Expression of the chloride channel ClC-2 in the murine small intestine epithelium

Katalin Gyömörey1, Herman Yeger1, Cameron Ackerley2, Elizabeth Garami1, and Christine E. Bear1

1 Programme in Cell Biology and 2 Department of Pathology in the Research Institute, Hospital for Sick Children, Toronto, Ontario, Canada M5G 1X8


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The chloride channel ClC-2 has been implicated in neonatal airway chloride secretion. To assess its role in secretion by the small intestine, we assessed its subcellular expression in ileal segments obtained from mice and studied the chloride transport properties of this tissue. Chloride secretion across the mucosa of murine ileal segments was assessed in Ussing chambers as negative short-circuit current (Isc). If ClC-2 contributed to chloride secretion, we predicted on the basis of previous studies that negative Isc would be stimulated by dilution of the mucosal bath and that this response would depend on chloride ion and would be blocked by the chloride channel blocker 5-nitro-2-(3-phenylpropylamino) benzoic acid but not by DIDS. In fact, mucosal hypotonicity did stimulate a chloride-dependent change in Isc that exhibited pharmacological properties consistent with those of ClC-2. This secretory response is unlikely to be mediated by the cystic fibrosis transmembrane conductance regulator (CFTR) channel because it was also observed in CFTR knockout animals. Assessment of the native expression pattern of ClC-2 protein in the murine intestinal epithelium by confocal and electron microscopy showed that ClC-2 exhibits a novel distribution, a distribution pattern somewhat unexpected for a channel involved in chloride secretion. Immunolabeled ClC-2 was detected predominantly at the tight junction complex between adjacent intestinal epithelial cells.

tight junction; immunofluorescence; hypotonic shock; chloride efflux


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

THE MAJOR CHLORIDE CHANNEL thought, to date, to mediate chloride secretion in the small intestine is the cystic fibrosis transmembrane conductance regulator (CFTR) (1). However, it remains unclear whether CFTR is the only chloride channel capable of mediating secretion in this tissue (3, 10, 12). A previous study in rat ileum showed that a secretory chloride current can be activated by hypotonicity, but the molecular basis for the chloride conductance path was not determined (3). It has been suggested that the hypotonicity-activated chloride channel, ClC-2, may be capable of mediating chloride secretion in rodent neonatal airways because ClC-2 protein can be detected in the apical membrane of rodent neonatal airway epithelial cells (17). In addition, ClC-2 has been suggested to play a role in gastric chloride secretion (15). So far, only ClC-2 message expression has been studied in the intestinal epithelium and has been detected in the human intestinal cell line T84, in rat intestinal tissue (22) as well as in the murine duodenum (12). Evaluation of the potential role for ClC-2 in intestinal secretion requires an assessment of subcellular ClC-2 protein localization in this tissue.

In the present study, using wild-type (WT) and CFTR knockout mice, we showed that in the murine ileum, hypotonic shock elicits chloride secretion through a non-CFTR chloride channel that exhibits the pharmacological properties expected for ClC-2-mediated currents. However, assessment of native localization of ClC-2 in this tissue by confocal and electron microscopy revealed that ClC-2 exhibits a distribution pattern that is unexpected for a channel involved in chloride secretion. We detected ClC-2 protein predominantly at the tight junction complex between adjacent intestinal epithelial cells with only diffuse labeling of the apical brush-border membrane.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Mice. Experiments were conducted on male and female WT and cystic fibrosis (CF) mice, matched for genetic background (BalbC/129), age (6-8 wk old), and diet (normal chow). The phenotype of CF mice, or CFTR knockout (CftrmHSC/CftrmHSC) mice, used in this study was described previously (18). Briefly, all but 30% of animals with this genetic background die before 6 wk of age due to intestinal obstruction. In the present study, we investigated the molecular basis of the swelling-activated chloride conductance in ileal segments obtained from surviving CF mice and their WT siblings.

