1Department of Cell Biology and Physiology, University of Pittsburgh School of Medicine, Pittsburgh, Pennsylvania 15261; 2Novartis Respiratory Research Center, Horsham RH125AB, United Kingdom; 3Institut National de la Santé et de la Recherche Médicale 94-04, Université de Nantes, F-44035 Nantes, France, and 4Department of Physiology and Biophysics, Case Western Reserve University, Cleveland, Ohio 44106
Submitted 19 December 2002 ; accepted in final form 15 September 2003
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ABSTRACT |
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capacitative calcium entry; membrane capacitance; chloride channels; fura 2
The Cl.16E subclone of the human colonic cancer cell line HT29 differentiates to a goblet cell-like epithelium with typically 50% of cells having large numbers of mucin granules in the apical cytoplasm (1). ATP has been shown to stimulate both chloride and mucin secretion in this line (20), allowing both regulatory pathways to be studied in a single model system. In single cells, whole cell patch clamping experiments have verified substantial exocytosis in response to ATP through measurement of membrane capacitance changes (11). Changes in capacitance reflect changes in net membrane area that occur during the fusion and retrieval of mucin granules at the plasma membrane, as well as the vesicular trafficking of membrane proteins. Stimulated capacitance increases are dominated by exocytosis during the initial seconds of the response, whereas later time points reflect endocytosis as well as exocytosis.
Several lines of evidence confirm that ATP-stimulated capacitance changes in Cl.16E correspond to mucin exocytosis. ATP stimulates the release of [3H]glucosamine-labeled mucin into the extracellular medium, and electron micrographs of basal and ATP-stimulated cells indicate massive, compound fusion of granules after ATP treatment (20). Given a typical granule diameter of 1 µm, previously observed capacitance changes in Cl.16E would correspond to the fusion of 130 mucin granules (11), a realistic number based on the electron micrographs. In contrast, the same change in capacitance would require the rapid fusion of 13,000 vesicles with the plasma membrane, given a vesicle diameter of 100 nm. Finally, ATP stimulation does not significantly increase capacitance in HT29-Cl.19A cells, which share similar chloride secretory responses with Cl.16E but lack mucin granules (11).
Purinergic stimulation in HT29-Cl.16E cells occurs through apical P2Y2 receptors (21) and induces a large, transient increase in intracellular calcium followed by a plateau phase lasting several minutes after agonist washout (10). A transient increase in intracellular calcium has been demonstrated to stimulate mucin secretion (2), although stimulated chloride secretion has been demonstrated in the absence of this intracellular calcium increase (10). Direct measurements of exocytosis under clamped intracellular calcium conditions were not performed in these early experiments, but a substantial portion of the purinergically stimulated chloride secretion was later shown to depend on granule fusion (11), suggesting that ATP-stimulated exocytosis itself might be calcium independent. These experiments utilized wortmannin, at concentrations known to block PI3-kinase (38), to inhibit exocytosis.
The identity of the purinergically stimulated chloride conductance in the HT29-Cl.16E subclone is not clear. Strong evidence exists for the presence of Ca2+-activated Cl channels (CaCC) in HT29 parental (undifferentiated) and Cl.19A subclone cells (reviewed in Ref. 18). Furthermore, 16E and 19A subclones show similar ATP-stimulated chloride secretory responses (10). However, DIDS is ineffective at blocking chloride secretion in Cl.16E, although di-Cl-DPC is very effective (77%, Ref. 20).
Niflumic acid (NA), a nonsteroidal anti-inflammatory agent, was originally determined to be a potent, reversible blocker of endogenous CaCC in Xenopus oocytes, with an apparent inhibition constant of 17 µM (37). NA has since been shown to block purinergically stimulated chloride currents in human bronchial cells (39), renal A6 cells (3), murine tracheal epithelial cells (8), and murine inner medullary collecting duct cells (31). Now regarded as a general blocker of CaCC, a growing body of evidence suggests that NA may also inhibit secretion, relax contraction, and potentiate potassium channels.
