Screening of a Human Renal cDNA Library.

A human kidney cDNA library was purchased from Invitrogen (Carlsbad, CA) and screened by colony hybridization with a 42-mer oligonucleotide (5′-CACTGCCGCCCGCCTGCCGATGCCAACCTCAGCAAGAACGGG-3′) corresponding to the 5′-end of clone expressed sequence tag (EST)58612 (GenBank accession number AA351032). The oligonucleotide was labeled with [³²P]dATP using T4 polynucleotide kinase. Bacterial colonies were transferred onto Hybond-N membranes (Amersham Pharmacia Biotech, Arlington Heights, IL) and hybridized overnight with the labeled probe in hybridization buffer (5× standard saline citrate (SSC), 5× Denhardt’s solution, 0.05% SDS) at 68°C. Final washing was performed twice with 0.1× SSC + 0.1% SDS. Filters were subjected to autoradiography overnight, and several positive colonies were identified. Plasmid DNA was purified from these clones, and the length of their cDNA inserts determined by polymerase chain reaction (PCR).

DNA Sequencing and Analysis.

The clone containing the largest cDNA insert (pOAT-8) was subjected to complete sequence analysis of both complementary strands by simultaneous “walking” from the 5′- and 3′-ends of the insert. Automatic sequencing was performed with Cy5-labeled oligonucleotide primers (Integrated DNA Technologies, Coralville, IA) using the Thermosequenase cycle sequencing kit (Amersham Pharmacia Biotech). All ambiguities were resolved by manual sequencing using the Thermosequenase radiolabeled terminator cycle sequencing kit (Amersham Pharmacia Biotech). If necessary, dideoxy-ITP was substituted for dideoxy-GTP to resolve sequence compressions. Initially, a BLAST search of the GenBank database identified mNKT(mROAT1) and rROAT1 as being related sequences, but no human homologs were found, and we designated this cDNA clone as hOAT1. All DNA and peptide sequence comparisons and database searches were done with the Wisconsin Package software (Genetics Computer Group, Madison, WI) with default settings.

Northern Blotting.

A human multiple tissue Northern blot (CLONTECH, Palo Alto, CA) was used for localization of hOAT1 expression in human tissues. An hOAT1-specific [³²P]dATP-labeled probe was generated by random priming the BsrGI/Bsu36I DNA fragment from pOAT-8 (nucleotides 420–854 of hOAT1 coding region). The membrane was hybridized for 1 h at 68°C in ExpressHyb hybridization buffer (CLONTECH) and then washed twice in 2× SSC with 0.05% SDS for 30 min at room temperature followed by a single wash in 0.1× SSC with 0.1% SDS at 50°C. The membrane was autoradiographed for 7 days at −70°C with intensifying screens. After stripping, the membrane was reprobed with a β-actin control probe provided by the manufacturer and autoradiographed for 3 h.

Tissue Localization by Reverse Transcription (RT)-PCR.

The multiple choice cDNA kits I and II containing tissue-specific cDNAs (Origene Technologies, Rockville, MD) and two sets of hOAT1-specific primers were used for PCR detection of hOAT1 expression. The oligonucleotides, 5′-CCCGCTGGCACTCCTCCTCCGGGAG-3′ (sense), and 5′-GTAGAGCTCGGCAGTCATGCTCACCA-3′ (antisense), were used to amplify a 606-bp fragment from the hOAT1 coding region (nucleotides 815-1420). In the independent set of PCR reactions, a 295-bp hOAT1 fragment [composed of the last 175 coding nucleotides and 120 nucleotides of the 3′-untranslated region (UTR)] was amplified using the oligonucleotides, 5′-CCAGCGCTGTCACTGTCCTCCTGC-3′ (sense), and 5′-AACCCCCACACTTGGGTCACCATTTCCTC-3′ (antisense).

