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Am J Physiol Gastrointest Liver Physiol 292: G1079-G1088, 2007. First published December 14, 2006; doi:10.1152/ajpgi.00354.2006
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MUCOSAL BIOLOGY

PAT-1 (Slc26a6) is the predominant apical membrane Cl–/HCO3– exchanger in the upper villous epithelium of the murine duodenum

Janet E. Simpson,1,2,3 Clifford W. Schweinfest,4 Gary E. Shull,5 Lara R. Gawenis,5 Nancy M. Walker,3 Kathryn T. Boyle,3 Manoocher Soleimani,6 and Lane L. Clarke1,3

Departments of 1Biomedical Sciences and 2Veterinary Pathobiology and 3Dalton Cardiovascular Research Center, University of Missouri, Columbia, Missouri; 4Department of Medicine, Medical University of South Carolina, Charleston, South Carolina; and Departments of 5Molecular Genetics, Biochemistry, and Microbiology and 6Medicine, University of Cincinnati, Cincinnati, Ohio

Submitted 1 August 2006 ; accepted in final form 12 December 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Basal HCO3– secretion across the duodenum has been shown in several species to principally involve the activity of apical membrane Cl–/HCO3– exchanger(s). To investigate the identity of relevant anion exchanger(s), experiments were performed using wild-type (WT) mice and mice with gene-targeted deletion of the following Cl–/HCO3– exchangers localized to the apical membrane of murine duodenal villi: Slc26a3 [down-regulated in adenoma (DRA)], Slc26a6 [putative anion transporter 1 (PAT-1)], and Slc4a9 [anion exchanger 4 (AE4)]. RT-PCR of the isolated villous epithelium demonstrated PAT-1, DRA, and AE4 mRNA expression. Using the pH-sensitive dye BCECF, anion exchange rates were measured across the apical membrane of epithelial cells in the upper villus of the intact duodenal mucosa. Under basal conditions, Cl–/HCO3– exchange activity was reduced by 65–80% in the PAT-1(–) duodenum, 30–40% in the DRA(–) duodenum, and <5% in the AE4(–) duodenum compared with the WT duodenum. SO42–/HCO3– exchange was eliminated in the PAT-1(–) duodenum but was not affected in the DRA(–) and AE4(–) duodenum relative to the WT duodenum. Intracellular pH (pHi) was reduced in the PAT-1(–) villous epithelium but increased to WT levels in the absence of CO2/HCO3– or during methazolamide treatment. Further experiments under physiological conditions indicated active pHi compensation in the PAT-1(–) villous epithelium by combined activities of Na+/H+ exchanger 1 and Cl–-dependent transport processes at the basolateral membrane. We conclude that 1) PAT-1 is the major contributor to basal Cl–/HCO3– and SO42–/HCO3– exchange across the apical membrane and 2) PAT-1 plays a role in pHi regulation in the upper villous epithelium of the murine duodenum.

bicarbonate secretion; intestine; cystic fibrosis transmembrane conductance regulator; Slc4; putative anion transporter 1; downregulated in adenoma; anion exhanger isoform 4

DUODENAL epithelial HCO3– secretion plays a critical role in the protection of the epithelium against damage from acidic chyme entering into the proximal duodenum (13). The process of active HCO3– secretion across the duodenum involves the apical membrane activities of the CFTR and anion exchange protein(s), which can mediate Cl– absorption in exchange for luminal HCO3– secretion (10, 32, 37). Under basal (unstimulated) conditions, pH stat studies in several species have indicated that Cl–/HCO3– exchange dominates duodenal HCO3– secretion and, in the CFTR– intestine, affords a degree of protection against acidic luminal pH (10, 11, 31, 37). However, the identity of the anion exchange protein(s) involved in duodenal HCO3– secretion and their functional contribution along the crypt-villus axis have yet to be resolved.

The principal candidates for this process are two members of the multifunctional anion exchanger (Slc26a) family, i.e., Slc26a3 [known as downregulated in adenoma (DRA)] and Slc26a6 [known as putative anion transporter 1 (PAT-1) or Cl–/formate exchanger] and one member of the HCO3– transporter family (Slc4a), i.e., Slc4a9 [known as anion exchanger isoform 4 (AE4)]. All three anion exchangers have been immunolocalized to the apical membrane of the intestinal epithelium, especially along the villous axis (18, 22, 45, 48). Studies utilizing recombinant proteins have shown that DRA transports HCO3– in exchange for luminal Cl–, SO42–, or oxalate (29). PAT-1 is more versatile, demonstrating Cl–, oxalate, SO42–, formate, HCO3–, and hydroxyl transport in Xenopus oocytes (29). Although controversial (7), recent studies have suggested that both DRA and PAT-1 exhibit electrogenic properties, leading to the proposal that that DRA and PAT-1 have apparent stoichiometries of 2 Cl–:1 HCO3– and 1 Cl–:2 HCO3–, respectively (33). Less well characterized is the AE4 protein, but a functional expression study (48) has shown that AE4 exhibits Cl–/HCO3– exchange (48).

Disease-causing loss-of-function mutations have not been reported for human PAT-1 (SLC26A6) or AE4 (SLC4A9). However, loss-of-function mutations in DRA (SLC26A3) are the causal factor in the inherited disease congenital chloride-losing diarrhea (CLD). These patients have defective intestinal Cl–/HCO3– exchange resulting in diarrhea with high Cl– content and systemic alkalinization (20, 28). Likewise, initial reports of clinical disease in the DRA knockout mouse model (i.e., diarrhea, elevated chloride stool concentration, and growth retardation) closely resemble that of human CLD (6). The severe intestinal manifestations of CLD may reflect an interaction between DRA and the major apical membrane Na+/H+ exchangers (NHE2 and NHE3) responsible for electroneutral NaCl absorption, as based on evidence of functional coupling of these exchangers in recombinant cell systems and changes in DRA expression in the NHE3 knockout intestine (26, 30).

Duodenal villi are directly exposed to gastric acid chyme from the stomach, but little is known regarding the physiology of HCO3– transport in the villous epithelium where DRA, PAT-1, and AE4 are reportedly expressed. Recently, we (35) have demonstrated the capability to directly measure Cl–/HCO3– exchange activity across the apical membrane of epithelial cells in the upper half of villi in the intact murine duodenal mucosa using BCECF-AM microfluorometry of intracellular pH (pHi). This investigation found robust Cl–/HCO3– exchange activity in the upper villus with exchange rates similar to or greater than those reported for exchanger isoforms expressed in heterologous cell systems (7, 38, 45); however, the in situ exchange process varied from recombinant protein studies by exhibiting relative insensitivity to distilbenes but not niflumic acid. It is important to examine the activity of specific anion exchangers in the native intestinal epithelium, but, unfortunately, there are no known inhibitors of anion exchangers that effectively discriminate between exchanger isoforms or lack additional effects on other anion transport proteins. Therefore, in the present study, we used the intestinal mucosa from mice with gene-targeted deletions of PAT-1, DRA, and AE4 and their wild-type (WT) littermates to assess the relative contributions of each exchanger to the process of apical Cl–/HCO3– exchange across the duodenal upper villous epithelium.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals. The experiments in this study were performed on mice with gene-targeted disruptions of the murine homologs of PAT-1 (44), DRA (6), or AE4 on a mixed genetic background. AE4-deficient mice were prepared by deleting a portion of exon 12 and all of exon 13, thereby eliminating codons 558–626 of mouse AE4 (Accession No. AK142545) and terminating the open reading frame at the beginning of the transmembrane domains (L. R. Gawenis and G. E. Shull, unpublished data). All comparisons of homozygous knockout (–/–) mice were made with gender- and age-matched (+/+; WT) siblings. Mutant mice were identified using a PCR-based analysis of tail-snip DNA, as previously described (9). All mice were maintained ad libitum on standard laboratory chow (Formulab 5008 Rodent Chow, Ralston Purina) and tap water. Mice were housed singly in a temperature (22–26°C)- and light (12:12-h light-dark cycle)-controlled room in the American Association for Accreditation of Laboratory Animal Care-accredited animal facility at Dalton Cardiovascular Research Center. Intestinal tissues for experiments were obtained from mice of 2–4 mo of age. These mice were fasted overnight prior to experimentation but were provided with water ad libitum. All experiments involving animals were approved by the University of Missouri Animal Care and Use Committee.

