A glucocorticoid-induced Na⁺ conductance in human airway epithelial cells identified by perforated patch recording

Cell culture

Standard methods were used to maintain stocks of H441 cells in RPMI 1640 culture media containing fetal bovine serum (FBS, 8.5%), newborn calf serum (NCS, 8.5%), glutamine (2 mm), insulin (5 μg ml⁻¹), transferrin (5 μg ml⁻¹), selenium (5 ng ml⁻¹) and an antibiotic/antimycotic mixture (Sigma Chemical Co., Poole, UK). For experiments cells were removed from the culture flasks using trypsin–EDTA, resuspended in standard medium and plated onto glass coverslips. Once cellular attachment had occurred (2–3 h), the medium was replaced with medium identical to that described above except that the FBS and NCS were replaced with FBS (8.5%) that had been dialysed to remove all hormones/growth factors. Unless otherwise stated, insulin and transferrin were thus the only hormones/growth factors present. Control cells were maintained in this medium for 24–30 h before being used in experiments whilst dexamethasone-treated cells were incubated for the same length of time in medium that had been supplemented with this synthetic glucocorticoid (0.2 μm) but were treated identically in all other respects.

Solutions

The pipette filling solution contained (mm): NaCl, 10; KCl, 18; potassium gluconate, 92; MgCl₂, 0.5; EGTA, 1; Hepes, 10. The pH of this solution was adjusted to 7.2 with KOH, which brought [K⁺] to 113.3 mm. Immediately before recording, a 100 mg ml⁻¹ stock of nystatin was prepared in DMSO and added to the pipette solution (0.5 mg ml⁻¹) which was then ultrasonicated and used within 2–3 h. The standard bath solution contained (mm): NaCl, 140; KCl, 4.5; MgCl₂, 1; CaCl₂, 2.5; Hepes, 10; glucose, 5. The pH of this solution was adjusted to 7.4 with NaOH which brought [Na⁺] to 144.4 mm. In some experiments, the composition of the external solution was modified by isosmotically replacing Na⁺ with K⁺, N-methyl-d-glucammonium (NMDG⁺) or Li⁺.

Electrophysiology

Coverslips bearing growing cells were placed in a small perfusion bath (volume ∼1 ml) attached to the stage of a Nikon Diaphot inverted microscope where the cells were superfused with standard bath solution and viewed using phase contrast optics. Experiments were carried out at room temperature and involved recording from single cells or groups of 2–6 cells. The electrophysiological properties of the cells were investigated using a modification of the whole cell recording configuration in which electrical access to the cell interior is gained by including nystatin in the pipette filling solution. This renders the patch of membrane spanning the pipette tip permeable to K⁺, Na⁺ and Cl⁻ and thus allows experimental control over the internal concentrations of these ions, but prevents the loss of higher molecular weight substances and allows intracellular Ca²⁺ to be regulated by normal mechanisms (Horn & Marty, 1988). Voltage/current clamp was achieved with an Axopatch 200B amplifier (Axon Instruments, Foster City, CA, USA) and data were recorded using a Digidata 1322A data acquisition board (Axon Instruments). The recording pipettes were manufactured from borosilicate glass (Clark Electromedical Instruments, Reading, UK) using a Flaming/Brown micropipette puller (P-97, Sutter Instruments Co., CA, USA) and had resistances of 4–7 MΩ when filled with the pipette solution. Seals were obtained by bringing the pipette into contact with the cell and applying gentle suction. The development of electrical access to the cell interior was monitored by following the nystatin-induced fall in access resistance (R_a) using the standard features of the pCLAMP 9 suite of programs. Experiments were initiated once R_a had fallen to a stable value below 35 MΩ, this typically took 10–15 min. At the onset of each experiment R_a and input capacitance (C_m) were noted and necessary compensations applied using the standard features of the Axon amplifier/software. These parameters were monitored throughout all experiments and minor adjustments to the compensation circuitry made as required. The development of a substantial change in R_a or C_m led to the discontinuation of the experiment and so all presented data are from cells/groups of cells in which these parameters remained stable. Although recording from larger groups of cells was usually associated with larger values of C_m, this was not always the case and, as C_m provides an indicator of membrane surface area, this suggests variability in the extent to which neighbouring cells are electrically coupled. The mean value of C_m in all preparations studied was 70.8 ± 6.7 pF (n = 73) but C_m for those data unambiguously derived from single cells was 39.3 ± 3.4 pF (n = 27). All recorded currents were normalized to this value of C_m so that all current data in this paper refer to the current flow associated with a standard cell (pA cell⁻¹). Liquid junction potentials were calculated using JPCalc, a computer program written by Professor P. Barry (University of Sydney, Australia) implementing procedures detailed elsewhere (Barry & Lynch, 1991) and appropriate corrections have been applied to all data. Data are shown as mean ±s.e.m.; values of n refer to the number of cells in each group, and the statistical significance of differences between mean values was assessed using Student's t test. All reported phenomena were observed in cells at three or more different passage numbers.

