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Human Molecular Genetics Advance Access originally published online on September 30, 2004
Human Molecular Genetics 2004 13(23):2979-2989; doi:10.1093/hmg/ddh317
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Human Molecular Genetics, Vol. 13, No. 23 © Oxford University Press 2004; all rights reserved

Genome-wide demethylation destabilizes CTG·CAG trinucleotide repeats in mammalian cells

Vera Gorbunova1,{dagger},{ddagger}, Andrei Seluanov1,{dagger},{ddagger}, David Mittelman1,2 and John H. Wilson1,2,*

1Verna and Marrs McLean Department of Biochemistry and Molecular Biology and 2Graduate Program in Structural and Computational Biology and Molecular Biophysics, Baylor College of Medicine, One Baylor Plaza, Houston, TX 77030, USA

Received August 13, 2004; Revised September 17, 2004; Accepted September 24, 2004


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Many neurological diseases, including myotonic dystrophy, Huntington's disease and several spinocerebellar ataxias, result from intergenerational increases in the length of a CTG·CAG repeat tract. Although the basis for intergenerational repeat expansion is unclear, repeat tracts are especially unstable during germline development and production of gametes. Mammalian development is characterized by waves of genome-wide demethylation and remethylation. To test whether changes in methylation status might contribute to trinucleotide repeat instability, we examined the effects of DNA methyltransferase inhibitors on trinucleotide repeat stability in mammalian cells. Using a selectable genetic system for detection of repeat contractions in CHO cells, we showed that the rate of contractions increased >1000-fold upon treatment with the DNA methyltransferase inhibitor 5-aza-deoxycytidine (5-aza-CdR). The link between DNA demethylation and repeat instability was strengthened by similar results obtained with hydralazine treatment, which inhibits expression of DNA methyltransferase. In human cells from myotonic dystrophy patients, treatment with 5-aza-CdR strongly destabilized repeat tracts in the DMPK gene, with a clear bias toward expansion. The bias toward expansion events and changes in repeat length that occur in jumps, rather than by accumulation of small changes, are reminiscent of the intergenerational repeat instability observed in human patients. The dramatic destabilizing effect of DNA methyltransferase inhibitors supports the hypothesis that changes in methylation patterns during epigenetic reprograming may trigger the intergenerational repeat expansions that lead to disease.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
A growing list of inherited human diseases, mostly neurological or neuromuscular in nature, arise as a consequence of expansion of a microsatellite repeat in a susceptible gene (1). Many such diseases are due to expansion of CTG·CAG trinucleotide repeat tracts beyond a threshold of about 25–30 repeats to a length that has pathogenic consequences (1). Inheritance of these diseases is typically characterized by a progressive worsening of the disease phenotype in subsequent generations, as the repeat tract continues to expand. The basis for this intergenerational repeat instability has been investigated by examining the stability of repeats in experimental systems that include Escherichia coli, yeast, mammalian cells and mice. These studies have identified cis-elements, such as tract length, sequence purity and orientation relative to replication origins (2–5), and trans-acting factors, such as DNA repair and replication proteins (6–12), as agents that can critically affect the stability of repeats. Nevertheless, the mechanism by which such agents collaborate to bring about intergenerational repeat instability remains unclear.

Examination of human patients with expansion diseases has pointed to two periods in development—early embryogenesis and germline development—at which repeats seem to be particularly unstable. Occasional fetal mosaicism and the existence of monozygotic twins with different repeat alleles have been interpreted as evidence for instability early in embryogenesis (13–15). More commonly, patients are observed to have limited variation in allele size in somatic tissue, typically blood cells, but extensive variation in germ cells, suggesting that repeat instability is significantly enhanced at some point after segregation of the germline (16). Analysis of male germline cells from Huntington patients showed that considerable variation already existed in diploid precursors of germ cells, indicating that instability must occur before meiosis (17,18). These two periods of repeat instability overlap the two major cycles of epigenetic reprograming that occur during mammalian development (19). Immediately after fertilization and again in the differentiating germline, genomic patterns of CpG methylation are largely erased genome-wide and then re-established in a tissue-appropriate manner.

