L. Photonics), images were acquired with a 20x Plan Neofluar (Zeiss) at a rate of 0.5 Hz at each excitation wavelength (100-ms excitation at 340 and 380 nm). CSA was calculated in the 340-nm excitation images. Because the cells did not shift laterally or vertically out of the observation area, CSA could be easily measured in the relaxed state as well as at all strain levels (see Fig. 4). Changes in cell surface area (%{Delta}CSA) were expressed according to %{Delta}CSA = [(CSAd - CSAu)/CSAu] x 100, where CSAu is CSA in the relaxed state and CSAd is CSA in the strained state. The strain rate was expressed as %{Delta}CSA per second.



View larger version (44K):
[in this window]
[in a new window]
 
Fig. 4. Dependence of cell response on strain amplitude. A: top from left to right: fura 2-labeled alveolar type II (AT II) cells (viewed at 340-nm excitation) at the indicated {Delta}CSA (%) (red lines indicate cell borders). Bottom: corresponding fura 2 ratio images, demonstrating distinct cytosolic Ca2+ concentration ([Ca2+]c) rises in individual AT II cells. Note that a lone cell (1 is seen in the upper right corner) did not respond to mechanical strain. B: % of cells showing a Ca2+ signal in response to mechanical strain as a function of increased strain amplitudes under various experimental conditions. Strain levels <10% {Delta}CSA only rarely produced a [Ca2+]c rise (threshold ~8% {Delta}CSA). Thapsigargin, preincubation (20–30 min) with 100 nM thapsigargin; calcium-free, lack of bath Ca2+ and addition of 1 mM EGTA; gadolinium, presence of 50 µM Gd3+ in the bath. Data (means ± SE) were pooled from at least 70 cells in a minimum of 3 experiments for each experimental condition.

 

Lactate dehydrogenase and ATP assay. Supernatants were collected 5 min before (control) and immediately after strain. Whole cell homogenates were prepared by adding a 2% Triton X-100 control solution. Aliquots of samples were stored at 4°C for the lactase dehydrogenase (LDH) measurement but immediately frozen in liquid nitrogen for the ATP analysis.

The LDH assay kit is based on the reduction of NAD+ to NADH/H+ by conversion of lactate to pyruvate. Diaphorase transfers H/H+ from NADH/H+ to a tetrazolium salt, which is reduced to the colored product formazan. Eighty microliters of diluted sample were incubated with one hundred microliters of reaction buffer in a 96-well plate for 30 min at room temperature and protected from light. The absorbance of the samples was determined at 492 nm (reference 630 nm) with a Tecan Spectra Thermo plate reader. Total LDH per sample was calculated by using the sum of homogenate and supernatant LDH activity. Because LDH release was expressed as a percentage of total LDH, before and after strain, and the measured LDH values were well within the range of the assay, it was not affected by the total number of cells.

ATP was determined with a luciferin-luciferase-based kit. Briefly, ATP standards and samples (10 µl) were pipetted into a white 96-well microplate. One hundred microliters of luciferin-luciferase reagent were added to the samples, and luminescence was immediately measured with a Tecan Spectra Fluor Plus. The detection limit of the assay was ~50 fmol of ATP per sample.

Simultaneous measurement of changes in [Ca2+]c and LB fusions. During experiments, cells were kept on membranes (fixed in homemade clamping devices) on the stage of an inverted Zeiss 135 TV Axiovert microscope at room temperature. Bath solution contained (in mM) 140 NaCl, 5 KCl, 1 MgCl2, 2 CaCl2, 5 glucose, and 10 HEPES, pH 7.4. Ca2+-free solutions contained no added CaCl2 and 1 mM EGTA.

For [Ca2+]c measurements, cells were loaded for 15–30 min at 37°C in DMEM with 1 µM fura 2-AM. At each excitation wavelength (340 and 380 nm) cells were illuminated for 50–100 ms at a rate of 0.2–2 Hz. [Ca2+]c values are expressed as baseline-corrected fura 2 ratios. Maximum amplitudes were calculated by subtracting the mean baseline values before strain (arbitrary units) from the maximum values of the strain-induced rise of the fura 2 ratio. In general, the sampling rate was high enough and lateral movement of the cell was small enough to ensure that errors of the fura 2 ratio due to pixel movement were minimal. In rare occasions when this occurred at the moving margins of a cell, the area of interest was restricted to the center part of the cell.

