1 Departments of Nutritional Sciences and 2 Medicine, University of Toronto, Toronto, Ontario, Canada, M5S 1A8, 3 Division of Gastroenterology, St Michael's Hospital, University of Toronto, Toronto, Ontario, Canada, M5B 1W8, 4 Department of Food Science, National Chung Hsing University, Taichung, Taiwan and 5 Department of Nutritional Sciences and Toxicology, University of California, Berkeley, CA 94720, USA
* To whom requests for reprints should be addressed at: Room 7258, Medical Sciences Building, University of Toronto, 1 King's College Circle, Toronto, Ontario, Canada, M5S 1A8, Tel: +1 416 978 1183; Fax: +1 416 978 8765; Email: youngin.kim{at}utoronto.ca
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Abstract |
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Abbreviations: CpG, cytosine-guanine dinucleotide sequences; DNMT, DNA methyltransferase; SAH, S-adenosylhomocysteine; SAM, S-adenosylmethionine
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Introduction |
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The methylation of cytosine located within the cytosine-guanine (CpG) dinucleotide sequences is a heritable, tissue and species-specific, post-synthetic, epigenetic modification of mammalian DNA (11,12). DNA methylation is an important epigenetic determinant in gene expression (an inverse relationship), maintenance of DNA integrity and stability, chromatin modifications and development of mutations (11,12). Neoplastic cells simultaneously harbor widespread genomic hypomethylation and more specific regional areas of hypermethylation (11,12). Genomic hypomethylation is an early and consistent event in the development of several cancers, including colorectal cancer (11,12), and is associated with genomic instability (13) and increased mutations (14). In contrast, site-specific hypermethylation at promoter CpG islands of tumor suppressor and mismatch repair genes is an important mechanism in gene silencing in carcinogenesis (15,16).
Diets deficient in different combinations of methyl group donors (choline, folate, methionine and vitamin B12) have been consistently observed to induce genomic and protooncogene (c-myc, c-fos, c-Ha-ras) DNA hypomethylation and elevated steady-state levels of corresponding mRNAs (1722) and site-specific p53 hypomethylation (2224) in rat liver. Methyl group donor deficiency has also been shown to upregulate the CpG DNA methyltransferase (DNMT) in rat liver (18,19,2325). Isolated folate deficiency appears to induce genomic and gene-specific DNA hypomethylation in rat liver (8,26), although this depends on the severity and duration of folate depletion (8,27). In contrast, isolated folate deficiency seems to have no significant effect on genomic and gene-specific DNA methylation in the rat colon (2734). One exception is that an isolated folate deficiency in conjunction with an alkylating agent may induce site-specific p53 hypomethylation in the rat colon (35). Some animal studies have suggested that an isolated folate deficiency may induce a transient genomic DNA hypermethylation in the rat colon (34) similar to the observation made in rat or mouse liver (8,36). In humans, folate depletion significantly decreases genomic DNA methylation (37,38) and folate supplementation can normalize pre-existing DNA hypomethylation (39) in peripheral leukocytes. However, there is no evidence that folate deficiency in humans induces significant aberrant genomic or gene-specific DNA methylation changes in the colon (40).
Although aberrant patterns and dysregulation of DNA methylation have been proposed as a leading mechanism by which folate depletion enhances the development of colorectal cancer, currently available data from animal models and human studies pertaining to the effect of isolated folate deficiency on methionine cycle intermediates (i.e. SAM and SAH) and DNA methylation in the colorectum are inconsistent (40). This is partly because of the lack of in vitro models of folate deficiency, imperfect animal models of folate deficiency currently available (e.g. species differences, different diet compositions, variable dose, time and duration of folate manipulations), incomplete understanding of the confounding effects of other methyl groups donors and the inability to precisely determine DNA methylation with currently available techniques (40). A mechanistic understanding of how folate status modulates colorectal carcinogenesis further strengthens the case for a causal relationship and provides insight into a possible chemopreventive role of folate. Therefore, it is important to clearly elucidate whether isolated folate deficiency would induce dysregulation and aberrant patterns of DNA methylation, in order to establish a mechanistic link between folate deficiency and colorectal carcinogenesis. In this study, we examined the effects of an isolated folate deficiency on methionine cycle intermediates, genomic and gene and site-specific DNA methylation and CpG DNMT in an in vitro model of folate deficiency, using both transformed and untransformed human and other mammalian cell lines.
