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Potential link between estrogen receptor-α gene hypomethylation and uterine fibroid formation

Potential link between estrogen receptor-α gene hypomethylation and uterine fibroid formation Abstract Uterine leiomyomas are the most common uterine tumors in women. Estrogen receptor-α (ER-α) is more highly expressed in uterine leiomyomas than in normal myometrium, suggesting a link between uterine leiomyomas and ER-α expression. DNA methylation is an epigenetic mechanism of gene regulation and plays important roles in normal embryonic development and in disease progression including cancers. Here, we investigated the DNA methylation status of the ER-α promoter region (−1188 to +229 bp) in myometrium and leiomyoma. By sodium bisulfite sequencing, 49 CpG sites in the proximal promoter region of ER-α gene were shown to be unmethylated in both leiomyoma and normal myometrium. At seven CpG sites in the distal promoter region of the ER-α gene, there was a variation in DNA methylation status in myometrium and leiomyoma. Further analysis of the DNA methylation status by bisulfite restriction mapping among 11 paired samples of myometrium and leiomyoma indicated that CpG sites in the distal region of ER-α promoter are hypomethylated in leiomyomas of nine patients. In those patients, ER-α mRNA levels tended to be higher in the leiomyoma than in the myometrium. In conclusion, there was an aberrant DNA methylation status in the promoter region of ER-α gene in uterine leiomyoma, which may be associated with high ER-α mRNA expression. DNA methylation, epigenetics, estrogen receptor-α, ER-α promoter, leiomyoma Introduction Uterine leiomyomas are the most common uterine tumors in women of reproductive age. Approximately 20–25% of women of reproductive age are afflicted with this disease (Vollenhoven et al., 1990). They frequently cause serious gynecological problems such as pelvic pain, menorrhagia, dysmenorrhea, reduced fertility and recurrent pregnancy loss (Bajekal and Li, 2000; Stewart, 2001). In addition, uterine leiomyoma is the most common indication for hysterectomy in Japan, as well as in the USA (Farquhar and Steiner, 2002). Despite the high prevalence rate and tremendous influence on reproductive women, the pathogenesis of uterine leiomyomas still remains to be elucidated. On the basis of the fact that uterine leiomyomas develop only after menarche and markedly shrink under hypoestrogenic conditions such as late menopause, it is presumed that their growth depends on estrogens (Stewart, 2001). Although the increased sensitivity to estradiol is important for the growth of uterine leiomyomas, high circulating estradiol levels are not a necessary requirement. The physiological effects of estrogen are mediated by estrogen receptors (ERs). Among them, ER-α is more highly expressed in uterine leiomyomas than in normal myometrium (Benassayag et al., 1999; Kovács et al., 2001), suggesting a possible link between uterine leiomyomas and ER-α expression level. Epigenetic mechanisms including DNA methylation and histone modification are known to play key roles in transcriptional regulation. DNA methylation occurs at cytosines within CpG dinucleotides that are clustered frequently in regions of ∼1–2 kb in length, called CpG islands, in or near promoter and first exon regions of genes (Esteller and Herman, 2002). In mammals, 60–70% of CpG sites are methylated in the genomic DNA (Boyes and Bird, 1992). DNA methylation is involved in various developmental processes by silencing, switching and stabilizing genes (Nan et al., 1998; Cho et al., 2001; Imamura et al., 2001; Li, 2002). Although there are differences in the frequency of CpGs in the gene regulatory regions, DNA methylation-dependent gene regulation has been previously reported (Razin and Cedar, 1991; Cho et al., 2001; Imamura et al., 2001). CpG methylation can down-regulate gene expression by preventing the binding of transcription factors or by recruiting repressor molecules (Bird, 1992; Ballestar and Wolffe, 2001). Accumulating evidence has indicated that increased methylation level of the CpG islands within the ER-α promoter region is highly negatively associated with ER-α expression in a variety of diseases including neoplastic and atherosclerotic lesions (Iwase et al., 1999; Post et al., 1999; Yoshida et al., 2000; Berger and Daxenbichler, 2002). Thus, down-regulation of the ER-α expression is caused by hypermethylation of the CpG islands within the ER-α promoter region. However, the fact that the ER-α expression was higher in uterine leiomyomas than in normal myometrium is different from other ER-α related diseases described earlier. This led us to assume that a different epigenetic abnormality might be involved in uterine leiomyomas. The present study was undertaken to investigate the methylation status of CpG sites within the promoter region of the human ER-α gene and to evaluate an association of aberrant DNA methylation status with ER-α gene expression in uterine leiomyomas and normal myometrium. Materials and Methods Tissue preparation Specimens of uterine leiomyomas and corresponding normal myometrium were obtained from 18 women, from 37 to 57 (mean 47.4) years of age, who underwent total hysterectomy. Normal myometrium was obtained from a woman without myoma (49 years, cervical cancer). Informed consent was obtained from all participating patients, and ethical approval was obtained from Yamaguchi University Graduate School of Medicine. Tissues were taken immediately after removal of the uterus, immersed in liquid nitrogen and stored at −80°C until used DNA/RNA extraction. For immunohistochemistry, the specimens were fixed immediately in 10% neutral formalin for ∼24 h, embedded in paraffin and cut into 4 μm thick sections. Immunohistochemistry The diagnosis of leiomyoma and normal myometrium was established on histological examination with hematoxylin and eosin staining. Immunohistochemistry was performed as described previously (Sugino et al., 2005) using an ER-α monoclonal antibody (ER1D5, mouse, Dako Japan Co. Ltd., Tokyo, Japan). Counterstaining was performed with Meyer's hematoxylin. Real-time RT–PCR analysis Total RNAs were isolated from tissues using Isogen reagent (Nippon Gene, Tokyo, Japan) and reverse-transcribed using an ExCript RT reagent kit (TaKaRa, Ohtsu, Japan) according to the manufacturer's protocol, respectively. For PCR amplification, first strand cDNA was synthesized from 1 µg total RNA with reverse transcriptase in 20 µl of reaction mixture. The oligonucleotide primers for ER-α (5′-TGTGCAATGACTATGCTTCA-3′ and 5′-GCTCTTCCTCCTGTTTTTA-3′; 149 bp amplified products) were designed from the human ER-α cDNA sequence (Matsuzaki et al., 2000). Internal control PCR primers for GAPDH (5′-AGGTGAAGGTCGGAGTCA-3′ and 5′-GGTCATTGATGGCAACAA-3′; 99 bp amplified products) were designed from the GAPDH cDNA sequence (Kaneda et al., 2004). Real-time PCR was performed using LightCycler (Roche Diagnostics, Indianapolis, IN, USA). The reaction mixture contained 10 µl SYBR Premix Ex Taq (TaKaRa), 0.2 µM each of primer sets of ER-α or GAPDH and 2 µl cDNA in a total volume of 20 µl. The thermocycling program was 40 cycles of 96°C for 5 s and 60°C for 20 s with an initial cycle of 96°C for 10 s. Sodium bisulfite genomic sequencing Genomic DNA was extracted using Genomic DNA kit (Qiagen, Tokyo, Japan) according to the manufacturer's protocol. The bisulfite reaction, in which unmethylated cytosine is converted to uracil and 5-methylcytsine remains non-reactive, was carried out as previously described (Cho et al., 2001; Imamura et al., 2001) with a slight modification: 2 µg of genomic DNA digested with Pvu II was denatured by incubation with NaOH at 42°C for 20 min. After the incubation, sodium metabisulfite and hydroquinone (Wako, Osaka, Japan) were added to the final concentrations of 2.0 M and 0.5 mM, respectively, and the mixture was incubated at 55°C for 16 h. The bisulfite reaction was terminated by incubation with NaOH again at 42°C for 20 min. The DNA fragments covering the transcriptional regulatory region of ER-α gene (−1297 to +279) were amplified by PCR using the following set of primers: Region