Ussing chamber analysis. Short-circuit current (Isc) measurements were performed on freshly excised WT and CF murine intestinal tissues from the ileum, defined as the segment 1-5 cm proximal to the cecum. Tissues were mounted in Ussing chambers with an aperture of 0.28 cm2. The buffer bathing the tissues was composed of 112 mM NaCl, 28 mM mannitol, 10 mM KHCO3, 1.2 mM K2HPO4, 2 mM CaCl2, 1.2 mM MgCl2, and 5 mM glucose in the apical bath and 5 mM mannitol in the serosal bath and was gassed with 95% O2 and heated to 37°C. After 5-8 min of measurement of basal current, the apical solution was changed to one lacking mannitol (28 mM) for 20% hypotonic shock [or 80% isotonicity, determined with a Wescor 5500 vapor pressure osmometer (Johns Scientific, Toronto, ON, Canada)]. The low-chloride buffer was prepared as above, except that 112 mM NaCl was replaced with 112 mM sodium gluconate and 5.8 mM CaCl2 to account for the calcium-chelating effect of gluconate (3). Stock solutions of 5-nitro-2-(3-phenylpropylamino) benzoic acid (NPPB) and 4,4'-diisothiocyanato-stilbene-2,2'-disulfonic acid (DIDS) were prepared in DMSO and used at a final concentration of 0.5 mM, applied into the apical bath.

RT-PCR analysis. Total RNA was isolated from murine intestinal tissues with the use of the Trizol extraction protocol (GIBCO-BRL) followed by DNA removal with (1 unit) DNase 1 (Ambion, Austin, TX). Reverse transcription with oligo(dT) primers (using the Superscript preamplification kit; GIBCO-BRL) was followed by PCR analysis in a 20-µl volume containing 0.5 µM of each primer (murine beta -actin or murine ClC-2: 5'-AGT TCC TAG AAT ATG GAC AGA GCC-3' and 5'-AAA GAG GGA GAG GAA CT-3' spanning 498 bp), 0.2 mM of each deoxynucleoside, 1 µl of cDNA, and 0.6 units of Ultratherm thermophilic DNA polymerase (Eclipse Molecular Biologicals, Ontario, ON, Canada). The ClC-2 fragment was amplified with a GeneAmp PCR System 9700 (Perkin Elmer/Applied Biosystems) with 35 cycles of 30 s at 94°C, 63°C, and 72°C. The PCR products were ligated into pCR2.1 vector (Invitrogen) with Taq polymerase (GIBCO-BRL), and the sequence identity was confirmed by BLAST sequence database search.

Anti-ClC-2 antibody. The polyclonal antibody used for detecting and localizing ClC-2 protein in the murine intestinal epithelium was generated against a glutathione-S-transferase (GST)-fusion peptide containing amino acids (residues 31-74) of rat ClC-2 (rClC-2 cDNA; kindly provided by T. Jentsch). The ClC-2-specific antibody was affinity purified as previously described (24).

Western analysis. Intestinal epithelial scrapings from three WT mice were added to 15 ml of lysis buffer containing 15 mM Tris, 10 mM KCl, 1.5 mM MgCl2, and protease inhibitors (10 µg/ml leupeptin, 10 µg/ml aprotinin, 1 mM benzamidine, 10 µM E64, and 2 mM dithiothreitol). The suspension was vortexed and centrifuged for 10 min at 4°C at 4,000 rpm (JA-17, Beckman) to remove organelles. The supernatant was centrifuged for 2 h at 4°C at 100,000 g to isolate crude membrane preparation. Of this preparation, 50 µg were analyzed by SDS-PAGE (8% gel) with the use of anti-ClC-2 antibody at a concentration of 1 µg/ml. Immunoreactive protein was detected using the enhanced chemiluminescence system (Life Science, Chicago, IL).

Immunoperoxidase staining. Immunoperoxidase staining was performed on 5-µm paraffin sections of Formalin-fixed tissues. The ClC-2 antibody was applied at a concentration of 116 µg/ml and was detected with a peroxidase-conjugated streptavidin system (Vectastain Elite ABC kit; Vector Laboratories, Peterborough, UK) and counterstained with hematoxylin. Specificity of the antibody for ClC-2 was assessed by competition using a 100-fold excess of antigenic ClC-2 fusion protein. Slides were viewed with the use of a ×10 objective on an Olympus Bx microscope, and images were captured using the CoolSnap program (Roper Scientific).

