Cell and stage of transformation-specific effects of folate deficiency on methionine cycle intermediates and DNA methylation in an in vitro model

Joanne M. Stempak 1, *, Kyoung-Jin Sohn 2, En-Pei Chiang 4, 5, Barry Shane 5 and Young-In Kim 1–3  

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


    Abstract
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Folate is an essential co-factor in the remethylation of homocysteine to methionine, thereby ensuring the supply of S-adenosylmethionine, the methyl group donor for most biological methylations, including that of DNA. Aberrant patterns and dysregulation of DNA methylation are consistent events in carcinogenesis and hence, DNA methylation is considered to be mechanistically related to the development of cancer. Folate deficiency appears to increase the risk of several malignancies, and aberrant DNA methylation has been considered to be a leading mechanism by which folate deficiency enhances carcinogenesis. Although diets deficient in methyl group donors (choline, folate, methionine and vitamin B12) have been consistently observed to induce DNA hypomethylation, the effect of an isolated folate deficiency on DNA methylation remains highly controversial and unresolved. Whether or not isolated folate deficiency can modulate DNA methylation is an important issue because it would establish a mechanistic link between folate deficiency and cancer. We examined the effects of isolated folate deficiency on methionine cycle intermediates, genomic and site-specific DNA methylation and DNA methyltransferase in an in vitro model of folate deficiency, using untransformed NIH/3T3 and CHO-K1 cells, and human HCT116 and Caco-2 colon cancer cells. Our data demonstrate that the effect of folate deficiency on the methionine cycle pathway and DNA methylation in these cells is highly complex and appears to depend on the cell type and stage of transformation, and may be gene and site-specific. The direction of changes of methionine cycle intermediates in response to folate deficiency is not uniformly consistent with the known biochemical effect of folate on the methionine cycle pathway. Moreover, the effect of folate deficiency on DNA methylation appears to be mediated by both methionine cycle intermediate-dependent and independent pathways.

Abbreviations: CpG, cytosine-guanine dinucleotide sequences; DNMT, DNA methyltransferase; SAH, S-adenosylhomocysteine; SAM, S-adenosylmethionine


    Introduction
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
An accumulating body of evidence over the past decade suggests that folate, a water-soluble B vitamin, may play a significant modulatory role in the development and prevention of several malignancies, with the most convincing evidence existing for colorectal cancer (1,2). These studies collectively suggest an inverse, albeit not uniformly consistent, association between folate status and the risk of these malignancies (1,2). Several proposed and studied mechanisms by which folate deficiency enhances and supplementation suppresses carcinogenesis in normal tissues are related to the sole biochemical function known for folate: mediating the transfer of 1-carbon moieties (13). Folate is an essential co-factor for the de novo biosynthesis of purines and thymidylate and hence plays an important role in DNA synthesis, stability and integrity, and repair, aberrations of which are mechanistically related to carcinogenesis (13). A growing body of evidence from in vitro, animal and human studies indicates that folate deficiency is associated with DNA strand breaks, impaired DNA repair and increased mutations, and that folate supplementation could correct some of these defects induced by folate deficiency (49). Folate, in the form of 5-methyltetrahydrofolate, is also involved in the remethylation of homocysteine to methionine, which is a precursor of S-adenosylmethionine (SAM), the primary methyl group donor for most biological methylations, including that of DNA (10). After transfer of the methyl group, SAM is converted to S-adenosylhomocysteine (SAH), a potent inhibitor of most SAM-dependent methyltransferases (10). It has been widely accepted that folate status alone can affect DNA methylation and it has been further proposed that a mechanism by which folate deficiency enhances carcinogenesis, particularly that of the colorectum, might be through an induction of genomic and site and gene-specific DNA hypomethylation (13).

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.