I (−126 to +279) F: 5′-GTTGTGTTTGGAGTGATGTTTAAGTT-3′, R: 5′-CAATAAAACCATCCCAAATACTTTA-3′; Region II (−670 to −94) F: 5′-GGAAGGGTTTATTTATTTTGGGAGTA-3′, R: 5′-TAACATTAACTTAAACATCACTCC-3′; Region III (−1297 to −731) F: 5′-TTGGGTGTTTGGGATAGTAATTAAA-3′, R: 5′-CTTAATCCCATTAAAAATTCTCATA-3′. The PCR conditions were 95°C for 10 min, and 40 cycles of 94°C for 30 s, 55°C for 30 s and 72°C for 1 min, with a final extension at 72°C for 10 min. The resulting products were subjected to agarose gel electrophoresis and purified using a QIAquick gel extraction kit (Qiagen). The PCR products were cloned into pGEM-T easy vector (Promega, Tokyo, Japan), and 10 or more clones were randomly picked from each of two independent PCRs were sequenced to determine the methylation status. Sequencing was performed using an ABI automated sequencer with BigDye terminators (Applied Biosystems, Foster City, CA, USA). Bisulfite restriction mapping The bisulfite-converted DNA was amplified by PCR with a set of primers (−1120 to −645), F: 5′-TATATATATGTGTGTGTGTATGTG-3′ and R: 5′-TACTCCCAAAATAAATAAACCCTTCC-3′. One half of the PCR products were digested with 5 U of Taq I (TaKaRa) at 65°C for 3 h. The remaining half was used for undigested control without Taq I treatment. Taq I recognizes 5′-TCGA-3′ sequences. Because only unmethylated cytosine sites are changed to thymine by sodium bisulfite PCR, PCR fragments from unmethylated genomic DNA are resistant to Taq I, whereas those from methylated DNA are digested by the enzyme. The resulting products of the bisulfite restriction mapping were assessed by agarose gel electrophoresis. Cell culture, demethylation and RT–PCR analysis Primary human uterine smooth muscle cells and cell culture reagents were purchased from Cambrex, Inc. (Walkersville, MD, USA). Cells were grown to 90% confluence and maintained in SMGM2 medium, which consists of smooth muscle basal medium supplemented with 5% fetal bovine serum, 2 ng/ml recombinant human fibroblast growth factor-β, 5 µg/ml insulin, 0.5 ng/ml recombinant human epidermal growth factor, 50 µg/ml gentamicin sulfate and 50 µg/ml amphotericin B. Medium was changed every other day and all experiments were completed with cells derived from passages 2. For treatment with a demethylating agent, 5-aza-dC (Sigma, St Louis, MO, USA), that inhibits DNA methylation, cells were seeded at a density of 1 × 106 cells/25 cm2 tissue culture flask. After 24 h of incubation, cells were cultured with treatment medium containing 1 μM 5-aza-dC for 72 h. The medium was changed daily. After treatment, cells were used for RNA isolation and ER-α mRNA levels were measured by RT–PCR as reported previously (Sugino et al., 1998). Total RNAs were isolated from the cells according to the protocol mentioned above. For PCR amplification, first-strand cDNA was synthesized from 1 μg total RNA with reverse transcriptase in 20 µl of reaction mixture. The oligonucleotide primers for ER-α (5′-TGTGCAATGACTATGCTTCA-3′ and 5′-GCTCTTCCTCCTGTTTTTA-3′; 149 bp amplified products) were designed from the human ER-α cDNA sequence (Matsuzaki et al., 2000). Internal control PCR primers for ribosomal protein L19 (5′-CTGAAGGTCAAAGGGAATGTG-3′ and 5′-GGACAGAGTCTTGATGATCTC-3′; 194 bp amplified products) were designed from the L19 cDNA sequence (Sugino et al., 1998). PCR amplification was performed using a programmed temperature control system (PC808, ASTEC, Fukuoka, Japan). The reaction mixture contained 4 µl cDNA, 1 µM each of primer sets of ER-α or L19, GeneAmp 10× PCR buffer, 0.2 mM deoxynucleotide triphosphate, 2.5 mM MgCl2 and 0.05 U AmpliTaq DNA polymerase (Applied Biosystems) in a total volume of 20 µl. The thermocycling program was an initial cycle of 94°C for 5 min, then 35 cycles of 94°C for 1 min, 60°C for 1 min, 72°C for 1 min, followed by 10 min of final extension at 72°C. The resulting products were subjected to agarose gel electrophoresis. The level of ER-α expression was determined by quantifying the intensities of the PCR product, compared with the L19 product, using NIH ImageJ software. Statistical analyses Wilcoxon signed-ranks test was used for paired samples. A value of P < 0.05 was considered significant. Results ER-α expression Immunohistochemical staining for ER-α expression was localized in nuclei of smooth muscle cells, and the staining distribution was homogenous in both leiomyoma and myometrium (Fig. 1A). Figure 1: View largeDownload slide ER-α expression in uterine leiomyoma and normal myometrium. (A) Immunohistochemical staining of ER-α in leiomyoma (leio) and myometrium (myo). Immunohistochemical staining for ER-α was performed on tissue samples obtained from three different patients. HE, hematoxylin–eosin staining, Bar; 50 µm. (B) ER-α mRNA expression in leiomyoma and myometrium. Specimens of leiomyomas and corresponding myometrium were obtained from 18 women. Total RNA was isolated from 18 pairs of leiomyomas and myometrium. ER-α mRNA levels were analyzed by SYBR Green I real-time quantitative RT–PCR. Relative ER-α expression normalized to GAPDH was calculated. Values are mean ± SEM. *P < 0.01 versus myometrium. Figure 1: View largeDownload slide ER-α expression in uterine leiomyoma and normal myometrium. (A) Immunohistochemical staining of ER-α in leiomyoma (leio) and myometrium (myo). Immunohistochemical staining for ER-α was performed on tissue samples obtained from three different patients. HE, hematoxylin–eosin staining, Bar; 50 µm. (B) ER-α mRNA expression in leiomyoma and myometrium. Specimens of leiomyomas and corresponding myometrium were obtained from 18 women. Total RNA was isolated from 18 pairs of leiomyomas and myometrium. ER-α mRNA levels were analyzed by SYBR Green I real-time quantitative RT–PCR. Relative ER-α expression normalized to GAPDH was calculated. Values are mean ± SEM. *P < 0.01 versus myometrium. Many investigators have reported that ER-α is more highly expressed in leiomyomas than in myometrium (Benassayag et al., 1999; Kovács et al., 2001), suggesting a possible link between leiomyomas and ER-α expression level. To investigate whether ER-α levels are altered in leiomyoma samples that we collected, the ER-α mRNA was measured in leiomyomas and myometrium by real-time RT–PCR. As shown in Fig. 1B, ER-α mRNA levels in the samples we examined were confirmed as significantly higher in leiomyomas than in myometrium (P < 0.01). Effects of 5-aza-dC on ER-α mRNA expression in human uterine smooth muscle cells To study the possibility that ER-α mRNA expression is under epigenetic regulation such as DNA methylation, human uterine smooth muscle cells were incubated with 5-aza-dC which inhibits DNA methylation. ER-α mRNA expression in human uterine smooth muscle cells was significantly (P < 0.05) increased by 5-aza-dC (Fig. 2). Figure 2: View largeDownload slide Effects of 5-aza-dC on ER-α mRNA expression in human uterine smooth muscle cells. Primary human uterine smooth muscle cells were incubated with a demethylating agent, 5-aza-dC (1 µM) that inhibits DNA methylation, for 72 h. After treatment, cells were used for RNA isolation and ER-α mRNA levels were measured by RT–PCR. The intensity of the signals of ER-α was normalized to that of the internal control L19 (the ratio of ER-α to L19). Data were expressed as a percentage of the control value in each incubation. Each bar represents the mean ± SEM of three different experiments. *P < 0.05 versus control. Figure 2: View largeDownload slide Effects of 5-aza-dC on ER-α mRNA expression in human uterine smooth muscle cells. Primary human uterine smooth muscle cells were incubated with a demethylating agent, 5-aza-dC (1 µM) that inhibits DNA methylation, for 72 h. After treatment, cells were used for RNA isolation and ER-α mRNA levels were measured by RT–PCR. The intensity of the signals of ER-α was normalized to that of the internal control L19 (the ratio of ER-α to L19). Data were expressed as a percentage of the control value in each incubation. Each bar represents the mean ± SEM of three different experiments. *P < 0.05 versus control. DNA methylation status of 5′-flanking region of ER-α gene Since ER-α mRNA expression seemed to be under the regulation of DNA methylation, we examined DNA methylation