Departments of 1Physiology, 2Hematology and Oncology, 3Internal Medicine, 5Anesthesiology and Critical Care Medicine; University of Innsbruck, A-6020 Innsbruck, Austria; and 4Department of Gynecology and Obstetrics, State University of New York, Buffalo, New York 14222
Submitted 25 September 2003 ; accepted in final form 29 December 2003
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ABSTRACT |
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alveolus; lamellar body; pulmonary; secretion; surface tension
The fluorophore FM 1-43 is ideally suited to monitor exocytosis in type II cells. This amphiphilic molecule passes from the external solution through the fusion pore, formed during exocytosis of lamellar bodies, into the lumen of these vesicles (16). The preferential staining of surfactant by FM 1-43 makes it possible to follow an LBP as it is further extruded through the fusion pore in the extracellular fluid. However, a better understanding of the extracellular surfactant's life cycle also requires detailed information on the fate of LBPs when they finally reach the air-liquid interface. Thus we designed a cell culture system allowing direct assessment of the transformation of FM 1-43-labeled LBPs immediately after release by the type II cells and subsequent insertion in an air-liquid interface.
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METHODS |
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Surface tension measurements. For static surface tension measurements of cell supernatants, we adopted and modified the vertical pull method (Fig. 1B). According to this method, the maximum change in weight of a fluid meniscus, suspended by a cylindrical stainless steel rod, is recorded and, together with sample density and rod radius, used to calculate surface tension from a set of published equations (2). Although detachment methods tend to overestimate surface tension, especially in the presence of insoluble films, we found the vertical pull method to be both sufficient and ideally suited for measurements of cell supernatants. Furthermore, we verified the accuracy of this method with a number of solutions and surfactant preparations, e.g., methanol (60% vol/vol): vertical pull 31.5 mN/m, standard value 32 mN/m (36), or Curosurf (2 mg/ml): vertical pull 26.7 ± 0.6, n = 3, vs. 23.3 ± 1.7, n = 13, independently measured by captive bubble (26).
For a functional evaluation of the LBP's ability to form a surfactant film at an air-liquid interface, we used a capillary surfactometer. As described in detail (5), this instrument adds up the periods during which a glass capillary is blocked by liquid and gives as a result the percentage of a 2-min period during which the capillary is open. Values close to 100% are indicative for a functional surfactant exerting high surface pressures. Frozen cell supernatants from several experiments were thawed, filtered, and pooled. Aliquots (1 ml each) of adjusted LBP concentrations were centrifuged at 40,000 g for 1 h. The resulting pellet (3% of the initial sample volume) was resuspended and directly used. Thus the indicated LBP concentrations (Fig. 2F) refer to the concentrations before centrifugation.
Quantifying LBPs. Filtered LBP suspensions were pumped at a constant rate of 0.28 µl/min through a micropipette (inner diameter 28 ± 2 µm), as illustrated (Fig. 1A). The pump (Cyclobios) consisted of a step motor, which was driving a high-precision screw micrometer pushing the piston of a 50-µl gas-tight Hamilton syringe (Hamilton). The syringe was connected by a Teflon tubing with the micropipette and mounted on the stage of a microscope. The micropipettes were prepared from borosilicate glass capillaries by a vertical puller and glued to coverslips (0.17 mm thickness) with Entellan (Merck) so that their narrowest portion was within the focal plane of the objective (Fluar x40 oil; Zeiss). They were replaced after each use. The passage of single FM 1-43-labeled LBPs was detected by a photomultiplier tube (Hamamatsu), and the signals were recorded with the software Pulse (Heka). Data were analyzed by an automated peak detection program. Particle size was calculated from the integrated peak area as shown. The limit of detectability was tested by fluorescent beads and found to be 200 nm. The advantages of this setup compared with flow cytometry or Coulter Counter techniques were the highly constant flow rate combined with a low detection limit.