Fluorescence measurements of pHi and image analysis. The method used for imaging villous epithelial cells in the intact murine intestine has been previously described (16, 35). Briefly, after the mouse had been euthanasized, the proximal duodenum was isolated, stripped of the outer muscle layers by blunt dissection, and mounted luminal side up in a horizontal Ussing-type perfusion chamber where luminal and serosal surfaces of the tissue were independently bathed. The luminal superfusate was an isethionate-bicarbonate-Ringer (IBR) solution containing (in mmol/l) 140.0 Na+, 55.0 Cl–, 55.0 isethionate–, 25.0 HCO3–, 5.2 K+, 5.0 TES, 4.8 gluconate–, 2.8 PO43–, 1.2 Ca2+, 1.2 Mg2+, 10.0 glucose, and 6.8 mannitol that was gassed with 95% O2-5% CO2 at 37°C (pH 7.4). The serosal superfusate was a Cl–-free IBR solution (with Cl– replaced with isethionate) gassed with 95% O2-5% CO2 at 37°C (pH 7.4). All duodena were treated with indomethacin (1 µM, bilateral) and TTX (0.1 µM, serosal) to minimize the effect of endogenous prostaglandins and neural tone, respectively (5, 34). During superfusion, the duodenum was treated on the serosal side with 1 µM EIPA to block the activity of NHE1. Villi immobilized by a fine nylon mesh overlay were incubated for 5 min with a luminal solution containing 100 µM DL-DTT to remove mucus and then incubated on the luminal side with 16 µM BCECF-AM for 10 min before superfusion of the luminal surface. Using a x40 water-immersion objective (Olympus, Melville, NY), 10 epithelial cells from the mid to upper region of a single villus were selected for ratiometric analysis. Changes in pHi were measured by dual-excitation wavelength techniques (440 and 495 nm), and villi were imaged at 535-nm emission. Ratiometric images were obtained at 20-s intervals with a Sensi-Cam digital camera (Cooke, Auburn Heights, MI) and processed using Axon Imaging Workbench 2.2 (Axon Instruments, Union City, CA). The 495-to-440-nm ratios were converted to pHi using a standard curve generated by the K+/nigericin technique (4, 40). The intrinsic buffering capacity (betai) of duodenal villous cells was estimated by the ammonium prepulse technique, and the total buffering capacity (betatotal) was calculated from the equation betatotal = betai + betaHCO3– = betai + 2.3 x [HCO3–]i, where betaHCO3– is the buffering capacity of the HCO3–/CO2 system and [HCO3–]i is the intracellular concentration of HCO3–. The rate of pHi change during the initial 90-s period of linear {Delta}pH/{Delta}t change was converted to the transmembrane flux (J) of HCO3– or H+ (JHCO3– and JH+; measured in mM/min) using the following equation: J = {Delta}pH/{Delta}t x betatotal (46).

Measurement of apical Cl–/HCO3– and Na+/H+ exchange. For measurements of Cl–/HCO3– exchange, duodenal preparations were superfused with IBR solution on the luminal side, and pHi alkalinization was induced by replacement of Cl– with isethionate– on an equimolar basis. After a stable pHi (~2 min) had been attained, pHi recovery was initiated by the readdition of Cl– to the luminal superfusate. In some experiments, SO42– was used to replace Cl– on an equimolar basis during pHi recovery. For methazolamide experiments, 100 µM methazolamide was initially added to the superfusate from a 10 mM stock in Cl–-free IBR solution. For bilateral Krebs-bicarbonate-Ringer (KBR) experiments, the luminal superfusate was KBR solution, which contained (in mM) 140.0 Na+, 5.2 K+, 25 HCO3–, 5 TES, 2.8 PO43–, 110 Cl–, 1.2 Ca2+, 1.2 Mg2+, 4.8 gluconate–, and 16.8 mannitol and was gassed with 95% O2-5% CO2 at 37°C (pH 7.4). The serosal perfusate was similar in composition to the luminal solution except that 10 mM mannitol was replaced (equimolar) with glucose. Rates of anion exchange during alkalinization and recovery ({Delta}pHi/min) were calculated from a linear regression of the values from the first 90 s of the initial pHi changes during Cl– removal and replacement, respectively, and these rates were converted to JHCO3–.

For measurements of apical membrane Na+/H+ exchange activity, the proximal duodenum was superfused with nominally CO2/HCO3–-free solutions where NaHCO3 was replaced equimolar with NaTES and gassed with 100% O2. Experiments to measure Na+/H+ exchange consisted of pHi acidification induced by replacement of luminal Na+ with NMDG+ on an equimolar basis. After a stable pHi had been obtained (~2 min), pHi recovery was initiated by replacing NMDG+ with Na+. Rates of anion exchange during Na+/H+ exchange were calculated from a linear regression of the values from the first 90 s of the initial pHi changes during Na+ replacement, and these rates were converted to JH+.

Experiments of recombinant murine DRA. Murine (m)DRA (a gift of J. Melvin, University of Rochester) was inserted into the multiple cloning site of a bicistronic pIRES plasmid (Invitrogen, Carlsbad, CA) containing DSRed2, a red fluorescent protein (RFP). The resulting plasmid was transformed into DH5{alpha}-competent cells (Invitrogen) by heat shock, and single colonies were picked for plasmid isolation (after overnight growth) by the alkaline lysis method. The plasmid was digested with HincII to confirm the proper orientation of the DRA insert. Plasmid containing mDRA or empty vector (control) was transfected using Superfect transfection reagent (Qiagen, Valencia, CA) according to the manufacturer's directions into CHO cells grown on coverslips (50% confluent). Cells expressing RFP were selected for pHi experiments 24 h after transfection.

