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Published ahead of print on February 14, 2008, doi:10.1165/rcmb.2007-0227OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 39, pp. 36-44, 2008
© 2008 American Thoracic Society
DOI: 10.1165/rcmb.2007-0227OC

Electrophysiological Characterization of Rat Type II Pneumocytes in Situ

Vadim Shlyonsky1, Arnaud Goolaerts1, Frédérique Mies1 and Robert Naeije1

1 Université Libre de Bruxelles, Brussels, Belgium

Correspondence and requests for reprints should be addressed to Dr. Shlyonsky Vadim, Université Libre de Bruxelles, Laboratoire de Physiologie et Physiopathologie, Campus Erasme, CP 604, 808, route de Lennik, 1070 Bruxelles, Belgium. E-mail: vshlyons{at}ulb.ac.be


   Abstract
Top
 Abstract
CLINICAL RELEVANCE
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
Optimal aeration of the lungs is dependent on an alveolar fluid clearance, a process that is governed by Na+ and Cl– transport. However, the specific contribution of various ion channels in different alveolar cell types under basal or stimulated conditions is not exactly known. We established a novel functional model of rat lung slices suitable for nystatin-perforated whole-cell patch-clamp experiments. Lung slices retained a majority of live cells for up to 72 hours. Type II pneumocytes in situ had a mean capacitance of 8.8 ± 2.5 pF and a resting membrane potential of –4.4 ± 1.9 mV. Bath replacement of Na+ with NMDG+ decreased inward whole-cell currents by 70%, 21% and 52% of which were sensitive to 10 µM and 1 mM of amiloride, respectively. Exposure of slices to 0.5 µM dexamethasone for 1 hour did not affect ion currents, while chronic exposure (0.5 µM, 24–72 h) induced an increase in both total Na+-entry currents and amiloride-sensitive currents. Under acute exposure to 100 µM cpt-cAMP, Type II cells in situ rapidly hyperpolarized by 25-30 mV, due to activation of whole-cell Cl–-currents sensitive to 0.1 mM of 5-Nitro-2-(3-phenylpropylamino)benzoic acid. In addition, in the presence of cpt-cAMP, total sodium currents and currents sensitive to 10 µM amiloride increased by 32% and 70%, respectively. Thus, in Type II pneumocytes in situ: (1) amiloride-sensitive sodium channels contribute to only half of total Na+-entry and are stimulated by chronic exposure to glucocorticoids; (2) acute increase in cellular cAMP content simultaneously stimulates the entry of Cl– and Na+ ions.

Key Words: alveolar epithelium • ion transport • amiloride • patch-clamp



   CLINICAL RELEVANCE
Top
 Abstract
 CLINICAL RELEVANCE
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
The findings described in this article contribute to understanding the ion transport function by lung alveolar cells in their native environment.

 
The efficiency of transepithelial water transport in alveoli is dependent on the expression and the activity of ion transport proteins, including amiloride-sensitive sodium channels (ENaC), nonspecific cation channels, cystic fibrosis transmembrane regulator (CFTR), and Na,K-ATPase (1–4). However, the specific contributions of ion channels to alveolar fluid clearance in different alveolar cell types under basal or stimulated conditions is not entirely clear. Alveolar type II cells play important role in normal lung function (1, 2). As complex nature of the lung architecture limits the accessibility to the alveolar epithelium, cultures of alveolar cells have been widely used as reliable in vitro model for the studies of ion transport (1, 3–6). However, both primary cultures of Type II cells and cell lines of alveolar epithelium derived from tumors or from immortalization techniques exhibit various features that differ from the differentiated phenotype of in vivo cells (4, 7). These include morphologic and biochemical changes, lack of native cell-to-cell communications, and low expression of tight-junction proteins (7), all of which can result in deviations from the characteristics of native cells (reviewed in Ref. 4). Two recent studies attempted to circumvent these shortcomings using lung slice preparations (8, 9). The authors succeeded in recording single ion channels from the apical membranes of Type I alveolar cells in situ. Their observations led to reconsider the role of Type I cells in alveolar ion and fluid absorption. In this report we re-evaluate the ion transport function of Type II cells in situ using a modified rat slice model that permits routine cell identification and whole-cell patch clamp measurements. Our results indicate that highly selective epithelial amiloride-sensitive channels are not the major pathways for Na+ entry in Type II pneumocytes from freshly prepared rat lung slices, while low-selective ENaCs mediate up to half of the Na+ entry. Consistent with other studies, in situ alveolar sodium transport is regulated by glucocorticoids. Our data also support the hypothesis that acute increase in cellular cAMP content stimulates simultaneous entry of chloride and sodium ions into Type II alveolar cells.


   MATERIALS AND METHODS
Top
 Abstract
 CLINICAL RELEVANCE
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 References
 
Reagents
Unless otherwise stated, all the reagents were from Sigma-Aldrich (Bornem, Belgium). Quinacrine, amiloride, acridine orange, propidium iodide, and trypsine stock solutions (x1,000) were prepared in water, dexamethasone in Me2SO.

