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CELLULAR |
1 Department of Physiology
2 Department of Pathology and Laboratory Medicine, Division of Medical Genetics, University of Pennsylvania, Philadelphia, PA 19104, USA
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Abstract |
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(Received 12 March 2007;
accepted after revision 21 May 2007;
first published online 24 May 2007)
Corresponding author J. K. Foskett: Department of Physiology, B39 Anatomy-Chemistry Bldg, 414 Guardian Drive, University of Pennsylvania, Philadelphia, PA 19104–6085, USA. Email: foskett{at}mail.med.upenn.edu
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Introduction |
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A fundamental model of submucosal gland fluid secretion (reviewed in Ballard & Inglis, 2004) proposes that a primary watery secretion emanates from the serous acini, which then washes over the mucous cells and hydrates released mucin granules, creating a mixture of ion-rich fluid and mucus that accumulates in the collecting duct. The fluid and mucus are propelled to the airway surface epithelium by combined actions of hydrostatic pressure, cilial beating and contraction of surrounding myoepithelial cells. The complex morphology and small gland size have limited experimental studies of the functions of the various cell types that comprise the glands. Current data have been derived from studies of intact glands in vivo or in excised pieces of mucosa. As a result, the functions and relative contributions of the various individual cell types to fluid and mucus secretion are not well understood.
Submucosal gland serous cells may be the major site of expression in the lung of the cystic fibrosis (CF) transmembrane conductance regulator (CFTR) Cl– channel (Engelhardt et al. 1992; Jacquot et al. 1993), and thus may be crucial to CF (reviewed in Ballard & Inglis, 2004; Inglis & Wilson, 2005), the most common lethal genetic recessive disease in the United States. In CF, the major lethal pathology involves the lung, where morbidity is manifested as mucus plugging of airways and chronic inflammation and infection (Boucher, 2004). It has been proposed that defective salt, fluid and mucus homeostasis is the underlying physiological consequence of impaired CFTR function in CF. Because the relative contributions of surface epithelial cells and submucosal glands to overall lung fluid homeostasis are unknown, it has been hypothesized that defective fluid homeostasis in CF may be partly due to defects in the amount or composition of submucosal gland fluid secretions (reviewed in Ballard & Inglis, 2004; Inglis & Wilson, 2005). The importance of CFTR in the secretory functions of submucosal glands is supported by observations of occluded mucus-filled gland ducts, as well as gland hypertrophy, hyperplasia and infection in lungs of CF patients (Ornoy et al. 1987; Jeffery & Brain, 1988). Pharmacological inhibition of glandular fluid secretion in porcine airways mimics some of the manifestations of CF pathology, including disrupted mucociliary clearance without reduction in ciliary beat frequency (reviewed in Ballard & Inglis, 2004). These results suggest that inhibition of glandular fluid secretion may be sufficient to cause CF-like pathology, independent of possible hyper-absorption of fluid by surface epithelial cells in the absence of CFTR function. Since glands contribute significantly to fluid secretion in large airways, it has been hypothesized that the absence of a functional CFTR Cl– conductance in CF glands may lead to defective salt and water secretion and contribute to CF lung pathology (reviewed in Ballard & Inglis, 2004; Inglis & Wilson, 2005).
Submucosal glands in airway tissue from individuals with CF appear to be defective in their ability to secrete fluid in response to agonists such as vasoactive intestinal peptide (VIP) that elevate intracellular concentrations of cyclic-AMP ([cAMP]i) (Joo et al. 2002a; Salinas et al. 2005; Song et al. 2005). However, freshly isolated CF glands still secrete significant volumes of fluid in response to cholinergic agonists that increase intracellular Ca2+ concentration ([Ca2+]i) (Joo et al. 2002a). In these studies, the cAMP-stimulated pathway appears to be the minor pathway of the two, as VIP induced sustained glandular secretion at a maximal rate that was only 
30–40% of the secretion rate elicited by cholinergic stimulation (Trout et al. 2001; Joo et al. 2001b, 2002a,b). Thus, it is not obvious how diminished cAMP-dependent secretion contributes to CF pathology. The implications of observations of intact glands are not clear, largely because there are few data regarding the cellular and molecular mechanisms that underlie gland secretion. This knowledge is not only critical for our understanding of the general airway pathology of CF and other lung diseases, but would also be very important for the development of therapeutic agents that may target submucosal glands.
The experiments described below represent a significant step toward filling this important gap by helping to elucidate the molecular mechanisms of submucosal gland serous cell ion and fluid transport. Submucosal serous cells from the mouse airway were isolated, identified and studied using simultaneous bright-field differential interference contrast (DIC) imaging of cell volume and quantitative fluorescence microscopy of ion indicator dyes, to examine the molecular mechanisms of serous cell ion transport and its regulation in single cells. These results provide insights into the molecular mechanisms of regulated fluid secretion in murine submucosal gland serous cells, and they suggest that secretion can be mediated by Cl– efflux through a non-CFTR anion conductance in murine serous acinar cells that express CFTR.