Immunofluorescence labeling. Immunofluorescence labeling was performed on 5-µm cryosections of intestinal tissues fixed in methanol at 4°C. After 2.5 h of incubation at 25°C with the zonula occludens-1 (ZO-1)-specific monoclonal antibody (2.5 ug/ml; Chemical International, Temecula, CA) and the affinity-purified polyclonal antibody against ClC-2 (58 µg/ml), sections were washed with PBS and incubated with FITC-conjugated anti-mouse or anti-rabbit secondary antibodies (0.02 mg/ml; Molecular Probes, Eugene, OR). Slides were viewed with a ×100 objective on an Olympus Vanox AHBT3 microscope, and with the use of epifluorescence, images were captured with the Image-Pro program (Media Cybernetics). For confocal microscopy, sections were viewed with a ×100 objective on a Leica TCS 4D microscope, and with the use of epifluorescence, images were captured using the SCANware 5.01 program.

Immunogold electron microscopy. Subcellular localization of immunogold-labeled ClC-2 in villus tip cells was detected by transmission electron microscopy. Paraformaldehyde- and glutaraldehyde-fixed tissues from the ileum of a WT mouse were infused with sucrose, frozen, and then substituted in absolute methanol containing uranyl acetate. Samples were then warmed, infiltrated in Lowicryl HM20 resin, embedded in gelatin capsules, and polymerized under ultraviolet light. Ultrathin sections were mounted on Formvar-coated nickel grids, incubated with affinity-purified ClC-2 antibody (580 µg/ml) followed by goat anti-rabbit IgG 10-nm gold complex, and stained with saturated aqueous uranyl acetate and lead citrate. Controls included omission of the primary antibody or competition with a 100-fold excess of the GST-ClC-2 fusion peptide or a 100-fold excess of GST alone.

Gold particle density determination. Random images of immunogold-labeled sections were captured with the use of a charge-coupled device camera (AMT, Danvers, MA) in the transmission electron microscope (JEOL 1200EXII; JEOL USA, Peabody, MA). A minimum of 3 fields from 25 epithelial cells were examined from 5 samples. Particle density was determined using NIH Image 1.51 (National Institutes of Health, Bethesda, MD). Density of labeling was determined for the apical cell junctions (particles/µm), apical membrane (particles/µm), and cytoplasm (particles/µm2). Data were expressed as means ± SE.

Statistics. The significance of differences between paired experimental groups was assessed using the Student's t-test for paired data.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

A non-CFTR chloride channel contributes to swelling-activated chloride secretion in the murine ileum. A previous study showed that hyposmotic media elicits chloride secretion in the rat ileum through an unidentified chloride channel presumably expressed in the apical membrane (3). To determine whether a similar swelling-induced chloride conductance is expressed in the apical membrane of the murine ileum, we applied hypotonic shock to the apical epithelial membrane of the ileum while monitoring the transepithelial Isc.

As previously reported by others (9), a luminally directed negative Isc was spontaneously generated by the intestinal epithelia of WT mice before hypotonic shock stimulation (i.e., -17.0 ± 2.7 µA/cm2, n = 10). In addition, intestinal segments from CFTR knockout mice also exhibited a spontaneous, luminally directed negative Isc (i.e., 12.2 ± 1.4 µA, n = 6). This basal current in CFTR knockout mice reflects chloride secretion, because it was decreased in response to apical application of the chloride channel blockers NPPB (500 µM), by 75.5 ± 9.1% (n = 7, P < 0.01) and glibenclamide (500 µM), by 73.3 ± 6.5% (n = 6, P < 0.02 ), as well as basolateral application of the Na+-K+-2Cl- cotransporter inhibitor bumetanide (200 µM), by 52.1 ± 14.4% (n = 5, P < 0.05). However, DIDS (500 µM) failed to inhibit this current, i.e., the change with DIDS was not significant (4.5% ± 2.98%, n = 6, P > 0.05). Therefore, a DIDS-insensitive, non-CFTR chloride channel likely contributes to basal chloride secretion across the ileum of CFTR knockout mice.

Hypotonic shock elicited a significant increase in luminally directed negative Isc across the ilea of WT mice (n = 10, P < 0.05; Fig. 1, A and B). This current was dependent on chloride, because there was no significant increase to hypotonic shock in low (15.6 mM)-chloride solutions in both compartments of the Ussing chamber (n = 9, P > 0.05). Furthermore, this current was rapidly reversed by apical application of the chloride channel blocker NPPB, but not the chloride channel blocker DIDS (Fig. 1, A and C). CFTR knockout mice also exhibited a significant (n = 6, P < 0.05) response to hypotonic shock in the ileum, which was sensitive to NPPB but not to DIDS (Fig. 1, A and B). There was a trend for the hypotonic shock response in the ileal segments obtained from CFTR knockout mice to be less than that observed in segments from WT mice, but the difference was not significant. Therefore, a non-CFTR chloride channel contributes to both hypotonic shock-evoked and basal chloride secretion across the ileum of CFTR knockout mice. ClC-2 is a possible candidate for mediating this secretion because it is known to be activated by hypotonic shock, inhibited by NPPB, and lacking in sensitivity to DIDS (24).


