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ABSORPTION, DISTRIBUTION, METABOLISM, AND EXCRETION

Genetic Variability at the Human FMO1 Locus: Significance of a Basal Promoter Yin Yang 1 Element Polymorphism (FMO1^*6)

Department of Pediatrics, Birth Defects Research Center and Department of Pharmacology/Toxicology, Medical College of Wisconsin, Milwaukee, Wisconsin

Received for publication April 30, 2003
Accepted June 6, 2003.

	Abstract

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The flavin-containing monooxygenases (FMOs) are important for the disposition of a variety of toxicants, therapeutics, and dietary components. Although FMO1 is the dominant isoform in fetal liver and adult kidney and intestine and despite up to a 10-fold intersubject variation in expression, a paucity of information is available on FMO1 genetic variability. To address this issue, 24 samples from the Coriell DNA Polymorphism Discovery Resource Panel were sequenced revealing 10 common single nucleotide polymorphisms (SNPs): four located upstream of the structural gene; three within exonic sequences; one within the intron 1 splice donor site; and two with the 3'-untranslated region. Six of these variants are novel. Compared with other FMO loci within the chromosome 1q23-25 cluster, FMO1 seems more highly conserved. Of the identified FMO1 SNPs, only a C>A transversion 9,536 base pairs upstream of the exon 2 ATG start codon (g.-9,536C>A) would likely affect function, because it lies within the conserved core binding sequence for the yin yang 1 (YY1) transcription factor. Electrophoretic mobility shift assays demonstrated that the g.-9,536C>A transversion eliminated YY1 binding. Furthermore, data from transient expression assays in HepG2 cells suggested this SNP could account for a 2- to 3-fold loss of FMO1 promoter activity. Genotype analysis revealed a g.-9,536A allele (FMO1^*6) frequency of 13 and 11% in African- and northern European-Americans, respectively, but a significantly higher frequency of 30% in Hispanic-Americans. Thus, the FMO1^*6 variant may account for some of the observed interindividual variation in FMO1 expression.

FMO1-4 are encoded by distinct genes, oriented identically and localized in a single 254-kbp cluster on human chromosome 1q23-25. The gene encoding FMO5 is localized more centromeric at 1q21 [PDB] . A recent study reported a prevalent nonsense mutation in exon 3 of a sixth FMO gene, FMO6, within the 1q23-25 cluster (Furnes et al., 2003). These data, combined with the previous report from this laboratory demonstrating FMO6 alternative processing to transcripts incapable of translation to a functional enzyme (Hines et al., 2002), suggest FMO6 is a pseudogene in most if not all individuals (thus, FMO6P). A nonsense variant also was found to be prevalent in the FMO2 gene (Whetstine et al., 2000) and more recently, a frameshift polymorphism in FMO2 exon 4 (Furnes et al., 2003). These variants would suggest an active FMO2 enzyme, the predominant pulmonary isoform in other mammalian species, would be rare in most human populations. Two other FMO family members, FMO4 and FMO5, are only detectable at low levels in those tissues that have been examined (Gasser, 1996). In contrast, FMO1 and FMO3 are highly expressed in the human. FMO1 is readily detectable in the fetal liver, but is suppressed shortly after birth. In contrast, FMO1 seems to be the dominant extrahepatic enzyme in the adult (Yeung et al., 2000; Krause et al., 2002). FMO3 essentially is nondetectable in fetal liver, but is observed in most individuals by 1 to 2 years of age (Koukouritaki et al., 2002) and is expressed in the adult human liver at levels that approach those reported for CYP3A4 (Wrighton et al., 1990; Overby et al., 1997).

Several studies have documented up to a 10-fold or more range in interindividual FMO1 and FMO3 expression (Overby et al., 1997; Yeung et al., 2000; Koukouritaki et al., 2002). Such interindividual variation is unlikely due to environmental influences because FMO expression is not generally affected by exogenous agents. Thus, genetic variability is a more likely underlying mechanism. Due to its strong linkage with trimethylaminuria (Mitchell and Smith, 2001), considerable work has been done on the contribution of genetic variability to interindividual differences in FMO3 expression and activity (e.g., Park et al., 2002), but much less is known about variation at the FMO1 locus. Although a recent study by Furnes et al. (2003) reported on human FMO1 genetic variability, this study was limited in sequence coverage, the populations included, and functional analysis. Thus, the objective of the current study was to identify common FMO1 sequence variants in DNA samples representing multiple ethnic groups and to examine the identified variants for their functional significance and frequency in different populations.