    Materials and methods
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Cell lines and culture
Two untransformed mammalian cell lines, NIH/3T3 mouse fibroblast cells and CHO-K1 Chinese hamster ovary cells, and two human colon adenocarcinoma cell lines, HCT116 and Caco-2 cells, were obtained from the American Type Culture Collection (Manassas, VA) and were cultured in standard RPMI 1640 medium (Invitrogen, Gaithersburg, MD) containing 0 (deficient) or 2.3 (control) µM folic acid. Growth medium was supplemented with 10% fetal bovine serum, 1% penicillin–streptomycin and 0.1% fungizone; dialyzed fetal bovine serum (Invitrogen) was added to the folate-deficient medium in order to eliminate folic acid in the serum. The cells were maintained at 37°C in 95% humidity and 5% CO2 and passaged every 4 days. Growth rates were determined by cell counts. NIH/3T3 and CHO-K1 cells were harvested after 12 days of growth while HCT116 and Caco-2 cells were harvested after 20 days of growth. No untransformed human colonic epithelial cells were used in the present study because of the lack of commercially available immortalized normal human colonic epithelial cells.

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 {lambda}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: 0–0.5 min, 100% A; 0.5–20 min, linear gradient to 75% A and 25% B; 20–30 min, 25% B; 30–45 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 Lowry–Bensadoun 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(I–C).d(I–C)] 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.


    Results
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 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
Effect of folate deficiency on cellular growth
Both untransformed and transformed cells cultured in folate-deficient medium for 12 and 20 days, respectively, grew despite the complete absence of folic acid in the medium. On comparison with corresponding cells cultured in folate-sufficient medium, however, the folate-deficient cells demonstrated a significant progressive retarded growth (data not shown). Cells cultured in folate-sufficient medium demonstrated widely different rates of growth with the fastest rate in CHO-K1 cells, intermediate rates in NIH/3T3 and HCT116 cells, and the slowest rate in Caco-2 cells (data not shown).

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 88–98%) 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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Table I. Intracellular folate, SAM and SAH concentrations and SAM:SAH ratios

 
Effect of folate deficiency on methionine cycle intermediates
Folate deficiency had cell-specific effects on the intracellular SAM and the SAH concentrations and the SAM:SAH ratio in both untransformed and transformed cell lines. In untransformed cell lines, the folate-deficient NIH/3T3 cells had significantly higher intracellular SAM (P = 0.043) and lower SAH (P = 0.006) concentrations, and an increased SAM:SAH ratio (P = 0.043) compared with the folate-sufficient cells (Table I). In contrast, the folate-deficient CHO-K1 cells had significantly lower intracellular SAM (P < 0.001) but no change in SAH concentrations, and a reduced SAM:SAH ratio (P < 0.001) compared with the corresponding folate-sufficient cells (Table I). In transformed cells, SAM concentrations and SAM:SAH ratios were significantly higher in the folate-deficient HCT116 cells than in the corresponding folate-sufficient cells (P = 0.002), whereas SAH concentrations were not significantly different between these cells (Table I). In contrast, SAM concentrations and SAM:SAH ratios were significantly lower in the folate-deficient Caco-2 cells than in the corresponding folate-sufficient cells (P < 0.001 and P = 0.006, respectively), whereas SAH concentrations were not significantly different between these cells (Table I).

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 SAM–SAH 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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Table II. The effect of folate deficiency on genomic DNA methylation and on CpG DNMT activity

 
Effect of folate deficiency on satellite DNA methylation
The effect of folate deficiency on specific satellite region DNA methylation was determined by Southern blot analyses after digesting the genomic DNA with restriction endonucleases that cleave only sequences lacking methylation at CpG residues within their recognition sites. In untransformed NIH/3T3 cells, folate deficiency induced hypomethylation in a mouse centromeric minor satellite repeat sequence as evidenced by the lower molecular weight HpaII-digested fragments present in the folate-deficient cells, compared with the folate-sufficient cells (Figure 1A). The HpaII-digestion patterns of the folate-deficient cells were similar to those of the folate-sufficient cells treated with a demethylating agent 5-aza-dC, although the intensity of the lower molecular weight fragments bands was higher in the 5-aza-dC treated samples than in the folate-deficient cells. The effect of folate deficiency on satellite region DNA methylation was not determined in CHO-K1 cells because of the lack of information related to satellite region sequences.