View larger version (30K):
[in this window]
[in a new window]
 
Fig. 1.   Hypotonicity of the mucosal bath activates chloride secretion in the ileum of wild-type (WT) and cystic fibrosis (CF) mice. A: sample traces showing the response to mucosal hypotonic shock (HTS) in the ileum of a WT (top) and a CF mouse (bottom). An increase in negative short-circuit current (Isc), reflecting electrogenic Cl- secretion across segment of ileum from WT and CF mice, is activated by exchange of the mucosal bath with one lacking mannitol (28 mM) for 20% hypotonic shock (HTS), as indicated by horizontal bar. In both WT and CF mice, the stimulated current is not inhibited by mucosal addition of the chloride channel blocker DIDS (0.5 mM) but is inhibited by the chloride channel blocker 5-nitro-2-(3-phenylpropylamino)benzoic acid (NPPB; 0.5 mM). B: mean changes in Isc caused by HTS in WT mice (filled bars, n = 10) and their CF siblings (open bars, n = 6). In both groups, HTS caused a significant increase in Isc in the ileum (P < 0.05, Student's paired t-test). C: %inhibition of the HTS response caused by DIDS (D) and NPPB (N) in the ileum of WT mice (filled bars) and their CF siblings (open bars). NPPB caused a significant decrease in the transepithelial current following HTS in the ileum of WT (P < 0.05, n = 8) and CF mice (P < 0.05, n = 5); DIDS did not affect this current in the ileum of either WT (P > 0.05, n = 10) or CF mice (P > 0.05, n = 6) (Student's paired t-test). In all cases, NPPB not only completely inhibited the HTS response but also inhibited basal Isc.

ClC-2 message and protein are expressed in the murine small intestinal epithelium. To determine whether ClC-2 expression is consistent with chloride secretion in the ileum, we first assessed ClC-2 mRNA and protein expression in the intestine of WT and CFTR knockout mice. Using RT-PCR analysis with sequence-specific primers to murine ClC-2 (see MATERIALS AND METHODS), we detected ClC-2 message in the ileum of adult WT and CFTR knockout mice (Fig. 2A). Therefore, ClC-2 message is expressed in the small intestine of both WT and CFTR knockout animals.


View larger version (33K):
[in this window]
[in a new window]
 
Fig. 2.   ClC-2 message and protein expression in the murine intestine. A: RT-PCR, with murine-specific ClC-2 primers spanning 498 bp, shows ClC-2 mRNA expression in the ileum of 2 WT and 2 CF mice. Negative controls without cDNA (H2O) or without reverse transcriptase (-RT) were performed. The PCR sequence identity was confirmed as ClC-2 by BLAST sequence database search. Murine beta -actin expression was equal in each RT reaction as confirmed with murine beta -actin-specific primers spanning ~800 bp. Size of RT-PCR products (in bp) is indicated at right. B, left: Western blot analysis shows that the polyclonal anti-ClC-2 antibody generated against a glutathione-S-transferase (GST)-fusion peptide containing amino acids (residues 31-74) of rat ClC-2 recognizes a broad band that corresponds to molecular mass of ~90-97 kDa (close to the predicted mass of rat ClC-2) in murine brain tissue (left lane). This 90- to 97-kDa band (not other minor bands) is competed with a 100-fold excess of the antigenic fusion peptide (middle lane), but not with GST alone (right lane), confirming antibody specificity. B, right: immunoblot of murine small intestinal epithelial scrapings shows that the ClC-2 antibody recognizes a single band of ~90-97 kDa, corresponding to the expression of ClC-2 protein in this tissue. The ClC-2 specific antibody was affinity purified as previously described (24).