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Materials and methods |
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Intracellular folate assay
Intracellular folate concentrations were determined by a standard microtiter plate assay using Lactobacillus casei as described previously (41). All analyses were performed in triplicate and repeated using three different cell lysates.
Deoxyuridine suppression test
Deoxyuridine suppression test was used to verify that the intracellular folate depletion was functionally significant as described previously (42). Deoxyuridine suppression test assesses the de novo synthesis of thymidylate on the basis of the competition between two pathways: the salvage pathway and the de novo pathway (42). The salvage pathway consists of phosphorylation of thymidine by thymidine kinase. The de novo pathway generates thymidylate by methylating deoxyuridine monophosphate. Since the enzyme for the latter reaction, thymidylate synthase, requires methylenetetrahydrofolate as a substrate, deoxyuridine suppression test has been used as a functional assay for determining folate status at the cellular level, including the colonic epithelial cells (42). In folate-replete cells, the incorporation of [3H]thymidine into DNA is suppressed by exogenous deoxyuridine, whereas in folate-deficient cells, the degree of suppression is less pronounced because of an impaired de novo synthesis of thymidylate and greater use of the salvage pathway (i.e. higher [3H]thymidine incorporation) (42). Each experiment was performed in triplicate and repeated using three different cell lysates.
Determination of SAM and SAH concentrations
SAM and SAH were determined by reversed-phase high-performance liquid chromatography by a modification of the previously described procedure (43). The cells were centrifuged and washed with cold phosphate-buffered saline (PBS) twice while being kept on ice. PBS was carefully aspirated and the cell pellets were resuspended in 150 µl 0.4 M ice-cold perchloric acid. The cell pellets were hand-homogenized on ice with a hand-held mini pestle. Homogenates were centrifuged at 10 000 g for 10 min at 4°C and the supernatants were collected and stored at 80°C until they were analyzed. The supernatant of each sample was filtered through 0.45 µM (millipore) and then loaded onto a C18 column (250 x 4.6 mm) fitted with a matched guard column operated by a Variant Vista 5500 chromatography system connected to an ultraviolet detector. Absorption of eluted compounds was monitored using ex = 254 nm. A two-buffer elution system was used: mobile phase A and B both contain 10 mM ammonium formate, 4 mM 1-heptanesulfonic acid (pH 4). Mobile phase B containing 50% acentonitrile by volume. Elution of SAM and SAH was achieved at a flow rate of 1 ml/min with the following parameters: 00.5 min, 100% A; 0.520 min, linear gradient to 75% A and 25% B; 2030 min, 25% B; 3045 min, 100% A. Chromatograms were recorded with a Hewlett-Packard HP3394 integrator with quantification accomplished by automatic peak area integration. SAM and SAH standards were used to identify the elution peaks and for the preparation of the standard curve. SAM and SAH values were normalized to cellular protein content that were determined using the LowryBensadoun method (44). All analyses were performed in triplicate and repeated using three different cell lysates.
Genomic DNA isolation
Total genomic DNA was extracted by a standard technique using proteinase K followed by organic extraction (45). The size of DNA estimated by agarose gel electrophoresis was >20 kb in all instances. No RNA contamination was detected on agarose gel electrophoresis. The final preparations had a ratio of A260:A280 between 1.8 and 2.0. The concentration of each DNA sample was determined as the mean of three independent spectrophotometric readings.
Genomic DNA methylation
The methylation status of CpG sites in genomic DNA was determined by the in vitro methyl acceptance assay using [3H-methyl]SAM (NEN Life Sciences) as a methyl donor and a prokaryotic CpG DNMT, Sss1 (New England Biolabs, Beverly, MA), as described previously (26,34). The manner in which this assay is performed produces a reciprocal relationship between the endogenous DNA methylation status and exogenous [3H-methyl] incorporation. All analyses were performed in duplicate.
