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GASTROINTESTINAL, HEPATIC, PULMONARY, AND RENAL

Genetic Variants of the Human H⁺/Dipeptide Transporter PEPT2: Analysis of Haplotype Functions

Department of Pharmacology Program in Pharmacogenomics, College of Medicine and Public Health, The Ohio State University, Columbus, Ohio (J.P., W.S.); and Molecular Biopharmaceutics, Department of Pharmaceutics, Danish University of Pharmaceutical Sciences, Copenhagen, Denmark (C.U.N.)

Received June 24, 2004; accepted July 27, 2004.

	Abstract

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PEPT2 is a high-affinity H⁺/dipeptide transporter expressed in kidney, brain, lung, and mammary gland. The physiological role of PEPT2 in kidney is to reabsorb small peptides generated by luminal peptidases. PEPT2 is also a transporter for peptide-like drugs such as penicillins and cephalosporins. We have conducted a haplotype analysis of 27 single nucleotide polymorphisms located in or near exons of the human gene encoding hPEPT2 (SLC15A2), using genotyping data from 247 genomic DNA samples from the Coriell collection. Our analysis reveals that hPEPT2 has a >6-kilobase sequence block with at least 10 abundant polymorphisms in almost complete linkage disequilibrium. As a result, only two main hPEPT2 variants exist (hPEPT2*1 and *2) with several phased amino acid substitutions, present in substantial frequencies in all ethnic groups tested. When expressed in Chinese hamster ovary cells, hPEPT2*1 and *2 displayed similar V_max values for glycyl-sarcosine (Gly-Sar), but they differed significantly in their K_m values (83 ± 16 and 233 ± 38 µM, respectively). Moreover, hPEPT2*1 and *2 differed in their pH sensitivity for H⁺/Gly-Sar transport. In addition, hPEPT2*1 and *2 generated varying levels of mRNA in nine heterozygous kidney tissue samples, including one allele expressing no detectable mRNA, suggesting the presence of cis-acting polymorphisms affecting transcription or mRNA processing. The results indicate that polymorphisms in the gene encoding hPEPT2 can alter substrate transport and therefore could affect drug disposition in vivo.

The physiological role of PEPT2 in kidney is to reabsorb small peptides left behind by luminal peptidases. It has been suggested that PEPT2 may be involved in drug disposition in the kidneys via reabsorption of -lactam antibiotics (Inui et al., 2000a,b). Reabsorption of some cephalosporins, glycyl-sarcosine (Gly-Sar) and Gly-Sar-Sar into the kidneys from the ultrafiltrate has been demonstrated in vivo (Arvidsson et al., 1979; Garrigues et al., 1991; Minami et al., 1992). Brain tissues expressing hPEPT2 include astrocytes, ependymal cells, and choroid plexus epithelial cells (Berger and Hediger, 1999). In the central nervous system, hPEPT2 may play a role in neuropeptide signaling. Neuropeptide signaling is terminated by extracellular peptidases that breakdown peptides into smaller fragments that are then removed by oligopeptide transporters. Based on PEPT2 substrate requirements and localization, it has been suggested that PEPT2 may be involved in the removal of neuropeptide fragments from the brain and in the regulation of neuropeptide levels in the cerebrospinal fluid (Nielsen et al., 2002). PEPT2 is present at the apical membrane of choroidal plexus epithelial cells and may function in the efflux of peptides across the blood-brain barrier from cerebrospinal fluid to blood (Shu et al., 2002). In mammary gland epithelia, PEPT2 is thought to be involved in the reuptake of small peptides that accumulate from the hydrolysis of milk proteins (Groneberg et al., 2002). PEPT2 is located in alveolar type II pneumocytes, bronchial epithelium, and endothelium of small vessels (Groneberg et al., 2001) and thought to be responsible for uptake of di- and tripeptides in the lung (Groneberg et al., 2001). Moreover, PEPT2 transports a number of drugs such as penicillins, cephalosporins, and angiotensin-converting enzyme inhibitors (Daniel and Adibi, 1993; Ganapathy et al., 1995, 1997, 1998; Zhu et al., 2000), and it may be responsible for the relatively low levels of these drugs detected in breast milk of lactating women who are taking them.

PEPT2 (SLC15A2, solute carrier 15A2) is 729 amino acids in size with 12 proposed transmembrane spanning regions, and a large extracellular loop between transmembrane domains 9 and 10 (Fig. 1). Located on chromosome 3 at 3q21.1, hPEPT2 contains 22 exons spread over 47 kilobases (Fig. 2). When the gene was cloned in 1995 by screening a human cDNA kidney library with a rabbit PEPT2 cDNA (Liu et al., 1995), a single transcript was identified and accepted as the consensus sequence. Numerous single nucleotide polymorphisms (SNPs) have been detected in the SLC15A2 gene (see PharmGKB, http://www.pharmgkb.org/views/ and http://pharmacogenetics.ucsf.edu/). The functional significance of these polymorphisms remains unknown, but recently, a rare nonsynonymous polymorphism (R57H) has been shown to disrupt PEPT2 function (Terada et al., 2004).