Lung Slice Preparation
Lung slices were prepared using previously reported methods (8, 9) with modification. Male Wistar rats 7 to 12 weeks of age were used, as after this age, tight pipette seals were obtained less readily (our observations). All experiments were carried out with the approval of the local ethical committee and in accordance with the Guidelines for Animal Care. The animals were deeply anaesthetized by an intraperitoneal sodium pentobarbital injection (10 mg/kg of body weight), then secured on preparation table. After tracheotomy, the trachea was cannulated with a 1- to 2-mm tube. The peritoneal cavity was opened and inferior vena cava and renal artery were cut to exsanguinate the animal. The chest was opened and blood was removed from pulmonary circulation by perfusion with physiological solution via pulmonary artery. The lungs were then inflated to a submaximal capacity with warm (35°C) 6% gelatin (300 bloom, cell culture tested) prepared in serum-free RPMI media. The excised lungs were chilled in ice-cold physiological solution containing (in mM) 144 NaCl, 5 KCl, 10 HEPES, 1 CaCl2, 1 MgCl2, 15 D-glucose, pH 7.3, and 300 mOsm to solidify the gelatin. A small transverse block of tissue was separated from the left lobe of the lung and mounted onto a VT1000S vibratome (Leica Microsystems, Bensheim, Germany) filled with ice-cold physiological solution, and transverse lung slices (200–250 µm) were cut. Before microscopic examination and electrophysiological recording, individual slices were kept for up to 4 hours in chilled serum-free RPMI 1640 media containing 0.5 µM dexamethasone and 100 µg/ml of antibiotic Primocin (Invitrogen, Merelbeke, Belgium), and immediately before experimentation were placed between a pair of 1- x 1-cm HAPW polycarbonate filters (pore diameter 0.45 µm; VWR International, Leuven, Belgium) with a pre-cut aperture of 1 x 2 mm and washed three times with warm physiological solution to melt and remove the gelatin within the aperture. Alternatively, after sectioning, individual slices were fixed between polycarbonate filters as described above and were placed on whatman paper saturated with RPMI 1640 media as described above and kept at 37°C in a humidified incubator gassed with 5% CO2/95% air for a minimum of 1 hour and a maximum of 7 days.

Cell Identification
Microscopy cell viability was assessed using differential staining of healthy undamaged cells by acridine orange (AO) and staining of damaged, late apoptotic, and necrotic cells by propidium iodide (PI) (10). AO readily enters living cells and is accumulated by normal nuclei, where it emits green fluorescence in blue light illumination. PI, being bound to nucleic acids in fluorescence blue light, has red emission. Lung tissue slices were treated with 0.1 µM of AO for 10 minutes at 37°C and with 0.1 µM of PI for 5 minutes at 37°C and washed three times with physiological solution before subsequent imaging.

Rat Type II pneumocytes within the slices were identified on the basis of quinacrine (QA) uptake. QA is a fluorescent vital dye, which was previously used to monitor formation of lamellar bodies and surfactant production in cultured alveolar epithelial cells (11, 12). It should be noted that bronchoalveolar Clara cells, which also produce surfactant, become quinacrine-positive within the slices as well. However, if one considers that rat Type II cells constitute 10 to 16% of alveolar cells (13) and Clara cells constitute 25% of rat bronchiolar cells (14), while the number of alveolar cells is approximately 18 times higher than that of bronchiolar cells (15), then under our conditions only 1 out of 10 QA-positive cells within the rat slice is of Clara cell type. Slices were incubated with 0.1 µM quinacrine for 20 to 30 minutes at 37°C before microscopical examination. For routine Type II cell identification for electrophysiological recordings, 0.05 µM of quinacrine was added to gelatin used to inflate the lung during isolation procedure. In order to confirm that Type II cells are not subject to damage and/or necrosis, rat slices treated with quinacrine were incubated with 0.1 µM of PI for 5 minutes before subsequent imaging. In some experiments quinacrine was replaced by the vital dye Nile red to identify Type II cells. In those cases, slices were incubated 5 minutes in physiological solution containing 0.1 µg/ml of Nile red (1,000-fold dilution of a stock solution prepared in ethanol).

Microscopy
Differential interference contrast and fluorescence-microscopy images were obtained using an Axioplan Zeiss microscope (Carl Zeiss, Zaventem, Belgium) with a x20 or x40 objective. Illumination for epifluorescence was achieved with a 100-W Xenon lamp. A long-pass fluorescence blue filter set was used for simultaneous visualizing of green and red fluorescence. For cell counting, images were taken with x20 objective and green and red epifluorescence was visualized using standard FITS and TRITS filter sets, respectively. Images were acquired with a charge-coupled device camera (Zeiss AxioCam) and stored on computer disk using Zeiss AxioCam software. Field view area was measured by means of objective micrometer (Swift) and was equal to 430 µm x 345 µm. Number of cells on images was counted independently by two persons and a mean value was then calculated.

Electrophysiological Recordings
Nystatin-perforated whole-cell patch-clamp experiments were conducted as described previously (12, 16). The choice of nystatin-perforated whole-cell configuration instead of conventional whole-cell configuration was dictated by a 100% loss of sealing during the procedure of mechanical patch rupture. Briefly, polycarbonate filters holding tissue slices as described above were placed in a chamber on an inverted microscope and perfused with physiological solution described in lung slice preparation procedure. In low Na bathing media, 134 mM of NaCl was replaced with 134 mM of N-Methyl-D-Glucamine (NMDG), pH = 7.3 adjusted with HCl (final [Cl] = 135 mM). For whole-cell measurements, the pipette solution contained (in mM): NaCl 10, KCl 10, K-gluconate 130, HEPES 10, D-glucose 15, pH = 7.2 adjusted with KOH (final [K] = 144 mM); the osmolarity was 300 mOsm. Stock solution of nystatin (40 mg/ml in DMSO) was prepared daily. The patch pipettes were double-step pulled from borosilicate glass capillaries (Hilgenberg, Malsfeld, Germany) using a vertical puller (PB-7; Narishige International, London, UK). Filled pipettes had resistances of 5-8 MOhms. Seals with at least 5 GOhms in cell-attached configuration were selected for further studies. Whole-cell patch-clamp configuration induced by nystatin permeabilization was achieved within 15 to 20 minutes at room temperature. Access resistance (Rs < 35 MOhms) was stable for at least 20 minutes after stabilization and it was compensated by 70%. Despite the relatively high access resistance, the use highly resistive pipettes for nystatin-perforated patches (5–8 MOhms) allowed for fair mechanical stability of the patches during perfusion changes. Indeed, the majority of seals obtained with pipettes with resistances of 2 to 4 MOhms was lost during perfusion changes, even though the transition was exceptionally smooth. Slices were used within 60 minutes after being taken from the incubator. The holding potential was 0 mV. Voltage ramp protocols consisted of a –100 mV step (200 ms in duration) followed by a ramp ranging from –100 mV to +100 mV over 1 seconds. Ramps were applied every 14 seconds. Values of whole-cell currents at chosen voltages were obtained by points averaging on the I/V curve within a 5-mV window around this voltage. Resting membrane voltages were taken from the amplifier readings in whole-cell zero current-clamp mode.