Previously, we published a description of an affinity-purified polyclonal antibody prepared against the amino terminus (residues 31-74) of rClC-2 (24), a region that is not conserved among different members of the ClC family of chloride channels (13). We showed that this antibody recognized recombinant ClC-2 expressed in Sf9 cells (24). In the present studies, we first assessed the specificity of this antibody in whole mouse brain lysates because ClC-2 is known to be abundantly expressed in brain tissue (19, 22). We showed that, in the murine brain, this polyclonal antibody recognized a broad band of ~90 kDa in molecular mass by Western blot analysis (Fig. 2B). This signal was not competed by GST alone, but it was competed by the fusion peptide of the ClC-2 amino terminus with GST, attesting to its specificity. In addition, this antibody recognized a broad band of ~90-97 kDa in epithelial scrapings from ileal segments from WT mice (Fig. 2C) and CFTR knockout mice, consistent with the expression of ClC-2 protein in the small intestine.

In agreement with our Western blot analysis, immunoperoxidase staining of Formalin-fixed tissues using the same polyclonal anti-ClC-2 antibody indicated that there is pronounced staining of the epithelium in the ileum of the small intestine of both WT and CFTR knockout mice. ClC-2 was expressed primarily at the villus tip of the mucosa lining the ileal section obtained from WT and CFTR knockout mice (Fig. 3, A and B, respectively). No difference in ClC-2 protein expression pattern was detected between WT and CFTR knockout mice. The specificity of this staining pattern was confirmed by competition with the antigenic peptide to which the antibody was generated and also by omitting the primary antibody (Fig. 3, C and D, respectively).


View larger version (46K):
[in this window]
[in a new window]
 
Fig. 3.   A: ClC-2 protein is primarily localized at the surface epithelium of the small intestine. B: immunoperoxidase stained cross section of ileum from a WT mouse shows ClC-2-specific staining predominantly at the tips of the intestinal villi with weak staining in crypts. Sections were counterstained with hematoxylin. C: ClC-2 immunostaining in the ileum of a CF mouse resembles that of WT ileum. Staining is specific for ClC-2 because it was competed by preincubation with a 100-fold excess of antigenic ClC-2 fusion protein. D: immunostaining was absent when the anti-ClC-2 antibody was omitted. Images were viewed with a ×10 objective.

The subcellular distribution of ClC-2 protein decorated by immunofluorescence in the villus tips of the murine small intestine was assessed first by epifluorescence microscopy. ClC-2 immunolabeling of the intestinal epithelium predominantly showed a punctate pattern with hot spots at the apical intercellular junction (Fig. 4A, top). This pattern of labeling resembles that of the tight junction protein ZO-1 (Fig. 4A, middle). The specificity of the ClC-2 immunofluorescence pattern was confirmed by competition with the ClC-2 antigenic peptide (Fig. 4A, bottom). Figure 4B shows the results of double-labeling of murine ileal epithelium with the ClC-2 polyclonal antibody (Fig. 4B, top, red) and the ZO-1 antibody (Fig. 4B, middle, green), which revealed that the two proteins colocalize at the tight junction (Fig. 4B, bottom, yellow). Furthermore, we performed confocal microscopy studies of ileal segments double-labeled using ClC-2 (red) and ZO-1 (green) antibodies and confirmed colocalization (yellow) of these proteins in a single confocal section in the region of the tight junctions (Fig. 4C). The subcellular localization of ClC-2 was identical for intestinal segments obtained from both WT and CFTR knockout mice.


View larger version (76K):
[in this window]
[in a new window]
 
Fig. 4.   Subcellular localization of ClC-2 protein. A: fluorescent immunostaining of epithelial cells at the villus tip in a WT mouse (top), with the polyclonal anti-ClC-2 antibody previously described. This expression pattern resembles that obtained with a monoclonal antibody against the tight junction protein ZO-1 (middle). ClC-2 staining pattern in the murine small intestine is specific because it can be competed by preincubation with 100-fold excess of the ClC-2 fusion protein (bottom). B, top: sections obtained from a WT mouse, double-labeled with the anti-ClC-2 antibody (red) and ZO-1 (green). The yellow image shows colocalization of these proteins in WT ileum. Bottom: sections of ileum from a CF mouse, double-labeled with the anti-ClC-2 antibody (red) and a monoclonal anti-ZO-1 antibody (green). The yellow image shows that these proteins colocalize in CF mouse ileum. Images were viewed with a ×40 objective. C: confocal microscopy showing colocalization (yellow signal) of ClC-2 (red) and ZO-1 (green) in the same horizontal optical section. Images were viewed with a ×100 objective.