Current clamp studies

Figure 1A shows continuous records of V_m made by monitoring zero current potentials in control and dexamethasone-treated cells. Analysis of data collected at the onset of these experiments, when the cells were superfused with standard saline, showed that treating the cells with this synthetic glucocorticoid caused significant depolarization (control: −54.4 ± 4.2 mV, dexamethasone-treated: −26.7 ± 9.4 mV, P < 0.02). Acute application of amiloride (10 μm) had no effect on control cells but hyperpolarized the dexamethasone-treated cells. This response (−16.8 ± 6.1 mV; P < 0.05) occurred with no discernible latency and V_m settled at its new value within 10–15 s. This hyperpolarization was reversible, although recovery was relatively slow, requiring 2–3 min (Fig. 1A). Analysis of the data collected in the presence of amiloride showed that the values of V_m measured in control (−55.2 ± 4.0 mV) and dexamethasone-treated (−43.5 ± 5.9 mV) cells did not differ significantly (P < 0.1). Amiloride thus abolishes the dexamethasone-induced depolarization.

The data in Fig. 1B are from dexamethasone-treated cells and this figure includes data showing the change in V_m evoked by transiently (20 s) lowering [Na⁺]_o to 10 mm, the concentration in the pipette filling solution, by isosmotically replacing this cation with NMDG⁺. In each of these experiments, we also measured the effect of amiloride (10 μm, 20 s pulse) and these data confirm the hyperpolarizing action of this drug (Fig. 1B). Replacing Na⁺ with NMDG⁺ also hyperpolarized the cells but this response was smaller than the response to amiloride. Parallel studies of control (i.e. dexamethasone-free) cells confirmed that V_m was insensitive to amiloride but, despite this, exposure to the NMDG⁺-containing solution did hyperpolarize these cells although the response (−10.3 ± 2.2 mV, n = 5, P < 0.05) was smaller (P < 0.005) than that measured in age-matched dexamethasone-treated cells at identical passage number (Fig. 1B). Further experiments showed that the Li⁺-containing saline caused a small depolarization in dexamethasone-treated cells (Fig. 1B) but had no effect on V_m in control cells (n = 5).

Although amiloride hyperpolarized all dexamethasone-treated cells in which it was tested (Fig. 1A and B), there was variability both in the magnitude of this response and in the values of V_m measured during exposure to the standard saline (Fig. 1A). Indeed, positive values of V_m were recorded in some instances, and these clearly lie outside the normal physiological range. Further analysis revealed a correlation between V_m and the magnitude of the amiloride-evoked fall in V_m (correlation coefficient r²= 0.766, P < 0.01); the largest responses to amiloride thus occur in the most strongly depolarized cells (Fig. 1C).

Properties of the dexamethasone-induced conductance

The data in Fig. 2 are from cells held under voltage clamp and show the current needed to hold V_m at −113 mV (I_-113mV) in control and dexamethasone-treated cells. This potential was chosen to maintain a large driving force for Na⁺ entry, and analysis of these data showed that dexamethasone increased the magnitude of this current ∼6-fold (P < 0.001). Although amiloride had no effect on I_-113mV in control cells, it reversibly inhibited the current in dexamethasone-treated cells (Fig. 2) and, in the presence of amiloride, there was no significant difference between the currents recorded from control and dexamethasone-treated cells (control: 24.4 ± 5.7 pA cell⁻¹; dexamethasone-treated: 44.0 ± 8.5 pA cell⁻¹, P < 0.1). Dexamethasone thus increases the amiloride-sensitive component of I_-113mV.

