Mucosal Inflammation Research Group and Department of Physiology and Biophysics, University of Calgary, Calgary, Alberta, Canada
Submitted 21 September 2004 ; accepted in final form 20 June 2005
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ABSTRACT |
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ion transport; inflammation; secretion
The CFTR is a key protein involved in ion-channel regulation and chloride secretion in several epithelia, including that of the intestine (5). Regulation of CFTR-mediated chloride secretion occurs through three processes: phosphorylation leading to channel opening, trafficking of CFTR to the membrane, and CFTR endocytosis. The control of these processes is complex and has been recently reviewed (6). The role of cAMP in the gating function of apical membrane-bound CFTR has been well described (see Ref. 14 for a review). Similarly, trafficking of CFTR to the apical membrane of intestinal epithelial cells appears to be a cAMP-dependent process, because T84 colonic cells stimulated with the adenylate cyclase activator forskolin exhibit increased CFTR immunoreactivity at the apical surface (30). Inhibition of trafficking is associated with the blockade of the cAMP-induced increase in CFTR-mediated chloride secretion (28).
Although we have shown that iNOS-derived NO inhibits cAMP-dependent chloride transport by intestinal epithelial cells, we have not determined whether this is an effect on phosphorylation of CFTR present in the apical plasma membrane or on CFTR trafficking. On the basis of the observation that increases in intracellular cAMP stimulated insertion of CFTR into the apical plasma membrane of epithelial cells (28, 30), we have tested the hypothesis that NO inhibits cAMP-dependent CFTR translocation into the apical plasma membrane, thereby reducing chloride secretion, using the SCBN intestinal crypt epithelial cell line.
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MATERIALS AND METHODS |
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Immunofluorescence. Cells were grown on Transwell membranes as described in Cell line. Confluence was determined by light microscopy. Cells were treated with the adenylate cyclase activator forskolin (10 µM) as described below. At various times after exposure to forskolin, they were washed with PBS and fixed in cold 100% methanol for 20 min at 20°C. The fixed cells were then washed three times with PBS and permeabilized in 0.1% Triton X-100 in PBS for 25 min. The 0.1% Triton X-100 in PBS was removed, and blocking solution was added consisting of 5% BSA and 10% goat serum in 0.1% Triton X-100 in PBS. The blocking solution was removed after 30 min, and the primary antibodies were added. Preparations without the primary antibodies added were used to control for specificity of immunostaining. The primary antibodies consisted of monoclonal anti-human CFTR (R domain specific) antibody (R&D Systems, Minneapolis, MN) at 1:100 and rabbit anti-Zonula Occludens (ZO)-1 (Zymed, San Francisco, CA) at 1:100. ZO-1 staining was conducted to visualize tight junctions and was thus used as a marker to distinguish between apical and basolateral plasma membranes. The negative controls used were mouse IgG1-negative control (Dako, Mississauga, Ontario, Canada) and preimmune rabbit serum. The antibodies were diluted in the same blocking solution used above and applied to the cells for 1 h. The cells were then washed five times with PBS, and secondary antibodies were applied for 1 h. The secondary antibodies consist of goat anti-mouse-Cy3 (Jackson Immunoresearch, West Grove, PA) to visualize the CFTR and goat anti-rabbit-FITC (Molecular Probes, Invitrogen, Burlington, Ontario, Canada) to visualize ZO-1. The cells were then washed five times in PBS.
After the staining procedure was complete, the Transwell filters were cut from the wells and embedded on edge in paraffin such that the tissue could be visualized in cross section following sectioning. The samples were cut in 5-µm sections using a microtome (Leica Microsystems RM2125, Richmond Hill, Ontario, Canada). The sections were mounted on silane-treated slides and placed on a slide warmer overnight. The slides were then deparaffinized using NeoClear (EM Sciences, Gibbstown, NJ), rehydrated and placed on a coverslip using FluorSave Reagent (Calbiochem, San Diego, CA) as a mounting medium.
The fixed, permeabilized cell cross sections were visualized using fluorescent microscopy on a Delta-Vision deconvolution microscopy system (Applied Precision, Issaquah, WA). The CFTR was visualized using a rhodamine filter. The ZO-1 was visualized using an FITC filter. Images were taken using a x100 objective, and adjacent pixels were averaged with a bin setting of 2 to allow for clearer images. The apical-to-basolateral ratio of CFTR immunofluorescence was measured using a modification of the procedure described by Moyer and Stanton (22) as shown in Fig. 1.