The potential link between epigenetic reprograming and repeat instability is strengthened by experimental observations that suggest a relationship between CpG methylation and repeat instability (20). Several di-, tri- and penta-nucleotide repeats, cloned on plasmids and tested in bacteria, were stabilized by CpG methylation (21). Surprisingly, CpG methylation of plasmid sequences adjacent to long CTG·CAG repeat tracts, which are devoid of CpG methylation sites, also enhanced repeat stability (21). Additional observations in humans show a correlation between CpG methylation and repeat stability. In fragile X patients, who carry an expanded CGG·CCG repeat, large methylated repeats are more stable than similar-sized unmethylated alleles (22–25). Early studies of Huntington's and myotonic dystrophy loci revealed no clear correlation between methylation at nearby CpG sites in the patient and age of onset or severity of the disease (26,27). However, in some severely affected myotonic dystrophy patients, who carry a highly expanded CTG·CAG repeat, hypermethylation of a CpG island located within 2 kb of the repeat tract is correlated with stabilization of the repeats in somatic tissues (28). Collectively, these observations are consistent with the hypothesis that human trinucleotide repeats could be destabilized during the waves of demethylation that occur during human development.

To test a key element of this hypothesis, we examined the effects of genome-wide demethylation on the stability of chromosomal CTG·CAG trinucleotide repeats in mammalian cells. To decrease DNA methylation genome-wide, we treated cells with 5-aza-deoxycytidine (5-aza-CdR) and with hydralazine, which inhibit DNA methyltransferase activity by different mechanisms (29,30). As detected by a selectable assay for repeat contractions at the APRT locus in CHO cells, these treatments dramatically destabilize long CTG·CAG repeat tracts. These results were confirmed by showing that 5-aza-CdR treatment causes a comparable destabilization of CTG·CAG repeats in the DMPK gene in cells from patients with myotonic dystrophy. As determined by GeneScan analysis, the instability induced by demethylation in human cells is biased toward repeat expansions. These results support the hypothesis that epigenetic reprograming during development may induce repeat instability.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
To analyze the effects of genome-wide DNA demethylation on the stability of chromosomal CTG·CAG trinucleotide repeats in mammalian cells, we utilized the selectable genetic system we developed at the APRT locus in CHO cells (31) (Fig. 1). In this system, a CTG·CAG repeat tract from the myotonic dystrophy gene is cloned into the second intron of the hamster APRT gene, in the orientation that gives CAG sequences in the APRT transcript. When the repeat tract is longer than 33 triplets, it is incorporated into the APRT mRNA as a new exon, rendering the encoded protein non-functional. Shorter repeat tracts, which are not efficiently incorporated into mRNA, do not affect APRT function. Therefore, contractions of long repeats can be monitored by selecting for APRT+ colonies from among the APRT– parental population. For these studies, we used the APRT– CHO cell lines, APRT(CAG)61 and APRT(CAG)95, which contain 61 and 95 repeats, respectively. In these cell lines, contraction to 33 or fewer repeats gives an APRT+ phenotype, providing a sensitive measure of repeat stability (31).



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  		Figure 1. Selectable assay for CTG·CAG repeat instability. Repeat tracts were cloned into the second intron of the APRT gene. Tracts equal to or longer than 34 repeats inactivate the APRT gene. Contractions of long repeat tracts to a length of fewer than 34 repeats restore gene activity and can be monitored by the appearance of APRT+ colonies.

 
In order to induce genome-wide demethylation, we treated cells with 5-aza-CdR, which is a well-characterized inhibitor of DNA methylation. An analog of cytidine, 5-aza-CdR is incorporated into DNA and inhibits DNA methylation by trapping DNA methyltransferases (31). Treatment of APRT(CAG)61 and APRT(CAG)95 cells with 0.5 µM 5-aza-CdR causes substantial genomic demethylation, as measured globally for CCGG (HpaII/MspI) sites or locally at a specific CCGG site adjacent to the APRT gene (Fig. 2). Global demethylation of CCGG sites was estimated to be nearly complete from the digestion patterns generated by methylation-insensitive (MspI) and methylation-sensitive (HpaII) restriction enzymes. In the absence of 5-aza-CdR treatment, MspI digestion produces a much more intense smear of low molecular weight fragments than does HpaII digestion; whereas after treatment with 0.5 µM 5-aza-CdR, the intensities are almost equal (Fig. 2C). We estimated that the percentage of methylated HpaII sites was reduced from 30 to 0.5% in APRT(CAG)61 cells and from 30 to 1.5% in APRT(CAG)95 cells following 5-aza-CdR treatment (see Materials and Methods). At the CCGG site adjacent to the APRT gene, 5-aza-CdR treatment reduced methylation from 100 to about 30% in APRT(CAG)61 cells and to about 50% in the APRT(CAG)95 cells, as judged from the intensities of bands corresponding to methylated and unmethylated HpaII restriction sites (Fig. 2B). These two assays indicate that 5-aza-CdR treatment significantly reduces DNA methylation in our CHO cells.