Two distinct mechanisms have been suggested to account for the action of NA on secretion and contraction. The first, supported by the action of NA on angiotensin II inhibition of renin secretion (23), stimulated lysozyme release in trachea submucosal glands (9), and inhibition of 5-hydroxytryptamine-induced contraction of rat trachea (34), is that CaCC play a direct role in the control of secretory events, where activation of CaCC and concomitant membrane depolarization could evoke Ca2+ influx via L-type calcium channels. Nifedipine, a blocker of L-type Ca2+ channels, was shown to mimic the effect of NA in rat trachea (34), and this mechanism could account for CaCC control of secretion in excitable cells. Depolarization-induced Ca2+ influx is not considered a significant mechanism for stimulating exocytosis in epithelia, however (12). The second, supported by NA inhibition of histamine-stimulated phospholipase C in bovine chromaffin cells (28), serotonin release in mast cells (27), and relaxation of endothelin-1-induced pulmonary artery constriction (17), is that NA blocks Ca2+ influx independent of CaCC block, for example, by blocking capacitative calcium entry. In support of this, Reinsprecht et al. (27) found La3+ as effective as NA at inhibiting secretion from mast cells. Potentiation of potassium channels by NA, in particular the large-conductance, calcium-activated potassium channel, has also been noted (13, 22, 24). This effect occurs at NA > 50 µM, whereas blockade of CaCC occurs below 50 µM.
The ability of NA to inhibit both chloride channels and exocytotic secretion makes it a candidate for probing the signaling divergence of mucin and chloride secretion during compound exocytosis in the intestinal goblet cell model HT29-Cl.16E. In the studies reported here, we have used whole cell patch clamping with capacitance measurements to characterize purinergically stimulated exocytosis and chloride secretion in the presence of NA and fura 2 fluorescence to characterize the corresponding intracellular calcium changes.
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MATERIALS AND METHODS |
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Whole cell patch clamping. Three different methods of single-cell capacitance measurement were used in the results described below: 1) a 4-sinewave estimation based on the earlier 2-sinewave method of Rohlicek and Schmid (29) using an EPC-7 amplifier (List Medical), previously described in detail (33); 2) the standard membrane test available in pClamp V 8.0 software using an Axopatch 200A amplifier (Axon Instruments); and 3) the square pulse technique of Thompson et al. (36), implemented as follows. From a holding potential of 20 mV, a +5 mV, 2-ms square pulse was generated using an S-95 Tri-level stimulator (Medical Systems) and applied to the external command input of a 200B Axopatch amplifier (Axon Instruments). The stimulator also generated a trigger signal 1 ms before the pulse, which was used to initiate acquisition of the resulting current signal by Clampex 8.1 software through a Digidata 1322A acquisition board (both from Axon Instruments). With the use of a sampling rate of 50 kHz, 5 ms of data were acquired after the trigger. Pulses were repeated at 50-ms intervals.
The current transients resulting from the square pulse were fit with exponentials using the published fast fitting routine (36), programmed in Matlab software (Mathworks). Averaging of 510 consecutive cycles was performed to improve signal-to-noise before fitting. Estimates of membrane capacitance (Cm), membrane resistance (Rm), and access resistance (Ra) were performed offline.
Four different protocols were used for the results reported below: capacitance measurements, with a holding potential of 20 mV and a sample pulse of 0.5 mV (1), 10 mV (2), or 5 mV (3); a current-voltage (I-V) protocol, with a holding potential of 20 mV; a voltage-clamp protocol, with an alternating pulse between ECl (5 mV) and EK (95 mV), holding potential = membrane potential (10 or 20 mV); and current-clamp protocol, with Im clamped at 0 pA. All reported capacitance peaks occurred within 5 s from the onset of the response, where capacitance is dominated by exocytosis.