Determination of LB fusions by FM 1-43 fluorescence (FFM1-43) was recently described in detail (8). In short, this method is based on the cell-impermeant, surfactant-staining properties of FM 1-43, resulting in localized fluorescence after fusion as FM 1-43 enters LB through the exocytotic fusion pore. Importantly, FM 1-43 is nonfluorescent in aqueous solutions, permitting fusion to be monitored in the continuous presence of the dye in the bath. For exocytosis response time histograms, vesicle fusion was measured as the onset of localized FM 1-43 fluorescence (see Fig. 7A) as previously described (8), excited at 480 nm for 10 ms.



View larger version (22K):
[in this window]
[in a new window]
 
Fig. 7. Dependence of lamellar body (LB) fusions on the Ca2+ signal. A: FM 1-43 image of the same cells as in Fig. 4 before (a) and 500 s after (b) strain (22% {Delta}CSA). New LB fusions (= stimulated fusions) can be seen as additional fluorescent spots. B: % of cells with stimulated LB fusions as a function of the strain amplitude in different cell populations. "Ca2+-responders" (n = 120) and "non-Ca2+ responders" (n = 336), cells with or without a Ca2+ signal after strain, respectively (compare with Fig. 4B). Each bar represents at least 10 cells. C: no. of LB fusion events per cell within different cell populations, as described in B, under different experimental conditions (see Fig. 4B). Data from all strain amplitudes were pooled. *P < 0.05, significant difference vs. unstrained control.

 

Owing to the low number of single exocytotic events per cell, data from many single cells were pooled to establish exocytosis response time histograms (actual numbers of cells is given in Fig. 8). Each bar in a histogram represents all fusion events that occurred within a defined period of time after the onset of stimulation (expressed as % of all fusions within 12 min of total observation time). Additionally, all cells showing a rapid, irreversible loss of intracellular fura 2 concentration and/or influx of the hydrophobic dye FM 1-43 at any time during the experiment were omitted from data analysis because of an assumed loss of cell membrane integrity.



View larger version (18K):
[in this window]
[in a new window]
 
Fig. 8. LB fusion response time histogram in Ca2+ responders. Each bar in the histogram represents all fusion events occurring within a defined period of time after the onset of strain (expressed as % of all fusions within 12 min). Time 0 denotes onset of strain or onset of strain-induced [Ca2+]c increase (inset). Data were pooled from 37 cells.

 

LysoTracker Green DND-26 (LTG) fluorescence experiments were performed as recently described (13). LTG fluorescence provides an alternative means to study LB fusion with the plasma membrane and was used for validation of the FM 1-43 experiments.

Chemicals. The fluorescent dyes fura 2-AM and FM 1-43 and the luciferin-luciferase kit for the ATP assay were purchased from Molecular Probes (Leiden, The Netherlands). The Boehringer Mannheim LDH assay kit was from Roche Diagnostics (Vienna, Austria). All other chemicals were from Sigma (Vienna, Austria).

Statistical analysis. Results are represented as means ± SE except where otherwise noted. For statistical analysis we used unpaired Student's t-test, two-tailed paired Student's t-test, ANOVA, and linear regression. The mode of statistical analysis is noted when used. Differences were regarded as significant when P < 0.05.


    RESULTS
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Characterization of the strain device. The {epsilon}{theta} and {epsilon}r values within each membrane (n = 5) at different strain levels revealed no significant difference (P > 0.3, 2-tailed paired Student's t-test), demonstrating an equibiaxial strain field over the whole operational range. Additionally, the dependence of {epsilon}{theta} on r and {epsilon}r on {theta} was not significantly different (P > 0.3, ANOVA) within five membranes at any strain level (Fig. 2A shows {epsilon}{theta} and {epsilon}r values at a ring indention of 5.2 mm), again demonstrating a uniform deformation of the membrane. To characterize the operational range of the device, five membranes were used to establish calibration curves, relating the amount of ring indention to the increase in {Delta}MSA at three radial locations (Fig. 2B). No significant differences in %{Delta}MSA were found at different radial positions (P > 0.5, ANOVA) at any strain amplitude.

Cell viability is affected neither by strain amplitude nor by strain rate. Viability of strained cells was investigated by LDH measurements (n = 15). ATP measurements (n = 14) served as an additional parameter for cell injury and to test whether ATP is released during strain in an autocrine fashion. LDH and ATP release, determined at different strain amplitudes and strain rates, is shown in Fig. 3. Linear regression revealed no significant increase in LDH release, neither in dependence on strain amplitude (r2 = 0.0017; P = 0.88) nor in dependence on strain rate (r2 = 0.0215; P = 0.60). ATP release was also unaffected by strain amplitude (r2 = 0.00004; P = 0.98) and strain rate (r2 = 0.0741; P = 0.35). The mean ATP concentration in the supernatant was 17.1 ± 2.8 nM, a concentration far lower than that required for a paracrine purinergic activation of type II cells (25, 26).



