Calcium oxalate urolithiasis in mice lacking anion transporter ... - Nature

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LETTERS

Calcium oxalate urolithiasis in mice lacking anion transporter Slc26a6 Zhirong Jiang1, John R Asplin2,3, Andrew P Evan4, Vazhaikkurichi M Rajendran1, Heino Velazquez5, Timothy P Nottoli6, Henry J Binder1,7 & Peter S Aronson1,7

Urolithiasis is one of the most common urologic diseases in industrialized societies. Calcium oxalate is the predominant component in 70–80% of kidney stones1, and small changes in urinary oxalate concentration affect the risk of stone formation2. SLC26A6 is an anion exchanger expressed on the apical membrane in many epithelial tissues, including kidney and intestine3–6. Among its transport activities, SLC26A6 mediates Cl–-oxalate exchange5–9. Here we show that mutant mice lacking Slc26a6 develop a high incidence of calcium oxalate urolithiasis. Slc26a6-null mice have significant hyperoxaluria and elevation in plasma oxalate concentration that is greatly attenuated by dietary oxalate restriction. In vitro flux studies indicated that mice lacking Slc26a6 have a defect in intestinal oxalate secretion resulting in enhanced net absorption of oxalate. We conclude that the anion exchanger SLC26A6 has a major constitutive role in limiting net intestinal absorption of oxalate, thereby preventing hyperoxaluria and calcium oxalate urolithiasis. Heterologous expression studies have demonstrated that mouse Slc26a6 and human SLC26A6 can function in multiple transport modes, including Cl–-formate exchange, Cl–-oxalate exchange, sulfate-oxalate exchange, Cl–-OH– exchange and Cl–-HCO–3 exchange5–9. To evaluate the role of Slc26a6-mediated anion exchange processes in organ function under in vivo conditions, we generated congenic Slc26a6-null mice (hereafter ‘null mice’). We disrupted the gene Slc26a6 in 129S6/SvEv embryonic stem cells by homologous recombination using a targeting vector in which parts of exons 2 and 5 and all of exons 3 and 4 were replaced with a neomycin resistance (neo) cassette (Fig. 1a). We injected three independent homologous recombinant clones into C57Bl/6 blastocysts to generate chimeric mice, which we bred with 129S6/SvEv mice. We achieved germline transmission and produced heterozygous mice. Homozygous Slc26a6-null mice were generated from the heterozygous mice with the expected

frequency of approximately 25%. We confirmed the genotypes of the mice by DNA blot analysis (Fig. 1b). RNA blot analysis of kidney RNA demonstrated complete loss of the major 2.7-kb Slc26a6 transcript in Slc26a6/ mice and the presence of a mutant 4.1-kb transcript containing the neo sequence in Slc26a6/ and Slc26a6+/ mice (Fig. 1c). Immunoblotting of kidney membrane proteins demonstrated complete loss of Slc26a6 protein in null mice and reduced expression in heterozygotes (Fig. 1d). As mentioned earlier, heterologous expression studies have indicated that SLC26A6 and Slc26a6 can function in multiple anion exchange modes, although there are species differences in ability to mediate Cl– transport9. However, both mouse and human isoforms have a high affinity for oxalate and avidly mediate Cl–-oxalate exchange7–9. To assess directly the role of Slc26a6 as a Cl–-oxalate exchanger, we compared Cl–-oxalate exchange activity in brush border membrane vesicles isolated from the renal cortices of wild-type and Slc26a6-null mice. Imposition of an outward Cl– gradient strongly stimulated the influx of oxalate into brush border vesicles from wildtype mice (Fig. 2a), indicating Cl–-oxalate exchange. However, the ability of an outward Cl– gradient to stimulate oxalate influx was completely abolished in Slc26a6-null mice. These findings demonstrate directly that Slc26a6 accounts for all of the Cl–-oxalate exchange activity in the brush border membrane of proximal tubule cells. These results are consistent with previous microperfusion studies using an independently generated line of Slc26a6-null mice that showed a defect in oxalate-stimulated Cl– absorption in the proximal tubule of the kidney10. In contrast to the complete loss of Cl–-oxalate exchange activity, we saw a partial but significant defect in sulfateoxalate exchange and no significant defects in Cl–-formate exchange and formate-OH– exchange in renal brush border membrane vesicles from Slc26a6-null mice (Fig. 2b–d). In addition, we did not see any detectable change in Na+-H+ exchange activity in Slc26a6-null mice (Fig. 2e). Taken together, these transport studies underscore the essential role of Slc26a6 as an apical membrane Cl–-oxalate exchanger in epithelial cells.