A phosphate-buffered bath solution (in mM: 145 NaCl, 0.4 KH2PO4, 1.6 K2HPO4, 1.0 MgCl2, 1.5 CaCl2, and 5 glucose) was used for general perfusion in most experiments. For low-extracellular calcium experiments, the phosphate-buffered bath was switched to a low-calcium bath (in mM: 145 NaCl, 0.4 KH2PO4, 1.6 K2HPO4, 1.0 MgCl2, 1.0 EGTA, and 5 glucose) for the indicated time interval. A phosphate-buffered pipette solution (in mM: 125 KCl, 1.2 NaH2PO4, 4.8 Na2HPO4, 1.03 MgCl2, 5 glucose, and 0.1 EGTA) was used during experiments utilizing phosphate bath solutions. For low-intracellular chloride experiments, 95 mM KCl in the phosphate-buffered pipette solution was replaced with K-gluconate. A subset of experiments was performed in HEPES-buffered bath (in mM: 140 NaCl, 5 KCl, 10 HEPES, 1 MgCl2, and 2 CaCl2) and pipette (in mM: 145 KCl, 1 MgCl2, 10 HEPES, and 0.1 EGTA) solutions, as explicitly noted. The low concentration of EGTA in all pipette solutions was used to set the free Ca2+ in the pipette to 100 nM without significantly buffering or preventing cytosolic Ca2+ transients (10). All solutions were at pH 7.3; bath solutions were maintained at 37°C and perfused at 3.0 ml/m using a Valvelink 8 (Automate Scientific) controlled by the Clampex 8.1 software. ATP was omitted from the pipette solution in all cases because it activated cells during seal formation. Pipettes using standard wall thickness borosilicate glass to decrease stray capacitance were pulled to tip diameters of 12 µm (access resistance <4 M
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Fluorescence measurements. Bath solutions were the same as for patch clamping. Coverslips were incubated in bath solution containing 5 µM fura 2-AM for 30 min at 25°C, followed by an additional 30 min in bath solution alone. The coverslips were mounted in an open perfusion chamber for imaging and were perfused with 37°C bath solution ± agonist at 1 ml/min. The chamber was mounted on the stage of an inverted microscope (Nikon Diaphot) and imaged with a x40 objective (DApo 40UV 1.30). Ratiometric fluorescence measurements were performed with excitation at 340/380 nm and emission at 510 nm. Typical exposure time per excitation wavelength was 0.16 s. Images were captured using a cooled charge-coupled device camera (model C4742-95, Hamamatsu).
Each experiment started with 4 min of perfusion with bath solution alone. Fluorescence measurements were performed at 30-s intervals during this time and then switched to 2-s intervals roughly 30 s before the addition of agonist solution (relative to chamber). The 2-s interval was maintained for at least 30 s after the start of the response. After this time, measurements were performed at 10-s intervals for an additional 3 min and then 30-s intervals for the remainder of the experiment. The duration of agonist perfusion was 3 min in all cases, followed by bath solution for 58 min.
Images were acquired and analyzed using Simple PCI software (Compix). A region of interest (ROI) was defined for each cell, and the mean fluorescent intensity at 340 and 380 was divided to give the ratio. Plots of the ratio values vs. time for individual ROIs indicated that cells in a cluster behaved similarly, but different clusters on a single coverslip exhibited greater variability. The ratio values for each cluster were averaged and then normalized by the mean of the 10 values (F0) preceding the start of the response. The normalized ratios were used in analysis; ratio values were not converted into intracellular calcium because of the difficulties in obtaining an accurate Kd value for the fura 2 inside the cells.
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RESULTS |
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In the presence of 20 µM NA, ATP-stimulated capacitance increases were markedly diminished, as shown in Fig. 1B. Addition of 200 µM ATP plus 20 µM niflumic acid stimulated a 3 ± 1% increase in membrane capacitance and a secretory current of 380 ± 255 pA (n = 4). The reduction in peak current did not reach statistical significance, however. To better account for variations in cell size and membrane area increases during individual experiments, peak current was normalized by peak membrane capacitance. ATP-stimulated cells held at 20 mV had an average peak current of 39 ± 10 pA/pF, whereas ATP stimulation in the presence of 20 µM NA produced an average peak current of 34 ± 22 pA/pF, indicating that NA was not blocking chloride secretion at the peak of response in these cells.
The lack of strong chloride secretion block was unexpected, because unstimulated HT29-Cl.16E cells typically display outward rectification, a characteristic associated with CaCC channels (18). A repetitive step protocol was performed to assess the I-V characteristics during the typical response. Protocol timing was automated to repeat the 80-ms-long step protocol every 10 s, with voltage clamp at 20 mV maintained in between. The I-V curves at several time points before and during ATP stimulation, as well as the corresponding membrane current during the voltage-clamp intervals for representative experiments (n 3, each condition), are shown in Fig. 2. At the peak of ATP stimulation (Fig. 2A), the I-V plot indicates a large, outwardly rectified current with a reversal potential very near the chloride equilibrium potential of 5 mV. In the presence of NA (Fig. 2B), an equally large current is stimulated by ATP but has a linear response again with a reversal potential very near 5 mV. Before stimulation, the basal curve in Fig. 2B demonstrates outward rectification. Clearly, if NA is blocking CaCC channels, HT29-Cl.16E possess alternate chloride transport paths that compensate any block at the peak of stimulation.