View larger version (51K):
[in this window]
[in a new window]
 
Fig. 3. A and B: cell viability determination by measurement of lactate dehydrogenase (LDH) and ATP release after strain. Changes in LDH release (n = 15) as well as ATP release (n = 14) were measured at different strain amplitudes and strain rates. Circles represent data of single experiments. Increasing strain amplitudes and strain rates had no significant effect on LDH (A) or ATP (B) release after strain. {Delta}CSA, change in cell surface area. C: determination of resealing mechanisms in a wounded cell by leakage of fura 2 and FM 1-43. Loss of fura 2 fluorescence (top) and increase of FM 1-43 fluorescence (bottom) occur in 1 of several cells during and after strain. Images were selected at the indicated times of the time course shown in the graph. Inset: time course and amount (%{Delta}CSA) of cell strain.

 

Additional parameters for cell integrity were the lack of fura 2 leakage out of the cells and the lack of FM 1-43 leakage into the cells. This occurred only on very rare occasions (<1%) at strain levels below 40% {Delta}CSA, with a slightly increased probability in the Ca2+-free bath solution. Using membrane-impermeant fluorescence molecules, Vlahakis et al. (35) reported that in a fraction of cells resealing of the plasma membrane occurred after stress failure. Using FM 1-43 and fura 2 as tools to assess membrane integrity, we found no evidence for complete resealing after relaxation from strain among the few injured cells. This is shown in Fig. 3C: the leakages of fura 2 and FM 1-43 exhibit different time courses of diffusion. It is evident that FM 1-43 leakage into the cell persists for a long time after relaxation of strain, continuing even after complete loss of fura 2 from the cell, suggesting a persistently increased membrane permeability for FM 1-43 (although occasional FM 1-43 uptake also occurred in the absence of strain, this process was, if present, far slower). Naturally, "resealing" may not be an all-or-none event but may depend on the molecular properties of the permeating substance used.

Single-cell Ca2+ response to strain revealed dependence on extracellular Ca2+ and strain amplitude. A single strain above a threshold of 8% {Delta}CSA led to a transient rise of [Ca2+]c concentration in some AT II cells (we refer to these cells as "Ca2+ responders"). The incidence of a rise in [Ca2+]c after strain increased significantly (P < 0.05, paired t-test) with enhanced strain amplitudes (Fig. 4B) and reached a maximum at ~40% increase in CSA (>80% responding cells). Higher strain amplitudes did not cause additional cells to respond with a Ca2+ signal (data not shown). This strain-induced Ca2+ response was completely inhibited by removal of bath Ca2+ (n = 174 cells from 21 independent experiments) but unaffected by the presence of 50 µM Gd3+, a blocker of various cation channels, particularly mechanosensitive channels, in the bath (Fig. 4B). Importantly, the lack of a [Ca2+]c elevation in the Ca2+-free bath solution was not a result of empty Ca2+ stores, because subsequent addition of 10 µM ATP to the bath elicited a Ca2+ signal, as previously described (8). Moreover, strain-induced Ca2+ signals could be evoked by a second strain directly following readdition of Ca2+ to the bath in cells lacking this response under prior Ca2+-free conditions (data not shown). Data were obtained from at least 70 cells within a minimum of three independent experiments.

Despite the clear relationship between strain and number of Ca2+ responders, Ca2+ signals varied considerably between individual cells (Fig. 5). In general, Ca2+ signals shared a transient initial peak, even in case of a static strain (Fig. 5C). Further analysis revealed that the [Ca2+]c elevations were not a function of the strain amplitudes except for those between 30% and 40% {Delta}CSA (Fig. 6A). We speculated that this might be due to various degrees of membrane unfolding that are not sensed by the cells. Only at higher strain amplitudes is strain assumed to exert a force on a strain sensor, presumably the Ca2+ channel or structures associated with it (membrane stress). To test this hypothesis, analysis was limited to the strain amplitude above threshold (i.e., above the strain amplitude initiating the [Ca2+]c rise) in each single cell, and the CSA increase above this threshold value was defined as "effective strain." Figure 6B reveals a clear correlation between the effective strain and the maximum [Ca2+]c elevation (P < 0.05, paired t-test). This graded response to effective strain was not affected by the presence of 50 µM Gd3+ in the bath solution (P > 0.05, unpaired t-test) but was inhibited by thapsigargin pretreatment (100 nM; Fig. 6C). Thapsigargin is a specific blocker of the Ca2+-ATPase in the endoplasmic reticulum and commonly used to deplete this important intracellular Ca2+ store.



