Interfacial sensing by alveolar type II cells: a new concept in lung physiology?

Cell isolation and culture conditions.

Cell preparations were conducted in conformity with the Austrian rules for animal care and testing (a license from the Austrian Government has been granted to T. Haller). The AT II cells were isolated from male Sprague-Dawley rats according to standard protocols described elsewhere (12, 24). For the microplate experiments (Figs. 2 and 3), isolated cells were plated in sterile multiwell tissue culture plates, for the studies with the inverted interface (Fig. 4) onto Petri dishes, and for the experiments in Fig. 5 onto glass coverslips, all left for 24 h in DMEM supplemented with 100 U/ml penicillin, 100 μg/ml streptomycin, 24 mM NaHCO₃, and 10% FCS (Biochrom) in a humidified 5% CO₂ atmosphere of 37°C. Before the inverted interface experiments, cells were detached from the dishes by mild trypsinization (0.25%; 4 min), followed by centrifugation (1,000 rpm, 7 min) and resuspension in buffer.

Cytotoxicity.

Cells were seeded in 24-well plates (Greiner) and grown to confluence. They were gently washed three times with experimental solution (see below) and exposed to air for the respective times by aspiration of the fluid out of the wells, followed by readdition of 600 μl experimental solution at 25°C and a relative humidity (rH) of 50%. After 2 h incubation, supernatants were collected and stored at 4°C. Samples were analyzed for LDH release according to kit instructions (Roche). Each sample of 50 μl was transferred into a 96-well plate and mixed with 100 μl assay reagent. The reaction mixture was incubated for 15 min at room temperature under light protection. Absorbance was measured at λ = 492 nm with a microplate reader (GENios Plus, Tecan, Austria).

Exocytosis.

Cells grown in 96-wells (Sarstedt) were incubated with 1 μmol/l LysoTracker Green DND-26 (LTG) for 30 min at 37°C in DMEM. This water-soluble dye specifically, dose and time dependently, accumulates in lamellar bodies (i.e., secretory vesicles of AT II cells) where it is stably trapped by protonation, but discharged into the extracellular space once the fusion pore has formed (22, 24). After the 30-min incubation, cells were rinsed three times with experimental solution (see Solutions and reagents) to remove unattached cells and remaining dye. The cells were then exposed to air by sucking the entire fluid out of the wells, and left, for the indicated times, at room temperature in an atmosphere of constant relative humidity (50%) controlled by an electronic hygrometer. Experimental solution of 200 μl was readded to the cells, which was removed after 1 h and centrifuged for 3 min at 2,000 rpm. Supernatants were transferred into clean 96 wells and fluorescence of released LTG measured in a microplate reader (GENios Plus) with λ_exc = 485 nm and λ_em = 535 nm and multiple reads/well.

Phospholipid release.

Cell supernatants were collected and centrifuged as above and analyzed for phospholipid (PL) content using coupled enzymatic reactions as described (18). Briefly, supernatants were added to a buffered solution containing phospholipase D (1 U/ml), choline oxidase (0.2 U/ml), horseradish peroxidase (2 U/ml), and Amplex Red (0.1 mM), and resorufin formation was recorded with the plate reader at λ_exc = 540 nm, λ_em = 595 nm, and T = 37°C. End points of the kinetic reaction were taken after 90 min.

Ca²⁺ measurements.

Cells were preincubated with 4 μmol/l fura 2-AM for 20 min at 37°C in DMEM, washed twice and placed into a Tecan M200 microplate reader. After 5 min, measurements were started using λ_exc = 335/380 nm and λ_em = 510 ± 20 nm. After the recording of cytosolic Ca²⁺ concentrations ([Ca²⁺]_c) under submerged conditions, the plate was shortly moved out of the instrument, and the supernatants were gently removed.

Inverted interface setup and experimental procedure.