Isolation of villous epithelium and RT-PCR. Duodenal segments were resected, opened longitudinally along the mesenteric border, and washed in oxygenated physiological Ringer solution containing 100 µM DTT. Under a dissecting microscope, the segment was placed mucosal side up over a parallel ridge (1-mm height) on a plastic surface and drawn beneath a no. 11 scalpel blade (perpendicular to the villi), thereby scraping the villous epithelium from the villous cores. Isolated villi were dispersed in a small volume (0.5 ml) of oxygenated Ringer solution containing 800 U/ml RNAse inhibitor (Qiagen) using a large-bore transfer pipette and immediately frozen in liquid nitrogen. Total RNA was isolated using TRI-Reagent (Molecular Research Center, Cincinnati, OH) according to the manufacturer's instructions and dissolved in 100 µl RNAse-free water. The Superscript preamplification system (Invitrogen) was used for first-strand cDNA synthesis according to the manufacturer's instructions. Each PCR of 25 µl contained 1 µl cDNA, 1x PCR Mg2+-free buffer, 2 mM MgCl2, 0.2 mM dNTP, 0.4 mM primer, and 0.5 U/reaction Taq DNA polymerase (Promega, Madison, WI). A master mix was made and added to 0.2-ml reaction tubes containing either cDNA (±reverse transcriptase) or water (as the negative control). Primers for DRA, PAT-1, and AE4 were designed based on murine sequences from the National Center for Biotechnology Information database. The primers for DRA were 5'-GGTTTAGCATTTGCTCTGCTGG-3' and 5'-TTACAGTCATGATGAGTTCGATG-3'. The primers for PAT-1 were 5'-GCGACTCTCTGAAAGAGAAGTG-3' and 5'-TCAGAGTTTGGTGGCCAAAACA-3'. The primers for AE4 were 5'-CATGCCTGGTCAAGAAAGCTAG-3' and 5'-CACTCATGTTACTGGGCCTGGTGG-3'. RT-PCR was performed on samples using RT-PCR cycles consisting of a 5.0-min denaturation at 94°C followed by 60 cycles of 94°C for 30 s, 60°C for 1.0 min, and 72°C for 1.0 min and ended with 7.0 min at 72°C using a MWG Primus 96 plus thermocycler (MWG-Biotech, Highpoint, NC). RT-PCR products were electrophoresed on a 2% agarose gel for analysis. To evaluate genomic DNA contamination of the RNA samples, RT-PCR was performed on samples using the following primers designed to amplify genomic DNA and cDNA of beta-actin: 5'-TGTTACCAACTGGGACGACA-3' and 5'-TCTCAGCTGTGGTGGTGAAG-3'. The absence of an amplicon in the no reverse transcriptase sample indicated minimal genomic DNA contamination. mRNA expression for each exchanger was compared against a similarly treated tissue control reported to express high levels of the specific transcript.

Western blot analysis of NHE3. For Western blot analysis, duodenal microsomal membranes were prepared from age-matched PAT-1(–) and WT adult mice as previously described (3, 41, 45). Briefly, membrane preparations were separated by SDS-PAGE and transferred to nitrocellulose membranes, blocked with 5% milk proteins, and probed with affinity-purified primary antibodies against NHE3 or beta-actin. The monoclonal anti-NHE3 antibody was directed against the COOH-terminal 131 amino acids of rabbit NHE3 (41), and beta-actin monoclonal antibodies were purchased from Alpha Diagnostics (San Antonio, TX). The secondary antibody was an anti-rabbit IgG conjugated to horseradish peroxidase (GIBCO-BRL, Gaithersburg, MD), which was visualized using chemiluminescence (SuperSignal Substrate, Pierce) and captured on light-sensitive imaging film (Kodak). Bands corresponding to NHE3 and beta-actin proteins were quantified by densitometric analysis (UN-SCAN-IT gel software, Silk Scientific, Orem, UT) and were expressed as percentages of control. The equity in protein loading in all blots was verified by gel staining using Coomassie brilliant blue R-250 (Bio-Rad, Hercules, CA).

Materials. The fluorescent dye BCECF-AM was obtained from Invitrogen. TTX was obtained from BioMol. All other materials were obtained from either Sigma-Aldrich (St. Louis, MO) or Fisher Scientific (Springfield, NJ).

Statistics. All values are reported as means ± SE. Data between two treatment groups were compared using a two-tailed unpaired Student t-test assuming equal variances between groups. A probability value of P < 0.05 was considered statistically significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Expression of DRA, PAT-1, and AE4 in the isolated duodenal villous epithelium. DRA, PAT-1, and AE4 have previously been localized to the apical membrane of the intestinal epithelia (18, 22, 45, 48). To verify their expression in murine duodenal villi, mRNA expression in isolated murine duodenal villi was determined by RT-PCR and compared against a tissue control with high expression levels of the particular transcript (the cecum for DRA and the kidney for PAT-1 and AE4) (21, 22, 26, 45, 47). As shown in Fig. 1, RT-PCR of the microdissected villous epithelium from the murine duodenum showed prominent expression of DRA and PAT-1 and lesser expression of AE4 compared with high-expressing tissue controls.


Figure 1
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  		Fig. 1. mRNA expression of downregulated by adenoma (DRA), putative anion transporter 1 (PAT-1), and anion exchanger isoform 4 (AE4) in microdissected murine duodenal villi. RT-PCR showed mRNA expression of DRA, PAT-1, and AE4 in the isolated villous epithelium (v) from wild-type (WT) duodena (representative of 3 experiments). Positive tissue controls were cDNA from the cecum (Ce) for DRA and the kidney (k) for PAT-1 and AE4. Water (w) and no reverse transcriptase RT(–) samples were negative controls. The absence of a band for beta-actin in the RT(–) control lanes indicates minimal contamination of the sample with genomic DNA.

 
Anion exchange in the AE4(–) upper villous epithelium of the murine duodenum. A functional study (48) in Xenopus oocytes has indicated that AE4 exhibits Cl/HCO3– exchange activity. Therefore, we compared anion exchange activity in the AE4(–) duodenal villous epithelium to that in the WT littermate epithelium. As shown in Fig. 2, A and B, there were no significant differences in the rates of HCO3– influx during Cl– removal (i.e., Cl–out/HCO3–in exchange) and HCO3– efflux during Cl– replacement (i.e., Cl–in/HCO3–out exchange) in the AE4(–) epithelium compared with the WT epithelium. These experiments did not detect significant differences in the baseline pHi of the upper villous AE4(–) epithelium compared with the WT epithelium (Fig. 2C). An anion substitution study (35) in murine duodenal villi has indicated the presence of apical membrane anion exchanger(s) that can readily perform SO42–in/HCO3–out exchange (35). Therefore, we examined this property in the AE4(–) epithelium compared with the WT epithelium. These experiments also did not demonstrate a significant difference in the rate of SO42–-dependent HCO3– efflux in the AE4(–) duodenum compared with the WT duodenum (Fig. 2D).


Figure 2
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  		Fig. 2. Anion exchange activity in the duodenal villous epithelium of WT and AE4(–) mice. A and B: summary of HCO3– influx (A) and HCO3– efflux (B) rates during luminal Cl– removal and replacement, respectively, in duodenal villous epithelial cells from WT and AE4(–) mice (n = 5). Cumulative data showed no significant differences between the rates of Cl–/HCO3– exchange in AE4(–) compared with WT mice. C: summary of baseline intracellular pH (pHi) in duodenal villous epithelial cells of WT and AE4(–) mice (n = 5). Cumulative data showed no significant differences between the baseline pHi in AE4(–) compared with WT mice. D: summary of SO42–-dependent HCO3– efflux rates in duodenal villous epithelial cells of WT and AE4(–) mice (n = 3). Cumulative data showed no significant differences between the rates of SO42–/HCO3– exchange in AE4(–) compared with WT mice.

 
Anion exchange in the DRA(–) upper villous epithelium of the murine duodenum. A previous study (18) has indicated that DRA is a major contributor to electroneutral HCO3– secretion in the duodenum of several species; therefore, we examined apical Cl–/HCO3– exchange in the DRA(–) upper duodenal villous epithelium. Cumulative data for several pHi experiments on DRA(–) and WT epithelia demonstrated that the rates of HCO3– influx during Cl– removal and HCO3– efflux during Cl– replacement were significantly reduced in DRA(–) villi relative to WT villi by 30–40% (Fig. 3, A and B). Interestingly, the baseline pHi was significantly increased in DRA(–) villi under the conditions of our study (Fig. 3C), which may reflect unopposed activity of apical NHE3 if DRA is coupled to NHE3 for electroneutral NaCl absorption in the upper villous epithelium.