PCR reactions were carried out using the Expand High Fidelity PCR system (Roche Molecular Biochemicals) in a total volume of 25 μl containing 1 μg of tissue-specific cDNA. Thirty-five amplification cycles (95°C for 40 s, 58°C for 1 min, and 72°C for 45 s) were performed. A negative control without cDNA template was included in each set of reactions. As an internal control, a β-actin fragment was amplified using primers provided by the manufacturer.

In Vitro Expression of hOAT1.

For efficient in vitro expression of hOAT1, the plasmid pOAT-8/ITT was constructed as follows. A fragment containing the first 500 bp of the hOAT1 coding region was amplified from pOAT-8 by the PCR using the sense primer 5′-ATGCCTGGATCCAAGCTATTTAGGTGACACTATAGAATACTCAAGCTTGTTTTTATTTTTAATTTTCTTTCAAATACGTCCACCATGGCCTTTAATGACC-3′ to introduce a BamHI restriction site, the SP6 promoter, a truncated form of the 5′-UTR from alfalfa mosaic virus, and a favorable initiation context (these elements are underlined in the same order in the primer sequence). The oligonucleotide 5′-CGATGCCTGGCCTTTGCCCATGGTCAGCTC-3′ was used as an antisense primer. The PCR product was digested with BamHI andBsrGI, and subcloned intoBamHI/BsrGI-digested pOAT-8. Plasmid pOAT-8/ITT was expressed in vitro with the TNT SP6 Coupled Reticulocyte Lysate system (Promega, Madison, WI). A 25-μl reaction, containing 2 μg of circular plasmid and other components according to the manufacturer’s protocol including complete amino acid mixture, was incubated for 2 h at 30°C and subjected to immunoblot analysis as described below.

Immunoblot Analysis of hOAT1.

Rabbit anti-hOAT1 polyclonal antibody was prepared at AnaSpec (San Jose, CA) using standard immunological techniques. Briefly, a peptide corresponding to hOAT1 amino acids 515 to 528 was conjugated to keyhole limpet hemocyanine through a cysteine residue added to the C terminus of the peptide. Animal serum was collected after four immunizations in the presence of complete Freund’s adjuvant and affinity-purified against the immunogenic peptide immobilized on a Sepharose resin.

For immunoblot analysis, human kidney cortex was obtained from the International Institute of Advanced Medicine (Exton, PA), frozen in liquid nitrogen, ground to a powder, and mixed with 10-fold excess (w/v) of extraction buffer (20 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% Nonidet P-40, 0.5% deoxycholate, 0.1% SDS). After homogenization in the presence of complete proteinase inhibitor cocktail (Roche Molecular Biochemicals) the extract was clarified by high-speed centrifugation. An aliquot was deglycosylated by incubation in the presence of peptide:N-glycosidase F (New England Biolabs, Beverly, MA) for 2 h at 37°C without prior denaturation. Tissue extracts (50 μg of protein) and in vitro expression reactions (10 μl) were separated by electrophoresis on an 8% SDS-polyacrylamide gel and electroblotted onto a nitrocellulose membrane (Millipore, Bedford, MA). The membrane was subsequently blocked in PBS/5% dry milk (PBS-M) for 1 h, washed three times in PBS/0.05% Tween 20, and incubated overnight in PBS-M with the anti-hOAT1 antibody. Following washing in PBS/0.05% Tween 20, the membrane was incubated in PBS-M with goat anti-rabbit antibody conjugated to horseradish peroxidase (Zymed, South San Francisco, CA). After an additional wash and incubation with a chemiluminescent substrate (Amersham Pharmacia Biotech), the immunoblot was exposed to X-ray film.

Xenopus Oocyte Expression Assay.