The inverted interface was previously used to analyze interfacial phenomena related with surfactant adsorption and surface film formation. For the details we thus refer to the following references: 7, 23, and 44. Briefly, a fluid is kept within a 200-μm spherically and sharply edged aperture of a sapphire cone, forming an essentially flat interface with the air below (Fig. 1A). To improve control over experimental conditions, the setup was modified as follows: A glass coverslip confined a space underneath the interface in which temperature and humidity could be controlled by a convective flow of air. Air flow was generated by a peristaltic pump connected to the inlet of the chamber. It delivered gases at rates (<2 ml/min) high enough to obtain rapid gas exchange but low enough to avoid perturbation of the interface. This was verified by fluorescent beads (2 μm) embedded at the interface showing no movements despite continuous gas-superfusion (not shown). Humidified (∼100% rH) and dehumidified (∼0% rH) air was produced by passing ambient air through either a water reservoir or a package of silica gel. In addition, housing of the interface and the entire stage of the microscope were thermostated (Tempcontrol 37, PeCon, Germany) yielding a measured temperature of 37 ± 0.1°C in the air and fluid sides of the interface, respectively. Before all experiments, the chamber was cleaned with water and acetone in an ultrasonic cleaner, thoroughly flushed with double-distilled water and dried with a stream of sterile air. After transfer to the microscope, the chamber was filled with 1 ml buffered solution, which immediately formed a “clean” (free of PL) I_AL at the aperture plane below. The aperture was aligned and focus adjusted to the interface (7) before the cells were added on top of the buffered solution. We usually applied 50 μl of a cell suspension containing 5–20 cells. Thus contact with an I_AL was accomplished by mere sedimentation of the cells toward that interface, circumventing any mechanical agitation. Costaining of cells (Fig. 4) was obtained by overnight incubation with DiOC (2 μM) and 10 min incubation with fura 2 (4 μM).

Dynamic interface.

A cylindrical rod (Ø 4 mm) of stainless steel was mounted on a micromanipulator (Narishige, Japan) and positioned ∼100 μm above the cells grown, at low density, on glass coverslips. By suction, buffer solution was removed from the cells except from those right underneath the rod, so to form a central remaining drop of fluid as depicted in Fig. 1C. By modulating the height and/or lateral position of the rod, the I_AL could be precisely moved while the cell(s) remained in focus. Interference resulting from light (550 nm) reflected by the interface and the upper surface of the glass (= reference beam) was used to calculate the actual thickness and steepness (slope) of the water front (see Interferometry).

Microscopy.

Details of the microscopic setup including the episcopic illumination are described (7, 44). Briefly, we used a Zeiss 100 inverted microscope (Zeiss) equipped with a monochromator (Polychrom II, TILL Photonics) and a cooled CCD camera (Imago-SVGA; TILL Photonics), both controlled by TillVision software. Dry objectives with long-working distances were used: A ×20 Plan-Neofluar numerical aperture (NA) 0.5 to image the aperture completely, a ×40 LD-Achroplan NA 0.6 for higher resolution imaging, and a Fluar ×20 NA 1.3 for the dynamic interface experiments (all objectives from Zeiss). For fluorescence, appropriate combinations of excitation wavelengths and filter sets were used. Fluorescence images are displayed in false colors.

Interferometry.

Interferometry was used to measure the topology (Fig. 4) and orientation (Fig. 5) of the I_AL. Optical path lengths were derived from interferograms that resulted from the superposition of light being reflected by the I_AL and a static reference plane. For the experiments involving static interfaces, such a reference plane was naturally provided by a reflective surface inside the objective (Fig. 1B). The static situation allowed the application of phase stepping, where the phase of the reference beam was repeatedly stepped by a known increment (25). This results in a set of fringe patterns that can be mathematically combined to extract the optical path length. For each situation, three interferograms were recorded with reference phase values of 0, 2π/3, and 4π/3 radians. The phase stepping was performed by controlled axial movements of the objective lens in the submicrometer range. To ensure a high accuracy of the objective movements, a special apparatus involving a level and a manual micrometer stage was attached to the microscope's focus knob. A He-Ne laser (633 nm) was used for these experiments.