ClC-2 protein localizes primarily to the tight junction complex and diffusely to the apical membrane. As previously mentioned, our immunofluorescence studies show that ClC-2 is predominantly situated in proximity to the tight junction complex. Similarly, electron microscopy revealed that ClC-2 decorated by immunogold was located predominantly, but not exclusively, in clusters at the apical aspect of the tight junction complex (Fig. 5). Gold particles were counted in a minimum of 3 fields from 25 epithelial cells from 5 different intestinal sections. In proximity to the tight junction, we found 4.2 ± 1.3 particles of gold per micrometer of membrane. On the other hand, in the brush border, we found 0.04 ± 0.03 particles of gold per micromillimeter of membrane, similar to the number of grains detected per square micrometer of cytoplasm (0.03 ± 0.01). This labeling pattern was effectively competed with the GST-ClC-2 peptide against which the ClC-2 antibody was raised (Fig. 5B), but not by GST alone (data not shown).


View larger version (143K):
[in this window]
[in a new window]
 
Fig. 5.   Subcellular localization of immunogold-labeled ClC-2 in villus tip cells by transmission electron microscopy. A: image of a section of the brush border and apical cell junction of WT mouse ileum. Immunogold-labeled ClC-2 is concentrated at the border between cells (arrowhead). Inset: tight junction region (*). B: signal obtained with the anti-ClC-2 antibody could be competed with the antigenic ClC-2 peptide. Bar, 0.2 µm. C: higher magnification of ClC-2 (decorated with immunogold) in the membranes at the tight junction (*). Bar, 0.1 µm. All grids were photographed in the transmission electron microscope.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

In this work we described a non-CFTR chloride secretory current in mouse small intestine that is activated by hypotonic solutions. This chloride secretory current shares properties previously reported for a secretory current described by Diener et al. (3) in the rat small intestine, namely, its activation by hypotonicity, inhibition by NPPB, and lack of sensitivity to DIDS. We suggest that this activity is mediated by ClC-2 because ClC-2 is known to be activated by hypotonic solutions in heterologous expression systems (22, 24). Furthermore, we found that ClC-2 protein is expressed endogenously in the epithelium of intestinal villi. While the classic model of secretion postulates that chloride secretion occurs primarily from the intestinal crypts (23), there are several reports suggesting that chloride secretion can also occur from the intestinal villi (2, 4, 20). Using a combination of fluorescence and electron microscopy, we determined that ClC-2 protein in the murine intestinal epithelium is concentrated close to the membranes of the tight junction complex. Therefore, ClC-2 has a novel subcellular distribution pattern that has not been described previously for chloride channels involved in anion secretion. Hence, further study will be required to determine the role of ClC-2 in chloride secretion by the intestinal epithelium.

As previously mentioned, ClC-2 is known to be activated by hypotonicity and, hence, may mediate the secretory chloride response observed in CFTR knockout mice. However, other non-CFTR chloride channels may possibly participate in this response. Our pharmacological analyses suggest that the response is relatively insensitive to the chloride channel blocker DIDS. Because DIDS, in concentrations comparable to those employed in the present study, is known to inhibit calcium-activated currents in intestinal epithelia (possibly mediated by murine calcium-sensitive chloride channel) (1, 10), the ubiquitous swelling-activated volume-sensitive organic osmolyte/anion channel, and ClC-3 (5, 21), these channels are not likely to play a role in the response. Ultimately, it will be necessary to generate a ClC-2 knockout mouse to define the relative contribution of ClC-2 to the secretion observed.

The subcellular distribution of ClC-2 is unique for an anion channel because it is concentrated close to the tight junction complex. The tight junction, a dynamic structure in epithelial tissue, plays a key role in regulating paracellular transport and the barrier functions of the intestine. Hence, we plan to directly assess the role for ClC-2 in these functions in our future studies. Furthermore, it remains to be determined whether specific interactions exist between ClC-2 and scaffolding and/or cytoskeletal proteins that reside in the tight junction complex, i.e., the ZO family of proteins, which participate in the targeting or retention of ClC-2 at this site in the small intestine (6-8, 16). We suggest that such interactions may be relatively specific for the small intestine; as Murray et al. (17) localized ClC-2 to the apical membrane of airway epithelial cells and in the mouse colon, ClC-2 is localized at the basolateral membrane (Gyömörey K and Bear CE, unpublished observations).

