DNA methylation analysis

Pharmacology & Therapeutics 84 (1999) 389–400

Associate editor: P.K. Chiang

DNA methylation analysis: a review of current methodologies Edward J. Oakeley* Hatherly Laboratories, University of Exeter, Prince of Wales Road, Exeter EX4 4PS, UK

Abstract The relationship between levels of DNA methylation and gene activity has been known for some time. Many of the early procedures developed gave only somewhat limited information about methylation patterns, for example, the total level of 5-methyl cytosine in the genome or the frequency of methylation of cytosines within certain restriction sites. However, in the last few years, there has been an explosion of interest in DNA methylation, and with it, many new and powerful techniques have been developed to facilitate its study. In this paper, the key techniques currently available are reviewed and the advantages, disadvantages, and potential artifacts of each are discussed. © 1999 Elsevier Science Inc. All rights reserved. Keywords: Sodium bisulphite; Sodium bisulfite; HPLC; TLC; Immunohistochemistry Abbreviations: COBRA, COmbined Bisulphite Restriction Analysis; FITC, fluorescein isothiocyanate; m5c, 5-methyl cytosine; MS-SnuPE, methylationsensitive single nucleotide primer extension; PCR, polymerase chain reaction; SAM, S-adenosyl methionine.

Contents 1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2. Nonspecific DNA methylation analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. Reverse-phase high-performance liquid chromatography . . . . . . . . . . . . . . . . . . . . . 2.2. Thin-layer chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. SssI methyltransferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. The chloracetaldehyde reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. Immunology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Gene-specific methylation analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Restriction endonucleases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Ligation-mediated-polymerase chain reaction/hydrazine reaction/permanganate reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. The sodium bisulphite reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.1. DNA denaturation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.2. DNA depurination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3.3. Interpretation of data . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Combined bisulphite restriction analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.5. Methylation-sensitive single nucleotide primer extension. . . . . . . . . . . . . . . . . . . . . 3.5.1. Experimental pitfalls: primer design . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Conclusions. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction 5-Methyl cytosine (m5C) is the most frequently modified base found in eukaryotic genomes (Bestor & Coxon, 1993; * Corresponding author. Tel.: 144-1392-263787; fax: 144-1392263700. E-mail address: [email protected] (E.J. Oakeley)

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Chiang et al., 1996; Christman, 1982). m5C is created in situ by DNA methyltransferase enzymes, which can transfer a methyl group from the universal methyl donor S-adenosyl methionine (SAM) to position 5 of the cytosine ring (Schmitt et al., 1997). There are four methylation processes that can occur within the nucleus: the first is de novo methylation, where previously unmethylated cytosines, usually in

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the symmetrical sequence context CpG, become methylated (CpNpG and nonsymmetrical CpX methylation also occur in plants and fungi). The second is maintenance methylation, where the strand symmetry of hemimethylated DNA is maintained after replication by the methylation of the newly synthesised strand (Bestor & Verdine, 1994). The third is passive demethylation, where the maintenance methylation activity is suppressed, resulting in a 50% decrease in methylation during each round of DNA replication. The fourth is active demethylation, where methylation levels are decreased, in the absence of DNA replication, via an enzymatic process (Fremont et al., 1997; Weiss et al., 1996). The best-characterised active demethylation process is the removal of the modified base, using a DNA glycosylase enzyme, followed by a mismatch repair process (Fremont et al., 1997). Other demethylation activities have been reported, where either the entire CpG site is excised and repaired (Weiss et al., 1996) or the C2C bond between the cytosine and the methyl group is broken by a hydrolytic attack (Bhattacharya et al., 1999). Of these active processes, only the Bhattacharya hydrolytic attack will work on symmetrically methylated DNA. The Fremont and Weiss enzymes both result in phosphodiester bond cleavage, which would cause a double-strand DNA break, if it were permitted to occur on symmetrically methylated DNA. Thus, it seems likely that one class of enzymes exists to remove methylation in conjunction with cell division (the Fremont or Weiss enzymes), while the other allows the demethylation of DNA in nondividing cells (the Bhattacharya enzyme). It is clear that the cell expends considerable effort in its quest to selectively methylate and demethylate regions of the genome, and begs the question of what purpose this modification plays in the function of the nucleus. The function of m5C in prokaryotes is well understood, as it forms the basis of the restriction-modification system that protects bacteria from foreign DNA (Arber & Dussoix, 1962; Dussoix & Arber, 1962). Its role in eukaryotes is somewhat more complex, as it is involved in the fine control of gene expression. This can be either by the methylation of transcription factor binding sites (Comb & Goodman, 1990; Huntriss et al., 1997) or by inducing heterochromatin formation mediated by methyl cytosine-dependent histone deacetylation (Grunstein, 1997; Razin, 1998). In addition, methylation in eukaryotes can also act as a defence against molecular parasites (Jones et al., 1998) by inhibiting the activity of viruses and transposons. In certain fungal species, it has also been shown to be critical for the enzyme-catalysed conversion of C to T in repeated sequences via a process known as repeat-induced point mutation (Rountree & Selker, 1997). The purpose of this review is to provide a summary of the most commonly used techniques available today for the study of m5C in complex genomes. Each section will provide a general outline of the basic principles behind a technique and some of the potential pitfalls that should be considered when interpreting the data generated by these