View larger version (30K): [in this window] [in a new window]   Fig. 1. Predicted hPEPT2 secondary structure. There are 12 transmembrane domains and a large extracellular domain located between transmembrane domains 8 and 9. This image was modified from an image generated using TOPO, a transmembrane protein display program, by S. J. Johns (University of California, San Francisco) and R. C. Speth (Washington State University). The three most abundant nonsynonymous amino acid changes are shaded black.

View larger version (7K): [in this window] [in a new window]   Fig. 2. The genomic structure of SLC15A2, the gene encoding the H+/peptide transporter PEPT2.

A haplotype is the combination of several sequence variants on a single chromosome at a specific locus. Haplotypes may more accurately reflect genetic diversity in the human population. Here, using haplotype analysis, we show that hPEPT2 exists in multiple variants and that two of these variants (hPEPT2*1 and *2) are significantly present in all ethnic groups we have tested. In vitro analysis indicates that the *1 and *2 protein variants differ in functional properties.

Genetic variations can affect protein structure on the one hand and mRNA processing, mRNA stability, and cis-regulatory mechanisms on the other. The latter variations have one common outcome: they affect the amount and nature of the mRNA generated from one allele versus another. To determine whether such cis-acting polymorphisms exist in hPEPT2, we have measured allele-specific mRNA expression of hPEPT2 in heterozygous samples from kidney tissues. Together, the results indicate significant functional variations are present in hPEPT2. This work provides a basis for exploring the pharmacological relevance of genetic variability.

	Materials and Methods

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Genotyping
Genomic DNA samples were obtained from the Coriell Institute (Camden, NJ). Primers for polymerase chain reaction (PCR) were synthesized to amplify each exon and a minimum of 35 bases of 5' and 3' flanking sequence. Sequencing and genotyping of the Coriell samples was performed in the genetics core facility at the University of California, San Francisco (San Francisco, CA). Primers for exons and adjoining intron regions (ca. 50 base pairs) for hPEPT2 (National Center for Biotechnology Information reference sequence NM 021082.1) were designed using the Virtual Genome center Web site at http://alces.med.umn.edu/VGC.html and ordered from Operon (Alameda, CA). A collection of 247 ethnically identified genomic DNA samples was used to screen for hPEPT2 variants. PCR was performed in split 20-µl reactions using TaqGold (PerkinElmer Life and Analytical Sciences, Boston, MA) on the GeneAmp 9700 thermocycler (PerkinElmer Life and Analytical Sciences). One of three PCR protocols was chosen based on an optimization performed before amplifying all 247 samples. Occasionally, it was necessary to add glycerol or dimethyl sulfoxide or both to a reaction to optimize it. Agarose gel electrophoresis (2%, 150 V, 35 min) was performed to verify the PCR products. Samples were pooled 3-fold, and denaturing high-pressure liquid chromatography (HPLC) was performed on Varian HPLC machines with Varian Helix columns (Varian, Palo Alto, CA) to detect the presence of sequence variants. The denaturing HPLC melt program described at http://insertion.stanford.edu/melt. html was used to approximate optimal temperature conditions for each run. Typically, the highest recommended temperature was run along with at least one lower temperature, depending on the complexity of the melt profile. The results were scored manually. If a well scored positive, the three PCRs corresponding to that well were cleaned with 2 units of shrimp alkaline phosphatase and Exonuclease I from USB (Cleveland, OH) and sequenced using ABI BigDye version 2 (Applied Biosystems, Foster City, CA), cleaned with 96-well gel filtration blocks from Edge BioSystems (Gaithersburg, MD), and run on an Applied Biosystems 3700 DNA analyzer. Sequences were scored in Sequencher version 4. All singleton SNPs were verified with a fresh PCR reaction and sequence.

Haplotype Analysis
Genotype data of the 14 most common SNPs from all 247 samples were converted into "A" or "B". AA is the homozygous major allele sequence, AB is heterozygous, and BB represents the homozygous minor allele. The data were formatted for Fallin and Schork's SNPEM.2001 program (Fallin and Schork, 2000; Fallin et al., 2001), which was compiled on a LINUX operating system. Up to 10 SNPs were run at a time over several runs, for example SNPs 5a, 9, 12c, 13b, 13c, 15, 17b, 20, 21a, and 22. The numbers in the SNP names correspond to exon position, and the lowercase letter corresponds to SNP number in the exon (Table 1). In exons with only a single SNP, no letter is specified. For further information, refer to the SNP description of hPEPT2 genotyping data at http://pharmacogenetic-s.ucsf.edu/.