Further studies were therefore undertaken which involved recording the currents evoked by driving V_m through a series of voltage ramps (−113 mV to 87 mV in 1.75 s) from a holding potential of −53 mV. Figure 3A shows currents recorded in this way from dexamethasone-treated cells. Under standard conditions the current displays outward rectification at positive potentials and the reversal potential (V_Rev) is −22.9 ± 2.5 mV (Fig. 3A). Application of amiloride (10 μm) reduced the inward currents flowing at negative potentials whilst having little effect on the outward current (Fig. 3A); V_Rev was shifted (P < 0.001) to −54.0 ± 3.3 mV. Data recorded 5–6 min after amiloride was washed from the bath showed that these effects were reversible (not shown). Analogous studies of control cells showed that V_Rev was more negative (−60.8 ± 3.1 mV, P < 0.001) than in dexamethasone-treated cells and established that this parameter was insensitive to 10 μm amiloride (−64.8 ± 3.2 mV). As V_Rev serves as an indicator of V_m, these data confirm that dexamethasone evokes amiloride-sensitive depolarization of V_m (see Fig. 1A).

Figure 3B shows estimates of membrane conductance calculated by regression analysis (δI/δV_m) of the data collected at negative holding potentials and this analysis confirms (see Fig. 2) that dexamethasone increases membrane conductance (P < 0.001). Amiloride had no effect on the conductance of control cells but in dexamethasone-treated cells this drug reduced the conductance to a value that did not differ from control (P < 0.1). Dexamethasone thus acts by evoking the expression of an amiloride-sensitive conductance with a magnitude of ∼300 pS cell⁻¹. However, despite this clear result, there was variability both in the magnitude of this dexamethasone-induced conductance and in the value of V_Rev. Regression analysis showed that the most strongly depolarized cells were those with the largest amiloride-sensitive conductances (Fig. 3C). Moreover, this analysis also predicted that V_Rev in the absence of such a conductance would be ∼−50 mV (Fig. 3C), and this is very similar to the value actually measured after the dexamethasone-induced conductance had been blocked with amiloride (Fig. 3A).

Ionic selectivity of the dexamethasone-induced conductance

The experimental records in Fig. 4A show the current–voltage relationship for the amiloride-sensitive component of the membrane currents recorded from dexamethasone-treated cells using the voltage ramp protocol described above. Such currents were first recorded from cells bathed with the standard external solution (i.e. [Na⁺]= 144 mm) and a further series of such paired measurements were then made once external [Na⁺] had been successively lowered to 99 mm, 49 mm and 14 mm. This reduction in [Na⁺] was achieved by isosmotically substituting K⁺. Analysis of the data recorded under standard conditions showed that current flow was inward across almost the entire range of membrane potentials tested, and established that V_Rev was ∼65 mV, a value close to E_Na (68 mV). Lowering external [Na⁺] caused a reduction of the inward current recorded at hyperpolarizing potentials and a progressive, rightward shift in V_Rev (Fig. 4A). Analysis of these data revealed a linear relationship between V_Rev and E_Na and showed that the line fitted to these data by least squares regression could not be distinguished from the theoretical relationship predicted (Nernst equation) for a perfectly selective Na⁺ conductance (Fig. 4B). The amiloride-sensitive conductance expressed by the dexamethasone-treated cells is thus Na⁺ selective.

Figure 5 shows the results of further experiments in which amiloride-sensitive currents were recorded under standard conditions and after Na⁺ had been lowered to 10 mm, the concentration in the pipette filling solution, by isosmotically replacing this ion with NMDG⁺ or Li⁺. The control experiments confirmed, once again, that current flow was predominantly inward in the presence of external Na⁺ and that V_Rev (69.5 ± 11.8 mV) was essentially identical to E_Na. Replacing Na⁺ with NMDG⁺, and thus shifting E_Na to 0 mV, caused V_Rev to fall to a value that did not differ significantly from zero. In contrast, replacing Na⁺ with Li⁺ caused V_Rev to shift to a more positive value (79.1 ± 9.6 mV) that differed significantly (P < 0.02) from E_Na.