The dynamic interface required an alternative method for fringe pattern evaluation. There, the thickness of the water layer was changed quickly by movements of the steel rod (Fig. 1C); hence, there was not enough time to perform phase stepping. The high density of fringes and the relatively smooth phase topography of the water layer, however, allowed determining the layer thickness from a single interferogram using Hilbert phase demodulation (28). From an obtained phase map, the mean slope of the I_AL was extracted. This parameter was used as an estimate for the thickness of the water layer that separates the cell from the surrounding air; i.e., the smaller the slope, the thinner is the remaining aqueous layer, and the higher the force exerted on the cell. Absolute slope was normalized between 1 (fringes not detectable: thickness of water layer exceeding the coherence length of the interfering light) and 0 (distance between single fringes approaching ∞: interface parallel to the glass coverslip). Normalization was necessary to eliminate differences in the lateral extension of the water drop (contact angle) together with differences in the heights of the cells and was done by developing own Matlab algorithms. A reduction in the normalized slope thus denotes a progressive steepening of the water front, no change denotes an interface that cannot be flattened further because of the resistance exerted by the cell(s) underneath. These measurements were combined with ratiometric fura 2 measurements as described above.

Solutions and reagents.

The standard experimental solution contained (in mM) 140 NaCl, 5 KCl, 1 MgCl₂, 2 CaCl₂, 5 glucose, and 10 HEPES (pH 7.4 at 25°C). The Amplex Red Phospholipase D assay kit, BAPTA-AM, BCECF-AM, 2-(4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene-3-dodecanoyl)-1-hexadecanoyl-sn-glycero-3-phosphocholine (BODIPY-PC), 3,3′-dioctadecyloxacarbocyanine perchlorate (DiOC₁₈), calcein-AM, FM 1–43, fura 2-AM, LysoTracker Green DND-26, and phorbol 12-myristate 13-acetate (PMA) were purchased from Invitrogen Molecular Probes (Austria); ATP, gadolinium, thapsigargin, EGTA, DMEM, IgG, phospholipase D, Triton X-100, and trypsin were from Sigma-Aldrich; FCS was from Biochrom; the LDH detection kit was from Roche; elastase (for the AT II cell preparation) was from Elastin Products; and PFD (perfluorodecalin) was from F2 Chemicals.

Statistic and image analysis.

Data are shown as means ± SE (unless differently specified). Image acquisition and first analysis was conducted using TillVision. Further analysis was done using ImageJ, Excel, and GraphPad. Interferometric analysis of the I_AL was performed by self-developed algorithms implemented in Matlab program. The algorithm preprocessed every fringe pattern by noise filtering and offset subtraction. Subsequently, the Hilbert transform of every image line was calculated and combined with the preprocessed fringe pattern to obtain a complex mathematical function (the analytic signal), the phase of which represents a map of the local water layer thickness.

Exposure to air is deleterious.

AT II cells, grown to confluence on a glass coverslip, were preincubated for 30 min with calcein-AM, and the coverslip was placed for 20 min in vertical position into buffered solution so that half of it was exposed to air. Afterwards, cells were rinsed and stained with FM 1–43. The result (Fig. 2A) demonstrated a loss in calcein (green) and a gain in FM 1–43 (red) in the air-exposed region. Since calcein is retained in intact cells and FM 1–43 enters damaged cells, the combination of both is an indicator of plasma membrane integrity and compellingly demonstrates the deleterious effect of air contact. However, this simple approach did not allow a reconstruction of the position of the I_AL nor a quantitative description of its effects, and thus has not been pursued further. Instead, we performed microplate experiments to analyze cell damage more quantitatively and on a time-resolved basis. LDH release in AT II cells was not measurable before 20 min air exposure (Fig. 2B). Notably, at this time, cells were already partially exsiccated (see Fig. 2C). Longer exposure times (40 min) increased LDH up to amounts comparable to those released by 1% Triton X-100 (equals 100% cytotoxicity). This proved, on a time-resolved basis, a similar extent of cell damage as shown before (Fig. 2A). Taking these findings together, we conclude that AT II tolerate air contact for more than 10 min after which cell membrane integrity is lost.

Exposure to air is a stimulus.