status of the ER-α promoter region between leiomyoma and myometrium. The 5′-upstream region around the first exon (between about −500 and +200 bp) of the ER-α gene is most importantly involved in the regulation of ER-α expression (Lapidus et al., 1996; Yan et al., 2001; Giacinti et al., 2006). Furthermore, a series of three estrogen response elements (EREs), which lie from −892 to −420 in the ER-α upstream region, is identified as a region with functional importance for ER-α gene transcription (Furguson et al., 1997; Li et al., 2000). Therefore, ∼1 kb upstream region together with a part of the first exon of the ER-α gene is considered as important for ER-α gene transcription, and the methylation status of this region was compared between leiomyoma and myometrium. Figure 3A shows the distribution of CpGs within the transcriptional regulatory region of the human ER-α gene. The ER-α gene has two CpG islands in the promoter region and in the first exon. According to the registered nucleotide sequence of the ER-α (GenBank accession no. AB090237), there were 56 CpG sites in the 1.5 kb genomic sequence including the promoter and exon 1 of the human ER-α gene (Fig. 3A). In the present study, these CpG sites were divided into two regions, the proximal and distal regions. The proximal region included 49 CpG sites around the transcription start site (−556 to +229, designed as Regions I and II), and the distal region included seven CpG sites in the upstream region (−1188 to −790, designed as Region III). First, the methylation status of all the CpG sites between −1188 and +229 (56 CpG sites) were analyzed by sodium bisulfite genomic sequencing for a paired sample of leiomyoma and myometrium (Fig. 3B, Case 1). The proximal region (Regions I and II containing a total of 49 CpG sites) was unmethylated in both tissues (Fig. 3B, Case 1). In the distal promoter region (Region III, seven CpG sites), 47 CpG sites (24.7%) in a total of 190 examined CpG sites were methylated in myometrium, whereas only nine CpG sites (4.9%) in a total of 184 examined CpG sites were methylated in leiomyoma (Fig. 3B, Case 1). There was a significant difference in the frequency of DNA methylation between leiomyoma and myometrium (chi-squared test, P < 0.05). ER-α mRNA level in the leiomyoma tissue was 6.4 times higher than that in the myometrial tissue in this patient (data not shown). Figure 3: View largeDownload slide DNA methylation status of the ER-α promoter region in uterine leiomyoma and normal myometrium. (A) Distribution of CpG sites in the promoter region and the first exon of ER-α gene. The position of the transcription start site is designated as +1. The diagram shows a detailed map of ∼1.5 kb region around the transcription start site (arrow), in which the ‘vertical lines’ indicate positions of CpG sites. Thick horizontal lines indicate the region identified as CpG islands. Thin horizontal lines indicate the regions analyzed by bisulfite sequencing (Regions I, II and III). (B) DNA methylation status of CpG sites in the promoter region and the first exon of the ER-α gene. Methylation status of all the CpG sites between −1188 and +229 (56 CpG sites) was analyzed by sodium bisulfite genomic sequencing in a paired sample of leiomyoma and myometrium from an individual with myoma (Case 1), normal myometrium from an individual without myoma (Case 2) and myometrium from another individual with myoma (Case 3). Open and filled circles indicate unmethylated and methylated CpG status, respectively. Figure 3: View largeDownload slide DNA methylation status of the ER-α promoter region in uterine leiomyoma and normal myometrium. (A) Distribution of CpG sites in the promoter region and the first exon of ER-α gene. The position of the transcription start site is designated as +1. The diagram shows a detailed map of ∼1.5 kb region around the transcription start site (arrow), in which the ‘vertical lines’ indicate positions of CpG sites. Thick horizontal lines indicate the region identified as CpG islands. Thin horizontal lines indicate the regions analyzed by bisulfite sequencing (Regions I, II and III). (B) DNA methylation status of CpG sites in the promoter region and the first exon of the ER-α gene. Methylation status of all the CpG sites between −1188 and +229 (56 CpG sites) was analyzed by sodium bisulfite genomic sequencing in a paired sample of leiomyoma and myometrium from an individual with myoma (Case 1), normal myometrium from an individual without myoma (Case 2) and myometrium from another individual with myoma (Case 3). Open and filled circles indicate unmethylated and methylated CpG status, respectively. Second, methylation status was analyzed for normal myometrium from an individual without myoma (Fig. 3B, Case 2). The proximal region (Regions I and II) was unmethylated, and only 3.6% in a total of 84 examined CpG sites were methylated in the distal promoter region (Region III) (Fig. 3B, Case 2). Since there was a difference in DNA methylation status between myometrium from an individual with myoma and without myoma, methylation status was further analyzed in myometrium from another individual with myoma (Fig. 3B, Case 3). The proximal region (Regions I and II) was unmethylated, and in the distal promoter region (Region III), 15.5% in a total of 84 examined CpG sites were methylated (Fig. 3B, Case 3). Relationship between DNA methylation status of the distal promoter region and mRNA expression in uterine leiomyoma and normal myometrium Since there was a variation in DNA methylation status in the distal promoter region by sodium bisulfite genomic sequencing, DNA methylation status of this region (six CpG sites; −1096 to −790), a part of the distal promoter region, was analyzed by sodium bisulfite restriction mapping in 11 paired samples of leiomyoma and myometrium. The results from the 11 patients showed two different methylation patterns. Of the 11 cases, nine showed unmethylated status in leiomyomas and a methylated status in myometrium for this region, which is represented as Pattern I in Fig. 4B. The other two cases showed methylated status in both leiomyomas and myometrium, which is represented as Pattern II in Fig. 4B. In the cases who showed Pattern I, ER-α mRNA levels tended to be higher in leiomyoma than those in myometrium (Table I). These results suggest that CpG sites in the distal region of ER-α promoter are hypomethylated in leiomyomas in most of the patients, and this may be associated with higher mRNA expression in leiomyomas than in myometrium. There seemed to be no relationship between location and size of the leiomyoma and DNA methylation status (Table I). Figure 4: View largeDownload slide DNA methylation status of the distal promoter region in uterine leiomyoma and normal myometrium. (A) Diagram of CpG sites (‘vertical lines’) and Taq I recognition sites (‘filled triangles’) in the distal promoter region. (B) Formation of fragmented DNA after Taq I treatment indicates methylated status in this region. Pattern I: unmethylated status in leiomyoma (leio) and methylated status in myometrium (myo) judging by DNA fragmentation in myometrium but not in leiomyoma. Pattern II: methylated status in both leiomyoma and myometrium judging by DNA fragmentation in both myometrium and leiomyoma. (−) undigested control without Taq I, (+) Taq I treatment. The white arrow heads indicate the fragmented DNA. Figure 4: View largeDownload slide DNA methylation status of the distal promoter region in uterine leiomyoma and normal myometrium. (A) Diagram of CpG sites (‘vertical lines’) and Taq I recognition sites (‘filled triangles’) in the distal promoter region. (B) Formation of fragmented DNA after Taq I treatment indicates methylated status in this region. Pattern I: unmethylated status in leiomyoma (leio) and methylated status in myometrium (myo) judging by DNA fragmentation in myometrium but not in leiomyoma. Pattern II: methylated status in both leiomyoma and myometrium judging by DNA fragmentation in both myometrium and leiomyoma. (−) undigested control without Taq I, (+) Taq I treatment. The white arrow heads indicate the fragmented DNA. Table I. Profile of the samples used for bisulfite restriction mapping. Patients  Age  