1Department of Internal Medicine, Yale University School of Medicine, New Haven, Connecticut 06520, USA. 2Litholink Corporation, 2250 W. Campbell Park Drive, Chicago, Illinois 60612, USA. 3Renal Section, University of Chicago, Illinois 60612, USA. 4Department of Anatomy and Cell Biology, Indiana University School of Medicine, Indianapolis, Indiana 46223, USA. 5Veterans Administration Connecticut Healthcare System, West Haven, Connecticut 06516, USA. 6Department of Comparative Medicine and 7Department of Cellular and Molecular Physiology, Yale University School of Medicine, New Haven, Connecticut 06520, USA. Correspondence should be addressed to P.S.A. ([email protected]).

Received 12 January 2005; accepted 10 February 2006; published online 12 March 2006; doi:10.1038/ng1762

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Figure 1 Targeted disruption of the Slc26a6 gene. (a) The mouse Slc26a6 gene is shown with the targeting vector and the targeted allele. Homologous recombination resulted in deletion of full exons 3 and 4, deletion of part of exons 2 and 5 and insertion of the neomycin resistance gene (neo). The SalI site used to linearize the construct, the outside probe (filled bar) and the expected sizes of NcoI fragments used for analysis of genomic DNA are indicated. (b) Genotyping the Slc26a6 allele by DNA blot analysis. The expected bands of B5.6 kb for the wild-type Slc26a6 allele and B4.0 kb for the mutant Slc26a6 allele were observed. (c) RNA blot analysis of total RNA (10 mg per lane) from kidney of Slc26a6+/+, Slc26a6+/ and Slc26a6/ adult mice. The blot was analyzed with an Slc26a6 cDNA probe containing exons 4 to 13 and a neo probe. The mutant mRNA which contains the neo gene is B4.1 kb. (d) Protein blots of kidney membrane proteins from Slc26a6+/+, Slc26a6+/ and Slc26a6/ adult mice probed with polyclonal rabbit antibodies against Slc26a6 and b-actin.

40 20 0 WT Null

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[Cl – ]i = [Cl – ]o

[Cl – ]i > [Cl – ]o

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[Sulfate]i = [Sulfate]o [Sulfate]i > [Sulfate]o

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Oxalate uptake (pmol/mg)

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in urine pH or urine concentrations of creatinine, sodium, potassium, calcium, magnesium, citrate and sulfate between wild-type and null mice. Urinary phosphate concentration was moderately higher and urinary osmolality was moderately lower in null mice than in wildtype mice. The decreased urine osmolality may have resulted from the renal medullary crystal deposition in null mice. In contrast, we observed pronounced hyperoxaluria in the Slc26a6-null mice, as the urinary oxalate concentration was 4.1-fold higher than in wild-type mice (Fig. 4a). Indeed, the relative urinary supersaturation of calcium oxalate11 (the ratio of the free ion activity product of calcium and oxalate to the solubility of calcium oxalate) was 62.5 in the null mice, compared with 16.7 in wild-type mice. The value of 62.5 is greatly elevated12, implicating hyperoxaluria as the critical factor most likely responsible for calcium oxalate urolithiasis in Slc26a6-null mice. We did not find any significant gender differences in urinary oxalate concentration that might explain the higher prevalence of urinary stones in male Slc26a6-null mice compared with female null mice. However, we observed that urinary calcium concentration was 2.6-fold higher in males than in females, independent of genotype (34.9 mg dl–1 versus 13.2 mg dl–1, P o 0.02). We did not observe any other significant gender differences in any of the other parameters listed in Table 1. Therefore, it is possible that the higher urinary calcium excretion in male mice, although not sufficient to cause urolithiasis in wild-type mice, aggravates the propensity toward stone formation in hyperoxaluric Slc26a6-null mice.