Construction of hPEPT2*2 Variant
A 1.2-kilobase (kb) fragment of hPEPT2 was amplified by polymerase chain reaction from human cerebellum QUICK-Clone cDNA (BD Biosciences Clontech, Palo Alto, CA) (forward primer E7F, GAGAAGACTGCTATGCATTGG; reverse primer E20R, ACCTGTGACAGAGAACATGACCTC). The Quick-Clone cDNA was made from the cerebella of 24 pooled male and female Caucasians. The collection of PCR fragments, representing multiple hPEPT2 variants, was ligated into the SmaI site of pBluescript (Stratagene, La Jolla, CA). Several DNA minipreps were screened by sequencing to obtain a 1.2-kb fragment of the hPEPT2*2 variant. The NsiI/XbaI fragment of hPEPT2*2 was cut out and ligated into the pTLN2 vector containing the hPEPT2*1 variant, obtained from M. Hediger (Harvard University, Cambridge, MA), with its NsiI and XbaI fragment removed. The final construct was verified by sequence analysis.

Identification of Partial Chimpanzee Sequence
Primers used to amplify a 1.2-kb fragment from human cDNA (same as described above) were used to amplify the same fragment in a first strand synthesis cDNA library made from chimpanzee (Pan troglodyte) mRNA (Lena 10-77). Chimpanzee RNA was kindly provided by Daniel J. Birmingham (The Ohio State University, Columbus, OH). The amplified fragment was sequenced using internal human gene specific primers.

Sequence Alignments
Alignments were conducted online using the ClustalW algorithm. Input sequences were in FASTA format. Accession number for sequences are as follows: reference sequence (human hPEPT2*1) NM_021082 [GenBank] ; hPEPT2*2 XM00292; rhesus monkey (Macaca mulatto) AAQ54588 [GenBank] and chimpanzee (P. troglodytes hPEPT2) genomic sequence from Washington University sequencing center.

Cell Culture, Transfection, and Uptake Experiments
CHO cells were cultured under standard conditions. The cells were seeded onto 12-well plates and transfected once they reached approximately 50% confluence. pcDNA 3.1 containing wild-type hPEPT2 and its variant were transfected into CHO cells using Jet-PEI (Qbiogene, Carlsbad, CA). Twenty-four hours after transfection, the cell monolayers were used for experiments.

Uptake Experiments. Cells were grown on TC-treated 12-well plates (Corning, Acton, MA). Uptake of [¹⁴C]Gly-Sar (PerkinElmer Life and Analytical Sciences) with a specific activity 49.9 mCi ± mmol^–1) was measured in Hanks' balanced salt solution supplemented with 0.05% bovine serum albumin and buffered with 10 mM 2-[N-morpholino] ethanesulfonic acid (MES) with pH adjusted to 6.0. Cells were placed on a shaking plate, preheated to 37°C, and allowed to equilibrate for 10 min in the buffer solution. The experiment was started by adding 0.5 ml of fresh buffer per well containing Gly-Sar (total concentration from 40 to 1000 µM), including 2 µCi/ml [¹⁴C]Gly-Sar, and 1 µCi/ml [³H]mannitol (specific activity 51.5 mCi ± mmol^–1; PerkinElmer Life and Analytical Sciences) as a marker of extracellular space. The uptake of Gly-Sar in transfected cells was linear for more than 30 min, tested at extracellular Gly-Sar concentrations of 40 and 1000 µM. Uptake experiments were terminated after 30 min by removing the uptake medium, followed by three washes of the monolayers with ice-cold Hanks' balanced salt solution. The cells were detached from the well with 0.1% Triton X (polyethylene glycol tert-octylphenyl ether) in phosphate-buffered saline. Radioactivity was determined in a liquid scintillation counter. Experiments were performed using four to nine individual transfections.

pH-Dependent Uptake. We investigated the pH dependence of the Gly-Sar transport via the two PEPT2 variants. To ensure that the intracellular pH of the CHO cells was not determined by the extracellular pH and to eliminate the proton-motive force across the cell membrane, we cultured the CHO cells on permeable polycarbonate filter support with a cell growth area of 1.1 cm². This allowed us to incubate the cells with a pH 7.4 buffer before the experiments and to change pH on the donor side, while maintaining the pH of 7.4 on the trans-side, and thus minimizing the rate of intracellular pH change caused by the buffer. The lower chamber contained 1 ml of HEPES buffer [N-(2-hydroxyethyl)piperazine-N'-(2-ethanesulfonic acid); 4-(2-hydroxyethyl)piperazine-1-ethanesulfonic acid] (pH 7.4), and the upper chamber 0.5 ml of 40 µM [¹⁴C]Gly-Sar solution in 10 mM MES buffer of varying pH values. Uptake into the cells was measured over a 5-min period. Experiments were performed with six individual transfections, all occurring after seeding. To validate the method we took samples from the trans-side, and here we did not detect any [¹⁴C]Gly-Sar counts, indicating that the donor and trans-solutions did not mix during the experiment.

Affinity Studies. The affinity experiment was initiated by adding 0.5 ml of MES buffer (pH 6.0) containing 2.0 µCi/ml [¹⁴C]Gly-Sar (40 µM) and various amounts of unlabeled test compounds to the transfected CHO cells. From this point on, the affinity experiment was similar to the Gly-Sar uptake experiment.