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Monolayers were stimulated using 10 µM forskolin added to the basolateral side of the monolayers. The NO donor PAPA-NONOate was also added to the basolateral side of the monolayers. PAPA-NONOate was chosen because it releases NO at a steady rate (2 mol/mol of parent compound), with a half-life of 15 min at 37°C (20). The dose of 100 mM was chosen as a maximally effective dose in terms of inhibition of cAMP-dependent ion transport (13). Nocodazole, when used, was added to both the apical and basolateral sides of the monolayers.
Measurement of permeability.
To test the effect of NO on the permeability of SCBN monolayers, we conducted experiments to assess the transepithelial movement of FITC-labeled Dextran 3000 as previously described (11). Briefly, SCBN cells were grown on Transwell filters to a resistance of >1,000 /cm2. They were pretreated with 100 µM PAPA-NONOate and FITC-Dextran 3000 (Molecular Probes, Eugene, OR) was added to the apical side to a final concentration of 10 µM. After a 3-h incubation at 37°C, aliquots (300 µl) were removed and assessed for fluorescence in a microplate fluorometer (Spectra Max Gamini, Molecular Devices, Sunnyvale, CA). Wavelengths for fluorometry were as follows: excitation, 496 nm; emission, 524 nm; and cutoff, 530 nm. Permeability was calculated as fluorescence on the basolateral side expressed as a percentage of the fluorescence of an equal volume sampled from the apical side of the monolayer and normalized to 1 cm2.
Measurement of cAMP. Cells were grown to confluence in 75-cm2 flasks. Confluence was determined by light microscopy. Cells were washed with sterile PBS, and 4 ml FBS- and antibiotic-free DMEM was added to each flask. Cells were lifted using a cell scraper. Cell suspensions were aliquoted (1 ml) into microfuge tubes and placed in a shaking water bath at 37°C. PAPA-NONOate (100 µM) or the vehicle was added to each tube. After 20 min, the phosphodiesterase inhibitor B-8279 (100 µM) was added followed immediately by 10 µM forskolin or vehicle. After being incubated for 5 min, the microfuge tubes were placed in liquid nitrogen to stop the reaction. The cells were lysed by cycling the tubes between liquid nitrogen and a 40°C water bath three times. Tubes were then centrifuged for 5 min at 15,000 g at 4°C. The concentration of cAMP in the supernatants of each tube was determined using a commercially available ELISA kit (Cayman Chemical, Ann Arbor, MI) according to the manufacturer's instructions and expressed as picomoles per milliliter.
Materials. All drugs were purchased from Sigma-Aldrich (Oakville, Ontario, Canada) unless otherwise stated. Routine chemicals for the preparation of buffers were purchased from BDH (Toronto, Ontario, Canada).
Statistics. Data are expressed as means ± SE. Comparisons between two groups were made using Student's t-test for unpaired data. Comparisons of more than two groups were made using analysis of variance with a post hoc Tukey test. Statistical analysis was carried out using GraphPad Instat version 3.00 software (GraphPad Software, San Diego, CA).
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RESULTS |
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To test the mechanism leading to the increase in apical to basolateral CFTR fluorescence ratio, the PKA inhibitor H89 was used. After pretreatment with 5 µM H89, 10 µM forskolin stimulation for 2 min resulted a significant decrease in fluorescence ratio compared with cells treated with forskolin alone (Fig. 3). Pretreatment with H89 did not affect the apical-to-basolateral CFTR fluorescence ratio in cells not stimulated with forskolin (Fig. 3).
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To determine whether there was an interaction among forskolin-stimulated CFTR trafficking, NO, and microtubules in terms of chloride secretion, SCBN cells were the grown on Snapwell filters and mounted in modified Ussing chambers. The cells were pretreated with 100 µM PAPA-NONOate, 33 µM nocodazole, or both. The cells were stimulated with 10 µM forskolin, and Isc was observed. The forskolin-induced
Isc was reduced 32% following pretreatment with nocodazole and 32% following pretreatment with PAPA-NONOate (P < 0.05 for both compared with vehicle controls). After pretreatment with NOC and PAPA-NONOate,
Isc was reduced by 36% (P < 0.05 compared with vehicle control; Fig. 5).
