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MUCOSAL BIOLOGY
1Department of Pathology, Immunology, and Laboratory Medicine, College of Medicine, University of Florida, Gainesville, Florida 2Department of Medicine, University of Cincinnati, Cincinnati, Ohio
Submitted 11 October 2005 ; accepted in final form 13 December 2005
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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), whereas in KO mice JnetOx was significantly absorptive (75 ± 10 pmol·cm–2h·h–1), which was the result of a smaller serosal-to-mucosal oxalate flux (JsmOx) and a larger mucosal-to-serosal oxalate flux (JmsOx). Mucosal DIDS (200 µM) reduced JsmOx in WT mice, leading to reversal of the direction of net oxalate transport from secretion to absorption (
) , but DIDS had no significant effect on KO ileum. In WT mice in the absence of mucosal Cl–, there were small increases in JmsOx and decreases in JsmOx that led to a small net oxalate absorption. In KO mice, JnetOx was 1.5-fold greater in the absence of mucosal Cl–, due solely to an increase in JmsOx. Urinary oxalate excretion was about fourfold greater in KO mice compared with WT littermates. We conclude that PAT1 is DIDS sensitive and mediates a significant fraction of oxalate efflux across the apical membrane in exchange for Cl–; as such, PAT1 represents a major apical membrane pathway mediating JsmOx.
putative anion transporter 1; hyperoxaluria; anion exchange; serum oxalate; 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid
The molecular identification of the individual pathways involved in transepithelial oxalate transport has become more promising with the recent characterization of a gene family (Slc26) encoding anion exchange proteins that accept a variety of monovalent and divalent substrates (4, 17, 27, 35). At least one-half of the 10 functional genes in this family have the ability to transport oxalate when functionally characterized in heterologous expression systems (1, 24, 27, 30, 31, 39), and several of these oxalate transporters are present in the intestine (1, 5, 8, 17, 18, 21, 27, 28, 30–32, 35, 36, 38). For example, Slc26a3 (DRA) and Slc26a6 [putative anion transporter (PAT1)] are expressed along the length of the alimentary system, with DRA being more abundant in apical membranes of large intestine (3, 16, 26, 35, 36) and PAT1 being localized to the apical membranes of villar enterocytes in the duodenum (35, 36, 38) and in acinar cells of the pancreas (23). Because PAT1 is relatively abundant in small intestinal enterocytes, it is an attractive candidate mediator for transepithelial oxalate transport in this intestinal segment, but this proposal has not been tested directly.
To establish the role of PAT1 in intestinal oxalate transport, we have compared ileal oxalate transport and urinary oxalate excretion in wild-type (WT) and Slc26a6 null [knockout (KO)] mice developed by targeted gene disruption (36). We report here that PAT1 is, indeed, an important component of vectorial oxalate transport in the mouse distal ileum as judged by several criteria. First, whereas WT mice show a small net ileal secretion of oxalate, tissues from PAT1 null mice exhibit large absorptive net oxalate fluxes. Second, 200 µM mucosal DIDS promoted net oxalate absorption in WT ileum by a reduction in the serosal-to-mucosal flux of oxalate but was without effect in the KO mouse ileum. Third, Cl– removal from the mucosal media increased the net absorption of oxalate in both groups, but in PAT1 null mice net ileal oxalate absorption was two times that measured in the presence of luminal Cl–. Fourth, urinary oxalate excretion in PAT1 null mice was fourfold that of the WT. Based on these results, we propose that the PAT1 gene product in the mouse ileum mediates apical efflux of oxalate in exchange for Cl– and is therefore a significant component of the transcellular serosal-to-mucosal unidirectional oxalate flux.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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Transepithelial transport measurements.
