Tracing surfactant transformation from cellular release to insertion into an air-liquid interface

T. Haller,1 P. Dietl,1 H. Stockner,1 M. Frick,1 N. Mair,1 I. Tinhofer,2 A. Ritsch,3 G. Enhorning,4 and G. Putz5

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


    ABSTRACT
 TOP
 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Pulmonary surfactant is secreted by alveolar type II cells as lipid-rich, densely packed lamellar body-like particles (LBPs). The particulate nature of released LBPs might be the result of structural and/or thermodynamic forces. Thus mechanisms must exist that promote their transformation into functional units. To further define these mechanisms, we developed methods to follow LBPs from their release by cultured cells to insertion in an air-liquid interface. When released, LBPs underwent structural transformation, but did not disperse, and typically preserved a spherical appearance for days. Nevertheless, they were able to modify surface tension and exhibited high surface activity when measured with a capillary surfactometer. When LBPs inserted in an air-liquid interface were analyzed by fluorescence imaging microscopy, they showed remarkable structural transformations. These events were instantaneous but came to a halt when the interface was already occupied by previously transformed material or when surface tension was already low. These results suggest that the driving force for LBP transformation is determined by cohesive and tensile forces acting on these particles. They further suggest that transformation of LBPs is a self-regulated interfacial process that most likely does not require structural intermediates or enzymatic activation.

alveolus; lamellar body; pulmonary; secretion; surface tension


LUNG SURFACTANT, THE ESSENTIAL biosynthetic product of alveolar type II cells, is stored in lamellar bodies before exocytotic release in the alveolar lining fluid (27). Once released, it exists in various distinct physical forms as follows: as densely packed lamellar body-like particles (LBPs), as multilayered packages, as lattice-like structures (tubular myelin), as oligolamellar vesicles, and as a thin lipid-protein film covering the air-liquid interface (9, 38). These forms are considered to be metabolically related and to represent different steps of conversion of the extracellular material. Thus LBPs are most likely the primary exocytotic product from the type II cells, and tubular myelin is suggested as an intermediary, but not obligatory, form feeding directly into the final film. The oligolamellar vesicles might represent spent surface material awaiting reuptake by type II cells, and the lipid-protein film is the functionally active state of lung surfactant (11, 37). However, the mechanisms promoting transformation of these distinctly different forms into surface-active components and the precise sequence of these events are still under debate (14, 15, 34). In particular, a surfactant convertase in conjunction with a cycling interface are the proposed mechanisms. Yet, no information is available about the fate of LBPs immediately after release by alveolar type II cells and their behavior when they finally reach an air-liquid interface.

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.


    METHODS
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 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Cell preparation and sampling of LBPs. Alveolar type II cells were isolated from Sprague-Dawley rats according to Dobbs et al. (4), with minor modifications (17). After isolation, cells were seeded on glass coverslips in petri dishes (Ø 35 mm), cultured in DMEM, and used the following day. For sampling LBPs, the adherent cells were washed and stimulated by ATP (100 µM) and phorbol 12-myristate 13-acetate (100 nM) in 2 ml of experimental solution (see below) supplemented with 5 mM glucose, 0.1 mg/ml streptomycin, and 100 U/ml penicillin. The secretagogues were given in combination to obtain a maximal stimulatory effect (6). After 3 h at 37°C, the cell culture dishes were shaken vigorously, and the supernatants were filtered through 8-µm cellulose nitrate membrane filters (Sartorius) to remove cell debris and nonadherent cells. The collected supernatants, containing freshly released LBPs, were immediately used, or they were frozen at -70°C in polypropylene tubes (Greiner). Pooled LBP suspensions were obtained by thawing and mixing frozen supernatants. Repeated freezing/thawing did not affect the LBP concentration of these samples. For the experiments in Fig. 2E, LBPs were stored at ambient temperature in experimental solution containing NaN3 (0.2 mg/ml). The storage was in either glass or Teflon vials, and the data from triplicate assays were pooled.



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Fig. 2. LBP release and surface tension. A: release from type II cells. Laser scanning microscope images demonstrating fused lamellar bodies (white spots) releasing their content as spherical (left) and tubular (center) protrusions. Released surfactant (right) appears as compact or as partially unraveled particles (= LBPs). Cell border is delineated by arrows at the presumptive site of the fusion pore. *Extracellular space. Bars = 5 µm. B: no. and size of LBPs released from stimulated cells. Time course of release followed a single exponential function with time constant ({tau}) = 122 min. Particle size increased within 1st h after stimulation. No. and mean particle size determined as described in Fig. 1A. au, Arbitrary units; n = 6 experiments. C: LBPs analyzed for phospholipid content. D: surface tension of supernatant from stimulated cell cultures measured as described in Fig. 1B. Surface tension was quickly lowered despite ongoing LBP release. Values at time 0 obtained from cell-free solutions; n = 6. E: no. and size of LBPs pooled from stimulated cell culture supernatants and corresponding surface tension during prolonged storage. LBP recovery was decreased on day 2 and remained constant thereafter. Mean size of LBPs remained unchanged. Surface tension was consistently lowered to ~52 mN/m and unaffected by initial drop in LBP recovery; n = 6. F: functional measurement of surface activity of LBPs in a capillary surfactometer. The capillary was open in 77% above ~4 x 107 LBPs/ml; n = 3.