View larger version (16K):
[in this window]
[in a new window]
 
Fig. 5. Strain-induced [Ca2+]c signals varied significantly between individual cells. Insets: strain profile (%{Delta}CSA) during the experiment. A: examples of [Ca2+]c responses (cells A and B) to a transient single strain of ~40% {Delta}CSA. B: example of a [Ca2+]c response to a sustained (static) strain of 40% {Delta}CSA.

 


View larger version (12K):
[in this window]
[in a new window]
 
Fig. 6. Dependence of the [Ca2+]c peak amplitude on the strain amplitude above threshold. A: change ({Delta}) in fura 2 ratio as a function of absolute {Delta}CSA [*significant difference (P < 0.05), unpaired t-test]. B: {Delta} in fura 2 ratio as a function of effective {Delta}CSA (see RESULTS for definition). C: effect of experimental conditions as in Fig. 4B. Data were pooled for 2 different strain amplitudes from at least 10 cells within a minimum of 5 experiments. Bars indicate means ± SE.

 

Cell-cell contacts affected strain-induced Ca2+ signal. The generation of a strain-induced Ca2+ signal strongly depended on the cell location within a monolayer. Only cells within a group, but never lone cells (n = 55 cells, 16 independent experiments), exhibited a Ca2+ signal after strain. This nonresponsiveness of lone cells was independent of strain amplitude (0–45% {Delta}CSA) and strain rate (0–2% {Delta}CSA/s).

Strain-induced Ca2+ signaling triggered LB fusions with plasma membrane. Strain-induced fusion events were observed in a distinct number of cells (exemplified in Fig. 7A). As we showed previously, exocytosis in type II cells is triggered above a [Ca2+]c threshold of ~320 nM (11) and the overall fusion response is tightly correlated with the time course of the Ca2+ signal (8). Here we examined strain-induced fusion events in Ca2+ responders and cells without a rise in [Ca2+]c ("non-Ca2+ responders"). Significant differences (P > 0.05, unpaired t-test) between these two groups of cells were observed (Fig. 7B). Within the Ca2+ responders (n = 120), the percentage of cells exhibiting fusion events was clearly related to the strain amplitude, whereas in non-Ca2+ responders (n = 336) the percentage of cells responding with fusion remained unchanged. Data were obtained from at least 10 cells within a minimum of three experiments.

Furthermore, in non-Ca2+ responders (n = 194), strain-induced fusions per cell were not significantly different from unstrained controls (n = 289), whereas Ca2+ responders (n = 120) exhibited a significantly (P > 0.05, unpaired t-test) enhanced fusion activity (Fig. 7C). This was not affected by the presence of 50 µM Gd3+ (n = 71 for non-Ca2+ responders and n = 39 for Ca2+ responders; Fig. 7C). Data on strain-induced LB fusions after thapsigargin treatment are not presented because thapsigargin per se (i.e., without strain) elicited a transient elevation of [Ca2+]c (data not shown) and stimulated LB fusions, resulting in LB depletion before application of strain.

Fusion response histogram. The LB fusion response histogram for strain and for the strain-induced Ca2+ signal is shown in Fig. 8. The data reveal that maximum fusion activity is delayed by 2–3 min after application of strain or 1–2 min after the Ca2+ signal and that stimulated fusion activity terminates after ~10 min.

Validation of FFM1-43-based LB fusion during cell strain. In the absence of cell strain, the FFM1-43 method has a high specificity and reliability for the detection of LB fusion in AT II cells (12, 13, 30). During conditions of cell strain, however, this could be different. To examine this, we performed experiments with LTG fluorescence (FLTG). As recently outlined in detail (13), FLTG is restricted to LBs owing to their acidic pH before fusion with the plasma membrane. After LB fusion, release of LTG into the extracellular space results in localized FLTG loss. An example of FLTG in a single cell in response to strain is shown in Fig. 9: loss of FLTG occurred exactly and exclusively at those sites that stained with FM 1-43. Hence, we conclude that the specificity of the FM 1-43 method is maintained under conditions of strain.



