Coordinated release of nucleotides and mucin from human airway epithelial Calu-3 cells

Abstract

The efficiency of the mucociliary clearance (MCC) process that removes noxious materials from airway surfaces depends on the balance between mucin secretion, airway surface liquid (ASL) volume, and ciliary beating. Effective mucin dispersion into ASL requires salt and water secretion onto the mucosal surface, but how mucin secretion rate is coordinated with ion and, ultimately, water transport rates is poorly understood. Several components of MCC, including electrolyte and water transport, are regulated by nucleotides in the ASL interacting with purinergic receptors. Using polarized monolayers of airway epithelial Calu-3 cells, we investigated whether mucin secretion was accompanied by nucleotide release. Electron microscopic analyses of Calu-3 cells identified subapical granules that resembled goblet cell mucin granules. Real-time confocal microscopic analyses revealed that subapical granules, labelled with FM 1-43 or quinacrine, were competent for Ca²⁺-regulated exocytosis. Granules containing MUC5AC were apically secreted via Ca²⁺-regulated exocytosis as demonstrated by combined immunolocalization and slot blot analyses. In addition, Calu-3 cells exhibited Ca²⁺-regulated apical release of ATP and UDP-glucose, a substrate of glycosylation reactions within the secretory pathway. Neither mucin secretion nor ATP release from Calu-3 cells were affected by activation or inhibition of the cystic fibrosis transmembrane conductance regulator. In SPOC1 cells, an airway goblet cell model, purinergic P2Y₂ receptor-stimulated increase of cytosolic Ca²⁺ concentration resulted in secretion of both mucins and nucleotides. Our data suggest that nucleotide release is a mechanism by which mucin-secreting goblet cells produce paracrine signals for mucin hydration within the ASL.

The airway surface liquid (ASL) is a protective and dynamic liquid film essential for ciliary beating and mucin hydration, which are key components of the mucociliary clearance (MCC) process that removes noxious materials from the lung (Boucher, 2007). The superficial human airway epithelium that mediates MCC is composed mainly of ciliated cells with a small component of mucin-secreting goblet cells. ASL volume regulation by ciliated cells and gel-forming mucin secretion from goblet cells are exquisitely coordinated to maintain physiological viscoelastic properties of the two ASL layers (i.e. the peri-ciliary and mucus layers). While it is recognized that mucus hydration and other MCC activities are regulated in part by signals generated within the ASL (Tarran et al. 2006), the mechanisms by which ion/water secretion and mucin secretion rates are coordinated is poorly understood.

Compelling evidence suggests that nucleotides and nucleosides present in ASL provide endogenous signalling to MCC functions. For example, electrophysiological studies with human airway epithelial Calu-3 cells have illustrated that the cystic fibrosis transmembrane conductance regulator (CFTR) Cl⁻ channel activity was regulated by extracellular adenosine derived from released ATP (Huang et al. 2001). Furthermore, removal of adenosine from the ASL bathing primary cultures of well-differentiated non-cystic fibrosis (CF) bronchial epithelial cells resulted in a phenotype (i.e. ASL depletion) that resembled the ion/water secretion deficiency characteristic of CF epithelia (Lazarowski et al. 2004b; Tarran et al. 2005). While adenosine generated from ATP promotes Cl⁻ secretion via CFTR activation, ATP itself activates a Ca²⁺-activated Cl⁻ channel that can, at least in part, compensate for CFTR deficiency (Tarran et al. 2005).

The extracellular actions of adenosine, ATP and other nucleotides are mediated by three classes of purinergic receptors. The G protein-coupled P1 receptor family, comprising four members (A₁, A_2a, A_2b and A₃), is selectively activated by the nucleoside adenosine (Fredholm et al. 2001). The ligand-gated ion channel P2X receptors, including seven species (P2X_1–7) are selectively activated by ATP (North, 2002). The G protein-coupled P2Y receptors, comprising eight species (P2Y₁, P2Y₂, P2Y₄, P2Y₆, P2Y₁₁, P2Y₁₂, P2Y₁₃ and P2Y₁₄), are activated by adenine and/or uridine nucleotides or nucleotide sugars (Abbracchio et al. 2006). The A_2b, P2Y₂, P2Y₆ and possibly some P2X receptors are functionally expressed on the apical surface of airway epithelial cells (reviewed by Tarran et al. 2006). Physiological, pharmacological and genetic evidence suggests that A_2b and P2Y₂ receptors are major transducers of nucleotide-/nucleoside-regulated ion and water transport, ciliary beating and mucin secretion in the airway epithelia (Tarran et al. 2006). Given the importance of the responses triggered by extracellular nucleotides in the airways, understanding how these molecules are released in a regulated fashion has important physiological implications for airway defence. However, little is known about the mechanisms involved in the release of nucleotides into ASL.

Recently, using polarized monolayers of Calu-3 cells, we demonstrated tonic release of ATP into the mucosal but not basolateral extracellular compartment, suggesting that ATP release may be controlled by mechanisms that are associated with apical membrane sorting (Lazarowski et al. 2004b). We have also discovered that most cells, including airway epithelial cells, tonically release UDP-glucose (UDP-Glc), which is a substrate for glycosylation reactions within the secretory pathway (Lazarowski et al. 2003b). Together, these observations suggested a link between nucleotide release and export of apically targeted glycoconjugates. In the present study, we used polarized monolayers of Calu-3 cells to test the hypothesis that Ca²⁺-regulated vesicle/granule exocytosis of glycoproteins (e.g. gel-forming mucins) occurs concomitantly with nucleotide release in mucin-secreting cells. If present, such coordinated release may provide a mechanism by which ‘condensed’ mucins secreted from goblet cells (Verdugo, 1990) are appropriately hydrated on airway surfaces.

Methods

Cell culture

Airway epithelial Calu-3 cells are derived from pleural effusion associated with a human lung adenocarcinoma (Shen et al. 1994). Unless otherwise indicated, Calu-3 cells were grown on 12-mm Transwell supports (Corning) and maintained at air–liquid interface for at least 3 weeks, as previously described (Lazarowski et al. 2004b). Rat tracheal epithelial SPOC1 cells were grown on 12-well cluster plates (Costar, Cambridge, MA, USA) as previously described (Abdullah et al. 1997). Cells were used for experiments at 18–22 days post confluence.

Electron microscopy

Calu-3 cultures were fixed and processed for electron microscopy (EM), as previously described (Kreda et al. 2000).