For batch oocyte isolation, adult female Xenopus laevis (Xenopus One, Ann Arbor, MI) were anesthetized by hypothermia and decapitated. Ovaries were removed and divided into clumps of ∼50 to 100 oocytes. Oocyte clumps were placed into a 50-ml conical tube containing oocyte Ringer’s 2 (OR-2: 82.5 mM NaCl, 2.5 mM KCl, 1 mM Na₂HPO₄, 3 mM NaOH, 1 mM CaCl₂, 1 mM MgCl₂, 1 mM pyruvic acid, and 5 mM HEPES, pH 7.6) until they reached the 4-ml mark. Oocytes were rinsed three times with 10 ml of calcium-free OR-2 (82.5 mM NaCl, 2 mM KCl, 1 mM MgCl₂, and 5 mM HEPES, pH 7.6), placed in collagenase solution [5 mg/ml collagenase A (Roche Molecular Biochemicals) and 1 mg/ml trypsin inhibitor type III-O (Sigma Chemical Co., St. Louis, MO) in calcium-free OR-2] and swirled at 100 rpm for 2 to 2.5 h at 18°C. After collagenase treatment, the oocytes were rinsed five times with OR-2 containing 1 mg/ml BSA and placed in a phosphate/BSA solution for 1 h (100 mM K₂HPO₄ with 1 mg/ml BSA, pH 6.5). Oocytes were agitated every 15 min to remove the follicle and then rinsed five times with OR-2 + BSA. Stages V and VI oocytes were collected and maintained at 18°C in OR-2 containing 0.05 mg/ml gentamycin sulfate, 2.5 mM sodium pyruvate, and 5% heat-inactivated horse serum. Overnight recovery was allowed before injection.

Capped cRNA for microinjection was synthesized from linearized plasmid DNA using Ambion’s mMessage mMachine in vitro transcription kit (Ambion, Inc., Austin, TX). The cRNA products were quantitated in a spectrophotometer and diluted before injection to allow delivery of 20 ng of cRNA/oocyte in 16 nl with a 1- to 2-s injection.

Uptake assay was performed as described previously (Sweet et al., 1997). Briefly, 3 days after injection, the oocytes were divided into experimental groups (containing 10 oocytes each) and incubated at 18–22°C for 20 or 60 min in OR-2 containing various substrates in the absence or presence of inhibitors, as indicated in the figure legends. After uptake, oocytes were rapidly rinsed three times with ice-cold OR-2 and placed into individual scintillation vials containing 0.5 ml of 1 M NaOH, incubated at 65°C for 20 min, and neutralized with 0.5 ml of 1 M HCl. Finally, 4.7 ml of Ecolume (ICN Biomedical, Cleveland, OH) was added and radioactivity was determined in a Packard 1600TR liquid scintillation counter with external quench correction. Uptake was calculated in picomoles per oocyte.

Statistics.

Data are presented as mean ± S.E. Differences in mean values were considered to be significant whenp ≤ .05 as determined by one-sample or paired Student’s t test. The degree of significance was as indicated in the figure legends.

Chemicals.

[³H]PAH (3.7 Ci/mmol) was obtained from NEN Life Science Products (Boston, MA). [¹⁴C]Tetraethylammonium (53 mCi/mmol) was obtained from American Radiolabeled Chemicals, Inc. (St. Louis, MO). Adefovir (30 Ci/mmol), cidofovir (56 mCi/mmol), [³H]9-(2-phosphonylmethoxyethyl)guanine (PMEG; 34 Ci/mmol), and [³H]9-(2-phosphonylmethoxyethyl)diaminopurine (PMEDAP; 32 Ci/mmol) were purchased from Moravek Biochemicals (Brea, CA). All unlabeled nucleoside phosphonates were synthesized at Gilead Sciences according to previously published procedures (Holy and Rosenberg, 1987). Unlabeled PAH, probenecid, and glutarate were obtained from Sigma.

Cloning of hOAT1 cDNA.

To identify potential cDNA sequences encoding a hOAT homolog, a BLAST search of the GenBank EST database was performed with the full-length rROAT1 cDNA sequence (Sweet et al., 1997). Two EST cDNA clones from human infant brain (accession numberR25797 and AA351032), whose 5′-end regions exhibited a high level of sequence homology to rROAT1 between coding nucleotides 390 and 730, were identified. Therefore, an oligonucleotide probe derived from EST clone AA351032 was used to screen a human kidney cDNA library, and a positive clone was identified (pOAT-8). We refer to this cDNA clone as hOAT1.