The unidirectional and net fluxes of oxalate across isolated ileal segments were measured essentially as described previously (10, 12, 13, 15). Immediately after death and exsanguination of the mice, 
4 cm of the distal ileum was removed and thoroughly cleansed by flushing with ice-cold 0.9% NaCl. With the use of a dissecting microscope, remnant tags of connective tissue were removed, and the intestinal segment was opened along the mesenteric border. Flat sheets of tissue were mounted in modified Ussing chambers having an exposed tissue area of 0.34 cm2. Unidirectional fluxes of [14C]oxalate were measured across tissues bathed on both sides by 4 ml of buffered saline (pH 7.4) at 37°C circulated by vigorously bubbling with 95% O2-5% CO2. The total oxalate concentration of mucosal and serosal buffers was 1.25 µM. The standard saline contained the following solutes (mmol/l): 139.4 Na+, 5.4 K+, 1.2 Mg2+, 123.2 Cl–, 21.0 HCO3–, 1.2 Ca2+, 0.6 H2PO4–, 2.4 HPO42–, and 10 glucose for the serosal buffer or 10 mannitol for mucosal buffer. Addition of 10 mM glucose to the mucosal side at the end of the experiments initiates electrogenic, Na+-dependent glucose absorption, which provides an additional comparative measure of tissue integrity. This standard buffer also contained 5 µM indomethacin to inhibit prostanoid production during flux measurements. Cl–-free buffers were prepared by substituting gluconate salts for Cl– salts, and calcium in this buffer was elevated to 5.0 mM to offset chelation of calcium by gluconate. The magnitude and direction of the net oxalate flux (JnetOx) was determined by calculating the difference between the two measured unidirectional fluxes [mucosal to serosal (JmsOx) and serosal to mucosal (JsmOx)] at 15-min intervals for up to 105 min under short-circuit conditions using an automatic voltage clamp (VCCMC6; Physiologic Instruments, San Diego, CA). Typically, we have divided this total measurement time into two periods: an initial period (Per I) representing the average of the first three 15-min flux intervals (0–45 min) and a second period (Per II) representing the average of the last three 15-min flux intervals (60–105 min). The electrical parameters of the tissue were also recorded at 15-min intervals throughout the entire experiment. Tissue conductance (GT; mS/cm2) was calculated as the ratio of the open-circuit potential (VT; mV) to the short-circuit current (Isc; µA/cm2) and net fluxes were computed on conductance-matched tissues pairs (GT within 15% of one another).
Urine collection. Mice were housed in metabolic cages and had free access to food and water; 24-h urine collections were made under mineral oil in vessels containing 10 µl of 2% sodium azide as a preservative (for pH, osmolarity, and Cl–) or in vessels containing 100 µl of 3.5 N HCl to ensure complete solubilization of crystallized oxalates.
Analytical methods. Urinary Cl– concentrations were determined with a chloridometer (Labcono, Kansas City, MO). Urine osmolality was measured with a freezing point osmometer (Fiske Associates, Norwood, MA) and urine pH with a pH electrode (Accumet; Fisher Scientific, Philadelphia, PA). Creatinine was determined in the urine and serum samples using a modification of the Jaffé reaction as described previously (7). Oxalate was measured in both serum and urine using a coupled enzymatic (oxalate decarboxylase and formate dehydrogenase) assay procedure routine in our laboratory (9). Mouse blood collected by cardiac puncture was handled immediately with the appropriate precautions to prevent oxalogenesis, and serum pools from two to five mice were prepared for oxalate determination.
Fecal oxalobacter.
Fecal samples were collected from mouse large intestine at the time tissue was being prepared for the flux studies. The presence of Oxalobacter sp. in the feces was determined by inoculating anerobically sealed vials, containing 20 mM oxalate, with 20 mg of fecal material. The potential loss of oxalate by microbial action in these vials at 37°C was determined 1 wk later by our routine enzymatic oxalate assay.
RNA isolation and real-time PCR.
A 50- to 100-mg segment of the distal ileum from each mouse was preserved in RNAlater (Ambion, Austin, TX), and total cellular RNA was isolated with TRIzol (Invitrogen, Carlsbad, CA) according to the manufacturer’s recommendations. Residual genomic DNA contamination was removed from the total RNA samples with the TURBO DNA-free kit (Ambion). Gene-specific oligonucleotide primers were designed with Primer3 (29) from murine reference nucleotide sequences retrieved from GenBank and are shown in Table 1. Real-time PCR was performed using the QuantiTect SYBR Green RT-PCR kit (Qiagen, Valencia, CA) with the DNA Engine Opticon continuous fluorescence detection system (MJ Research, San Francisco, CA). Briefly, 100 ng of total RNA was added to 25 µl of 2X QuantiTect SYBR Green RT-PCR Master Mix, 250 ng of each gene-specific primer, and 0.5 µl of QuaniTect RT mix. The reactions were adjusted to 50 µl total volume with RNase-free water and incubated at 50°C for 30 min. RT was deactivated, and the HotStarTaq DNA polymerase was activated by incubation at 95°C for 15 min. Reactions were then subjected to 45 cycles of denaturation at 94°C for 15 s, annealing at 55°C for 30 s, and extension at 72°C for 30 s. Fluorescence data were collected after each extension step. After each PCR run, a melting-curve analysis was performed using the built-in software of the DNA Engine Opticon System to verify specificity of the RT-PCR product. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as an internal reference to normalize for amount of RNA added to the reverse transcription reaction. Data are presented as the difference in critical threshold cycles between the gene of interest and that of a reference gene [
C(T)] as previously described (22).