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Fig. 1. The effect of folate deficiency on specific satellite region DNA methylation was determined by Southern blot analysis after digesting genomic DNA with restriction endonucleases at CpG residues within their recognition sites. MspI (M) and HpaII (H) 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 methylation status. BstBI (B) cleaves the sequence 5'-TT^CGAA-3' if cytosine is unmethylated. Hybridization was performed with probes derived from a mouse centromeric minor satellite repeat sequence (pMR 150) for NIH/3T3 cells (A), from a human juxtacentromeric satellite 2 sequence from chromosome 1 for HCT116 (B) and Caco-2 (C) cells, and from a human centromeric alphoid satellite sequence from chromosome 18 for HCT116 (D) and Caco-2 (E) cells. Genomic DNA from 5-aza-2'-deoxycytidine (5-aza-dC; a demethylating agent) treated cells grown in control medium was used as a positive control. +F and –F denote DNA samples from cells cultured in control and folate-deficient medium, respectively.

 
In contrast, folate deficiency did not produce any appreciable degree of hypomethylation in specific satellite sequences in transformed HCT116 and Caco-2 colon cancer cells. Probes for classical satellite 2 and 3 sequences at the juxtacentromeric region of chromosomes 1, 9, 15 and 16 (Figure 2B and C) and for alphoid satellite sequences at the centromeres of chromosomes 3, 4, 7, 10, 12, 17, 18, 22 and X (Figure 2D and E) failed to reveal differences in the HpaII-digestion patterns between the folate-deficient and sufficient cells. Based on the comparison with the HpaII-digestion patterns of the sample treated with 5-aza-dC and the observation that high molecular weight fragments after HpaII digestion were present in both folate-deficient and sufficient cells, it appears that both centromeric and juxtacentromeric regions are densely methylated in CCGG sites.



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Fig. 2. The methylation status of individual CpG sites within the promoter CpG island (–695 bp to start) of the human Estrogen Receptor (ER) gene was determined by the sodium bisulfite-sequencing assay. Pictorial representation of the methylation status at each CpG site (total 13 sites) of each of 20 subclones for HCT116 (A) and Caco-2 (B) cells. Each row represents one subclone. Open circles represent unmethylated cytosines in CpG sites, whereas filled circles represent methylated cytosines in CpG sites. Proportion of methylated subclones at each CpG site in HCT116 (C) and Caco-2 (D) cells. * denotes statistically significant difference between folate deficiency and sufficiency at P < 0.05. The effect of the observed folate-deficiency-induced site-specific alterations in the promoter CpG island region of ER in HCT116 and Caco-2 cells on the ER protein expression was determined by western blot analysis (E). MCF-7 breast cancer cells express abundant ER and were used as a positive control. To confirm equal loading, the membranes were stripped and reprobed with a human anti-ß-actin antibody. Folate deficiency did not alter ER expression in either HCT116 (ratio of ER:ß-actin expression: +F = 1.58 versus –F = 1.37) or Caco-2 (ratio of ER:ß-actin expression: +F = 1.09 versus –F = 0.94) cells.

 
Similarly, the BstBI-digestion patterns of the folate-deficient and sufficient HCT116 and Caco-2 cells were similar for satellite 2 and 3 (Figure 2B and C) and alphoid satellite (Figure 2D and E) sequences, suggesting that folate deficiency failed to induce appreciable hypomethylation in TTCGAA sites present in these satellites. Comparison of the BstBI-digestion patterns between 5-aza-dC-treated and untreated samples reveals that the intrinsic degree of methylation in TTCGAA sites are different between the centromeric and juxtacentromeric satellite sites (Figure 2B–E). It appears that the degree of methylation in TTCGAA sites is higher in juxtacentromeric than in the centromeric satellite regions because the BstBI-digestion pattern was similar between 5-aza-dC-treated and untreated samples for centromeric satellite region (Figure 2D and E), whereas a more complete degree of BstBI-digestion was evident in 5-aza-dC-treated samples when compared with untreated samples for the juxtacentromeric satellite region (Figure 2B and C).