Figure 3
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  		Fig. 3. Anion exchange activity in the duodenal villous epithelium of WT and DRA(–) mice. A. and B: summary of HCO3– influx (A) and HCO3– efflux (B) rates in duodenal villous epithelial cells of WT (n = 10) and DRA(–) (n = 7) mice. Cumulative data showed a moderate but significant reduction in the rates of Cl–/HCO3– exchange following luminal Cl– removal (HCO3– influx) and replacement (HCO3– efflux) in DRA(–) compared with WT mice. C: summary of baseline pHi in duodenal villous epithelial cells of WT (n = 10) and DRA(–) (n = 7) mice. Cumulative data showed significantly increased baseline pHi in DRA(–) compared with WT mice. D: summary of SO42–-dependent HCO3– efflux rates in duodenal villous epithelial cells of WT and DRA(–) mice (n = 3). Cumulative data showed no significant differences between the rates of SO42–/HCO3– exchange in DRA(–) compared with WT mice. E: representative pHi trace from murine (m)DRA-transfected CHO cells during extracellular Cl– removal and replacement followed by a second Cl– removal period and extracellular SO42– addition (representative of 4 experiments). Removal of extracellular Cl– resulted in intracellular alkalinization, and the readdition of Cl– led to pHi recovery, demonstrating Cl–/HCO3– exchange. However, SO42– replacement following alkalinization did not lead to appreciable pHi recovery. *Significantly different from WT.

 
The examination of apical membrane SO42–/HCO3– exchange between the DRA(–) epithelium compared with the WT epithelium revealed no significant differences in the rate of SO42–-dependent HCO3– efflux (Fig. 3D). These findings indicate that the degree of SO42–/HCO3– exchange attributable to mDRA may be minimal, as suggested previously (29). To investigate this possibility, we expressed recombinant mDRA in CHO fibroblasts.1 As shown by the representative experiment in Fig. 3E, CHO cells expressing mDRA alkalinized upon removal of extracellular Cl–, and pHi recovery occurred upon the readdition of extracellular Cl–, thus indicating robust Cl–/HCO3– exchange mediated by mDRA. In contrast, after a second period of pHi alkalinization induced by extracellular Cl– removal, the pHi did not significantly recover in the presence of extracellular SO42–, indicating that mDRA does not efficiently exchange SO42– for HCO3–.

Cl–/HCO3– exchange in the PAT-1(–) upper villous epithelium of the murine duodenum. Previous studies conducted in upper murine duodenal villous epithelia have revealed characteristics of basal anion exchange (e.g., high rates of SO42– transport and enhanced Cl–/HCO3– exchange activity during epithelial cell depolarization) that were most consistent with PAT-1 activity based on recombinant protein expression studies (19, 23, 35, 45, 47). To evaluate the contribution of PAT-1, we compared the rates of apical membrane Cl–/HCO3– exchange activity between PAT-1(–) and WT duodenal villi. As shown by the experiment in Fig. 4A, the rates of alkalinization and recovery during Cl– substitution/replacement were significantly reduced in the PAT-1(–) duodenum compared with the WT duodenum. Cumulative data from several pHi experiments on PAT-1(–) and WT epithelia demonstrated that the rates of HCO3– influx during Cl– removal and HCO3– efflux during Cl– replacement were significantly reduced by 65% and 80%, respectively, in PAT-1(–) villi relative to WT villi (Fig. 4, B and C). Paradoxically, the baseline pHi was significantly more acidic in PAT-1(–) villi relative to WT villi under the conditions of our study (Fig. 4D).


Figure 4
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  		Fig. 4. Comparison of Cl–/HCO3– exchange activity and baseline pHi in the duodenal villous epithelium of WT and PAT-1(–) mice. A: representative pHi traces of WT and PAT-1(–) duodenal villous epithelial cells during luminal Cl– removal and replacement. B and C: summary of HCO3– influx (B) and HCO3– efflux (C) rates in duodenal villous epithelial cells from WT and PAT-1(–) mice (n = 5). Cumulative data showed significantly reduced rates of Cl–/HCO3– exchange following luminal Cl– removal (HCO3– influx) and replacement (HCO3– efflux) in PAT-1(–) compared with WT mice. D: summary of baseline pHi in duodenal villous epithelial cells of WT and PAT-1(–) mice (n = 5). Cumulative data showed significantly reduced baseline pHi in PAT-1(–) compared with WT mice. *Significantly different from WT.

 
To assess the contribution of PAT-1 to SO42–/HCO3– exchange, we measured SO42–-dependent HCO3– transport rates in PAT-1(–) and WT duodenal villous epithelia. As shown by the experiment in Fig. 5A, after pHi alkalinization induced by luminal Cl– removal, luminal SO42– application resulted in pHi recovery in the WT villous epithelium. In contrast, the PAT-1(–) villous epithelium exhibited no discernable pHi recovery with SO42– application. Cumulative data for experiments on PAT-1(–) and WT epithelia showed that the rate of SO42–-dependent HCO3– efflux (Fig. 5B) was essentially abolished in the PAT-1(–) epithelium. Thus, PAT-1 is the major anion exchanger responsible for SO42– transport in the murine duodenal villous epithelium.


Figure 5
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  		Fig. 5. Comparison of SO42–/HCO3– exchange in the duodenal villous epithelium of WT and PAT-1(–) mice. A: representative pHi traces of WT and PAT-1(–) duodenal villous epithelial cells. Following alkalinization secondary to luminal Cl– removal, SO42– was introduced in the luminal superfusate for SO42–in/HCO3–out exchange. B: summary of HCO3– efflux rates induced by luminal SO42– application in duodenal epithelial cells from WT (n = 3) and PAT-1(–) (n = 5) mice. Cumulative data showed significantly reduced rates of SO42–/HCO3– exchange in PAT-1(–) compared with WT mice. *Significantly different from WT.

 
Investigation of the reduced pHi in the PAT-1(–) upper villous epithelium of the murine duodenum. As shown in Fig. 4, A and D, baseline pHi was significantly reduced in PAT-1(–) duodenal villous epithelial cells under the conditions of our study. This result was surprising given that PAT-1 is considered a cellular base exporter (44). We hypothesized that the reduced pHi may result from the loss of a coupled interaction between PAT-1 and NHE3, resulting in inhibition of NHE3 expression and/or activity. Therefore, NHE3 protein expression and activity were measured in WT and PAT-1(–) duodena. First, as shown in Fig. 6A, Western blot analysis did not indicate that NHE3 protein expression was different between PAT-1(–) and WT duodena. Second, apical membrane Na+/H+ exchange activity was measured as the Na+-dependent recovery of pHi after cellular acidification in WT and PAT-1(–) duodenal villi. These experiments revealed no significant differences in the rates of Na+/H+ exchange (i.e., H+ efflux) between the PAT-1(–) and WT duodenal villous epithelium (Fig. 6B), indicating that the acidic pHi in the PAT-1(–) epithelium is not due to loss of NHE3 activity. An alternative explanation for the acidic pHi is that PAT-1 may act as a HCO3–-loading process (Cl–out/HCO3–in) in the villous epithelium under the conditions of our study. To determine whether the process was HCO3– dependent, the baseline pHi was measured in the absence of CO2/HCO3– in the PAT-1(–) and WT epithelium. This maneuver essentially eliminated the difference in baseline pHi between the PAT-1(–) and WT villous epithelium, as shown in Fig. 7A. Since both a functional and physical relationship between cytosolic carbonic anhydrase II (CA II) and PAT-1 has been shown (1, 25), we also examined the effect of the CA inhibitor methazolamide (100 µM) on baseline pHi in PAT-1(–) and WT duodenal villous epithelial cells in the presence of CO2/HCO3–. As shown in Fig. 7B, methazolamide treatment eliminated the difference between the baseline pHi of the PAT-1(–) and WT villous epithelium. Since isolation of the apical membrane transporters in our study required the inhibition of basolateral transporters including Na+-K+-2Cl– cotransporter 1 (NKCC1) and the Cl–/HCO3– exchanger AE2 (with basolateral Cl–-free conditions) and the pHi-compensating effects of basolateral NHE1 (with 1 µM EIPA), we asked whether removal of this inhibition would normalize the pHi in the PAT-1(–) villous epithelium2 . Therefore, the activity of NHE1 was restored by removing basolateral EIPA, and pHi was measured. As shown in Fig. 7C, baseline pHi tended to increase but remained acidic in the PAT-1(–) epithelium compared with the WT epithelium. Finally, untreated duodenal mucosal preparations were superfused with physiological KBR solution bilaterally, thereby restoring Cl– to the basolateral transporters. These experiments revealed that under physiological conditions, pHi was similar between the PAT-1(–) and WT epithelium (Fig. 7D).