Measurement of vesicle exocytosis by real-time confocal microscopy

FM 1-43-associated granule fluorescence

Secretory granules were labelled as previously described (Cochilla et al. 1999; Brumback et al. 2004). Briefly, Calu-3 cultures grown on glass cover slips were incubated with 3 μm FM 1-43 for 30 min, exposed to a brief (1 min) hypotonic shock (67% tonicity) to initiate endocytotic uptake of the fluorophore, and incubated in minimum essential medium (MEM)–Hepes in the absence of FM 1-43 for an additional 4 h at 37°C. Cells were mounted on the stage of a Leica SP2 AOBS confocal microscope equipped with an HCX Apo L63 × NA 0.9 immersion lens and a 488-nm Ar laser (Leica, Germany). Cells were bathed in at least 2 ml of MEM–Hepes, or Hank's balanced salt solution (HBSS)–Hepes with 1.6 mm or nominally 0 Ca²⁺ for 15 min, and then challenged as described in the figure legends. Real-time recording was performed every 10 or 30 s in the xz axes by galvo-stage xz scanning, or in face by xy scanning of the subapical domain (uppermost 3 μm). The fluorescence intensity of all the pixels contained within a granule was quantified. The data from all granules of ∼1 μm diameter were measured at each time point, normalized to basal values (time 0) and averaged using Leica software. To confirm the cellular location of FM 1-43-loaded granules, the corresponding differential interference contrast (DIC) confocal planes were recorded in parallel and a full z-stack was generated at the end of each experiment.

Quinacrine and acridine orange granule fluorescence

Labelling of granules with 10 μm quinacrine or 5 μm acridine orange was achieved by incubation of Calu-3 cells at 37°C for 15–30 min with either dye. When indicated, 2 μm bafilomycin A1 was added to cells 30 min prior to the addition of quinacrine or acridine orange. Confocal microscopy analysis was carried out as described above using a 488-nm Ar laser and emission band widths of 500–540 nm (quinacrine) or 600–700 nm (acridine orange). Data acquisition and analysis were performed as above. Additionally, granules presenting similar fluorescence decay kinetics were grouped and averaged.

Mucin mRNA expression

RT-PCR analyses of human transcripts for MUC1 (362 forward (F), gccagtagcactcaccatagctcg; 762 reverse (R), tgacagacagccaaggcaatgag), MUC2s (418 F, agc gat gcc tac acc aaa gt; 652 R, gat cct cac aca cca cat cg), MUC4 (625 R, cag gct gtg ttc acc ata g; 122 F, cat cag gaa tga caa cac c), MUC5AC (1781 F, tct atg agg gct gcg tct tt; 1984 R, cgt agc agt agg agg ggt tg), MUC5B (9839 F, tac caa agt gcc gac tac c; 10203 R, gtg cta gag gag ggt gtt gc), MUC16 (1878 F, agt tct gta ccc acc acc a; 1910 R, gca ccg ttc ttc aca gac ct) and aquaporin 5 (953 F, cat ctt cgc ctc cac tga ct; 1145 R, ccc tac cca gaa aac cca gt) were performed at the UNC-CH Cystic Fibrosis Center Molecular Biology Core Laboratory using standardized protocols. Amplified PCR products were subsequently identified by sequence analysis at the UNC-CH DNA sequencing facility.

Immnofluorescence and confocal microscopy

Calu-3 cells were fixed in 4% paraformaldehyde, permeabilized in ice-cold methanol for 2 min, and subjected to immunofluorescence staining and confocal microscopic analysis, as previously described (Kreda et al. 2005). Two different and well-characterized antibodies were independently used against human CFTR (monoclonal antibodies 596 and 769 (Mall et al. 2004; Kreda et al. 2005)), human MUC5AC (monoclonal antibody 45M1 and polyclonal antibody MAN-5AC1 (Sheehan et al. 2000; Kreda et al. 2005), and human MUC1 (monoclonal antibody M2C5 (Okada et al. 2006) and polyclonal antibody (epitope aa 239–255)). In all experiments, negligible fluorescence was observed when the primary antibody was replaced by either an equivalent dilution of species homologue non-immune IgG or blocking solution (not shown).

The efficacy of cytochalasin D treatment (30 min at 37°C) in disrupting actin fibres was tested at the end of each experiment by fixing and staining the cells with fluorescent phalloidin, as previously described (Kreda et al. 2005).

Mucin and lysozyme secretion

To remove deposited surface mucus, Calu-3 cell cultures were rinsed three times and incubated at 37°C for 1.5 h with HBSS/Hepes. Subsequently, the mucosal and basolateral media were replaced with 300 μl HBSS/Hepes (with or without test drugs) and after 10 min at 37°C, 200 μl aliquots were removed bilaterally, centrifuged to eliminate potential cell debris, and analysed by slot blot. Cells were promptly fixed and subjected to immunofluorescence staining and confocal microscopic analysis as above. For slot blot analyses, 50–100 μl sample aliquots were blotted on nitrocellulose membranes (Bio-Rad, Hercules, CA, USA), which were developed first with an anti-human lysozyme antibody, and subsequently immunostained with monoclonal antibody 45M1 against human MUC5AC (Kreda et al. 2005). For cell enzyme-linked immunosorbent assay (ELISA), Calu-3 cells grown in 96-well plates were challenged and processed for immunostaining as above with MUC1 antibodies, except that permeabilization was omitted, secondary antibodies were labelled with IRDye 800CW, and TO-PRO-3 iodide was utilized to stain nuclei. Plates were analysed using an infrared LI-COR Odyssey scanner (Lincoln, NE, USA). MUC1 fluorescence integrated intensity was normalized using nuclear fluorescence, and the results were expressed as arbitrary fluorescence units (AFUs). To measure mucin secretion from rat SPOC1 cells, the mucin enzyme-linked lectin assay (ELLA) was performed as previously described (Abdullah et al. 1997).

Real-time luciferin–luciferase assay for ATP

Real-time ATP measurements in thin ASL were performed as described recently (Okada et al. 2006). Briefly, Calu-3 cell cultures were bilaterally rinsed twice, transferred to a Turner TD-20/20 luminometer (Turner Biosystems, Sunnyvale, CA, USA), and 50 μl HBSS/Hepes containing 0.5 μg luciferase (specific activity, 15–30 × 10⁶ light units (mg protein)⁻¹) and 150 μm luciferin was added to the mucosal compartment. This addition resulted in a 28 μm high apical liquid film. The basolateral compartment was incubated in the presence of 1 ml HBSS/Hepes. Luminescence was integrated every 4 s and recorded as arbitrary light units at 30 s intervals (unless indicated otherwise). Calibration curves using known concentrations of ATP were performed at the end of each assay.

Quantification of adenyl purines via etheno-derivatization and HPLC analysis

Calu-3 cell cultures were preincubated in 0.4 ml basolateral, 0.3 ml mucosal HBSS, and SPOC1 cells were preincubated in 0.5 ml HBSS for 90 min before challenge. Aliquots (150 μl) were removed, heated to 95°C to inactivate secreted nucleotidases, and then derivatized with chloroacetaldehyde; the resulting fluorescent etheno-species were analysed by HLPC as previously described (Lazarowski et al. 2004b).

Quantification of UDP-Glc

Calu-3 cells were incubated bilaterally in 0.4 ml MEM. SPOC1 cells were incubated in 0.5 ml MEM. UDP-Glc was assayed in 100 μl aliquots in the presence of 25 mm Hepes (pH 7.4), 0.5 U ml⁻¹ UDP-Glc pyrophosphorylase and 200 nm 0.2 μCi [³²P]pyrophosphate. Incubations were terminated by addition of 0.3 mm pyrophosphate. The UDP-Glc-dependent conversion of [³²P]pyrophosphate to [³²P]UTP was quantified by HLPC via a Nova-Pack C18 column, as previously described (Lazarowski et al. 2003b).