First indication of increased exocytosis (i.e., release of LTG through exocytotic fusion pores, see materials and methods) was at 30 s (Fig. 3A). This suggests that air contact stimulates the cells and is effective at a time when cell damage can be ruled out. It also suggests that cells respond before they may start to actually “dry out” (Fig. 2C). The profound increase in LTG release after cell damage (20 and 40 min) can be explained by a loss of plasma membrane integrity together with that of the limiting LB membrane with subsequent discharge of LTG into the cell exterior. This is also supported by the signals that considerably overrun those produced by ATP + PMA, two potent secretagogues that usually yield a high exocytotic response (53).

Similar with the time course of exocytosis was the release of PL (Fig. 3B). The first measurable increase was at 30 s, the highest (within the period of cell survival) at 10 min, and all values are in the range of response elicited by ATP + PMA. However, the time course of PL release appeared biphasic, with a return to control values at 2 and 5 min (the same tendency may be seen in exocytosis). A further difference is the massive LTG release at times 20 and 40 min, which is not reflected by a similarly pronounced increase in the amount of released PL. A likely explanation is the kind of membrane damage: it may be extensive enough to discharge the low-molecular-weight compound LTG out of lamellar bodies (and LDH out of the cytosol) but not extensive enough to release the bulky and essentially water-insoluble surfactant aggregates (∼2 μm) into the extracellular space.

The stimulus is Ca²⁺ dependent.

The exocytosis assay was repeated with BAPTA-loaded cells (Fig. 3C). The result revealed that in the absence of an air stimulus, constitutive secretion was identical in control and BAPTA-treated cells. After 1 min air exposure, however, BAPTA-treated cells exhibited a significantly reduced secretory response, suggesting a regulated exocytosis induced by contact with air. However, BAPTA was unable to block exocytosis completely and was more effective in ATP-stimulated cells. Both results suggest a significant contribution of an additional, Ca²⁺-independent path. Astonishingly, BAPTA also blocked exocytosis at time 40 min, when cell damage was considerable (Fig. 2B). The obvious Ca²⁺ dependence was further analyzed by continuous measurement of the [Ca²⁺]_c using fura 2-loaded cells (Fig. 3D). All measurements showed a small but significant [Ca²⁺]_c elevation immediately after suction of the fluid, probably responsible for the early initiation of exocytosis. A second, more sustained and pronounced [Ca²⁺]_c elevation occurred after ∼5 min, reaching values exceeding those of the Ca²⁺ agonist ATP (not shown). The source of the bulk Ca²⁺ change was clearly extracellular: Washout of Ca²⁺ (Ca²⁺-free buffer) before air exposure abolished the rise in the fura 2 ratio, similar to the effect of gadolinium in Ca²⁺-containing buffer. Furthermore, thapsigargin (tg), which depletes intracellular Ca²⁺ stores, had no inhibitory effect. Perfluorodecalin (PFD) was used in these experiments as a model for a liquid-liquid interface. It had no effect except the initial rise in [Ca²⁺]_c.

Microscopy of cell-interface interactions.

So far, the microplate experiments revealed a highly significant, time- and Ca²⁺-dependent exocytotic response of AT II cells to air contact followed by massive cell damage. But what are the mechanisms, and what are the signals? To approach these questions, we used a static, inverted I_AL (Fig. 1A) for the following reasons: 1) Cells, applied on top, sediment toward that interface and contact it with the least possible mechanical force (7). 2) Motions of cells abruptly stop when they reach the focal plane, allowing to precisely determine the instance of contact. 3) Cells can be imaged by brightfield, reflection, epifluorescence, and interference without focal shifts or loss of cells out of view. 4) Temperature (37°C), humidity (∼100 or ∼0%), and gas composition (ambient or N₂) in the air compartment could be adjusted. 5) Finally, and importantly, problems associated with osmotic changes can be ruled out because of a constant and large (1 ml) fluid volume in the chamber.