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Methods |
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AlexaFluor (AF)-labelled anti-mouse and anti-rabbit IgG secondary antibodies, 6-methoxy-N-(3-sulfopropyl)quinolinium (SPQ), ionomycin, A23187, and acetoxymethylester (AM)-derivatives of fura-2, 1,2-bis(o-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA), and 2',7'-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein (BCECF), and calcein were purchased from Molecular Probes (Eugene, OR, USA). RNasin RNase inhibitor was purchased from Promega (Madison, WI, USA). All other molecular biology reagents were from Invitrogen (Carlsbad, CA, USA). CFTRinh172 was a gift from Dr A. S. Verkman and also purchased from Calbiochem (San Diego, CA, USA). Anti-CFTR 24–1 monoclonal antibody was obtained from R & D Systems (Minneapolis, MN, USA). Anti-lysozyme polyclonal antibody was purchased from Dakocytomation (Carpinteria, CA, USA). Recombinant CFTR C-terminal peptide was a kind gift from Dr V. Raghuram (NIH/NHLBI; Bethesda, MD, USA). Microscope filters were obtained from Chroma Technologies, Inc. (Rockingham, VT, USA). All other reagents were obtained from Sigma (St Louis, MO, USA).
Submucosal gland serous acinar cell isolation
All animal handling procedures were performed in accordance with regulations of the Institutional Animal Care and Use Committee (IACUC) of the University of Pennsylvania. Nasal turbinate and septum were obtained from wild-type (WT), cftrtm1Unc–/– knockout, or heterozygote mice on a congenic C57BL/6 strain background housed in a pathogen-free facility after killing the animals by CO2 asphyxiation and cervical dislocation. Isolated tissue was first placed in a physiological saline buffer containing (mM): 125 NaCl, 5 KCl, 1.2 MgCl2, 1.2 CaCl2, 1.2 NaH2PO4, 11 glucose, 15 Hepes, pH 7.4. The tissue was mechanically minced with scissors and then incubated for 25 min at room temperature in medium containing (mM): 125 NaCl, 5 KCl, 1.2 MgCl2, 1.2 NaH2PO4, 11 glucose, and 25 NaHCO3, gassed with 95% O2–5% CO2, supplemented with 0.8 mg ml–1 collagenase (Worthington Type II), 2 mM L-glutamine, 1 x MEM-vitamins, 1 x MEM-amino acids, and 1% BSA. After washing via gentle centrifugation, cells were plated on glass coverslips and allowed to adhere for 10–20 min. The isolation protocol yielded acini, single cells and strings of cells. Cells were identified based on visible morphology (size, polarized secretory granules, acinar structures) under DIC optics using a 40x oil-immersion objective lens (Nikon S Fluor 1.3 NA) on an inverted Nikon microscope. WT and cftrtm1Unc–/– mice used in these experiments ranged in age from 3 to 12 months and 3 to 7 months, respectively. No age-dependent differences were observed in cell viability (assessed by visible morphology and responses to agonists), yield, or responses to agonist (volume, [Ca2+]i, or [Cl–]i) between WT and cftrtm1Unc–/– mice.
Confocal immunofluorescence microscopy
After isolation, cells were plated for 30 min on poly L-lysine (poly K) coated coverslips (1–2 mg ml–1) and fixed in 4% formaldehyde for 20 min at 4°C. Blocking/primary antibody incubation was performed overnight at 4°C in Dulbecco's phosphate buffered saline (DPBS; with Ca2+ and Mg2+) containing 1% BSA, 2% goat serum, and 0.1% saponin. Following washing (3 x 5 min in DPBS), cells were incubated with AlexaFluor (AF)-conjugated secondary antibodies for 2 h at 4°C. After washing, cells were mounted in DPBS containing 40% glycerol and 1 mg ml–1 p-phenylenediamine. Imaging and excitation of fluorescently labelled secondary antibodies was performed using the 488 and 568 nm lines of an Ar/Kr-ion laser (50% intensity) attached to a Perkin-Elmer spinning-disc confocal system coupled to an inverted Nikon microscope equipped with a 60x oil-immersion objective lens (Nikon Plan Apo 1.4 NA). Imaging and image overlays were performed using Perkin-Elmer Ultraview software.
Cell harvesting/aRNA amplification
Cells were plated on poly K-coated coverslips for 15 min and then gently washed to remove free-floating debris. Cells were harvested via gentle suction with a patch-clamp pipette with a 5 µm diameter opening. Care was taken to aspirate three- to four-cell structures of morphologically distinct acinar cells and to avoid contamination from surrounding cells and/or cellular debris. The cells were ejected into a solution of 15 mM DTT and 5–6 U µl–1 RNasin RNAse inhibitor and frozen overnight at –80°C. Three- to five-cell sheets of ciliated epithelial cells were isolated using identical procedures. Two rounds of antisense RNA (aRNA) amplification were carried out with a poly dT-oligo containing a T7 RNA polymerase promoter (Van Gelder et al. 1990). Control RNA was isolated from nasal turbinate, brain and kidney using TRIzol reagent according to the manufacturer's protocol. Reverse transcription PCR (RT-PCR) was carried out for 30 cycles, using denaturation, annealing, and extension temperatures of 94°C, 57°C and 72°C, respectively. Primers and expected product sizes are listed in online supplemental material, Supplemental Table 1.