In view of the role of Slc26a6 as an oxalate transporter, we were intrigued by the observation of a high frequency of urinary stones in the bladders of Slc26a6-null mice (Fig. 3a). Visibly detectable stones were present in 88% of the male (23/26) and 25% of the female (5/20) null mice examined at age 3–6 months. We did not observe any bladder stones in wild-type mice (0/42). Histological examination of the kidney with the Yasue metal substitution method showed birefringent deposits primarily in the lumen of cortical tubules (Fig. 3b,c) and in the urinary space (Fig. 3d,e), with an occasional deposit in the lumen of the inner medullary collecting duct (Fig. 3e) of the Slc26a6null mice. We observed no such birefringent deposits in the kidneys of wild-type mice (data not shown). Analysis of two bladder stones by infrared spectroscopy indicated that they were predominantly (99– 100%) composed of calcium oxalate: one was 100% calcium oxalate dihydrate, and the other was 53% calcium oxalate monohydrate, 46% calcium oxalate dihydrate and 1% calcium phosphate (hydroxyapatite). Thus, these findings indicate a notable disease phenotype of calcium oxalate urolithiasis in Slc26a6-null mice. To understand the mechanisms underlying calcium oxalate urolithiasis in Slc26a6-null mice, we measured plasma and/or urine concentrations of electrolytes and organic substances relevant to kidney function and risk for urinary stone formation (Table 1). We did not find any significant differences in plasma concentrations of urea, Na+, Cl–, HCO–3, calcium or phosphate between wild-type and Slc26a6-null mice. Similarly, we did not find any significant differences

Oxalate uptake (pmol/mg)


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Figure 2 Transport activities in renal brush border membrane vesicles from wild-type and Slc26a6-null mice. (a) Cl–-oxalate exchange assayed as the increment in oxalate uptake induced by an outward Cl– gradient. (b) Sulfate-oxalate exchange assayed as the increment in oxalate uptake induced by an outward sulfate gradient. (c) Cl–-formate exchange assayed as the increment in formate uptake induced by an outward Cl– gradient. (d) Formate-OH– exchange assayed as the increment in formate uptake induced by an outward OH– gradient. (e) Na+-H+ exchange assayed as the increment in Na+ uptake induced by an outward H+ gradient. Data presented are mean ± s.e.m. for n ¼ 9. The difference between wild-type and null was significant only for oxalate uptake measured with Cl– concentration inside greater than Cl– concentration outside ([Cl–]i 4 [Cl–]o; P o 5 10–10) and for oxalate uptake measured with [sulfate]i 4 [sulfate]o (P o 0.01).

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Figure 3 Urolithiasis phenotype in Slc26a6-null mice. (a) Stones found in the bladder of a Slc26a6-null mouse as compared with wild-type bladder without stones. (b) Yasue-positive crystals (arrow) filling the lumen of an atrophic cortical tubule with surrounding inflammation noted by presence of lymphocytes (asterisk). Several nearby proximal tubular segments possess cells that vary in their morphology from normal (brush border noted at high magnifications, not shown) to hyperplastic (double arrow). Magnification, 300. (c) Adjacent dilated cortical tubules with flattened lining cells (arrowhead) that possess Yasue-positive crystals (arrow). A tubule shows an intraluminal crystal mass that is lined by its own cell layer (double arrowheads). Magnification, 250. (d) A large mineral mass (double arrows) in the urinary space pressing against the side of the renal papilla. Magnification, 50. (e) Several small mineral masses (double arrow) in the urinary space at the fornix. A mineral plug is seen in the lumen of a nearby inner medullary collecting duct (arrow). Magnification, 250.

Wild-type Blood urea nitrogen (mg dl–1) Plasma Na (mM) Plasma Cl (mM) Plasma HCO3 (mM) Plasma phosphate (mg dl–1) dl–1)

Plasma Ca (mg Plasma Mg (mg dl–1)

Null

18.9 ± 1.4

18.3 ± 1.8

147.0 ± 1.0 115.0 ± 0.8

146.9 ± 0.6 114.5 ± 0.8

24.3 ± 1.9 9.4 ± 0.9

20.0 ± 0.6 9.0 ± 0.6

9.2 ± 0.2 2.1 ± 0.1

9.6 ± 0.2 1.8 ± 0.1

Urine creatinine (mg dl–1)

29.8 ± 2.3

31.6 ± 2.3

Urine Na (mM) Urine K (mM)

120 ± 12 257 ± 13

118 ± 15 227 ± 27

Urine Ca (mg Urine Mg (mg dl–1)