	Materials and Methods

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Materials. Common chemical reagents, cell culture medium, and fetal bovine serum were purchased from Sigma-Aldrich (St. Louis, MO). Taq polymerase was purchased from Invitrogen (Carlsbad, CA) whereas restriction endonucleases and other DNA-modifying enzymes were obtained from either Invitrogen or New England Biolabs (Beverly, MA). QIAmp DNA blood MIDI kits were procured from QIAGEN (Valencia, CA) and NE-PER nuclear extraction kits were from Pierce Chemical (Rockford, IL). The ExoSAP-IT mix, containing both exonuclease I and shrimp alkaline phosphatase, as well as shrimp alkaline phosphatase alone was purchased from USB (Cleve-land, OH). ABI Prism big dye terminator cycle sequencing and SNaPshot kits were obtained from Applied Biosystems (Foster City, CA) and CEQ SNP-Primer Extension kits were from Beckman Coulter, Inc. (Fullerton, CA). The pGL-3 luciferase reporter system and luciferase assay kits were purchased from Promega (Madison, WI), whereas the LumiGal assay system for -galactosidase activity was obtained from BD Biosciences Clontech (Palo Alto, CA). The hepatocyte nuclear factor 1 (HNF1) expression plasmids pBJ5HNF1, pBJ5HNF1 (Mendel et al., 1991a), and pBJ5DCoH (Mendel et al., 1991b) were generous gifts from Dr. Gerald R. Crabtree (Stanford University School of Medicine, Stanford, CA). [-³²P]Deoxyribonucleotide triphosphates (3,000 Ci/mmol) were purchased from PerkinElmer Life Sciences (Boston, MA). Antibodies to yin yang 1 (YY1) (H-414), HNF1 (C-19), HNF1 (C-20), and octamer binding protein 1 (Oct1) (C-21) were obtained from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA), whereas antibody to Ku (p80) was obtained from NeoMarkers (Fremont, CA). Custom oligonucleotides were synthesized by Integrated DNA Technologies, Inc. (Coralville, IA). For single nucleotide polymorphism (SNP) discovery, a panel of DNA samples from 24 unrelated individuals and representative of the human population's ethnic diversity (Collins et al., 1998) was obtained from the Coriell Institute (Camden, NJ). A sample of human fetal liver total RNA was purchased from Stratagene (La Jolla, CA).

Patient Recruitment and DNA Isolation. Patients representing various ethnic groups were recruited to provide DNA samples as described previously (McCarver et al., 1997; Whetstine et al., 2000; Zheng et al., 2003). After obtaining informed consent, blood (2-4 ml) was obtained by venipuncture, and DNA was extracted using the QIAamp DNA blood MIDI kit according to the manufacturer's directions. Using DNAzol reagent (Invitrogen), DNA samples from individuals of known ethnicity also were isolated from tissue obtained from the University of Miami and University of Maryland Brain and Tissue Banks for Developmental Disorders that are in contract to the National Institute for Child Health and Development, NOI-HD-8-3284 and NOI-HD-8-3283, respectively. These two DNA sources resulted in 208 samples from individuals of African descent, 184 samples from individuals of Hispanic descent (145/184 from Latin or South American countries, 5/184 from the Caribbean, 34/184 unknown), and 134 samples from individuals of northern European descent. Finally, DNA was isolated from fetal liver samples obtained from the Central Laboratory for Embryology Research, University of Washington (HD-0-0836) that had been previously characterized for FMO1- and FMO3-specific content (Koukouritaki et al., 2002). These research protocols were approved by all Institutional Review Boards involved.

DNA Amplification and Sequence Analysis for SNP Discovery. PCR amplification of FMO1 DNA was accomplished using primers designed to amplify the entire coding and flanking splice acceptor and donor site sequences for each of FMO1's nine exons. Due to its size, exon 9 was analyzed as three overlapping regions to yield product sizes no more than 450 bp each. The 5'-flanking region covering approximately 1,000 bp also was amplified using two primer sets. Reactions were performed using 0.4 µg of genomic DNA and 1.25 U of Taq polymerase in a final volume of 50 µl containing 20 mM Tris-HCl (pH 8.4), 50 mM KCl, 1.5 mM MgCl₂, 0.2 mM each deoxynucleotide triphosphate, and 0.5 µM of each primer. Table 1 lists each primer set and the corresponding amplification conditions. Each of the 24 samples from the Coriell DNA Polymorphism Discovery Resource Panel and 53 of the samples isolated from tissue obtained from the University of Miami and University of Maryland Brain and Tissue Banks for Developmental Disorders were analyzed independently. Amplification products from 100 unrelated individuals of African-American descent were analyzed as pools of five individuals.

Before sequencing, a 5-µl aliquot of the DNA amplification reaction was transferred to a 96-well plate and treated with exonuclease I and shrimp alkaline phosphatase (ExoSAP-ITM) to degrade any residual single stranded DNA and hydrolyze remaining deoxynucleotide triphosphates. Sequencing reactions were performed on an MJ Research (Waltham, MA) thermal cycler after the addition of Big Dye reagent to each well and the appropriate sequencing primer [either the forward or reverse PCR primer (Table 1) at the concentration recommended in the Big Dye kit protocol].