Kinetic Analysis
Concentration-Dependent Gly-Sar Uptake. Uptake of Gly-Sar in transfected CHO cells was corrected for noncellular uptake using mannitol as described previously (Bravo et al., 2004), and the cellular uptake as a function of apical Gly-Sar concentration was fitted to a Michaelis-Menten type equation (eq. 1):

(1)

where V is uptake rate (pmol x cm^–2 x min^–1), V_max is maximum uptake rate (pmol x cm^–2 x min^–1), K_m is the Michaelis-Menten constant (µM), [S] is Gly-Sar concentration (µM), and n is Hill factor. The Hill factor was between 0.94 and 0.96.

Affinity Experiments. Affinity for hPEPT2 in transfected CHO cells was determined from inhibition curves of 40 µM [¹⁴C]Gly-Sar uptake in the presence of varying concentrations of test compound. The degree of inhibition was fitted to a Michaelis-Menten type equation (eq. 2):

(2)

where U is uptake of [¹⁴C]Gly-Sar, U₀ is uptake of [¹⁴C]Gly-Sar at zero inhibitor concentration, IC₅₀ is the concentration required to inhibit the maximum uptake by 50% (µM), [I] is compound concentration (µM), and n is Hill factor. The Hill factor was between 0.96 and 1.14. K_i values were calculated as described by Cheng and Prusoff (1973) using the K_m values for Gly-Sar obtained for the two variants.

Kidney Tissue Preparation for Allele-Specific Assays
Genomic DNA and RNA were prepared from frozen kidney and tissue autopsy samples (obtained from The Ohio State University tissue bank). Frozen tissue samples were pulverized under liquid nitrogen. The frozen powder was portioned into aliquots for DNA and RNA extractions. DNA was prepared by digestion of the pellet or frozen powder with sodium dodecyl sulfate and proteinase K followed by sodium chloride precipitation of proteins (Miller et al., 1988). The DNA in the supernatant was further purified and recovered by ethanol precipitation. For RNA preparation, the starting material was homogenized in TRIzol reagent (monophasic solution of phenol and guanidine isothiocyanate) (Invitrogen, Carlsbad, CA), and extracted with chloroform. The RNA was recovered by precipitation with isopropanol followed by centrifugation. For additional purification, the RNA precipitate was dissolved in RNase-free water or QIAGEN buffer (QIAGEN, Valenia, CA), and then extracted using QIAGEN RNeasy columns according to the manufacturers instructions. Complimentary DNA (cDNA) was generated from the mRNA by Superscript II reverse transcriptase (Invitrogen) using oligo dT as a primer, to selectively bind to the polyA tail of mRNA. This ensured that all of the gene transcripts were represented at a rate that reflected the original abundance of each gene product.

SNaPshot Assay and Quantitative Analysis of Allelic Ratios in Genomic DNA and mRNA
This is a primer extension method originally intended for use with the ABI 3100 sequencer. We have successfully adapted this genotyping assay for the ABI 3730, with a few modifications that were needed to accommodate data handling. A stretch of genomic DNA (70 base pairs) or cDNA was amplified by PCR, and the allelic ratio was measured by primer extension using fluorescently labeled terminator nucleotides. We assume that this amplifies both alleles equally. Genomic DNA serves as an internal control compared with amplification of cDNA of the same samples. Standard PCR conditions were used on 15-µl reactions. Amplification conditions consisted of 35 cycles of denaturation at 95°C for 30 s, and then primer annealing at 60°C for 1 min, followed by extension at 72°C for 1 min (hPEPT2 exon 14 amplification primer pairs were forward primer AGGAAAATGGCTGTTGGTATGATC and reverse primer CGCAACTGCAAATGCCAG) After amplification, the reactions were treated with exonuclease I and bacterial Antarctic alkaline phosphatase (New England Biolabs, Beverly, MA). For the primer extension, a gene-specific primer was designed with its 3'-end one base from the SNP position (hPEPT2 exon 14 extension primer GCTGTTGGTATGATCCTAGC). Primers used to analyze the SNP in exon 15 were forward primer GAAATGGCCCCAGCCC, reverse primer CATCTGCCAGATTCAAGACTTGTAG, and extension primer AACCTCCTGGGGACCTG. SnaPshot reagent from Applied Biosystems was used to incorporate a single fluorescently labeled dideoxynucleotide to the 3' end of the primer in a template dependent manner. The final primer extension reactions were analyzed using an ABI 3730 capillary electrophoresis DNA instrument, and calculated with Gene Mapper 3.0 (Applied Biosystems) software. The data for each incorporated fluorescently labeled nucleotide was measured as a peak area, which is proportional to the amount of amplified allele. Heterozygous samples have two differently labeled peaks at approximately the same position. Since differing fluorophores may influence nucleotide incorporation and migration rates, the peak areas are not identical between two alleles present in equal abundance. Therefore, peak area ratios of genomic DNAs assumed to be present in equal amounts (ratio = 1) were used to normalize average genomic and cDNA ratios of heterozygous samples. The mRNA results in Fig. 5b are averages of four separate experiments, normalized to genomic DNA. Ratios are derived by dividing peak areas of each of the two alleles of hPEPT2 (B/A) where *1 is the B allele and *2 is the A allele. Deviations from unity in the normalized peak area ratios between alleles in the cDNA are attributed to differences in allele-specific mRNA levels.