Molecular Characterization of hOAT1.

The complete DNA sequence of both strands of hOAT1 was determined, and the sense strand sequence is presented in Fig. 2A. The hOAT1 cDNA is 2118-bp long, including: 257 bp of 5′-UTR; a single, large open reading frame 1650-bp long; and 211 bp of 3′-UTR. The ATG at position 258 to 260 is the first start codon and has a correlation with Kozak’s consensus for the translation initiation (Kozak, 1987) giving a predicted protein length of 550 amino acids with a molecular mass of 60.3 kDa. Hydropathy analysis (Kyte and Doolittle, 1982) predicted 12 potential α-helical membrane-spanning domains with both the amino and carboxyl termini located intracellularly (Fig. 2). According to this analysis, there is a large extracellular loop between transmembrane domains 1 and 2. Within this loop are five potentialN-linked glycosylation sites at Asn39, Asn56, Asn92, Asn97, and Asn113 and four cysteine residues at positions 49, 78, 105, and 128 that could participate in disulfide bond formation (Fig. 2A). A significant intracellular loop between transmembrane domains 6 and 7, from approximately amino acid residues 271 to 336, contains four protein kinase C (PKC) consensus sites located at Ser271, Ser278, Thr284, and Thr334 and a potential casein kinase II (CKII) site at Ser325 (Fig. 2A). Finally, within the carboxyl-terminal domain there is another putative PKC site at Thr521, two potential CKII sites at Thr515 and Ser543, and a tyrosine kinase consensus site at Tyr536. Whether any or all of these consensus sites participate in protein modification or function requires further experimentation.

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Figure 2

DNA and predicted amino acid sequence of hOAT1. A, complete nucleotide sequence of the sense strand of the hOAT1 cDNA and the predicted amino acid sequence of the single large open reading frame is shown. Approximate locations of the 12 predicted transmembrane domains are indicated below the peptide sequence by bars. Cysteine residues (●) and potential N-linked glycosylation sites (∗) within the first extracellular loop are indicated. Potential intracellular phosphorylation sites for PKC (♦), CKII (▪), and TK (▵) are identified. Numbers to the left of the sequences indicate nucleotide and amino acid position, respectively. B, Kyte-Doolittle hydropathy plot. Proposed transmembrane domains are indicated by number.

During the course of our study, other reports have appeared in the literature concerning the cloning of hROATs designated hROAT1 (Reid et al., 1998), hOAT1–1 and hOAT1–2 (Hosoyamada et al., 1999), and human p-aminohippurate transporter (hPAHT) (Lu et al., 1999). Unfortunately, the DNA sequences for hOAT1–1 (accession numberAB009697), hOAT1–2 (accession number AB009698), and hPAHT (no accession number reported) were not yet available in the GenBank database to compare with our hOAT1. Although there are some minor differences between our DNA sequence and that for hROAT1 (accession number AF057039), the predicted peptide sequence of our clone is identical with hROAT1 except for the final amino acid residue, Phe in hROAT1 versus Leu in hOAT1. At this time, no functional data has been published for hROAT1. Comparison with the published peptide sequence of hOAT1–2 confirms that our clone is the same, however, no characterization of hOAT1–2 was presented (Hosoyamada et al., 1999). Similarly, comparison with the published hPAHT sequence reveals that the sequence of our clone is identical except for the amino acid at position 14 (Ser in hPAHT versus Gly in hOAT1).

Alignment of the hOAT1 DNA sequence with hROAT1, rROAT1, mNKT(mROAT1), and fROAT showed a 99.7, 83.8, 83, and 59.5% identity, respectively. A summary of potential modification sites for these transporters is presented in Table 1. It is our opinion that hOAT1–2 and hPAHT have all of the same sites as those listed for hOAT1, yet, there was no mention of PKA, CKII, or tyrosine kinase (TK) consensus sites for hOAT1–2 (Hosoyamada et al., 1999) nor any CKII or TK sites for hPAHT (Lu et al., 1999). In contrast to our findings, Lu et al. (1999) reported four PKA consensus sites within the hPAHT sequence, however, these sites do not conform to the consensus used in our motif search (R,K)2X(S,T).