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Although NA shows very little inhibitory effect on chloride secretion during the initial transient response, differences are more pronounced at later time points. The typical ATP I-V response shows little difference between 30 s and 60 s (Fig. 2A); the cell remains partially activated. In contrast, cells stimulated with ATP in the presence of NA indicate a return to basal levels at 60 s (Fig. 2B). The reversal potential in both cases at 60 s has shifted slightly positive. Current-clamp experiments verified a significant difference at later time points in the presence of NA (Fig. 3). ATP stimulation induced an immediate, rapid depolarization during the initial transient response in all cases, but in the presence of NA, cells failed to repolarize (Fig. 3A). When a low-chloride pipette solution was used, ATP stimulated a rapid hyperpolarization, followed by a return to basal levels (Fig. 3B), indicating that chloride ions are predominantly involved in the stimulatory response. ATP stimulated the same capacitance change observed in Fig. 1 when using a low-chloride pipette solution (data not shown). Although NA may block chloride channels involved in recovery after ATP stimulation, there is no evidence to indicate that it has a significant chloride channel effect during the initial transient response.
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To assess whether potassium channel potentiation might be contributing to the observed responses, a pulse protocol was performed to measure current flow at the chloride (ECl) and potassium (EK) equilibrium potentials. In ATP-stimulated cells, the majority of current flows when the cells are clamped at the EK potential of 95 mV, shown in Fig. 4A. Very little current flow is observed at the ECl potential of 5 mV. Including 20 µM NA with ATP does not significantly change the response (data not shown). Increasing the concentration of NA to 100 µM results in a small increase in potassium current (Fig. 4B), but the current at EK is still significantly greater.
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Effect of NA on intracellular calcium dynamics. The time to peak capacitance change after ATP stimulation noted above occurs during the initial transient response when NA exerts minimal chloride channel block. This suggests that NA must be acting on a different target to effect the significant reduction of exocytosis observed. To assess whether NA might be exerting its effect on exocytosis through a calcium-mediated pathway, fura 2 imaging was performed. Stimulation with 200 µM ATP induced a rapid, transient elevation in intracellular calcium, followed by an elevated plateau phase lasting several minutes (Fig. 5). The peak amplitude was not significantly different in the presence of NA; however, a significant delay in the time required to reach the peak was observed (Fig. 5B). NA frequently depressed the plateau phase, but this effect did not reach statistical significance.
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The observed delay in the transient elevation of calcium (Fig. 5) confirms that NA affects intracellular calcium dynamics but does not necessarily account for its effect on exocytosis. The time required to reach a calcium peak with ATP alone is 37 ± 3 s (Fig. 5B), whereas the time required to reach a capacitance peak is 2.4 ± 0.3 s (above, Fig. 1A). The curves in Fig. 5A indicate a subtle difference in the initial few seconds of the calcium transient, however, with timing comparable to stimulated exocytosis. Both curves show an initial, rapid calcium pulse occurring in the first 2 s but with different kinetics. In the presence of NA, the initial peak is delayed 12.5 ± 1.7%, with a significant (P < 0.05) attenuation in amplitude of 8 ± 2% (n = 7). With ATP alone, calcium continues to rise after an almost imperceptible drop, but in the presence of NA, a very pronounced depression occurs before calcium again increases to its peak value, presumably due to the attenuated response during the rapid, initial pulse.
Removing extracellular calcium mimics the effect of NA on capacitance and intracellular calcium dynamics. Previous work in HT29-Cl.16E indicated that when ATP stimulation occurred in the presence of low extracellular calcium, the transient calcium response was delayed and the plateau phase was absent (10). This previous study (10) did not use a sampling interval fast enough to detect changes in the initial calcium spike but nonetheless indicated that extracellular calcium had a role in shaping the intracellular calcium response to ATP. In these studies, whole cell capacitance measurements were performed with varying intervals of calcium-free bath application before stimulation with ATP. With 0 s and 20 s pretreatment in calcium-free bath, ATP stimulation was normal (n = 3 each, data not shown). When calcium was removed from the bath for 40 s before ATP stimulation, peak capacitance change was reduced by 50% whereas peak secretory current was typical at 772 ± 420 pA (n = 3). The responses (Fig. 6) were very similar to ATP stimulation in the presence of NA (Fig. 1B).