Location of leiomyoma  Diameter of the leiomyoma (cm)  ER-α mRNA levels   BRM pattern  myo  leio  1  41  Intramural  5  6  42  I  2  49  Intramural  16  31  50  I  3  37  Intramural  8  31  47  I  4  45  Subserosal  7  103  128  I  5  55  Intramural  10  119  120  I  6  53  Intramural  15  29  220  I  7  49  Intramural  7  59  68  I  8  44  Intramural  8  116  133  I  9  50  Intramural  3  100  101  I  10  46  Intramural  15  100  80  II  11  52  Subserosal  9  147  83  II  Patients  Age  Location of leiomyoma  Diameter of the leiomyoma (cm)  ER-α mRNA levels   BRM pattern  myo  leio  1  41  Intramural  5  6  42  I  2  49  Intramural  16  31  50  I  3  37  Intramural  8  31  47  I  4  45  Subserosal  7  103  128  I  5  55  Intramural  10  119  120  I  6  53  Intramural  15  29  220  I  7  49  Intramural  7  59  68  I  8  44  Intramural  8  116  133  I  9  50  Intramural  3  100  101  I  10  46  Intramural  15  100  80  II  11  52  Subserosal  9  147  83  II  DNA methylation status of promoter region (six CpG sites; −1096 to −790), a part of the distal promoter region, was analyzed by sodium bisulfite restriction mapping (BRM) in 11 paired samples of leiomyoma (leio) and myometrium (myo). The results from 11 patients showed two different methylation patterns. Nine of the 11 cases showed unmethylated status in leiomyomas and methylated status in myometrium of this region, which is represented as Pattern I in Fig. 4B. The remaining two of the 11 cases showed methylated status in both leiomyomas and myometrium, which is represented as Pattern II in Fig. 4B. View Large Discussion The present study demonstrated that there were differences in DNA methylation status of the ER-α promoter region between uterine leiomyomas and normal myometrium, suggesting that epigenetic aberration actually occurs in uterine leiomyomas. We found DNA hypomethylation in the distal promoter region of ER-α (−1188 to −790) in a uterine leiomyoma compared with the myometrium by sodium bisulfite sequencing, and further confirmed hypomethylation status of this region in 9 patients out of the 11 patients by bisulfite restriction mapping. Moreover, in these patients who showed unmethylated status of this region in uterine leiomyomas and methylated status in myometrium, ER-α mRNA levels tended to be higher in leiomyoma than in myometrium. Thus, the pathological feature of uterine leiomyomas could be supported by our finding that the aberrant DNA hypomethylation was associated with an increased expression of ER-α that mediates sensitivity to estradiol. It is not surprising that there are cases in which DNA methylation status of the ER-α promoter region is not consistent with ER-α mRNA expression, because DNA methylation may occur heterogeneously and/or gradually and the change in DNA methylation varies among individuals. However, further studies with more samples are needed regarding the relevance of the promoter methylation pattern on ER-α mRNA expression. A number of reports have addressed the association between aberrant DNA hypermethylation of the ER-α promoter and the ER-α inactivation in a variety of neoplasms such as breast cancer (Lapidus et al., 1996; Yoshida et al., 2000; Yan et al., 2001; Berger and Daxenbichler, 2002; Giacinti et al., 2006), prostate cancer (Li et al., 2000), esophagus adenocarcinoma (Eads et al., 2000), hematopoietic neoplasms (Issa et al., 1996), brain tumors (Li et al., 1998) and colon cancer (Ahuja et al., 1998). In breast cancers, the increased incidence of DNA methylation in the CpG island in the proximal promoter or the first exon of ER-α was highly associated with the loss of ER-α expression (Lapidus et al., 1996; Iwase et al., 1999). In uterine leiomyomas, however, the DNA hypomethylated status was observed in the distal promoter region of ER-α outside the CpG island. In this regard, the epigenetic aberration that we found in uterine leiomyomas is different from those in the previous reports on other tumors or cancers. This is not surprising because it has been reported that DNA methylation of the CpG sites other than CpG islands in the promoter region regulates transcription (Razin and Cedar, 1991; Cho et al., 2001; Imamura et al., 2001). It is also suggested that DNA methylation is involved in the regulation of gene expression regardless of richness of CpGs (Shiota, 2004). The proximal promoter region and the first exon of ER-α are the most important region for ER-α expression (McPherson et al., 1997; Reid et al., 2002). The proximal region is the minimal core promoter of the ER-α gene and determines on–off switching of the ER-α transcription. On the other hand, two EREs, ERE 2 and 3, are present in the distal promoter region and can also regulate transcription of ER-α gene (Treilleux et al., 1997). In addition, the distal promoter region has been reported to contain two ER-α upstream binding factor-1 binding sites, which have a strong transcriptional enhancer activity (Cohn et al., 1999). Thus, the distal promoter region is considered as important for the modulation of ER-α transcriptional level. In fact, the present in vitro study revealed that ER-α mRNA expression was increased by 5-aza-dC that inhibits DNA methylation, suggesting that ER-α mRNA expression is under epigenetic regulation. However, further studies including promoter activity assay with methylated reporter constructs are needed to demonstrate that DNA methylation of the distal promoter region actually controls ER-α mRNA expression. It is of interest to note that DNA methylation of the distal promoter region of ER-α was observed in several sequenced clones of myometrium and that the extent of DNA methylation in this region of the myometrium varies among individuals, suggesting that DNA methylation occurs heterogeneously in the normal tissue, which may be a part of physiological changes in a certain cell type such as smooth muscle cells in myometrium. In fact, there is also a variation in DNA methylation status of the promoter region of ER-β in human endometrial stromal cells among individuals (Xue et al., 2007). Alternatively, DNA methylation seen in the myometrium may be caused by some factors that induce aberrant DNA methylation such as aging, chronic inflammation and possibly viral infection (Ushijima and Okochi-Tanaka, 2005). This is the first report demonstrating that in uterine leiomyomas there is aberrant DNA hypomethylation in the ER-α promoter, especially outside the CpG island that has been well studied in other clinical cases. Decreased mRNA expression of DNA methyltransferase-3 (DNMT-3) with genome-wide DNA hypomethylation has been reported in uterine leiomyomas compared with myometrium (Li et al., 2003). This suggests that epigenetic alterations are involved in the development of uterine leiomyomas. The aberrant hypomethylation of the ER-α gene could also be caused by the decreased DNMT-3 level. Recent data have shown that altered expression of a variety of genes contributes to pathogenesis of uterine leiomyomas (Skubitz and Skubitz, 2002; Tsibris et al., 2003; Luo et al., 2005). Taken together with our finding, potential epigenetic alterations such as aberrant DNA hypomethylation are strongly suggested to be involved in pathogenesis of uterine leiomyomas. In conclusion, there seems to be a potential link between aberrant DNA methylation level and ER-α expression in uterine leiomyomas, and this is the first example that the ER-α promoter region is aberrantly hypomethylated in human disease cases. Funding This work was supported in part by Grants-in-Aid 17791121, 18791158, 19791153 and 20591918 for Scientific Research from the Ministry of Education, Science, and Culture, Japan. 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Published by Oxford University Press on behalf of the European Society of Human Reproduction and Embryology. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org http://www.deepdyve.com/assets/images/DeepDyve-Logo-lg.png Molecular Human Reproduction Oxford University Press

Potential link between estrogen receptor-α gene hypomethylation and uterine fibroid formation

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Oxford University Press