Satellite DNA methylation analysis
CpG methylation status in highly repetitive satellite DNA sequences at the centromeric and juxtacentromeric regions was determined by a methyl-sensitive restriction digestion method as described previously (46). MspI and HpaII are isoschizomers which cleave the sequence, 5'-C^CGG-3'. HpaII will cleave the sequence only when the internal cytosine is unmethylated, while MspI will cleave irrespective of the methylation status. BstBI cleaves the sequence 5'-TT^CGAA-3' if the cytosine is unmethylated. Digested DNA was separated on a 1% agarose gel and transferred to a nylon membrane (Roche Diagnostics, Laval, Quebec, Canada) using a Southern blot technique under standard conditions (36). A mouse centromeric minor satellite repeat sequence derived from plasmid MR150 (pMR 150; generously provided by Dr Janet Rossant, Samuel Lunenfeld Research Institute, Toronto, Ontario, Canada) and human satellite oligonucleotide probes derived from classical satellite 2 and 3 sequences (S.2, S.3) at the juxtacentromeric region of chromosomes 1, 9, 15 and 16 and from alphoid satellite sequences (A.0) at the centromeres of chromosomes 3, 4, 7, 10, 12, 17, 18, 22 and X were used to hybridize the restricted DNA fragments in DIG Easy Hybridization Solution (Roche Diagnostics), according to the manufacturer's protocol. The hybridized membranes were washed and following conjugation with anti-DIG antibody (Roche Diagnostics), digestion patterns were detected with CSPD chemiluminescent substrate (Roche Diagnostics), according to the manufacturer's protocol. pMR150 was labeled with digoxigenin-11-dUTP using the PCR DIG Probe Synthesis kit (Roche Diagnostics) and S.2, S.3 and A.0 probes were 3' end labeled with digoxigenin-11-ddUTP by terminal transferase (Roche Diagnostics), according to the manufacturer's protocol.
As a positive control, control RPMI 1640 medium supplemented with 5 µM 5-aza-2'-deoxycytidine (5-aza-dC) (Sigma Aldrich), a demethylating agent, was added to log-phase cells every 24 h over a 72-h period before isolating DNA as described previously (46). 5-Aza-dC incorporates into DNA and results in an irreversible binding of the DNMT enzymes, thereby leading to DNA demethylation (47). Genomic DNA from 5-aza-dC treated cells grown in the control medium was analyzed for satellite DNA methylation using the methyl-sensitive restriction digestion method as described above.
Sodium bisulfite-sequencing assay
The methylation status of individual CpG sites within the promoter CpG island of the human MLH1, Estrogen Receptor (ER), p16INK4a genes was determined by the sodium bisulfite-sequencing assay as described previously (34). This method is based on the fact that the treatment of denatured DNA with sodium bisulfite converts all the cytosine residues to uracil, which are then amplified as thymines in the PCR reactions (48). In contrast, 5-methylcytosine is resistant to bisulfite deamination under the reaction conditions and is amplified as cytosine (48). Sequencing of bisulfite-modified DNA thus allows the positive identification of all methylated cytosine residues within a defined gene sequence (48). After modification with bisulfite, the top strand of the bisulfite-treated DNA was PCR amplified for each of the promoter CpG islands of the human MLH1, ER, and p16INK4a genes as described previously (34), using the following primers: MLH1: 5'-GAT TTT TAT TTT GTT TTT TTT GGG-3' (forward) and 5'-AAA ATA CCT TCA ACC AAT CAC CTC AAT AAC-3' (reverse); ER: 5'-TGT GTT TAA ATA TTG TAA TAT TGG GGG -3' (forward) and 5'-AAA AAA TAC CCT ATA CTT TCT ACT ACC-3' (reverse); and p16INK4a: 5'-GAT TTT TTA AAA GGA ATT TTT TGA ATT AGG-3' (forward) and 5'-CAC CCT CTA ATA ACC AAC CAA CCC-3' (reverse). These primers were specifically designed to amplify the sodium bisulfite-modified template based on the published sequences, according to the recommendations of Clark and Frommer (48) and synthesized by ACGT Corporation (Toronto, Ontario, Canada). The product from the first PCR reaction was re-amplified by PCR using the following nested primers under the same conditions (34): MLH1: 5'-ACA CTC GAA TTC GGG AGG TTA TAA GAG TAG GG-3' (forward) and 5'-CTC ACA CTC GAG ACT ATT AAT TAA ACA ACT TAA ATA CCA ATC-3' (reverse); ER: 5'-ACA CTC GAA TTC TTT TAG TAA TTG TAT AGT GTT TTA GGG-3' (forward) and 5'-CTC ACA CTC GAG CAA ACT TAC TAT AAA TCA TAA TCT TAC-3' (reverse); and p16INK4a: 5'-ACA CTC GAA TTC GGT GGG GTT TTT ATA ATT AGG AAA G-3' (forward) and 5'-CTC ACA CTC GAG CTA TCC CTC AAA TCC TCT AAA AAA ACC-3' (reverse). The sequences of the nested primers for the second PCR reaction, which contain flanking sequences of EcoRI and XhoI restriction sites on the sense and antisense primers, respectively, to facilitate subcloning into a vector, was constructed based on the published sequence and synthesized by ACGT.