Measurement of UDP-[³H]Glc metabolism

Calu-3 cells were incubated for the indicated times in 0.5 ml mucosal and basolateral MEM in the presence of 0.1 μCi UDP-[³H]Glc added bilaterally, and the resulting species were analysed by HPLC as previously described (Lazarowski et al. 2003b).

CFTR short circuit current (I_sc) measurement

Calu-3 cell bioelectrical activities were assessed in modified Ussing chambers as described (Kreda et al. 2005).

Reagents

Bafilomycin A₁, cytochalasin D, ionomycin, thapsigargin and forskolin were purchased from Calbiochem (San Diego, CA, USA). Acridine orange, luciferase, luciferin, purified lysozyme, NTP-pyrophosphatase, quinacrine, UDP-Glc pyrophosphorylase and uridine monophosphate (UMP) were purchased from Sigma (St Louis, MO, USA). BAPTA-AM, fluorescently labelled phalloidin, FM 1-43 and TO-PRO-3 iodide were purchased from Molecular Probes (Eugene, OR, USA). [γ³²P]ATP (3000 Ci mmol⁻¹) and UDP-d-[6-³H] glucose (27 Ci mmol⁻¹) were purchased from Amersham Biosciences (Piscataway, NJ, USA). Anti-aquaporin 5 antibody (Kreda et al. 2001) was obtained from Chemicon (Temecula, CA, USA). Anti-human CFTR monoclonal antibodies 596 and 769 (Mall et al. 2004; Kreda et al. 2005) were a kind gift from Dr John Riordan (UNC-CH). Antibodies against MUC5AC MAN-5AC1 (Kreda et al. 2005) and MUC5B were a kind gift from Dr John Sheeham (UNC-CH). Anti-human MUC1 monoclonal M2C5 (Okada et al. 2006) and polyclonal (epitope aa 239–255) antibodies, and antihuman MUC5AC monoclonal antibody 45M1 (Kreda et al. 2005) were purchased from LabVision (Fremont, CA, USA) and Abcam. Anti-human lysozyme antibody was from BioGenex (Netherlands). Syntaxin 3 and 4 antibodies were purchased from Abcam and BD Biosciences Pharmingen (San Jose, CA, USA), respectively. All secondary antibodies were from Jackson ImmunoResearch Laboratories (West Grove, PA, USA), except those labelled with IRDye 800CW, which were purchased from LI-COR (Lincoln, NE, USA). CFTR_inh-172 (Ma et al. 2002) was synthesized in house. Other chemicals were of the highest purity available and from sources previously reported.

Statistics

Student's paired ttest was performed using Sigma Plot 8.0; P < 0.01 was accepted to indicate statistical significance.

Results

Calu-3 cells express mucin granules released by regulated exocytosis

Our studies with polarized cultures of Calu-3 cells suggested that the release of ATP was linked to an apical plasma membrane secretion process (Lazarowski et al. 2004b). EM studies indicated that Calu-3 cell monolayers express ∼1 μm-diameter round granules that were apically localized. These granules, observed in ∼30–40% of the cells, were electron-translucent with an electron dense core (Fig. 1), resembling mucin granules of airway mucous goblet cells in organization and size (Davis & Randell, 2001). No electron dense granules, characteristic of native submucosal gland acinar serous cells, were observed, as previously reported (Shen et al. 1994). The presence of these mucin-like granules in Calu-3 cells prompted us to examine the potential relationship between granule exocytosis/mucin secretion and nucleotide release in these cells.

Figure 1. Calu-3 cells express goblet cell-like mucin granules.

EM micrograph of a Calu-3 cell monolayer displaying ∼1 μm diameter dense-core electron-translucent granules; the area within the frame is shown at higher magnification in the right panel; bar, 1 μm.

The styryl fluorophore FM 1-43 was used to assess exocytosis of secretory granules in real time by confocal microscopy. FM styryl probes are virtually non-fluorescent in aqueous solution, but their fluorescence quantum yields greatly increase upon their reversible partitioning into the hydrophobic environment of cell membranes. Using a modified protocol designed to load FM 1-43 into dense-core granules in neuro-endocrine cells (Cochilla et al. 1999; Brumback et al. 2004), incorporation of FM 1-43 into ∼1 μm-diameter, apically organized granules was achieved (Fig. 2A, left panel and Fig. 2B, top panel). Fluorescently labelled granules were present in 30–40% of the cells (Fig. 2A, left panel) and, in the absence of stimuli, remained unchanged for at least 60 min (data not shown).

Figure 2. Real-time exocytosis of FM 1-43-labelled granules.

Calu-3 cell granules were preloaded with FM 1-43 and monitored by confocal microscopy analysis as described in the Methods. Cells were challenged with vehicle (CONT), 10 μm ionomycin in 1.6 mm (IONO) or 0 extracellular Ca²⁺ (0 Ca), 10 μm forskolin (FSK), 10 μm ionomycin in the presence of 5 μm cytochalasin D (CYTD) or 1 μm thapsigargin (THAP). A, identical xy focal plane illustrating cells at time 0 and after 5 min of ionomycin treatment; bar, 2 μm. Insets, confocal DIC plane corresponding to the area delimited by a white rectangle in the fluorescence panel, encompassing at least one cell; asterisks indicate the same position in all panels; bar, 2 μm. Note, after ionomycin treatment, loss of fluorescence in the red channel is accompanied by absence of granule images in the DIC channel. B, identical galvo-stage xz focal plane displaying subapical localization of fluorescent granules, before and after 10 s and 5 min of ionomycin treatment. Arrowheads indicate plasma membrane position; asterisks denote single fluorescent granules that loose fluorescence after 10 s of ionomycin treatment; bar, 20 μm. C, Quantification of changes in overall granule-associated fluorescence over time in cells incubated with indicated agents. Fluorescence changes associated with all 1 μm granules (relative to time 0, i.e. 100%) are expressed as mean ±s.e.m. Similar results were obtained in at least three experiments performed in duplicate.

The Calu-3 cells used in our laboratory do not express Ca²⁺-mobilizing P2Y₁ or muscarinic receptors (not shown), and therefore ionomycin was used to raise intracellular Ca²⁺. Addition of ionomycin in the presence of calcium promoted loss of granule-associated fluorescence (Fig. 2A, right panel, and Fig. 2B, middle and bottom panels), which was observed as early as 10 s post stimulation (the earliest reproducible time point recorded; Fig. 2B, middle panel). Parallel recording in face of the corresponding DIC xy confocal plane (displaying the absence of granule images; Fig. 2A, insets), and a thorough scan of additional xy focal planes at the end of the incubation period (data not shown), indicated that the loss of granule-associated fluorescence reflected disappearance of fluorescent granules from the cells. Ionomycin-promoted changes in granule-associated fluorescence occurred in the presence, but not in the absence of extracellular Ca²⁺ (Fig. 2C). Thapsigargin, which increases intracellular Ca²⁺ concentration by a different mechanism and by different kinetics compared with ionomycin (Pedrosa Ribeiro et al. 2000), also elicited loss of fluorescently labelled granules in Calu-3 cells. Thus, Calu-3 cells exposed for 5 min to thapsigargin (1 μm) lost 74 ± 5% (n= 3) of FM 1-43 granule-associated fluorescence. Because Ca²⁺-regulated exocytosis depends on the integrity of the cytoskeletal network (Richards et al. 2004), the disruption of actin cytoskeleton by cytochalasin D was investigated. The ionomycin-elicited decrease in granule-fluorescence was blocked by cytochalasin D (Fig. 2C). Collectively, the results in Fig. 2 indicate that ionomycin-promoted loss of granule fluorescence reflected Ca²⁺-trigered exocytosis of apically localized granules. In contrast to ionomycin, forskolin produced negligible changes in granule-associated fluorescence (Fig. 2C).