Statistical Analysis
Cell capacitance and cell membrane resting potential data are presented as box plots. Upper and lower hinges (quadrilles), interquadrile range (IQR), and medians were calculated using Microsoft Excel (Microsoft, Redmond, WA). Mild outliers situate 1.5 to 3 IQR away from the hinge, and extreme outliers are more than 3 IQR away from hinges. Ion current data are expressed as mean ± SEM. Paired or nonpaired two-tailed t tests were performed when appropriate. In all comparisons, a P < 0.05 was considered significant.


   RESULTS
Top
 Abstract
 CLINICAL RELEVANCE
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 References
 
Lung Slices Keep Intact Morphology and Retain Live Type II Cells
Explants of rat lung tissue produced by slicing gelatin-inflated lung kept intact alveoli, as shown by differential interference contrast microscopy (Figure 1A). Figure 1B represents cell staining with a combination of live/dead dyes, AO, and PI. It shows that lung slices also retain a majority of live cells. Some apoptotic and/or damaged cells appear to have some remainder of double-stranded nucleic acids that could be stained by AO, and this fact results in co-localization of green and red fluorescence in these cells (Figure 1B). The same image depicts bright yellow granules around green nucleus that presumably represent lamellar bodies in Type II alveolar cells.


Figure 1
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  		Figure 1. Cell identification and evolution of cell numbers in rat lung slices. Differential interference contrast (A and C) and corresponding epifluorescence (B and D) images of nonfixed rat lung slices. (A and B) Focus within the plane of a slice treated with a combination of live/dead stains, acridine orange (AO) (green fluorescence) and propidium iodide (PI) (red fluorescence), respectively. (C and D) Focus on the surface of a slice treated with a combination of quinacrine (QA), lamellar bodies marker (green fluorescence), and PI, dead cell marker (red fluorescence). Bar = 20 µm. Gamma correction was applied to B and D to enhance the signal of red fluorescence. White arrows indicate Type II cells situating in the corners of alveoli on the surface of the slice. These observations suggest that lung slices keep intact morphology and retain live Type II cells. (E and F) Evolution of the numbers of cells with (E) lamellar bodies and (F) dead cells in rat lung slices in culture. Slices were cultured in the absence (open circles) and in the presence of 0.5 µM dexamethasone (solid circles). Dexamethasone supports normal appearance of lamellar bodies of Type II cells, which remain accessible in its presence for up to 72 hours in culture. Each data point represents a mean of 36 view field counts from 18 slices isolated from four different animals. #P < 0.05 versus time 0 until the end of observation, nonpaired t test, *P < 0.05 versus corresponding control condition, nonpaired t test.

 
Lamellar bodies in Type II pneumocytes in slices were specifically identified using fluorescent vital dye QA, which was previously used to monitor surfactant production in ATII cultured cells (11, 12). Figure 1D depicts fluorescent green inclusions in Type II cells resulting from accumulation of quinacrine by lamellar bodies. The corresponding differential interference contrast image focused on the surface of the slice is shown in Figure 1C. These representative pictures show that each slice presents Type II cells on its surface, and these cells are easily accessible (Figures 1C and 1D, white arrows). Another important observation is that QA-positive cells never co-localized with PI-positive nucleus of dead cells. Incubation of slices with 0.1 µM QA did not result in increased cell mortality. The number of dead cells per field of view were 19.0 ± 5.1 (n = 19) and 21.1 ± 4.8 (n = 36) in the absence and in the presence of quinacrine, respectively (P > 0.05, nonpaired t test), confirming that quinacrine is a safe staining agent of cells with lamellar bodies.

The number of Type II cells and of dead cells in slices was monitored for up to 1 week in culture. Slices were cultured in control media and in the presence of 0.5 µM dexamethasone. The number of Type II cells was stable for 24 hours after slice preparation, which was followed by a significant decrease at 48 hours (Figure 1E). After 72 hours, the number of QA-positive cells in slices cultured in control media decreased to less than 15% of initial counts, and it was significantly lower compared with slices cultured in the presence of dexamethasone (~ 40%; Figure 1E). On the other hand, dexamethasone treatment slightly affected significantly cell viability throughout the observation period (Figure 1F). Under both control and dexamethasone conditions, number of dead cells remained stable for up to 72 hours, indicating the absence of necrotic process. After 72 hours cell mortality increased gradually under both conditions (Figure 1F).

Thus (1) lung tissue slicing neither damaged cells nor induced immediate necrosis; (2) dexamethasone supported normal appearance of lamellar bodies of Type II cells identified by quinacrine; and (3) live quinacrine-positive Type II cells remained accessible for patch-clamp examination for up to 72 hours in culture.