Genotype Analysis. Validation of the functional FMO1 SNP was accomplished using a single base extension assay essentially as described by Lindblad-Toh et al. (2000) and following the recommended protocol included in the SNaPshot multiplex kit or the CEQ SNP-primer extension kit. The primer used for this assay was 5'-(C)₁₇TCT GGT ATT AGC AGA GAT CAA AAT-3'. The inclusion of the 5' nonhomologous poly (C) tail is due to this primer originally being designed as part of a 8mer multiplex set. However, because only one of these variants seemed to be functional, the complete multiplex analysis was limited to a restricted number of DNA samples and is not shown. Single base extension reactions were carried out for 25 cycles of 96°C for 10 s, 50°C for 5 s, and 60°C for 30 s using an MJ Research DNA engine tetrad thermal cycler. After treatment with shrimp alkaline phosphatase to eliminate interference from unincorporated dideoxynucleotide triphosphates, reactions were analyzed on an Applied Biosystems Prism 3700 DNA Analyzer or a Beckman Coulter, Inc. CEQ-8000 genetic analysis system.

Plasmids. Human FMO1 sequences were initially isolated using a Genome Walker kit (BD Biosciences Clontech), resulting in two plasmids, pRNH666 and pRNH667, containing FMO1 sequences from position -2330 to -760 and -821 to +57, respectively, cloned into the pCRII vector. The identity of these original clones was verified by sequence analysis. The above-mentioned coordinates are based on the most 5' transcription start site located 9,594 bp upstream of the FMO1 ATG start codon (National Center for Biotechnology Locus Link ID 2326, based on contig NT_029874.7, from build 31). This transcription start site was identified by 5'-rapid amplification of cDNA ends performed as described previously (Luo and Hines, 1997) using total fetal liver RNA (Stratagene) (data not shown; see Fig. 1 for mapped sites) and agrees with that reported in the Database of Transcriptional Start Sites (http://elmo.ims.u-tokyo.ac.jp/dbtss). Sequences from these two plasmids, along with a short, amplified fragment isolated from a genomic DNA sample, were used to generate pRNH759, pRNH757, pRNH760, pRNH761, pRNH765, and pRNH758, which contain human FMO1 sequences from position -22 to +112, -68 to +112, -262 to +112, -364 to +112, -1668 to +112, and -2268 to +112, respectively, directing luciferase expression within the pGL3basic backbone. pRNH766, pRNH717, pRNH767, pRNH768, pRNH770, and pRNH719 are identical to the above-mentioned plasmids, respectively, except that they contain a cytosine-to-adenine base substitution at position +59 that was introduced using the QuikChange II site-directed mutagenesis kit (Stratagene).

View larger version (19K): [in this window] [in a new window]   Fig. 1. YY1, Oct1, and HNF1 sites in the human FMO1 basal promoter. The position and orientation of the overlapping YY1, Oct1, and HNF1 sites within human FMO1 exon 1 are depicted. The YY1 site is a perfect match to the YY1 consensus. The shaded residues represent mismatches relative to the Oct1 consensus sequence, whereas the underlined residues represent mismatches to the HNF1 consensus sequence. The boxed residues depict the position of the g.-9,536C>A transversion. Transcription start sites, as determined by 5'-rapid amplification of cDNA ends, are shown by arrowheads, the closed arrow at position +16 indicating the major start site.

Cell Culture and Transient Reporter Gene Assays. The HepG2 human hepatoma cell line was a gift from Dr. Barbara Knowles (Jackson Laboratories, Bar Harbor, ME). The cells were cultured in Eagle's minimal essential medium supplemented with 10% fetal bovine serum, 50 U of penicillin/ml, and 50 µg of streptomycin/ml and were incubated in a humidified incubator at 37°Cinan atmosphere of 5% CO₂, 95% air. The cells were subcultured (1:4 ratio) every 3 to 4 days and used in the present study at passages 12 to 20. Transient transfections and reporter gene assays were performed as described previously (Luo and Hines, 1996).