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  		Figure 2. Demethylating effect of 5-aza-CdR in APRT(CAG)61 and APRT(CAG)95 CHO cells. (A) Schematic representation of the APRT gene, showing the location of the CTG·CAG repeat track, the position of the probe and the methylated HpaII site, H*. B, BamHI site; H/M, unmethylated HpaII/MspI site; H*/M, methylated HpaII/MspI site. (B) Southern blot analysis of APRT(CAG)61 and APRT(CAG)95 cell lines before and after treatment with 0.5 µM 5-aza-CdR. B, BamHI digestion; H, HpaII+BamHI digestion; M, MspI+BamHI digestion. Arrows at 1.8 and 1.2 kb show the positions of the bands that arise when the H* site is methylated or not methylated, respectively. (C) Global genome analysis of demethylating effect of 5-aza-CdR. Total genomic DNA from APRT(CAG)61 and APRT(CAG)95 cells was digested with HpaII and MspI enzymes and separated on an agarose gel. The intensity of the smear produced by HpaII digestion reflects the extent of methylation at CCGG sites. Comparisons of intensities were made at positions indicated by empty arrows. H, HpaII digestion; M, MspI digestion.

 
The effect of genome-wide DNA demethylation on CAG repeat stability was measured by allowing the APRT(CAG)61 and APRT(CAG)95 cell lines to proliferate in the presence of increasing concentrations of 5-aza-CdR. Cells were then plated in the absence of selection to measure cell survival and in the presence of selection to measure the frequency of APRT+ colonies. Genome demethylation was accompanied by a dramatic dose-dependent increase in APRT+ colonies among cells that survived the treatment (Fig. 3A). At 0.5 µM 5-aza-CdR, which corresponds to the extensive demethylation shown in Figure 2, the rates of APRT+ colony formation in the APRT(CAG)61 and APRT(CAG)95 cell lines increased by 140- and 1400-fold, respectively, above their background rates in the absence of treatment. The 10-fold difference between the two cell lines is accounted for by about a 3-fold higher rate in treated APRT(CAG)95 cells and about a 3-fold lower spontaneous rate in untreated APRT(CAG)95 cells. As we have speculated previously, the lower spontaneous rate in APRT(CAG)95 cells relative to APRT(CAG)61 cells may be due to the larger contraction required to reduce the (CAG)95 repeat to a length compatible with APRT gene expression (31).



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  		Figure 3. Effects of various drug treatments on the stability CTG·CAG repeat tracts and on cell survival. (A) Effects of the DNA methyltransferase inhibitor 5-aza-CdR. (B) Effects of the DNA methyltransferase inhibitor hydralazine. (C) Effects of the histone deacetylase inhibitor sodium butyrate (NaButyrate). APRT(CAG)61 cells (open bars) and APRT(CAG)95 cells (filled bars) were treated with increasing concentrations of the indicated drugs, as described in Materials and Methods, and incubated in the absence of selection to score cell survival and in the presence of APRT+ selection to measure the number of APRT+ clones. Approximately 2x107 cells were used in each treatment. All the treatments were repeated at least three times and the standard deviations are shown.

 
A control treatment with 1.0 µM deoxycytidine did not increase APRT+ colony formation, nor were surviving colonies observed in 5-aza-CdR-treated DEPO1 cells (32), which carry a deletion in the APRT gene (data not shown). Contractions of the CTG·CAG repeats in 5-aza-CdR-treated cells were confirmed by DNA sequence analysis of individual APRT+ colonies. Interestingly, the distributions of contraction events for spontaneous and 5-aza-CdR-induced APRT+ colonies were significantly different [P({chi}2)=0.0005], with more extensive contractions evident in the drug-treated cells (Fig. 4). These results indicate that 5-aza-CdR treatment strongly destabilizes CTG·CAG repeat tracts.