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Other solutions and chemical resources. [14C]oxalate (4.14 GBq/mmol) was purchased from Amersham (Piscataway, NJ). Concentrated stock solutions of 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid (DIDS; Molecular Probes, Eugene, OR) were prepared daily in DMSO. The final concentration of DMSO in the mucosal chamber after DIDS addition was <0.1%. Dibutyryl-cAMP (dB-cAMP) and all other reagent-grade salts were purchased from Sigma-Aldrich (St. Louis, MO).
Statistical analyses.
Comparisons between WT and KO mice were made using Student’s t-test (2-tailed, unpaired), whereas comparisons between Per I and Per II variates within a given experimental design were made by a paired t-test. Results are presented as means ± 1 SE, and differences between means were judged significant if P < 0.050. To compare differences in gene expression levels, C(T) values were rank transformed before ANOVA.
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RESULTS |
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Stilbene sensitivity of ileal oxalate transport. Oxalate transport pathways have long been known to be sensitive to disulfonic stilbenes (11, 19); consequently, we evaluated the effects of 200 µM DIDS, added to the mucosal compartment, on the unidirectional and net fluxes of oxalate across WT and KO mouse ileum. Mucosal DIDS had no significant effect on Isc or GT in either animal groups (data not shown). In WT mouse ileum, mucosal DIDS produced a reversal of JnetOx(from –11.7 ± 3.6 to 17.6 ± 9.1 pmol·cm–2·h–1) by a significant reduction (33%) in JsmOx and an increase in JmsOx that was not statistically significant (Fig. 2A). In contrast to these observations on WT mouse ileum, mucosal DIDS had no effect on the unidirectional or net fluxes of oxalate in the KO mouse ileum (Fig. 2B).
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Cl– dependence of ileal oxalate transport. Because the previous experiments suggested that PAT1 mediates oxalate efflux in the mouse ileum, we evaluated the possibility that mucosal Cl– is a countertransport partner for PAT1-mediated oxalate efflux by removing mucosal Cl–. This maneuver generates bi-ionic diffusion potentials across the epithelium that contribute to the transepithelial potentials that must be clamped to establish the short-circuiting condition (VT = 0). Consequently, the electrical parameters measured in both WT (Fig. 3, B and C) and KO (Fig. 4, B and C) mice during Per I (0 mucosal Cl–) are not a measure of active, electrogenic ion transport in the absence of mucosal Cl–. However, the unidirectional flux measurements presented in these figures can be compared with other flux measurements, since in all experiments the electrical driving forces are nominally clamped at VT = 0 mV.
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In the KO mouse ileum, Cl–-free media produced changes in the electrical characteristics (Fig. 4, B and C) of the tissues that were similar to that observed in WT ileum, and the same constraints apply to interpretations of the significance of these observations. Oxalate fluxes measured on adjacent pieces of ileum from a KO mouse are shown in Fig. 4A in the absence and presence of Cl–. Unlike the WT ileum, where mucosal Cl– altered both unidirectional oxalate fluxes, readmission of mucosal Cl– only changed JmsOx, reducing it by 34% (Fig. 3D). The net effect was a significant reduction in JnetOx(46%; Fig. 4F). As with WT mouse ileum, the unidirectional and net fluxes observed in Per II are similar to those presented previously for KO tissues, suggesting that these tissues remain viable throughout this experimental protocol.
Based on these findings, it is clear that mucosal Cl– is important in mediating apical membrane oxalate exchange in both WT and KO mouse ileum. In WT tissues, the presence of mucosal Cl– depressed JmsOx, which may reflect a competition between Cl– and oxalate for an exchanger at the luminal side of the apical membrane. The fact that in WT mucosal Cl– significantly increases JsmOx suggests that Cl– is an exchange partner for oxalate efflux across the apical membrane (i.e., an apical component of JsmOx). If the PAT1 exchanger represents this oxalate efflux pathway, as suggested in the DIDS experiments, then in KO mouse ileum lacking PAT1 JsmOx should be unaffected by restoration of mucosal Cl–, an expectation that was observed experimentally.