View larger version (45K):
[in this window]
[in a new window]
 
Fig. 9. LB fusion measured with Lyso-Tracker Green DND-26 fluorescence (FLTG). Top images: this cell was used for analysis (graph). Top lane: fluorescence intensities of fura 2 + LTG (gray scale) at various times before, during, and after strain. Middle lane: fura 2 ratios (pseudocolors as in Fig. 4A) of the same cell. Bottom lane: LTG fluorescence (true color) of this cell. FM 1-43 (orange fluorescence) was added as control for fused LBs (loss of FLTG) to the bath shortly before image at 12 min was taken. Graph shows time course of fura 2 ratio (black line), measured over the area of the entire cell, and FLTG (colored lines), measured over localized areas as indicated by arrows in the last 2 images. Inset: {Delta}CSA (%).

 


    DISCUSSION
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Methodological aspects of cell strain. The basic principle of the strain system is similar to a number of previously described techniques (15, 16, 29, 31, 32) in which a flexible substrate of the cells acts as a diaphragm that is mechanically deformed by a piston mechanism. In contrast to these setups, our system is exclusively designed to meet the requirements for live imaging of cells during the whole strain process. To achieve this, cells must be kept in the same focal plane throughout the experiment and lateral shifting of cells during strain must be minimized. Although we use objectives with high magnification, these problems occur only at high strain amplitudes due to membrane flattening, shifting the cells slightly up- or downward in the focal plane. However, strain meets the theoretical predictions by being uniform (equibiaxial) over the part of the Silastic substrate that remains in the field of interest.

Because changes in CSA could be directly determined for each single cell, the relationship between {Delta}MSA and {Delta}CSA after strain, as used in most previous studies, was not a necessary prerequisite for analysis and {Delta}MSA was not routinely assessed. Nevertheless, comparison of {Delta}MSA with {Delta}CSA in a few experiments revealed no apparent difference (data not shown). In this context, it is worth noting that AT II cells attached well to Silastic membranes within 24 h after isolation and did not lose contact during application of strain unless strain amplitudes exceeded 40% {Delta}CSA (unpublished observation). This was based on the finding that CSA after relaxation of strain roughly equaled CSA before strain, indicating maintenance of cell shape and attachment. These findings were in line with the findings of Tschumperlin and Margulies (32).

Cell viability. Cell viability, defined as plasma membrane integrity, was affected neither by strain amplitude nor by strain rate when strain protocols that closely mimic physiological conditions were used. Even extended strain amplitudes up to 45% {Delta}CSA affected cell viability in <10% of cells. This low incidence of cell injuries found by us is probably due to the low strain rates we used (0–2% {Delta}CSA/s), consistent with the finding of Vlahakis et al. (35) that cell membrane injury correlates well with the strain rate at constant strain amplitude. The findings presented here, and those by other authors using A549 cells (31), suggest that room temperature has no negative influence on strain tolerance. On the other hand, the temperature may considerably affect the ability of wounded cells to reseal, consistent with the effect of temperature on lipid trafficking (35). Although freshly seeded AT II cells (24 h) were reported to exhibit significantly increased mortality even at low strain amplitudes (32), we did not observe enhanced mortality in our primary cultures between 28 and 40 h after isolation. This apparently high strain tolerance appears to be an important protective mechanism against hyperinflation-induced lung injury, even though previous stress analysis of the alveolus (9) revealed that AT II cells in vivo may be protected against large-scale deformations (because of their preferential location in the alveolar corners). This ability is likely caused by membrane unfolding and cell flattening, because the plasma membrane can sustain only small amounts of strain between 2% and 3% (in the plane of the membrane) before it breaks (reviewed in Refs. 20, 34). The amount of "recruitable membrane" would also explain the strong cell-to-cell variability of Ca2+ signaling and its dependence on cell-cell contacts (see also Strain-induced Ca2+ signaling and LB fusions), indicating that the Ca2+ signal is possibly a most sensitive parameter of membrane stress.

Strain-induced Ca2+ signaling and LB fusions. Results from combined [Ca2+]c and LB fusion measurements demonstrated that strain of AT II cells above a threshold of 8% {Delta}CSA resulted in a transient rise of [Ca2+]c (in agreement with Ref. 37), with enhanced strain amplitudes leading to an increased number of Ca2+ responders. As a result, fusion activity in AT II cells increased in a dose-dependent way. This threshold value of 8–10% {Delta}CSA is unlikely to be accomplished by normal tidal lung volumes, because within this range alveolar distension is due to unfolding of alveolar structures rather than single-cell strain (2, 21, 33). At higher volumes, e.g., during exercise, however, the graded response would adjust the supply of surfactant to the increased demand (22). The graded response of the Ca2+ signal to strain, which was already described by Wirtz and Dobbs (37), is most likely the result of increased plasma membrane stress with increasing cell distension. In fact, the correlation between effective strain as defined here (see RESULTS) and the amplitude of the Ca2+ signal (Fig. 6B) suggest that Ca2+ channel activation may be not only the most sensitive parameter for membrane stress but also the structure that defines the threshold of "physiologically relevant" strain. The graded response of LB fusion to strain is also entirely consistent with previous analyses revealing the number of fusion events to be a function of the integrated [Ca2+]c elevation over time (8). This relationship is probably an important principle for repetitive strains, which were not the subject of this study.