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With the use of a similar interval of pretreatment with calcium-free bath, fura 2 imaging was repeated. As shown in Fig. 7, ATP stimulation in the presence of normal extracellular calcium produced a very sharp, initial calcium spike with amplitude comparable to the second, slower peak rise. When calcium was removed from the bath for 50 s, the initial spike was absent. Overall, the absence of extracellular calcium delayed the initial transient by 28.5 ± 1.7% (P < 0.05, n = 10). Low extracellular calcium muted the intracellular plateau phase as well, but these effects are again well past the response time for exocytosis. Under normal conditions, this initial calcium spike occurs extremely fast, reaching a peak in 4 s. The timing of this initial transient and its significant delay under conditions shown to reduce ATP-stimulated capacitance increases suggest its involvement in signaling exocytosis.
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Blockers of capacitative calcium entry duplicate the effect of NA on capacitance and intracellular calcium dynamics. To test for the involvement of capacitative calcium entry in ATP-stimulated exocytosis, whole cell patch-clamp capacitance measurements were repeated in the presence of La3+ and SKF-96365, potent blockers of several channels thought to be involved in capacitative calcium entry (25). Because lanthanum is precipitated by phosphate, HEPES-based solutions were used for the following experiments (micromolar concentrations of LaCl3 had no effect on ATP-stimulated exocytosis in phosphate-buffered solutions, data not shown). The ATP ± NA experiments (Fig. 1) were also repeated in HEPES-buffered solutions to rule out any solution based effects. As shown in Fig. 8, 20 µM LaCl3, 10 µM SKF-96365, or 20 µM NA were equally effective in attenuating ATP-stimulated exocytosis (70% attenuation over ATP alone, P < 0.05). None of the blockers had a statistically significant effect on stimulated current, and for SKF-96365 and NA, average stimulated currents were greater than with ATP alone.
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To test whether capacitative calcium entry was responsible for the timing and magnitude of the initial calcium transient observed in Figs. 5 and 7, fura 2 imaging was repeated in HEPES-buffered solutions using ATP ± LaCl3. In the presence of 20 µM LaCl3, both the timing and magnitude of the initial calcium transient were significantly affected, with a delay of 36.3 ± 1.9% and a reduction in peak magnitude of 12.2 ± 1.9% (n = 7, P < 0.05). Thus capacitative calcium entry contributes to a rapid calcium transient coincident with ATP-stimulated mucin exocytosis.
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DISCUSSION |
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In the absence of an effect on CaCC channels, the effect of NA on ATP-stimulated intracellular calcium dynamics was assessed using fura 2. NA increased the amount of time required for intracellular calcium to reach a peak after ATP stimulation without affecting the peak value itself (Fig. 5). This result clearly demonstrated that NA was altering intracellular calcium dynamics. The time to reach the ATP-stimulated and NA-delayed calcium peaks extended beyond the initial transient phase during which exocytosis occurs, however, suggesting that bulk intracellular calcium elevation alone was not a sufficient trigger of exocytosis.
The subtle difference between the ATP-stimulated and NA-delayed calcium transients during the first 10 s of response, shown in Fig. 5A, occurred during the same time frame as exocytosis. This result suggested that a rapid calcium increase might be required for stimulating maximum exocytosis and that NA might exert its effect in part by interfering with the timing of the initial calcium transient. Removing extracellular calcium for 40 s inhibited the ATP-stimulated capacitance increase with minimal effect on the current, similar to observations with NA (Fig. 1B vs. Fig. 6), whereas measurement of ATP-stimulated intracellular calcium changes after 50 s in extracellular calcium free bath indicated that the initial peak calcium transient was markedly diminished (Fig. 7).
The rapid influx of calcium across the plasma membrane in epithelia is most commonly associated with capacitative calcium entry, triggered by depletion of intracellular calcium stores. The mechanisms responsible for such entry are still unclear, and competing hypotheses exist (26). One family of channels thought to be involved with capacitative calcium entry is the transient receptor potential, or TRP channels (26). The observed influx block by NA or low extracellular calcium shown in Figs. 5 and 7 occurs in the initial phase of the calcium response, presumably before stores are depleted. TRP channel activation has been demonstrated in the absence of store depletion (5); thus TRP channels seem a possible target for NA. Reinsprecht et al. (27) also demonstrated that La3+, known to block TRP channels (25), mimicked the effect of NA in mast cells.