21.1 ± 5.4 48.3 ± 3.4

27.0 ± 7.0 43.3 ± 5.6

Urine phosphate (mg dl–1) Urine citrate (mM)

122 ± 13 9.0 ± 0.8

166 ± 17 7.2 ± 1.1

55.0 ± 3.0 2,164 ± 183

48.3 ± 3.7 1,512 ± 114

5.38 ± 0.07

5.25 ± 0.09

dl–1)

Urine sulfate (mM) Urine osmolality (mOsmol kg–1) Urine pH

Values are mean ± s.e.m. n ¼ 10–20 for each group. Difference between wild-type and null was significant only for urine phosphate (P o 0.05) and for urine osmolality (P o 0.01).

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c Fecal oxalate (µmol/g)

Table 1 Plasma and urine chemistries in wild-type and Slc26a6-null mice

concentration in Slc26a6-null mice might be explained by reduced oxalate secretion along the gastrointestinal tract. Consistent with this hypothesis, fecal oxalate concentration was significantly lower in Slc26a6-null mice than in wild-type mice (Fig. 4c). Although net secretion of oxalate by the intestine can occur in the setting of renal failure17, under normal conditions there is net absorption of approximately 10% of ingested oxalate18. Our findings raised the possibility that this net absorption may mask a large secretory flux of oxalate in the gastrointestinal tract. In the mouse, Slc26a6 is expressed at high levels in the small intestine (at equivalent levels in the duodenum and jejunum, and at lower levels in the ileum), with far lower levels of expression in the large intestine6. Accordingly, to evaluate directly the possible role of Slc26a6 in intestinal oxalate secretion, we measured the magnitude of unidirectional and net oxalate fluxes across duodenal epithelium in vitro. Slc26a6-null mice had a significant defect in the serosa-to-mucosa flux of oxalate (JSM), resulting in conversion of net oxalate transport (JNET) from net secretion to net absorption (Table 2). These findings are consistent with the hypothesis that reduction of enteric secretion of oxalate in Slc26a6-null mice greatly enhances the net amount of oxalate absorbed, thereby elevating plasma oxalate and causing hyperoxaluria. To test this hypothesis, we evaluated the effect of removing dietary oxalate. An oxalate-free diet greatly ameliorated the elevations in urinary and plasma oxalate in Slc26a6-null mice (Fig. 4), confirming that dietary oxalate is the source of most of the excess oxalate in the urine and plasma of Slc26a6-null mice. Nevertheless, although the

Plasma oxalate (µM)

Previous findings indicating stimulation of proximal tubule Cl– absorption by oxalate have been interpreted to mean that apical membrane Cl–-oxalate exchange mediates Cl– absorption and oxalate secretion13–15. Loss of apical membrane Cl–-oxalate exchange in the proximal tubule of Slc26a6-null mice might therefore be expected to cause reduced rather than increased urinary oxalate excretion. However, although net tubular secretion of oxalate can contribute to urinary oxalate excretion under some conditions, oxalate is predominantly excreted by glomerular filtration under normal conditions16. Accordingly, the hyperoxaluria in Slc26a6-null mice could be explained if there were a substantial increase in the plasma concentration of oxalate in Slc26a6-null mice, leading to enhanced filtration of oxalate. In fact, we observed that the plasma oxalate concentration was 2.1-fold higher in Slc26a6-null mice than in wild-type mice (Fig. 4b). Slc26a6 is expressed on the apical membrane in many gastrointestinal tissues, including the pancreas and small intestine4,6. If Slc26a6 functions as a Cl–-oxalate exchanger and mediates oxalate secretion in one or more of these tissues, then the increased plasma oxalate

Urine oxalate (mM)


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Figure 4 Abnormal oxalate homeostasis in Slc26a6-null mice. Urine (a), plasma (b) and fecal (c) oxalate concentrations were compared between wild-type and Slc26a6-null mice after 7-d equilibration on a control oxalatecontaining diet or an oxalate-free diet. Data represent mean ± s.e.m. for n ¼ 9 to 10 for each group. Differences between null mice and wild-type mice were significant for urine (P o 2 10–8), plasma (P o 0.0002) and fecal oxalate (P o 2 10–7) on the control diet and for urine (P o 0.003) and plasma oxalate (P o 0.002) on the oxalate-free diet.