Cell volume determinations
Cells were imaged under continuous perfusion with the HCO3– buffer described above (except lacking collagenase, vitamins, amino acids, glutamine and BSA but containing 1.2 mM CaCl2) at 37°C (chamber volume 300 µl, flow rate 2 ml min–1) with continuous gassing. Experiments performed in solutions lacking extracellular Ca2+ (0-Ca2+ solution) were done in the same buffer with the 1.2 mM CaCl2 omitted and 1 mM EGTA added. To avoid CO2 loss, the perfusion solutions were kept in heated overhead glass reservoirs connected by glass tubing to glass heating coils near the perfusion chamber. Changes in the extracellular solution were controlled by an electronic pinch-valve system (Warner Instruments; Hamden, CT, USA). No differences in the magnitude of cell volume changes were observed between single acinar cells and small acini composed of two to four cells. Large acinar clumps exhibited qualitatively similar volume responses but their complex structure prevented quantitative volume determinations. Since the isolated cells and small acini were approximately spherical and volume changes occurred in the same proportion in all directions (described below and in Supplemental Fig. 1), the cell outline (traced using ImageJ software; W. Rasband, NIH/NIMH, Bethesda, MD, USA) was used to determine volume by taking the cross-sectional area of the centre of the imaged cell raised to the power 3/2. Volume at time 0 (Vo) was normalized to 1, and relative volume expressed as V/Vo. All data are reported as means ± standard error of the mean (S.E.M.) unless otherwise noted. All graphing and linear regression fits were performed with Igor Pro software (Wavemetrics, Inc., Lake Oswego, OR, USA). P-values were determined in Excel using Student's two-tailed t test.
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Acinar cells were loaded with calcein by incubation in 2 µM calcein-AM for 10–15 min, followed by incubation in calcein-free medium for 5–10 min to allow cells to recover/de-esterify the loaded dye. Cells were illuminated with the 488 nm line of an Ar/Kr laser attached to a Perkin-Elmer spinning disc confocal Nikon microscope equipped with a 40x, 1.3 NA objective lens and Prior Optiscan z-stepper focal motor controlled by Ultraview software. Fluorescence was collected by a CCD camera after passing through a 525/50 nm band-pass (bp) filter. Images were taken with a 0.5 µm z-step size. Images were taken in 305 mosmol l–1 extracellular solution and after the medium was supplemented with 100 mM sucrose (407 mosmol l–1 by vapour pressure osmometry), since it was observed that this concentration induced a cell shrinkage similar to the maximal shrinkage observed during exposure to 100 µM CCh (as measured by DIC; see Supplemental Fig. 2A). Image stacks were rendered into 3-D volumes and measurements were made of cell volume and x-, y-, and z-axis diameter using Volocity software (Improvision, Lexington, MA, USA).
Simultaneous single cell volume and [Ca2+]i determinations
Cells were loaded with the Ca2+-sensitive dye fura-2 by incubation in a medium containing 0.5–1 µM fura-2-AM for 10–15 min, followed by 5 min incubation in fura-2-free solution to allow cells to recover and to de-esterify the loaded dye. Simultaneous DIC and fura-2 fluorescence imaging was performed as previously described (Foskett, 1988, 1990b; Foskett & Melvin, 1989) except the analyser was housed in the emission filter wheel and moved out of the light path during measurements of fluorescence. To minimize toxicity, fluorescence was sampled as infrequently as possible while still accurately tracking changes in [Ca2+]i. The ratio of emitted light upon excitation with 340 nm and 380 nm light was used to determine [Ca2+]i, as described (Foskett & Melvin, 1989).
Simultaneous single cell volume and [Cl–]i determinations
Cells were loaded with SPQ by incubation in medium containing 10 mM SPQ for 30–40 min. SPQ fluorescence was excited with a 340/10 nm filter, with emission collected with a 450/50 nm filter. Simultaneous DIC and SPQ fluorescence imaging was performed as described (Foskett, 1990a) except that DIC illumination was filtered at 530/30 nm. SPQ properties were calibrated in vivo in a separate group of calibration cells as previously described (Foskett, 1990a). Osmotic shrinkage of serous acinar cells was accomplished using Hepes-buffered solutions containing (mM) 150 K+, 65 Cl– (determined to be the resting [Cl–]i, described below), 85 gluconate, supplemented with nigericin and tributyltin (to clamp [Cl–]i at 65 mM), and containing varying sucrose concentrations (0, 50, 100, or 150 mM). Acinar cells were exposed to the 0-sucrose nigericin–tributyltin solution (308 mosmol l–1 by osmometry) until a stable SPQ fluorescence level was achieved, followed by exposure to one of three hyperosmotic solutions (356, 407 and 453 mosmol l–1) to induce cell shrinkage while keeping [Cl–]i clamped at 65 mM. Due to possible effects of volume regulatory mechanisms, each experiment was limited to one hyperosmotic exposure. The cells shrank rapidly (within 10–20 s) upon exposure to hyperosmotic medium and remained shrunken until re-exposure to 308 mosmol l–1 solution, upon which they swelled back to resting volume.