Phospholipids were measured enzymatically using the phospholipid kit MPR2 (Roche), as described previously (33) with the following modification: 20 µl of sample in PBS was mixed with 80 µl reagent and incubated for 25 min at room temperature. Precinorm L (Roche) was used as an internal standard. Absorbance was read at 495 nm with a Spectrophotometer DU 640 (Beckman).
Chamber with inverted air-liquid interface. A chamber was built to visualize insertion of LBPs in an air-liquid interface (Fig. 1C). It had a conical interior (volume 400 µl) with a wide opening at the top (Ø 12 mm) and a small, sharply edged aperture at the bottom (Ø <300 µm). At the lower aperture, liquid in the chamber formed an interface with the air below. The chamber was cast from polyester resin. A conically shaped and polished stainless steel rod was used as a mold. After polymerization, the steel rod was removed, and the bottom of the chamber was ground and polished until the desired width of the aperture was achieved. The conical shape caused LBPs to slide down along the chamber's inner wall, whereby they became concentrated when arriving at the narrow air-liquid interface. Furthermore, the small aperture helped maintain a minimally domed surface (radius of curvature 2 mm). That we actually focused on the surface and not the subphase was confirmed by the fact that sedimenting LBPs, randomly moving at high velocity when approaching the interface from above, abruptly slowed down when reaching the focal plane. Moreover, below that plane structures were never detected.
Imaging techniques. The images shown in Fig. 2A were obtained with a Zeiss Laser Scan Microscope (LSM 410 invert; Zeiss). For all other experiments, we used an inverted microscope (Zeiss 100) equipped for polychromatic illumination and image analysis (TILL Photonics). For the measurements at the air-liquid interface we used a long-distance objective (x40 LD-Achroplan). The charge-coupled device camera (Imago-SVGA; TILL Photonics) was operated at an image acquisition rate of 1 Hz with a symmetrical binning factor of two.
Solutions and materials. The experimental solution contained, in mM: 140 NaCl, 5 KCl, 1 MgCl2, 2 CaCl2, and 10 HEPES (pH 7.4). FM 1-43 was purchased from Molecular Probes, and Curosurf was from Nycomed (Linz). Other chemicals were obtained from Sigma (Munich, Germany). Experiments were performed at room temperature, except those shown in Fig. 2, B and F (temperature = 37°C).
Statistics. Data are presented as arithmetic means ± SD.
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RESULTS |
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Fate of LBPs after release and contribution to surface tension. After stimulation of cultured type II cells (Fig. 2B; 10 µM ATP at time 0), aliquots of cell supernatants were collected for further investigating the fate of released LBPs. We found that accumulation of LBPs was approaching equilibrium with a time constant () of 122 min. After 6 h at 37°C, cell supernatants contained 2.3 ± 0.7 x 106 LBPs/ml, corresponding to 0.55 µg phospholipid/ml (Fig. 2C) or to a mean of 7.3 LBPs released/single cell. In addition, it was found that mean LBP size increased rather than decreased (Fig. 2B). These findings showed that LBPs do not progressively disperse in experimental solution. To further investigate if LBPs transform in extracellular solution after release from the cells, we measured surface tension of cell supernatants in relation to LBP release. Surface tension was quickly lowered to
58 mN/m but then remained at that value despite ongoing LBP release (Fig. 2D). This result showed that some surface active material was present in the extracellular solution. To test whether a further decrease in surface tension could be obtained with higher LBP densities or, alternatively, after prolonged storage, we increased their concentration to
55 x 106/ml (Fig. 2E). This was achieved by collecting filtered supernatants from numerous stimulated cell cultures followed by storage at -70°C. Before analysis, these samples were thawed, pooled, and gently centrifuged (4,000 g for 1 h). The surface tension of these samples was
52 mN/m and thus only slightly lower than the value obtained when the LBP density was considerably less (Fig. 2D). However, these values are in the range of data obtained for suspension of lamellar bodies prepared from homogenized lungs (7, 25). When the samples were reanalyzed after 2 and 5 days, the number of LBPs/ml was reduced to
43 x 106 (Fig. 2E). This was probably the result of adherence of LBPs on the wall of the vials. Yet static surface tension was still
53 mN/m. The size and shape of LBPs after 2 and 5 days of storage were not different from what they were right after thawing (Fig. 2E). These results suggest that a significant fraction of LBPs had not transformed during this period of time and that surface tension attains an equilibrium that is largely independent of LBP subsurface concentration or time of storage.