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 2A–D). In HCT116 cells, all the 13 CpG sites were 70–100% 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 60–100% 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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Fig. 3. DNMT1 and DNMT3a protein expression was determined in NIH/3T3 (A) and HCT116 and Caco-2 (B) cells cultured in folate-sufficient (+) or deficient (–) medium by western blot analysis. To confirm equal loading, the membranes were stripped and reprobed with a mouse or human anti-ß-actin antibody. Folate deficiency decreased DNMT1 and DNMT3a protein expression in NIH/3T3 (ratio of Dnmt1:ß-actin expression: +F = 0.92 versus –F = 0.80; ratio of Dnmt31:ß-actin expression: +F = 0.89 versus –F = 0.49), HCT116 (ratio of DNMT1:ß-actin expression: +F = 0.63 versus –F = 0.50; ratio of DNMT3a:ß-actin expression: +F = 0.73 versus –F = 0.60) and Caco-2 (ratio of DNMT1:ß-actin expression: +F = 0.95 versus –F = 0.87; ratio of DNMT3a:ß-actin expression: +F = 0.81 versus –F = 0.26) cells.

 
DNMT activity was not significantly different between the folate-deficient and sufficient HCT116 colon cancer cells (Table II). In contrast, DNMT activity was 28% lower in the folate-deficient Caco-2 colon cancer cells than in the folate-sufficient cells (Table II; P = 0.013). In both HCT116 and Caco-2 cells, DNMT1 and 3a protein expression was significantly lower in the folate-deficient cells than in the folate-sufficient cells (Figure 3B). DNMT3b protein expression was not determined because of the lack of commercially available DNMT3b antibody.


    Discussion
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 
We have developed an in vitro model of folate deficiency using both transformed and untransformed human and other mammalian cell lines to circumvent the limitations associated with animal models and to clearly elucidate the effect of an isolated folate deficiency on DNA methylation under stringent experimental conditions. We deliberately chose a severe degree of folate deficiency, albeit physiologically and nutritionally not applicable to in vivo situations, to maximize the chance of observing any effect of isolated folate deficiency on DNA methylation. Both untransformed mammalian cell lines and human colon cancer cell lines cultured in a medium completely lacking folic acid grew for 12 and 20 days, respectively, although these cells demonstrated a significant progressive retarded growth. A significant degree of functional folate depletion was observed in the folate-deficient cells compared with the corresponding folate-sufficient cells, thereby confirming the appropriateness of this in vitro model to study the effect of isolated folate deficiency on DNA methylation.

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.


    Acknowledgments
 
This project has been supported in part by a grant from the Canadian Institutes of Health Research (Grant # MOP-14126; to Y.-I. K.) and a grant from the National Science Council of Taiwan (Grant #NSC 93-2320-B005-006; to E.-P.C.). Young-In Kim is a recipient of a Scholarship from the Canadian Institutes of Health Research. Presented in part at the 2002 and 2003 American Association for Cancer Research Meeting, April 2002, San Francisco, CA and July 2003, Washington, DC, respectively, and published in abstract form in Proceedings of the American Association for Cancer Research 2002; 43: 521 (abst #2584) and 2003; 44: 890 (abst #3892).


    References
 Top
 Abstract
 Introduction
 Materials and methods
 Results
 Discussion
 References
 