Basic Electrical Properties of Type II Cells In Situ
We used the nystatine-perforated whole-cell patch clamp technique to examine basic electrical properties of Type II cells in situ—cell capacitance, cell membrane resting potential, and membrane resistance. Capacitance of Type II cells from freshly prepared lung slices varied between 5 and 26 pF; however actual data values were not normally distributed. In fact, 90% of values fell in the range of 5 to 15 pF, with a median of 8.5 pF (Figure 2A). Three outlier points seen above the first box in Figure 2A presumably represent contamination of measured values from the capacitance of Clara cells, which usually have a larger size than Type II pneumocytes. This 10% cell contamination (3 out of 28) correlates with our estimations of the ratio between Clara cells and Type II pneumocytes in rat lungs (see MATERIALS AND METHODS). The same box plot analysis shows that cells with a capacitance equal or higher 16.5 pF (upper hinge plus 1.5 interquadrile range) do not belong to the Type II alveolar cells. Thus, we excluded these cells from the analysis of the properties of Type II pneumocytes in situ in freshly prepared lung slices. Mean capacitance values after exclusion of contaminating cells was 8.8 ± 2.5 pF (n = 25). Values obtained for Type II cells cultured for 1 hour in the presence of dexamethasone were identical, Cm = 8.3 ± 2.5 pF, n = 22. We monitored electrical capacitance of cells for up to 3 days in the presence of dexamethasone (Figure 2A). After 24 hours of culture we observed a slight decrease of the median value in Type II cell capacitance (7 pF versus 8.5 pF), with no change in the mean value (Cm = 8.3 ± 3.7 pF, n = 13, P = 0.1 versus Day 0, nonpaired t test). After 72 hours we observed a significant increase in cell capacitance (Cm = 17.3 ± 7.9 pF, n = 8, P = 0.03 versus Day 0, nonpaired t test). One can note that at 72 hours cell capacitance varied in a wide range (7–28 pF) with near-normal distribution of data points. Thus, we conclude that this capacitance increase represents a real phenomenon and accordingly we could not exclude at this stage cells with higher capacitance.


Figure 2
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  		Figure 2. Evolution of cell capacitance (A) and resting membrane voltage (B) of Type II cells in rat lung slices in the presence of dexamethasone. Box plots were constructed as explained in MATERIALS AND METHODS. Mild outlier points are indicated as open symbols, and extreme outliers as solid symbols. Slices were cultured in the presence of 0.5 µM dexamethasone. Changes in cell capacitance suggest transdifferentiation of Type II cells into Type I cells. *P < 0.04 versus time 0, nonpaired t test.

 
Several studies report the presence of connexins and functional dye coupling between alveolar epithelial cells (17, 18). Gap junctions allow electrical communication between cells, and if current passes between cells via connexins, this would provide an overestimation of cell capacitance values. To test for electrical coupling between cells we measured cell capacitance in the presence of heptanol, a gap junction inhibitor. After 5 minutes of exposure to heptanol (1 mM), capacitance decreased from 8.4 ± 3.6 pF to 7.8 ± 3.1 pF (n = 5, P = 0.08, paired t test). Thus, it appears that alveolar Type II cells in freshly prepared lung slices are not electrically coupled or coupled in a very limited way. We have also tested whether the increase in cell capacitance after 72 hours culture in the presence of dexamethasone was due to establishment of strong cell-to-cell communications. In the presence of 1 mM heptanol, capacitance of these cells decreased slightly from 18.1 ± 8.3 pF to 16.2 ± 7.6 pF (n = 4, P > 0.05, paired t test). Thus, it is clear that doubling of cell capacitance at 72 hours cannot be explained by cell coupling that does not exceed 10%.

Mean membrane resistance of Type II cells in situ determined in response to 5-mV step from 0 mV holding voltage was 0.99 ± 0.34 GOhms (n = 5). It decreased significantly at 72 hours of culture to 0.52 ± 0.15 GOhms (n = 4, P = 0.037, nonpaired t test).

In zero current clamp mode, resting membrane voltage of these cells varied from –12 mV to +8 mV with a mean Vm of –4.4 ± 1.9 mV (n = 8). There was no statistical difference in Vm values between cells cultured in the presence and in the absence of dexamethasone as well as throughout the observation period in the presence of dexamethasone (figure 2B).

Whole-Cell Voltage and Current Clamp Studies
The fluorescent dye QA that we used for cell identification is known to be a nonselective inhibitor of potassium channels, ACh channels, and prion channels at concentrations of 30 to 50 µM and also acts as a mitochondrial uncoupler at micromolar concentrations. Although the concentration used in our study to identify lamellar bodies in Type II cells is at least two orders of magnitude lower than those producing an inhibitory effect, we conducted control experiments, to assess whether QA affects whole-cell currents in A549 cells, a well-established cellular model of alveolar ion transport. QA at concentrations up to 20 µM did not have any effect (n = 3, data not shown), consistent with previously published data (19).

Since perforated whole-cell configuration is characterized by high access resistance (Ra), we evaluated whether high Ra introduces limitations in our experiments. First, high Ra causes a current-dependent voltage error in the I/V relationships. The highest value of Ra in our study was 10.5 MOhms (35 MOhms compensated by 70%). Therefore, 100 to 200 pA (maximal currents observed in our study) would result in a voltage error of only 1 to 2 mV. The second limitation of high Ra is a slower exponential settling of steady-state values of holding membrane voltages and currents in the circuit. According to the maximal values of Ra (10.5 MOhms) and Cm (26 pF), the time constant of this process is 273 microseconds, which should not interfere with the much slower sodium and potassium currents. The voltage ramps used in our study are the result of 400 rising steps of 0.5 mV in amplitude each and 2.5 milliseconds in duration. The calculations show that for at least 1.2 milliseconds at the end of each mini-step, membrane holding voltage value is 99% of that of the command voltage. Thus, we conclude that at the Ra observed in our study, applied voltage ramps reflect in a fair steady-state membrane voltage settling and ion current readings.