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  		Figure 4. Comparison of spontaneous and 5-aza-CdR-induced repeat contractions in CHO cells. Empty bars represent spontaneous contractions and filled bars represent 5-aza-CdR-induced events. Repeat length was analyzed in 48 spontaneous and 37 5-aza-CdR-induced APRT+ colonies by PCR and sequencing as described in Materials and Methods. The spontaneous contractions comprised 27 that have been described previously (31) and 21 from this study. Neither the previous analysis nor this one revealed any differences in the distribution of contractions observed in APRT(CAG)61 cells and APRT(CAG)95 cells; thus, the results from both cell lines have been pooled in this figure.

 
Although 5-aza-CdR is a specific inhibitor of DNA methyltransferases, it also has a mild mutagenic effect (33–35). To determine whether its mutagenic properties might account for the instability of repeat tracts, we measured the rates of APRT mutation in the presence and absence of 5-aza-CdR in wild-type CHO cells. Treatment with 5-aza-CdR increased the rate of formation of APRT– colonies to 3.8x10–6, about three times the spontaneous rate (1.2x10–6). This level of mutagenesis seems unlikely to account for the huge increase in repeat instability observed in the cell lines containing trinucleotide repeats. Furthermore, no mutations other than repeat contractions were found in the sequences of the repeat tract and its flanking regions in the 5-aza-CdR-induced APRT+ cells.

As an independent assessment of whether the demethylating effects of 5-aza-CdR are in fact responsible for the observed instability of CTG·CAG repeats in treated cells, we tested another demethylating agent that acts through a different mechanism than 5-aza-CdR. Hydralazine inhibits expression of DNA methyltransferase I (DNMT1) mRNA by inhibiting the ERK signaling pathway (36). Treatment with 300 µM hydralazine reduced global methylation of CCGG sequences from 30 to 6% in APRT(CAG)61 cells and from 30 to 7% in APRT(CAG)95 cells (data not shown). In comparison, treatment with 5-aza-CdR reduced methylation in both cell lines to ~1% (Fig. 2C) (see earlier), consistent with hydralazine being a milder inhibitor of methylation than 5-aza-CdR (36). Treatment of the APRT(CAG)61 and APRT(CAG)95 cell lines with 300 µM hydralazine increased the rate of APRT+ colony formation 41- and 54-fold, respectively (Fig. 3B). The ability of two distinct demethylation agents, acting by different mechanisms, to stimulate contraction of CTG·CAG repeats in rough proportion to the extent of DNA demethylation argues strongly that genomic demethylation is responsible for the observed destabilization of CTG·CAG repeat tracts.

Methylated DNA is often associated with more compact forms of chromatin by virtue of its ability to recruit histone modifying enzymes such as histone deacetylase (37). This functional link raises the possibility that the destabilizing effect of genomic demethylation might be mediated through a loosening of chromatin structure, secondary to the loss of CpG methylation. To explore this possibility, we treated APRT(CAG)61 and APRT(CAG)95 cells with the histone deacetylase inhibitor sodium butyrate, which causes chromatin to adopt the unfolded nucleosome structure typical of transcribed chromatin (38). As shown in Figure 3C, this treatment had no effect on the rate of formation of APRT+ colonies in the APRT(CAG)61 cell line and produced less than a 3-fold increase in rate in the APRT(CAG)95 cell line. The minimal destabilizing effect of sodium butyrate treatment suggests that demethylation itself, rather than a change in chromatin structure, is the cause of CTG·CAG repeat instability.

Because of the nature of the selectable system at the APRT locus, we are able to use it to examine the effects of demethylation only on contraction of CTG·CAG repeat tracts. In human patients, however, disease-associated repeat instability is invariably linked to repeat expansions (1). To determine whether 5-aza-CdR treatment can induce expansions in CHO cells, as well as contractions, we PCR amplified the repeats in populations of cells, separated the products and used GeneScan to analyze the spectrum of repeat lengths present among the genomes from treated and untreated APRT(CAG)61 and APRT(CAG)95 cells. In addition to the major peak at 61 or 95 repeats, several new peaks were observed in the 5-aza-CdR-treated samples for APRT(CAG)61 and APRT(CAG)95 cells, respectively (Fig. 5). As summarized in Table 1, all the new peaks were contractions. These results indicate that demethylation-induced destabilization of CTG·CAG repeats in CHO cells is significantly biased toward contraction.