In light of this, the effect of LaCl3, as well as the more specific TRP channel blocker SKF-96365, was tested during ATP stimulation of exocytosis and found to inhibit ATP-stimulated capacitance increases by 76 and 65%, respectively (Fig. 8). As with NA, the inhibition of exocytosis was independent of chloride secretion. Furthermore, LaCl3 significantly delayed and attenuated the initial peak transient of the calcium response similar to NA or removal of extracellular calcium, implicating TRP channels as the source for extracellular calcium entry critical for initiating maximum exocytosis.
Previous studies in HT29-Cl.16E demonstrated purinergically stimulated chloride secretion was unaffected when intracellular calcium was clamped by 10 mM BAPTA in the pipette solution (10). These data support the conclusion that either CaCC are not strongly active in HT29-Cl.16E during the transient capacitance response or that alternate chloride channels are activated in the absence of CaCC activity. In a later study using wortmannin to inhibit exocytosis, Guo et al. (11) found that the inhibition of 80% of the peak capacitance change was accompanied by a 50% reduction in peak secretory current and concluded that a significant portion of the purinergically stimulated chloride channels resided in granules. Several recent publications have identified CaCC expression in airway goblet cells (14) and, more explicitly, in the mucin granule membranes of murine intestinal and airway goblet cells (19) and rat pancreatic zymogen granule membranes (35). If CaCC also reside on the mucin granules of HT29-Cl.16E, it might be expected that inhibiting 80% of mucin granule exocytosis should have a measurable effect on chloride secretion.
In our study, an 80% reduction in capacitance change does not diminish chloride secretory currents. Our initial data suggested a 50% reduction in current during ATP stimulation in the presence of NA before normalization (Fig. 1B), and the data do exhibit a large standard error. The lack of strong chloride channel block was later confirmed by 1) strong chloride secretion in the I-V curves for ATP stimulation ± NA, especially at the potassium equilibrium potential (95 mV) where chloride currents are isolated (Fig. 2); 2) similar current tracings at the ECl and EK equilibrium potentials during ATP stimulation ± NA (Fig. 4); and 3) demonstration that ATP ± NA was equally rapid and effective at depolarizing the membrane, having also shown that the depolarization was due to chloride secretion (Fig. 3). Thus we cannot conclude that the purinergically stimulated chloride current results from channels located in granules.
Although our results appear to conflict with expression of CaCC on mucin granule membranes, there is a general lack of published data correlating endogenous CaCC expression with measurable chloride secretion under physiological conditions (16). Granule membrane CaCC could function during synthesis or condensation of mucins into granules (19) rather than during secretion. Thévenod et al. (35) recently found that CaCC expressed in isolated zymogen granules responded to elevated calcium with an increase in conductance rather than chloride. Our data, and earlier results in HT29-Cl.16E showing a lack of DIDS inhibition (20), suggest that CaCC do not play a critical role in the transient secretory current stimulated by ATP.
Our data suggest that a localized calcium signal is important in the stimulation of compound mucin granule exocytosis during ATP stimulation. Of particular interest is the importance of the timing of the transient, because bulk intracellular calcium eventually reaches the same peak in the presence of NA or in the absence of extracellular calcium. Localized calcium signals have been shown to play an important role in controlling secretory events in epithelia (4), and the initial calcium transient observed in our studies might be significantly greater within the apical space surrounding granules than evident from fura 2 measurements; that is, greater than the bulk calcium peak observed at later time points. Most importantly, our data indicate a divergence in the signaling of mucin exocytosis and chloride secretion in this intestinal cell model that may provide a pharmacological target for controlling excessive mucin release.
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DISCLOSURES |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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REFERENCES |
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---|
2. Augeron C, Voisin T, Maoret JJ, Berthon B, Laburthe M, and Laboisse CL. Neurotensin and neuromedin N stimulate mucin output from human goblet cells (Cl.16E) via neurotensin receptors. Am J Physiol Gastrointest Liver Physiol 262: G470G476, 1992.
3. Banderali U, Brochiero E, Lindenthal S, Raschi C, Bogliolo S, and Ehrenfeld J. Control of apical membrane chloride permeability in the renal A6 cell line by nucleotides. J Physiol 519: 737751, 1999.