Electrophoretic Mobility Shift Assays (EMSAs). Nuclear extract was prepared from the human hepatoma cell line HepG2 essentially as described by Boucher et al. (1993) or by using the NE-PER nuclear extraction kits (Pierce Chemical) following the manufacturer's recommended protocol. EMSA were performed as described by Luo and Hines (2001). The double-stranded oligonucleotide probes used in this study represent human FMO1 sequences from position +40 to +74 (relative to the 5'-most transcription start site), 5'-agctc ATA CTG ATT CAT TTT GAT CTC TGC T-3', and 5'-agctc ATA CTG ATT AAT TTT GAT CTC TGC T-3', the latter representing the human FMO1 g.-9,536A¹ variant identified in this study (adenine for cytosine transversion at underlined position +59; italicized residues indicate nonhomologous 5' ends added to facilitate radionucleotide labeling using a polymerase fill-in protocol). Double-stranded oligonucleotides used in competitive EMSA included these same human FMO1 g.-9,536C and g.-9,536A variants listed above and consensus sequences for HNF1 (5'-agct GGT TAA TGA TCT ACA-3') (Tronche and Yaniv, 1992), Oct1 (5'-TGT CGA ATG CAA ATC ACT AGA A-3'), and YY1 (5'-agctt CGC TCC GCG GCC ATC TTG GCG GCT GGT-3') (consensus sequences underlined), the latter two being commercially available (Santa Cruz Biotechnology, Inc.). Antibodies used for supershift experiments were used at 2.0 µg for YY1, HNF1, HNF1, and Oct1 and at 0.2 µg for Ku (p80).

Data Analysis. To identify FMO1 SNPs, DNA sequencing results were initially analyzed with PolyPhred (Nickerson et al., 1997) to identify potential discrepancies relative to the reference sequence (National Center for Biotechnology Information Locus Link ID 2326 based on contig NT_029874.7, build 31) and subsequently with the SeqManII software from DNAstar, Inc. (Madison, WI). The latter allowed for elimination of any poor quality data, contiguous sequence assembly, an automated assessment of trace quality, and finally, a visual inspection of any suspected variants. Sequence variations within splice sites were analyzed and scored for their relative fit to a consensus sequence using the Splice Site Predictor Program authored by Dr. Martin G. Reese (University of California, Berkeley, CA) and available at the Berkeley Drosophila Genome Project Web site (http://www/fruitfly.org/seq_tools/splice.html) (Reese et al., 1997).

	Results

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FMO1 SNP Discovery. Past studies on FMO expression and genetic variability have focused primarily on FMO3, because the expression of this gene predominates in the adult liver (Lomri et al., 1992). However, more recent studies have demonstrated that FMO1 is the major isoform expressed in the human fetal liver, adult intestine, and adult kidney (Yeung et al., 2000; Koukouritaki et al., 2002; Krause et al., 2002). Given this expression pattern, FMO1's potential impact on xenobiotic metabolism in these tissues, and the previously observed 10- to 20-fold variation in FMO1 expression (Koukouritaki et al., 2002), the possibility of FMO1 genetic variability takes on increased significance. To address this problem, the sequence of approximately 1 kbp of FMO1 5' upstream information, as well as the sequence of all nine exons and the immediate 5' and 3' exon flanking information was determined in DNA samples from 24 unrelated individuals obtained from the Coriell Polymorphism Discovery Resource Panel (Camden, NJ), 53 DNA samples of known ethnicity from the University of Maryland and University of Miami Brain and Tissue Banks for Developmental Disorders, and 100 individuals of African-American descent. The results from this effort are summarized in Table 2. Similar to the reported utility of the Coriell Polymorphism Discovery Resource Panel for identifying common variants in the CYP3A gene family (Kuehl et al., 2001), all of the common variants identified in the 100 African-American samples and the 53 samples from the University of Maryland and University of Miami also were identified in the Coriell Polymorphism Discovery Resource Panel. For completeness, Table 2 also includes FMO1 SNPs that have been identified and reported in the GeneSNPs database (http://www.genome.utah.edu/genesnps/) and those recently reported by Furnes et al. (2003). None of the 5' upstream variants (g.-10,361A, g.-10,330T, g.-10,046G, or g.-9,782T) fall within any of the previously identified regulatory elements that have been implicated in FMO1 regulation (Luo and Hines, 2001). Of the intronic variants, two fall within consensus splice sequences. The g.-11C variant lies within the intron 1 splice acceptor site, whereas the g.+17,248C variant lies within the intron 3 splice acceptor site. However, these substitutions represent conservative changes of one pyrimidine for another within the poly-pyrimidine tract and have a minimal effect on the splice site score, i.e., 0.91 (T) to 0.88 (C) for the FMO1^*1H² allele and 0.96 (T) to 0.93 (C) for the FMO1^*1A allele. As such, the g.-11C and g.+17,248C variants are unlikely to affect splicing efficiency. Similarly, the two 3'-untranslated region variants (g.+27,568T and g.+27,664T) do not seem to fall within functional sequences, e.g., those known to regulate mRNA stability. Of the three exonic variants, two (g.+22,828C>T in exon 6, T249T, FMO1^*1B; and g.+25,061A>G in exon 8, V396V, FMO1^*1C) represent synonymous changes and also do not seem to involve known exon splicing enhancer elements. Thus, one would predict that these two sequence variants also would not affect FMO1 function. However, the g.-9,536C>A transversion in the 5'-untranslated region of exon 1 lies within a core YY1 binding sequence (Shi et al., 1997). Previous studies in our laboratory demonstrated a role for this element in regulating rabbit FMO1 expression and the ability of the orthologous human FMO1 sequence to effectively compete with the rabbit element for YY1 binding (Luo and Hines, 2001). Thus, we questioned whether this variant might be capable of altering transcription efficiency and contribute to interindividual variability in human FMO1 expression.