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  		Figure 5. GeneScan analysis of repeat instability in CHO APRT(CAG)61 and APRT(CAG)95 cell lines. (A) Examples of GeneScan traces for APRT(CAG)61 cells in the absence of treatment and after treatment with 0.5 µM 5-aza-CdR. (B) Examples of GeneScan traces for APRT(CAG)95 cells in the absence of treatment and after treatment with 0.5 µM 5-aza-CdR. Major peaks of 61 or 95 repeats are indicated by empty arrows. Additional peaks corresponding to repeat contractions are indicated by filled arrows. The approximate lengths of the repeat tracts were determined by reference to internal size standards.

 

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  		Table 1. Summary of GeneScan analyses of repeat tracts in CHO and human cells
 
To determine whether demethylation destabilizes CTG·CAG repeats in human cells and whether there was a bias for contraction or expansion, we measured the effects of 5-aza-CdR treatment on the stability of repeat tracts in the native DMPK gene in human cells. Two fibroblast cell lines from myotonic dystrophy patients, which contain expansions of 80 and 150 repeats, were treated with 5-aza-CdR and analyzed by GeneScan (Fig. 6). Demethylation significantly destabilized the repeats in these cells, as evidenced by the appearance of multiple additional peaks in the 5-aza-CdR-treated samples (Fig. 6A and B). In contrast to the results in CHO cells, however, demethylation-induced instability of CTG·CAG repeats in human cells, like spontaneous instability, is biased toward expansions, with 60–70% of the new peaks being longer than the major peak (Fig. 6C, Table 1). A comparison of GeneScan results in these cells line indicates that CTG·CAG repeat tracts are more unstable in human cells than in CHO cells in the absence of treatment, and that repeats in human cells are 10–20-fold more sensitive to 5-aza-CdR treatment (Table 1). Differences in the stability of trinucleotide repeats in humans and rodents have been noted previously (39).



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  		Figure 6. GeneScan analysis of the repeat instability in human primary fibroblasts from myotonic dystrophy patients. (A) Examples of GeneScan traces from MD9 and MD8 cells in the absence of treatment (control) and after treatment with 0.5 µM 5-aza-CdR (CdR). Major peaks corresponding to the mutant alleles in the patient cell lines are indicated by empty arrows. Additional peaks corresponding to contractions and expansions are indicated by filled arrows. The approximate lengths of the repeat tracts were determined by reference to internal size standards. Each GeneScan sample corresponds to an independently treated plate of MD9 or MD8 cells. (B) Number of GeneScan traces containing novel peaks. The number of additional peaks per GeneScan trace is distributed evenly, showing that the results are not the consequence of a few ‘jackpot’ samples. Only peaks with a height >10% of the major peak were included, although many smaller peaks were observed. (C) Distribution of repeat track sizes in contractions and expansions. Each bar represents one or more peaks corresponding to contractions or expansions within 20-triplet windows on either side of the major peak, which corresponds to the original mutant allele (indicated by empty arrows).