The PCR products from the second PCR reaction were gel purified using the Qiaex II Agarose Gel Extraction Kit (Qiagen, Mississauga, Ontario, Canada), according to the manufacturer's protocol, re-extracted and dissolved in 50 µl of double-distilled H2O. The PCR products were subcloned into pBluescript II KS(+) vector (Stratagene, Cambridge, UK) at EcoRI and XhoI sites. Over 100 subclones were screened for each sample and 20 positives were sequenced using the Dideoxy Terminator Label Cycle Sequencing Kit (Applied Biosystems, Foster City, CA) and an Applied Biosystems 373 sequencer (Applied Biosystems) as described previously (34) to yield the final percent methylation results. In all reactions, the bisulfite-mediated deamination of non-methylated cytosines to uracil was >95% efficient and methylated cytosine remained >95% resistant to deamination under these conditions.
Western blot analysis
Cell lysates containing 100 µg of cellular protein were separated on an 8% SDS-PAGE and transferred onto nitrocellulose membranes in Trans-Blot® transfer medium (Bio-Rad). The membranes were blocked with PBS containing 5% skim milk for 2 h at room temperature. To detect DNMT1 protein (200 kDa) expression, the membranes were incubated with a rabbit polyclonal antibody against human and mouse DNMT1 (New England Biolabs) at a dilution of 1:2000. To detect DNMT3a protein (120 kDa) expression, the membranes were incubated with a mouse monoclonal antibody against human and mouse DNMT3a (Imgenex, San Diego, CA) at a concentration of 2 µg/µl. To detect ER protein (67 kDa), the membranes were incubated with a rabbit polyclonal antibody against human ER, C-terminus antibody (Research Diagnostics, Inc., Flanders, NJ) at a dilution of 1:200. The DNMT1, DNMT3a and ER proteins were visualized by an enhanced chemiluminescence system (Amersham Pharmacia Biotech, Piscataway, NJ). To confirm that the proteins were loaded equally, the membranes were stripped and reprobed with a human or mouse anti-ß-actin antibody (Sigma Aldrich) at a dilution of 1:3000. Western analyses were repeated using three different cell lysates. Densitometry of bands were determined using the public domain ScionImage program available on the Internet at http://rsb.info.nih.gov/nih-image.
Methyltransferase enzyme activity assay
DNMT activity was measured by incubating cell lysates containing 10 µg of protein with 0.5 µg of poly[d(IC).d(IC)] template (Amersham Pharmacia Biotech) and 3 µCi [3H]SAM (NEN Life Sciences) for 2 h at 37°C as described previously (49). The reaction was terminated, and the DNA template was purified by organic extraction and ethanol precipitation. The pellets were resuspended in 30 µl of 0.3 M NaOH, incubated at 37°C for 1 h, spotted onto GF/C Whatman filter papers, and processed for liquid scintillation counting. DNMT activity was determined for five cellular lysates obtained from each treatment (control and folate-deficient). Each reaction was performed in triplicate and the assay was repeated three times.
Statistics
Comparisons of means between the control and folate-deficient groups were determined using the Student's t-test. Statistical analyses were performed using SigmaStat 2.03 for Windows (Access Softek Inc., San Rafael, CA). Results are expressed as mean ± SD. Logistic regression was performed to determine if there were any site-specific alterations in the CpG DNA methylation owing to folate deficiency. The Fisher's exact test was used to analyze sites where the logistic regression failed as a result of one group of clones in either treatment (control or folate-deficient) being either completely methylated or unmethylated, thereby preventing a percent methylation to be calculated. These analyses were performed using SPSS 10.0 for Windows (SPSS Inc., Chicago, IL). For all analyses, the results were considered statistically significant if two-tailed P-values were <0.05.