Figure 6
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  		Fig. 6. Comparison of Na+/H+ exchanger isoform 3 (NHE3) expression and apical membrane Na+/H+ exchange activity in WT and PAT-1(–) small intestines. A, top: representative Western blot of NHE3 and beta-actin protein (as a loading control) in duodenal microsomes of gender-matched WT and PAT-1(–) littermate pairs. Bottom, densiometric analysis of protein expression for NHE3 normalized to expression of beta-actin (n = 4 mouse pairs). These data showed no significant differences of NHE3 protein expression in PAT-1(–) compared with WT brush-border membrane vesicles. B: summary of Na+-dependent H+ efflux rates in duodenal villous cells of WT and PAT-1(–) mice (n = 3). Cumulative data showed no significant differences between the rates of Na+/H+ exchange in PAT-1(–) compared with WT duodenal villi.

 

Figure 7
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  		Fig. 7. Baseline pHi experiments in the duodenal villous epithelium of WT and PAT-1(–) mice. A: summary of baseline pHi in duodenal villous epithelial cells of WT and PAT-1(–) (n = 3) mice in the absence of CO2/HCO3–. Cumulative data showed no significant differences in baseline pHi of PAT-1(–) compared with WT mice. B: summary of baseline pHi in duodenal villous epithelial cells of WT and PAT-1(–) (n = 3) mice in the presence of bilateral methazolamide (100 µM) to inhibit carbonic anhydrase activity. Cumulative data showed no significant differences in baseline pHi of PAT-1(–) compared with WT mice. C: summary of baseline pHi in duodenal villous epithelial cells of WT and PAT-1(–) (n = 5) mice in the absence of serosal EIPA. Cumulative data showed significantly reduced baseline pHi in PAT-1(–) compared with WT mice. D: summary of baseline pHi in duodenal villous epithelial cells of WT and PAT-1(–) (n = 3) mice with bilateral Krebs-bicarbonate-Ringer (KBR) solution. Cumulative data showed no significant differences in baseline pHi of PAT-1(–) compared with WT mice under physiological conditions. *Significantly different from WT.

 

   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
The process of basal HCO3– secretion across the duodenum as measured by pH stat has been shown in several species to principally involve the activity of apically located Cl–/HCO3– exchanger(s) (10, 32, 37). Therefore, in an effort to resolve the identity of the anion exchangers involved in these processes, we examined anion exchange properties in epithelial cells located on the upper one-half of duodenal villi in murine knockout models with deletions of the apical membrane transporters PAT-1, DRA, and AE4 (18, 22, 45, 48). RT-PCR of isolated murine duodenal villi demonstrated the expression of PAT-1 and DRA and a low level of AE4 expression. Phenotypically, investigations of PAT-1(–) (44) and AE4(–) (L. R. Gawenis and G. E. Shull, unpublished observations) mice have revealed no overt disease and normal growth rates. In contrast, DRA(–) mice have chronic mild diarrhea (i.e., soft feces), a propensity to rectal prolapse, decreased body weights, and a mortality rate that exceeds that of PAT-1(–) and AE4(–) mice (6).

In the present investigation, BCECF microfluorometry experiments of the intact PAT-1(–) duodenum revealed that apical Cl–/HCO3– exchange was significantly reduced by 65–80% compared with the WT duodenum. This finding confirms the results of our previous study (35), which revealed characteristics of anion exchange in the upper villous epithelium that were most consistent with PAT-1 activity. In addition to PAT-1, a finite rate of Cl–/HCO3– exchange was still present in the PAT-1(–) duodenum, indicating the contribution of other anion exchangers such as DRA and AE4. The examination of AE4(–) mice compared with WT mice revealed minimal involvement of AE4 in basal Cl–/HCO3– exchange in the upper villous epithelium. In contrast, the examination of Cl–/HCO3– exchange activity in DRA(–) mice compared with WT mice demonstrated a 30–40% reduction in exchange rates. Thus, these data indicate that PAT-1 is the principal Cl–/HCO3– exchanger in the apical membrane of the upper villous epithelium with a smaller but significant contribution by DRA and minimal contribution from AE4.

The results of the functional experiments suggested that there was limited alteration in the apical surface activity of the remaining anion exchangers in each knockout mouse. In this regard, we noted a minor discrepancy in the estimates for the relative contribution of DRA to the total Cl–/HCO3– exchange activity in the murine upper villous epithelium. Taking the residual Cl–/HCO3– exchange activity in the PAT-1(–) duodenum as a measure of DRA activity, we estimate a minimum contribution of ~20% to total Cl–/HCO3– exchange activity (Fig. 4C). On the other hand, the reduction of Cl–/HCO3– exchange activity in the DRA(–) villous epithelium yields a maximal estimate of ~40% (Fig. 3B). This discrepancy suggests decreased expression/activity of the remaining anion exchanger in each knockout, e.g., DRA in the PAT-1(–) duodenum and PAT-1 in the DRA(–) duodenum. Thus, compensatory increases in anion exchanger activity are not apparent in the villous epithelium. Precise estimation of the contribution of DRA to the total Cl–/HCO3– exchanger activity will require measurements of anion exchanger expression specifically in the upper villous epithelium, e.g., by using laser capture microdissection or differential cell isolation techniques, which are beyond the scope of the present investigation.

The present investigation found that a distinguishing feature of PAT-1 is its ability to transport SO42–, because SO42–/HCO3– exchange was essentially eliminated in the PAT-1(–) villous epithelium compared with the WT villous epithelium. Both PAT-1 and DRA have been reported to be SO42– transporters (29); however, recent studies using recombinant protein expression systems have shown that human DRA does not efficiently transport SO42–, i.e., the rate of SO42– transport is two to three orders of magnitude less than the rate of Cl–/HCO3– exchange (8, 29). In the present study, SO42–/HCO3– exchange in DRA(–) duodenal villi was not significantly different from that observed in WT villi, indicating that mDRA also does not appreciably transport SO42–. To verify that mDRA, like human DRA, does not effectively transport SO42–, we measured SO42–/HCO3– exchange activity of recombinant mDRA expressed in CHO fibroblasts. These experiments revealed robust Cl–/HCO3– exchange but little evidence of SO42–/HCO3– exchange in mDRA-transfected cells. In addition, the examination of SO42– transport in the surface epithelium of the cecum, where DRA is the dominant apical membrane exchanger, also revealed no detectable SO42– transport (data not shown). Rates of SO42–/HCO3– exchange were also measured in the AE4(–) duodenal villous epithelium because studies examining AE4 exchange properties are limited (48). Like the experiments with the DRA(–) epithelium, these experiments revealed no significant differences in SO42–-dependent HCO3– efflux in the AE4(–) villous epithelium compared with the WT villous epithelium. Thus, these data show that a distinguishing characteristic of PAT-1 is the capability for SO42–/HCO3– exchange in the upper villous epithelium of the murine duodenum.