Effects of cAMP. Addition of dB-cAMP (0.5 mM to both sides) at the end of Per I significantly increased Isc in Per II for both WT (n = 6, from 0.68 ± 0. 21 to 5.46 ± 0.42 µeq·cm–2·h–1) and KO (n = 6, from 0.87 ± 0.16 to 6.60 ± 0.48 µeq·cm–2·h–1) mice. cAMP also produced a significant 23% reduction of JmsOx(from 89.7 ± 8.1 to 70.1 ± 6.0 pmol·cm–2·h–1) in KO ileum which led to a significant fall in JnetOx(from 58.8 ± 11.9 to 35.9 ± 11.1 pmol·cm–2·h–1). In WT mice, although dB-cAMP did tend to reduce JmsOx 16%, neither this, nor the other fluxes, was statistically different from Per I values. The observation that JsmOx was not increased in either WT or KO mice suggests that PAT1-mediated oxalate efflux is not regulated by increases in cellular cAMP in mouse ileum.
Ileal PAT1 protein and mRNA. The absence of PAT1 protein and mRNA in Slc26a6 null mice was verified by immunoblot and real-time RT-PCR, respectively. As shown in Fig. 5, PAT1 protein was not detected in ileal luminal membrane homogenates from Slc26a6–/– (KO) mice at 25 or 50 µg total protein but abundantly expressed in Slc26a6+/+ (WT) mice at both protein concentrations. Furthermore, we were unable to detect PAT1 mRNA in KO mouse ileum but did find it in the WT mice. These findings are in agreement with immunofluorescent labeling and mRNA expression patterns of PAT1 in the duodena of KO and WT mice reported earlier (36).
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C(T) is an inverse logarithmic expression of the abundance of a given mRNA; the higher the C(T) the lower the quantity of a given mRNA. As shown in Table 2, there were no significant differences in the mRNA expression profiles between WT and KO mouse ileum for DRA, SAT1, DTDST, or CFTR as determined by real-time PCR, suggesting that these genes are not differentially expressed in the KO mouse ileum. It should be noted that, although comparisons between WT and KO mice for a given gene are valid, comparisons of relative mRNA abundance of different mRNAs is not necessarily suitable because PCR reaction efficiencies for different genes may not be identical. Consequently, we do not consider the differences in C(T) among different genes as a meaningful measure of their relative abundance.
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Enteric oxalate-degrading bacteria. We also tested for the presence of oxalate-degrading bacteria in the large bowel of WT and KO mice, since a relative loss of these organisms in KO mice could conceivably increase the enteric oxalate burden and contribute to the observed hyperoxaluria in KO mice. However, no evidence was found for the presence of oxalate-degrading bacteria in either the WT or KO mouse large intestine.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONGRANTSREFERENCES |
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PAT1 is expressed, in varying degrees, throughout the mouse gastrointestinal system, yet it is particularly abundant in the small intestine, a distribution that is the opposite of that of DRA (Slc26a3; see Ref. 35). Immunohistochemical staining of PAT1 in mouse duodenum labels the apical aspect of the villar, but not crypt, epithelium (35). In the duodenum, PAT1 functions as a apical Cl–/HCO3– exchanger mediating basal HCO3– secretion, as evidenced by reduced baseline HCO3– secretion in PAT1 null mouse duodenum and reduced Cl–/HCO3– exchange in duodenal brush-border membrane vesicles from KO mice (in the presence of an outward pH and HCO3– gradient) compared with WT mice (36). Slc26a6 has been shown to have an affinity for a variety of anions, including Cl–, oxalate, formate, sulfate, and HCO3–, and can function in a number of exchange modes involving different pairs of these ions (17, 35). In mouse duodenum, the Cl–/HCO3– exchange mode is important to HCO3– secretion, and it also appears, based on the present results, that PAT1 mediates Cl–/oxalate exchange in the ileum.