View larger version (26K): [in this window] [in a new window]   Fig. 5. Allele-specific mRNA analysis of Pept2 mRNA from heterozygous kidney samples. a, allele-specific quantification of DNA and mRNA from heterozygous samples. Genomic DNA is shown for one sample K009, but it is representative of all of the genomic DNA samples. Allele-specific levels in cDNA in sample K020 are not significantly different from DNA; however, levels in K009 deviate significantly. B, allelic ratios showing *1/*2 (or B/A) are calculated for genomic DNA and RNA. A ratio of 1 indicates that both alleles are expressed equally. Ratios greater than 1 indicate that the *1 allele is expressed more than *2. Ratios under 1 indicate that the *2 allele is expressed more. RNA samples with significant allelic differences are marked (#).

	Results

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Genotype Analysis and Identification of Variants. Samples (247) of human genomic DNA from different ethnic groups were genotyped. The samples include 100 Caucasian, 100 African American, 30 Asian, 10 Mexican, and 7 Pacific Islander individuals. Samples were genotyped over all 22 exons of hPEPT2, and 27 SNPs were detected. The genotyping results are summarized in Table 1. Of the 27 SNPs, eight were in coding regions with four SNPs being synonymous and four nonsynonymous.

SNP Frequencies. Seventeen SNPs occurred with frequencies greater than 1%, 14 SNPs were detected with allele frequencies greater than 7%, and 10 of these had frequencies of 48 to 49%. Of the latter, three were nonsynonymous with a frequency of 49%. There are some notable ethnic differences with respect to genotype frequency of some SNPs. For example, the synonymous SNP in exon 9 ranges from 36.7% in Asians up to a frequency of 71.4% in Pacific Islanders, but all of the abundant SNPs are present in all ethnic groups we have tested. Frequencies and occurrence of the rare SNPs varied between ethnic groups. For example the second SNP in exon 21 is present in 3% of Asians and 1.8% of Caucasians, but it is missing in the African American, Mexican, and Pacific Islander populations tested.

The abundant nonsynonymous SNPs affect the following amino acids: F350L (exon 13, rs2257212), located in transmembrane domain 8 near the cytoplasmic side of the membrane; and S409P (exon 15, rs1143671) and K509R (exon 17, rs1143672), both located in the large extracellular domain. All three of these amino acids are indicated by black shading in Fig. 1.

Haplotype. Haplotypes were inferred using the estimation maximization algorithm (SNPEM.2001) (Fallin and Schork, 2000; Fallin et al., 2001). Fourteen SNPs with frequencies over 7% were included with the analysis. For simplicity, all of the genotype data for each SNP in every sample were converted to A or B, where A usually represented the major allele, but in some cases the designation was arbitrary. Figure 3 depicts predicted haplotypes with frequencies over 1%. The haplotypes are grouped by similarity. The haplotype with the highest frequency (number 1) has the allele designation of A in all 14 SNPs. Since its frequency (38%) is much larger than that of other haplotypes, we considered haplotype 1 to be the reference. However, haplotype 1 does not represent the hPEPT2 sequence cloned by Liu et al. (Liu et al., 1995), which we have termed here hPEPT2*1, because it was published first while not representing the most abundant haplotype variant.

View larger version (35K): [in this window] [in a new window]   Fig. 3. Summary of the haplotype data for the 14 most abundant SNPs with frequencies above 7%. Each column, beginning with the one labeled 5a represents a SNP. The top row specifies the exon (number) and position of the SNP (lowercase letter if more than one per exon was identified; see Table 1). The second row specifies the base pair change for each SNP. The letter on the left is the reference sequence and the letter on the right is the variant. The third row from the top indicates whether the SNP is intronic or if coding and nonsynonymous, the amino acid change caused by the SNP. The row at the bottom lists the genotype frequencies obtained for each SNP. The first column on the left identifies each haplotype with a number, 1 through 12. The second column from the left indicates which protein variant, *1, *2, or *3, the haplotype encodes. The column on the far right lists the haplotype frequencies obtained from the SNPEM.2001 algorithm. Frequencies in bold text indicate haplotypes that are high frequency. The middle blocks represent each SNP. White blocks labeled A are the reference sequence. Gray blocks labeled B are variants.