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LETTERS Table 2 Duodenal oxalate fluxes in wild-type and Slc26a6-null mice JMS (pmol h–1 cm–2)

JSM (pmol h–1 cm–2)

JNET (pmol h–1 cm–2)

ISC (mA cm–2)

GTE (mSi cm–2)

Wild-type

84.6 ± 19.1

157.9 ± 38.5

73.3 ± 23.8

53.0 ± 23.0

13.3 ± 1.5

Null

73.4 ± 9.2

45.3 ± 5.7

28.2 ± 6.3

40.9 ± 13.0

11.8 ± 3.5


Values for mucosa to serosa flux (JMS), serosa to mucosa flux (JSM), net flux (JNET), short-circuit current (ISC) and tissue conductance (GTE) are mean ± s.e.m. n ¼ 6 for each group. The difference between wild-type and null was significant for JSM (P o 0.01) and JNET (P o 0.02).

absolute levels of plasma and urinary oxalate excretion were greatly reduced on the oxalate-free diet, these values were still significantly higher in the null than in the wild-type mice. Fecal oxalate concentration was reduced by 30% in the null mice under these conditions, suggesting that reduced enteric secretion of endogenously produced oxalate was the underlying mechanism, but this conclusion could not be established with certainty because the difference in fecal oxalate did not reach statistical significance. Although Cl–-oxalate exchange is expected to function in the direction of mediating oxalate secretion in the proximal tubule13–15, sulfate-oxalate exchange, which was partially defective in Slc26a6-null mice (Fig. 2b), would be expected to mediate reabsorption of oxalate15,19. Thus, a defect in renal tubular reabsorption of oxalate might also contribute to the hyperoxaluria of Slc26a6-null mice. In fact, the relative increases in urine oxalate in null mice on the control and oxalate-free diets (4.1-fold and 3.7-fold, respectively) were disproportionate to the increases in plasma oxalate (2.1-fold and 1.6-fold, respectively), suggesting that the hyperoxaluria of null mice could not be attributed entirely to the increased filtration of plasma oxalate. However, a primary renal leak of oxalate cannot be the predominant defect leading to hyperoxaluria because such a defect would reduce rather than elevate the plasma oxalate. In partial support of our findings, a recently published report has described enhanced net ileal absorption of oxalate and hyperoxaluria in an independently developed line of Slc26a6-null mice20. However, phenotypes of urolithiasis or renal crystal deposition were not reported in this study, nor were the authors able to reach a conclusion regarding whether the hyperoxaluria was primarily renal or enteric in origin. In summary, we have demonstrated that the anion transporter Slc26a6 has a major constitutive role in limiting net intestinal absorption of oxalate, thereby preventing hyperoxaluria and calcium oxalate urolithiasis. Defective expression of Slc26a6 leads to elevated plasma oxalate, hyperoxaluria and calcium oxalate urolithiasis. Thus, defects in function or regulation of SLC26A6 are potential molecular mechanisms causing calcium oxalate urolithiasis, and SLC26A6 represents a novel therapeutic target for modifying urinary oxalate excretion. METHODS Generation of Slc26a6 knockout mice. We replaced an 0.8-kb fragment of the mouse Slc26a6 gene containing part of exons 2 and 5 and all of exons 3 and 4 with a PGK-neo cassette. This targeting vector in pKO scrambler (Stratagene), which also contains a MC1-TK gene, was linearized at the SalI site and electroporated into the 129S6/SvEv embryonic stem cell line TC-1 (ref. 21). Cells resistant to G418 and ganciclovir were selected, and seven Slc26a6 knockout clones from two independent transfections were identified by DNA blot analysis after NcoI digestion using a PCR-generated probe located downstream of the short arm (Fig. 1a). We microinjected three correctly targeted clones into C57Bl/6 blastocysts and bred the resulting chimeras with 129S6/ SvEv mice (Taconic) to produce congenic mouse lines. We achieved germline transmission and identified heterozygous animals by DNA blot analysis (Fig. 1b) and/or PCR (data not shown). We bred both Slc26a6 knockout