  1. Choi,S.W. and Mason,J.B. (2002) Folate status: effects on pathways of colorectal carcinogenesis. J. Nutr., 132, 2413S–2418S.[Abstract/Free Full Text]
  2. Kim,Y.I. (1999) Folate and carcinogenesis: evidence, mechanisms, and implications. J. Nutr. Biochem., 10, 66–88.[CrossRef][ISI][Medline]
  3. Lamprecht,S.A. and Lipkin,M. (2003) Chemoprevention of colon cancer by calcium, vitamin D and folate: molecular mechanisms. Nature Rev. Cancer, 3, 601–614.[CrossRef][ISI][Medline]
  4. Blount,B.C., Mack,M.M., Wehr,C.M., MacGregor,J.T., Hiatt,R.A., Wang,G., Wickramasinghe,S.N., Everson,R.B. and Ames,B.N. (1997) Folate deficiency causes uracil misincorporation into human DNA and chromosome breakage: implications for cancer and neuronal damage. Proc. Natl Acad. Sci. USA, 94, 3290–3295.[Abstract/Free Full Text]
  5. Branda,R.F. and Blickensderfer,D.B. (1993) Folate deficiency increases genetic damage caused by alkylating agents and gamma-irradiation in Chinese hamster ovary cells. Cancer Res., 53, 5401–5408.[Abstract]
  6. Duthie,S.J. and Hawdon,A. (1998) DNA instability (strand breakage, uracil misincorporation, and defective repair) is increased by folic acid depletion in human lymphocytes in vitro. FASEB J., 12, 1491–1497.[Abstract/Free Full Text]
  7. Duthie,S.J., Narayanan,S., Blum,S., Pirie,L. and Brand,G.M. (2000) Folate deficiency in vitro induces uracil misincorporation and DNA hypomethylation and inhibits DNA excision repair in immortalized normal human colon epithelial cells. Nutr. Cancer, 37, 245–251.[CrossRef][ISI][Medline]
  8. Kim,Y.I., Pogribny,I.P., Basnakian,A.G., Miller,J.W., Selhub,J., James,S.J. and Mason,J.B. (1997) Folate deficiency in rats induces DNA strand breaks and hypomethylation within the p53 tumor suppressor gene. Am. J. Clin. Nutr., 65, 46–52.[Abstract]
  9. Kim,Y.I., Shirwadkar,S., Choi,S.W., Puchyr,M., Wang,Y. and Mason,J.B. (2000) Effects of dietary folate on DNA strand breaks within mutation-prone exons of the p53 gene in rat colon. Gastroenterology, 119, 151–161.[ISI][Medline]
  10. Selhub,J. and Miller,J.W. (1992) The pathogenesis of homocysteinemia: interruption of the coordinate regulation by S-adenosylmethionine of the remethylation and transsulfuration of homocysteine. Am. J. Clin. Nutr., 55, 131–138.[Abstract]
  11. Jones,P.A. and Laird,P.W. (1999) Cancer epigenetics comes of age. Nat. Genet., 21, 163–167.[CrossRef][ISI][Medline]
  12. Jones,P.A. and Baylin,S.B. (2002) The fundamental role of epigenetic events in cancer. Nature Rev. Genet., 3, 415–428.[ISI][Medline]
  13. Lengauer,C., Kinzler,K.W. and Vogelstein,B. (1997) DNA methylation and genetic instability in colorectal cancer cells. Proc. Natl Acad. Sci. USA, 94, 2545–2550.[Abstract/Free Full Text]
  14. Chen,R.Z., Pettersson,U., Beard,C., Jackson-Grusby,L. and Jaenisch,R. (1998) DNA hypomethylation leads to elevated mutation rates. Nature, 395, 89–93.[CrossRef][ISI][Medline]
  15. Esteller,M., Corn,P.G., Baylin,S.B. and Herman,J.G. (2001) A gene hypermethylation profile of human cancer. Cancer Res., 61, 3225–3229.[Abstract/Free Full Text]
  16. Toyota,M., Ahuja,N., Suzuki,H., Itoh,F., Ohe-Toyota,M., Imai,K., Baylin,S.B. and Issa,J.P. (1999) Aberrant methylation in gastric cancer associated with the CpG island methylator phenotype. Cancer Res., 59, 5438–5442.[Abstract/Free Full Text]