Several patterns emerge in Fig. 3. The 10 SNPs with frequencies over 49% are in near complete linkage disequilibrium and form the core of several related haplotypes. The core haplotype block begins at exon 9 and continues through the middle of exon 17, spanning at least 6500 base pairs of genomic DNA. These haplotypes include the three most abundant nonsynonymous SNPs— each allele present in nearly equal abundance—that affect amino acid positions 350, 409, and 509; two synonymous SNPs at amino acid positions 284 (exon 9, rs2293616) and 387 (exon 14, rs1143670); and at least five additional SNPs located in introns. Haplotypes 7 through 12 contain B alleles in all 10 SNPs within the 6.5-kb block. Together, they constitute one variant: hPEPT2*1. The hPEPT2*1 protein contains the amino acids L350, P409, and R509, as cloned by Liu et al. (1995). Haplotypes 1 through 5 are closely related, containing the reference genotype in all 10 SNPs within the 6.5-kb block and comprising the second variant hPEPT2*2. The hPEPT2*2 protein contains amino acids F350, S409, and K509. Both the *1 and *2 variants are distributed roughly equally in the population at about 44 to 47%. There is a third protein sequence variant represented with lower abundance (1.6%) (haplotype 6) termed hPEPT2*3. It is the only haplotype where the three abundant nonsynonymous SNPs are not in linkage disequilibrium.

Conservation of Human Coding Region Polymorphisms across Primate Sequences. We found five common SNPs that differ in the cDNA of the three human PEPT2 variants. The approximately even distribution of the hPEPT2*1 and hPEPT2*2 alleles in the population suggests that the two alleles and their corresponding core haplotypes are ancient, raising the question how these two variants have evolved, carrying multiple-phased SNPs. We compared the cDNA of the three human variants with cDNA from chimpanzee (P. troglodytes) and rhesus monkey (M. mulatto) (Table 2). There were eight single base-pair differences in the coding region between the chimpanzee and human sequences: the five SNPs that differ between the human variants and three additional chimpanzee-specific SNPs. Two of the three in common nonsynonymous SNPs match the hPEPT2*2 variant (positions 409 and 509) and the other SNP (350) matches the hPEPT2*1 variant. Hence, the chimpanzee sequence resembles the human hPEPT2*3 variant.

The cDNA from rhesus monkey had 38 base-pair changes that were not found in either human or chimpanzee sequence, but overall the cDNA was 98.3% identical. Using the eight nucleotides that differ between chimpanzee and human (Table 2), a comparison of the five primate sequences allowed the construction of an ancestral primate PEPT2 allele, which resembles neither the *1 or *2 variant. SNPs in exon 14 (Syn) and 15 (S409P) most likely diverged from the ancestral allele before chimpanzees and humans diverged, generating a haplotype that encodes *3. The exon 13 SNP, found only in the *2 variant, originated later, as did the SNPs in exon 9 and exon 17, which are specific to the *1 variant. Overall, the *1 variant is more similar to the ancestral primate allele, whereas the *2 variant is more similar to the chimpanzee sequence.

Functional Analysis of the hPEPT2*1 and *2 Variants. To test for the presence of functional differences between the two main protein variants hPEPT2*1 and *2, the cDNAs for both variants were subcloned into pcDNA3.1 expression vector and transiently transfected into CHO cells. Analysis of Gly-Sar uptake in transfected CHO cells (Fig. 4a) showed that the two variants had similar V_max values (hPEPT2*1, 0.96 ± 0.07 = pmol ± cm^–2 ± min^–1; hPEPT2*2, 1.11 ± 0.07 pmol ± cm^–2 ± min^–1. The *1 variant had a K_m of 83 ± 16 µM, whereas the *2 variant had a significantly higher K_m value of 233 ± 38 µM (p < 0.05; n = 4–9). Since the transport of substrates by hPEPT2 is partly driven by the cotransport of protons, we investigated the effect of extracellular pH on the transport of Gly-Sar. For hPEPT2*1, the highest uptake was seen at pH 6.0 (Fig. 4b), whereas for hPEPT2*2 uptake at pH 6.0 was significantly lower (p < 0.01; n = 6–8). At the other extracellular pH values investigated no difference in uptake was observed. The affinities of cephalexin and kyotorphin for hPEPT2 were also investigated in CHO cells transfected with hPEPT2*1 and *2 (Fig. 4c). The K_i values obtained for *1 and *2 were similar for both cephalexin (98 ± 23 and 102 ± 3 µM, respectively) and kyotorphin (5.1 ± 0.3 and 6.0 ± 1.2 µM, respectively).

View larger version (18K): [in this window] [in a new window]   Fig. 4. a, Gly-Sar concentration-dependent effect of apical uptake of [14C]Gly-Sar in CHO cells transfected with hPEPT2*1 () or hPEPT2*2 (). The apical uptake of Gly-Sar was measured over a 30-min incubation period. The lines describe the best fit to the experimentally obtained points using the Michaelis-Menten equation. Each data point represents the mean ± S.E. of four to nine individual transfections. hPEPT2*1: Km = 82.9 ± 16, Vmax = 0.96 ± 0.07 pmol/(cm2 x min), n = 1.01 ± 0.16. hPEPT2*2: Km = 233 ± 38, Vmax = 1.11 ± 0.07 pmol/(cm2 x min), n = 0.94 ± 0.07. b, pH dependence of Gly-Sar uptake. CHO cells were grown on filter support. The lower chamber contained a HEPES buffer (pH 7.4) solution, and the upper chamber had a 40 µM Gly-Sar solution in MES buffer with varying pH values. Uptake was measured over a 5-min period. Each point represents average ± S.E. of three to nine individual transfections. c, inhibition of Gly-Sar uptake. The uptake of 40 µM Gly-Sar in MES-buffer at an extracellular pH of 6.0 was measured in the presence of varying amount of kyotorphin and cephalexin in transfected CHO cells. The lines describe the best fit to the experimentally obtained points using eq. 2. Each data point represents the mean ± S.E. of six individual transfections.