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and wild-type control mice continuously on a 129S6/SvEv genetic background. We performed RNA blot analysis after agarose-formaldehyde gel electrophoresis of total kidney RNA. We then transferred the RNA in the gel to a Hybond-N+ nylon membrane and hybridized with a Slc26a6 cDNA probe containing exons 4 to 13 and with a neo coding sequence probe. We performed immunoblotting using standard procedures described previously22. The primary antibody used was a rabbit antiserum raised against a peptide in the C-terminal hydrophilic domain (residues 503–524) of opossum SLC26A6 (CPDTDIYRDVTEYKEAKE VPGVK). Rabbit antibody to b-actin was from Santa Cruz Biotechnology. Protocols for use of experimental animals were approved by the Yale Institutional Animal Care and Use Committee. Preparation of renal brush border membrane vesicles. We prepared microvillus membrane vesicles from mouse renal cortices by homogenization, magnesium aggregation and differential centrifugation as described23. The homogenization medium contained 200 mM mannitol, 41 mM KOH and 80 mM HEPES, pH 7.5. We suspended the final membrane pellets in this medium to a protein concentration of 30–50 mg ml–1. The vesicle preparations were enriched 10- to 15-fold in specific activity of the luminal membrane marker enzyme g-glutamyltranspeptidase. Measurement of ion transport in renal brush border membrane vesicles. We assayed the rates of uptake of [14C]oxalate, [14C]formate and 22Na into renal brush border membrane vesicles by rapid filtration as described previously19,24,25. For assays of Cl–-oxalate exchange and Cl–-formate exchange, vesicles were washed twice and then preloaded with chloride by preincubation in a pH 8.2 buffer containing 120 mM potassium gluconate, 30 mM potassium chloride and 15 mM HEPES for 2 h at 20 1C. We then performed timed 30-s uptake of 20 mM [14C]oxalate or 40 mM [14C]formate after 1:20 dilution of the vesicles into either 120 mM potassium gluconate, 30 mM potassium chloride and 15 mM HEPES, pH 8.2 (where the Cl–concentration inside equals the Cl– concentration outside ([Cl–]i ¼ [Cl–]o)) or 150 mM potassium gluconate and 15 mM HEPES, pH 8.2 ([Cl–]i 4 [Cl–]o). Similarly, to assay oxalate-sulfate exchange, we preincubated vesicles with 5 mM sulfate and then measured 30-s uptake of 20 mM [14C]oxalate after 1:20 dilution of the vesicles into a solution with ([sulfate]i ¼ [sulfate]o) or without ([sulfate]i 4 [sulfate]o) 5 mM sulfate. To assay formate-OH– exchange, we pre-equilibrated vesicles in pH 7.5 medium and then measured 30-s uptake of 40 mM [14C]formate after 1:5 dilution of the vesicles into pH 7.5 medium ([OH–]i ¼ [OH–]o) or into a medium with final pH 6.0 ([OH–]i 4 [OH–]o). To assay Na+-H+ exchange, we pre-equilibrated vesicles in pH 6.0 medium and then measured 30-s uptake of 1 mM 22Na after 1:5 dilution of the vesicles into pH 6.0 medium ([H+]i ¼ [H+]o) or into a medium with final pH 7.5 ([H+]i 4 [H+]o). We performed each transport assay three times on each of three different membrane preparations prepared from the renal cortices of ten null or wild-type mice. Renal histology. Mouse kidneys were routinely processed for paraffin embedment and positioned in order to obtain complete cross sections of the kidney at the level of the renal papilla. We cut 15 serial sections at 4 mm per kidney and stained alternating sections with either hematoxylin and eosin, for routine histological examination, or by the Yasue metal substitution histochemical method26 to detect the presence of calcium crystals in renal tissue. Measurement of plasma, urine and fecal chemistries. We measured plasma Na+, K+, Cl–, HCO–3, BUN, Ca2+, Mg2+ and inorganic phosphorus with the COBAS MIRA system (Roche Diagnostics). We put individual mice into metabolic cages to collect urine and fecal samples. We measured urine pH and osmolality with a Jenco pH meter and an Advanced Micro-osmometer