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© The Author 2008. Published by Oxford University Press on behalf of the European Society of Human Reproduction and Embryology. All rights reserved. For Permissions, please email: journals.permissions@oxfordjournals.org
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1360-9947
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1460-2407
DOI
10.1093/molehr/gan045
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18701604
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Abstract

Abstract Uterine leiomyomas are the most common uterine tumors in women. Estrogen receptor-α (ER-α) is more highly expressed in uterine leiomyomas than in normal myometrium, suggesting a link between uterine leiomyomas and ER-α expression. DNA methylation is an epigenetic mechanism of gene regulation and plays important roles in normal embryonic development and in disease progression including cancers. Here, we investigated the DNA methylation status of the ER-α promoter region (−1188 to +229 bp) in myometrium and leiomyoma. By sodium bisulfite sequencing, 49 CpG sites in the proximal promoter region of ER-α gene were shown to be unmethylated in both leiomyoma and normal myometrium. At seven CpG sites in the distal promoter region of the ER-α gene, there was a variation in DNA methylation status in myometrium and leiomyoma. Further analysis of the DNA methylation status by bisulfite restriction mapping among 11 paired samples of myometrium and leiomyoma indicated that CpG sites in the distal region of ER-α promoter are hypomethylated in leiomyomas of nine patients. In those patients, ER-α mRNA levels tended to be higher in the leiomyoma than in the myometrium. In conclusion, there was an aberrant DNA methylation status in the promoter region of ER-α gene in uterine leiomyoma, which may be associated with high ER-α mRNA expression. DNA methylation, epigenetics, estrogen receptor-α, ER-α promoter, leiomyoma Introduction Uterine leiomyomas are the most common uterine tumors in women of reproductive age. Approximately 20–25% of women of reproductive age are afflicted with this disease (Vollenhoven et al., 1990). They frequently cause serious gynecological problems such as pelvic pain, menorrhagia, dysmenorrhea, reduced fertility and recurrent pregnancy loss (Bajekal and Li, 2000; Stewart, 2001). In addition, uterine leiomyoma is the most common indication for hysterectomy in Japan, as well as in the USA (Farquhar and Steiner, 2002). Despite the high prevalence rate and tremendous influence on reproductive women, the pathogenesis of uterine leiomyomas still remains to be elucidated. On the basis of the fact that uterine leiomyomas develop only after menarche and markedly shrink under hypoestrogenic conditions such as late menopause, it is presumed that their growth depends on estrogens (Stewart, 2001). Although the increased sensitivity to estradiol is important for the growth of uterine leiomyomas, high circulating estradiol levels are not a necessary requirement. The physiological effects of estrogen are mediated by estrogen receptors (ERs). Among them, ER-α is more highly expressed in uterine leiomyomas than in normal myometrium (Benassayag et al., 1999; Kovács et al., 2001), suggesting a possible link between uterine leiomyomas and ER-α expression level. Epigenetic mechanisms including DNA methylation and histone modification are known to play key roles in transcriptional regulation. DNA methylation occurs at cytosines within CpG dinucleotides that are clustered frequently in regions of ∼1–2 kb in length, called CpG islands, in or near promoter and first exon regions of genes (Esteller and Herman, 2002). In mammals, 60–70% of CpG sites are methylated in the genomic DNA (Boyes and Bird, 1992). DNA methylation is involved in various developmental processes by silencing, switching and stabilizing genes (Nan et al., 1998; Cho et al., 2001; Imamura et al., 2001; Li, 2002). Although there are differences in the frequency of CpGs in the gene regulatory regions, DNA methylation-dependent gene regulation has been previously reported (Razin and Cedar, 1991; Cho et al., 2001; Imamura et al., 2001). CpG methylation can down-regulate gene expression by preventing the binding of transcription factors or by recruiting repressor molecules (Bird, 1992; Ballestar and Wolffe, 2001). Accumulating evidence has indicated that increased methylation level of the CpG islands within the ER-α promoter region is highly negatively associated with ER-α expression in a variety of diseases including neoplastic and atherosclerotic lesions (Iwase et al., 1999; Post et al., 1999; Yoshida et al., 2000; Berger and Daxenbichler, 2002). Thus, down-regulation of the ER-α expression is caused by hypermethylation of the CpG islands within the ER-α promoter region. However, the fact that the ER-α expression was higher in uterine leiomyomas than in normal myometrium is different from other ER-α related diseases described earlier. This led us to assume that a different epigenetic abnormality might be involved in uterine leiomyomas. The present study was undertaken to investigate the methylation status of CpG sites within the promoter region of the human ER-α gene and to evaluate an association of aberrant DNA methylation status with ER-α gene expression in uterine leiomyomas and normal myometrium. Materials and Methods Tissue preparation Specimens of uterine leiomyomas and corresponding normal myometrium were obtained from 18 women, from 37 to 57 (mean 47.4) years of age, who underwent total hysterectomy. Normal myometrium was obtained from a woman without myoma (49 years, cervical cancer). Informed consent was obtained from all participating patients, and ethical approval was obtained from Yamaguchi University Graduate School of Medicine. Tissues were taken immediately after removal of the uterus, immersed in liquid nitrogen and stored at −80°C until used DNA/RNA extraction. For immunohistochemistry, the specimens were fixed immediately in 10% neutral formalin for ∼24 h, embedded in paraffin and cut into 4 μm thick sections. Immunohistochemistry The diagnosis of leiomyoma and normal myometrium was established on histological examination with hematoxylin and eosin staining. Immunohistochemistry was performed as described previously (Sugino et al., 2005) using an ER-α monoclonal antibody (ER1D5, mouse, Dako Japan Co. Ltd., Tokyo, Japan). Counterstaining was performed with Meyer's hematoxylin. Real-time RT–PCR analysis Total RNAs were isolated from tissues using Isogen reagent (Nippon Gene, Tokyo, Japan) and reverse-transcribed using an ExCript RT reagent kit (TaKaRa, Ohtsu, Japan) according to the manufacturer's protocol, respectively. For PCR amplification, first strand cDNA was synthesized from 1 µg total RNA with reverse transcriptase in 20 µl of reaction mixture. The oligonucleotide primers for ER-α (5′-TGTGCAATGACTATGCTTCA-3′ and 5′-GCTCTTCCTCCTGTTTTTA-3′; 149 bp amplified products) were designed from the human ER-α cDNA sequence (Matsuzaki et al., 2000). Internal control PCR primers for GAPDH (5′-AGGTGAAGGTCGGAGTCA-3′ and 5′-GGTCATTGATGGCAACAA-3′; 99 bp amplified products) were designed from the GAPDH cDNA sequence (Kaneda et al., 2004). Real-time PCR was performed using LightCycler (Roche Diagnostics, Indianapolis, IN, USA). The reaction mixture contained 10 µl SYBR Premix Ex Taq (TaKaRa), 0.2 µM each of primer sets of ER-α or GAPDH and 2 µl cDNA in a total volume of 20 µl. The thermocycling program was 40 cycles of 96°C for 5 s and 60°C for 20 s with an initial cycle of 96°C for 10 s. Sodium bisulfite genomic sequencing Genomic DNA was extracted using Genomic DNA kit (Qiagen, Tokyo, Japan) according to the manufacturer's protocol. The bisulfite reaction, in which unmethylated cytosine is converted to uracil and 5-methylcytsine remains non-reactive, was carried out as previously described (Cho et al., 2001; Imamura et al., 2001) with a slight modification: 2 µg of genomic DNA digested with Pvu II was denatured by incubation with NaOH at 42°C for 20 min. After the incubation, sodium metabisulfite and hydroquinone (Wako, Osaka, Japan) were added to the final concentrations of 2.0 M and 0.5 mM, respectively, and the mixture was incubated