procedures. The current methodologies may be broadly classed into gene-specific and nonspecific technologies, and it is under these major headings that they will be grouped. 2. Nonspecific DNA methylation analysis 2.1. Reverse-phase high-performance liquid chromatography Large-scale genome-wide changes in cytosine methylation levels are probably best monitored by reverse-phase HPLC. This procedure is one of the oldest available for methylation analysis (Kuo et al., 1980; Christman, 1982; Gomes & Chang, 1983). It relies on the quantitative hydrolysis of DNA using DNase I and nuclease P1 (Kuo et al., 1980) or snake venom phosphodiesterase (Gomes & Chang, 1983), followed by alkaline phosphatase treatment. The liberated deoxyribonucleosides are then separated by standard reverse-phase HPLC and the different bases identified by monitoring their UV absorbances at 254 and 280 nm. Further specificity may be achieved by combining HPLC separation with mass spectrometry so as to provide positive identification of the separated bases (Del Gaudio et al., 1997). The amount of material required for this analysis is typically quite small (,1 mg), making it suitable for the routine analysis of tissue samples. However, problems may occur if only small numbers of cells are available, as may be the case in paraffin-embedded histological samples, for example. 2.2. Thin-layer chromatography Not all laboratories have access to HPLC equipment, and a number of alternative procedures have been developed for studying genome-wide methylation levels. As has been discussed in Section 1, most vertebrate methylation occurs in the symmetrical context CpG. The restriction enzyme MspI has a target site of CCGG, and will cut this site whether or not the internal cytosine is methylated. The 59-phosphate of the internal cytosine may then be labelled using [g-32P]ATP and polynucleotide kinase (Bestor et al., 1984; Schmitt et al., 1997). The DNA is then hydrolysed to mononucleotides using nuclease P1, and separated on cellulose thin-layer chromatography plates (Kuchino et al., 1987; Schmitt et al., 1997), using isobutyric acid:ammonia:water (66:1:33 v/v/v) as the first dimension solvent and isopropanol:37% HCl:water (70:15:15 v/v/v) for the second dimension. The relative intensity of the spots may then be determined using a Molecular Dynamics PhosphorImager (or similar). The identity of each spot is confirmed by running control plates containing cytosine monophosphate or 5-methyl cytosine monophosphate markers. The relative intensity of the C to m5C spots will show the proportion of MspI sites that are methylated in the genome. In theory, only C and m5C should give spots in this experiment. However, we actually observed signals corresponding to A, G, and T as well. This is because random nicks in the DNA, caused by shearing forces, may also become labelled (Fig. 1). Digesting the DNA

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Another important source of error in the SssI procedure is the measurement of DNA concentration. Genomic DNA can be extremely difficult to dissolve to homogeneity, and as a result, the apparent OD260 might not accurately reflect the amount of DNA included in the reaction. Concentration errors of this kind can be at least as great as real differences in genome-wide methylation. The best way to minimise such errors is to digest the DNA with a restriction enzyme that does not contain a CpG sequence in its recognition site and then phenol:chloroform extract it prior to measuring the DNA concentration. Such digestion will reduce the viscosity of the DNA and ensure that a more homogeneous solution can be obtained. 2.4. The chloracetaldehyde reaction Fig. 1. Schematic representation of thin-layer chromatography of deoxynucleoside monophosphates. This diagram shows the typical spacing and pattern of spots obtained after kinase-labelling an MspI digest of genomic DNA and then degrading the DNA to deoxynucleoside monophosphates and separating the products by two-dimensional thin-layer chromatography. The positions of dCMP and m5dCMP are labelled as “C” and “mC,” respectively. The unlabelled circles represent the positions of the other deoxynucleoside monophosphates that are often visible due to labelling of nonspecific breakages in the DNA.

to free deoxynucleosides (in a similar way to the HPLC experiment) prior to labelling is not recommended, as it makes the C spot extremely hot, and this can obscure the m5C spot. It is a good idea to run a control plate in parallel where no MspI digestion has been performed, as this will allow a measure of the nonspecific C and m5C signals to be estimated. 2.3. SssI methyltransferase The enzyme SssI DNA methyltransferase is able to catalyse the de novo methylation of CpG sites (Shapiro et al., 1970b) using the universal methyl donor SAM. The principle of this assay is very simple: the enzyme is used to transfer a tritium-labelled methyl group from SAM to unmethylated cytosines in CpG sites of genomic DNA (Schmitt et al., 1997; Wu et al., 1993). The DNA is then immobilised on DEAE paper and the unincorporated SAM washed off. The amount of incorporation may then be quantitated using a scintillation counter. The more radioactive the sample, the less CpG methylation there was in the DNA sample. This procedure can be used to quantitate small global changes in methylation; however, the absolute number of counts recorded can vary a great deal from experiment to experiment. This is because both the SAM and the SssI enzyme are somewhat unstable. Typically, the error that is observed between repetitions performed at the same time is less than 5% of the mean. The difference between the mean values of an experiment performed on one day with those of an experiment performed on a different day can be as high as 50% (unpublished observation). If it is impractical to analyse all of the samples at the same time, then an internal control should be used to normalise the data between days.