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LETTERS (Advanced Instrument), respectively. We measured urine calcium, magnesium, sodium, potassium, phosphorus and creatinine using a Beckman Synchron autoanalyzer. We measured oxalate, citrate and sulfate by ion chromatography using a Dionex ICS 2000 system (Dionex Corporation) equipped with an AG-11 guard column and AS-11 analytical column in series27. The mobile phase was potassium hydroxide at a flow rate of 2 ml min–1. We detected ion peaks using a conductivity meter, with the eluent background conductivity suppressed using an anion self-regenerating suppressor (ASRS Ultra II, Dionex). We measured plasma oxalate by diluting the plasma ultrafiltrate 1:3 with 0.3 M boric acid and loading onto the column with a 100-ml injection loop. For urine and fecal samples, we prepared an ultrafiltrate using a 10-kDa centrifugal ultrafiltration device. We loaded the samples using a 25-ml injection loop. Urine and fecal sample dilutions varied from 1:10 to 1:80 so that the peak area fell in the range of the calibration. We used Chromeleon Software (version 6.5; Dionex) to calculate the concentration of the oxalate peak. We performed assays of chow and fecal oxalate concentration on extracts prepared by heating samples at 60 1C for 3 h in 5 N HCl (ref. 28). We performed all measurements on 12-week-old gender-matched mice that were fed control chow (Harlan), which was measured to contain oxalate at 12 mmol g–1. We also performed urine, plasma and fecal oxalate measurements on a separate group of 12-week-old gender-matched mice that we fed for 7 d an oxalate-free diet (Harlan, Purified Basic Diet) that did not contain detectable oxalate (o0.4 mmol g–1). Kidney stone analysis by infrared spectroscopy was performed by Beck Laboratories. Measurement of oxalate fluxes across duodenum. Duodenal segments (3 cm) adjacent to stomach were opened longitudinally along the mesenteric border and mounted as an intact sheet in a modified Ussing chamber that had an exposed surface area of 0.11 cm2. We bathed mucosal and serosal surfaces of the duodenal segments with 10 ml of warmed (37 1C), oxygenated bicarbonate Ringer’s solution (139.4 mM Na+, 123.2 mM Cl–, 5.4 mM K+, 1.2 mM Mg2+, 21 mM HCO–3, 0.6 mM HPO–4, 2.4 mM H2PO2– 4 and 10 mM glucose at pH 7.4). We measured transepithelial short-circuit current (Isc) and total tissue conductance (GTE) as described earlier29. We measured fluxes of [14C]oxalate by previously described methods17. We added 2 mM [14C]oxalate either to mucosal or to serosal bath. After a 30-min equilibration, we collected a 60-min sample to calculate unidirectional mucosa to serosa (JMS) and serosa to mucosa (JSM) fluxes. We derived net oxalate flux (JNET) by determining the difference between JMS and JSM. Positive JNET indicates net absorption, whereas negative values indicate net secretion. We performed all flux studies under voltage clamp conditions using a DVC 1000 (World Precision Instruments). Statistical analysis. Results in all experiments are given as the mean ± s.e.m. for the indicated number of experiments. The results were analyzed by unpaired two-tailed Student’s t-test. ACKNOWLEDGMENTS This work was supported by US National Institutes of Health grants DK33793, DK17433, DK56788 and DK60699. COMPETING INTERESTS STATEMENT The authors declare that they have no competing financial interests. Published online at http://www.nature.com/naturegenetics Reprints and permissions information is available online at http://npg.nature.com/ reprintsandpermissions/