In the search for the molecular identity of the mechanosensor, the initial event of Ca2+ entry activation intuitively directs attention toward the plasma membrane and the Ca2+ channel. Naturally, this channel may also be activated downstream of another mechanosensor located apart from the plasma membrane.

Lone cells never elicited [Ca2+]c signals, regardless of strain amplitudes and strain rates, revealing cell-cell connections as a prerequisite for the generation of Ca2+ signals. This might be a result of different strain "distributions" within lone cells compared with grouped cells. Whereas lone cells may have a cell architecture and membrane structure that allow an easy membrane recruitment, grouped cells may exhibit an inhomogeneous strain field due to junctional complexes, affecting a possible recruitment of membrane reservoirs.

In an elegant study using the intact alveolus in situ, Ashino et al. (1) recently suggested that alveolar type I (AT I) cells rather than AT II cells are the main site of mechanotransduction because alveolar expansion evoked [Ca2+]c oscillations in AT I cells, which communicated to AT II cells. We agree with their conclusion that surfactant secretion may not be entirely self-regulated by AT II cells. It appears reasonable to assume that neither AT I nor AT II cells are exclusive in their function as mechanosensors but that the modes of regulation depend on various factors such as cell location within the alveolus, distinct stress distributions within the whole lung, and possibly many other factors.

The data presented in Fig. 6C strongly suggest two distinct mechanisms to account for the strain-induced elevation of [Ca2+]c: 1) a Ca2+ entry pathway with a high sensitivity to strain (this mechanism operates at low effective strain amplitudes and is not affected by pretreatment with thapsigargin) and 2) intracellular Ca2+ release from Ca2+ stores. The second mechanism is activated at high effective strain amplitudes and is completely inhibited by prior depletion of the Ca2+ stores with thapsigargin. It operates in addition to Ca2+ entry and, importantly, is entirely dependent on Ca2+ entry. This is evidenced by the total lack of Ca2+ signaling in the absence of Ca2+ in the bath. It clearly indicates that Ca2+ entry is the prime event triggered by strain and a prerequisite for Ca2+ release to follow. This is an extension to previous findings (37), because Ca2+ signaling was reported to result from intracellular Ca2+ release exclusively. The authors of the previous study did not discuss how the release mechanism may be triggered. From our data we suggest that Ca2+-induced Ca2+ release is the underlying mechanism. However, we do not have an explanation for the reduced number of Ca2+ responders at each strain amplitude after store depletion by thapsigargin (Fig. 4B). It should be considered, however, that thapsigargin activates an additional store-operated Ca2+ entry (SOCE) pathway in AT II cells (unpublished findings) and the interaction between SOCE and strain-induced Ca2+ entry is unknown.

Mechanical stimulation increases Ca2+ influx through stretch-activated ion channels in fetal lung cells (17). Irrespective of whether this Ca2+ influx occurs through Ca2+-selective or nonselective cation channels, they certainly belong to a class of lanthanide-independent channels because they could not be inhibited by Gd3+. Importantly, we can essentially exclude that the strain-induced Ca2+ entry proceeds via L-type Ca2+ channels because they are not functionally present in AT II cells (8). Consequently, the nature of the strain-induced Ca2+ entry pathway in the AT II cell will have to be determined in further studies.

All data presented here suggest Ca2+ to be the major, if not exclusive, second messenger for strain-induced surfactant secretion, consistent with other published findings (37). Naturally, we cannot infer a direct action of Ca2+ on the exocytotic fusion machinery, and other second messengers, activated downstream or in parallel with Ca2+ entry (such as protein kinase C), may be involved. The response time of LB fusions to strain, as presented here (Fig. 8), is relatively short (a few minutes) compared with strain-induced surfactant secretion (~30 min; Ref. 37). This dissociation between fusion and release is not unexpected, considering the unique release properties of these cells, which have been described in detail previously (12, 13, 30). With regard to exocytosis, two major events take place during and after a single distension of type II cells: 1) strain-induced fusion pore expansion (30) (this effect probably accounts for the rapid release of those LBs that had already fused before strain either constitutively or by stimulation) and 2) strain-induced LB fusions (this occurs, as shown here, after a certain delay and serves to keep a certain number of LBs in the fused state, ready to quickly release their contents during the next deep breath).