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Results |
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Serous and mucous cells isolated from the lung are difficult to culture, and they rapidly de-differentiate and begin to express markers of the opposite cell type (Marin & Culp, 1986; Sommerhoff & Finkbeiner, 1990; Yamaya et al. 1991; Chopra et al. 1994; Emery et al. 1995). Consequently, data obtained from cultured airway gland cells must be interpreted with caution. The Calu-3 cell line has been frequently used as a surrogate model for serous cell function, because it is a lung-derived secretory cell that expresses high levels of CFTR (Shen et al. 1994), and Calu-3 cells have proven valuable for understanding CFTR-mediated fluid secretion. However, this carcinoma-derived cell line is anneuploid and expresses some markers of goblet cells while lacking some serous cell markers (Duszyk, 2001; Dubin et al. 2004), indicating that extrapolation of data obtained from Calu-3 cells to submucosal gland serous cell function must also be done cautiously. To examine fluid transport properties of lung submucosal gland acinar cells directly, we developed a protocol to isolate living primary submucosal gland cells from murine nasal turbinate and septum. The mouse was chosen because it is currently the only animal available with CFTR mutant and knockout models (reviewed in Grubb & Boucher, 1999), which was felt to be valuable for deciphering the role of CFTR in submucosal gland function.
In mice, submucosal glands are present only in the nasal cavity and the upper cartilaginous rings of the proximal trachea (Yamaya et al. 1991; Song & Verkman, 2001; Widdicombe et al. 2001). Preliminary experiments determined that acinar cells were more efficiently isolated from nasal tissue than from tracheal tissue, and therefore nasal turbinate and septum were used as a source of submucosal gland cells. Previous immunohistochemical characterization (Groneberg et al. 2003; Martinez-Anton et al. 2006), as well as functional measurements of intact human nasal gland secretion (Salinas et al. 2005; Song et al. 2005) have indicated no significant differences in the properties of upper airway and tracheal or bronchial glands.
Serous cells were first identified by the appearance of small apically localized secretory granules, their ‘smooth’ basolateral side, and the acinar morphology of isolated cell clumps (Fig. 1A). To confirm the morphological identification, confocal microscopy and immunocytochemistry were employed using antibodies directed against CFTR and serous cell-specific markers (Klockars & Reitamo, 1975). Fixed acinar cells displayed bright immunofluorescence at the apical region using anti-CFTR monoclonal antibody 24–1 (Fig. 1B), which was substantially reduced when purified recombinant C-terminal CFTR peptide (peptide sequence N-(LC-Biotin)-QIAALKEETEEEVQDTRL-C, containing the 24–1 epitope) was added to the primary antibody incubation solution (Fig. 1E). In contrast, the CFTR peptide did not affect lysozyme immunofluorescence (below), indicating that the reduced CFTR staining was caused by competition of the peptide for specific antibody binding. CFTR immunofluorescence was absent in cells isolated from cftrtm1Unc–/– mice (Fig. 1C), confirming that the antibody detected CFTR specifically. The morphologically identified isolated acinar cells also displayed lysozyme immunofluorescence (Fig. 1D) that consistently appeared to be localized to the secretory granules, and was reduced to near-background levels when 0.2 mg ml–1 recombinant human lysozyme was added to the primary antibody incubation solution (not shown). Ciliated epithelial cells from WT mice displayed CFTR immunofluorescence at the apical membranes (Fig. 1B and G) but did not display lysozyme immunofluorescence (Fig. 1G), supporting specific acinar localization.
To further confirm the identity of the cells that we believed were CFTR-expressing serous acinar cells, three- to four-cell acini were harvested for mRNA amplification via the Eberwine antisense RNA (aRNA) method (Van Gelder et al. 1990; reviewed in Eberwine, 2001). Because this method uses the mRNA poly A tail for amplification (via an oligo dT primer), it selectively amplifies expressed mRNA and not genomic DNA. This sensitive method allows amplification and detection of as few as two to three copies of a single mRNA species. After two rounds of amplification (106-fold), reverse transcription (RT)-PCR revealed expression of mRNA sequences for CFTR, NKCC1 and lysozyme in the cells identified as serous acinar cells (n
= 4 reactions from three WT mice; Fig. 2A). Neither the 
-subunit of the epithelial sodium channel (ENaC) nor CaBP1 (a neuronal Ca2+ binding protein) was detected. RNA amplification performed on two to three isolated ciliated cells revealed expression of mRNA transcripts for CFTR and NKCC1 as well as ENaC, but not lysozyme (n
= 4 reactions). As a control, aRNA amplified from salivary gland acinar cells revealed expression of CFTR, NKCC1 and lysozyme mRNA, but not ENaC mRNA, as expected. To validate the integrity/function of all primers, RT-PCR was performed on RNA extracted from murine nasal turbinate tissue. CFTR, NKCC1, lysozyme, lactoferrin and ENaC transcripts were detected, but not mRNA for the neuronal CaBP1. RNA extracted from mouse brain revealed expression of CFTR, NKCC1, CaBP1, lysozyme and ENaC transcripts, as expected. No PCR products were detected in control reactions run for all primer sets and RNA samples with the reverse transcription step omitted (data not shown), suggesting that the products detected were not amplified contaminating genomic DNA.