Effects of cAMP on the dexamethasone-induced conductance

To explore the extent to which the amiloride-sensitive conductance expressed by the dexamethasone-treated cells is subject to acute regulation via the cAMP-dependent signalling pathway, membrane currents were recorded under standard conditions and in the presence of 10 μm amiloride as described above. In one group of cells, these measurements were simply repeated after 20 min exposure to the standard salt solution and analysis of the total membrane currents showed that I_-113mV did not change during this period (0 min: −70.7 ± 15.7 pA cell⁻¹; 20 min: 62.9 ± 15.7 pA cell⁻¹). Similarly, there were no discernible changes in the amiloride-sensitive component of the total current, I_Amil. (Fig. 6A and C) and so it is clear that spontaneous changes in membrane conductance do not occur over the time scale of these experiments. The data in Fig. 6B are from age-matched cells at identical passage that were exposed to a cocktail of compounds (10 μm forskolin, 100 μm isobutylmethylxanthine, 1 mmN⁶, 2′-O-dibutyryladenonsine 3′5′-cyclic monophosphate) designed to provoke maximal activation of the cAMP-dependent signalling pathway once the first measurements of membrane conductance were completed. Data collected after 20 min exposure to these compounds revealed an increase in I_-113mV(0 min: 82.5 ± 23.6 pA cell⁻¹; 20 min: 125.8 ± 31.4 pA cell⁻¹, P < 0.05, Student's paired t test) and this response could be attributed to a ∼2-fold increase in I_Amil (Fig. 6A and C). Moreover, this response occurred with no change in V_Rev, indicating that the increase in conductance is not associated with a change in ionic selectivity. However, examination of the total membrane current records showed that the cAMP-activating drugs caused a depolarizing shift (18 ± 4 mV, P < 0.05) in V_Rev and so the cAMP-evoked increase in conductance is associated with a depolarization of V_m.

The dexamethasone-induced depolarization

In the present study, the first indication that glucocorticoids could modify the conductive properties of H441 cells came from the observation that dexamethasone-treated cells were depolarized by ∼25 mV. Acute application of 10 μm amiloride, a drug that blocks epithelial Na⁺ channels, reversibly abolished this effect and regression analysis showed that the largest amiloride-evoked hyperpolarizations occurred in the most strongly depolarized cells. Furthermore, amiloride had no effect on V_m in control cells and replacing external Na⁺ with NMDG⁺ caused a larger hyperpolarization in the dexamethasone-treated cells. Taken together, these data suggest strongly that the depolarization reflects the expression of an amiloride-sensitive Na⁺ conductance. However, the hyperpolarization that occurred when the cells were exposed to the NMDG⁺-containing solution was smaller than that evoked by amiloride, and this difference raised the possibility that the amiloride-sensitive conductance might permit a depolarizing influx of NMDG⁺. However, it is important to remember that 10 mm Na⁺ was present in the NMDG⁺-containing saline and that there will be a driving force for Na⁺ entry during exposure to this solution. In contrast, the concentration of amiloride used was sufficient to block Na⁺ influx completely (see for example Collett et al. 2002; Ramminger et al. 2003) and this predicts that amiloride would cause the largest hyperpolarization. It is interesting, however, that lowering external [Na⁺] by isosmotically replacing this cation with NMDG⁺ did cause a small hyperpolarization in control cells, which thus appear to express an amiloride-insensitive Na⁺ conductance. The molecular basis of this conductance is unknown but it may contribute to the larger responses seen when dexamethasone-treated cells were exposed to the NMDG⁺-containing solution.

In contrast to the effects of NMDG⁺, replacing Na⁺ with Li⁺ depolarized dexamethasone-treated cells but had no effect on control cells. These data thus suggest that the dexamethasone-induced conductance is more permeable to Li⁺ than Na⁺. It is interesting in this context that studies of other Na⁺-transporting epithelia suggest that electrogenic Na⁺ transport is mediated by the epithelial Na⁺ channel (ENaC) which characteristically displays a high permeability to Li⁺. Such channels are composed of three subunits (α-, β- and γ-ENaC), each encoded by a separate gene, and dexamethasone has been reported to increase the abundance of these mRNA species in H441 cells as well as in certain other lung-derived epithelial cell types (Tchepichev et al. 1995; Venkatech & Katzberg, 1997; Sayegh et al. 1999; Pitkänen et al. 2001; Itani et al. 2002).