Measurements of pHi in the DRA(–) and PAT-1(–) villous epithelium revealed opposite effects of these transporters on pHi regulation. In the DRA(–) villous epithelium, baseline pHi was significantly increased compared with the WT epithelium. This change is consistent with the loss of a Cl–/HCO3– exchanger that is normally coupled to NHE3 for electroneutral NaCl absorption, i.e., continued proton efflux via NHE3 without concurrent HCO3– efflux would increase pHi. Although NHE3 may not normally contribute to pHi regulation in the intestine (14), the genetic ablation of a partnering anion exchanger possibly removes an element of normal regulation for NHE3. Additional studies of DRA(–) mice using the jejunum, where electroneutral NaCl absorption predominates, will be necessary to test this hypothesis. In contrast to the deletion of DRA, measurements of pHi in the PAT-1(–) villous epithelium revealed a significantly reduced baseline pHi compared with the WT epithelium. This phenomenon was investigated further because cell acidification resulting from deletion of a base exporter is counterintuitive. One potential explanation is that loss of a coupled interaction between PAT-1 and NHE3 reduces the activity or expression of the NHE. However, apical membrane Na+/H+ exchange activity and NHE3 protein expression were not different between PAT-1(–) and WT epithelia. An alternative hypothesis is that PAT-1 acts as a base importer (i.e., Cl–out/HCO3–in exchange) under the conditions of our study. Consistent with this hypothesis was the demonstration that removal of CO2/HCO3– or treatment with the CA inhibitor methazolamide normalized pHi between PAT-1(–) and WT epithelia. Based on evidence that PAT-1 may functionally and physically interact with CA II (1), the latter finding suggests a novel interaction between PAT-1 and intracellular CA, i.e., a "reversed" HCO3– transport metabolon (25, 39) that facilitates HCO3– influx to protect the epithelial cell against intracellular acidification. This may be of physiological importance as the upper villous epithelium of the duodenum is exposed to acid challenge not only from gastric effluent but also from H+ influx during nutrient absorption (e.g., apical H+/peptide cotransporter 1) (12). Given the high PCO2 in gastric effluent (17), the source of HCO3– for PAT-1 Cl–out/HCO3–in exchange may be dependent on extracellular CA activity (27). Although rapid diffusion of CO2 through cell membranes induces pHi acidification in the duodenal villous epithelium during luminal exposure to a high-PCO2/low-HCO3– solution (pH 6.4) (15), we found that a greater extent of pHi acidification occurs during exposure to high-PCO2/low-HCO3– solution (pH 5.4) in the PAT-1(–) duodenum compared with the WT duodenum (WT {Delta}pHi = –0.9 ± 0.1 and PAT-1(–) {Delta}pHi = –1.3 ± 0.2, n = 3; J. Simpson and L. Clarke, unpublished observations). The recent proposal that PAT-1 is electrogenic with a stoichiometry of 1 Cl–:2 HCO3– (33) would also favor the HCO3– influx activity of PAT-1 during depolarization of the villous epithelium, e.g., during Na+- or H+-coupled nutrient absorption. The resting apical membrane potential and intracellular Cl– concentrations of the upper villous epithelium under the conditions of our study are presently unknown, so whether the electrochemical gradient favors Cl–out/HCO3–in activity by PAT-1 cannot be determined at this time. However, as we (35) have shown previously, depolarization of the upper villous epithelium using high extracellular K+ concentration specifically increases the rate of Cl–out/HCO3–in exchange. Together, these considerations raise the possibility that PAT-1 plays a role in pHi regulation, especially during nutrient absorption, by providing a HCO3– influx pathway across the apical membrane of the villous enterocyte.

In our study, isolation of apical membrane acid-base transporters required inhibition of basolateral membrane NHE1 and Cl–/HCO3– exchange by EIPA treatment and removal of basolateral Cl–, respectively. We first investigated whether the activity of NHE1 could normalize basal pHi in the PAT-1(–) epithelium by removal of basolateral EIPA treatment. Although a trend toward increased pHi was noted, the PAT-1(–) villous epithelium remained acidic compared with the WT epithelium, indicating that activity of NHE1 alone is apparently unable to compensate for the loss of PAT-1. Next, a physiological Cl– concentration was included in the basolateral bath in addition to removal of basolateral EIPA treatment. This change corrected the pHi difference between PAT-1(–) and WT villous epithelia. Thus, under physiological conditions, pHi of the upper villous epithelium can be normalized in the absence of PAT-1 by a complex process apparently requiring the activity of basolateral transporters.

The activity of PAT-1 in the upper villous epithelium is likely affected by the presence of low levels of CFTR. In our previous study (35) of the CFTR(–) mouse intestine, it was shown that both Cl–in/HCO3–out and Cl–out/HCO3–in exchange rates were reduced in the upper villous epithelium compared with the WT mouse. The mechanism of decrease was traced to the loss of a CFTR Cl– leak pathway that facilitated apical membrane Cl–/HCO3– exchange activity. The role of CFTR was considered passive and not due to a direct intermolecular interaction with the anion exchanger(s) (24) because CFTR is expressed at low levels in the upper villous epithelium (thereby minimizing the opportunity for 1:1 stoichiometry between CFTR and a Slc26a exchanger) and the effect could be reproduced by acute CFTR channel blockade. Given the dominance of PAT-1 activity (Fig. 4), it is reasonable to conclude that the CFTR Cl– leak pathway primarily facilitates PAT-1 and perhaps lesser amounts of DRA activity in the upper villous epithelium. Since CFTR demonstrates ohmic conductance of Cl– (2), it is reasonable that the Cl– leak pathway may operate to the advantage of PAT-1 Cl–/HCO3– exchange activity independent of the direction of transport, i.e., providing Cl– uptake during Cl–out/HCO3–in exchange and Cl– exit during Cl–in/HCO3–out exchange.

Although we (31) have previously suggested that the reduced Cl–/HCO3– exchange activity of the upper villous epithelium contributes to deficient basal HCO3– secretion across the CFTR(–) duodenum, a subsequent pH stat study (42) of PAT-1(–) mice showed that PAT-1 only contributes ~20% to basal HCO3– secretion in the WT duodenum. Thus, a 30% reduction in PAT-1 activity in the upper villous epithelium of the CFTR(–) duodenum would have a negligible effect on basal HCO3– secretion. Apparently, most transepithelial HCO3– secretion under basal conditions comes from the lower villous and crypt epithelium where the activities of DRA or AE4 may provide a major contribution. Given evidence that PAT-1 plays a minor role in transepithelial HCO3– secretion under basal conditions, it is possible that the abundant PAT-1 activity in the upper villous epithelium serves other functions, such as pHi regulation, as suggested above.