Having established that Calu-3 cells are competent for Ca²⁺-regulated exocytosis, we investigated whether Calu-3 cells produced and secreted mucins. An initial screening by RT-PCR of mucins commonly expressed in airway epithelia indicated that Calu-3 cells expressed transcripts for MUC5AC and MUC5B, two secretory mucins highly expressed in goblet airway epithelial cells (Buisine et al. 1999; Kreda et al. 2005), and the membrane-tethered MUC1 (Fig. 3A). MUC4, a tethered mucin expressed abundantly in ciliated airway cells (Buisine et al. 1999), and MUC2, an intestinal goblet cell secretory mucin (Mall et al. 2004), were not amplified from Calu-3 cells. Aquaporin 5, a water channel abundantly expressed in native submucosal gland acinar serous cells (Kreda et al. 2001), was amplified from Calu-3 cells, but its signal was weak relative to that of total lung (Fig. 3A).

Figure 3. Mucin expression in Calu-3 cells.

A, RT-PCR analysis of mRNA extracts from Calu-3 cells, and human lung and colon using specific primers for the indicated transcripts (see Methods). Brackets indicate the 500 bp DNA standard band. S, standard DNA ladder; L, lung; C, Calu-3 cells; Co, colon; (–), none. B, confocal microscopy analysis of Calu-3 cells co-immunostained with monoclonal and polyclonal antibodies against MUC5AC (45M1, red) and MUC1 (green), respectively. In face (xy scanning) confocal microscopy images at low magnification (a) and high resolution (b) of the uppermost 3 μm of the cell are shown. c, galvo-stage xz scanning of a monolayer illustrating that MUC5AC staining (red) is associated with ∼1 μm diameter granules localized towards the cell apex; bar, 20 μm. C, confocal microscopy xy (left) and xz (right) images of Calu-3 cells co-immunostained with monoclonal antibody 769 against human CFTR (green) and a polyclonal MUC5AC antibody (MAN-5AC1, red). CFTR is expressed in the apical membrane of most cells, except in those exhibiting MUC5AC immunostaining. Note, the MAN-5AC1 polyclonal antibody reacts against ‘relaxed’ MUC5AC from granules disrupted using the current fixation/permeabilization conditions. Arrowheads indicate plasma membrane position; bar, 20 μm.

Utilizing the monoclonal antibody 45M1, which selectively recognizes MUC5AC in native human airways (Kreda et al. 2005), strong anti-MUC5AC immunoreactivity was visualized in up to 40% of the cells by confocal microscopy (Fig. 3Ba). MUC5AC immunostaining was concentrated in round granules (∼1 μm diameter) that were localized towards the apex of the cell (Fig. 3Bb and c). No MUC5AC signal was observed in the basolateral domains (Fig. 3c). Figure 3B also illustrates that Calu-3 cells displayed MUC1 immunoreactivity, which was associated with the apical plasma membrane. MUC1 immunostaining signal was stronger in those cells exhibiting weak or negative MUC5AC immunoreactivity (Fig. 3B). The Cl⁻ channel CFTR is abundantly expressed in Calu-3 cells (Shen et al. 1994). Immunoreactivity towards CFTR was evident at the apical domain of Calu-3 cell monolayers, but MUC5AC and CFTR immunostaining signals segregated to different cells (Fig. 3C). CFTR and MUC1 colocalized in the surface of MUC5AC-negative (non-mucous) Calu-3 cells (data not shown). The results suggest that MUC5AC is produced by a subset of Calu-3 cells different from CFTR-expressing ‘non-mucous’ cells. Aquaporin 5 was detected in the apical domain of non-mucous Calu-3 cells, but staining levels were much lower than in native gland acinar serous cells (data not shown), which is consistent with the RT-PCR findings (Fig. 3A).

Next, we examined whether MUC5AC immunoreactive granules in Calu-3 cells were competent for Ca²⁺-triggered exocytosis. Treatment of the cells with ionomycin resulted in marked reduction of granule-associated MUC5AC immunostaining (Fig. 4A). The effect of ionomycin was dependent on the presence of extracellular Ca²⁺ (Fig. 4A) and was inhibited by cytochalasin D (see below). Forskolin induced no change in granule-associated MUC5AC immunostaining (Fig. 4A). In contrast to the effect of ionomycin on MUC5AC, assessment of cell surface MUC1 by cell ELISA indicated no difference between control (1.15 ± 0.03 AFU) and ionomycin-stimulated cells (1.19 ± 0.03 AFU; n= 2). Similar results were obtained by immunofluorescence and confocal microscopy analysis (Fig. 4B).

Figure 4. Regulated exocytosis of MUC5AC from Calu-3 cells.

Calu-3 cells were stimulated for 10 min with either, vehicle (CONT), 10 μm forskolin (FSK) or 10 μm ionomycin (IONO) in 1.6 mm or 0 extracellular Ca²⁺, and were promptly fixed for immunostaining as in Fig. 3B. A, confocal microscopy xy images illustrating immunostaining with a MUC5AC monoclonal antibody (45M1, green). Insets, 3-dimensional reconstruction of a high-resolution xy confocal microscopy scanning of multiple planes comprising the whole cell. MUC5AC signal is associated with granules in control conditions (left), and non-granular structures following ionomycin addition (right; the green channel was enhanced to reveal MUC5AC staining in detail). Nuclei were stained with DAPI (blue); bar, 40 μm; inset bar, 8 μm. B, confocal microscopy images obtained by xy (upper panels) and xz galvo-stage (lower panels) scanning depicting immunostaining with a MUC1 polyclonal antibody (red); arrowheads indicate plasma membrane position; bar, 20 μm.

To verify that the reduction in granule-associated MUC5AC immunostaining in response to ionomycin reflected exocytosis of mucin-containing granules, slot blot analyses of Calu-3 cell apical and basolateral extracellular media were performed. As illustrated in Fig. 5A (left panel), the mucosal, but not the basolateral, medium of resting Calu-3 cells displayed strong MUC5AC immunoreactivity, which increased ∼8-fold after treatment of the cells with ionomycin (the absolute MUC5AC concentration could not be assessed because of the unavailability of purified standards of this mucin; Fig. 5A and B, left panels). Consistent with the results shown in Fig. 4, ionomycin-promoted MUC5AC secretion into the apical compartment was dependent on the presence of extracellular Ca²⁺ and inhibited by cytochalasin D (Fig. 5B, left panel). Thapsigargin, but not forskolin, mimicked the effect of ionomycin on MUC5AC secretion (Fig. 5A and B, left panels). The absence of effect of forskolin on MUC5AC secretion in Calu-3 cells is in agreement with studies showing lack of involvement of CFTR on mucin expression/secretion in native intestinal and airway epithelia goblet cells (Lethem et al. 1993; Mall et al. 2004; Kreda et al. 2005).