at 55°C for 16 h. The bisulfite reaction was terminated by incubation with NaOH again at 42°C for 20 min. The DNA fragments covering the transcriptional regulatory region of ER-α gene (−1297 to +279) were amplified by PCR using the following set of primers: Region I (−126 to +279) F: 5′-GTTGTGTTTGGAGTGATGTTTAAGTT-3′, R: 5′-CAATAAAACCATCCCAAATACTTTA-3′; Region II (−670 to −94) F: 5′-GGAAGGGTTTATTTATTTTGGGAGTA-3′, R: 5′-TAACATTAACTTAAACATCACTCC-3′; Region III (−1297 to −731) F: 5′-TTGGGTGTTTGGGATAGTAATTAAA-3′, R: 5′-CTTAATCCCATTAAAAATTCTCATA-3′. The PCR conditions were 95°C for 10 min, and 40 cycles of 94°C for 30 s, 55°C for 30 s and 72°C for 1 min, with a final extension at 72°C for 10 min. The resulting products were subjected to agarose gel electrophoresis and purified using a QIAquick gel extraction kit (Qiagen). The PCR products were cloned into pGEM-T easy vector (Promega, Tokyo, Japan), and 10 or more clones were randomly picked from each of two independent PCRs were sequenced to determine the methylation status. Sequencing was performed using an ABI automated sequencer with BigDye terminators (Applied Biosystems, Foster City, CA, USA). Bisulfite restriction mapping The bisulfite-converted DNA was amplified by PCR with a set of primers (−1120 to −645), F: 5′-TATATATATGTGTGTGTGTATGTG-3′ and R: 5′-TACTCCCAAAATAAATAAACCCTTCC-3′. One half of the PCR products were digested with 5 U of Taq I (TaKaRa) at 65°C for 3 h. The remaining half was used for undigested control without Taq I treatment. Taq I recognizes 5′-TCGA-3′ sequences. Because only unmethylated cytosine sites are changed to thymine by sodium bisulfite PCR, PCR fragments from unmethylated genomic DNA are resistant to Taq I, whereas those from methylated DNA are digested by the enzyme. The resulting products of the bisulfite restriction mapping were assessed by agarose gel electrophoresis. Cell culture, demethylation and RT–PCR analysis Primary human uterine smooth muscle cells and cell culture reagents were purchased from Cambrex, Inc. (Walkersville, MD, USA). Cells were grown to 90% confluence and maintained in SMGM2 medium, which consists of smooth muscle basal medium supplemented with 5% fetal bovine serum, 2 ng/ml recombinant human fibroblast growth factor-β, 5 µg/ml insulin, 0.5 ng/ml recombinant human epidermal growth factor, 50 µg/ml gentamicin sulfate and 50 µg/ml amphotericin B. Medium was changed every other day and all experiments were completed with cells derived from passages 2. For treatment with a demethylating agent, 5-aza-dC (Sigma, St Louis, MO, USA), that inhibits DNA methylation, cells were seeded at a density of 1 × 106 cells/25 cm2 tissue culture flask. After 24 h of incubation, cells were cultured with treatment medium containing 1 μM 5-aza-dC for 72 h. The medium was changed daily. After treatment, cells were used for RNA isolation and ER-α mRNA levels were measured by RT–PCR as reported previously (Sugino et al., 1998). Total RNAs were isolated from the cells according to the protocol mentioned above. For PCR amplification, first-strand cDNA was synthesized from 1 μg total RNA with reverse transcriptase in 20 µl of reaction mixture. The oligonucleotide primers for ER-α (5′-TGTGCAATGACTATGCTTCA-3′ and 5′-GCTCTTCCTCCTGTTTTTA-3′; 149 bp amplified products) were designed from the human ER-α cDNA sequence (Matsuzaki et al., 2000). Internal control PCR primers for ribosomal protein L19 (5′-CTGAAGGTCAAAGGGAATGTG-3′ and 5′-GGACAGAGTCTTGATGATCTC-3′; 194 bp amplified products) were designed from the L19 cDNA sequence (Sugino et al., 1998). PCR amplification was performed using a programmed temperature control system (PC808, ASTEC, Fukuoka, Japan). The reaction mixture contained 4 µl cDNA, 1 µM each of primer sets of ER-α or L19, GeneAmp 10× PCR buffer, 0.2 mM deoxynucleotide triphosphate, 2.5 mM MgCl2 and 0.05 U AmpliTaq DNA polymerase (Applied Biosystems) in a total volume of 20 µl. The thermocycling program was an initial cycle of 94°C for 5 min, then 35 cycles of 94°C for 1 min, 60°C for 1 min, 72°C for 1 min, followed by 10 min of final extension at 72°C. The resulting products were subjected to agarose gel electrophoresis. The level of ER-α expression was determined by quantifying the intensities of the PCR product, compared with the L19 product, using NIH ImageJ software. Statistical analyses Wilcoxon signed-ranks test was used for paired samples. A value of P < 0.05 was considered significant. Results ER-α expression Immunohistochemical staining for ER-α expression was localized in nuclei of smooth muscle cells, and the staining distribution was homogenous in both leiomyoma and myometrium (Fig. 1A). Figure 1: View largeDownload slide ER-α expression in uterine leiomyoma and normal myometrium. (A) Immunohistochemical staining of ER-α in leiomyoma (leio) and myometrium (myo). Immunohistochemical staining for ER-α was performed on tissue samples obtained from three different patients. HE, hematoxylin–eosin staining, Bar; 50 µm. (B) ER-α mRNA expression in leiomyoma and myometrium. Specimens of leiomyomas and corresponding myometrium were obtained from 18 women. Total RNA was isolated from 18 pairs of leiomyomas and myometrium. ER-α mRNA levels were analyzed by SYBR Green I real-time quantitative RT–PCR. Relative ER-α expression normalized to GAPDH was calculated. Values are mean ± SEM. *P < 0.01 versus myometrium. Figure 1: View largeDownload slide ER-α expression in uterine leiomyoma and normal myometrium. (A) Immunohistochemical staining of ER-α in leiomyoma (leio) and myometrium (myo). Immunohistochemical staining for ER-α was performed on tissue samples obtained from three different patients. HE, hematoxylin–eosin staining, Bar; 50 µm. (B) ER-α mRNA expression in leiomyoma and myometrium. Specimens of leiomyomas and corresponding myometrium were obtained from 18 women. Total RNA was isolated from 18 pairs of leiomyomas and myometrium. ER-α mRNA levels were analyzed by SYBR Green I real-time quantitative RT–PCR. Relative ER-α expression normalized to GAPDH was calculated. Values are mean ± SEM. *P < 0.01 versus myometrium. Many investigators have reported that ER-α is more highly expressed in leiomyomas than in myometrium (Benassayag et al., 1999; Kovács et al., 2001), suggesting a possible link between leiomyomas and ER-α expression level. To investigate whether ER-α levels are altered in leiomyoma samples that we collected, the ER-α mRNA was measured in leiomyomas and myometrium by real-time RT–PCR. As shown in Fig. 1B, ER-α mRNA levels in the samples we examined were confirmed as significantly higher in leiomyomas than in myometrium (P < 0.01). Effects of 5-aza-dC on ER-α mRNA expression in human uterine smooth muscle cells To study the possibility that ER-α mRNA expression is under epigenetic regulation such as DNA methylation, human uterine smooth muscle cells were incubated with 5-aza-dC which inhibits DNA methylation. ER-α mRNA expression in human uterine smooth muscle cells was significantly (P < 0.05) increased by 5-aza-dC (Fig. 2). Figure 2: View largeDownload slide Effects of 5-aza-dC on ER-α mRNA expression in human uterine smooth muscle cells. Primary human uterine smooth muscle cells were incubated with a demethylating agent, 5-aza-dC (1 µM) that inhibits DNA methylation, for 72 h. After treatment, cells were used for RNA isolation and ER-α mRNA levels were measured by RT–PCR. The intensity of the signals of ER-α was normalized to that of the internal control L19 (the ratio of ER-α to L19). Data were expressed as a percentage of the control value in each incubation. Each bar represents the mean ± SEM of three different experiments. *P < 0.05 versus control. Figure 2: View largeDownload slide Effects of 5-aza-dC on ER-α mRNA expression in human uterine smooth muscle cells. Primary human uterine smooth muscle cells were incubated with a demethylating agent, 5-aza-dC (1 µM) that inhibits DNA methylation, for 72 h. After treatment, cells were used for RNA isolation and ER-α mRNA levels were measured by RT–PCR. The intensity of the signals of ER-α was normalized to that of the internal control L19 (the ratio of ER-α to L19). Data were expressed as a percentage of the control value in each