In conclusion, our study of PAT-1(–), DRA(–), and AE4(–) mice indicated that PAT-1 is the major contributor to basal Cl–/HCO3– and SO42–/HCO3– exchange across the apical membrane of the upper villous epithelium in the murine duodenum. Based on past studies of CFTR activity in this region of the villus, it is likely that PAT-1 activity is facilitated indirectly by a CFTR Cl– leak pathway that sustains the gradient for Cl–/HCO3– exchange. However, additional experiments in the PAT-1(–) duodenum revealed a propensity for PAT-1 to operate in Cl–out/HCO3–in mode for HCO3– influx, thereby providing a potentially important mechanism for protecting the upper villous epithelial cell from intracellular acidification. DRA was also found to contribute a smaller but significant portion of Cl–/HCO3– exchange in the upper villous epithelium. DRA deletion resulted in intracellular alkalinization, which may indicate unregulated activity of Na+/H+ exchange and evidence of DRA's involvement in coupled NaCl absorption in this locale. Therefore, DRA may not significantly contribute to net HCO3– secretion under basal conditions from the upper villous epithelium, whereas a recent report (36) from our laboratory suggested that DRA may have a different role in the lower villus epithelium. Thus, it will be necessary to examine anion transport in the lower villus/crypt epithelium where CFTR is predominantly expressed to fully evaluate the contributions of AE4, DRA, and PAT-1 to transepithelial HCO3– secretion.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by National Institutes of Health Grants T32-RR-07004 (to J. E. Simpson), DK-48816 (to L. L. Clarke), DK-50594 (to G. E. Shull), CA-95172 (to C.W. Schweinfest), and DK-62809 (to M. Soleimani) and by Cystic Fibrosis Foundation Grant CLARKE05G0 (to L. L. Clarke).


   FOOTNOTES
 

Address for reprint requests and other correspondence: L. L. Clarke, 324D Dalton Cardiovascular Research Center, 134 Research Park Dr., Univ. of Missouri, Columbia, MO 65211 (e-mail: clarkel{at}missouri.edu)

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.

1 No endogenous Cl–/HCO3– exchange was observed in mock-transfected CHO cells, whereas inconsistent levels of activity were found when mock-transfected NIH 3T3 and HEK-293 cells were examined. Back

2 Pharmacological inhibition of NHE1 by 1 µM EIPA added to the basolateral superfusate during a similar time period, i.e., 20 min, has been previously demonstrated in this preparation by a study (43) showing significant acidification of the pHi in the upper villous epithelium of the NHE3(–) duodenum during 1 µM EIPA treatment. Back