Representative whole-cell currents obtained in Type II pneumocytes in situ in response to voltage ramps from –100 mV to +100 mV are shown in Figure 3A. In normal physiological solution, QA-positive Type II cells in slices displayed outwardly rectifying currents at positive voltages (Figure 3A, curve 1). Addition of 10 µM amiloride induced small inhibition in inward currents (Figure 3A, curve 2), while addition of 1 mM amiloride blocked both inward and outward currents, consistent with the inhibition of nonselective ENaCs (Figure 3A, curve 3). There was no difference in the magnitude of amiloride effect in cells investigated directly after slicing and those kept in the incubator at 37°C for 1 hour with and without dexamethasone, hence these data were pooled. Whole-cell currents averaged between –95 and –90 mV (I-95mV) are summarized in Figure 2B. Currents decreased from –3.46 ± 0.45 pA/pF to –3.02 ± 0.50 pA/pF in the presence of 10 µM of amiloride (n = 6, P = 0.018, paired t test) and to –1.91 ± 0.50 pA/pF in the presence of 1 mM of amiloride (n = 6, P = 0.002, paired t test). Replacement of Na+ in the bath with a non-permeant cation N-methyl-D-glucamine (NMDG+) decreased inward current to –1.08 ± 0.18 pA/pF (Figure 3A, curve 4, and Figure 3B; n = 6, P = 0.001 versus control, paired t test). In zero current clamp experiments, cell membrane resting potential hyperpolarized from –4.3 ± 2.1 mV to –5.2 ± 1.6 mV in the presence of 10 µM of amiloride (n = 8, P = 0.047 paired t test) and to –18.2 ± 3.7 mV in the low Na+ bath (n = 8, P = 0.008 versus control, paired t test).


Figure 3
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  		Figure 3. Electrophysiological data from Type II cells in rat lung slices. (A) Representative whole-cell voltage ramp current records. Curves: 1, currents in normal bath; 2, currents in the presence of 10 µM of amiloride; 3, currents in the presence of 1 mM of amiloride; 4, currents in low-sodium media. Shown as an insert: voltage ramp protocol ranged from –100 mV to +100 mV over 1 second. Ramps were applied every 15 seconds. (B) Summary of inhibition of inward whole-cell currents. Bars represent means of six experiments. Inward currents were averaged from whole-cell current voltage ramp records between –95 mV and –90 mV. *P < 0.05 versus control, paired t test, **P = 0.003 versus control, paired t test. (C) Representative whole-cell voltage ramp current records. Curves: 1, currents in normal bath; 2, currents in the presence of 1 mM of LaCl3; 3, currents in low-sodium media. Shown as an insert: voltage ramp protocol. (D) Pharmacologic profile of inward whole-cell currents. Bars represent means of four to six experiments. Effects of inhibitors were normalized to the effect of sodium bath replacement set as 100%. (E) Representative whole-cell voltage ramp current records. Curves: 1, currents in normal bath; 2, currents in the presence of 10 mM of TEA. Shown as an insert: voltage ramp protocol. (F) Summary of inhibition of outward whole-cell currents. Bars represent means of four experiments. Outward currents were averaged from whole-cell current voltage ramp records between +65 mV and +70 mV. *P < 0.05 versus control, paired t test.

 
We have recently shown the sensitivity of whole-cell sodium currents in alveolar cells to trivalent cation La3+ (12), a broad inhibitor of nonspecific cation channels. Figure 3C pictures the representative whole-cell current records showing inhibitory effect of 1 mM LaCl3 compared with the replacement of Na+ with NMDG+. The pharmacological profile of inward sodium current sensitivity to amiloride and lanthanum, normalized to the effect of sodium bath replacement is shown in Figure 3D. These results indicate that in freshly prepared lung slices, 21%, 29%, and 52% of total inward sodium currents in Type II pneumocytes can be inhibited by 10 µM, 100µM, and 1 mM of amiloride, respectively, while 1 mM of La3+ blocks up to 75% of these currents.

Application of the nonspecific potassium channels inhibitor, tetraethylammonium (TEA), did not have any effect on inward current but inhibited 30% of outward currents, indicating K-channel activity in Type II cells in situ (Figures 3E and 3F). Currents averaged between +70mV and +75 mV (I+70mv) decreased from 5.73 ± 1.20 pA/pF to 3.95 ± 1.19 pA/pF (n = 9, P = 0.03, paired t test).

Total sodium currents and amiloride-sensitive currents in Type II cells cultured in the absence of dexamethasone were stable for up to 48 hours of culture, at which point it became difficult to the cellular type (n = 4 and n = 3 for 24 h and 48 h, respectively, data not shown). In contrast, in the presence of 0.5 µM of dexamethasone, total sodium currents increased after 24 hours, while there was a non significant trend for amiloride-sensitive currents to increase as well (Figure 4). After 72 hours of culture, both types of currents were further elevated and amiloride-sensitive currents became significantly different from those measured on Day 0 (Figure 4A). We have tested the effect of heptanol on whole-cell currents. The results suggest that it does not affect ion currents in cells in freshly prepared lung slices (n = 3, data not shown). In contrast, in cells cultured in the presence of 0.5 µM dexamethasone for 72 hours, heptanol inhibited 25.2 ± 4.6% of inward sodium currents (n = 3, Figure 4B). We have repeated the experiments in single A549 cells chronically treated with 0.5 µM dexamethasone and we have obtained similar inhibition by heptanol (n = 4, data not shown), which indicates that this effect is not due to disruption of cell coupling. Together with the lack of strong cell coupling, these results suggest that the change in heptanol sensitivity of whole-cell currents is related to the shift in the properties of amiloride-sensitive sodium channels in the presence of dexamethasone, most likely due to the increase in the number of highly selective ENaCs sensitive to submicromolar concentrations of amiloride (20).