Expression of hOAT1 in Human Tissues.

Northern blot analysis was used to examine hOAT1 expression in different human tissues with an hOAT1-specific DNA probe. A strong signal was detected in kidney corresponding to a transcript 2.5-kb in size (Fig.3a, top). In addition, there was a very weak signal of about 3.5 kb in liver that was detectable only after prolonged exposure. No positive signal was found in brain, heart, skeletal muscle, colon, thymus, spleen, small intestine, placenta, lung, or peripheral blood leukocytes. The lack of a detectable signal in brain was an unexpected result because, as mentioned above, two EST clones from a human infant brain cDNA library were identified with sequence identical to a 340-bp region of hOAT1. Therefore, we used the more sensitive technique of RT-PCR amplification to examine hOAT1 expression in various human tissues. Each sample was analyzed independently using two distinct sets of primers with proper negative controls (Fig. 3B, top and middle). A strong positive signal was consistently detected in kidney with both sets of primers. In contrast with the Northern analysis, RT-PCR amplification consistently yielded a product from brain. Interestingly, a strong positive signal was also detected in skeletal muscle, although only with one set of hOAT1-specific primers.

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Figure 3

Detection of hOAT1 expression in human tissues. A, Northern analysis of human tissue-specific poly(A)+ RNA. After hybridization with a 430-bp, ³²P-labeled hOAT1 fragment, a 2.5-kb transcript was detected in kidney (top). The membrane was stripped and reprobed with a ³²P-labeled human β-actin-specific probe (bottom). B, analysis of hOAT1 tissue distribution using RT-PCR amplification of 606- and 295-bp hOAT1-specific regions (top and middle, respectively). As a control, a β-actin fragment was amplified from each tissue sample (bottom).

Immunoblot Analysis of hOAT1.

A polyclonal antibody was generated in rabbits immunized with a synthetic peptide derived from a region of hOAT1 with predicted high antigenicity. In the immunoblot analysis of an extract from human kidney cortex, the antibody recognized a heterogeneous product with an apparent molecular mass of approximately 80 to 90 kDa (Fig.4). This was significantly larger than the predicted molecular mass from the hOAT1 amino acid sequence. However, when the cortex extract was treated with peptide/N-glycosidase F, which specifically cleavesN-linked oligosaccharide chains, a 60-kDa homogeneous product was detected on the immunoblot. To determine whether this treatment completely removes all modifications from the hOAT1 peptide, we attempted to generate a control for the unmodified polypeptide by in vitro expression of hOAT1 cDNA in rabbit reticulocyte lysate. At first, no product of in vitro expression was detected in the presence of pOAT-8 plasmid. To achieve efficient expression, the original hOAT1 5′-UTR was replaced by a truncated form of the alfalfa mosaic virus RNA4 5′-UTR, which has been previously shown to significantly stimulate in vitro expression in reticulocyte lysate (Cihlar et al., 1997). In addition, the sequence surrounding the starting ATG codon was modified to match an optimal initiation context (Kozak, 1987). As shown in Fig.4, the modified plasmid yielded a single polypeptide whose size was identical to that of the deglycosylated hOAT1. Together, these findings indicate that hOAT1 is modified by abundant N-glycosylation, which could account for up to 30% of its total molecular weight. Importantly, because all potential N-glycosylation sites reside within the large loop between the first and second putative transmembrane domain, it also shows that this loop is indeed located extracellularly as predicted from the hOAT1 hydropathy plot.