The chloracetaldehyde reaction is a fluorescent assay of DNA methylation levels (Oakeley et al., 1999). The DNA is depurinated by treatment with sulphuric acid, and the purines then are removed by precipitation with silver or by column chromatography. The depurinated DNA may then be reacted with sodium bisulphite (see Section 3.3), which converts dC into dU, but has minimal effect on dm5C (Frommer et al., 1992). Incubation of the sample with chloroacetaldehyde results in the formation of the ethenocytosine derivative of m5C, which is intensely fluorescent (Oakeley et al., 1999). This fluorescence may be quantified using a fluorimeter and is proportional to the level of m5C in the genome. This technique provides a good alternative to the other approaches discussed in Sections 2.1–2.3, and is able to detect m5C in any sequence context. Because neither the chloroacetaldehyde nor the DNA themselves are fluorescent, the technique does not require extensive downstream purification. The removal of purines, however, is critical to the experiment because dA can also react with chloroacetaldehyde to form a fluorescent adduct. There are two major disadvantages with this procedure: firstly, it is somewhat time-consuming, as it requires one 4-hr and one overnight incubation; secondly, the chloroacetaldehyde reagent is quite toxic, and so, special care should be taken to minimise exposure to it (perform experiments in a fume hood). 2.5. Immunology The final genome-wide approach to the study of DNA methylation makes use of the highly specific reaction between monoclonal antibodies and m5C. The simplest approach is to immobilise denatured DNA samples onto DEAE membranes and then incubate them with a monoclonal antibody directed against m5C (Oakeley et al., 1997). The antibody may then be detected using a fluorescein isothiocyanate (FITC)-linked secondary followed by fluorescence scanning using a Molecular Dynamics PhosphorImager. The intensity of the staining is proportional to the degree of methylation in the DNA (Fig. 2). As an internal control, the membrane may be stained with ethidium bromide

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after the antibody signal has been quantified. The ethidium bromide fluorescence will be proportional to the amount of DNA loaded on the membrane, and thus, this value can be used to normalise the data from different samples. The major disadvantage with this strategy is that many common membrane types exhibit autofluorescence when scanned in the PhosphorImager (unpublished observation). Therefore, researchers are advised to scan blank membrane samples before risking valuable DNA samples. We have found that the membrane that gives the best signal:noise response is DEAE paper (Schleicher & Schuell NA 45, Dassel, Germany). This membrane binds both single- and double-stranded DNA in 50 mM NaCl. The DEAE should be pre-wetted by soaking for at least 1 hr in wash buffer (50 mM NaCl/10 mM Tris-HCl/1 mM EDTA, pH 7.5) 1 1% Triton X-100. The DNA was adsorbed onto the membrane using a slot-blot apparatus (BioRad, Hemel Hempstead, Hertfordshire, UK) and then washed for 15 min in wash buffer. The membrane was incubated with a 1:1000 dilution of a mouse monoclonal antibody directed against 5-methylcytidine (Podestà et al., 1993) for 2 hr. The membrane was then washed 3 times for 15 min in wash buffer 1 1% Triton X-100, and then incubated for an additional hour with an FITClinked goat anti-mouse secondary antibody (Sigma-Aldrich Co.

Fig. 2. Immunological identification of m5c. (a) A PhosphoImager scan of a DEAE membrane with 1, 2, 3, 4, or 5 mg of a synthetic oligonucleotide, containing m5C, immobilised on it. The m5C was detected by incubation with a mouse anti-m5C monoclonal antibody (Podesta et al., 1993) coupled to an FITC-anti-mouse secondary. (b) An image of an immature tobacco pollen grain undergoing mitosis. The pollen wall is visible because it possesses considerable autofluorescence. The chromosomes have been stained using the antibody described in (a) and detected with an FITC-linked secondary antibody using the procedure outlined in Oakeley et al. (1997). The scale bar is 10 mm in length.