 



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Fig. 1. Methods. A: lamellar body-like particle (LBP) detection: Aliquots of FM 1-43-labeled cell supernatants pumped through a micropipette (left). No. of passing LBPs (center) counted by automated peak analysis (right). Peak areas reflect particle size. PMT, photomulitplier tube. B: vertical pull method for surface tension measurement of cell culture supernatant. Maximum negative weight of a suspended fluid is used to calculate surface tension (2). C: schematic drawing of fluid-filled conical chamber with inverted air-liquid interface. The interface at bottom of chamber is inspected by bright-field or epifluorescence microscopy. Experimental solution inside chamber can be supplemented from top with either isolated LBPs or, as shown here, cultured cells. Freshly released LBPs are observed when approaching and inserting in the inverted interface. Ex, excitation; Em, emission. Dimensions not to scale.

 
Fluorescence staining protocol. LBPs and surface films were stained with FM 1-43 (Molecular Probes). As described (1), this styryl dye preferentially partitions into, but does not penetrate, lipid/water interfaces because of its hydrocarbon tail connected to a dicationic head group. The quantum efficiency of FM 1-43 is low in water but enhanced several hundredfold after insertion in lipids (1). Fluorescence of FM 1-43 was excited at 480 nm and emission monitored at >530 nm. For imaging exocytosis (see Fig. 2A) and LBP insertion in an air-liquid interface (see Fig. 3), FM 1-43 was used at 1 µM. At this concentration, the dye did not lower surface tension as measured by the vertical pull method (71.8 ± 0.3 mN/m). For LBP quantitation (Fig. 1A), FM 1-43 was used at higher concentration (40 µM) to increase the signal-to-noise ratio.



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Fig. 3. LBP interactions at air-liquid interface. A: cumulative incorporation of FM 1-43-labeled LBPs in inverted interface when observed at x20 magnification and 480 nm excitation. B–G: visualization of LBP transformations at x40 magnification. LBPs approaching the interface are out of focus until surface contact (*). B: single LBP transforming into a dark area, which successively brightens to fluorescence intensity above background but retains a dark structure (arrow). This LBP splits into two particles of gradually fading fluorescence. C: LBP entirely disappearing after contact. D: LBP transforming simultaneously into 2 particles and a bright area. E: transformation of LBP giving rise to a cycle of dark and bright areas. F: LBP splitting into several particles of fading fluorescence and lack of prominent area formation. Experiments B–F were performed in the presence of 1 µM FM 1-43. G: phase-contrast imaging of LBP demonstrating instantaneous and complete disappearance after surface contact. Intensity scalings individually adjusted to optimize image contrast. Bars = 20 µm.

 

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.


    RESULTS
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 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Surfactant transformation during release from stimulated cells. After intracellular lamellar bodies had fused with the plasma membrane, their content slowly protruded in the extracellular space. During this process, compression of the material was always apparent, as it squeezed out through the exocytotic fusion pore (Fig. 2A, left). Sometimes the released material attained a tubular structure resembling that of concentric, retractable telescope tubes (Fig. 2A, center). After complete release (= detachment from the cells), surfactant material was present as spherical and usually compact (>90%) or partially unraveled (<10%) particles (= LBPs; Fig. 2A, right). On rare occasions, we observed that LBPs adsorbed on the surface of the coverslip, forming disc-like structures. However, we never observed that LBPs, or parts of them, disintegrated when suspended in water. They always remained particulate, even showing distinct viscoelastic properties (31). From these observations, we conclude that structural transformations of LBPs take place at the earliest possible stage, during their exocytotic release from the cells.

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 ({tau}) 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 ~40–50 mN/m. From these observations, we conclude that the surface tension is an important determinant for LBP transformation.



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Fig. 4. Quantitation of LBP transformations at the inverted interface. A: cumulative appearance of LBPs over time. LBP suspensions (200 µl), collected from stimulated cell supernatants, were added to experimental solution (200 µl) containing FM 1-43. B: no. of transformed LBPs as a function of LBPs that arrived at the interface. Transformations were defined as sudden, distinct area formations or splitting of particles (cf. Fig. 3). On average, transformation was ~64% for the first 10 LBPs and strived toward 0% with ongoing sedimentation. C: LBP transformations as a function of surface tension. Chamber with inverted interface was filled with solutions of given surface tensions. No. of LBP transformations with regard to first 20 LBPs arriving in the interface were recorded. Transformations ceased at surface tensions below ~40–50 mN/m.

 


    DISCUSSION
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 ABSTRACT
 METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
In this paper, we demonstrate that surfactant, the prime secretory product of the alveolar epithelium, is transformed during release but then primarily forms spherical particles in extracellular solution. We also demonstrate that, if the surface area remains constant, only a fraction of released LBPs spontaneously transforms into a surface film, whereas the bulk remains stable. This latter transformation of LBPs, when directly observed by fluorescence microscopy at an air-liquid interface, was found to be an instantaneous self-regulated process driven by surface forces. Finally, when exposed to cycling area changes in a capillary surfactometer, we demonstrate that LBPs are able to generate a film exerting high surface pressure.

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 (~20–30 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 ~20–30 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.


    ACKNOWLEDGMENTS
 
We are grateful to Dr. J. A. Clements for critically reading the manuscript.

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.


    FOOTNOTES
 

Address for reprint requests and other correspondence: T. Haller, Dept. of Physiology, Univ. of Innsbruck, Fritz-Pregl-Str. 3, A-6020 Innsbruck, Austria (E-mail: thomas.haller{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.


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