 
In both human cells and CHO cells, GeneScan analyses of CTG·CAG repeat instability suggest that 5-aza-CdR treatment induces discrete jumps in the lengths of repeats (Figs 5 and 6). These scans are incompatible with the notion that 5-aza-CdR induces a high frequency of small changes that accumulate to form larger changes. As shown explicitly in Figure 5, 5-aza-CdR treatment has no effect on the spectrum of small length differences that make up the major peak. Thus, the CTG·CAG repeat instability induced by 5-aza-CdR treatment mirrors this characteristic aspect of the disease process in humans.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
In this study, we have used two different inhibitors of methylation, 5-aza-CdR and hydralazine, to decrease DNA methylation genome-wide. We show that these treatments are potent inducers of CTG·CAG repeat instability in both a selectable system in CHO cells and in primary human cells from myotonic dystrophy patients. In CHO cells, where we can readily quantify the effects of treatment, we show that genome-wide demethylation in cells that contain a tract of 95 repeats increases repeat instability >1000-fold. We are unaware of any chemical treatment or genetic defect in any organism or model system that destabilizes repeats so dramatically (3,40,41). GeneScan analysis of cells from myotonic dystrophy patients indicates that repeats in human cells are even more susceptible to the effects of demethylation. Furthermore, the demethylation-induced changes in repeat length in human cells are biased toward expansion, which is the hallmark of the human diseases and similar to the patterns of instability observed in human sperm (17,42). Moreover, demethylation-induced changes appear to occur in jumps rather than by the accumulation of small changes, as is reminiscent of the appearance of full-mutation alleles in affected families (1,43). The extreme sensitivity of repeats to demethylation and the resulting bias toward expansion support the hypothesis that epigenetic reprograming during development contributes to the intergenerational instability of repeats in human disease. Although our results forge a potential link between epigenetic reprograming and trinucleotide repeat instability, others have observed germline instability during post-meiotic sperm maturation (11,17) and aging of the adult (42,44), which may or may not be related to the changes in methylation during development.

Our results do not address the specific mechanism that connects the removal of methyl groups to the instability of repeats. Demethylation of DNA might destabilize repeat sequences indirectly through global changes in gene expression patterns (45–47). By altering the levels or species of enzymes involved in DNA metabolism, demethylation might change the way cells deal with repeat sequences, heightening their instability. Alternatively, demethylation might destabilize repeats directly through local effects on the repeats, themselves. These two categories of explanation are not mutually exclusive. Here, we discuss the possible local effects of demethylation.

Demethylation might directly alter repeat stability by removal of methyl groups from the repeat, itself, or from adjacent sequences. On plasmids in bacteria, several unmethylated microsatellite repeats were shown to be less stable than their methylated counterparts, and CGG·CCG repeats were shown to be more stable in their methylated form in extrachromosomal assays in mammalian cells (21). Although CTG·CAG repeats do not contain CpG sequences, they could be modified via non-CpG methylation (48–50). This possibility seems unlikely, however, because non-CpG methylation is thought to be rare.

The more likely explanation for a local effect is that demethylation of nearby CpG elements is responsible for induction of repeat instability. The possibility that methylation of flanking CpG sequences might confer stability on nearby CTG·CAG repeats has been demonstrated directly for plasmids in bacteria (21). It has also been noted previously that many unstable trinucleotide repeats lie next to or within CpG-rich DNA segments (1,20,51). For example, sequence analysis of 1.5 kb on either side of the repeat at the myotonic dystrophy locus revealed 202 CpG dinucleotides, clustered so densely that most of the sequence can be classified as a CpG island. Methylation of these sequences in some severely affected patients is correlated with a dramatic stabilization of the repeats in somatic tissues (28). Similarly, the CTG·CAG repeats cloned into the APRT gene in our CHO cells lines are surrounded by 3 kb of DNA that contains 93 CpG dinucleotides, one of which we have shown to be 100% methylated in untreated cells. Thus, the demethylation-induced repeat instability we have observed in CHO cells and in cells from myotonic dystrophy patients could be due to methylation changes in flanking CpG dinucleotides.

Structural studies indicate that the effect of methylation may extend well beyond the methylatable sites. For example, methylation of DNA can induce local distortions (52) and increase duplex melting temperature (53). These effects may alter the ability of nearby repeats to form non-B DNA structures, such as hairpins, cruciforms, triplexes and slipped mispairs, which are the presumptive mutagenic precursors to expanded and contracted repeats (20). These altered structures are thought to mediate instability by changing the fidelity of DNA polymerases or by affecting DNA recombination.

Several observations indicate that DNA methylation may inhibit homologous recombination. Demethylation by treatment with 5-aza-CdR has been shown to increase the frequency of sister-chromatid exchange, even well after the drug was removed (30), to stimulate recombination among an array of transgenes (54), and to increase the frequency of dimerization of double-minute chromosomes (55). Knockout of DNMTI in ES cells, which leads to hypomethylation, increases genomic instability via pathways that may involve both homologous and nonhomologous recombination (56). Similarly, mice carrying a hypomorphic allele of DNMTI show increased chromosomal instability especially at the centromeric or pericentric regions, which are rich in repetitive elements (57). The genomes of mammalian germ cells, which undergo obligatory meiotic recombination during gametogenesis, are hypomethylated (19). In the mouse and human major histocompatibility complex, there is a high frequency of gene conversion in regions that have elevated CpG content; moreover, these recombination events are thought to occur in the male germline at a time when the CpG sequences are not methylated (58). The most direct evidence for inhibition of recombination by CpG methylation comes from experiments in Ascobolus immerses, which showed that meiotic recombination between methylated genes was reduced several 100-fold (59). These observations, and others, have led to the suggestion that CpG methylation, in addition to its role in transcription regulation, may serve to preserve genome integrity through its repressive effects on homologous recombination (56,60).