4. Bootman MD, Lipp P, and Berridge MJ. The organisation and functions of local Ca2+ signals. J Cell Sci 114: 22132222, 2001.[ISI][Medline]
5. Clapham DE, Runnels LW, and Strubing C. The TRP ion channel family. Nature Reviews 2: 387396, 2001.
6. Davis CW. Regulation of mucin secretion from in vitro cellular models. In: Mucus Hypersecretion In Respiratory Disease, edited by Chadwick DJ and Goode JA. Chichester: John Wiley and Sons, 2002, p. 292.
7. Forstner G. Signal transduction, packaging and secretion of mucins. Annu Rev Physiol 57: 585605, 1995.[CrossRef][ISI][Medline]
8. Gabriel SE, Makhlina M, Martsen E, Thomas EJ, Lethem MI, and Boucher RC. Permeabilization via the P2X7 purinoreceptor reveals the presence of a Ca2+-activated Cl conductance in the apical membrane of murine tracheal epithelial cells. J Biol Chem 275: 3502835033, 2000.
9. Griffin A, Newman TM, and Scott RH. Electrophysiological and ultrastructural events evoked by methacholine and intracellular photolysis of caged compounds in cultured ovine trachea submucosal gland cells. Exp Physiol 81: 2743, 1996.[Abstract]
10. Guo X, Merlin D, Harvey RD, Laboisse C, and Hopfer U. Stimulation of Cl secretion by extracellular ATP does not depend on increased cytosolic Ca2+ in HT29-Cl.16E. Am J Physiol Cell Physiol 269: C1457C1463, 1995.
11. Guo XW, Merlin D, Laboise C, and Hopfer U. Purinergic agonists, but not cAMP, stimulate coupled granule fusion and Cl conductance in HT29-C1.16E. Am J Physiol Cell Physiol 273: C804C809, 1997.
12. Hille B, Billiard J, Babcock DF, Nguyen T, and Koh DS. Stimulation of exocytosis without a calcium signal. J Physiol 520: 2331, 1999.
13. Hogg RC, Wang Q, and Large WA. Action of niflumic acid on evoked and spontaneous calcium-activated chloride and potassium currents in smooth muscle cells from rabbit portal vein. Br J Pharmacol 112: 977984, 1994.[Abstract]
14. Hoshino M, Morita S, Iwashita H, Sagiya Y, Nagi T, Nakanishi A, Ashida Y, Nishimura O, Fujisawa Y, and Fujino M. Increased expression of the human Ca2+-activated Cl channel 1 (CaCC1) gene in the asthmatic airway. Am J Respir Crit Care Med 165: 11321136, 2002.
15. Hwang TH, Schwiebert EM, and Guggino WB. Apical and basolateral ATP stimulates tracheal epithelial chloride secretion via multiple purinergic receptors. Am J Physiol Cell Physiol 270: C1611C1623, 1996.
16. Jentsch TJ, Stein V, Weinreich F, and Zdebik AA. Molecular structure and physiological function of chloride channels. Physiol Rev 82: 503568, 2002.
17. Kato K, Evans MA, and Kozlowski RZ. Relaxation of endothelin-1-induced pulmonary arterial constriction by niflumic acid and NPPB: mechanism(s) independent of chloride channel block. J Pharmacol Exp Ther 288: 12421250, 1999.
18. Kidd JF and Thorn P. Intracellular Ca2+ and Cl channel activation in secretory cells. Annu Rev Physiol 62: 493513, 2000.[CrossRef][ISI][Medline]
19. Leverkoehne I and Gruber AD. The murine mCLCA3 (alias gob-5) protein is located in the mucin granule membranes of intestinal, respiratory, and uterine goblet cells. J Histochem Cytochem 50: 829838, 2002.
20. Merlin D, Augeron C, Tien XY, Guo X, Laboisse CL, and Hopfer U. ATP-stimulated electrolyte and mucin secretion in the human intestinal goblet cell line HT29-Cl.16E. J Membr Biol 137: 137149, 1994.[ISI][Medline]
21. Merlin D, Guo X, Martin K, Laboisse C, Landis D, Dubyak G, and Hopfer U. Recruitment of purinergically stimulated Cl channels from granule membrane to plasma membrane. Am J Physiol Cell Physiol 271: C612C619, 1996.