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Results |
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Effect of folate deficiency on intracellular folate concentrations
Intracellular folate concentrations of both untransformed and transformed cells cultured in folate-deficient medium were significantly lower (by 8898%) than those of the corresponding cells cultured in folate-sufficient medium (P < 0.002; Table I). Despite the complete absence of folic acid in the medium, the folate-deficient cells demonstrated measurable levels of intracellular folate (Table I). To determine whether the observed degree of intracellular folate depletion in the folate-deficient cells was functionally significant, the deoxyuridine suppression test was performed. The folate-deficient cells were significantly less suppressed by exogenous deoxyuridine (by 24% in NIH/3T3 cells; by 30% in CHO-K1 cells; by 18% in HCT116 cells; and by 7% in Caco-2 cells; P < 0.01), resulting in significantly higher [3H]thymidine incorporation into DNA, compared with the corresponding folate-sufficient cells, suggesting significant functional intracellular folate depletion in the folate-deficient cells. This effect was abolished by preincubation with folinic acid.
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Effect of folate deficiency on genomic DNA methylation
The effect of folate deficiency on genomic DNA methylation was specific for the stage of transformation. In both untransformed NIH/3T3 and CHO-K1 cell lines, the extent of exogenous [3H-methyl] incorporation into genomic DNA was 20% higher in the folate-deficient cells than in the corresponding folate-sufficient cells (P < 0.002; Table II), suggesting a significantly lower degree of genomic DNA methylation in the folate-deficient cells, compared with the folate-sufficient cells. In contrast, in the transformed HCT116 and Caco-2 colon cancer cell lines, there was no significant difference in genomic DNA methylation between the folate-deficient and sufficient cells (Table II). The effect of folate deficiency on genomic DNA methylation also appears to be independent of the SAMSAH pathway. Despite significantly higher SAM (by 15%) and lower SAH (by 40%) concentrations and a higher SAM:SAH ratio (by 90%), the folate-deficient NIH/3T3 cells had a significantly lower extent of genomic DNA methylation than the folate-sufficient cells. In contrast, the observed genomic DNA hypomethylation in the folate-deficient CHO-K1 cells occurred in the setting of 28% lower SAM concentrations and a 31% lower SAM: SAH ratio and no significant change in SAH concentrations when compared with the folate-sufficient cells.
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Effect of folate deficiency on promoter CpG island methylation
The effect of folate deficiency on promoter CpG island methylation was determined in three prototypic tumor suppressor (ER, p16) and mismatch repair (MLH1) genes that are silenced in association with CpG-island methylation (11,12). In both HCT116 and Caco-2 cells, the promoter CpG island region (445 to 164 bp) of MLH1 was almost completely unmethylated, and folate deficiency did not alter this methylation pattern (data not shown). In contrast, the promoter CpG island region of p16 was almost completely methylated in both HCT116 and Caco-2 cells, and folate deficiency did not change this methylation pattern (data not shown). The effect of folate deficiency on the promoter CpG island region (695 bp to start) of ER appeared to be site-specific in both HCT116 and Caco-2 cells (Figure 2AD). In HCT116 cells, all the 13 CpG sites were 70100% methylated and no significant difference in methylation was observed at each CpG site except for site 10 between the folate-deficient and sufficient cells (Figure 2A and C). The folate-deficient HCT116 cells were slightly hypermethylated at site 10 compared with the folate-sufficient cells (Figure 2C; 100 and 75%, respectively; P = 0.047). In Caco-2 cells, all the 13 CpG sites were 60100% methylated and no significant difference in methylation was observed at each CpG site except for the sites 2, 6 and 11 between the folate-deficient and sufficient cells (Figure 2B and D). Folate deficiency induced non-significant hypermethylation at site 2 (95 and 75%, respectively; P = 0.066) and significant hypomethylation at site 6 (75 and 100%, respectively; P = 0.047) and site 11 (75 and 100%, respectively; P = 0.047) (Figure 2D).
Although the observed methylation changes in the promoter CpG island region of ER appeared to very modest, prior studies have suggested that methylation changes of one or two target CpG sites is sufficient to alter the activity of a promoter (50) and that methylation of as few as 7% of the CpG sites could effectively modulate gene expression (51). Therefore, to determine whether the observed folate-deficiency-induced site-specific alterations in the promoter CpG island region of ER in HCT116 and Caco-2 cells had any functional ramification, the effect of folate-deficiency on the ER protein expression was determined. In both HCT116 and Caco-2 cells, folate deficiency did not significantly alter the ER protein expression (Figure 2E).