Figure 5. Secretion of MUC5AC and lysozyme from Calu-3 cells.

Aliquots from lumenal and basolateral baths from the experiments described in Fig. 4 were analysed by a slot blot immunodetection-based assay using 45M1 MUC5AC antibody, as indicated in the Methods. A, representative slot blot results of mucosal (AP) and basolateral (BL) samples from cells incubated with vehicle (CONT), forskolin (FSK) or ionomycin (IONO) (see Fig. 4 for details). For calibration, immunoreactivity displayed by the indicated volume of a MUC5AC-rich sample (purified MUC5AC standard was unavailable) and the indicated amount of a lysozyme standard is shown (ST). B, digital densitometry analysis of slot blots indicating changes in MUC5AC and lysozyme secretions in the mucosal (filled bars) and basolateral (open bars) baths of cells incubated under the indicated conditions. Thapsigargin (THAP, 1 μm) was added for 10 min, and cytochalasin D (CYTD, 5 μm) was added for 30 min before ionomycin addition. The results are expressed as percentage change relative to control samples (control, 100%), and are expressed as the mean ±s.e.m. from at least three experiments performed in quadruplicate; *P < 0.01.

The effect of Ca²⁺-triggered exocytosis on other secreted proteins was also examined. It has been reported previously that Calu-3 cells released lysozyme tonically onto the mucosal compartment, and that forskolin, and to a lesser extent thapsigargin, elicited enhanced lysozyme release, but only after a prolonged exposure of the cells to agonists (8 h (Dubin et al. 2004) and 0.5–2 h (Joo et al. 2004)). By performing slot blot analysis for lysozyme secretion, we observed that 10 min incubation in the absence of agonist resulted in release of 4.7 ± 1 and 3.5 ± 2 ng lysozyme cm⁻² to the mucosal and basolateral medium, respectively. Thus, Calu-3 cells appear to release lysozyme constitutively not only to the mucosal compartment but also to the basolateral medium. In contrast to the above-mentioned long-term effects of cyclic AMP and Ca²⁺ agonists, a short (10 min) exposure of Calu-3 cells to ionomycin or forskolin resulted in negligible changes in lysozyme secretion (Fig. 5A and B, right panels).

In sum, the results in Figs 4 and 5 indicate that MUC5AC, but not MUC1 or lysozyme, undergoes substantial Ca²⁺-regulated exocytosis in Calu-3 cells.

Ca²⁺-promoted release of ATP from Calu-3 cells

The extent to which mucin granule secretion was accompanied by nucleotide release was next investigated. To monitor ATP release in real time, the luciferin–luciferase assay was applied to cells bathed in a 28-μm high ASL thin film (44 μl cm⁻² culture), as recently reported (Okada et al. 2006). In non-stimulated Calu-3 cells, a baseline ATP concentration (0.5–3 nm) was established within 15–30 min after addition of the luciferase cocktail. Ionomycin promoted a robust increase in ATP release, which was detected as early as 10 s after stimulation (the earliest reproducible recordable time point, data not shown) and reached the maximum at ∼2 min (Fig. 6A). Thapsigargin also promoted rapid and robust mucosal release of ATP from Calu-3 cells. By contrast, no ATP release was detected in response to forskolin (Fig. 6A). Incubation of Calu-3 cells in nominally Ca²⁺-free medium or preincubating the cells with cytochalasin D, two conditions that inhibited exocytosis of FM 1-43-labelled granules and mucin secretion (Figs 2, 4, and 5), also reduced ionomycin-promoted mucosal ATP release (Fig. 6B). Ionomycin-promoted ATP release was not affected by the CFTR_inh-172 (Fig. 6B), which effectively blocked forskolin-stimulated Cl⁻ secretion in these cells (Fig. 6C).

Figure 6. Real-time assessment of Ca²⁺-promoted ATP release.

Polarized Calu-3 cells were incubated in the presence of a 28-μm high mucosal film (50 μl well⁻¹, 44 μl cm⁻²) containing the luciferin–luciferase cocktail. A, ATP was assessed in real-time in response to 5 μm ionomycin, 1 μm thapsigargin or 10 μm forskolin. B, quantitative illustration of mucosal ATP released from Calu-3 cells in response to 5 μm ionomycin added to cells in the presence of either 0 Ca²⁺ or 1.6 mm CaCl₂, or after a 30 min preincubation with 10 μm BAPTA (in 0 Ca²⁺), 4 μm bafilomycin A₁ (BAFI), 5 μm cytochalasin D (CYTD) or 10 μm CFTR_inh-172. The data indicate percentage changes in ATP levels (at peak values) relative to changes with ionomycin in 1.6 mm CaCl₂ (100%). The data are expressed as the mean ±s.d. from one experiment performed with quadruplicate samples. Similar results were obtained with n= 2–5 independent experiments. C, representative tracing of measurements of short circuit current (I_sc) in Calu-3 cells illustrating forskolin (10 μm)-promoted Cl⁻ secretion and its inhibition by 10 μm CFTR_inh-172.

To assess the possibility that a vesicular ATP pool contributed to Ca²⁺-stimulated ATP release in Calu-3 cells, the effect of bafilomycin A₁ was examined. Bafilomycin A₁, an inhibitor of the H⁺-ATPase that loads ATP into specialized granules in secretory cells (Bankston & Guidotti, 1996), markedly impaired ionomycin-promoted ATP release (Fig. 6B).

Ca²⁺-promoted depletion of putative ATP storage granules

Quinacrine and acridine orange are fluorescent dyes that bind to nucleic acids and ATP and have been extensively employed to label ATP storage granules in secretory cells (Mitchell et al. 1998; Bodin & Burnstock, 2001; Sorensen & Novak, 2001; Coco et al. 2003). Incubation of Calu-3 cells with quinacrine (or acridine orange) resulted in incorporation of the dye into granules in up to ∼40% of the cells (Fig. 7A, left). Labelling of Calu-3 cell granules with quinacrine was completely inhibited by bafilomycin A₁ (data not shown). Quinacrine-labelled granules were identified at the cell apex by confocal microscopy xz scanning (Fig. 7A) and were typically ∼1 μm in diameter (Fig. 7B). Addition of ionomycin resulted in a decrease in fluorescence of quinacrine-labelled granules. Ionomycin-induced loss of quinacrine-associated fluorescence was readily detected at 10 s post stimulation (Fig. 7B and C), but most fluorescence was released with a half-life of 2 min post stimulation (Fig. 7C and D). Ca²⁺-dependent loss of fluorescence was accompanied by disappearance of the corresponding granule image in the DIC confocal plane (Fig. 7B, lower panels). Scanning of alternative focal planes confirmed that quinacrine-fluorescent granules were absent from cells, strongly suggesting that decrease in fluorescence intensity reflected granule exocytosis (i.e. granule fusion with the plasma membrane and release of quinacrine) rather than laser photo-bleaching, or chemical quenching of the dye due to changes in luminal pH. As with ionomycin, addition of thapsigargin to quinacrine-labelled Calu-3 cells resulted in marked decrease of granule-associated fluorescence (Fig. 7D). Labelling Calu-3 cell granules with acridine orange resulted in similar results (data not shown).