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Figure 4

Immunoblot analysis of hOAT1. Samples were separated by electrophoresis on an 8% SDS-polyacrylamide gel, electroblotted onto nitrocellulose membrane, and probed with an hOAT1-specific polyclonal antibody followed by chemiluminescent detection. Kidney cortex extract before (lane 1) and after (lane 2) treatment with peptide/N-glycosidase F, and in vitro transcription/translation reaction in the presence (lane 3) and absence (lane 4) of pOAT-8/ITT plasmid.

Functional Characterization of hOAT1 Transport.

That the hOAT1 cDNA does indeed encode an organic anion transporter was confirmed byXenopus oocyte expression assay. After cRNA injection, hOAT1-mediated uptake of 50 μM [³H]PAH was equivalent to that seen with the previously identified basolateral organic anion transporter rROAT1 (Fig. 5;Sweet et al., 1997). Addition of 1 mM probenecid reduced PAH uptake by 98%, proving that virtually all of the uptake was carrier mediated. Water-injected oocytes showed no appreciable uptake and were unaffected by probenecid.

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Figure 5

Functional activity of hOAT1 inXenopus oocytes. Three days after injection with rROAT1 or hOAT1 cRNA, oocytes were incubated in buffer containing 50 μM [³H]PAH for 60 min, and uptake was determined in the absence (No Inhibitor) or presence (+ Inhibitor) of 1 mM probenecid. Data are mean ± S.E. values from a representative animal (10 oocytes/treatment).

The effects of various compounds and reduced temperature on hOAT1-mediated PAH uptake were also examined (Fig.6). The organic anions probenecid, α-ketoglutarate (α-KG), bromcresol green, and 2,4-dichlorophenoxyacetic acid, as well as excess unlabeled PAH, each reduced [³H]PAH uptake by at least 95% at 1 mM concentration. Interestingly, unlike the results reported for rROAT1 (Sweet et al., 1997), the organic anion urate (1 mM) was a fairly effective inhibitor of hOAT1, reducing PAH uptake by 75%; however, when used as substrate, there was no detectable transport of urate (data not shown). Transport was unaffected by the cation tetraethylammonium (1 mM) or by the P-glycoprotein inhibitor cyclosporin-A (10 μM). Furthermore, the ability of α-KG tocis-inhibit PAH uptake is indicative of a basolateral location for hOAT1, because the basolateral dicarboxylate/organic anion exchangers (e.g., rROAT1) should be inhibited by external α-KG as opposed to the luminal PAH carriers, which should be insensitive to this inhibition (Pritchard and Miller, 1993).

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Figure 6

Inhibition profile of hOAT1-mediated PAH uptake. cRNA from hOAT1 was injected into oocytes, and 50 μM [³H]PAH transport was measured after 60 min in the presence of several organic anions and cations. Data are presented as percentage of control uptake. Values are mean ± S.E. for two animals (10 oocytes/treatment/animal). *p ≤ .05, **p ≤ .01, and ***p ≤ .001, as determined by one sample Student’s t test.

rROAT1 has been shown to function as a basolateral dicarboxylate/organic anion exchanger. If hOAT1 functions in the same way in human kidney, increasing the intracellular concentration of dicarboxylate should induce trans-stimulation of exogenous substrate uptake (Pritchard and Miller, 1993). For this determination, glutarate is the preferred counterion, because, in contrast to α-KG, it is not extensively metabolized (Pritchard, 1990). As shown in Fig.7, incubating the oocytes in 5 mM glutarate for 90 min before exposure to PAH (i.e., preloading) significantly (p < .0001) stimulated PAH uptake in hOAT1-expressing oocytes, as compared with nonpreloaded oocytes. Moreover, glutarate preloading had no effect on water-injected oocytes (data not shown). Thus, glutarate induced trans-stimulation of hOAT1-mediated PAH uptake, suggesting that the uptake occurs in conjuction with the countertransport of dicarboxylate.