Effect of folate deficiency on DNMT protein expression and total cellular CpG DNA DNMT activity
Dnmt activity was 28% lower in the folate-deficient NIH/3T3 cells than the folate-sufficient cells (Table II; P = 0.013). In contrast, no significant difference in Dnmt activity was observed between the folate-deficient and sufficient CHO-K1 cells (Table II). Consistent with Dnmt activity observed in NIH/3T3 cells, Dnmt 1 and 3a protein expression was significantly lower in the folate-deficient NIH/3T3 cells than in the folate-sufficient cells (Figure 3A).
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Discussion |
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The effect of isolated folate deficiency on methionine cycle intermediates was cell-specific and did not appear to be dependent on the extent of folate depletion or the stage of transformation. In CHO-K1 and Caco-2 cells, folate deficiency significantly decreased intracellular SAM concentrations and the SAM:SAH ratios as expected, but did not significantly change the intracellular SAH concentrations. Paradoxically, in NIH/3T3 and HCT116 cells, folate deficiency significantly increased the intracellular SAM concentrations and the SAM: SAH ratios. Folate deficiency significantly decreased the intracellular SAH concentrations in NIH/3T3 cells but not in HCT116 cells. These effects of folate deficiency observed in NIH/3T3 and HCT116 cells are unexpected and counterintuitive. Previous animal studies have shown a predictable decreased SAM and increased SAH concentrations and reduced SAM:SAH ratios associated with folate deficiency, even at less severe degrees than that employed in the present study, in the brain (52), kidney (52), pancreas (53) and liver (8,26,27,29,30,54,55) in rats. However, folate deficiency failed to induce significant changes in the colonic SAM and SAH concentrations and the SAM:SAH ratios in rat colon (27,28,30,34), except in an extremely severe folate-deficient state associated with a significant growth retardation (34) and in old animals (33). In these instances, folate deficiency significantly increased the colonic SAH concentrations (33,34) and decreased the SAM:SAH ratios (34) but did not significantly change the colonic SAM concentrations (33,34). Therefore, it has been suggested that the tenacious resistance of the colonic epithelial cells to altered SAM and SAH in response to folate deficiency might be a primary reason for the inability of folate deficiency to induce DNA methylation changes in the colon (34,40). The cell-specific effect of folate deficiency on the methionine cycle intermediates may be related to cell-specific differences in the presence and the relative activity of the following in response to folate deficiency: (a) a compensatory upregulation of betaine:homocysteine methyltransferase utilizing choline and betaine for remethylation of homocysteine to methionine (54); (b) a coordinated regulation by SAM of transmethylation (e.g. inhibition of methylenetetrahydrofolate reductase) and transsulfuration (e.g. activiation of cystathionine ß-synthase) of homocysteine (10); (c) an upregulation of glycine-N-methyltransferase for the transfer of the methyl group of SAM to glycine forming sarcosine, which is inhibited by 5-methyltetrahydrofolate (56); and (d) other, as yet undetermined, biochemical responses of methionine cycle enzymes such as cystathionine ß-synthase, SAH hydrolase and methionine adenosyltransferase (57).
It appears that the effect of folate deficiency on genomic DNA methylation is specific for the stage of transformation. In both untransformed NIH/3T3 and CHO-K1 cells, folate deficiency induced a significant, albeit modest, 20% reduction in genomic DNA methylation, which in the case of NIH/3T3 cells, was confirmed by significant hypomethylation in a centromeric minor satellite repeat sequence. In the case of CHO-K1 cells, genomic DNA hypomethylation in response to folate deficiency occurred in the setting of reduced SAM concentrations and SAM:SAH ratios as expected. However, genomic DNA hypomethylation occurred in the absence of a significant elevation of SAH, a potent inhibitor of most SAM-dependent methyltransferases (10). This is an unexpected finding because SAH levels appear to be a more accurate predictor of genomic DNA methylation than SAM concentrations or SAM:SAH ratios (58,59). Furthermore, genomic DNA hypomethylation in the folate-deficient CHO-K1 cells was not associated with a significant reduction in total Dnmt activity, which is probably explained by the lack of SAH-mediated Dnmt inhibition. In contrast, folate deficiency induced significant genomic DNA hypomethylation in NIH/3T3 cells despite a significant increase in SAM concentrations and SAM:SAH ratios and decrease in SAH concentrations, but was associated with an expected reduced Dnmt1 and Dnmt3a protein expression and total cellular Dnmt activity. This observation suggests that folate deficiency-induced genomic DNA hypomethylation might have been through a SAM and SAH-independent pathway in NIH/3T3 cells.