Figure 7. Exocytosis of quinacrine-labelled granules.

Calu-3 cell monolayers grown on glass coverslips were loaded with quinacrine and monitored by confocal microscopy (see Methods). A, image composition of equivalent xz galvo-stage confocal microscopy recordings. Left, live Calu-3 cells displaying subapical quinacrine fluorescent granules. Right, to verify polarized morphology of Calu-3 cell monolayers grown on glass substrate, fixed cells were co-immunostained for syntaxins 3 (red) and 4 (green), which are selectively expressed in the apical and basolateral compartments of polarized epithelial cells, respectively. Arrowheads indicate the position of the apical plasma membrane; bar, 5 μm. B, time course of Ca²⁺-promoted loss of quinacrine fluorescence in a single cell. Cells were preincubated for 15 min in the presence of 10 μm ionomycin and 0 Ca²⁺, and images were taken just before (0 s) and after the addition of 1.6 mm CaCl₂. Upper and lower images depict an identical xy scanning focal plane (within the uppermost 3 μm of the cell) in the fluorescence and DIC channels (to identify granule position), respectively. Arrowheads indicate a granule displaying a fast fluorescence loss kinetics (i.e. loss of 90% of initial fluorescence at 10 s post stimulation); bar, 5 μm. C, data represent relative loss of fluorescence from individual granules (n= 10) displaying either fast (∼10 s, ♦) or slow (> 2 min, □) fluorescence loss kinetics. The results are expressed as the mean ±s.e.m. and are representative of two independent experiments performed in duplicate. D, quantification of overall granule-associated quinacrine fluorescence changes after 5 min treatment with vehicle (CONT, 100%), ionomycin (IONO) in either 0 or 1.6 mm extracellular Ca²⁺, or 1 μm thapsigargin (THAP). The data depict the mean ±s.e.m. of all granules recorded in two experiments performed with duplicate samples. *P < 0.01.

Ca²⁺-promoted vectorial release of ATP and UDP-sugars

To investigate the polarity of Ca²⁺-promoted ATP release from Calu-3 cells, etheno-derivatization and HPLC analysis was applied to samples from the mucosal and basolateral compartments. We have reported that this protocol quantifies ATP and its metabolites with subnanomolar sensitivity (Lazarowski et al. 2004b). In resting Calu-3 cells, ATP concentration in the mucosal medium was 2 ± 0.3 nm (Fig. 8A), and levels of ADP (41 ± 11 nm), AMP (46 ± 14 nm) and adenosine (17 ± 3 nm) were ∼10- to 20-fold higher than ATP levels (Fig. 8A). The robust accumulation of ADP, AMP and adenosine relative to ATP is consistent with previous findings that ATP metabolites are more stable than ATP on the mucosal surface of Calu-3 cells (Lazarowski et al. 2004b). Adenine nucleotide/nucleoside accumulation in the basolateral compartment of Calu-3 cells was modest relative to mucosal accumulations (Fig. 8A, and Lazarowski et al. 2004b). Addition of ionomycin to Calu-3 cells resulted in nearly 10-fold increase of mucosal ATP levels (18 ± 4 nm; Fig. 8A). ATP concentrations measured after ionomycin challenge probably underestimated the total mass of ATP released from stimulated cells, as suggested by the increase (although statistically not significant) of ADP, AMP and adenosine observed in ionomycin-treated cells (Fig. 8A). In contrast to the mucosal surface, ionomycin elicited no change in the level of ATP or ATP metabolites in the basolateral medium (Fig. 8A). As we have previously established that ATP metabolism rates at the basolateral and mucosal surfaces of Calu-3 cells are similar (Lazarowski et al. 2004b), the results in Fig. 8A strongly suggest that a specialized process associated with the apical but not basolateral plasma membrane is involved in Ca²⁺-promoted ATP release.

Figure 8. Vectorial release of adenine nucleotides and UDP-glucose (UDP-Glc) in polarized Calu-3 cells.

Cells were preincubated for 90 min in Hank's balanced salt solution followed by the bilateral addition of 5–10 μm ionomycin (filled bars) or vehicle (open bars). Incubations continued for an additional 5 min, and the resulting adenyl purines (A) and UDP-Glc (B) released into the extracellular solutions were quantified as described in the Methods. UMP (300 μm) was added to mucosal solutions, as indicated. The data represent the mean ±s.d. from at least three experiments performed in triplicate. *P= 0.01.

Our data collectively suggest a correlation between ATP release and granule exocytosis. To further examine the possibility that Calu-3 cells release nucleotides from the secretory pathway, we investigated whether ionomycin promoted the release of UDP-sugars. UDP-sugars serve as monosaccharide donors in oligo/polysaccharide chain elongation on exportable glycoproteins, glycolipids and proteoglycans. They are synthesized in the cytosol and translocated into the lumen of the endoplasmic reticulum and the Golgi apparatus (reaching concentrations of up to 20-fold higher than cytosolic levels (Hirschberg et al. 1998)). UDP-sugar/UMP antiporters, which are members of the SLC35 nucleotide sugar transporter family, translocate cytosolic UDP-sugars into the secretory pathway (Ishida & Kawakita, 2004). We reasoned that vesicle exocytosis mediating glycoprotein export (e.g. Ca²⁺-promoted mucin secretion) may provide a pathway for the release of molecules such as UDP-sugars, in addition to ATP. Therefore, we took advantage of a recently developed assay that quantifies UDP-Glc in the low nanomolar concentration range (Lazarowski et al. 2003b) to investigate whether ATP released from ionomycin-stimulated Calu-3 cells was accompanied by UDP-Glc release. Figure 8B illustrates that ionomycin promoted mucosal but not basolateral release of UDP-Glc from Calu-3 cells. The effect of ionomycin on UDP-Glc release was not affected by the presence of 300 μm UMP in the mucosal medium (Fig. 8B), arguing against the possibility that cytosolic UDP-Glc is translocated to the extracellular milieu via a SLC35 transporter. Bilateral addition of radiotracer UDP-[³H]Glc to Calu-3 cell cultures resulted in only minor hydrolysis of the radiolabelled nucleotide-sugar; 91 ± 2% and 89 ± 4% (n= 4) of the label was recovered unchanged in the mucosal and basolateral media, respectively, after 2 h. These results strongly suggest that, as shown above for ATP, mucosal versus basolateral differences in release rates rather than metabolism rates accounted for the selective detection of UDP-Glc in the mucosal compartment.