Figure 4
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  		Figure 4. Evolution of inward sodium currents of Type II cells in rat lung slices in the presence of dexamethasone. (A) Slices were cultured in the presence of 0.5 µM dexamethasone. Current density inhibited by 10 µM amiloride (solid triangles), current density inhibited by low Na+ bath (solid squares), and total inward currents (open circles) were measured in at least four cells. *P < 0.05 versus time 0, nonpaired t test; **P = 0.001 versus time 0, nonpaired t test. (B) Representative whole-cell voltage ramp current records in cells cultured for 72 hours in the presence of 0.5 µM dexamethasone. Curves: 1, currents in normal bath; 2, currents in the presence of 1 mM of heptanol; 3, currents in the presence of 10 µM of amiloride; 4, currents in low-sodium media. Shown as an insert: voltage ramp protocol ranged from –100 mV to +100 mV over 1 second. Ramps were applied every 15 seconds.

 
To test the functional response of Type II cells in situ we used cpt-cAMP, a cell-permeable analog of cyclic-AMP. Cyclic AMP is a known modulator of ion currents in alveolar cells (21–23). Representative whole-cell records are shown in Figure 5A. Within 5 minutes of cpt-cAMP addition, we observed an increase in whole-cell currents, whose reversal potential shifted to more negative voltages compared with control (Figure 5A, curves 1 and 2). Hyperpolarization of cells was confirmed in zero current clamp experiments, Vm changed from –1.5 ± 5.2 mV to –31.1 ± 1.1 mV (n = 3, P = 0.005, paired t test). In the presence of cpt-cAMP, the cell membrane resting potential tended to approach the equilibrium potential for chloride ions under the ionic conditions used in our study (ECl = –50 mV). This observation suggests that the main effect of cAMP is on chloride transport pathways; however, other ion transport pathways (possibly sodium) are activated as well, and this explains the deviation of the cell membrane resting potential from the equilibrium potential for chloride ions. As can be seen on curve 3 of Figure 5A, whole-cell currents activated by cpt-cAMP were largely inhibited by 5-Nitro-2-(3-phenylpropylamino) benzoic acid (NPPB), a blocker of Cl-channels, including cAMP-dependent Cl-channel, CFTR. Whole-cell currents at the exclusion of sodium currents averaged between 65 mV and 70 mV (the range of the equilibrium sodium potential for the ionic conditions used, ENa = 68 mV) are shown in Figure 5B. The figure depicts a modest but significant inhibitory effect of NPPB under control conditions, while outward whole-cell currents stimulated by cpt-cAMP could be largely suppressed by this drug.


Figure 5
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  		Figure 5. Effect of cAMP on whole-cell currents of Type II cells in rat lung slices. (A) Representative whole-cell voltage ramp current records. Curves: 1, currents in normal bath; 2, currents in the presence of 100 µM of cpt-cAMP; 3, currents in the presence of 100 µM of cpt-cAMP and 100 µM of 5-Nitro-2-(3-phenylpropylamino) benzoic acid (NPPB). Shown as an insert: voltage ramp protocol ranged from –100 mV to +100 mV over 1 second. Ramps were applied every 15 seconds. (B) Summary of whole-cell currents. Data points represent means of four experiments. In order to exclude the contribution of sodium currents, data points were obtained by averaging whole-cell current voltage ramp records near the equilibrium sodium potential (between +65 mV and +70 mV). *P < 0.04 versus control, paired t test, **P = 0.005 versus control, paired t test, §P = 0.006 versus cpt-cAMP alone, paired t test.

 
The curve 3 in Figure 5A shows that inward currents remain elevated compared with control after inhibition of chloride channels by NPPB, and these residual inward currents reversed at voltages close to zero millivolts. This observation together with the deviation of cell membrane resting potential in the presence of cAMP from the equilibrium chloride potential suggests that cAMP also activates cation transport pathways. We thus explored further this observation and compared the effects of amiloride and of sodium substitution in the absence and in the presence of cAMP activation. To study the effect of cAMP on sodium currents, whole-cell currents were analyzed at the voltage close to the value of the equilibrium chloride potential under ionic conditions used in our study (between –52 mV and –47 mV) in order to exclude the contribution of chloride currents. Figure 6A shows the effect of amiloride and sodium substitution with NMDG+ in a resting cell. Figure 6B shows activated whole-cell currents in the same cell after a 5-minute stimulation with 100 µM cpt-cAMP and the effects of amiloride and sodium replacement on these currents. A summary of 4 experiments is presented in Figure 6C. These results show that inward currents sensitive to sodium replacement are increased by 30% (open circles, P < 0.05) and this activation coincides with the rise in total whole-cell currents (filled squares, P < 0.05). Sodium currents sensitive to 10 µM amiloride are increased by 70% (open squares, P < 0.05).


Figure 6
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  		Figure 6. Effect of cAMP on inward sodium whole-cell currents of Type II cells in rat lung slices. (A) Representative whole-cell voltage ramp current records. Curves: 1, currents in normal bath; 2, currents in the presence of 10 µM of amiloride; 3, currents in low Na bath. Shown as an insert: voltage ramp protocol ranged from –100 mV to +100 mV over 1 second. Ramps were applied every 15 seconds. (B) Whole-cell voltage ramp current records from cell shown in A after cAMP stimulation. Curves: 1, currents in normal bath; 2, currents in the presence of 10 µM of amiloride; 3, currents in low-sodium bath. Shown as an insert: voltage ramp protocol. (C) Summary of whole-cell currents. Individual data points of total whole-cell currents, currents sensitive to 10 µM of amiloride and currents sensitive to sodium bath replacement with NMDG+ are shown. In order to exclude contribution of chloride whole-cell currents, currents were averaged from whole-cell current voltage ramp records near the equilibrium Cl– potential (between –47 mV and –52 mV). *P < 0.05 versus control, paired t test.