  17. Zapisek,W.F., Cronin,G.M., Lyn-Cook,B.D. and Poirier,L.A. (1992) The onset of oncogene hypomethylation in the livers of rats fed methyl-deficient, amino acid-defined diets. Carcinogenesis, 13, 1869–1872.[Abstract]
  18. Wainfan,E., Dizik,M., Stender,M. and Christman,J.K. (1989) Rapid appearance of hypomethylated DNA in livers of rats fed cancer-promoting, methyl-deficient diets. Cancer Res., 49, 4094–4097.[Abstract]
  19. Wainfan,E. and Poirier,L.A. (1992) Methyl groups in carcinogenesis: effects on DNA methylation and gene expression. Cancer Res., 52, 2071S–2077S.[Medline]
  20. Dizik,M., Christman,J.K. and Wainfan,E. (1991) Alterations in expression and methylation of specific genes in livers of rats fed a cancer promoting methyl-deficient diet. Carcinogenesis, 12, 1307–1312.[Abstract]
  21. Christman,J.K., Sheikhnejad,G., Dizik,M., Abileah,S. and Wainfan,E. (1993) Reversibility of changes in nucleic acid methylation and gene expression induced in rat liver by severe dietary methyl deficiency. Carcinogenesis, 14, 551–557.[Abstract]
  22. Pogribny,I.P., Basnakian,A.G., Miller,B.J., Lopatina,N.G., Poirier,L.A. and James,S.J. (1995) Breaks in genomic DNA and within the p53 gene are associated with hypomethylation in livers of folate/methyl-deficient rats. Cancer Res., 55, 1894–1901.[Abstract]
  23. Pogribny,I.P., Poirier,L.A. and James,S.J. (1995) Differential sensitivity to loss of cytosine methyl groups within the hepatic p53 gene of folate/methyl deficient rats. Carcinogenesis, 16, 2863–2867.[Abstract]
  24. Pogribny,I.P., Miller,B.J. and James,S.J. (1997) Alterations in hepatic p53 gene methylation patterns during tumor progression with folate/methyl deficiency in the rat. Cancer Lett., 115, 31–38.[CrossRef][ISI][Medline]
  25. Wainfan,E., Kilkenny,M. and Dizik,M. (1988) Comparison of methyltransferase activities of pair-fed rats given adequate or methyl-deficient diets. Carcinogenesis, 9, 861–863.[Abstract]
  26. Balaghi,M. and Wagner,C. (1993) DNA methylation in folate deficiency: use of CpG methylase. Biochem. Biophys. Res. Commun., 193, 1184–1190.[CrossRef][ISI][Medline]
  27. Kim,Y.I., Christman,J.K., Fleet,J.C., Cravo,M.L., Salomon,R.N., Smith,D., Ordovas,J., Selhub,J. and Mason,J.B. (1995) Moderate folate deficiency does not cause global hypomethylation of hepatic and colonic DNA or c-myc-specific hypomethylation of colonic DNA in rats. Am. J. Clin. Nutr., 61, 1083–1090.[Abstract]
  28. Kim,Y.I., Salomon,R.N., Graeme-Cook,F., Choi,S.W., Smith,D.E., Dallal,G.E. and Mason,J.B. (1996) Dietary folate protects against the development of macroscopic colonic neoplasia in a dose responsive manner in rats. Gut, 39, 732–740.[Abstract]
  29. Le Leu,R.K., Young,G.P. and McIntosh,G.H. (2000) Folate deficiency diminishes the occurrence of aberrant crypt foci in the rat colon but does not alter global DNA methylation status. J. Gastroenterol. Hepatol., 15, 1158–1164.[CrossRef][ISI][Medline]
  30. Le Leu,R.K., Young,G.P. and McIntosh,G.H. (2000) Folate deficiency reduces the development of colorectal cancer in rats. Carcinogenesis, 21, 2261–2265.[Abstract/Free Full Text]
  31. Davis,C.D. and Uthus,E.O. (2003) Dietary folate and selenium affect dimethylhydrazine-induced aberrant crypt formation, global DNA methylation and one-carbon metabolism in rats. J. Nutr., 133, 2907–2914.[Abstract/Free Full Text]
  32. Duthie,S.J., Narayanan,S., Brand,G.M. and Grant,G. (2000) DNA stability and genomic methylation status in colonocytes isolated from methyl-donor-deficient rats. Eur. J. Nutr., 39, 106–111.[CrossRef][ISI][Medline]