Ltd., Gillingham, Dorset, UK). The sample was then scanned using a Molecular Dynamics PhosphoImager (Fig. 2a). Immunological approaches have also been used to visualise the chromosomal patterns of DNA methylation in individual cells by fluorescence microscopy (Buzek et al., 1998; Rougier et al., 1998; Oakeley et al., 1997). An example of such an application is shown in Fig. 2b, which shows an image of a tobacco pollen grain that is undergoing a mitotic division to generate the vegetative and generative cells found in mature pollen. The major limitation with antibody labelling of DNA is that it is only quantitative when the m5C is not basepaired—a situation that is very difficult to achieve in condensed chromosomes. The best way to achieve this objective is to treat the samples with sulphuric acid. The acid will depurinate the DNA (Maxam & Gilbert, 1980) and thus make it impossible, or at least much less likely, for m5C to be involved in base pairing.

3. Gene-specific methylation analysis 3.1. Restriction endonucleases As mentioned in Section 1, the major function of m5C in prokaryotes is as a defence against foreign DNA. The bacterial restriction/modification system makes use of DNA methyltransferases that introduce a methylation “signature” into newly synthesised DNA on C and A bases. Restriction endonucleases check any DNA in the cell for this “signature,” and any sequence that lacks the correct methylation pattern will be cleaved. In this way, most foreign DNA will be identified and degraded before it has a chance to be expressed (Bickle & Kruger, 1993). Restriction endonuclease isoschizomers that have different sensitivities to cytosine methylation provide a simple tool for the study of methylation changes within their recognition sites. Essentially, the methylation of a site will render it insensitive to cleavage by one enzyme, but not to the other. This will give a size difference on a Southern blot that easily can be observed and scored. This procedure is extremely popular with researchers who have large numbers of samples to analyse because of its relative simplicity, low cost, and ease of interpretation (Fig. 3). Where large quantities of DNA (.5 mg) are available, a standard Southern blot may be performed as follows: 5–10 mg of CsCl-purified (or equivalent quality) genomic DNA is digested with 10 U of the restriction enzyme HpaII overnight at 378C. A further 5–10 mg of DNA is digested in parallel, using the enzyme MspI. The digested DNA samples are then analysed by electrophoresis through a 0.8% agarose gel, transferred to a nylon membrane by Southern blotting, and then probed with the gene of interest. MspI will cut the sequence CCGG, whether or not the internal cytosine is methylated, whereas HpaII will only cut if the internal cytosine is unmethylated. Thus, if the hybridising bands are the same size in both HpaII and MspI digests, then this base is not

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Fig. 3. A schematic outline of the results expected from HpaII-MspI digestion of genomic DNA. (a) The patterns of bands expected from a single, methylated CCGG site within the target gene. HpaII will not cut this site, but MspI will. Thus, on a Southern blot (Blot), we will observe a big hybridising band in the HpaII track and a smaller band in the MspI track (the probe in this case would only extend as far as the CCGG site in one direction). In the case of PCR, a product will be observed only if the DNA is methylated. No band should ever appear in the MspI track, and this can be taken as a control for complete digestion. (b) The same outline, but in this case, the CCGG site is not methylated, so both enzymes cut.

methylated. If the band is bigger in an HpaII digest, then the site is methylated. In the case of partial methylation, you will see both a big and a small band, and their relative intensities will be proportional to the degree of methylation present at that site. Clearly, this procedure is only useful for probing a very limited number of potential methylation sites, so the amount of useful information that can be gained from it is somewhat limited. A correlation between the methylation status of a CCGG site and transcriptional activity can be useful, but if no correlation is observed, then it does not mean that DNA methylation is unimportant in the regulation of this gene, but rather, that CCGG methylation may be unimportant. One further complication that can occur in plant DNA analysis is that the outer cytosine can also be methylated by CXG methyltransferases. If this happens, then the band might be cut with HpaII (internal cytosine unmethylated), but not with MspI (outer cytosine methylated). Perhaps worse, if both cytosines are methylated, then neither enzyme will cut and at first glance, this would look the same as an unmethylated site (i.e., both bands the same size). This technique can be made more sensitive by the use of the polymerase chain reaction (PCR) (Fig. 3) to amplify products from small quantities of digested DNA (SingerSam et al., 1990). If a single CCGG lies between the PCR primers, then a product will only be amplified if the site is

not cleaved (Fig. 3). The only problem with this is that a partial digest will be indistinguishable from cytosine methylation. An alternative version of this procedure, known as COmbined Bisulphite Restriction Analysis (COBRA) (Xiong & Laird, 1997) and somewhat more informative, is given in Section 3.3.3. 3.2. Ligation-mediated-polymerase chain reaction/ hydrazine reaction/permanganate reaction DNA treated with hydrazine will undergo hydrazinolysis at C and T residues, but not at m5Cs. Partial hydrazinolysis can be used to map the locations of C and T residues by Maxam-Gilbert sequencing (Maxam & Gilbert, 1980). m5C may be identified by its absence from the sequencing ladder (Pfeifer et al., 1989). A related procedure makes use of permanganate oxidation at acidic pH (pH 4.1). In this reaction, m5C and T are sensitive to permanganate oxidation, and subsequent pyridinolysis will cleave the DNA at these sites, giving a positive display of the position of m5C residues (Fritzsche et al., 1987). In principle, these two procedures are equivalent and complementary, but in practice, the permanganate reaction should be avoided unless reaction artifacts from hydrazine are suspected. The reason for this is that the reaction between permanganate and m5C is highly context-dependent and highly variable under conditions that discriminate well between C and m5C (Fritzsche et al., 1987; Ru-