Model for ileal brush-border oxalate transport. Slc26a6–/– mice exhibit ileal oxalate transport characteristics that are decidedly different from those of their Slc26a6+/+ littermates. These observations, together with previous findings regarding the apical localization (36) and exchange modes of PAT1 (4, 17, 27, 38), suggest a working model (Fig. 6) of some components of oxalate transport across the brush-border membrane of ileal enterocytes. Under the conditions of the present experiments, we suggest that PAT1 mediates the efflux (cytosol to lumen) of oxalate across the apical membrane of ileal enterocytes in exchange for luminal Cl–. The first piece of evidence supporting this hypothesis is derived from a simple comparison of the unidirectional fluxes of oxalate in WT and KO mice (Fig. 1) under control conditions. Deletion of PAT1 caused a significant reduction (50%) in JsmOx, whereas JmsOx actually increased (2-fold) in KO mice. The most simple explanation of these observations is that PAT1 mediates a component of JsmOx, and in its absence this unidirectional flux is diminished, whereas the increase in JmsOx is secondary to the decreased secretory flux of oxalate through the epithelium (as elaborated below). Second, 200 µM mucosal DIDS (Fig. 2) had no effect on oxalate transport in KO mouse ileum, but this stilbene significantly reduced JsmOx in WT mouse ileum, implying that mucosal DIDS sensitivity resides in PAT1 and suggests, again, that this protein mediates a component of the secretory flux of oxalate. The failure of mucosal DIDS to inhibit JmsOx does not necessarily mean that the apical uptake mechanism is DIDS insensitive but rather that, at the concentration employed (200 µM), only the PAT1 exchanger was affected. Third, in WT mice, both unidirectional fluxes were affected by mucosal Cl– removal; JmsOx was greater and JsmOx was smaller than that in the presence of luminal Cl–. In contrast, in KO mice only, JmsOx was affected by Cl– removal (Fig. 4), and, like the WT mice, this flux was greater in the absence of Cl–. The fact that in both WT and KO mice JmsOx increased significantly in the absence of mucosal Cl– might also imply a competitive interaction between oxalate and Cl– at the apical membrane of the enterocyte in the course of apical oxalate uptake.
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The model presented in Fig. 6 assumes that the differences in oxalate transport between WT and KO mice originate solely from the lack of Slc26a6 in the KO mouse. However, it is always possible that there are compensatory adaptations of other transporters that directly or indirectly affect oxalate transport. For example, it has been reported that DRA (Slc26a3) is upregulated in Na+/H+ exchanger 3 null mouse colon (26). The over twofold enhancement of JmsOx in PAT1 null mice (compared with WT; Fig. 1D) raised the possibility that this increase may be the result of an overexpression of the apical transporter mediating oxalate absorption. The molecular identity of the brush-border oxalate uptake pathway remains to be identified, but other members of the Slc26a family are possible candidates. In an earlier report, no differences in DRA (Slc26a3) mRNA were observed between WT and KO mouse small intestine (36), a result that we have confirmed here. In addition to our finding that DRA mRNA levels in WT and KO mouse ileum were similar, we found no evidence (Table 2) that CFTR or other members of the Slc26a family (SAT1, DRA, and DTDST) are upregulated in the KO mouse ileum. How other exchangers, like members of the Slc4 and Slc13 families, are involved in ileal oxalate transport and how they are impacted by PAT1 deletion needs to be assessed.
In the KO mouse ileum, JmsOx was significantly greater (2-fold) than that measured in WT ileum and, quantitatively, this elevation in JmsOx was mostly responsible for the significant net absorption of oxalate in the KO (Fig. 1) mouse ileum. If the exchangers mediating apical efflux (PAT1) and the unidentified apical influx exchanger(s) are located in the same cell, then the enhanced absorption of oxalate might be explained as a reduction in shunting (or recycling) of oxalate across the apical membrane in KO mouse ileum. That is, in the absence of the PAT1 oxalate efflux pathway, more of the oxalate that is taken up by the apical influx pathway will be available for export across the basolateral membrane in the KO mouse ileum. Again, identification of the molecular nature of the apical exchanger(s) mediating oxalate uptake will be most useful in resolving these issues.
The model presented in Fig. 6 applies to the specific experimental conditions employed in our study; incorporation of additional transport elements not considered here may reveal alternative interpretations. For example, in sulfate-containing buffers, it is plausible that oxalate is also absorbed via oxalate-sulfate exchange, with luminal oxalate exchanging for cytosolic sulfate. The sulfate could then be recycled via the Na+-SO42– cotransporter (25), which is abundantly expressed in the ileum of both WT and KO mice (data not shown). According to this scheme, in WT animals, the PAT1-mediated Cl–/oxalate exchange would work in parallel with the oxalate/sulfate exchanger, again resulting in the recycling of oxalate with minimal net oxalate transport. Further studies are required to evaluate the role of PAT1 in oxalate absorption under additional experimental conditions.