The physiological stimulus that activates ClC-2 in native tissue is not known. In heterologous expression systems, ClC-2 currents appear to require activation by stressors such as hypotonic shock, low pH, or hyperpolarization to -100 mV (11). On the other hand, ClC-2 endogenously expressed in neuronal cells is basally active at resting membrane potentials (19), suggesting that ClC-2 regulation may vary with the host cell type. We observed that the ileal epithelium of CFTR knockout mice is capable of supporting NPPB-sensitive, DIDS-insensitive chloride secretion under resting conditions and suggest that ClC-2 may be partially active under basal conditions in this tissue, as in neuronal cells (19). In our studies, luminal hypotonic shock appeared to stimulate ClC-2 activity. Osmotic change may in fact be a physiological regulator of ClC-2 in the intestine because absorptive epithelia experience local changes in osmolarity as a result of nutrient transport (14). In fact, expression of ClC-2 at the villus tip enterocytes of the jejunum overlaps with the expression of the sodium-glucose cotransporter SGLT1 at this site (25). It also has been suggested that phosphorylation by protein kinase A and protein kinase C can modulate ClC-2 currents (15, 19). The role of these signaling pathways in the regulation of ClC-2 in native epithelial cells will be the subject of future studies.


    ACKNOWLEDGEMENTS

CFTR knockout animals and WT siblings were generously provided by Lap-Chee Tsui and Richard Rozmahel (Genetics Department, Research Institute).


    FOOTNOTES

This research was supported by operating grants to C. E. Bear from the Canadian Cystic Fibrosis Foundation (CCFF), Medical Research Council (MRC), and National Institutes of Health Specialized Center of Research Grant 01 P50 DK-490096-06. K. Gyömörey was supported by a CCFF Studentship Award, and C. E. Bear was supported by an MRC Scientist Award.

Address for reprint requests and other correspondence: C. E. Bear, Division of Cell Biology, Research Institute, Hospital for Sick Children, Dept. of Physiology, Univ. of Toronto, 555 Univ. Ave., Toronto, Ontario, Canada M5G 1X8 (E-mail: bear{at}sickkids.on.ca).

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.

Received 5 May 2000; accepted in final form 27 July 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Anderson, MP, Sheppard DN, Berger HA, and Welsh M. Chloride channels in the apical membrane of normal and cystic fibrosis airway and intestinal epithelia. Am J Physiol Lung Cell Mol Physiol 263: L1-L4, 1992[Abstract/Free Full Text].

2.   Butt, AG, Bowler JM, and McLaughlin CW. Villus and crypt cell composition in the secreting mouse jejunum measured with x-ray microanalysis. J Membr Biol 162: 17-29, 1998[ISI][Medline].

3.   Diener, M, Bertog M, Fromm M, and Scharrer E. Segmental heterogeneity of swelling-induced Cl- transport in rat small intestine. Pflügers Arch 432: 293-300, 1996[ISI][Medline].

4.   Donowitz, M, and Madara JL. Effect of extracellular calcium depletion on epithelial structure and function in rabbit ileum: a model for selective crypt or villus epithelial cell damage and suggestion of secretion by villus epithelial cells. Gastroenterology 83: 1231-1243, 1982[ISI][Medline].

5.   Duan, D, Winter C, Cowley S, Hume JR, and Horowitz B. Molecular identification of a volume-regulated chloride channel. Nature 390: 417-421, 1997[ISI][Medline].

6.   Fanning, AS, and Anderson JM. Protein molecules as organizers of membrane structure. Curr Opin Cell Biol 11: 432-439, 1999[ISI][Medline].

7.   Fanning, AS, Jameson BJ, Jesaitis LA, and Anderson JM. The tight junction protein ZO-1 establishes a link between the transmembrane protein occludin and the actin cytoskeleton. J Biol Chem 273: 29745-29753, 1998[Abstract/Free Full Text].

8.   Furuse, M, Fujita K, Hiiragi T, Fujimoto K, and Tsukita S. Claudin-1 and -2: novel integral membrane proteins localizing at the tight junction with no sequence similarity to occludin. J Cell Biol 141: 1539-1550, 1998[Abstract/Free Full Text].