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bin & Schmid, 1980). On the other hand, the hydrazine reaction is much more uniform and provides clear and informative data. The visualisation of the reaction products is best achieved by the application of ligation-mediated-PCR, where first-strand synthesis is performed on the modified DNA using a gene-specific primer. A linker is then ligated onto this product at the hydrazine/permanganate cleavage site, and an exponential PCR is then used to replicate the DNA between the linker and the gene-specific primer (Grange et al., 1997; Mueller & Wold, 1989; Pfeifer et al., 1989). In this way, the reaction products may be simply resolved on a sequencing gel and identified. 3.3. The sodium bisulphite reaction The reaction between pyrimidines and sodium bisulphite has received considerable attention in recent years because it permits the rapid identification of m5C in any sequence context. However, despite its apparent technical superiority to other procedures available, it is prone to a number of important reaction artifacts. The reaction between pyrimidines and sodium bisulphite was first described in the early 1970s (Hayatsu et al., 1970; Shapiro et al., 1970a, 1970b). Early kinetic studies used uracil, uridine, thymine, and deoxycytidine as substrates, and it was demonstrated that they all underwent a rapid and reversible sulphonation at position 6 of the pyrimidine ring (Shapiro et al., 1970b). Subsequent analysis showed that m5C was also able to be sulphonated in this way (Hayatsu & Shiragami, 1979; Hayatsu et al., 1970; Wang et al., 1980). The amino group at position 4 of C and m5C is destabilised by this sulphonation so that these bases will then deaminate to either U or T, respectively, with a pH optimum of 5.8 (Hayatsu et al., 1970; Shapiro et al., 1970b). In the classic paper of Frommer et al. (1992), this reaction was used to distinguish between C and m5C in DNA. Frommer made use of the fact that the deamination reactions of sulphonated C and m5C proceed at very different rates, such that the deamination of C will be complete before substantial m5dC deamination has occurred (Frommer et al., 1992).

3.3.1. DNA denaturation Probably the most critical step in this reaction is the denaturation of the DNA. Sodium bisulphite can only react with pyrimidines that are not involved in base-pairing because only bases that can adopt a syn conformation can undergo substitutions at position 6 (Schweizer et al., 1971), and in double-stranded DNA, this is not possible because base-pairing locks the ring in the unreactive anti conformation (Shapiro et al., 1973; Fig. 4). Once the pyrimidines have reacted, base-pairing is no longer possible (Schweizer et al., 1971), so the melting temperature of the DNA will fall. Unfortunately, the high salt concentration in a standard reaction (3–5 M) makes double-stranded DNA a particularly unfavourable conformation, and as the strands come back together, the risk of incomplete reaction increases (Rein et al., 1997). Various technical modifications to this procedure have been attempted to reduce strand annealing, such as digesting the DNA prior to modification and then performing the reaction in a thermocycler with repeated heating steps to 958C (Rein et al., 1997). This is a clever approach, but suffers from the problem that DNA is very prone to acid-catalysed depurination, and repeated heating to 958C combined with a reaction pH of 5 can cause severe DNA degradation (see Section 3.3.2). A further solution that neatly avoids the problem of strand annealing is to embed the DNA in low melting point agarose blocks (Olek et al., 1996). The DNA is denatured, using NaOH, in solution and is then mixed with molten agarose. The agarose is cooled rapidly, and the DNA is frozen in a denatured form. All subsequent reaction and purification steps are performed on the agarose block, which greatly reduces DNA losses from complicated downstream purification procedures. The only problem that we have found with this procedure is that the agarose appears to be quite hydrophobic after following the recommended embedding procedure (Olek et al., 1996), and we found that we could only obtain reproducible reactions if a wetting agent such as 1% Nonidet P40 was added to the subsequent solutions. With this proviso aside, it is an excellent solution to a difficult problem.

Fig. 4. Steric impediment to the reaction of dC and m5dC in double-stranded DNA. (a) In double-stranded DNA, the bases are locked in the “anti-” conformation in which carbon 6 is sterically blocked so that it cannot react. (b) In single-stranded DNA, the bases can rotate about the C2N bond to deoxyribose. When they enter the “syn-” conformation, they can react with bisulphite.