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Serous acinar cells respond to cholinergic stimulation with a rapid decrease in cell volume
Submucosal glands are cholinergically innervated (Yu et al. 1989; Hejal et al. 1993; reviewed in Rogers, 2001a), and the greatest glandular response to fluid-secreting agonists is seen upon cholinergic stimulation. The responses of isolated acinar cells to the acetylcholine (ACh) agonist carbachol (CCh) were examined using DIC imaging to track changes in cell volume. The rationale for imaging cell volume is based on the fact that water is almost always at equilibrium across membranes, so changes in cell solute content as a consequence of alterations of ion transport rates result in parallel changes of cell volume. Real time measurements of cell volume, combined with appropriate pharmacology and ion substitutions, can provide insights into the molecular mechanisms of ion transport. Furthermore, when combined with simultaneous measurements of ion-indicator dyes, this method affords an approach to correlate concentrations of transported solutes and signalling molecules with secretion in intact cells. Optical studies of cell volume combined with simultaneous quantitative low-light-level fluorescence microscopy using ion indicator dyes has been successfully employed in primary exocrine cells to characterize the molecular mechanisms involved in cholinergic-induced salivary gland ion and fluid secretion (Foskett, 1988, 1990a; Foskett & Melvin, 1989). Confocal three-dimensional reconstruction of isolated calcein-loaded serous acinar cells indicated that the cells behaved as spheres that changed size approximately equally in all dimensions (Supplemental Fig. 1) validating the use of changes in the area of a single optical section to track changes in cell volume. Stimulation with CCh caused cell volume to decrease by 20 ± 2% (volume/original volume (V/Vo) = 0.80 ± 0.02) within 62 ± 5 s (n = 11; Fig. 3). Upon agonist removal, the cells swelled back to the resting volume. We hypothesized that these changes in cell volume were reflective of changes in ionic content in the isolated serous cells as a consequence of changes in their secretory state, and thus set out to determine the utility of this technique for monitoring serous cell fluid secretion.
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Submucosal gland fluid secretion in response to cholinergic stimulation is believed to result from agonist-induced increases in [Ca2+]i (reviewed in Ballard & Inglis, 2004). To examine the role of [Ca2+]i in the observed CCh-induced cell volume responses, an inverted microscope was modified to carry out simultaneous DIC and fluorescence microscopy of the ratiometric Ca2+ indicator dye fura-2 (Fig. 4A). Resting [Ca2+]i was 89 ± 2 nM (n
= 19). Stimulation with 100 µM CCh caused [Ca2+]i to rapidly increase to 447 ± 15 nM within 25 ± 2 s (n
= 19; Fig. 4B), typically followed by a plateau phase of increased [Ca2+]i (240 ± 10 nM; n
= 15). Upon washout of agonist, [Ca2+]i relaxed to resting levels (Fig. 4B). Reapplication of CCh caused a similar spike (468 ± 15 nM within 25 ± 2 s) and plateau (230 ± 6 nM) of increased [Ca2+]i (n
= 15). These agonist-induced changes in [Ca2+]i were associated with profound changes in cell volume observed simultaneously by DIC microscopy (Fig. 4C). The initial CCh-induced peak of increased [Ca2+]i was accompanied by a maximal decrease in cell volume of 21 ± 1% within 62 ± 3 s (n
= 19 fura-2-loaded cells; Fig. 4B). Initiation of cell shrinkage occurred after the onset of the rapid rise of [Ca2+]i. Although the sustained [Ca2+]i response to CCh was somewhat variable between cells, cell volume was nevertheless tightly correlated with [Ca2+]i. Thus, an elevated plateau phase of [Ca2+]i was associated with sustained cell shrinkage, whereas more substantial relaxation of [Ca2+]i toward resting levels was accompanied by swelling of cell volume toward resting levels. In 20% of cells and acini, stimulation with 100 µM CCh caused only a transient increase of [Ca2+]i (Fig. 4D). The transient rise of [Ca2+]i was accompanied by a transient cell shrinkage (Fig. 4D). In this event, wash out and reintroduction of extracellular Ca2+ in the continued presence of CCh caused another transient spike of [Ca2+]i that induced another transient cell shrinkage.
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To examine the role of extracellular Ca2+, serous acinar cells were stimulated in a Ca2+-free solution containing no added Ca2+ and 1 mM EGTA (0-Ca2+ solution). Stimulation with 100 µM CCh in 0-Ca2+ solution caused a transient peak increase in [Ca2+]i (283 ± 6 nM within 31 ± 3 s; n = 10) that returned to resting levels (below 100 nM) within 190 ± 25 s (Fig. 6A). This was accompanied by a transient cell shrinkage (21 ± 1% within 62 ± 5 s) followed by swelling back to resting volume within 200 ± 11 s. In the continued presence of CCh, reintroduction of extracellular Ca2+ caused a sustained increase in [Ca2+]i and associated sustained cell shrinkage (Fig. 6A). In a second protocol (Fig. 6B), cells were similarly stimulated in 0-Ca2+ solution, causing a transient rise of [Ca2+]i and cell shrinkage. Washout and re-application of CCh was without effect on [Ca2+]i, indicating that intracellular Ca2+ stores had been depleted by the first exposure to CCh in the absence of extracellular Ca2+. This lack of CCh-induced [Ca2+]i signal was associated with no change of cell volume (Fig. 6B). In contrast, reintroduction of Ca2+ during a third exposure to CCh induced a rapid increase in [Ca2+]i accompanied by cell shrinkage (Fig. 6B). These data suggest that influx of extracellular Ca2+ is required for both the sustained increase of [Ca2+]i and the sustained cell shrinkage. After depletion of [Ca2+]i by prolonged CCh stimulation (> 300 s) in 0-Ca2+ solution and subsequent washout of CCh, reintroduction of [Ca2+]o alone was sufficient to cause a small transient increase in [Ca2+]i that caused a transient decrease in cell volume (Fig. 6C). This suggests that the increased [Ca2+]i caused by Ca2+-influx across the plasma membrane is sufficient to induce cell shrinkage independently of the presence of an agonist.