Measurements with the capillary surfactometer showed that the ability of LBPs to develop a film with high surface pressure was related to LBP concentration. When it was lower than 2 x 107/ml, the ability to maintain capillary patency was close to zero. However, it improved significantly as the LBP concentration was increased, and above
4 x 107 LBPs/ml the capillary was open in 77 ± 13% (Fig. 2F). From these observations, we conclude that LBPs, when released from the cells in the extracellular space, are remarkably stable. However, a fraction of LBPs is likely to be transformed and provided as surface-active material that can reach the air-liquid interface and exert a significant interfacial effect.
LBP transformations at the inverted interface. When the chamber was filled with experimental solution containing ATP (10 µM) and FM 1-43 (1 µM), and a coverslip with adherent cells was added on top (as depicted in Fig. 1C), released LBPs appeared at the inverted interface because of ongoing exocytosis and sedimentation (Fig. 3A). Similarly, when aliquots of filtered cell supernatants containing freshly released LBPs were added on top of the experimental solution containing FM 1-43, the same observation was made. When we then investigated the insertion of LBPs in the air-liquid interface, we found that LBPs transformed instantaneously (<1 s) as soon as they hit the interface (video available at: http://physiologie.uibk.ac.at/dietl). In some cases, LBPs expanded into large, dark (Fig. 3, B and C) or bright (Fig. 3, D and E) areas. Occasionally, dark spots remained within bright areas (Fig. 3B). In other cases (Fig. 3, B and F), they split into several smaller particles before they slowly disappeared. To exclude the possibility that LBP transformation was caused by FM 1-43, we repeated these experiments in the absence of dye by phase-contrast microscopy (Fig. 3G). Again, it was found that LBPs disappeared after they hit the surface. However, in contrast to the observations with fluorescent LBPs, no expanded remnant structures could be detected. From these observations, we conclude that LBPs spontaneously transform upon insertion in the air-liquid interface.
When we inspected the surface several minutes later, it was filled with numerous LBPs (Fig. 4A) that had not transformed despite surface contact. Importantly, transformations were observed in the first few minutes of accumulation only. We reasoned that the number of transformations was a function of material already inserted in the interface rather than of time. Thus the number of transformations within a predefined area was related to the cumulative number of surface contacts (including transformed and nontransformed particles; Fig. 4B). It was found that the number of transformations leveled off with increasing numbers of LBPs present in the interface. This suggests that LBPs inserted early into the air-liquid interface are exposed to high surface tension (= that of experimental solution) and thus transform more frequently than those arriving later when surface tension is probably low. Because it is difficult to obtain direct dynamic measurements of surface tension at the inverted interface, we decided to use solutions of Triton X-100, preadjusted to different surface tensions, to determine the percentage of transformations of LBPs. Importantly, at the concentrations used, Triton had no solubilizing effects on LBPs. The results (Fig. 4C) revealed that LBP transformations gradually decreased with decreasing surface tension and ceased below values 4050 mN/m. From these observations, we conclude that the surface tension is an important determinant for LBP transformation.
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DISCUSSION |
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Extracellular surfactant metabolism starts with release from type II cells (38). This process is different compared with release of soluble exocytotic products because a large, insoluble aggregate has to move through a narrow pore (16). During this process, the material undergoes considerable structural deformation (18). Our results obtained by fluorescence imaging of vital cells confirm these ultrastructural findings. It has been proposed that this deformation could be necessary for a later transformation into surface-active components (20) or even a self-decomposition of the material (29). However, despite striking differences in the physical appearance of surfactant during release (spherical and tubular), it still remains unclear whether structural changes at this early stage of extracellular metabolism are a prerequisite for rapid film formation.