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Chemical procedures have also been employed, such as the addition of 6 M urea to the bisulphite solution (Paulin et al., 1998). The authors found that the presence of urea destabilised base-pairing in the DNA and ensured that the reaction proceeded to completion (Paulin et al., 1998). This variant is too new to have been tested extensively by other groups, but in our laboratory, it did not appear to work as efficiently as the agarose procedure. We found it difficult to dissolve the urea in the presence of bisulphite, and we failed to obtain any PCR amplification of the DNA after the reaction was complete. As a result, we were unable to determine whether or not the bisulphite reaction had proceeded as anticipated. The value of performing a urea/bisulphite reaction in a thermocycler with 20 cycles of heating up to 958C (30 sec for each) seemed excessive, given the chaotropic nature of urea. Also, the rate of depurination is temperature-dependent, so incubation at elevated temperatures would be expected to cause a significant amount of additional DNA damage (Maxam & Gilbert, 1980), together with accelerating the thermal decomposition of the urea (McHenry, 1997). 3.3.2. DNA depurination As discussed in the previous section, DNA degradation due to partial acid-catalysed depurination can result in severe damage to template sequences, which subsequently may be difficult to amplify (Raizis et al., 1995). Under the moderately acidic conditions (pH 5) employed in this reaction, depurination proceeds relatively slowly and is only a serious problem where overnight incubations are employed (Raizis et al., 1995). The standard reason offered for an overnight incubation is that this ensures that the reaction proceeds to completion; however, kinetic justifications for such prolonged incubation times are obscure. Indeed, one potential problem with overnight incubations comes from several observations that up to 5% of the m5C in the genome will be deaminated by such a treatment (Clark et al., 1994; Wang et al., 1980), giving an underrepresentation of m5C in PCR products. As several groups have claimed that 4- to 5-hr incubations with bisulphite are sufficient for complete conversion (Olek et al., 1996; Paulin et al., 1998; Raizis et al., 1995; Rein et al., 1997; Shapiro et al., 1970b), we performed bisulphite reactions essentially as described in Raizis et al. (1995) using either poly-dC or poly-m5dC oligonucleotides (Microsynth AG, Buchs, Switzerland). As controls, we also reacted poly-dU and poly-dT oligonucleotides. The reactions were stopped at regular intervals by the addition of 2.5 vol of 2 M NaOH (this is required to overcome the pH buffering effect of 5 M bisulphite), and the desulphonation reaction was allowed to proceed for 30 min at room temperature. The residual salts were removed by gel filtration through a Nap-10 (Sephadex G50) column (Amersham Pharmacia plc, Amersham, England). The extent of reaction was measured by monitoring the chromatic shift that occurs as cytosine deaminates to uracil (Fig. 5a) or methylcytosine to thymine (Fig. 5c). The UV absorbances of the samples at pH 7 were measured at 262 nm and 278 nm (C reaction) and

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at 266 nm and 282 nm (m5C reaction). The relative concentrations of C to U and m5C to T were estimated using the technique of Kerr et al. (1949), and the results are plotted in Fig. 5b and 5d. From these data, the pseudo first order rate constants for these reactions may be estimated as 2.1 3 1023 sec21 (a half-life of 5.5 min) for C and 5.57 3 1026 sec21 (a half-life of 34.6 hr) for m5C. Thus, if the DNA is completely denatured, then overnight incubation should not have any beneficial effect on the reaction. However, overnight incubations may be required if the DNA is not completely denatured because the acid-catalysed depurination of the DNA will proceed to a much greater extent in samples treated in this way. Depurinated DNA does not base-pair as efficiently as undamaged DNA, so this loss of base-pairing may permit access of the bisulphite to cytosines located in difficult to denature sequences. However, the extensive DNA damage associated with long incubations may make the subsequent PCR of the modified DNA more difficult to perform. 3.3.3. Interpretation of data The most common procedure for interpreting bisulphite data is DNA sequencing. This can be done either by cloning and sequencing or by direct sequencing of PCR products. Cloning of products provides useful information about the context of methylated bases. For example, are certain sites always methylated together or is methylation of these sites an either/or affair? There are two major problems with cloning and sequencing: the first is the time factor, as a large number of clones (ideally .20) should be sequenced in order to obtain a high degree of confidence in the data; the second is the risk of artifacts, as incomplete conversion may appear as nonsymmetrical CpX methylation. The alternative approach is direct sequencing, which, while technically more difficult than plasmid sequencing, is less likely to show up incomplete conversion artifacts. Care should be taken when interpreting band intensities from a direct-sequencing experiment, however. The presence of a band in the C-track indicates the presence of m5C in the genome; if a band also appears in the T-track, then this indicates partial methylation (or incomplete bisulphite treatment). However, the relative intensities of these two bands will not reflect the proportion of m5C to C in the genome, if enzymatic sequencing is used. The reason for this is that the number of extending molecules present in the reaction determines band intensities. The T-track is likely to contain a lot of bands, and so, the number of molecules that terminate early will be high and the number that extend through the full sequence consequently will be very low, and this will reduce the signal strength of T-bands as the distance from the primer increases. The reverse is true for the C-track. Because very few cytosines are expected to be methylated, the number of extending molecules that encounter each C will be very high, and this will make the band intensity of each C unrealistically strong. Thus, the ratio of C:T on the sequencing gel will not reflect the true genomic m5C:C ratio (Tabor & Richardson, 1987, 1989, 1995).