In contrast to the effect of folate deficiency on DNA methylation in untransformed cells, folate deficiency failed to induce significant genomic and satellite-specific DNA hypomethylation in human colon adenocarcinoma HCT116 and Caco-2 cells. One possible explanation for this observation is that the colon epithelial cells are resistant to the hypomethylating effect of folate deficiency compared with other cell types. Another explanation is that the transformed colonic epithelial cells are resistant to the hypomethylating effect of folate deficiency compared with untransformed cells, which is suggested by a recent study demonstrating a 25% lower degree of genomic DNA methylation in immortalized normal human colonic epithelial HCEC cells cultured in folate-deficient medium (<1 ng/l folic acid) for 14 days compared with cells cultured in control medium (4 mg/l folic acid) (7). Colon cancer cells are probably hypomethylated already and thus, may not be amenable to further changes. In this respect, a recent study has shown that HCT116 cells lacking DNMT1 exhibited only a modest 20% decrease in the overall genomic DNA methylation despite the markedly decreased cellular DNMT activity (46). In this model, although juxtacentromeric satellites became significantly demethylated, centromeric satellite loci and the promoter CpG island of the p16 gene remained fully methylated (46). Only when both the DNMT1 and DNMT3b genes were disrupted, genomic DNA methylation was reduced by >95% and significant hypomethylation of satellite sequences and several promoter CpG islands, including that of the p16 gene, was observed (60). These observations suggest that it is extremely difficult to alter DNA methylation in certain transformed cell lines such as HCT116 cells. The fact that an almost complete abolishment of DNMT activity by disruption of both the DNMT1 and DNMT3b genes is required to produce significant DNA hypomethylation in HCT116 cells (60) suggest that folate deficiency alone, which was not associated with a significant change in DNMT activity in HCT116 cells, and a modest 28% decrease in DNMT activity in Caco-2 cells in the present study, is unlikely to be a sufficient predisposing condition to produce significant DNA hypomethylation in these cells. However, there is evidence suggesting that the effect of isolated folate deficiency on DNA methylation may be cell-specific in transformed cells because human nasopharyngeal carcinoma KB cells grown in a folate-deplete medium (<10 nM folic acid) was associated with the hypomethylation of the folate binding protein gene (61) and paradoxical hypermethylation in a CpG island of the H-cadherin gene (62), compared with cells grown in a folate-replete medium (2.0 µM folic acid).
Isolated folate deficiency did not produce significant changes in the promoter CpG island methylation of the p16 and MLH1 genes in HCT116 and Caco-2 cells. In contrast, certain sites in the promoter CpG island of the ER gene demonstrated modest, albeit statistically significant, changes in CpG methylation in response to folate deficiency, which were not associated with significant functional consequences, as evidenced by the lack of significant change in the ER protein expression. However, our data do not exclude the possibility that folate deficiency may induce sequence-specific alterations in the promoter CpG island DNA methylation in other genes that are silenced in association with the CpG-island methylation. A more comprehensive genome-wide analysis of CpG island methylation is warranted to elucidate the potential gene and site-specific effect of folate deficiency on CpG island methylation in the human colonic epithelial cells.
In summary, our data indicate that the effect of folate deficiency on the methionine cycle intermediates is cell-specific. The direction of changes of these intermediates in response to folate deficiency is not uniformly consistent with the known biochemical effect of folate on the methionine cycle pathway. Our data further indicate that the effect of folate deficiency on DNA methylation depends on the stage of transformation; a modest degree of DNA hypomethylation was observed in untransformed mammalian cells in response to folate deficiency, whereas transformed human colonic epithelial cells were resistant to folate deficiency. In untransformed cells, the effect of folate deficiency on DNA methylation appears to be both SAM and SAH-dependent and independent. The effect of folate deficiency on DNMT activity also appears to be cell-specific, and the contribution of SAH inhibition on DNMT is not readily apparent in our in vitro system. The fact that a profound degree of folate deficiency produced only a very modest degree of DNA hypomethylation in mammalian cells suggest that a more moderate, physiologically and clinically relevant degree of folate deficiency is unlikely to induce a significant degree of DNA hypomethylation.
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Acknowledgments |
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References |
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