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Because we had hypothesized that the NO donor was having its inhibitory effect through the inhibition of adenylate cyclase, we assessed the effect of PAPA-NONOate on forskolin-induced generation of cAMP in SCBN cells. Forskolin (10 µM) caused an approximately sixfold increase in cAMP (P < 0.001; Fig. 6). Preincubation with PAPA-NONOate inhibited forskolin-induced cAMP by 50% (P < 0.05). PAPA-NONOate did not affect basal cAMP levels in cells not stimulated with forskolin (Fig. 6).
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DISCUSSION |
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The insertion of CFTR into the apical plasma membrane has been observed after stimulation of adenylate cyclase in cell lines (28, 30) and, more recently, in the rat jejunum (1). In the present study, we showed that exposure to 10 µM forskolin on the basolateral side of SCBN monolayers mounted on Transwells caused an increase in cAMP and in CFTR immunoreactivity in the apical plasma membrane. To assess intracellular translocation of CFTR, we used an adaptation of the method of Moyer and Stanton (22). The apical plasma membrane was demarcated in cross sections of SCBN monolayers using staining for the tight junctional protein, ZO-1. CFTR staining was calculated as the ratio of apical to basolateral staining, where "basolateral" refers to the fluorescence intensity in the basolateral membrane minus the background fluorescence intensity as measured in the nuclear compartment. The distribution of CFTR in unstimulated SCBN cells was different from that observed in other intestinal epithelial cell lines. For example, in unstimulated T84 cells, CFTR exists in a subapical cytoplasmic pool, whereas HT-29 cells have a baseline CFTR distribution that is intermediate between SCBN and T84 (18). These different patterns of intracellular distribution of CFTR likely represent inherent differences among these cell lines.
We have investigated the relationship between forskolin-induced CFTR trafficking to the plasma membrane and increases in Isc. Interestingly, the period of peak insertion of CFTR into the plasma membrane following exposure to forskolin preceded the peak change in Isc elicited by forskolin. Caution must be exercised when interpreting an observation of this sort, because a temporal correlation does not prove a cause-and-effect relationship. Nevertheless, this association is what would be expected if insertion of CFTR into the membrane played a part in the secretory response to forskolin, as opposed to the response being due only to phosphorylation and gating of CFTR already present in the plasma membrane. In our study, the peak apical-to-basolateral ratio of CFTR occurred at 30 s following exposure to forskolin, whereas the forskolin-induced change in Isc occurred at 2 min. This observation, together with the fact that disrupting microtubules with nocodazole also reduced forskolin-induced Isc, suggests that CFTR trafficking plays a role, together with the phosphorylation of CFTR already resident in the apical membrane, in the rapid secretory response to activation of adenylate cyclase.
We also observed that forskolin-induced CFTR trafficking into the apical plasma membrane was a transient event. The apical-to-basolateral ratio of CFTR immunofluorescence was not significantly above baseline at 4 or 10 min after exposure to forskolin. This observation is consistent with previous reports showing that endocytosis and, hence, inactivation of CFTR is also a rapid event, such that there is no difference in apical membrane CFTR by 8 min after exposure of T84 cells to forskolin, compared with unstimulated cells (24). Thus, in our experiments, increased apical-to-basolateral ratio of CFTR following forskolin is likely a combination of increased insertion into the membrane and followed by endocytosis of CFTR out of the apical plasma membrane. Although the endocytotic event is slowed by forskolin (24), it may still be a key regulator determining the apical membrane content of CFTR in our study.
One of the novel and important findings of the present study is the observation that an NO donor blocked forskolin-induced CFTR trafficking. It has long been known that endogenous NO can promote fluid absorption in in vivo models. In the rat small intestine, inhibition of NOS with N-nitro-L-arginine methyl ester enhanced basal absorption (27) and inhibited the secretion caused by PGE2, cholera toxin, 5-HT, or heat-stable enterotoxin (7, 8, 27). Increased iNOS expression is characteristic of inflammatory bowel disease (16, 29), where the secretory function of the epithelium is impaired (2, 19, 26). Indeed, we have shown that iNOS-derived NO mediates inflammation-induced suppression of responses to cAMP-dependent secretagogues in trinitrobenzene sulfuric acid-induced colitis (21) and radiation enteropathy in mice (12). In a rat model of mucosal recovery from inflammation, the response to cAMP-dependent secretagogues was still suppressed 6 wk after a bout of inflammation, a time when iNOS expression was still elevated (3). Interestingly, secretory function as measured in vitro returned to normal when tissues were treated with an iNOS inhibitor (3). Of significance is the fact that inhibition of iNOS and return to normal secretory function in the postinflammatory bowel also prevented the increased bacterial translocation observed in this model (4). Recently, we showed that the effect of NO was due to inhibition of NO-sensitive isoforms of adenylate cyclase (13). What remained unknown was whether NO inhibited cAMP-dependent CFTR trafficking, gating, or both.