22. Miralles F, Marsal J, Peres J, and Solsona C. Niflumic acid-induced increase in potassium currents in frog motor nerve terminals: effects on transmitter release. Brain Res 714: 192200, 1996.[CrossRef][ISI][Medline]
23. Nabel C, Schweda F, Riegger GAJ, Kramer BK, and Kurtz A. Chloride channel blockers attenuate the inhibition of renin secretion by angiotensin II. Pflügers Arch 438: 694699, 1999.[CrossRef][ISI][Medline]
24. Ottolia M and Toro L. Potentiation of large conductance KCa channels by niflumic, flufenamic, and mefenamic acids. Biophys J 67: 22722279, 1994.[Abstract]
25. Putney JW Jr. The pharmacology of capacitative calcium entry. Molecular Interventions 1: 8594, 2001.
26. Putney JW Jr, Broad LM, Braun FJ, Lievremont JP, and Bird GSJ. Mechanisms of capacitative calcium entry. J Cell Sci 114: 22232229, 2001.[ISI][Medline]
27. Reinsprecht M, Rohn MH, Spadinger RJ, Pecht I, Schindler H, and Romanin C. Blockade of capacitive Ca2+ influx by Cl channel blockers inhibits secretion from rat mucosal-type mast cells. Mol Pharmacol 47: 10141020, 1995.[Abstract]
28. Roberts-Thomson EL, Palmer SM, Powis DA, and Bunn SJ. Histamine-stimulated phospholipase C activity in bovine adrenal medullary chromaffin cells: the effect of chloride-channel antagonists and low extracellular chloride concentrations. Neurosci Lett 278: 9396, 2000.[CrossRef][ISI][Medline]
29. Rohlicek V and Schmid A. Dual-frequency method for synchronous measurement of cell capacitance, membrane conductance and access resistance on single cells. Pflügers Arch 428: 3038, 1994.[ISI][Medline]
30. Specian RD and Oliver MG. Functional biology of intestinal goblet cells. Am J Physiol Cell Physiol 260: C183C193, 1991.
31. Stewart GS, Glanville M, Aziz O, Simmons NL, and Gray MA. Regulation of an outwardly rectifying chloride conductance in renal epithelial cells by external and internal calcium. J Membr Biol 180: 4964, 2001.[CrossRef][ISI][Medline]
32. Stutts MJ, Fitz JG, Paradiso AM, and Boucher RC. Multiple modes of regulation of airway epithelial chloride secretion by extracellular ATP. Am J Physiol Cell Physiol 267: C1442C1451, 1994.
33. Sun F, Hug MJ, Bradbury NA, and Frizzell RA. Protein kinase A associates with cystic fibrosis transmembrane conductance regulator via an interaction with ezrin. J Biol Chem 275: 1436014366, 2000.
34. Teixeira MCL, Coelho RR, Leal-Cardoso JH, and Criddle DN. Comparative effects of niflumic acid and nifedipine on 5-hydroxytrytamine-and acetylcholine-induced contraction of the rat trachea. Eur J Pharmacol 394: 117122, 2000.[CrossRef][ISI][Medline]
35. Thévenod F, Roussa E, Benos DJ, and Fuller CM. Relationship between a HCO3-permeable conductance and a CLCA protein from rat pancreatic zymogen granules. Biochem Biophys Res Commun 300: 546554, 2003.[CrossRef][ISI][Medline]
36. Thompson RE, Lindau M, and Webb WW. Robust, high-resolution, whole cell patch-clamp capacitance measurements using square wave stimulation. Biophys J 81: 937948, 2001.
37. White MM and Aylwin M. Niflumic and flufenamic acids are potent reversible blockers of Ca2+-activated Cl channels in Xenopus oocytes. Mol Pharmacol 37: 720724, 1990.[Abstract]
38. Yano H, Nakanishi S, Kimura K, Hanai N, Saitoh Y, Fukui Y, Nonomura Y, and Matsuda Y. Inhibition of histamine secretion by wortmannin through the blockade of phosphatidylinositol 3-kinase in RBL-2H3 cells. J Biol Chem 268: 2584625856, 1993.
39. Zegarra-Moran O, Sacco O, Romano L, Rossi GA, and Galietta LJV. Cl currents activated by extracellular nucleotides in human bronchial cells. J Membr Biol 156: 297305, 1997.[CrossRef][ISI][Medline]