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Figure 7

trans-Stimulation of rROAT1- and hOAT1-mediated uptake. rROAT1 or hOAT1 cRNA-injected oocytes, either nonpreloaded or preloaded by 90 min of incubation in 5 mM glutarate, were washed with glutarate-free medium and exposed to 50 μM [³H]PAH for 60 min in the absence (No Inhibitor) or presence (+ Inhibitor) of 1 mM probenecid. There was no effect on similarly treated water-injected oocytes (data not shown). Data are mean ± S.E. values from two animals (10 oocytes/treatment/animal). ***p ≤ .001, as determined by paired Student’s t test.

Nucleoside Phosphonate Analogs Are High-Affinity Substrates for hOAT1.

Finally, the ability of rROAT1 and hOAT1 to mediate the transport of therapeutically important antiviral nucleoside phosphonates was tested. Together with adefovir (10 μM) and cidofovir (36 μM), two other nucleoside phosphonates with broad-spectrum antiviral and antiproliferative activity, PMEG (10 μM) and PMEDAP (10 μM), were found to be substrates for both rROAT1 and hOAT1 (Fig.8). However, uptake of all antivirals tested was markedly higher for hOAT1 compared with rROAT1. Little to no uptake was measured in water-injected oocytes, indicating the complete lack of diffusive uptake of these nucleotide analogs. Like PAH, both hOAT1- and rROAT1-mediated uptake of all nucleoside phosphonates was efficiently blocked by 1 mM probenecid. In contrast to hOAT1 and rROAT1, the renal organic cation transporter 2, did not mediate detectable uptake of adefovir or cidofovir in renal organic cation transporter 2 cRNA-injected oocytes (data not shown). Finally, similar to PAH, preloading the oocytes in 5 mM glutarate for 90 min before exposure to adefovir significantly stimulated adefovir uptake in rROAT1 (5.3-fold increase, p < .001) and hOAT1 (1.5-fold increase, p < .01)-expressing oocytes, as compared with nonpreloaded oocytes. Thus, hOAT1-mediated adefovir uptake correlates with the countertransport of dicarboxylate.

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Figure 8

Uptake of antiviral nucleoside phosphonates mediated by hOAT1 and rROAT1. Xenopus oocytes were injected with hOAT1 or rROAT1 cRNA, and substrate accumulation was determined in the absence (No inhibitor) or presence (+ Inhibitor) of 1 mM probenecid. Substrate concentrations were as follows: 10 μM adefovir, 36 μM cidofovir, 10 μM PMEG, and 10 μM PMEDAP. Data are mean ± S.E. values from representative animals (10 oocytes/treatment/animal).

To find out whether the marked dissimilarity between rROAT1 and hOAT1 transport of nucleoside phosphonates is simply due to different levels of expression of the two transporters or actually due to lower substrate affinity of rROAT1, K _m values for cidofovir and adefovir were determined. Adefovir and cidofovir uptake by hOAT1-expressing oocytes increased steadily with time and was linear for about 1 h (data not shown). Using a 20-min incubation period to approximate initial rate, the transport was assessed by incubating cRNA-injected oocytes in medium containing 0.002 to 1 mM substrate concentrations. Double reciprocal plots of the saturation data were constructed, and linear regression analysis was performed to obtainK _m estimates (Fig. 9). Adefovir and cidofovir exhibited K _m values of 30 and 46 μM, respectively, for hOAT1-mediated transport (Table2). However, the correspondingK _m values for rROAT1-mediated transport were 5- and 9-fold higher, respectively, compared with hOAT1. A significant difference in K _m values between the two transporters was also observed for PAH (Table 2).

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Figure 9

Kinetic analysis of hOAT1- and rROAT1-mediated transport of cidofovir. hOAT1 and rROAT1 cRNA-injected oocytes were exposed to increasing substrate concentrations (0.002 to 1 mM depending on substrate and transporter) for 20 min. Saturation analysis was performed, and a representative double reciprocal plot with linear regression analysis is shown. The determined K _mvalues for all of the substrates tested are reported in Table 2. The data shown are mean ± S.E. values from three to four animals (10 oocytes/treatment/animal).