Functional Analysis of the FMO1 g.-9,536C>A Variant. An EMSA was performed to determine whether the putative human FMO1 YY1 element could specifically bind YY1 or other transcription factors and what effect the observed SNP might have on such binding. A double-stranded oligonucleotide probe representing the g.-9,536C FMO1 variant from position +40 to +74 (relative to most 5' transcription start site) was capable of forming six specific complexes with nuclear proteins isolated from the human hepatoma cell line HepG2 (Fig. 2A; compare lane 1 with lanes 2-4). Inclusion of an anti-Ku (p80) antibody in the binding reaction further retarded complex D, consistent with this complex representing a semispecific interaction with the ubiquitous Ku DNA binding protein (data not shown). As such, further studies on complex D were not pursued. The ability of a double-stranded oligonucleotide representing a consensus YY1 element to compete with proteins forming complexes E and F suggested that these complexes involved YY1 (Fig. 2A, lane 8). This was further confirmed by inclusion of a YY1 antibody in the binding reaction that further retarded the mobility of the DNA/protein complexes E and F (Fig. 2A, lane 9). However, competition for YY1 binding was not observed with a 100- to 400-fold molar excess of a double-stranded oligonucleotide representing the human FMO1 g.-9,536A variant (Fig. 2A, lanes 5-7). Sequence analysis of the FMO1 position +40 to +74 EMSA probe revealed potential Oct1 and HNF1 sites that overlap the YY1 element (Fig. 1). Indeed, formation of the much weaker specific DNA/protein complex C was inhibited by inclusion of excess consensus Oct1 binding element and its electrophoretic mobility further retarded by the inclusion of an anti-Oct1 antibody in the binding reaction (Fig. 2A, lanes 10 and 11, respectively). Similarly, formation of the specific complexes A and B with the radiolabeled probe was eliminated by inclusion of a molar excess of an HNF1 consensus sequence (Fig. 2A, lane 12). The electrophoretic mobility of complex A and B was further retarded by inclusion of an HNF1 antibody in the binding reaction, whereas the electrophoretic mobility of only complex B was retarded by inclusion of an antibody specific for HNF1 (Fig. 2A, lanes 13 and 14, respectively). These data suggest complex A involves an HNF1 homodimer and that complex B involves an HNF1/HNF1 heterodimer. The double-stranded oligonucleotide representing the human FMO1 g.-9,536A variant retained the ability to compete for Oct1, HNF1, and HNF1 binding (Fig. 2A, lanes 5-7). Overall, these data are consistent with the g.-9,536C>A FMO1 sequence variation potentially affecting transcription by significantly reducing YY1 binding.

View larger version (42K): [in this window] [in a new window]   Fig. 2. DNA protein interactions at the putative human FMO1 YY1 element. A, EMSA was used to characterize specific HepG2 nuclear protein binding to a radiolabeled fragment representing position +50 to +74 of the human FMO1 gene (g.+9,536C, FMO1*1 allele). The addition of a 100- to 400-fold molar excess of competitive DNA or antibody to specific transcription factors is indicated above each lane. Solid arrows to the left of the figure indicate specific DNA/protein complexes, whereas the open arrows on the right indicate DNA/protein complexes further retarded by the addition of antibody. B, EMSA was used to characterize specific HepG2 nuclear protein binding to a radiolabeled fragment representing position +50 to +74 of the human FMO1 gene (g.+9,536A, FMO1*6 allele). The addition of a 400-fold molar excess of competitive DNA or antibody to specific transcription factors is indicated above each lane. Solid arrows to the left of the figure indicate specific DNA/protein complexes, whereas the open arrows indicate the predicted position of the missing E and F complexes. The open arrows on the right indicate DNA/protein complexes further retarded by the addition of antibody.