incubation. Each bar represents the mean ± SEM of three different experiments. *P < 0.05 versus control. DNA methylation status of 5′-flanking region of ER-α gene Since ER-α mRNA expression seemed to be under the regulation of DNA methylation, we examined DNA methylation status of the ER-α promoter region between leiomyoma and myometrium. The 5′-upstream region around the first exon (between about −500 and +200 bp) of the ER-α gene is most importantly involved in the regulation of ER-α expression (Lapidus et al., 1996; Yan et al., 2001; Giacinti et al., 2006). Furthermore, a series of three estrogen response elements (EREs), which lie from −892 to −420 in the ER-α upstream region, is identified as a region with functional importance for ER-α gene transcription (Furguson et al., 1997; Li et al., 2000). Therefore, ∼1 kb upstream region together with a part of the first exon of the ER-α gene is considered as important for ER-α gene transcription, and the methylation status of this region was compared between leiomyoma and myometrium. Figure 3A shows the distribution of CpGs within the transcriptional regulatory region of the human ER-α gene. The ER-α gene has two CpG islands in the promoter region and in the first exon. According to the registered nucleotide sequence of the ER-α (GenBank accession no. AB090237), there were 56 CpG sites in the 1.5 kb genomic sequence including the promoter and exon 1 of the human ER-α gene (Fig. 3A). In the present study, these CpG sites were divided into two regions, the proximal and distal regions. The proximal region included 49 CpG sites around the transcription start site (−556 to +229, designed as Regions I and II), and the distal region included seven CpG sites in the upstream region (−1188 to −790, designed as Region III). First, the methylation status of all the CpG sites between −1188 and +229 (56 CpG sites) were analyzed by sodium bisulfite genomic sequencing for a paired sample of leiomyoma and myometrium (Fig. 3B, Case 1). The proximal region (Regions I and II containing a total of 49 CpG sites) was unmethylated in both tissues (Fig. 3B, Case 1). In the distal promoter region (Region III, seven CpG sites), 47 CpG sites (24.7%) in a total of 190 examined CpG sites were methylated in myometrium, whereas only nine CpG sites (4.9%) in a total of 184 examined CpG sites were methylated in leiomyoma (Fig. 3B, Case 1). There was a significant difference in the frequency of DNA methylation between leiomyoma and myometrium (chi-squared test, P < 0.05). ER-α mRNA level in the leiomyoma tissue was 6.4 times higher than that in the myometrial tissue in this patient (data not shown). Figure 3: View largeDownload slide DNA methylation status of the ER-α promoter region in uterine leiomyoma and normal myometrium. (A) Distribution of CpG sites in the promoter region and the first exon of ER-α gene. The position of the transcription start site is designated as +1. The diagram shows a detailed map of ∼1.5 kb region around the transcription start site (arrow), in which the ‘vertical lines’ indicate positions of CpG sites. Thick horizontal lines indicate the region identified as CpG islands. Thin horizontal lines indicate the regions analyzed by bisulfite sequencing (Regions I, II and III). (B) DNA methylation status of CpG sites in the promoter region and the first exon of the ER-α gene. Methylation status of all the CpG sites between −1188 and +229 (56 CpG sites) was analyzed by sodium bisulfite genomic sequencing in a paired sample of leiomyoma and myometrium from an individual with myoma (Case 1), normal myometrium from an individual without myoma (Case 2) and myometrium from another individual with myoma (Case 3). Open and filled circles indicate unmethylated and methylated CpG status, respectively. Figure 3: View largeDownload slide DNA methylation status of the ER-α promoter region in uterine leiomyoma and normal myometrium. (A) Distribution of CpG sites in the promoter region and the first exon of ER-α gene. The position of the transcription start site is designated as +1. The diagram shows a detailed map of ∼1.5 kb region around the transcription start site (arrow), in which the ‘vertical lines’ indicate positions of CpG sites. Thick horizontal lines indicate the region identified as CpG islands. Thin horizontal lines indicate the regions analyzed by bisulfite sequencing (Regions I, II and III). (B) DNA methylation status of CpG sites in the promoter region and the first exon of the ER-α gene. Methylation status of all the CpG sites between −1188 and +229 (56 CpG sites) was analyzed by sodium bisulfite genomic sequencing in a paired sample of leiomyoma and myometrium from an individual with myoma (Case 1), normal myometrium from an individual without myoma (Case 2) and myometrium from another individual with myoma (Case 3). Open and filled circles indicate unmethylated and methylated CpG status, respectively. Second, methylation status was analyzed for normal myometrium from an individual without myoma (Fig. 3B, Case 2). The proximal region (Regions I and II) was unmethylated, and only 3.6% in a total of 84 examined CpG sites were methylated in the distal promoter region (Region III) (Fig. 3B, Case 2). Since there was a difference in DNA methylation status between myometrium from an individual with myoma and without myoma, methylation status was further analyzed in myometrium from another individual with myoma (Fig. 3B, Case 3). The proximal region (Regions I and II) was unmethylated, and in the distal promoter region (Region III), 15.5% in a total of 84 examined CpG sites were methylated (Fig. 3B, Case 3). Relationship between DNA methylation status of the distal promoter region and mRNA expression in uterine leiomyoma and normal myometrium Since there was a variation in DNA methylation status in the distal promoter region by sodium bisulfite genomic sequencing, DNA methylation status of this region (six CpG sites; −1096 to −790), a part of the distal promoter region, was analyzed by sodium bisulfite restriction mapping in 11 paired samples of leiomyoma and myometrium. The results from the 11 patients showed two different methylation patterns. Of the 11 cases, nine showed unmethylated status in leiomyomas and a methylated status in myometrium for this region, which is represented as Pattern I in Fig. 4B. The other two cases showed methylated status in both leiomyomas and myometrium, which is represented as Pattern II in Fig. 4B. In the cases who showed Pattern I, ER-α mRNA levels tended to be higher in leiomyoma than those in myometrium (Table I). These results suggest that CpG sites in the distal region of ER-α promoter are hypomethylated in leiomyomas in most of the patients, and this may be associated with higher mRNA expression in leiomyomas than in myometrium. There seemed to be no relationship between location and size of the leiomyoma and DNA methylation status (Table I). Figure 4: View largeDownload slide DNA methylation status of the distal promoter region in uterine leiomyoma and normal myometrium. (A) Diagram of CpG sites (‘vertical lines’) and Taq I recognition sites (‘filled triangles’) in the distal promoter region. (B) Formation of fragmented DNA after Taq I treatment indicates methylated status in this region. Pattern I: unmethylated status in leiomyoma (leio) and methylated status in myometrium (myo) judging by DNA fragmentation in myometrium but not in leiomyoma. Pattern II: methylated status in both leiomyoma and myometrium judging by DNA fragmentation in both myometrium and leiomyoma. (−) undigested control without Taq I, (+) Taq I treatment. The white arrow heads indicate the fragmented DNA. Figure 4: View largeDownload slide DNA methylation status of the distal promoter region in uterine leiomyoma and normal myometrium. (A) Diagram of CpG sites (‘vertical lines’) and Taq I recognition sites (‘filled triangles’) in the distal promoter region. (B) Formation of fragmented DNA after Taq I treatment indicates methylated status in this region. Pattern I: unmethylated status in leiomyoma (leio) and methylated status in myometrium (myo) judging by DNA fragmentation in myometrium but not in leiomyoma. Pattern II: methylated status in both leiomyoma and