Departments of 1Physiology and 2Medical Physics, University of Innsbruck, A-6020 Innsbruck, Austria
Submitted 15 September 2003 ; accepted in final form 21 September 2003
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ABSTRACT |
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surfactant secretion; alveolar type II cells; mechanical strain; stretch
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.
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METHODS |
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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.
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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 () and radial (
r) strains (
= u/r,
r =
u/
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
=
r, the strain field is uniform (equibiaxial), indicating equal strains in all directions. Therefore, we compared the
r and
values at different strain levels. The dependence of average
amplitude on r and average
r amplitude on
(radial direction) were measured at every indention of the inner cylinder and calculated over five membranes.
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Membrane deformations were computed according to Tschumperlin and Margulies (32). The stretch ratio () in an equibiaxial strain field is defined as
=
r =
- 1 and can be computed according to
= (r + u)/r. We calculated the average
for every radial position at every strain level by averaging the
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 (%
MSA) were determined by the method described in detail by Tschumperlin and Margulies (32) and were calculated by %
MSA = (
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 (%CSA) were expressed according to %
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 %
CSA per second.
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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 1530 min at 37°C in DMEM with 1 µM fura 2-AM. At each excitation wavelength (340 and 380 nm) cells were illuminated for 50100 ms at a rate of 0.22 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.
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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.
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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.
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RESULTS |
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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).
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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% 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% 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% 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.
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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 (045% CSA) and strain rate (02%
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 23 min after application of strain or 12 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.
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DISCUSSION |
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Because changes in CSA could be directly determined for each single cell, the relationship between MSA and
CSA after strain, as used in most previous studies, was not a necessary prerequisite for analysis and
MSA was not routinely assessed. Nevertheless, comparison of
MSA with
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%
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% 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 (02%
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% 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 810%
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.
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ACKNOWLEDGMENTS |
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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.
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FOOTNOTES |
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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.
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REFERENCES |
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