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Fig. 5. Molar absorbance standard curves. (a) Plot of the molar absorbance for dC (solid line) and dU (dotted line) between 220–320 nm. (b) Semi-log plot of dC decay with time in minutes. The concentrations have been normalised ([dC]nor) to adjust for DNA loss during recovery. This normalisation was achieved by estimating [dC] and [dU] present in each sample and then plotting [dC]/([dC] 1 [dU]). (c) Plot of the molar absorbance for m5dC (solid line) and dT (dotted line) between 220–320 nm. (d) Semi-log plot of m5dC decay with time in minutes. The concentrations have been normalised ([m5dC]nor) to adjust for DNA loss during recovery. This normalisation was achieved by estimating [m5dC] and [dT] present in each sample and then plotting [m5dC]/([m5dC] 1 [dT]).

The major disadvantage of the bisulphite technique is that the modified DNA usually must be cloned and sequenced before useful data can be obtained. This can be extremely labour-intensive, so other approaches for the analysis of bisulphite products have been developed in recent years. These will be described in the following section. 3.4. Combined bisulphite restriction analysis One particularly powerful technique is known as COBRA, where DNA samples are reacted with bisulphite in a standard way. The sequence of interest is then amplified by PCR and subjected to restriction digestion with an enzyme that contains cytosines only within CG sites in its recognition sequence (such as BstUI [CGCG] or TaqI [TCGA]). If the cytosines are methylated, then the enzyme will still cut the site; if they were not methylated, then the restriction site will be lost (Xiong & Laird, 1997). As a control for complete bisulphite conversion, a second enzyme may be used that lacks CG sites within its recognition sequence. Hsp92II (CATG) is a good example of such an enzyme. Any cleavage by Hsp92II would indicate either incomplete conversion or nonsymmetrical methylation (Xiong & Laird, 1997). The reverse experiment is also possible; we have found that

the enzyme Tru9I is useful, as it provides a positive display of complete conversion. The sequence CCAA is not a restriction site and should almost never be methylated (except perhaps in the rare case of nonsymmetrical methylation). Complete bisulphite conversion will convert this sequence into TTAA, which can be cleaved by Tru9I. Partial conversion will not give a restriction site in the PCR product (Fig. 6). Taken together, Hsp92II and Tru9I give us a positive display of either partial conversion or complete conversion. The standard approach for this technique is: 1. Sequence the unmodified gene and identify useful restriction sites; 2. Perform a bisulphite reaction on the DNA; 3. Amplify the target gene by strand-specific PCR; 4. Perform a restriction map on the PCR product to locate the sites that are still present. Some example scenarios and their consequences are presented in Fig. 6. The COBRA technique is still limited to restriction sites and as such, cannot be used to probe all possible m5C residues. An alternative approach is provided by the technique of methylation-sensitive single nucleotide primer extension (MS-SNuPE).

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Fig. 6. COBRA analysis of DNA. (a) An example fragment of genomic DNA for analysis. It contains two restriction sites, CGCG (BSTUI) and TCGA (TAQI). In the centre is the sequence CCAA, which is not a restriction site, but can be converted into one (Tru9I) by the bisulphite reaction. The DNA can be subdivided into four parts labelled a, b, c, and d. In this example, we consider four potential outcomes. ①, no DNA methylation, complete reaction with bisulphite. The BstUI and TaqI sites are lost and a Tru9I site is created. ②, the BstUI site is fully methylated and the TaqI site is unmethylated. The reaction with bisulphite went to completion (Tru9I site created). ➂, both restriction sites fully methylated and the bisulphite went to completion. ➃, partial conversion has occurred. The absence of a Tru9I site indicates only partial bisulphite conversion, so the experiment should be repeated. Note that partial methylation of a BstUI site is indistinguishable from complete methylation. (b) The expected banding patterns from each of thesee outcomes in a triple digest. To the right of the gel is a key identifying the bands for clarity.

3.5. Methylation-sensitive single nucleotide primer extension MS-SNuPE (Gonzalgo & Jones, 1997) allows the user to probe the methylation status of a given cytosine in any sequence context, without having to clone and sequence PCR products. It is also based on the reaction of DNA with sodium bisulphite (Section 3.3). The basis of this technique is quite straightforward. After an initial bisulphite modification, the target sequence is amplified using strand-specific PCR primers (Section 3.3). The PCR product is then purified by gel electrophoresis and mixed with a primer that will hybridise to the modified PCR sequence immediately 59 to the cytosine to be probed. It is assumed that two possible outcomes may occur: either the site was methylated and the first base after the primer is unconverted (i.e., a C) or else it was unmethylated and was deaminated by the bisulphite (in which case, the PCR product will contain a T at this point). Two primer extension reactions are performed, one where radioactive [32P]dCTP is the only nucleotide in the reaction and one where [32P]dTTP is the only nucleotide. In the classical paper of Gonzalgo and Jones (1997), Taq polymerase is then used to extend the primer in each case. The products of the two labelling reactions may then be analysed by electrophoresis through a denaturing (7 M urea) polyacrylamide (15%) gel and quantified using a phosphorimager. If the base was methylated, then a band will be observed in the C reaction, but not in the T, and vice versa (Fig. 7).