 
Taken together our results thus suggest simultaneous activation of sodium and chloride whole-cell currents after cell stimulation with a permeable analog of cAMP.


   DISCUSSION
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 Abstract
 CLINICAL RELEVANCE
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
References
 
The present results obtained on a novel functional lung slice model to study whole-cell currents in Type II pneumocytes in situ suggest that (1) sodium amiloride-sensitive channels (ENaC) constitute only 50% of sodium entry pathways in in situ Type II pneumocytes; (2) ENaCs are stimulated by chronic glucocorticoid treatment; and (3) an acute increase in cellular cAMP content stimulates entry of both chloride and sodium ions into in situ Type II alveolar cells.

Our results on lung slice morphology and cell viability are consistent with previous studies showing that precision-cut lung slices cultured in serum-free media maintain normal orientation of alveolar spaces (7–9, 24, 25). The slices also keep epithelial differentiation for at least 48 to 72 hours (24, 25). The advantage of the lung slice culture model described in the present report consists in the fact that lung alveoli are void of the agarose inflating agent used in previous studies. Agarose does not re-melt at 37°C and interferes with the patch-pipette sealing process (8). Instead, we used gelatin that easily dissipates in culture and leaves alveolar spaces accessible by the patch-clamp pipettes. In contrast to slicing noninflated collapsed lung described in Ref. 8, we show the advantage of lung inflation step in slice preparation procedure, and slice culture between a pair of polycarbonate filters. In our experience, the procedure described ensures near complete success in the rate of sealings.

Rat Type II pneumocytes within the slices were identified on the basis of QA uptake, fluorescent vital dye, which was previously used to monitor formation of lamellar bodies and surfactant production in cultured cells (11, 12). One limitation for the use of this dye is the fact that bronchoalveolar Clara cells, which also secrete surfactant, become QA-positive in lung slices. However, if one considers that rat Type II cells constitute 10 to 16% of alveolar cells (13) and Clara cells constitute 25% of rat bronchiolar cells (14), while the number of alveolar cells is approximately 18 times higher than that of bronchiolar cells (15), then under our conditions 90% of QA-positive cells within the rat slice are the alveolar Type II cells, a value comparable with the purity of freshly isolated cells (6, 21, 26–33). Moreover, if one chooses the cells situated in the corners of alveolar structures as shown in Figure 1, Clara cells can be completely avoided. As a substitute to quinacrine, in some experiments we used vital dye Nile red to identify Type II cells. However, red fluorescence arising from Nile red staining interferes with dead cell staining using PI, which produces also red signal.

We compared the electrical capacitance of the Type II cells in situ with that published for primary cultured Type II cells (Table 1). Mean values observed immediately after slice preparation (8–9 pF) were slightly higher than those published for freshly isolated Type II cells (6–7 pF; see Table 1). The difference could be accounted for by a slight coupling of cells in situ; however, our experiments using the gap junction inhibitor heptanol did not suggest the existence of a strong cell coupling. It has been reported that the capacitance of Type II cells may increase by more than 20% during lamellar bodies exocytosis (26). It has been also shown that Type II cells in culture have a tendency to become bigger with time and their capacitance at Days 4 to 7 after isolation increased by a factor of 1.5 to 2 (27–29). This phenomenon was confirmed for the cells in situ in the present study, and it was shown not to be related to the re-establishment of strong cell-to-cell communications. On the other hand, the gradual change in cell morphology including increase in cell capacitance and disappearance of lamellar bodies raises the possibility that the lamellar body–containing cells in slices cultured for several days represent intermediate cell type between ATII and ATI cells.


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  		TABLE 1. COMPARISON OF BASIC ELECTRICAL PROPERTIES OF TYPE II CELLS IN SITU AND FRESHLY ISOLATED CELLS

 
The magnitude of inward whole-cell currents measured in Type II pneumocytes in situ—3 to 4 pA/pF—was comparable to the value of 4-5 pA/pF obtained by Jiang and colleagues from freshly isolated Type II cells under similar conditions, i.e. in perforated whole-cell patch-clamp mode (29). It should be noted that in classical whole-cell configuration inward currents in freshly isolated Type II cells are reported to be significantly larger. The values vary within the range of 7 to 10 pA/pF (30, 31) and can be as high as 30 pA/pF (32). Our results indicate that highly selective epithelial amiloride-sensitive sodium channels sensitive to submicromolar concentrations of amiloride are not the major pathway for Na+ entry in Type II pneumocytes in freshly prepared rat lung slices. We observed that less than 13% of inward currents and approximately 20% of total sodium entry currents in these cells could be inhibited by 10 µM amiloride. In contrast, millimolar concentration of amiloride blocked both inward and outward currents, consistent with the inhibition of nonselective ENaCs (1, 5). We estimate that more than 50% of inward sodium currents were sensitive to 1 mM of amiloride. In addition, large multivalent cation La3+, a broad inhibitor of nonselective cation channels, blocked up to 75% of inward sodium currents (Figure 3). Great variability was reported for amiloride sensitivity of whole-cell currents in freshly isolated Type II pneumocytes. A review of the literature shows values varying from near 100% sensitivity of these currents to amiloride (33), to 40–60% (28, 30) and 20–25% (31, 34). However, no direct comparison could be made with our results, because all the reported patch-clamp studies on Type II cells were performed in classic whole-cell mode and, in some cases, using modified bath and pipette solutions to mask certain ion conductances. In this mode, intracellular Ca and ATP contents are buffered, thus possibly modifying whole-cell currents. In contrast to that we used the nystatin-perforated whole-cell mode in normal physiological solution, thus keeping normal physiological ion gradients and intact cellular Ca2+ and ATP homeostasis.