9.   Grubb, BR. Ion transport across the jejunum in normal and cystic fibrosis mice. Am J Physiol Gastrointest Liver Physiol 268: G505-G513, 1995[Abstract/Free Full Text].

10.   Gruber, AD, Gandhi R, and Pauli BU. The murine calcium-sensitive chloride channel (mCaCC) is widely expressed in secretory epithelia and in other select tissues. Histochem Cell Biol 110: 43-49, 1998[ISI][Medline].

11.   Jentsch, TJ, Friedrich T, Schriever A, and Yamada H. The CLC chloride channel family. Pflügers Arch 437: 783-795, 1999[ISI][Medline].

12.   Joo, NS, Clarke LL, Han BH, Forte LR, and Kim HD. Cloning of ClC-2 chloride channel from murine duodenum and its presence in CFTR knockout mice. Biochim Biophys Acta 1446: 431-437, 1999[ISI][Medline].

13.   Jordt, SE, and Jentsch TJ. Molecular dissection in the ClC-2 chloride channel. EMBO J 16: 1582-1592, 1997[Abstract/Free Full Text].

14.   MacLeod, RJ, and Hamilton JR. Volume regulation initiated by Na(+)-nutrient cotransport in isolated mammalian villus enterocytes. Am J Physiol Gastrointest Liver Physiol 260: G26-G33, 1991[Abstract/Free Full Text].

15.   Malinowska, DH, Kupert EY, Bahinski A, Sherry AM, and Cuppoletti J. Cloning, functional expression, and characterization of a PKA-activated gastric Cl- channel. Am J Physiol Cell Physiol 268: C191-C200, 1995[Abstract/Free Full Text].

16.   Matter, K, and Balda MS. Occludin and the functions of tight junctions. Int Rev Cytol 187: 117-146, 1999.

17.   Murray, CB, Morales MM, Flotte TR, McGrath-Morrow SA, Guggino WB, and Zeitlin PL. ClC-2: a developmentally dependent chloride channel expressed in the fetal lung and downregulated after birth. Am J Respir Cell Mol Biol 12: 597-604, 1995[Abstract].

18.   Rozmahel, R, Wilschanski M, Matin A, Plyte S, Oliver M, Auerbach W, Moore Forstner J, Durie P, Nadeau J, Bear C, and Tsui LC. Modulation of disease severity in cystic fibrosis transmembrane conductance regulator deficient mice by a secondary genetic factor. Nat Genet 12: 280-287, 1996[ISI][Medline].

19.   Staley, K, Smith R, Schaack J, Wilcox C, and Jentsch T. Alteration of GABAA receptor function following gene transfer of the CLC-2 chloride channel. Neuron 17: 543-551, 1996[ISI][Medline].

20.   Stewart, CP, and Turnberg LA. A microelectrode study of responses to secretagogues by epithelial cells on villus and crypt of rat small intestine. Am J Physiol Gastrointest Liver Physiol 257: G334-G343, 1989[Abstract/Free Full Text].

21.   Strange, K, Emma F, and Jackson PS. Cellular and molecular physiology of volume-sensitive anion channels. Am J Physiol Cell Physiol 270: C711-C730, 1996[Abstract/Free Full Text].

22.   Thiemann, A, Gründer S, Pusch M, and Jentsch T. A chloride channel widely expressed in epithelial and non-epithelial cells. Nature 356: 57-60, 1992[ISI][Medline].

23.   Welsh, MJ, Smith PL, Fromm M, and Frizzell RA. Crypts are the site of intestinal fluid and electrolyte secretion. Science 218: 1219-1221, 1982[ISI][Medline].

24.   Xiong, H, Li C, Garami E, Wang Y, Ramjeesingh M, Galley K, and Bear CE. ClC-2 activation modulates regulatory volume decrease. J Membr Biol 167: 215-221, 1999[ISI][Medline].

25.   Yoshida, A, Takata K, Kasahara T, Aoyagi T, Saito T, and Hirano H. Immunohistochemical localization of Na(+)-dependent glucose transporter in the rat digestive tract. Histochem J 27: 420-426, 1995[ISI][Medline].


Am J Physiol Cell Physiol 279(6):C1787-C1794
0363-6143/00 $5.00 Copyright © 2000 the American Physiological Society