myometrium judging by DNA fragmentation in both myometrium and leiomyoma. (−) undigested control without Taq I, (+) Taq I treatment. The white arrow heads indicate the fragmented DNA. Table I. Profile of the samples used for bisulfite restriction mapping. Patients  Age  Location of leiomyoma  Diameter of the leiomyoma (cm)  ER-α mRNA levels   BRM pattern  myo  leio  1  41  Intramural  5  6  42  I  2  49  Intramural  16  31  50  I  3  37  Intramural  8  31  47  I  4  45  Subserosal  7  103  128  I  5  55  Intramural  10  119  120  I  6  53  Intramural  15  29  220  I  7  49  Intramural  7  59  68  I  8  44  Intramural  8  116  133  I  9  50  Intramural  3  100  101  I  10  46  Intramural  15  100  80  II  11  52  Subserosal  9  147  83  II  Patients  Age  Location of leiomyoma  Diameter of the leiomyoma (cm)  ER-α mRNA levels   BRM pattern  myo  leio  1  41  Intramural  5  6  42  I  2  49  Intramural  16  31  50  I  3  37  Intramural  8  31  47  I  4  45  Subserosal  7  103  128  I  5  55  Intramural  10  119  120  I  6  53  Intramural  15  29  220  I  7  49  Intramural  7  59  68  I  8  44  Intramural  8  116  133  I  9  50  Intramural  3  100  101  I  10  46  Intramural  15  100  80  II  11  52  Subserosal  9  147  83  II  DNA methylation status of promoter region (six CpG sites; −1096 to −790), a part of the distal promoter region, was analyzed by sodium bisulfite restriction mapping (BRM) in 11 paired samples of leiomyoma (leio) and myometrium (myo). The results from 11 patients showed two different methylation patterns. Nine of the 11 cases showed unmethylated status in leiomyomas and methylated status in myometrium of this region, which is represented as Pattern I in Fig. 4B. The remaining two of the 11 cases showed methylated status in both leiomyomas and myometrium, which is represented as Pattern II in Fig. 4B. View Large Discussion The present study demonstrated that there were differences in DNA methylation status of the ER-α promoter region between uterine leiomyomas and normal myometrium, suggesting that epigenetic aberration actually occurs in uterine leiomyomas. We found DNA hypomethylation in the distal promoter region of ER-α (−1188 to −790) in a uterine leiomyoma compared with the myometrium by sodium bisulfite sequencing, and further confirmed hypomethylation status of this region in 9 patients out of the 11 patients by bisulfite restriction mapping. Moreover, in these patients who showed unmethylated status of this region in uterine leiomyomas and methylated status in myometrium, ER-α mRNA levels tended to be higher in leiomyoma than in myometrium. Thus, the pathological feature of uterine leiomyomas could be supported by our finding that the aberrant DNA hypomethylation was associated with an increased expression of ER-α that mediates sensitivity to estradiol. It is not surprising that there are cases in which DNA methylation status of the ER-α promoter region is not consistent with ER-α mRNA expression, because DNA methylation may occur heterogeneously and/or gradually and the change in DNA methylation varies among individuals. However, further studies with more samples are needed regarding the relevance of the promoter methylation pattern on ER-α mRNA expression. A number of reports have addressed the association between aberrant DNA hypermethylation of the ER-α promoter and the ER-α inactivation in a variety of neoplasms such as breast cancer (Lapidus et al., 1996; Yoshida et al., 2000; Yan et al., 2001; Berger and Daxenbichler, 2002; Giacinti et al., 2006), prostate cancer (Li et al., 2000), esophagus adenocarcinoma (Eads et al., 2000), hematopoietic neoplasms (Issa et al., 1996), brain tumors (Li et al., 1998) and colon cancer (Ahuja et al., 1998). In breast cancers, the increased incidence of DNA methylation in the CpG island in the proximal promoter or the first exon of ER-α was highly associated with the loss of ER-α expression (Lapidus et al., 1996; Iwase et al., 1999). In uterine leiomyomas, however, the DNA hypomethylated status was observed in the distal promoter region of ER-α outside the CpG island. In this regard, the epigenetic aberration that we found in uterine leiomyomas is different from those in the previous reports on other tumors or cancers. This is not surprising because it has been reported that DNA methylation of the CpG sites other than CpG islands in the promoter region regulates transcription (Razin and Cedar, 1991; Cho et al., 2001; Imamura et al., 2001). It is also suggested that DNA methylation is involved in the regulation of gene expression regardless of richness of CpGs (Shiota, 2004). The proximal promoter region and the first exon of ER-α are the most important region for ER-α expression (McPherson et al., 1997; Reid et al., 2002). The proximal region is the minimal core promoter of the ER-α gene and determines on–off switching of the ER-α transcription. On the other hand, two EREs, ERE 2 and 3, are present in the distal promoter region and can also regulate transcription of ER-α gene (Treilleux et al., 1997). In addition, the distal promoter region has been reported to contain two ER-α upstream binding factor-1 binding sites, which have a strong transcriptional enhancer activity (Cohn et al., 1999). Thus, the distal promoter region is considered as important for the modulation of ER-α transcriptional level. In fact, the present in vitro study revealed that ER-α mRNA expression was increased by 5-aza-dC that inhibits DNA methylation, suggesting that ER-α mRNA expression is under epigenetic regulation. However, further studies including promoter activity assay with methylated reporter constructs are needed to demonstrate that DNA methylation of the distal promoter region actually controls ER-α mRNA expression. It is of interest to note that DNA methylation of the distal promoter region of ER-α was observed in several sequenced clones of myometrium and that the extent of DNA methylation in this region of the myometrium varies among individuals, suggesting that DNA methylation occurs heterogeneously in the normal tissue, which may be a part of physiological changes in a certain cell type such as smooth muscle cells in myometrium. In fact, there is also a variation in DNA methylation status of the promoter region of ER-β in human endometrial stromal cells among individuals (Xue et al., 2007). Alternatively, DNA methylation seen in the myometrium may be caused by some factors that induce aberrant DNA methylation such as aging, chronic inflammation and possibly viral infection (Ushijima and Okochi-Tanaka, 2005). This is the first report demonstrating that in uterine leiomyomas there is aberrant DNA hypomethylation in the ER-α promoter, especially outside the CpG island that has been well studied in other clinical cases. Decreased mRNA expression of DNA methyltransferase-3 (DNMT-3) with genome-wide DNA hypomethylation has been reported in uterine leiomyomas compared with myometrium (Li et al., 2003). This suggests that epigenetic alterations are involved in the development of uterine leiomyomas. The aberrant hypomethylation of the ER-α gene could also be caused by the decreased DNMT-3 level. Recent data have shown that altered expression of a variety of genes contributes to pathogenesis of uterine leiomyomas (Skubitz and Skubitz, 2002; Tsibris et al., 2003; Luo et al., 2005). Taken together with our finding, potential epigenetic alterations such as aberrant DNA hypomethylation are strongly suggested to be involved in pathogenesis of uterine leiomyomas. In conclusion, there seems to be a potential link between aberrant DNA methylation level and ER-α expression in uterine leiomyomas, and this is the first example that the ER-α promoter region is aberrantly hypomethylated in human disease cases. Funding This work was supported in part by Grants-in-Aid 17791121, 18791158, 19791153 and 20591918 for Scientific Research from the Ministry of Education, Science, and Culture, Japan. 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Molecular Human ReproductionOxford University Press

Published: Aug 13, 2008

Keywords: Keywords DNA methylation epigenetics estrogen receptor-α ER-α promoter leiomyoma

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