Here we also show that, after release from the cells, surfactant presents as spherical LBPs that do not readily disperse when exposed to an aqueous environment over days. This observation indicates that LBPs are remarkably stable. Although this is in contrast to reported findings (28), it is in agreement with other investigators who reported that LBPs are regularly found as constituents of the alveolar lining fluid (9), in the ultraheavy and heavy fractions of bronchoalveolar lavage (11), and in abundance in fluid-filled premature lungs and amniotic fluid (23). In addition, evidence of a pronounced structural stability was presented recently, where we could demonstrate that LBPs even withstand strong mechanical deformation imposed by laser tweezers (31). Finally, dispersion of LBPs within a highly polar medium like water is not to be expected because of their predominantly amphiphilic composition.
However, at some point, LBPs have to undergo structural transformations to provide material for a surface-active film. Several studies showed that large surfactant aggregates can turn into small aggregates when exposed to surface area cycling by an end-over-end rotating tube (34). This method, initially introduced by Gross and Narine (12), is widely used to study the regulation of extracellular surfactant transformation in vitro. It is assumed that this particle conversion is enzymatically driven and requires interfacial manipulation (13). Another assumption is that it could be caused by shear forces resulting from turbulences within the fluid (15). Nevertheless, though, the basic mechanism(s) underlying these processes, in particular the sequence of structural transformations between release and insertion of LBPs in an air-liquid interface, and the forces involved are unclear (14, 15).
To investigate whether additional possibilities exist to explain these important structural transformations, we designed a method allowing direct visualization of LBP interaction with an air-liquid interface. It is based on a simple chamber with a small aperture that can easily be combined with any kind of inverted microscope. In particular, this method uses an inverted interface to trace the fate of single LBPs, freshly released from alveolar type II cells, in real time. In addition, turbulences within the chamber and at the interface complicating interpretation of experimental data are negligible. Furthermore, movement of film out of the microscopic field, which can occur in Langmuir troughs (21), is not a problem in this system. Thus this method could be ideally suited to study interfacial events with high-resolution optical techniques.
Using this method in combination with FM 1-43, we observed that LBPs approached the air-liquid interface as spherical particles and instantaneously disintegrated into large, rapidly expanding areas upon contact with it. The fluorophore FM 1-43 has a high affinity for lipids (30). Furthermore, its fluorescence intensity is considerably increased at lipid-water boundaries of which LBPs are highly equipped (1). Thus LBPs in the presence of FM 1-43 fluoresce intensely (17). Interestingly, when they hit the interface and transform into large areas, LBP fluorescence intensity decreased substantially. This likely indicates a loss of densely packed particle content and suggests a concomitant flow of this material on the interface. Additional considerations also suggest that the expanding areas are formed from LBPs: based on our results we calculate a mean phospholipid content of 0.24 x 10-6 µg/LBP. This amount (if pure dipalmitoylphosphatidylcholine) covers an area of 150 µm2, assuming a molecular area of
75 Å2 at a surface pressure of 10 mN/m (10). This value is in agreement with the areas observed in our system. Thus we reason that these areas emerging from transformed LBPs most likely correspond to lipids and/or hydrophobic surfactant proteins.
Another observation was that brightly stained LBPs instantaneously developed into dark areas when surface contact was established. This was somewhat unexpected, because dilute lipid films in the presence of FM 1-43 should still be more fluorescent than the background. At present, we do not have a definitive explanation for this phenomenon. A likely explanation is that the high concentration of FM 1-43 in the lipid bilayer(s) of LBPs is abruptly decreased as LBP content is distributed over a large surface area. As a result, the apparently dark areas then would contain less FM 1-43 than the surrounding surface. However, at this point, a final explanation will also have to await more detailed information on the internal molecular arrangement of LBPs.