Allele-Specific mRNA Analysis in Kidney Tissue. To test whether the two variants of hPEPT2 are associated with differences in mRNA expression in their native tissue, we measured the quantity of each allele in heterozygous kidney samples using SNaPshot and compared them to each other. Figure 5a is an example of allelic peaks obtained for one DNA and two RNA samples. Allelic ratios of the synonymous SNP in exon 14 (rs1143670) and the nonsynonymous SNP in exon 15 (rs1143671) of hPEPT2 were tested separately in DNA and mRNA from heterozygous kidney samples. Of 20 kidney samples analyzed, nine were heterozygous for the *1 and *2 SNPs. Allele-specific measurements were precise and highly reproducible for the exon 14 SNP in genomic DNA, deviating from unity with a S.D. of only 4% (Fig. 5b). In contrast, significant deviations from unity were observed for allelic mRNA ratios (Fig. 5b). We also measured allelic ratios in DNA and mRNA using the SNP in exon 15 (data not shown). Whereas the variability of the results was somewhat greater than with the exon 14 SNP, the results were comparable, confirming the validity of the allele-specific analysis. In particular, only the B allele (representing variant*2) was detectable in mRNA of the heterozygous sample K018; this was confirmed with both SNPs located in exons 14 and 15, in independent experiments. Complete absence of one allele suggest the presence of a mutation in the regulatory region, or one that is affecting mRNA processing (e.g., splicing), or gene silencing by epigenetic effects. We found three samples: K004 (B/A ratio 1.18 ± 0.03), K005 (1.28 ± 0.03), and K009 (1.52 ± 0.22) where the *1 allele mRNA was significantly higher. The other five kidney samples did not exhibit significant allele-specific differences in gene expression

	Discussion

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A genetic and haplotype analysis of the gene encoding hPEPT2, SLC15A2, revealed the presence of two distinct, common variants that are present at a high frequency in all of the ethnic groups tested. The gene contains a block of 10 polymorphisms in a 6-kb region, including three nonsynonymous SNPs. All of the SNPs in this block have the same allele frequency and are in near complete linkage disequilibrium with each other. There are additional genomic SNPs outside of this block, which increases the number of haplotypes. The two common haplotype groups encode two protein variants, termed hPEPT2*1 and hPEPT2*2, that differ by three amino acids. hPEPT2*3 is a less frequent variant with a slightly different pattern of nonsynonymous SNPs.

The presence of several linked SNPs with similar allele frequencies that form a haplotype block with only two major variants, suggested that the haplotype block is ancient and highly resistant to crossover events. It also suggested the possibility that all of these polymorphisms arose together. To examine the evolution of the haplotypes, we compared the cDNA sequence of the three human PEPT2 variants with cDNA from chimpanzee and rhesus monkey. Humans and chimpanzees diverged from each other approximately 3.5 to 5.5 million years ago (Sarich and Cronin, 1976), and the genomes are 95% identical (Britten, 2002). In contrast, rhesus monkeys diverged from other primates approximately 50 million years ago (Martin, 1993), consistent with 38 additional base pair differences in the rhesus sequence absent in the chimpanzee or human sequences. We inferred an ancestral primate PEPT2 allele from sequence comparisons showing that the *1 variant more closely resembles the ancestral primate allele, whereas the *2 variant is more similar to the chimpanzee sequence. The comparison also indicated that the block of polymorphisms did not arise at the same time. However, if the SNPs arose independently, we would have expected to find multiple transitional haplotypes in the human population, which is not the case. The rare *3 variant encoded by haplotype 6 does fit the description of a transitional haplotype. Considering that the history of human evolution is full of bottle necks and dead ends, perhaps there are no additional human haplotypes in the PEPT2 block because only families carrying the three current hPEPT2 alleles survived.

Our in vitro experiments suggest that the two main variants differ significantly in biochemical properties. The Gly-Sar K_m value for the *1 variant (83 ± 16 µM) is similar to the K_m values previously obtained for hPEPT2 and rPEPT2 (from rat). The K_m for Gly-Sar uptake via hPEPT2 has previously been reported to be 74 and 110 µM (Ramamoorthy et al., 1995; Fujita et al., 2004) and 90 µM for rPEPT2. (Bravo et al., 2004). However, the K_m value for the *2 variant (233 ± 38 µM) is significantly higher than for the *1 variant. Yet, the inhibitory potencies of cephalexin and kyotorphin on Gly-Sar uptake were similar for both variants. The inhibitory potency of cephalexin for both variants (98 ± 23 µM for *1 and 102 ± 3 µM for *2) are similar to reported values for rPEPT2 (63 and 73 µM) (Ganapathy et al., 1995; Brandsch et al., 1997). On the other hand, the affinity of kyotorphin for rPEPT2 has been reported to be 29 µM, which is higher than the values obtained in this study for hPEPT2 (5.1 ± 0.3 µM for *1 and 6.0 ± 1.2 µM for *2). The reasons for this discrepancy remain to be clarified. Because the two variants differed in their pH dependence, yielding disparate Gly-Sar K_m values, whereas the affinities for cephalexin and kyotorphin did not differ, it seems that the functional difference between the two variants stems from the proton-dependent part of the translocation cycle. Alternatively, cephalexin and kyotorphin may bind differently from Gly-Sar to hPEPT2. Further studies are needed to determine kinetic differences for drug substrates between the hPEPT2 variants.