1. Asplin, J.R. Hyperoxaluric calcium nephrolithiasis. Endocrinol. Metab. Clin. North Am. 31, 927–949 (2002).

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2. Robertson, W.G. & Peacock, M. The cause of idiopathic calcium stone disease: hypercalciuria or hyperoxaluria? Nephron 26, 105–110 (1980). 3. Waldegger, S. et al. Cloning and characterization of SLC26A6, a novel member of the solute carrier 26 gene family. Genomics 72, 43–50 (2001). 4. Lohi, H. et al. Mapping of five new putative anion transporter genes in human and characterization of SLC26A6, a candidate gene for pancreatic anion exchanger. Genomics 70, 102–112 (2000). 5. Knauf, F. et al. 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). 6. Wang, Z., 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). 7. Jiang, Z., Grichtchenko, I.I., Boron, W.F. & Aronson, P.S. Specificity of anion exchange mediated by mouse Slc26a6. J. Biol. Chem. 277, 33963–33967 (2002). 8. Xie, Q., Welch, R., Mercado, A., Romero, M.F. & Mount, D.B. Molecular characterization of the murine Slc26a6 anion exchanger: functional comparison with Slc26a1. Am. J. Physiol. Renal Physiol. 283, F826–F838 (2002). 9. Chernova, M.N. et al. 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). 10. Wang, Z. et al. Renal and intestinal transport defects in Slc26a6-null mice. Am. J. Physiol. Cell Physiol. 288, C957–C965 (2005). 11. Werness, P.G., Brown, C.M., Smith, L.H. & Finlayson, B. EQUIL2: a BASIC computer program for the calculation of urinary saturation. J. Urol. 134, 1242– 1244 (1985). 12. Asplin, J. et al. Supersaturation and stone composition in a network of dispersed treatment sites. J. Urol. 159, 1821–1825 (1998). 13. Wang, T., Giebisch, G. & Aronson, P.S. Effects of formate and oxalate on volume absorption in rat proximal tubule. Am. J. Physiol. 263, F37–F42 (1992). 14. Wang, T., Segal, A.S., Giebisch, G. & Aronson, P.S. Stimulation of chloride transport by cAMP in rat proximal tubules. Am. J. Physiol. 268, F204–F210 (1995). 15. Wang, T., Egbert, A.L. Jr., Abbiati, T., Aronson, P.S. & Giebisch, G. Mechanisms of stimulation of proximal tubule chloride transport by formate and oxalate. Am. J. Physiol. 271, F446–F450 (1996). 16. Holmes, R.P., Ambrosius, W.T. & Assimos, D.G. Dietary oxalate loads and renal oxalate handling. J. Urol. 174, 943–947 (2005). 17. Hatch, M., Freel, R.W. & Vaziri, N.D. Intestinal excretion of oxalate in chronic renal failure. J. Am. Soc. Nephrol. 5, 1339–1343 (1994). 18. Holmes, R.P. & Assimos, D.G. The impact of dietary oxalate on kidney stone formation. Urol. Res. 32, 311–316 (2004). 19. Kuo, S.M. & Aronson, P.S. Pathways for oxalate transport in rabbit renal microvillus membrane vesicles. J. Biol. Chem. 271, 15491–15497 (1996). 20. Freel, R.W., Hatch, M., Green, M. & Soleimani, M. Ileal oxalate absorption and urinary oxalate excretion are enhanced in Slc26a6-null mice. Am. J. Physiol. Gastrointest. Liver Physiol. advanced online publication 22 December 2005 (doi:10.1152/ ajpgi.00481.2005). 21. Deng, C., Wynshaw-Boris, A., Zhou, F., Kuo, A. & Leder, P. Fibroblast growth factor receptor 3 is a negative regulator of bone growth. Cell 84, 911–921 (1996). 22. Kocinsky, H.S. et al. Use of phospho-specific antibodies to determine the phosphorylation of endogenous Na+/H+ exchanger NHE3 at PKA consensus sites. Am. J. Physiol. Renal Physiol. 289, F249–F258 (2005). 23. Karniski, L.P. et al. Formate-stimulated NaCl absorption in the proximal tubule is independent of the pendrin protein. Am. J. Physiol. Renal Physiol. 283, F952–F956 (2002). 24. McConnell, K.R. & Aronson, P.S. Effects of inhibitors on anion exchangers in rabbit renal brush border membrane vesicles. J. Biol. Chem. 269, 21489–21494 (1994). 25. Wu, M.S., Biemesderfer, D., Giebisch, G. & Aronson, P.S. Role of NHE3 in mediating renal brush border Na+-H+ exchange. Adaptation to metabolic acidosis. J. Biol. Chem. 271, 32749–32752 (1996). 26. Yasue, T. Histochemical identification of calcium oxalate. Acta Histochem. et Cytochem. 2, 83–95 (1969). 27. Hoppe, B., Kemper, M.J., Hvizd, M.G., Sailer, D.E. & Langman, C.B. Simultaneous determination of oxalate, citrate and sulfate in children’s plasma with ion chromatography. Kidney Int. 53, 1348–1352 (1998). 28. Palgi, N., Vatnick, I. & Pinshow, B. Oxalate, calcium and ash intake and excretion balances in fat sand rats (Psammomys obesus) feeding on two different diets. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 141, 48–53 (2005). 29. Vidyasagar, S., Rajendran, V.M. & Binder, H.J. Three distinct mechanisms of HCO3 – secretion in rat distal colon. Am. J. Physiol. Cell Physiol. 287, C612–C621 (2004).

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