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Alvarez BV, Vilas GL, Casey JR. Metabolon disruption: a mechanism that regulates bicarbonate transport. EMBO J 24: 2499–2511, 2005.[CrossRef][ISI][Medline]
  2. Anderson MP, Sheppard DN, Berger HA, Welsh MJ. Chloride channels in the apical membrane of normal and cystic fibrosis airway and intestinal epithelia. Am J Physiol Lung Cell Mol Physiol 263: L1–L14, 1992.[Abstract/Free Full Text]
  3. Barone S, Amlal H, Xu J, Kujala M, Kere J, Petrovic S, Soleimani M. Differential regulation of basolateral Cl–/HCO3– exchangers SLC26A7 and AE1 in kidney outer medullary collecting duct. J Am Soc Nephrol 15: 2002–2011, 2004.[Abstract/Free Full Text]
  4. Boyarsky G, Ganz MB, Sterzel RB, Boron WF. pH regulation in single glomerular mesangial cells. I. Acid extrusion in absence and presence of HCO3–. Am J Physiol Cell Physiol 255: C844–C856, 1988.[Abstract/Free Full Text]
  5. Bukhave K, Rask-Madsen J. Saturation kinetics applied to in vitro effects of low prostaglandin E2 and F2{alpha} concentrations on ion transport across human jejunal mucosa. Gastroenterology 78: 32–42, 1980.[ISI][Medline]
  6. Chapman JM, Kim JH, Henderson KW, Spyropoulos DD, Schweinfest CW. DRA, an intestinal anion transporter, suppresses cell growth and proliferation. Proc Am Assoc Cancer Res 43: 990, 2003.
  7. Chernova MN, Jiang L, Friedman DJ, Darman RB, Lohi H, Kere J, Vandorpe DH, Alper SL. Functional comparison of mouse slc26a6 anion exchanger with human SLC26A6 polypeptide variants: differences in anion selectivity, regulation, and electrogenicity. J Biol Chem 280: 8564–8580, 2005.[Abstract/Free Full Text]
  8. Chernova MN, Jiang L, Shmukler BE, Schweinfest CW, Blanco P, Freedman SD, Stewart AK, Alper SL. Acute regulation of the SLC26A3 congenital chloride diarrhoea anion exchanger (DRA) expressed in Xenopus oocytes. J Physiol 549.1: 3–19, 2003.
  9. Clarke LL, Harline MC. CFTR is required for cAMP inhibition of intestinal Na+ absorption in a cystic fibrosis mouse model. Am J Physiol Gastrointest Liver Physiol 270: G259–G267, 1996.[Abstract/Free Full Text]
  10. Clarke LL, Harline MC. Dual role of CFTR in cAMP-stimulated HCO3– secretion across murine duodenum. Am J Physiol Gastrointest Liver Physiol 274: G718–G726, 1998.[Abstract/Free Full Text]
  11. Clarke LL, Stien X, Walker NM. Intestinal bicarbonate secretion in cystic fibrosis mice. J Pancreas 2, Suppl 4: 263–267, 2001.
  12. Daniel H. Molecular and integrative physiology of intestinal peptide transport. Annu Rev Physiol 66: 361–384, 2004.[CrossRef][ISI][Medline]
  13. Flemstrom G. Gastric and duodenal mucosal bicarbonate secretion. In: Physiology of the Gastrointestinal Tract, edited by Johnson LR. New York: Raven, 1987, p. 1011–1030.
  14. Furukawa O, Bi LC, Guth PH, Engel E, Hirokawa M, Kaunitz JD. NHE3 inhibition activates duodenal bicarbonate secretion in the rat. Am J Physiol Gastrointest Liver Physiol 286: G102–G109, 2004.[Abstract/Free Full Text]
  15. Furukawa O, Hirokawa M, Zhang L, Takeuchi T, Bi LC, Guth PH, Engel E, Akiba Y, Kaunitz JD. Mechanism of augmented duodenal HCO3– secretion after elevation of luminal CO2. Am J Physiol Gastrointest Liver Physiol 288: G557–G563, 2005.[Abstract/Free Full Text]
  16. Gawenis LR, Franklin CL, Simpson JE, Palmer BA, Walker NM, Wiggins TM, Clarke LL. cAMP inhibition of murine intestinal Na+/H+ exchange requires CFTR-mediated cell shrinkage of villus epithelium. Gastroenterology 125: 1148–1163, 2003.[CrossRef][ISI]
  17. Holm M, Johansson B, Pettersson A, Fandriks L. Carbon dioxide mediates duodenal mucosal alkaline secretion in response to luminal acidity in the anesthetized rat. Gastroenterology 115: 680–685, 1998.[CrossRef][ISI][Medline]
  18. Jacob P, Rossmann H, Lamprecht G, Kretz A, Neff C, Lin-Wu E, Gregor M, Groneberg DA, Kere J, Seidler U. Down-regulated in adenoma mediates apical Cl–/HCO3– exchange in rabbit, rat, and human duodenum. Gastroenterology 122: 709–724, 2002.[CrossRef][ISI][Medline]
  19. Jiang Z, Grichtchenko II, Boron WF, Aronson PS. Specificity of anion exchange mediated by mouse Slc26a6. J Biol Chem 277: 33963–33967, 2002.[Abstract/Free Full Text]
  20. Kere J, Lohi H, Hoglund P. Genetic disorders of membrane transport. III. Congenital chloride diarrhea. Am J Physiol Gastrointest Liver Physiol 276: G7–G13, 1999.[Abstract/Free Full Text]
  21. Knauf F, Yang CL, Thomson RB, Mentone SA, Giebisch G, Aronson PS. Identification of a chloride-formate exchanger expressed on the brush border membrane of renal proximal tubule cells. Proc Natl Acad Sci USA 98: 9425–9430, 2001.[Abstract/Free Full Text]
  22. Ko SBH, Luo X, Hager H, Rojek A, Choi JY, Licht C, Suzuki M, Muallem S, Nielsen S, Ishibashi K. AE4 is a DIDS-sensitive Cl–/HCO3– exchanger in the basolateral membrane of the renal CCD and SMG duct. Am J Physiol Cell Physiol 283: C1206–C1218, 2002.[Abstract/Free Full Text]
  23. Ko SBH, Shcheynikov N, Choi JY, Luo X, Oshibashi K, Thomas PJ, Kim JY, Kim KH, Lee MG, Naruse S, Muallem S. A molecular mechanism for aberrant CFTR-dependent HCO3– transport in cystic fibrosis. EMBO J 21: 5662–5672, 2002.[CrossRef][ISI][Medline]
  24. Ko SBH, Zeng W, Dorwart MR, Luo X, Kim KH, Millen L, Goto H, Naruse S, Soyombo A, Thomas PJ, Muallem S. Gating of CFTR by the STAS domain of SLC26 transporters. Nat Cell Biol 6: 343–350, 2004.[CrossRef][ISI][Medline]
  25. McMurtrie HL, Cleary HJ, Alvarez BV, Loiselle FB, Sterling D, Morgan PE, Johnson DE, Casey JR. The bicarbonate transport metabolon. J Enzyme Inhib Med Chem 19: 231–236, 2004.[CrossRef][ISI][Medline]
  26. Melvin JE, Park K, Richardson L, Schultheis P, Shull GE. Mouse down-regulated in adenoma (DRA) is an intestinal Cl–/HCO3– exchanger and is up-regulated in colon of mice lacking the NHE3 Na+/H+ exchanger. J Biol Chem 274: 22855–22861, 1999.[Abstract/Free Full Text]
  27. Mizumori M, Meyerowitz J, Takeuchi T, Lim S, Lee P, Supuran CT, Guth PH, Engel E, Kaunitz JD, Akiba Y. Epithelial carbonic anhydrases facilitate PCO2 and pH regulation in rat duodenal mucosa. J Physiol 573.3: 827–842, 2006.
  28. Moseley RH, Hoglund P, Wu GD, Silberg DG, Haila S, de la Chapelle A, Holmberg C, Kere J. Downregulated in adenoma gene encodes a chloride transporter defective in congenital chloride diarrhea. Am J Physiol Gastrointest Liver Physiol 276: G185–G192, 1999.[Abstract/Free Full Text]
  29. Mount DB, Romero MF. The SLC26 gene family of multifunctional anion exchangers. Pflügers Arch 447: 710–721, 2004.[CrossRef][ISI][Medline]
  30. Musch MW, Arvans DL, Field M, Wu GD, Chang EB. DRA is apical intestinal anion exchanger functionally coupled to NHE2 and NHE3 to mediate electroneutral NaCl absorption (Abstract). Gastroenterology 124: A40, 2003.[CrossRef]
  31. Pratha V, Hogan DL, Martensson B, Isenberg JI. Cystic fibrosis (CF) patients have impaired duodenal mucosal transport. Gastroenterology 114: G1655, 1998.
  32. Pratha VS, Hogan DL, Martensson BA, Bernard J, Zhou R, Isenberg JI. Identification of transport abnormalities in duodenal mucosa and duodenal enterocytes from patients with cystic fibrosis. Gastroenterology 118: 1051–1060, 2000.[CrossRef][ISI][Medline]
  33. Shcheynikov N, Wang Y, Park M, Ko SB, Dorwart M, Naruse S, Thomas PJ, Muallem S. Coupling modes and stoichiometry of Cl–/HCO3– exchange by slc26a3 and slc26a6. J Gen Physiol 127: 511–524, 2006.[Abstract/Free Full Text]
  34. Sheldon RJ, Malarchik ME, Fox DA, Burks TF, Porreca F. Pharmacological characterization of neural mechanisms regulating mucosal ion transport in mouse jejunum. J Pharmacol Exp Ther 249: 572–582, 1988.[ISI]
  35. Simpson JE, Gawenis LR, Walker NM, Boyle KT, Clarke LL. Chloride conductance of CFTR facilitates Cl–/HCO3– exchange in the villous epithelium of intact murine duodenum. Am J Physiol Gastrointest Liver Physiol 288: G1241–G1251, 2005.[Abstract/Free Full Text]
  36. Simpson JE, Walker NM, Schweinfest CW, Clarke LL. Down-regulated in adenoma (DRA, Slc26a3) is the predominant Cl–/HCO3– exchanger in the lower villous epithelium of murine duodenum (Abstract). Gastroenterology 130: A375, 2006.
  37. Spiegel S, Phillipper M, Rossmann H, Riederer B, Gregor M, Seidler U. Independence of apical Cl–/HCO3– exchange and anion conductance in duodenal HCO3– secretion. Am J Physiol Gastrointest Liver Physiol 285: G887–G897, 2003.[Abstract/Free Full Text]
  38. Sterling D, Brown NJ, Supuran CT, Casey JR. The functional and physical relationship between DRA bicarbonate transporter and carbonic anhydrase II. Am J Physiol Cell Physiol 283: C1522–C1529, 2002.[Abstract/Free Full Text]
  39. Sterling D, Reithmeier RAF, Casey JR. A transport metabolon. Functional interaction of carbonic anhydrase II and chloride/bicarbonate exchangers. J Biol Chem 276: 47886–47894, 2001.[Abstract/Free Full Text]
  40. Thomas JA, Buchsbaum RN, Zimniak A, Racker E. Intracellular pH measurements in ehrlich ascites tumor cells utilizing spectroscopic probes generated in situ. Biochemistry 18: 2210–2218, 1979.[CrossRef][Medline]
  41. Thomson RB, Wang T, Thomson BR, Tarrats L, Girardi A, Mentone S, Soleimani M, Kocher O, Aronson PS. Role of PDZK1 in membrane expression of renal brush border ion exchangers. Proc Natl Acad Sci USA 102: 13331–13336, 2005.[Abstract/Free Full Text]
  42. Tuo B, Riederer B, Wang Z, Colledge WH, Soleimani M, Seidler U. Involvement of the anion exchanger Slc26a6 in PGE2- but not forskolin-stimulated murine duodenal HCO3 secretion. Gastroenterology 130: 349–358, 2006.[CrossRef][ISI][Medline]
  43. Walker NM, Simpson JE, Boyle KT, Clarke LL. Increased electrogenic bicarbonate secretion and uncoupled chloride/bicarbonate exchange activity are responsible for high rates of transepithelial bicarbonate secretion in the NHE3 knockout duodenum (Abstract). Gastroenterology 128: A365, 2005.
  44. Wang Z, Wang T, Petrovic S, Tuo B, Riederer B, Barone S, Lorenz JN, Seidler U, Aronson PS, Soleimani M. Renal and intestine transport defects in Slc26a6-null mice. Am J Physiol Cell Physiol 288: C957–C965, 2005.[Abstract/Free Full Text]
  45. Wang ZH, Petrovic S, Mann E, Soleimani M. Identification of an apical Cl–/HCO3– exchanger in the small intestine. Am J Physiol Gastrointest Liver Physiol 282: G573–G579, 2002.[Abstract/Free Full Text]
  46. Weintraub WH, Machen TE. pH regulation in hepatoma cells: roles for Na-H exchange, Cl-HCO3 exchange, and Na-HCO3 cotransport. Am J Physiol Gastrointest Liver Physiol 257: G317–G327, 1989.[Abstract/Free&nbs