The sequencing analysis had focused on coding regions of hPEPT2 and only portions of intronic splice regions and regulatory domains. Functional diversity is likely also to arise from polymorphisms that affect transcription or mRNA processing. However, responsible functional polymorphisms are difficult to identify. Therefore, we have developed an experimental approach to determine whether hPEPT2 harbors any such cis-acting polymorphisms. Allele-specific mRNA analysis in relevant target tissues is a potentially powerful tool for detecting cis-acting functional polymorphisms. To reflect the native regulatory environment of tissues where hPEPT2 is functionally expressed, this type of analysis must be performed on DNA and mRNA extracted from the target tissues. We have developed a primer extension assay capable of precisely measuring allelic differences in hPEPT2 mRNA levels in kidney tissues, using two indicator SNPS present in both genomic DNA and mRNA. Even though our sample was small (n = 9 kidneys heterozygous for the marker SNPs (of 20 kidney samples analyzed), significant allelic differences were observed in three samples. Moreover, only the hPEPT2*2 allele was expressed in a fourth heterozygous kidney sample, indicating the presence of a mutation in hPEPT2*1 that prevented mRNA accumulation. However, we cannot rule out gene imprinting or other epigenetic events. hPEPT2 allelic mRNA differences were not associated solely with one of the two main variants, suggesting that cis-acting polymorphisms affecting mRNA levels are present, but they are likely to be located in a separate haplotype block. Using allelic differences in mRNA expression, we can now begin to identify the mechanisms and functional polymorphisms underlying this observation. Given the magnitude of the expression differences, it is possible that a combination of allele-specific biochemical differences at the protein level in concert with cis-regulatory polymorphisms could exert strong effects on drug disposition of various hPEPT2 substrates in some individuals. Dose-dependent renal clearance of cefadroxil in healthy human males has been demonstrated, and the renal clearance was significantly increased in the presence of high doses of cephalexin (Garrigues et al., 1991). Although the contribution of hPEPT2 to the clearance was not specifically investigated, both cephadroxil and cephalexin are known substrates for hPEPT2. The cefadroxil plasma concentrations in this study were in micromolar range, which indicate that the transporter involved in the reabsorption of the compound is more likely to be the high-affinity transporter, hPEPT2, rather than hPEPT1. Increased urinary excretion of Gly-Sar-Sar in the presence of high amounts of the peptide has also been shown in perfused rat kidneys (Minami et al., 1992). In this study, the initial concentration of Gly-Sar-Sar in the perfusate was 100 µM, which again falls into the range of recognition by PEPT2 rather than PEPT1.

In summary, SLC15A22 encodes two main variants of the human di/tripeptide transporter hPEPT2 (*1 and *2) that differ in biochemical properties; moreover, cis-acting polymorphisms seem to affect expression levels in kidneys. Because both protein variants are highly abundant in all populations, it is likely that these variants have not been subject to selection pressures in evolution. However, the impact of hPEPT2 sequence variations on drug disposition could be substantial and needs to be studied further.

	Acknowledgements

 
We are grateful to Danielle Fallin (Johns Hopkins University, Baltimore, MD) for providing the SNPEM.2001 software; Audrey Papp for SNaPshot analysis and helpful discussions about the project; Gloria Smith for kidney sample DNA/RNA preparation; and Daniel J. Birmingham (The Ohio State University) for providing chimpanzee RNA. Maria Pedersen and Dr. Jan Amstrup (University of Copenhagen, Copenhagen, Denmark) have contributed to the functional analysis of the hPEPT2 variants.

	Footnotes

 
This work was in part supported by the University of California, San Francisco, Plasma Membrane Transporter Group (Grant GM61390 from General Medical Sciences, National Institutes of Health), by funds from The Ohio State University, and The Carlsberg Foundation (to C.U.N.).

doi:10.1124/jpet.104.073098.

ABBREVIATIONS: Gly-Sar, glycyl-sarcosine; SNP, single nucleotide polymorphism; kb, kilobase; PCR, polymerase chain reaction; HPLC, high-pressure liquid chromatography; CHO, Chinese hamster ovary; MES, 2-(N-morpholino)-ethanesulfonic acid.

Address correspondence to: Dr. Julia Pinsonneault, Department of Pharmacology, 333 West 10th Ave., The Ohio State University, Columbus OH 43210-1239. E-mail: pinsonneault.2{at}osu.edu

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