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We hypothesized that CCh increases [Ca2+]i via binding to a muscarinic receptor, based on previous evidence of M1 and M3 receptor expression in submucosal glands (Ishihara et al. 1990; reviewed in Rogers, 2001a). Muscarinic activation of receptor-coupled Gq proteins leads to production of InsP3 and release of Ca2+ from the endoplasmic reticulum via the InsP3 receptor. In agreement, atropine (100 µM), a competitive muscarinic receptor antagonist, blocked the normally sustained increase in [Ca2+]i stimulated during exposure to 10 µM CCh, causing both [Ca2+]i and cell volume to return to resting levels (Fig. 6F).
Together, the above data suggest that an increased [Ca2+]i is necessary to induce cell shrinkage. To determine whether elevated [Ca2+]i was sufficient, cells were treated with the calcium ionophores ionomycin or A23187. Both ionophores raised [Ca2+]i and induced similar cell shrinkage (Fig. 6G; maximum shrinkage was 25 ± 3% with ionomycin and 24 ± 2% with A23187; n
= 3 experiments with each ionophore). After application of either ionophore, the extent of cell shrinkage achieved a plateau despite the fact that [Ca2+]i continued to rise, suggesting that a saturating concentration of [Ca2+]i was reached and that a 20–25% volume decrease is a maximal response.
Histamine and ATP induce similar [Ca2+]i-dependent volume responses in murine serous acinar cells
To determine if other receptors similarly mobilized Ca2+ and induced cell-volume responses, acinar cells were perfused with solutions containing either histamine or ATP. Histamine is a local mediator of inflammatory responses; its release during inflammation in CF lungs could possibly stimulate gland secretion via serous cell H1 receptors. Histamine (100 µM) increased [Ca2+]i (peak 350 ± 16 nM within 25 ± 2 s; plateau 253 ± 5 nM; n = 9) that was accompanied by cell shrinkage (maximal shrinkage was 21 ± 1% within 62 ± 4 s) comparable to that elicited by CCh (Fig. 7A). ATP is a coneurotransmitter in adrenergic and cholinergic neurons and is also released from airway epithelial cells, where it may play a role in ASL homeostasis (Donaldson et al. 2000; Lazarowski et al. 2004). Exposure to 100 µM ATP elicited similar changes in [Ca2+]i and cell volume to that observed with CCh (peak [Ca2+]i = 348 ± 14 nM within 26 ± 3 s; plateau [Ca2+]i = 249 ± 3 nM; maximal shrinkage was 21 ± 1% within 57 ± 3 s; n = 8 (Fig. 7B). While the peak [Ca2+]i increase with both agonists was somewhat less than that observed with 100 µM CCh, both agonists increased [Ca2+]i sufficiently to result in a significant sustained cell shrinkage. We conclude that cell shrinkage is a generalized response of submucosal gland serous acinar cells in response to a variety of secretagogues that signal through changes in [Ca2+]i.
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We hypothesized that the observed changes in cell volume upon CCh-stimulation were due to changes in intracellular solute content caused by [Ca2+]i-dependent alterations of ion permeabilities, as previously shown in salivary acinar cells (Foskett, 1990a). However, other possible explanations exist, including loss of cell content during exocytosis of secretory granules, internalization of the plasma membrane, or cell shrinkage via contractile mechanisms. To determine if cell volume changes were caused by changes in cell solute content, we combined optical measurements of acinar cell volume with a fluorescent indicator to track the solute content of the cell. As discussed in Foskett (1990b), because Cl– is not the overwhelmingly predominant anion in cells (typically > 50% of cellular anionic content is made up of organic membrane-impermeant molecules), changes in Cl– content will be reflected in changes in [Cl–]i. Accordingly, changes in cell volume should be associated with changes in cell Cl– content that would be reflected in corresponding changes in [Cl–]i. To test this hypothesis, cell volume and [Cl–]i were monitored by simultaneous DIC imaging and fluorescence imaging of the halide-sensitive fluorophore SPQ. SPQ fluorescence is quenched by Cl– via a collisional mechanism, leading to a linear reciprocal relationship between its fluorescence and [Cl–].
CCh-induced cell shrinkage was paralleled quantitatively by increases in SPQ fluorescence (plotted inversely as fluorescence normalized to fluorescence at time 0 (F/Ft=0; Fig. 8A). In SPQ-loaded cells, 100 µM CCh caused a 20 ± 2% decrease in cell volume that was accompanied by a 55 ± 2% increase in SPQ fluorescence (n = 9). Under all conditions, cell volume was correlated tightly with the magnitude of SPQ fluorescence. Thus, partial recovery of cell volume towards prestimulation levels was associated with parallel decrease in SPQ fluorescence, and full recovery of cell volume following removal of CCh was associated with a decrease of SPQ fluorescence to prestimulation levels.