Additional experiments focused on the germline will be needed to confirm the hypothesis that epigenetic reprograming during development plays a part in the intergenerational repeat instability in human disease and to elucidate the molecular steps that link demethylation and repeat instability. These are important connections to be made because they suggest novel avenues for clinical management of repeat-expansion disease. For example, diets that promote DNA methylation may help to reduce intergenerational repeat expansion and slow down anticipation in affected families (61).


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Cell lines and growth conditions
Construction of the APRT(CAG)61 and APRT(CAG)95 CHO cell lines is described in detail elsewhere (31). CHO cells were grown in Dulbecco's modified Eagle's medium with high glucose supplemented with 10% fetal calf serum (FCS), non-essential amino acids, pyruvate, penicillin and streptomycin. Human primary fibroblasts from myotonic dystrophy patients MD9 (GM03991) and MD8 (GM03987) were obtained from the Coriell Cell Repository. Primary cells were grown in minimum essential medium with Earle's salts supplemented with 15% FCS, pyruvate, non-essential amino acids, penicillin and streptomycin.

Prior to drug treatments, CHO cells were grown continuously for 2 weeks in the presence of 8-aza-adenine (400 µM) to eliminate pre-existing APRT+ cells. For 5-aza-CdR, hydralazine and sodium butyrate treatment, the pre-selected CHO cells were plated at 7.5x105 cells per 100 mm dish, and then 24 h later the medium was replaced with medium containing one of the above inhibitors. Adding the inhibitors at low cell density is important, since they are most effective in dividing cells (30). After an additional 24 h, the medium was replaced again with fresh medium containing inhibitors. Inhibitors were removed 48 h later and cells were allowed to recover in fresh medium for 3 days and then plated in the absence of selection for cell survival and in the presence of ALASA selection for APRT+ colonies. The rate of formation of APRT+ colonies was calculated as described previously (31): the number of APRT+ colonies was adjusted for cell survival [APRT+ colonies/(surviving cells/plated cells)] and then divided by the number of population doublings that the cells underwent after treatment prior to applying APRT+ selection.

For 5-aza-CdR treatment, human primary cells were plated at a density of 5x105 cells per 100 mm dish, and then 24 h later medium was replaced with fresh medium containing 5-aza-CdR. On day 3, the cells were split to avoid reaching confluence. The 5-aza-CdR-containing medium was replaced daily for 5 days. Then 5-aza-CdR was removed and cells were allowed to recover in standard growth medium for 3 days and harvested.

APRT selection
Cells were plated at 5x105 cells per 100 mm plate in the presence of ALASA (25 µM alanosine, 50 µM azaserine, 100 µM adenine) to select for APRT+ cells, or with 8-aza-adenine (400 µM) for APRT– cells. Plates were incubated undisturbed for 3 weeks. The colonies were then picked for analysis or stained with 1% Coomassie blue for counting.

Sequencing of contracted repeat tracts from CHO cells
CTG·CAG repeat tracts were amplified from stained colonies or from isolated genomic DNA using Platinum Taq DNA polymerase (Invitrogen). The following primer pairs, which anneal within intron 2 of the APRT gene, were used to amplify and characterize contraction events: primers immediately flanking the repeat tract (5'-cctctagagtcgtccttgtagccgggaatg and 5'-gcctggccgaaagaaagaaatggtctgtgatcc); primers that anneal 100 bp away from the repeat tract (5'-gaaacaccctagggtcgctgaatgtccacc and 5'-tagcacatgtcagggctaccgaattcgcgg) and primers that anneal 200 bp away from the repeat tract (5'-taggagtagcacctaagatgaactagatgc and 5'-agttcagggtatatgtctggggtcacttcc). Following initial characterization of contracted repeats by PCR, the PCR products were cloned using a TOPO-2 PCR cloning kit (Invitrogen) and sequenced.