It is likely that these two distinct processes are of fundamental importance in the homeostasis of the surfactant system in vivo, because it is generally assumed that strain of AT II cells is the most physiologically relevant stimulus for surfactant secretion in vivo (7, 19, 28, 36, 38). The nature of the strain-induced Ca2+ entry pathway will have to be determined. It might serve as a useful pharmacological target against hyperventilation-induced lung injury.


    ACKNOWLEDGMENTS
 
The technical assistance of Irina Öttl and Gerlinde Siber is gratefully acknowledged.

Parts of this work were presented at the Meeting of the German Physiological Society, Bochum, Germany, 2003, and the Experimental Biology 2002 Meeting, New Orleans, LA, 2002.

GRANTS

This study was supported by Austrian Science Foundation grants P14263 [GenBank] , P15742, and P15743 [GenBank] , and the Austrian Bundesministerium für Bildung, Wissenschaft und Kunst.


    FOOTNOTES
 

Address for reprint requests and other correspondence: P. Dietl, Dept. of Physiology, Univ. of Innsbruck, Fritz-Pregl-Str. 3, A-6020 Innsbruck, Austria (E-mail: paul.dietl{at}uibk.ac.at).

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.


    REFERENCES
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. 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]
  2. 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]
  3. Chander A and Fisher AB. Regulation of lung surfactant secretion. Am J Physiol Lung Cell Mol Physiol 258: L241-L253, 1990.[Abstract/Free Full Text]
  4. Dobbs LG. Pulmonary surfactant. Annu Rev Med 40: 431-446, 1989.[CrossRef][ISI][Medline]
  5. Dobbs LG, Gonzalez RF, Marinari LA, Mescher EJ, and Hawgood S. The role of calcium in the secretion of surfactant by rat alveolar type II cells. Biochim Biophys Acta 877: 305-313, 1986.[ISI][Medline]
  6. 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]
  7. Edwards YS. Stretch stimulation: its effects on alveolar type II cell function in the lung. Comp Biochem Physiol A Mol Integr Physiol 129: 245-260, 2001.[CrossRef][ISI][Medline]
  8. Frick M, Eschertzhuber S, Haller T, Mair N, and Dietl P. Secretion in alveolar type II cells at the interface of constitutive and regulated exocytosis. Am J Respir Cell Mol Biol 25: 306-315, 2001.[Abstract/Free Full Text]
  9. Gefen A, Elad D, and Shiner RJ. Analysis of stress distribution in the alveolar septa of normal and simulated emphysematic lungs. J Biomech 32: 891-897, 1999.[CrossRef][ISI][Medline]
  10. Goerke J. Pulmonary surfactant: functions and molecular composition. Biochim Biophys Acta 1408: 79-89, 1998.[ISI][Medline]
  11. Haller T, Auktor K, Frick M, Mair N, and Dietl P. Threshold calcium levels for lamellar body exocytosis in type II pneumocytes. Am J Physiol Lung Cell Mol Physiol 277: L893-L900, 1999.[Abstract/Free Full Text]
  12. Haller T, Dietl P, Pfaller K, Frick M, Mair N, Paulmichl M, Hess MW, Furst J, and Maly K. Fusion pore expansion is a slow, discontinuous, and Ca2+-dependent process regulating secretion from alveolar type II cells. J Cell Biol 155: 279-289, 2001.[Abstract/Free Full Text]
  13. Haller T, Ortmayr J, Friedrich F, Volkl H, and Dietl P. Dynamics of surfactant release in alveolar type II cells. Proc Natl Acad Sci USA 95: 1579-1584, 1998.[Abstract/Free Full Text]
  14. Hollingsworth M and Gilfillan AM. The pharmacology of lung surfactant secretion. Pharmacol Rev 36: 69-90, 1984.[ISI][Medline]
  15. Hung CT and Williams JL. A method for inducing equibiaxial and uniform strains in elastomeric membranes used as cell substrates. J Biomech 27: 227-232, 1994.[ISI][Medline]
  16. Lee AA, Delhaas T, Waldman LK, MacKenna DA, Villarreal FJ, and McCulloch AD. An equibiaxial strain system for cultured cells. Am J Physiol Cell Physiol 271: C1400-C1408, 1996.[Abstract/Free Full Text]