3.5.1. Experimental pitfalls: primer design There are several potential difficulties with this protocol. These all focus on the design of the primer for use in primer extension. It is important to be clear as to which genomic strand your PCR primers will amplify after the bisulphite reaction. A genomic sequence may be represented as follows: 59-AAAACGTATA-39 [Strand A] 39-TTTTGCATAT-59 [Strand B] If this sequence is not methylated, then it will look like this after modification: 59-AAAAuGTATA-39 [Strand A] 39-TTTTGuATAT-59 [Strand B] If the PCR primers amplify Strand A, then the product will be: 59-AAAATGTATA-39 [Strand A9] 39-TTTTACATAT-59 [Strand A99] Whereas, if they amplify Strand B, the product will be: 59-AAAACATATA-39 [Strand B9] 39-TTTTGTATAT-59 [Strand B99] Therefore, it is only possible to probe the methylation status of the cytosine on the strand that was amplified. Thus, if we amplify Strand A, the MS-SNuPE primer used for the primer extension should be complementary to Strand A99 in

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extent of methylation of the target cytosine. Whilst it is best to avoid internal cytosines completely, it can be assumed that a C in a nonsymmetrical site usually will be converted into a U(T) and, therefore, the primer can contain a T at this position rather than a C. If a CG/CNG site must be located within the primer, then a degeneracy should be introduced at this point so that the primer contains both a C and a T. To avoid this complexity, the optimum target sequence in the PCR product should contain only C, T, and A bases and the primer should contain only G, A, and Ts. To clarify this point, this is the target sequence in a fully unmethylated region of DNA where we are interested in the CG site in bold: 59-AGACTTCGAACGTTTCTAAGTT-39 [Strand A] 39-TCTGAAGCTTGCAAAGATTCAA-59 [Strand B] If we have amplified Strand A, then we will obtain the following PCR product: 59-AGAtTTtGAAtGTTTtTAAGTT-39 [Strand A9] 39-TCTaAAaCTTaCAAAaATTCAA-59 [Strand A99]

Fig. 7. MS-SNuPE analysis of DNA. Flow chart showing the logical steps required to perform an MS-SNuPE analysis. Extreme care must be taken to ensure that the correct strand is amplified during the PCR. Amplification of the wrong strand will give a clear result, but it will always indicate methylation, even where none exists. As a control, it is a good idea to use a primer that is pointing in the same direction as the experimental primer, but which is immediately upstream of a nonsymmetric cytosine. This should always incorporate a T in the MS-SNuPE step. If instead it incorporates a C, then it is likely that the wrong strand has been amplified, or else that the bisulphite reaction did not run to completion.

the PCR product (i.e., 59-AAAA-39). Likewise, if we amplified Strand B in the PCR, then the MS-SNuPE primer must be complementary to Strand B9 in the PCR product (59TATA-39). In this way, the first nucleotide after the 39 end of the primer will be the cytosine that can change. If confusion arises and the wrong MS-SNuPE primer is used, then the experiment will appear to work, but the results will be invalid. In the example given, the use of the correct primer would incorporate a radioactive “T” into the primer, indicating no methylation, whereas the wrong primer will incorporate a radioactive “C” irrespective of the methylation status of the site. When designing the MS-SNuPE primers, it is also important to pay attention to the sequence around your target site. It is very important that the MS-SNuPE primer should not contain any Cs in CG (or CNG) sites prior to modification. If it does, then the annealing temperature of the primer will be dependent on the degree of conversion observed at each of these internal cytosines and thus, the incorporation of radioactivity will no longer be directly proportional to the

The MS-SNuPE primer contains two internal sites where sequence conversion can occur. The first is nonsymmetrical (italic) and can be assumed to always convert, but the second lies in a CG site (underlined) and so, might be methylated in certain situations. Thus, the following MS-SNuPE primer must be used: 59-AGAtTT(c/t)GAA-39 Even so, the data obtained from such a primer will only be semi-quantitative because of the uncertainty posed by this internal CG site.

4. Conclusions The importance of DNA methylation is clearly reflected in the vast amount of literature generated each year in this field. Many new techniques have been developed over the years to avoid certain limitations due to artifacts or the information content of the data. The fact that so many techniques exist presents a unique opportunity to researchers in this field, as it allows us to test hypotheses in several independent ways. This is particularly important when unexpected observations are made, as the data will always sit on a much firmer basis if it can be shown that the claimed methylation is present in more than one method of analysis. This is particularly important for the bisulphite reaction, where incomplete denaturation or partial reaction can lead to data that are very easily overinterpreted.

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