Our results are consistent with other studies showing that currents arising from the activity of ENaC in lung epithelial cells are regulated by glucocorticoids (12, 20, 35). The importance of constant levels of circulating endogenous cortisol in basal alveolar liquid clearance has been shown recently (36). These findings underscore the necessity of chronic glucocorticoid presence in maintaining basal rates of alveolar ion transport. We cannot rule out, however, that the ion transport up-regulation in the presence of dexamethasone seen in our study overlaps with the gradual transdifferentiation of Type II cells in slices into Type I cells.

Active absorption of sodium across the alveolar epithelium occurs by the standard mechanism involving entry of Na+ through the cell apical membrane down its electrochemical gradient followed by the active extrusion across the basolateral membrane by Na,K-ATPase. The driving force for net sodium entry, which generally represents the limiting step in the above process, is given by the difference between the resting membrane voltage (Vm) and the equilibrium potential for Na (ENa). Since the equilibrium sodium potential remains the same regardless of different conditions (ENa ~ +50 mV), it is important to determine Vm. A review of the literature shows that there is only limited number of studies describing Vm of freshly isolated Type II cells (33, 37). The comparison of published data with our results shows reasonable agreement in terms of the mean values (Table 1). The low values of cell membrane resting potential reported in our study (–4 mV) are not due to the cell damage during tissue slicing procedure, since differential cell staining shows the complete absence of co-localization of fluorescence signals from PI and QA, markers of damaged/apoptotic cells and lamellar bodies in Type II pneumocytes, respectively (Figure 1D). Analysis of available data on cell membrane resting potential suggests that since in Type II pneumocytes the cell membrane resting potential is close to zero, the driving force for net sodium entry in the basal state is determined mostly by ENa. It is also important to note that resting membrane voltage of these cells remains stable despite the up-regulation of amiloride-sensitive currents by dexamethasone (Figures 2 and 4). The hyperpolarization of these cells seen after application of cpt-cAMP in our study (~ 25 mV) represents approximately a 50% increase in this driving force and, thus, by itself could significantly modify the rate of sodium absorption. One could argue whether it is applicable to the in vivo conditions. Ionic contents of Type II cells and alveolar surface liquid have been reported (38, 39). Intracellular Na and Cl concentrations were 51 mM and 70 mM, respectively (38), and of these the alveolar fluid was –122 mM and 123 mM, respectively (39). From these values one can calculate the equilibrium potentials for conditions that approximate the in vivo situation (ENa ~+25 mV and ECl ~-15 mV). It is then plausible that the relative shift in the driving force for sodium entry due to hyperpolarization of Vm toward ECl after cAMP stimulation seen in our study and that for in vivo situation would be the same. This observation favors the hypothesis of an indirect role of Cl channel activity in modulation of active absorption of Na by the Type II pneumocytes after cAMP stimulation (29, 40). Similar hyperpolarization of Type II cells after cAMP stimulation has been observed by Brochiero and colleagues (41), although in their study amiloride-sensitive currents were silenced by amiloride. Together with direct activation of ENaCs by cAMP described by other groups in single-channel studies (21, 42) and in whole-cell study on lung epithelial cells (this article), the modulation of the driving force for sodium entry represents an additional mechanism for activation of sodium reabsorption by Type II cells. Thus, these two mechanisms of activation are cumulative. However, it may not be the case for rat Type I pneumocytes that have been shown to express ENaC channels but much lower levels of CFTR compared with Type II cells (6).

Our results support the hypothesis of the potential role of CFTR in the isoosmolar alveolar fluid transport (2, 43). We show that there is modest but significant Cl– entry into alveolar Type II cells under basal condition. This observation is consistent with the studies showing that inhibition of CFTR has a mild effect on alveolar liquid absorption under nonstimulated conditions and completely prevents its stimulation in the presence of cAMP (43, 44).

In summary, we present a novel functional lung slice model suitable for whole-cell currents measurements in Type II pneumocytes. This model preserves the alveolar architecture and keeps all phenotypic features typical of Type II cells in vivo, including tight junctions separating basolateral and apical cell membranes. It may prove useful in re-evaluation of the precise contribution of different ion channels to the alveolar ion and fluid transport under basal and stimulated conditions.


   Acknowledgments
 
The authors thank Dr. S. Sariban-Sohraby (National Eye Institute, Bethesda, MD) and Dr. I. Ismailov (Baylor College of Medicine, Houston, TX) for helpful criticism of the manuscript.


   Footnotes
 
This work was funded by grants from Université Libre de Bruxelles and Fonds Defay (to Dr. Sarah Sariban-Sohraby; currently managed by R.N.). V.S. was the recipient of a European Respiratory Society Fellowship (number 306). A.G. is a doctoral fellow from Fonds National de la Recherche Scientifique (Belgium).

Originally Published in Press as DOI: 10.1165/rcmb.2007-0227OC on February 14, 2008

Conflict of Interest Statement: None of the authors has a financial relationship with a commercial entity that has an interest in the subject of this manuscript.

Received in original form June 15, 2007

Accepted in final form January 29, 2008


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 Abstract
 CLINICAL RELEVANCE
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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D. H. Ingbar, M. Bhargava, and S. M. O'Grady
Mechanisms of alveolar epithelial chloride absorption
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