The double-stranded oligonucleotide representing the g.-9,536A variant also was used as an EMSA probe (Fig. 2B) to further elucidate the degree to which this sequence change impacts transcription factor binding. As seen in Fig. 2B, the g.-9,536C>A transversion results in a complete loss of complexes E and F, indicating the g.-9,536A sequence is no longer capable of binding YY1 (Fig. 2B, expected mobilities indicated by open arrows). Of interest, complexes A, B, and C, and in particular complex A, seem more intense (Fig. 2B, lane 1), consistent with an increased affinity of the variant sequence for Oct1, HNF1, and HNF1. Further evidence for such a change in binding affinity is observed in Fig. 2B, lanes 2 and 3. Using a 400-fold molar excess of either the unlabeled g.-9,536A or g.-9,536C double-stranded oligonucleotides, competition with the radiolabeled g.-9,536A probe for complexes A, B, and C was less effective with the g.-9,536C sequence. Competition with an excess of the Oct1 (Fig. 2B, lane 7) and HNF1 (Fig. 2B, lane 4) consensus sequences, as well as further retardation of electrophoretic mobility by the HNF1 and HNF1 antibodies (Fig. 2B, lanes 5 and 6, respectively) reconfirmed the identity of complexes A, B, and C. A new, high-molecular-weight-specific complex G is observed with the variant sequence (Fig. 2B). Complex G also was eliminated by competition with the unlabeled HNF1 consensus sequence (Fig. 2B, lane 4) and its mobility further retarded by inclusion of the HNF1, but not HNF1 antibody in the binding reaction. These data suggest the involvement of HNF1 in complex G as well.

To further explore the potential impact of the human FMO1 YY1 sequence variant, several luciferase expression plasmids containing segments of the FMO1 promoter representing either the FMO1^*1 (g.-9,536C) or FMO1^*6 (g.-9,536A) alleles were analyzed by transient transfection and expression analysis in HepG2 cells (Fig. 3). There were no significant differences between the constructs containing either the minimal promoter (position -22 to +112) or up to position -66 [relative to the most 5' transcription start site, the YY1 element is located at FMO1 position +57 to +63 with the C>A transversion at +59 (Fig. 1)]. However, in constructs containing further upstream information, a significant decrease in transcriptional activity was observed with the g.-9,536A variant that could account for a 2- to 3-fold difference in FMO1 expression between individuals. Based on the in vitro DNA/protein binding experiments presented above (Fig. 2, A and B), it was of interest to see what differential effect, if any, cotransfection with either or both HNF1 and HNF1 expression vectors might have on the function of the two FMO1 promoter sequences. However, increased expression of either HNF1 factor, or both factors together, was not able to reverse the negative impact of the g.-9,536C>A transversion on FMO1 promoter activity (data not shown).

View larger version (15K): [in this window] [in a new window]   Fig. 3. Functional analysis of the human FMO1*6 (g.-9,536A) variant. Reporter constructs containing various lengths of human FMO1 5'-flanking information directing luciferase expression were transiently transfected into HepG2 cells and analyzed for transcriptional activity. Pairs of plasmids (individual plasmids identified by numbers on the ordinate) containing identical lengths of 5'-flanking information, but containing either the FMO1*1 (g.-9,536C) or FMO1*6 (g.-9,536A) variant were compared. Relative to the most 5' transcription start site, the putative YY1 element is located at position +57 to +63 with the C>A transversion under study at position +59 (indicated by arrowheads under each promoter construct). The activity reported has been normalized for both transfection efficiency and protein content and represents the mean ± S.D. from four experiments. The significance of differences observed between variants was tested using an unpaired t test with a Welch post hoc correction. A p < 0.05 was accepted as significant and is indicated by an asterisk.

SNP Validation. Although a multiplex assay was developed for the FMO1 SNPs identified in this study and used to analyze DNA from several individuals, complete analysis is only reported for the g.-9,536C>A SNP because it seems to be the only functional sequence variant. The FMO1^*6 (g.-9,536A) allele was present at a frequency of 25% in the DNA samples obtained from the Coriell Polymorphism Discovery Resource Panel (Table 2). However, these samples are representative of the human population's ethnic diversity and cannot be used to assess potential differences in frequency among different ethnic groups (Collins et al., 1998). To address the frequency of the FMO1^*6 variant, genotyping was performed in defined African-, northern European-, and Hispanic-American populations. As shown in Table 3, the frequency of the FMO1^*6 allele in the African- and northern European-American populations is not significantly different (13.8 and 10.8%, respectively). However, the frequency of the FMO1^*6 allele in Hispanic-Americans, which were predominantly of Mexican descent, was significantly higher, i.e., 29.6% (Fisher's exact test, p < 0.001). For each population, the genotype frequencies were not significantly different from that predicted by the Hardy-Weinberg equilibrium (² test accepting an < 0.05 as significant).