Analysis of local and global CpG methylation
CpG methylation status was examined using the methylation-sensitive HpaII restriction enzyme, agarose gel electrophoresis and Southern blotting. HpaII enzyme cleaves only unmethylated CCGG sites, whereas MspI cleaves CCGG sites regardless of methylation status.

To examine global CpG methylation, genomic DNA was extracted from the APRT(CAG)61 and APRT(CAG)95 cell lines and digested with either HpaII or MspI. The digested DNA was then separated by gel electrophoresis. We estimated the global methylation at HpaII/MspI sites in each cell line before and after treatment with demethylating agents by comparing the intensities of the smears at 0.85, 1.0 and 1.6 kb, which were produced by HpaII and MspI digestion (Fig. 2C). Percentage of methylated CCGG sites was calculated as [1–(intensityHpaII/intensityMspI)]x100%. The NIH Image software package was used to quantify the intensity of the smears from the DNA gel.

To examine the demethylating effect of 5-aza-CdR at the APRT locus, we identified a methylated HpaII/MspI site located 1.5 kb downstream of the repeat track. Genomic DNA from the APRT(CAG)61 and APRT(CAG)95 cell lines was digested with BamHI and either HpaII or MspI, and was analyzed by Southern blotting. We used a 700 bp probe that annealed between the methylation-sensitive HpaII/MspI site and the downstream BamHI site (Fig. 2A). Depending on methylation status of the CpG site, the probe hybridized to either a 1.8 (methylated CpG site), or a 1.2 kb (unmethylated CpG site) DNA fragment in the BamHI and HpaII digest. Percentage of methylated CCGG sites was calculated as [intensity1.8 kb band/(intensity1.2 kb band+intensity1.8 kb band)]x100%. The NIH Image software package was used to quantify band intensities.

GeneScan analysis
DNA was prepared from CHO or human cells following 5-aza-CdR treatment using a QIAmp blood kit (Qiagen) according to the manufacturer's instructions. The APRT locus in CHO DNA (25 ng) was amplified using Platinum Taq DNA polymerase (Invitrogen) with primers (5'-6FAM-taggagtagcacctaagatgaactagatgc and 5'-agttcagggtatatgtctggggtcacttcc), which anneal 200 bp away from the repeat tract. The repeat tracts in the DMPK gene in human DNA (25 ng) were amplified using a GC-rich PCR amplification kit (Roche) with primers (5'-6FAM-gggacagacaataaataccgaggaatgtcg and 5'-cgtgcgagtggactaacaacagctgtaggc), which anneal to sites ~500 bp away from the repeat tract. An aliquot of 1 µl of the PCR reaction mixture was added to 0.5 µl of GeneScan-2500 size standard and 10 µl of formamide. The samples were run on the ABI Prism 3100 Genetic Analyzer to separate the dye-tagged PCR products, and analyzed using GeneScan 3.1 software with an upgrade for long template analysis.


   ACKNOWLEDGEMENTS
 
We thank Graham Scott, Christie Kovar and other members of the Human Genome Sequencing Center at Baylor College of Medicine for help with GeneScan analysis and operating the ABI Prism 3100 instrument. We also thank Elisabeth Casinova from ABI Inc. for help and advice on GeneScan software. We thank Vincent Dion and other members of the Wilson laboratory for stimulating discussions. This work was supported by a Human Frontier of Science fellowship to V.G. and by an NIH grant, GM38219, and a Muscular Dystrophy Association grant to J.H.W.


   FOOTNOTES
 
* To whom correspondence should be addressed at: Verna and Marrs McLean, Department of Biochemistry and Molecular Biology, Baylor College of Medicine, One Baylor Plaza, Houston, TX 77030, USA. Tel: +1 7137985760; Fax: +1 7137969438; Email: jwilson{at}bcm.tmc.edu

{dagger} The authors wish it to be known that, in their opinion, the first two authors should be regarded as joint First Authors. Back

{ddagger} Present address: Department of Biology, University of Rochester, River Campus Box 270211, Rochester, NY 14627, USA. Back


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
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