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20% (Supplemental Fig. 2B), a value that agrees with observations in other mammalian cells (discussed in Foskett, 1990b). The observed CCh-induced changes in SPQ fluorescence and their tight correlation with cell volume strongly suggested that agonist-induced changes in cell volume are caused by changes in cell solute content, reflected by quantitative changes in [Cl–]i. To quantify [Cl–]i, SPQ fluorescence was calibrated using the tributyltin and nigericin along with solutions of known [Cl–] to clamp [Cl–]i = [Cl–]o and [pH]i = [pH]o in a separate group of calibration cells (as described in Foskett, 1990a). Six calibration experiments were performed (an example is shown in Fig. 8B), with each consisting of two exposures of a cell or acinus to a range of [Cl–]. Only cells in which similar fluorescence intensities were observed during the first and second exposures were used, since these had no dye leakage during the course of the calibrations. The Stern–Volmer plot (Fig. 8C) indicated that the quenching constant (Ksv; the slope of the linear fit) was 17.6 M–1, nearly identical to the quenching constant of 17 M–1 previously measured in salivary gland acinar cells (Foskett, 1990a) and similar to the reported values of 13 M–1 found for both canine tracheal epithelial cells (Chao et al. 1990) and fibroblasts (Chao et al. 1989). Extrapolation of the average SPQ fluorescence intensity in the cells used in the calibration experiments before their exposure to the nigericin–tributyltin solutions indicated that resting [Cl–]i in isolated murine serous acinar cells is 64.5 ± 4 mM. Using this value and the Stern–Volmer plot, average changes in SPQ fluorescence during agonist stimulation were converted to quantitative changes in [Cl–]i. The average increase in SPQ fluorescence of 55% upon stimulation with CCh corresponds to a decrease of [Cl–]i from 64.5 in unstimulated cells to 28 mM during the peak cell shrinkage. The relationship between [Cl–]i and cell volume was plotted in aggregate for nine cells during CCh-induced stimulation (Fig. 8D). The linear relationship demonstrates that cell volume quantitatively tracks [Cl–]i. Thus, cell volume is a quantitative measure of cell solute content as well as [Cl–]i. Importantly, these results establish that cell volume changes are caused by changes in cell solute content.
K+ efflux is required for cell shrinkage and [Cl–]i decrease
Because the results suggested that changes in cell volume and Cl– content are tightly coupled, it is expected that a similar quantity of cation must accompany net Cl– fluxes to preserve electroneutrality. Because K+ is the predominant cation inside cells, we hypothesized that cell shrinkage predominately reflected loss of KCl. We therefore examined whether K+ efflux contributed to agonist-evoked cell shrinkage. As previously discussed (Foskett, 1990a), isosmotic loss of KCl will be expected to result in only negligible changes in [K+]i, and thus measurements of [K+]i would not be informative. Instead, we examined the effects of changing the driving forces for K+ efflux. When [K+]o in the medium was increased from 5 mM to 85 mM (by isosmotic replacement of Na+), normal CCh-induced cell shrinkage was prevented and no change in SPQ fluorescence was observed (Fig. 9A). This result is consistent with a requirement of K+ efflux during agonist-induced cell shrinkage. However, because exposure of the cell to 85 mM K+ is likely to have depolarized the plasma membrane potential, it was possible that the cells failed to shrink because the high [K+]o solution reduced the magnitude of the CCh-induced [Ca2+]i signal. However, the initial [Ca2+]i increase, which is primarily contributed to by release from intracellular stores, was substantially unaltered in the high K+ solution (Fig. 9B). The sustained plateau phase of increased [Ca2+]i was somewhat lower under these conditions (Fig. 9B), likely to be due to reduction of the driving force for Ca2+ entry across the plasma membrane during the sustained phase. Nevertheless, our other observations suggest that the magnitude of the [Ca2+]i during the sustained phase was still sufficient to induce cell shrinkage under normal conditions. Therefore, we conclude that the effects of high K+ to block normal cell shrinkage and efflux of intracellular Cl– were unrelated to effects on [Ca2+]i signals. Rather, the results are consistent with the hypothesis that K+ efflux from the cell is required for agonist-induced serous acinar cell shrinkage. Together with the SPQ data, our results suggest that the observed shrinkage is due to loss of cellular K+ and Cl– content in response to agonist-induced increased [Ca2+]i.