As Calu-3 cells constitute a mixed population of CFTR-expressing non-mucous cells and gel-forming mucin secreting goblet-like cells, we repeated key experiments in a more homogeneous goblet cell line (i.e. SPOC1 cells). SPOC1 cells are a well-defined rat airway goblet cell line, which secrete gel-forming mucins in response to P2Y₂ receptor stimulation (e.g. ATP or UTP) (Abdullah et al. 1997). We investigated whether P2Y₂ receptor-promoted mucin secretion in SPOC1 cells is associated with nucleotide release. Figure 9A indicates that, as expected, incubation of SPOC1 cells with the P2Y₂ receptor agonist ATP resulted in increased mucin secretion. Notably, this incubation also resulted in enhanced release of UDP-Glc (Fig. 9C). To assess ATP release in response to P2Y₂ receptor stimulation, UTP was used as the agonist and the extracellular accumulation of adenyl purines (ATP + ADP + AMP + adenosine) was quantified by etheno-derivatization. Figure 9B indicates that UTP promoted a significant increase in the extracellular accumulation of adenyl nucleotides in the lumenal bath of SPOC1 cells. By contrast, the P2Y₂ receptor-inactive nucleotide GTP failed to stimulate mucin, UDP-Glc or ATP release, indicating that these activities were regulated via P2Y₂ receptor and not ecto-nucleoside diphosphokinase-mediated generation of ATP (Fig. 9B).

Figure 9. P2Y₂ receptor-promoted mucin secretion and nucleotide release in goblet-like SPOC1 cells.

Cells were incubated for 30 min with 300 μm ATP (A and C) or UTP (B) and vehicle (A and C) or the P2Y₂ receptor-inactive nucleotide GTP (300 μm, B). The results represent the mean ±s.d. from one experiment performed in quadruplicate, and reproduced in at least two independent experiments. UDP-glucose and adenyl purines were assessed as described in Fig. 8 and mucins were measured by an enzyme-linked lectin assay as described in the Methods.

Discussion

Our study demonstrates that Calu-3 cells are competent for Ca²⁺-regulated exocytosis of gel-forming mucins (e.g. MUC5AC) and that mucin secretion from Calu-3 cells is accompanied by enhanced release of ATP and UDP-Glc. We have also shown that nucleotide release from an airway epithelial goblet cell model, SPOC1 cells, is associated with the primary function of these cells (i.e. mucin secretion).

ATP release is critical to ASL homeostasis, but the sources and mechanisms of nucleotide release are unknown. Airway epithelia have a complex cellular composition, and thus, several mechanisms and pathways may be involved in the release of nucleotides into ASL. Circumstantial evidence supports the involvement of both the secretory pathway and plasma membrane channels or transporters in the cellular release of nucleotides from non-excitatory tissues, but unambiguous evidence for either vesicular or conductive/transport mechanisms in airway epithelial cells is lacking.

We believe that it is unlikely that plasma membrane channels and/or transporters participated in the release of cytosolic ATP and UDP-Glc in Calu-3 cells. For example, CFTR, which is highly expressed at the apical membrane of Calu-3 cells (Figs 3 and 6), has been suggested to mediate ATP release in epithelial cells (Schwiebert, 1999). However, both our current (Fig. 6) and previously published data (Lazarowski et al. 2004b) illustrating that elevation of cyclic AMP levels and activation of CFTR, or inhibition of CFTR activity, did not affect the rates of ATP release from Calu-3 cells, argue against the involvement of CFTR in ATP release. These findings are consistent with those failing to show a role for CFTR in ATP release in well-differentiated human airway epithelial cell cultures (Donaldson et al. 2000; Lazarowski et al. 2004a; Okada et al. 2006; Button et al. 2007). Studies in glioma cells and leucocytes have suggested that connexin hemichannels may function as ATP channels (reviewed by Lazarowski et al. 2003a), but expression of connexin hemichannels on the apical plasma membrane of Calu-3 cells has not been reported. Moreover, removal of lumenal Ca²⁺, a condition favouring connexin hemichannel opening, resulted in no enhancement of ATP release in Calu-3 cells (Fig. 6B). In addition, a nucleotide channel/transporter could be inserted into the plasma membrane during Ca²⁺-triggered mucin secretion. Potential candidates to fulfil this function are members of the SLC35 family of Golgi UDP-sugar–UMP antiporters and their related ATP–AMP antiporters (Hirschberg et al. 1998; Ishida & Kawakita, 2004). However, these transporters appear not to be expressed/inserted in the plasma membrane of Calu-3 cells, as extracellular UMP (luminal antiporter substrate) failed to increase UDP-Glc release (Fig. 8).

Rather, our data suggest that Ca²⁺-regulated release of ATP from Calu-3 cells reflects a Ca²⁺-regulated exocytotic mechanism. First, Ca²⁺-mobilizing agents promoted robust ATP release and granule secretion, which were both selectively released onto the apical surface and inhibited by disruption of the actin cytoskeleton (Figs 4, 5, 6, and 8). Second, Ca²⁺-triggered ATP release was impaired by the v-ATPase inhibitor bafilomycin A₁, which causes ATP depletion inside secretory compartments by disrupting a vesicular H⁺-driven ATP transport (Bankston & Guidotti, 1996) (Fig. 6). Third, Ca²⁺-promoted ATP release was accompanied by enhanced release of UDP-Glc, a molecule actively transported (and concentrated) into the lumen of the secretory pathway (Hirschberg et al. 1998) (Figs 8 and 9). Last, Ca²⁺ mobilization following P2Y₂ receptor activation regulated mucin and ATP (and UDP-Glc) secretion from SPOC1 cells (Fig. 9) suggesting that the Ca²⁺ mobilization triggered by ionomycin in Calu-3 cells is likely to be a physiologically relevant stimulus.

Because nucleotide release in Calu-3 cells accompanies Ca²⁺-regulated MUC5AC secretion, but not MUC1 or lysozyme secretion, Ca²⁺-promoted nucleotide release appears to be selectively associated with secretion of gel-forming mucins (e.g. MUC5AC). An attractive hypothesis generated by our data is that nucleotides and gel-forming mucins are cotransported molecules within mucin secretory granules. Not only did mucin secretion and ATP release manifest the same polarity and modulation by pharmacological manoeuvres, but the kinetics of ATP release and mucin granule exocytosis were also comparable. Ionomycin-promoted ATP release was observed at the earliest measurable integration point (i.e. 10 s post stimulation), comparable to single granule loss of fluorescence in FM 1-43- and quinacrine-labelled cells (Figs 2 and 7). This time-frame is in accord with that reported for single mucin granule exocytosis, visualized by real-time video-imaging of airway goblet cells (Davis et al. 1992). It could be argued that ionomycin-promoted maximal ATP release (∼2 min, Fig. 6B) was faster than depletion of FM 1-43-labelled granules (5 min, Fig. 2B) and of the reported rate of cumulative mucin secretion (> 5 min; Davis et al. 1992; Conway et al. 2003). The rather slow diffusion rate of mucins (organized inside granules as a condensed meshwork embedded in a fluid phase; Perez-Vilar et al. 2006), as opposed to the fast diffusion rate of small cotransported molecules (e.g. ATP), may be a contributing factor for this apparent discrepancy. A similar phenomenon has been observed in studies of nucleotide release from dense-core granules in insulin-secreting cells (Obermuller et al. 2005) and of neurotransmitter release during ‘kiss-and-run’ mechanism of synaptic vesicles loaded with FM probes (tightly associated with the granule dense-core) (Cousin & Robinson, 1999).