The results from the static interface experiments can be grouped into 3 classes: The first is no response (Fig. 4A) and this is when a cell gets close to an I_AL at 100% rH (scheme). The cell neither deforms nor penetrates it (interferometry), the intracellular Ca²⁺ does not change, there is no dye leakage out of the cells (fura 2 ratio), and exocytosis is not measurable (% DiOC). However, it may still proceed at a slow pace, as it leads to detectable amounts of PL surrounding the still intact cells after 24 h (Fig. 4D). The lack of an immediate Ca²⁺ signal suggests that interface contact per se is not sensed by the cells, and precludes nanometer-scaled electrochemical or osmotic gradients [which are described in thermodynamic models of the interface (32)], surface-associated gradients of respiratory gases (tested by perfusion with N₂, not shown), or the surface tension itself as reasons for an interfacial sensing. These experiments also demonstrate that immediate (0–10 min) and intimate (nm-μm) interface contact is neither a threat nor a stimulus, even when surface tension γ of the interface is unphysiologically high (∼70 mN/m).

The second is response (Fig. 4B and supplemental video S1 shown online at the AJP-Cell Physiol website). When the distance to the I_AL is further reduced by applying 0% rH, the intracellular Ca²⁺ rose quickly, and secretion was enhanced in most but not all cells. Despite the sustained [Ca²⁺]_c, which did not fully revert to precontact levels, there were no signs of cell damage. We assume that evaporative water loss forces cell structures, such as the glycocalyx and other macromolecules that extend into the extracellular space, into ultimate close distances to the interface, probably beyond a remaining hydration shell (20). This configuration is highlighted by the distortion of interference fringes at the site of a cell. When analyzed, this amounted to actual surface deformations in the range of several (∼70) nm. Thus, in configuration B, the cell surface is in obvious contact with the interface and probably subject to forces resulting from these interfacial deformations. The positive effect of 0% rH was additionally confirmed by changing rH (100% to 0%) during continued perfusion of the air compartment (practically, cells in condition A were subject to condition B; not shown). Theoretically, changing to 0% humidity could also have influenced local T at the utmost surface zone of the I_AL. However, the experiments were performed in thermal equilibrium with a high heat transfer to the chamber (thermostat plus air convection), and AT II cells did not respond to a sudden cooling down when tested independently, thus essentially ruling out that [Ca²⁺]_c signals were evoked by a local change in T.

The third is rupture (Fig. 4C and supplemental video S2). In seldom scenarios (4–5 cells) and never at 100% rH, we have seen an actual penetration of the interface followed by cell rupture and loss of cytosolic dye. We therefore conclude that penetration of the interface exposes the entire cell, or part of it, to the surface tension that ultimately leads to irreversible cell rupture. The main force acting here (besides gravitation and buoyancy as in Fig. 4, A and B), is surface tension with the lateral component of it being sufficient to tear a cell apart (cell rupture, however, was often seen with other cells, e.g., erythrocytes and monocytes, not shown).

So far, we have shown that intimate interface contact per se (Fig. 4A) is not the stimulus. On the other hand, surface penetration (Fig. 4C) is a deleterious event.

Dynamic interface.

The hypothesis remains that the sensory event is associated with a bending or a corrugation of the interface (provoked, e.g., by situation in Fig. 4B), exerting a mechanical load. To test this assumption further, we applied a dynamic interface model. With this setup, the I_AL, visualized by interferometry, could be precisely moved with respect to the cells while monitoring [Ca²⁺]_c (Figs. 1C, 5, 6, and supplemental video S3). The results principally confirmed those of the static measurements: AT II cells do not respond up to a certain distance to the interface (white symbols in Fig. 5, A and B). Onset of [Ca²⁺]_c increase (lines in Fig. 5A) was observed, however, when the interface started to bend over the cells (gray symbols). Moving the rod farther from the cells lead to [Ca²⁺]_c increase of different magnitudes or even to cell rupture in rare cases (n = 2), demonstrating that the bended interface exerts a force. Moving back the waterfront toward the cells (black symbols) resulted in [Ca²⁺]_c recovery (except after cell rupture). Finally, and most intriguingly, when the I_AL, kept at a minimum distance to the cells, was repetitively moved back and forth in small increments, the cells responded with out-of-phase [Ca²⁺]_c oscillations (Fig. 5, C and D; and supplemental video S3).