	Discussion

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Considerable interindividual variability in human FMO1 expression has been noted: 3- to 6-fold in adult kidney (Krause et al., 2002), up to 6-fold in adult small intestine (Yeung et al., 2000), and 10- to 15-fold in fetal liver (Koukouritaki et al., 2002). Given the apparent absence of environmental influences on FMO1 expression, a thorough screen for possible genetic variability was undertaken to determine whether this might be a contributing factor to intersubject differences. Sequence analysis of approximately 1.0 kbp of upstream information, as well as all exons and immediate flanking sequences identified 10 single-base substitutions. An additional five single-base substitutions have been reported in the GeneSNP database located in intronic sequences not covered in the current study (Table 2). Overall, the amount of variability at the FMO1 locus seems considerably less than that reported in the GeneSNP database for either FMO2 (39 reported SNPs with 17 having a frequency equal to or greater than 1%) or FMO3 (44 reported SNPs with 29 having a frequency equal to or greater than 1%) (http://www.genome.utah.edu/genesnps). This is despite all three genes being localized within a single, approximate 256-kbp chromosome 1q cluster that also includes FMO4 and FMO6P. The conservation of FMO1 relative to FMO2 and FMO3 also is consistent with the data recently reported by Furnes et al. (2003) wherein a total of seven FMO1 SNPs were reported relative to 30 variants for FMO2 and 16 for FMO3 in 50 unrelated African-Americans. Two of the four rare FMO1 variants (1-2%) that would affect protein structure or expression reported by Furnes et al. (2003) were observed in the current study, but also were rare. The FMO1^*4 allele (g.+23,971T>C, I303T) was observed in two pooled African-American DNA samples and based on relative peak height, most likely represented single heterozygotes. A single African-American heterozygote was observed for FMO1^*3 (g.+23,970A>G, I303V). None of the FMO1^*2 (g.+9,614C>G, H97Q) or FMO1^*5 (g.+27,362C>T, R502X) variants were observed. Combining the data from the two studies suggests frequencies of 0.3% for FMO1^*2, 1.0% for FMO1^*3, 1.3% for FMO1^*4, and 0.3% for FMO1^*5 within the African-American population. The FMO1^*1B and FMO1^*1C synonymous variants reported by these investigators also were observed commonly in the current study (Table 2). However, Furnes et al. (2003) did not include FMO1 exon 1 or upstream sequences in their analysis, precluding comparisons for five of the SNPs identified herein (i.e., FMO1^*1D, ^*1E, ^*1F, ^*1G, and ^*6).

Among the single nucleotide variations identified in the current report, only the SNP located at position g.-9,536 (+59 relative to the 5'-most transcription start site, Fig. 1) seems to have functional significance. A single nucleotide change of C to A at this position eliminates binding of YY1 (or an unknown factor closely related to YY1), a transcription factor important for negatively regulating rabbit FMO1 expression (Luo and Hines, 2001). Similar to what was observed for the rabbit gene, functional studies on human FMO1 suggest the loss of YY1 binding has little or no effect on basal FMO1 promoter activity. However, in contrast to rabbit FMO1, loss of binding in conjunction with the presence of upstream regulatory sequences resulted in a 2- to 3-fold loss of human FMO1 promoter activity when examined in HepG2 cells. These data suggest that YY1 can positively regulate human FMO1 expression, whereas it can negatively regulate rabbit FMO1 expression, but in both cases, only in coordination with more distal regulatory elements. Such a dual role for YY1 is not surprising, because there are ample examples of both positive and negative gene regulation through YY1's interactions with cofactors possessing histone acetylase or histone deacetylase activity, respectively (Thomas and Seto, 1999). YY1 is a highly conserved member of GLI-Krüppel family of zinc-finger proteins (Shi et al., 1997) and is ubiquitously expressed both in the adult (Thomas and Seto, 1999) and the developing fetus (Donohoe et al., 1999). Thus, the inability of YY1 to participate in FMO1^*6 transcriptional regulation might be expected to impact expression levels during development and in the adult.

The EMSA analysis also suggests potential roles for Oct1, HNF1, and/or HNF1 in regulating FMO1 expression that would not be affected by the g.-9,536A variant, because this sequence change seems to enhance binding of these factors. Analysis of the +40 to +74 element identified potential Oct1 and HNF1 sites that are unique to the human FMO1 gene and overlap the previously identified YY1 element (Fig. 1), consistent with competitive binding between YY1 and these two factors. Indeed, the C-to-A transversion improves this element's identity with the core HNF1 consensus site (5'-GGTTAATNATTAMCA-3') (Tronche and Yaniv, 1992). This observation may contribute to an explanation for a previous, paradoxical observation made when FMO1 genotype was correlated with FMO1-specific content in a group of fetal liver samples of various gestational ages (Koukouritaki et al., 2002). Two individuals homozygous for the FMO1^*6 allele were identified. However, the FMO1 protein levels in these samples were in the upper quartile of their age bracket, rather than the lower quartile as one would have predicted based on the loss of YY1 binding. Enhanced binding of HNF1 and/or other transcription factors at the g.-9,536A variant element might compensate for YY1's absence. Although its impact would be less in extrahepatic and adult tissues, HNF1 is expressed at relatively high levels in the fetal liver and is important in the temporal- and tissue-specific expression of numerous genes. However, an increased role for HNF1 is unlikely the sole explanation for the unexpected high level of FMO1^*6 expression in these two fetal liver samples, because transient expression studies in HepG2 cells using FMO1^*1- or FMO1^*6-based reporter constructs and an HNF1 expression vector failed to show any sort of differential effect on transcription. A definitive explanation must await a better understanding of FMO1 regulation during development.