  33. Choi,S.W., Friso,S., Dolnikowski,G.G., Bagley,P.J., Edmondson,A.N., Smith,D.E. and Mason,J.B. (2003) Biochemical and molecular aberrations in the rat colon due to folate depletion are age-specific. J. Nutr., 133, 1206–1212.[Abstract/Free Full Text]
  34. Sohn,K.J., Stempak,J.M., Reid,S., Shirwadkar,S., Mason,J.B. and Kim,Y.I. (2003) The effect of dietary folate on genomic and p53-specific DNA methylation in rat colon. Carcinogenesis, 24, 81–90.[Abstract/Free Full Text]
  35. Kim,Y.I., Pogribny,I.P., Salomon,R.N., Choi,S.W., Smith,D.E., James,S.J. and Mason,J.B. (1996) Exon-specific DNA hypomethylation of the p53 gene of rat colon induced by dimethylhydrazine. Modulation by dietary folate. Am. J. Pathol., 149, 1129–1137.[Abstract]
  36. Song,J., Sohn,K.J., Medline,A., Ash,C., Gallinger,S. and Kim,Y.I. (2000) Chemopreventive effects of dietary folate on intestinal polyps in Apc+/–Msh2–/– mice. Cancer Res., 60, 3191–3199.[Abstract/Free Full Text]
  37. Jacob,R.A., Gretz,D.M., Taylor,P.C., James,S.J., Pogribny,I.P., Miller,B.J., Henning,S.M. and Swendseid,M.E. (1998) Moderate folate depletion increases plasma homocysteine and decreases lymphocyte DNA methylation in postmenopausal women. J. Nutr., 128, 1204–1212.[Abstract/Free Full Text]
  38. Rampersaud,G.C., Kauwell,G.P., Hutson,A.D., Cerda,J.J. and Bailey,L.B. (2000) Genomic DNA methylation decreases in response to moderate folate depletion in elderly women. Am. J. Clin. Nutr., 72, 998–1003.[Abstract/Free Full Text]
  39. Ingrosso,D., Cimmino,A., Perna,A.F., Masella,L., De Santo,N.G., De Bonis,M.L., Vacca,M., D'Esposito,M., D'Urso,M., Galletti,P. and Zappia,V. (2003) Folate treatment and unbalanced methylation and changes of allelic expression induced by hyperhomocysteinaemia in patients with uraemia. Lancet, 361, 1693–1699.[CrossRef][ISI][Medline]
  40. Kim,Y.I. (2004) Folate and DNA methylation: a mechanistic link between folate deficiency and colorectal cancer? Cancer Epidemiol. Biomarkers Prev., 13, 511–519.[Abstract/Free Full Text]
  41. Tamura,T. (1990) Microbiological assay of folates. In Piccairo,M.F., Stokstad,R. and Gregory,J.F. (eds) Folic Acid Metabolism in Health and Diseases. Wiley-Liss, New York, NY, pp. 121–137.
  42. Cravo,M.L., Mason,J.B., Selhub,J. and Rosenberg,I.H. (1991) Use of the deoxyuridine suppression test to evaluate localized folate deficiency in rat colonic epithelium. Am. J. Clin. Nutr., 53, 1450–1454.[Abstract]
  43. Fell,D., Benjamin,L.E. and Steele,R.D. (1985) Determination of adenosine and S-adenosyl derivatives of sulfur amino acids in rat liver by high-performance liquid chromatography. J. Chromatogr., 345, 150–156.[Medline]
  44. Bensadoun,A. and Weinstein,D. (1976) Assay of proteins in the presence of interfering materials. Anal. Biochem., 70, 241–250.[CrossRef][ISI][Medline]
  45. Laird,P.W., Zijderveld,A., Linders,K., Rudnicki,M.A., Jaenisch,R. and Berns,A. (1991) Simplified mammalian DNA isolation procedure. Nucleic Acids Res., 19, 4293.[ISI][Medline]
  46. Rhee,I., Jair,K.W., Yen,R.W., Lengauer,C., Herman,J.G., Kinzler,K.W., Vogelstein,B., Baylin,S.B. and Schuebel,K.E. (2000) CpG methylation is maintained in human cancer cells lacking DNMT1. Nature, 404, 1003–1007.[CrossRef][ISI][Medline]