We cannot rule out the possibility that nucleotides and mucins were released from different vesicular compartments within Calu-3 cells. Unequivocal evidence for ATP released from secretory granules has not been obtained due, in part, to problems of specifically labelling ATP-containing organelles. Acridine dyes have been employed often to fluorescently label ATP storage organelles in secretory cells (Bodin & Burnstock, 2001; Sorensen & Novak, 2001; Mitchell et al. 1998; Coco et al. 2003). We demonstrated that quinacrine (and acridine orange) labelled subapical granules in Calu-3 cells, in a bafilomycin A₁-sensitive manner. Furthermore, quinacrine-labelled (and acridine orange-labelled) granules were depleted in response to ionomycin or thapsigargin, mirroring ATP release rates (Fig. 7). One interpretation of these data is that acridine dyes labelled ATP storage granules competent for Ca²⁺-regulated exocytosis in Calu-3 cells. However, caution should be taken in interpreting these results because acridine dyes label acidic compartments within the secretory pathway, which may or may not store ATP.

The data from the homogeneous SPOC1 goblet cell line suggest that ATP and mucin are secreted from the same cell type (Fig. 9), a conclusion that could not be reached from studies with the heterogeneous Calu-3 cell line. Our studies of Calu-3 and SPOC1 cells, however, again do not distinguish between Ca²⁺-regulated release of mucin and ATP from a common vesicular compartment versus different Ca²⁺-regulated secretory vesicles, or indeed Ca²⁺-regulated vesicular secretion of mucin and release of ATP from the cytosol via an unknown Ca²⁺-regulated (plasma membrane-inserted) nucleotide transporter. Furthermore, nucleotides may be released from non-mucous (Calu-3) cells. Unambiguous assessment of the contribution of mucin secretory granules to nucleotide release awaits development of protocols for isolation of non-disrupted mucin granules, which remains a challenge (Singer et al. 2004).

Functional consequences of airway epithelial nucleotide release

There is general consensus that ATP and its metabolite adenosine promote Ca²⁺- and cyclic AMP-dependent MCC responses via activation of epithelial P2Y₂ and A_2b receptors, respectively (Tarran et al. 2006). Mucin, the principal polymeric species in the mucus matrix, is condensed inside secretory granules and is secreted without involving ion and water secretion from goblet cells (Verdugo, 1990; Davis et al. 1992). However, for effective mucin dispersion in the ASL, coordinated ion and water channel activities (mainly Na⁺ absorption and Cl⁻ secretion) are required to provide adequate water secretion onto the mucosal surface (Boucher, 2007). Our data suggest that nucleotide release that accompanies mucin exocytosis from goblet cells may provide the signalling for the activation of ion channels and ciliary beating on neighbouring ciliated cells that are necessary for mucin incorporation into the ASL. Regardless of the cellular source and compartments from which released nucleotides originate, it is worth noting that coordinated (regulated) release from separate compartments also provides a mechanism for insuring appropriate ion transport responses to fully hydrate and disperse newly secreted mucins.

Our findings that UDP-Glc is actively released with mucins from Calu-3 (Fig. 8B) and from SPOC1 goblet cells (Fig. 9) suggest a link between airway epithelial UDP-Glc release and mucin secretion. Notably, sputum specimens from CF patients, who exhibited airway inflammation and mucin hyper-secretion, contained remarkably high levels of UDP-Glc relative to specimens from healthy subjects (E.R. Lazarowski, S.H. Donaldson and R.C Boucher, unpublished results). However, the signalling actions of UDP-Glc in the lung are poorly understood. The Gi-coupled P2Y₁₄ receptor, the cognate receptor for UDP-Glc, is expressed at the mRNA level in human lung (Chambers et al. 2000), alveolar type II cells, BEAS-2B bronchial epithelial and A549 lung carcinoma cells (Muller et al. 2005). Addition of UDP-Glc to these cells resulted in robust and selective secretion of interleukin-8, a potent neutrophil chemo-attractant (Muller et al. 2005). Moreover, recent reports illustrated that P2Y₁₄ receptor is expressed both at mRNA levels and as a functional receptor in human neutrophils (Moore et al. 2003; Scrivens & Dickenson, 2006). Whether accumulation of UDP-Glc in ASL may occur as a result of mucin hypersecretion and whether ASL UDP-Glc promotes pro-inflammatory signalling in the lung remain to be investigated.

Calu-3 cells as a model of airway mucous cells

Initial studies suggested that acinar serous cells of the submucosal glands are the major site of CFTR expression in human airways (Engelhardt et al. 1992). Because of the relatively high levels of expression of CFTR observed in Calu-3 cells, they have been broadly used as a model of the airway submucosal gland acinar serous cell (e.g. Shen et al. 1994; Joo et al. 2004). However, this concept is at odds with the observation that Calu-3 cells produce gel-forming mucins (current study and Shen et al. 1994; Berger et al. 1999), and with recent reports illustrating that human native acinar serous cells express relatively low levels of CFTR (ciliated cells of the superficial and proximal glandular duct epithelia exhibit the highest level of CFTR expression in native human airways (Kreda et al. 2005)). Moreover, native acinar serous cells express electron dense granules (Shen et al. 1994), high levels of aquaporin 5 (Kreda et al. 2001, 2005) and low levels of MUC-1 (S.M. Kreda, unpublished results), characteristics that are not exhibited by Calu-3 cells (Figs 1 and 3). Our data may contribute to the resolution of this apparent conflict; that is, Calu-3 cell cultures display a heterogeneous cell composition. While up to 40% of the cells produce and secrete mucin granules, resembling airway goblet cells (Figs 1, 3, 4, and 5), the remaining (mucin granule-negative) cells express CFTR at the apical surface (Fig. 3). The presence of mucin-secreting cells within Calu-3 cell cultures was independent of the passage number and growth conditions (e.g. permeable supports versus glass coverslips). This cell culture heterogeneity was replicated with Calu-3 cells from diverse sources, such as our own repository at the UNC-CH Cystic Fibrosis Center (Huang et al. 2001), the UNC-CH Tissue Culture Facility, and the American type culture collection (ATCC). Collectively, these observations imply that differentiated Calu-3 cells do not resemble native submucosal acinar serous cells but represent a mix of phenotypes, with an important goblet cell component and a CFTR-expressing component of undefined lineage.

In conclusion, the results from our study suggest that regulated exocytotic nucleotide release is a feature of complex airway epithelia containing mucous (goblet) cells. We cannot distinguish between the hypotheses that this exocytotic process reflects the release of nucleotides and mucins together in mucin secretory granules, or the secretion of nucleotides and mucins from different, but coordinated and Ca²⁺-regulated, compartments. Regardless, coordinated nucleotide and mucin secretion is important for ASL physiology. As ASL nucleotides regulate ion channel activities in ciliated cells, nucleotide release occurring simultaneously with mucin secretion provides a signalling mechanism for ion and water transport (and cilia beating) necessary for mucin hydration and dispersion into the ASL.