  17. Liu M, Xu J, Tanswell AK, and Post M. Inhibition of mechanical strain-induced fetal rat lung cell proliferation by gadolinium, a stretch-activated channel blocker. J Cell Physiol 161: 501-507, 1994.[ISI][Medline]
  18. Mair N, Haller T, and Dietl P. Exocytosis in alveolar type II cells revealed by cell capacitance and fluorescence measurements. Am J Physiol Lung Cell Mol Physiol 276: L376-L382, 1999.[Abstract/Free Full Text]
  19. Mason RJ and Voelker DR. Regulatory mechanisms of surfactant secretion. Biochim Biophys Acta 1408: 226-240, 1998.[ISI][Medline]
  20. Matthay MA, Bhattacharya S, Gaver D, Ware LB, Lim LH, Syrkina O, Eyal F, and Hubmayr R. Ventilator-induced lung injury: in vivo and in vitro mechanisms. Am J Physiol Lung Cell Mol Physiol 283: L678-L682, 2002.[Abstract/Free Full Text]
  21. 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]
  22. Nicholas TE, Power JH, and Barr HA. Surfactant homeostasis in the rat lung during swimming exercise. J Appl Physiol 53: 1521-1528, 1982.[Abstract/Free Full Text]
  23. Nicholas TE, Power JH, and Barr HA. The pulmonary consequences of a deep breath. Respir Physiol 49: 315-324, 1982.[CrossRef][ISI][Medline]
  24. Pian MS, Dobbs LG, and Duzgunes N. Positive correlation between cytosolic free calcium and surfactant secretion in cultured rat alveolar type II cells. Biochim Biophys Acta 960: 43-53, 1988.[ISI][Medline]
  25. Rice WR, Dorn CC, and Singleton FM. P2-purinoceptor regulation of surfactant phosphatidylcholine secretion. Relative roles of calcium and protein kinase C. Biochem J 266: 407-413, 1990.[ISI][Medline]
  26. Rice WR and Singleton FM. P2Y-purinoceptor regulation of surfactant secretion from rat isolated alveolar type II cells is associated with mobilization of intracellular calcium. Br J Pharmacol 91: 833-838, 1987.[Abstract]
  27. Rooney SA. Lung Surfactant: Cellular and Molecular Processing. Georgetown, TX: Landes, 1998.
  28. Rooney SA. Regulation of surfactant secretion. Comp Biochem Physiol A Mol Integr Physiol 129: 233-243, 2001.[CrossRef][ISI][Medline]
  29. Schaffer JL, Rizen M, L'Italien GJ, Benbrahim A, Megerman J, Gerstenfeld LC, and Gray ML. Device for the application of a dynamic biaxially uniform and isotropic strain to a flexible cell culture membrane. J Orthop Res 12: 709-719, 1994.[ISI][Medline]
  30. Singer W, Frick M, Haller T, Bernet S, Ritsch-Marte M, and Dietl P. Mechanical forces impeding exocytotic surfactant release revealed by optical tweezers. Biophys J 84: 1344-1351, 2003.[Abstract/Free Full Text]
  31. 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.[Abstract/Free Full Text]
  32. 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]
  33. 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]
  34. Vlahakis NE and Hubmayr RD. Plasma membrane stress failure in alveolar epithelial cells. J Appl Physiol 89: 2490-2496, 2000.[Abstract/Free Full Text]
  35. Vlahakis NE, Schroeder MA, Pagano RE, and Hubmayr RD. Role of deformation-induced lipid trafficking in the prevention of plasma membrane stress failure. Am J Respir Crit Care Med 166: 1282-1289, 2002.[Abstract/Free Full Text]
  36. Wirtz H and Schmidt M. Ventilation and secretion of pulmonary surfactant. Clin Investig 70: 3-13, 1992.[ISI][Medline]
  37. Wirtz HR and Dobbs LG. Calcium mobilization and exocytosis after one mechanical stretch of lung epithelial cells. Science 250: 1266-1269, 1990.[ISI][Medline]
  38. Wirtz HR and Dobbs LG. The effects of mechanical forces on lung functions. Respir Physiol 119: 1-17, 2000.[CrossRef][ISI][Medline]
  39. Wright JR and Clements JA. Metabolism and turnover of lung surfactant. Am Rev Respir Dis 136: 426-444, 1987.[ISI][Medline]