A further observation was that LBPs split into smaller particles that eventually disappeared. It is known that lamellar bodies are quite complicated in architecture and can contain distinct lamellar whorls surrounded by a common envelope (24, 32). Perhaps, after surface contact, the envelope disrupts and the released lamellar whorls flow into the surface. Alternatively, individual LBPs could stick together and simply separate upon contact with the interface.
Finally, we observed that dark irregularly shaped material remained within fluorescent areas after transformation of LBPs, which did not further disintegrate. Proteins, depending on their degree of hydrophobicity, are stained by FM 1-43 to a much lesser degree than lipids. Whether this dark irregularly shaped material is a remnant of the dense matrix core, a material of essentially unknown but presumably proteinaceous composition found within lamellar bodies, remains unclear (24, 32, 37). However, this issue particularly needs further investigation, inasmuch as the release of proteins in the alveolar lining fluid is of considerable interest in lung function.
Interestingly, we found that numerous LBPs did not transform after reaching the interface. This was the case both when the surface was preoccupied with transformed LBPs and when surface tension of experimental solution was systematically lowered. In particular, LBPs sediment until one of them reaches the clean air-liquid interface (surface pressure 0 mN/m) and subsequently transforms. The transformation of that LBP indicates that it adheres to the water more strongly than its content coheres to itself (3) and is likely to be driven by mechanisms such as surface diffusion and surface-tension gradients (8). As a result, material from that LBP is deposited on the surface forming an initial film. This sequence of events is consistent with the two-step model for surfactant adsorption recently proposed by Walters et al. (35). Further LBP transformations cause deposition of additional material until surface pressure of film will eventually increase to equilibrium (2030 mN/m in this series of experiments). At this point, transformation of newly arriving LBPs comes to a halt. Consequently, they accumulate without structural change. Similar observations have been reported by Holm et al. (19), who found that surfactant extract was not inserted in the air-liquid interface when albumin films were present at a surface pressure of
20 mN/m, and by Hall et al. (15), who demonstrated that large surfactant particles were stabilized against conversion when the air-liquid interface was occupied by proteins. From these observations, we conclude that contact of LBPs with the air-liquid interface is an essential prerequisite for subsequent transformation. We further conclude that surface tension is the force that drives LBP transformation and that this event is self-regulated and stops when cohesive and tensile forces come to equilibrium.
The gradual decline of LBP transformations with an increased amount of adsorbed material agrees with many findings in the literature of surface balance studies. Usually, spreading stops when film pressure reaches so-called equilibrium spreading pressure, e.g., 45 mN/m (7, 10). Our measurements with suspended LBPs (Fig. 2, D and E) and with the inverted interface (Fig. 4C) suggest that a surface pressure of 2030 mN/m might already be the limit where spontaneous spreading of LBPs ceases. This difference to reported values could be explained by a deficiency in alveolar components like surfactant protein A (SP-A), which might be present in limiting amounts in our system (7).
It is assumed that LBPs, once released, feed their content in tubular myelin, which is then involved in film formation by promoting insertion of surface-active material in the air-liquid interface (9, 37). However, this concept is debatable, since SP-A knock-outs lack tubular myelin but have normal lung functions (22). In line with this, we could not find any evidence for the presence of tubular myelin in our system and conclude that LBPs are able to form a film by themselves. Perhaps the inverted interface, combined with advanced analytical methods, could be useful to study tubular myelin formation in a direct way.
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ACKNOWLEDGMENTS |
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Parts of this work were presented at the Federation of American Societies for Experimental Biology Summer Research Conference (Saxtons River, VT, 2002) and the Meeting of the German Physiological Sciences (Bochum, Germany, 2003).
GRANTS
This work was supported by Grants P15742 and P15743 [GenBank] from the Austrian Science Foundation, FWF, and by Grant 9640 from the Austrian National Bank.
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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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