  47. Christman,J.K. (2002) 5-Azacytidine and 5-aza-2'-deoxycytidine as inhibitors of DNA methylation: mechanistic studies and their implications for cancer therapy. Oncogene, 21, 5483–5495.[CrossRef][ISI][Medline]
  48. Clark,S.J. and Frommer,M. (1997) Bisulfite genomic sequencing of methylated cytosines. In Taylor,G.R. (ed.) Laboratory Methods for the Detection of Mutations and Polymorphisms in DNA. CRC Press, Boca Raton, pp. 151–162.
  49. Issa,J.P., Vertino,P.M., Wu,J., Sazawal,S., Celano,P., Nelkin,B.D., Hamilton,S.R. and Baylin,S.B. (1993) Increased cytosine DNA-methyltransferase activity during colon cancer progression. J. Natl Cancer Inst., 85, 1235–1240.[Abstract]
  50. Robertson,K.D., Hayward,S.D., Ling,P.D., Samid,D. and Ambinder,R.F. (1995) Transcriptional activation of the Epstein-Barr virus latency C promoter after 5-azacytidine treatment: evidence that demethylation at a single CpG site is crucial. Mol. Cell. Biol., 15, 6150–6159.[Abstract]
  51. Hsieh,C.L. (1994) Dependence of transcriptional repression on CpG methylation density. Mol. Cell. Biol., 14, 5487–5494.[Abstract]
  52. Ordonez,L.A. and Wurtman,R.J. (1974) Folic acid deficiency and methyl group metabolism in rat brain: effects of L-dopa. Arch. Biochem. Biophys., 160, 372–376.[CrossRef][ISI][Medline]
  53. Balaghi,M. and Wagner,C. (1992) Methyl group metabolism in the pancreas of folate-deficient rats. J. Nutr., 122, 1391–1396.[ISI][Medline]
  54. Kim,Y.I., Miller,J.W., da Costa,K.A., Nadeau,M., Smith,D., Selhub,J., Zeisel,S.H. and Mason,J.B. (1994) Severe folate deficiency causes secondary depletion of choline and phosphocholine in rat liver. J. Nutr., 124, 2197–2203.[ISI][Medline]
  55. Miller,J.W., Nadeau,M.R., Smith,J., Smith,D. and Selhub,J. (1994) Folate-deficiency-induced homocysteinaemia in rats: disruption of S-adenosylmethionine's co-ordinate regulation of homocysteine metabolism. Biochem. J., 298, 415–419.[ISI][Medline]
  56. Selhub,J. (1999) Homocysteine metabolism. Annu. Rev. Nutr., 19, 217–246.[CrossRef][ISI][Medline]
  57. Finkelstein,J.D. (1990) Methionine metabolism in mammals. J. Nutr. Biochem., 1, 228–237.[CrossRef][ISI][Medline]
  58. Yi,P., Melnyk,S., Pogribna,M., Pogribny,I.P., Hine,R.J. and James,S.J. (2000) Increase in plasma homocysteine associated with parallel increases in plasma S-adenosylhomocysteine and lymphocyte DNA hypomethylation. J. Biol. Chem., 275, 29318–29323.[Abstract/Free Full Text]
  59. Caudill,M.A., Wang,J.C., Melnyk,S., Pogribny,I.P., Jernigan,S., Collins,M.D., Santos-Guzman,J., Swendseid,M.E., Cogger,E.A. and James,S.J. (2001) Intracellular S-adenosylhomocysteine concentrations predict global DNA hypomethylation in tissues of methyl-deficient cystathionine beta-synthase heterozygous mice. J. Nutr., 131, 2811–2818.[Abstract/Free Full Text]
  60. Rhee,I., Bachman,K.E., Park,B.H., Jair,K.W., Yen,R.W., Schuebel,K.E., Cui,H., Feinberg,A.P., Lengauer,C., Kinzler,K.W., Baylin,S.B. and Vogelstein,B. (2002) DNMT1 and DNMT3b cooperate to silence genes in human cancer cells. Nature, 416, 552–556.[CrossRef][ISI][Medline]
  61. Hsueh,C.T. and Dolnick,B.J. (1993) Altered folate-binding protein mRNA stability in KB cells grown in folate-deficient medium. Biochem. Pharmacol., 45, 2537–2545.[CrossRef][ISI][Medline]
  62. Jhaveri,M.S., Wagner,C. and Trepel,J.B. (2001) Impact of extracellular folate levels on global gene expression. Mol. Pharmacol., 60, 1288–1295.[Abstract/Free Full Text]
Received October 29, 2004; revised January 21, 2005; accepted January 26, 2005.