Properties of the dexamethasone-induced conductance

Steady state measurements of I_-113mV showed that dexamethasone-treated cells expressed an amiloride-sensitive conductance that was not present in control cells, and this conclusion was supported by further experiments that analysed the currents evoked by ramp changes in V_m. These studies also confirmed that dexamethasone caused amiloride-sensitive depolarization, and regression analysis showed that the most strongly depolarized cells were those with the largest amiloride-sensitive conductance. These findings, in common with the direct measurements of V_m, thus suggest that the depolarization is secondary to the increased conductance. Analysis of the present data suggested that the magnitude of this dexamethasone-induced conductance was ∼300 pS cell⁻¹. It is interesting that parallel studies of H441 cells grown as confluent epithelial sheets also indicate that amiloride-sensitive conductance is negligible in control (i.e. hormone-free) cells but increased to ∼250 μS cm⁻² in dexamethasone-treated cells (Ramminger et al. 2004). As the cultured epithelia used in these studies contained ∼10⁶ cells cm⁻², the conductance measured in this preparation equates to single cell conductance of ∼250 pS cell⁻¹. Although they used very different methods, there is excellent agreement between the two studies.

Dexamethasone-evoked increases in amiloride-sensitive conductance have been reported in A549 cells, another lung-derived epithelial cell type (Lazrak et al. 2000b, c) but, in contrast to the situation described here, these cells expressed a significant amiloride-sensitive conductance if maintained under hormone-free conditions. Dexamethasone thus caused a relatively modest (∼3-fold) augmentation of this basal conductance (Lazrak et al. 2000b, c). Moreover further analysis of these data (Lazrak et al. 2000c) indicates that the amiloride-sensitive conductance of dexamethasone-treated cells is ∼20-fold greater than the value reported here (see also Ramminger et al. 2004). However, the present study also showed that V_m in dexamethasone-treated cells is ∼−25 mV and so, under standard conditions, there would be a ∼90 mV driving force for Na⁺ entry, and a conductance of 300 pS cell⁻¹ would thus allow a Na⁺ current of −27 pA cell⁻¹. When grown as confluent epithelial sheets, H441 cells spontaneously generate 20–30 μA cm⁻² of amiloride-sensitive short circuit current (Ramminger et al. 2004) and, as such cultures contain ∼10⁶ cells cm⁻², this corresponds to a current of 20–30 pA cell⁻¹. The conductance that we now describe is therefore large enough to account for the observed rate of Na⁺ transport. Interestingly, the amiloride-sensitive conductance of the Na⁺-absorbing amphibian urinary bladder epithelium also appears to be ∼200 μS cm⁻² (Gordon & MacKnight, 1991) and, if we assume that this epithelium also contains ∼10⁶ cells cm⁻², this conductance is very similar to that reported here (see also, Ramminger et al. 2004). Moreover, the amiloride-sensitive conductance of Na⁺-absorbing colonic epithelial cells appears to be 3–4 times larger than in H441 cells (Inagaki et al. 2004) and it is therefore interesting that this tissue generates 3–4 times more amiloride-sensitive short circuit current (Ramminger et al. 2004; Inagaki et al. 2004). It is therefore clear that physiologically relevant quantities of Na⁺ can be transported by epithelia with very modest amiloride-sensitive conductances (present study, Gordon & MacKnight, 1991; Inagaki et al. 2004).

Ionic selectivity of the dexamethasone-induced conductance

Heterologous coexpression studies have shown that expression of α-, β- and γ-ENaC is associated with the appearance of small (∼5 pS) Na⁺ channels that are essentially impermeable to K⁺ and such channels appear to underlie epithelial Na⁺ transport in several absorptive tissues (Canessa et al. 1993, 1994; Ishikawa et al. 1998; Inagaki et al. 2004). However, despite much effort, many workers have failed to identify such channels in distal airway epithelial cells. Indeed, many such studies indicate that the most abundant amiloride-sensitive channel in such cells is a 25 pS cation channel that cannot discriminate between Na⁺ and K⁺. It has therefore been proposed that distal airway Na⁺ transport is dependent on these channels rather than selective Na⁺ channels (Orser et al. 1991; MacGregor et al. 1994; Tohda et al. 1994; Marunaka, 1996, 1999; Matalon & O'Brodovich, 1999; Kemp et al. 2001). Moreover, there is evidence that these non-selective channels may reflect an alternative, stoichiometric arrangement of the ENaC subunits (Kizer et al. 1997; Jain et al. 1999, 2001), a hypothesis that reconciles a role for non-selective channels in pulmonary Na⁺ transport with the fact that this process is clearly dependent on α-ENaC (Hummler et al. 1996). However, not all data are consistent with this hypothesis as true Na⁺ channels have been identified by some authors (Voilley et al. 1994; Lazrak et al. 2000b, c; Jain et al. 2001; Itani et al. 2002; Lazrak & Matalon, 2003), and as electrometric studies of cells isolated from the alveolar region of the fetal (Baines et al. 2001) and adult (Jiang et al. 1998) lung show that the amiloride-sensitive conductance in the apical membrane is Na⁺ selective.