The results of the present study demonstrate that PAT1 mediates a significant fraction of oxalate efflux across the apical membrane of mouse ileum, which suggests that this specific avenue can contribute to net oxalate secretion by this intestinal segment. The observation of basal net oxalate secretion in WT mouse ileum agrees with previous findings of basal net oxalate secretion in the rat (14) and rabbit (6, 20) ileum under similar experimental conditions. In contrast to this phenomenon of basal net oxalate secretion in the small bowel, the distal colon of rats and rabbits typically exhibits basal net oxalate absorption (12, 15). As noted previously, the DRA protein is reported to be more abundant than PAT1 in the colon, whereas the inverse relationship applies to the small intestine (3, 26, 35, 36, 38). Whether the segmental differences in basal oxalate handling noted above can be simply attributed to differential expression of one or more anion exchangers remains to be established. Finally, It should also be noted that, when oxalate secretion is induced by secretagogues, transport avenues other than anion exchange mechanisms may predominate. For example, in rat and rabbit intestine, secretagogue-stimulated oxalate secretion exhibits many of the characteristics of cAMP-induced Cl– secretion (6, 11, 15).
Urinary oxalate excretion. The PAT1 gene product is also expressed in the proximal tubule where, like in the ileum (36), it is localized to the brush-border membrane (18). It is referred to as CFEX (Cl–/formate exchanger) in this tissue because it was originally suggested to represent the Cl–/formate exchanger (2, 17, 18, 38); however, CFEX is now known to have an affinity for several other anions, including oxalate and HCO3– and can function in a variety of exchange modes involving pairs of these anions (17, 35). A role for luminal oxalate in transcellular NaCl reabsorption was suggested from earlier microperfusion studies which demonstrated that micromolar concentrations of luminal oxalate promoted a DIDS-sensitive increase in volume and Cl– reabsorption (2, 33, 34). Oxalate-stimulated increases in volume reabsorption were not observed in Slc26a6 null mice (36), further supporting a role for Cl–/oxalate exchange in proximal tubule function. Repeated determinations of urinary volume, osmolarity, or Cl– excretion (Table 3) failed to reveal any significant differences between KO and WT mice, in agreement with studies performed on anesthetized mice (36). In addition, urine pH and urinary creatinine excretion were similar in the two groups as reported here. As noted elsewhere (36), downstream events may mitigate the expected salt-wasting effects resulting from a deficiency in oxalate-stimulated NaCl and volume reabsorption in the proximal tubule of KO mice.
In marked contrast, urinary oxalate excretion in KO mice was approximately fourfold greater than that of WT mice (Table 3). Whether this hyperoxaluric phenotype of the KO mouse is the direct result of an absence of Slc26a6 in renal epithelia, a consequence of the hyperabsorption of oxalate by the small intestine as observed here, or some combination of both, cannot be resolved from the present results. Serum oxalate levels were not significantly different between KO and WT mice, although there was a tendency toward hyperoxalemia (28% increase) in the KO mouse population studied here (Table 3). Remarkably, the degree (4x) of hyperoxaluria observed in KO mice vs. WT is similar to the degree of increase in urinary oxalate concentration reported for mice imbibing an oxalate precursor [1% ethylene glycol (EG)] for 4 wk (37) and comparable to that of rats drinking 0.75% EG for 2 wk (7, 10). This degree of hyperoxaluria in mice can lead to the formation of calcium oxalate crystals, the principal component of 75% of renal stones in humans; consequently, the Slc26a6 null mouse may also be useful as a model of nephrolithiasis that is independent of dietary manipulations.
In conclusion, our studies employing Slc26a6 null mice have identified, for the first time, a specific apical membrane protein involved in the transcellular transport of oxalate. The PAT1 anion exchanger plays a significant role in the transcellular secretory unidirectional flux of oxalate across the ileum by mediating oxalate efflux across the apical membrane. The hyperoxaluria observed in the PAT1 KO mice may be the result of enhanced ileal oxalate absorption or because of a defect in renal oxalate handling.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
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C(T) method. Methods 25: 402–408, 2001.[CrossRef][Web of Science][Medline]This article has been cited by other articles:
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) and reduces M-S JOx (
). In B-F, the mean electrical and flux measurements made in the absence of mucosal Cl– during period I (open bars) are compared with those measured in the presence of Cl– during period II (shaded bars). GT and Isc are depicted in B and C, respectively. Note that the reversal in Isc shown in C is because of the bi-ionic diffusion potential generated in the absence of mucosal Cl–, but after readmission of Cl– in period II, the currents were similar to those measured in WT (