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If cell volume decrease is due to loss of solute, cell swelling towards resting levels upon lowering of [Ca2+]i should reflect solute uptake. In a paradigm of fluid secretion by epithelial cells, solute uptake mechanisms in the basolateral membrane accumulate [Cl–]i above its electrochemical equilibrium value, enabling it to exit the cell across the apical membrane through Cl– channels there. The best-characterized Cl– uptake pathways involve the Na+/H+ exchanger (NHE) coupled in parallel to a Cl–/OH– exchanger (AE), as well as the Na+–K+–2Cl– cotransporter (NKCC). We hypothesized that the activities of either or both of these pathways were responsible for solute accumulation that accounted for cell swelling. To determine the relative contributions of these pathways to the observed cell volume increases, cells were exposed to dimethylamiloride (DMA) or bumetanide, specific inhibitors of NHE and NKCC, respectively. Application of either reagent was without effect on the steady-state volume of unstimulated cells for up to 5 min (not shown). This result suggests that steady-state maintenance of cell solute content (and therefore [Cl–]i) is not dependent on these pathways, indicating either that other transport mechanisms are involved, or that solute efflux and influx pathways are both inactive under resting conditions. Maximal shrinkage upon 100 µM CCh stimulation was 20 ± 1% within 64 ± 4 s (n = 17) in the presence of 30 µM DMA (Fig. 10A) and 22 ± 2% within 62 ± 4 s (n = 25) in the presence of 100 µM bumetanide (Fig. 10B). Thus, neither inhibitor affected the rate or extent of cell shrinkage during CCh stimulation. The agonist-stimulated changes in [Ca2+]i were also unaffected by either inhibitor. In DMA-treated cells stimulated with 100 µM CCh in 0-Ca2+ solution, the mean [Ca2+]i peak was 300 ± 7 nM within 23 ± 2 s, and [Ca2+]i returned to resting levels within 165 ± 18 s (n = 17). Bumetanide-treated cells stimulated with 100 µM CCh in 0-Ca2+ solution exhibited a rise in [Ca2+]i to 310 ± 8 nM within 27 ± 3 s, and [Ca2+]i returned to resting levels within 211 ± 15 s (n = 15).
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These results suggest that cell swelling following Ca2+-induced shrinkage is due to solute uptake through a bumetanide-sensitive pathway, most likely a NKCC transporter. The cotransporter involved is most likely NKCC1, as NKCC2 expression is believed to be restricted to the kidney (reviewed in Mount et al. 1998). Mouse nasal acinar cells express detectable levels of NKCC1 mRNA (Fig. 2). RNA amplification and RT-PCR of NKCC2 in mouse nasal acinar cells as well as mouse nasal tissue was employed to determine whether NKCC2 might play a role in serous cell solute uptake. Although several splice variants of NKCC2 exist in the kidney, most of them differ only in the exons encoding the second transmembrane domain, with the various transcripts having only two different C-terminal ends (one full length and one truncated; reviewed in Mount et al. 1998). Therefore, all NKCC2 isoforms should be detectable with only two 3'-biased primer sets (listed in Supplemental Table 1). Both primers sets detected expression of NKCC2 using RNA isolated from murine kidney (Fig. 10E). However, NKCC2 transcripts could not be detected in either isolated acinar cell aRNA or RNA extracted from mouse nasal tissue. Thus, NKCC1 appears to be the major NKCC isoform expressed in airway gland serous acinar cells. Taken together, our results indicate that solute uptake after Ca2+-induced acinar cell shrinkage is mediated by NKCC1.
Ca2+-evoked fluid secretion does not require CFTR
Since previous data (Engelhardt et al. 1992; Jacquot et al. 1993) as well as our own immunocytochemistry suggest that serous acinar cells are sites of CFTR expression, we explored the possible role of CFTR in the observed cell shrinkage and Cl– loss in response to CCh stimulation. To this end, we used the cftrtm1Unc–/– mouse, which lacks functional CFTR. The maximal cell volume decrease in acinar cells from heterozygote WT/cftrtm1Unc mice was 22 ± 2.5% within 61 ± 3 s (n = 7; data not shown) in response to 10 µM CCh and 21 ± 2% within 61 ± 4 s (n = 8; data not shown) in response to 100 µM CCh. Serous cells isolated from homozygous cftrtm1Unc–/– mice (cftr–/– cells) shrank robustly in response to both 10 µM CCh (19 ± 2% maximal shrinkage within 64 ± 4 s; n = 8, data not shown) and 100 µM CCh (20 ± 2% maximal shrinkage within 63 ± 4 s; n = 19, Fig. 11A). Cells from cftr–/– mice also shrank robustly in response to 100 µM histamine (maximal shrinkage 20 ± 1% within 65 ± 4 s; n = 7, data not shown) and 100 µM ATP (maximal shrinkage 20 ± 1% within 64 ± 3 s; n = 7, data not shown). Thus, no differences in either magnitude or rate of cell shrinkage were observed among cftr–/–, WT, or heterozygote WT/cftrtm1Unc cells. CCh-induced [Ca2+]i signals were nearly identical in cftr–/– and WT cells. Upon stimulation with 100 µM CCh, [Ca2+]i rose to 400 ± 19 nM within 30 ± 2 s and then typically remained at an elevated level of 248 ± 5 nM (n = 14; Fig. 11A). These data indicate that CCh-induced [Ca2+]i increases are capable of inducing cell volume changes in cells lacking CFTR in a manner quantitatively similar to CFTR-expressing WT acinar cells.
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A potential caveat in using cells from the cftrtm1Unc–/– mouse is that they may possess compensatory mechanisms in the absence of CFTR, for example the up-regulation of one or more alternative Cl– conductance(s). To address this possibility, the specific inhibitor CFTRinh172 was used to inhibit CFTR in WT cells. CFTRinh172 has been shown to block glandular fluid secretion in response to both forskolin and pilocarpine (Thiagarajah et al. 2004). WT serous acinar cells that had been pret