In the present study, we have shown that the NO donor PAPA-NONOate reduced the forskolin-induced increase in apical CFTR. Previously, we showed that PAPA-NONOate and another NO donor, sodium nitroprusside, reduced forskolin-induced accumulation of cAMP in both mouse colon and T84 colonic epithelial cells and that both of these NO donors inhibited changes in Isc induced by forskolin in segments of mouse colon mounted in Ussing chambers (13). The ability of PAPA-NONOate to block the forskolin-induced increase cAMP was confirmed in the present study. We observed that PAPA-NONOate reduced forskolin-induced cAMP by 50%, whereas the NO donor decreased forskolin-induced changes in the apical-to-basolateral CFTR ratio by 80%. This may be due to the fact that in the cAMP measurement experiments, a phosphodiesterase inhibitor was used so that we could increase the amount of forskolin-stimulated cAMP to be measured by enzyme immunosorbant assay. In the other experiments, in which no phosphodiesterase inhibitor was used, the actual inhibitory effect of PAPA-NONOate on adenylate cyclase activity may have been the same, but concurrent phosphodiesterase activity in the cells reduced cAMP levels further. Thus the combined effect of the NO donor and endogenous phosphodiesterase activity was evident as an enhanced reduction of cAMP-dependent CFTR trafficking.
Because many of the cellular effects of NO are mediated through activation of guanylate cyclase and increased cytoplasmic cGMP (25), we repeated the studies of PAPA-NONOate-induced blockade of forskolin-induced CFTR trafficking in cells pretreated with the guanylate cyclase inhibitor ODQ (15). Interestingly, ODQ pretreatment failed to block the PAPA-NONOate effect, suggesting that NO was not acting through guanylate cyclase. This is consistent with our previous work that showed that NO donors failed to increase cGMP in T84 cells or mouse colon (13). ODQ also had a small but statistically significant inhibitory effect on forskolin-stimulated apical-to-basolateral CFTR ratio. The mechanism underlying this is not known, but the observation does not negate the fact that ODQ was without effect on PAPA-NONOate-mediated inhibition of the effect of forskolin.
Microtubules have been shown to mediate the cytoplasm to apical plasma membrane trafficking of CFTR stimulated by increases in cAMP (30). That study and others (28) used the microtubule inhibitor nocodazole to demonstate cAMP-dependent trafficking of CFTR-containing membrane vesicles. To determine whether the effect of the NO donor was on microtubules, we pretreated SCBN monolayers, either processed for CFTR immunocytochemistry or mounted in Ussing chambers, with PAPA-NONOate or nocodazole or both, before challenge with forskolin. At the concentrations used, both nocodazole and PAPA-NONOate significantly reduced the forskolin-induced increases in apical-to-basolateral CFTR fluorescence and in Isc. Interestingly, the reduction of forskolin-induced effects by the combination of PAPA-NONOate and nocodazole was approximately the same. This observation suggests that PAPA-NONOate and nocodazole are acting at the same pathway or at points in series along the same pathway, rather than on distinct, parallel pathways. Additional studies will be necessary to determine precisely the mechanism underlying the interrelationships among NO, cAMP, and microtubules in CFTR trafficking. Nevertheless, it is clear that CFTR trafficking is responsible for a proportion of the change in Isc elicited by exposure to forskolin and that this is sensitive to NO.
In summary, we have shown that cAMP-dependent trafficking of CFTR from cytoplasmic stores to the apical plasma membrane can be rapidly induced by increasing cAMP in SCBN cells. This process can be blocked by NO in a guanylate cyclase-independent fashion. The ability of NO to disrupt the action of cAMP-dependent secretagogues has implications in terms of the ability of the intestinal epithelium to maintain the secretory component of its barrier function in conditions when mucosal iNOS is elevated.
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GRANTS |
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A. C. Skinn is the recipient of a MD-PhD Studentship awarded through the Canadian Institutes of Health Research Rx&D Research Program. W. K. MacNaughton is an Alberta Heritage Foundation for Medical Research Senior Scholar.
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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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