Previous studies have demonstrated considerable ethnic variability in the frequency of polymorphisms at other FMO loci (Whetstine et al., 2000; Cashman et al., 2001; Park et al., 2002). Consistent with these observations, the FMO1^*6 allele occurred at a frequency of 14 to 10% in the African- and northern European-American populations tested, respectively, but at 31% in the Hispanic-American population. Thus, it is likely the g.-9,536C>A polymorphism not only contributes to interindividual variation in FMO1 expression, but also may contribute to differences in expression between different subpopulations. Although their sample size was small, such a difference is consistent with a recent report by Krause et al. (2002).

Several studies have documented the importance of FMO1 in the metabolism of a variety of therapeutics, pathways that potentially would be impacted by the FMO1^*6 polymorphism. Because FMO1 is the dominant hepatic FMO isoform in mammalian species other than human, previous studies using liver preparations from animal models would have focused on FMO1 by default. Of the different FMO isoforms, FMO1 has the largest substrate access channel (or cleft), although human FMO1 is more restricted than the orthologous porcine enzyme (Kim and Ziegler, 2000). Nevertheless, human FMO1 seems to selectively N-oxidize the bulky tricyclic antipsychotic agents imipramine, chlorpromazine, and orphenadrine (Kim and Ziegler, 2000) and the antihistaminics promethazine and brompheniramine (Cashman et al., 1993a; Clement et al., 1993). Examples of therapeutic agents for which both FMO1 and FMO3 are capable of efficiently catalyzing heteroatom-oxidation include the H₂ receptor antagonists ranitidine and cimetidine (Cashman et al., 1993b; Overby et al., 1997), the gastroprokinetic agent itopride (Mushiroda et al., 2000), and the M₁-muscarinic agonist xanomeline (Ring et al., 1999). However, the selective expression of FMO1 in the adult small intestine and kidney would predict that this enzyme would be most important for intestinal first pass and renal metabolism, respectively. Similarly, the selective expression of FMO1 in the developing fetal liver would suggest this enzyme would be more important after exposure to such compounds in utero. Other examples of therapeutics and toxicants metabolized by this enzyme system, and as such, pathways potentially impacted by the FMO1^*6 polymorphism, can be found in the review by Hines (2003).

In summary, we have demonstrated that human FMO1 is more highly conserved than other members of the FMO gene family localized within the same cluster on chromosome 1. Coupled with the previous demonstration of fetal-specific FMO1 expression in human liver (Koukouritaki et al., 2002), this observation causes one to speculate about an important role for this enzyme during ontogeny. The identification of a common variant at the exon 1 YY1 element (FMO1^*6) is the first report of a functional FMO1 polymorphism. This variant results in a loss of YY1 binding, could account for up to a 2- to 3-fold decrease in FMO1 expression and may also contribute to differential expression among different subpopulations. Although the FMO1^*6 variant may well impact a variety of metabolic pathways (see examples above), its importance will depend on the relative role and abundance of the different transcription factors that are capable of binding this regulatory element in different tissues and at different times during development and aging.

	Footnotes

 
These studies were supported in part by U.S. Public Health Services Grant CA53106 from the National Cancer Institute and funds from the Children's Hospital Foundation of Wisconsin.

Article, publication date, and citation information can be found at http://jpet.aspetjournals.org.

DOI: 10.1124/jpet.103.053686.

ABBREVIATIONS: FMO, flavin-containing monooxygenase; kbp, kilobase pair; HNF1, hepatocyte nuclear factor 1; YY1, yin yang 1; Oct1, octamer binding protein 1; SNP, single nucleotide polymorphism; bp(s), base pair(s); PCR, polymerase chain reaction; EMSA, electrophoretic mobility shift assay.

¹ Allele nomenclature follows the guidelines adopted for the human cytochrome P450 gene family (http://www.imm.ki.se/CYPalleles/criteria.htm).

² Genetic variants (SNPs) are named based on the recommendations of the Nomenclature Working Group as outlined by den Dunnen and Antonarakis (2001). In brief, "g". indicates a reference genomic sequence followed by a gene coordinate based on the A of the ATG-translation initiation codon being assigned +1, followed by the nucleotide variation, e.g., C>T where the first nucleotide is the dominant allele.

Address correspondence to: Dr. Ronald N. Hines, Department of Pediatrics, Birth Defects Research Center, Medical College of Wisconsin, 8701 Watertown Plank Rd., Milwaukee WI 53226-4801. E-mail: rhines{at}mail.mcw.edu

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