Our data show clearly that, at least under the present conditions, the dexamethasone-induced conductance in H441 cells is highly selective for Na⁺ over K⁺, essentially impermeable to NMDG⁺, but has a substantial permeability to Li⁺. The dexamethasone-evoked currents are therefore carried by Na⁺ flowing through channels that are essentially identical to those associated with heterologous coexpression of α-, β- and γ-ENaC (Canessa et al. 1994; Ishikawa et al. 1998). There have been few previous electrophysiological studies of H441 cells but selective Na⁺ channels have been identified by some workers (Itani et al. 2002; Lazrak & Matalon, 2003) although we are unaware of a systematic quantification of their abundance. However, if we assume an open probability of ∼0.3 (see Lazrak & Matalon, 2003), then our data predict that only ∼180 such channels will be present at the cell surface. This very low density may explain why these channels have been so difficult to identify in distal airway epithelial cells.

Effects of cAMP

Although glucocorticoid hormones are important to the development of the lung's Na⁺-absorbing phenotype, pulmonary Na⁺ transport is also subject to acute regulation via cAMP-coupled agonists (see for example Olver et al. 1986; Collett et al. 2002), but the mechanisms underlying this control have not been agreed upon (see review by Widdicombe, 2000). Work from this laboratory (Collett et al. 2002; Ramminger et al. 2002) and elsewhere (Lazrak et al. 2000a; Lazrak & Matalon, 2003) suggests that the response is due to an increase in Na⁺ conductance; however, others favour a model in which G_Na remains constant. In this model, cAMP-dependent agonists are thought to stimulate Na⁺ transport by hyperpolarizing the cell and thus increasing the driving force for Na⁺ entry, and this hyperpolarization is attributable to the cAMP-induced activation of the cAMP-dependent Cl⁻ channel encoded by the gene that is defective in cystic fibrosis (CFTR channels) (Jiang et al. 1998; O'Grady et al. 2000; Jiang et al. 2001). The present data show clearly that a cocktail of cAMP-activating compounds can cause a 2-fold increase in the amiloride-sensitive conductance of dexamethasone-treated cells and this result, in common with data recently presented by Lazrak & Matalon (2003), supports the view that Na⁺ transport is controlled via changes in conductance. Interestingly, there is evidence that this control involves a mechanism that regulates the number of channels in the membrane (Morris & Schafer, 2002). The present study also provided no evidence to support the view (Jiang et al. 1998, 2001; O'Grady et al. 2000) that stimulation with cAMP-dependent agonists increases the driving force for Na⁺ entry. Indeed, all available data now show that such agonists cause depolarization so that the increase in conductance is accompanied by a fall in the driving force for Na⁺ entry (present study; Lazrak & Matalon, 2003). However, it is clear that, in certain experimental situations, compounds that characteristically block Cl⁻ channels can inhibit Na⁺ transport (O'Grady et al. 2000). Moreover, in vivo studies of mouse lungs have shown that lung liquid absorption is inhibited, both by compounds that block CFTR channels and by artificial knockout of the CFTR gene (Fang et al. 2002). There is thus good evidence that the control of Na⁺ conductance is in some way dependent on these channels, although in our view the mechanism of this interdependence is far from clear.

The present data thus suggest that distal airway Na⁺ transport occurs via selective (see for example Voilley et al. 1994) rather than non-selective (see for example Marunaka et al. 1999) Na⁺ channels, and indicates that the cAMP-dependent control of Na⁺ transport seen in these tissues reflects control over this conductance rather than modification of the driving force for Na⁺ entry. However, these data are derived from an adenocarcinoma cell line, and so it is now important to determine the extent to which these conclusions are applicable to cells acutely dissociated from the lung.