Electrochemical DNA Methylation Detection for Enzymatically

Sep 11, 2011 - 2011, 83, 7595-7599. LETTER pubs.acs.org/ac. Electrochemical DNA Methylation Detection for Enzymatically. Digested CpG Oligonucleotides...
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Electrochemical DNA Methylation Detection for Enzymatically Digested CpG Oligonucleotides Dai Kato,† Keisuke Goto,†,‡ Shin-ichiro Fujii,§ Akiko Takatsu,§ Shigeru Hirono,|| and Osamu Niwa*,†,‡ †

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Biomedical Research Institute, National Institute of Advanced Industrial Science and Technology (AIST), 1-1-1 Higashi, Tsukuba, Ibaraki 305-8566, Japan ‡ University of Tsukuba, 1-1-1 Tenno-dai, Tsukuba, Ibaraki, 305-8571 Japan § National Metrology Institute of Japan, AIST, 1-1-1 Umezono, Tsukuba, Ibaraki 305-8563, Japan MES-Afty Corporation, 2-35-2 Hyoe, Hachioji, Tokyo 192-0918, Japan ABSTRACT: We describe the electrochemical detection of DNA methylation through the direct oxidation of both 5methylcytosine (mC) and cytosine (C) in 50 -CG-30 sequence (CpG) oligonucleotides using a sputtered nanocarbon film electrode after digesting a longer CpG oligonucleotide with endonuclease P1. Direct electrochemistry of the longer CpG oligonucleotides was insufficient for obtaining the oxidation currents of these bases because the CG rich sequence inhibited the direct oxidation of each base in the longer CpG oligonucleotides, owing to the conformational structure and its very low diffusion coefficient. To detect C methylation with better quantitativity and sensitivity in the relatively long CpG oligonucleotides, we successfully used an endonuclease P1 to digest the target CpG oligonucleotide and yield an identical mononucleotide 20 deoxyribonucleoside 50 -monophosphate (50 -dNMP). Compared with results obtained without P1 treatment, we achieved 4.4 times higher sensitivity and a wider concentration range for mC detection with a resolution capable of detecting a subtle methylated cytosine difference in the CpG oligonucleotides (60mer).

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enomic DNA methylation, frequently observed at C5 of cytosine (C) in the 50 -CG-30 sequence (CpG), plays an important role in the regulation of gene transcription, embryogenesis, and various diseases such as cancer, without any change in the DNA sequence.14 CpG units with a high density constitute the main part of the promoter region of the gene (CpG island: several tens to hundreds of CpG repetitions), and the CpG methylation density is closely associated with the transcription level of a gene.14 In normal cells, most CpG islands spanning the promoter regions are unmethylated, and their downstream genes are transcriptionally active. In contrast, when CpG island promoters in cancer cells are methylated, their downstream genes such as tumor suppressor genes are consistently silenced.14 These phenomena clearly demonstrate that the CpG methylation density is a critical hallmark for research in this field. Therefore, the methylation quantification of the CpG repetition sequence is important with respect to understanding DNA methylation related diseases and developing diagnosis tools. A conventional sequencing technique cannot be used for a DNA methylation assay, since both methylcytosine (mC) and C bases exhibit identical WatsonCrick base-pair behavior. Thus, current DNA methylation assays involve technologies for distinguishing both mC and C in DNA, including bisulfites,510 restriction enzymes,11,12 and molecules with biological/chemical affinity.1319 In the bisulfite-based assay, C is deaminated and r 2011 American Chemical Society

then converted to uracil, whereas mC remains almost unchanged, owing to the extremely low reactivity.5 The use of the above bisulfite and restriction enzyme-based techniques commonly provides a C-positive assay. Nevertheless, the bisulfite-based methods are currently considered gold standard assay techniques because of their applicability, despite the fact they are complicated and require several steps for mC detection, and this is time-consuming. In contrast, certain recent studies have used antibodies,13 binding proteins,1416 and metal complexes1719 with high affinity for mC that can be read out as an mC-positive assay, as well as some analysis with HPLC. More recently, Flusberg and co-workers reported a new methodology consisting of the direct detection of DNA methylation during singlemolecule, real-time (SMRT) sequencing using a nanostructured zero-mode waveguide device. With this technique, the presence of mC in a DNA template affected polymerase kinetics during SMRT sequencing that made it possible to distinguish mC from C in DNA.20 Electrochemical techniques have been widely used for DNA analysis.2132 These techniques are very simple and inexpensive and so are expected to be used as one of the postlight DNA analysis methods, including coulometric detection,2124 Received: July 7, 2011 Accepted: September 10, 2011 Published: September 11, 2011 7595

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Analytical Chemistry amperometric detection with electroactive tags or intercalators,2530 or potentiometric detection using ion-sensitive field-effect transistor (ISFET).31,32 Recently, we developed a methylation detection method based on the nonlabel electrochemical detection of both mC and C in oligonucleotides using nanocarbon film electrodes.33,34 This film was formed by employing the electron cyclotron resonance (ECR) sputtering method that provided a nanocrystalline sp2 and sp3 mixed bond structure.3537 We previously reported that a nanocarbon film electrode can distinguish both mC and C bases individually, by measuring the peak potential differences (150 mV) caused by C methylation.33 This is because a nanocarbon film electrode has a wide potential window while maintaining the high electrode activity needed to oxidize both bases. The potential window of the nanocarbon is from 1.7 to 2.0 V (vs Ag/AgCl and current limit < (500 μAcm2).38 The anodic limit is almost equivalent to that of a commercially available boron-doped diamond (BDD) electrode. Moreover, this film electrode allowed us to realize the quantitative detection of DNA methylation ratios solely by measuring methylated CpG repetition oligonucleotides (60mers, MW > 15 000) while largely avoiding surface fouling.34 Although it has been reported that certain electrodes allow the direct oxidation of large molecules,39,40 the electrochemical reaction for longer DNA molecules is insufficient because such macromolecular analytes generally exhibit low electrochemical sensitivity because the average distance between each base and electrode surface increases as a result of conformational problems, and the diffusion coefficient decreases with increasing molecular size.4042 The longer DNA sequences also foul the electrode surface in a much shorter time since the interaction with the electrode surface increases as the size of the DNA molecule increases even when new electrode materials such as BDD and our nanocarbon film electrodes are used. Further improvement of the quantitative performance including sensitivity and stability against mC and C in these longer sequences will be necessary if we are to develop a DNA methylation assay technique without a separation process. An appropriate way to achieve this is the identical digestion of a target oligonucleotide since the sensitivity of smaller molecules in direct electrochemistry is superior to that of macromolecules with respect to the diffusion coefficient and the electrochemical distance as a result of the conformational structure. Here, we report the electrochemical detection of DNA methylation in enzymatically digested CpG oligonucleotides based on direct oxidation employing nanocarbon film electrodes. We used an ECR sputtered nanocarbon film electrode with optimized electrochemical properties that were realized by changing the ion acceleration voltage during sputtering and electrochemical pretreatment, respectively, depending on the analytes. Film with an sp2/(sp2 + sp3) ratio of 0.6 has a sufficiently wide potential window with high electrode activity and stability against fouling caused by the biomolecule oxidation.33,34,43 As stated above, a longer DNA molecule exhibits lower electrochemical reactivity in direct electrochemistry.4042 In fact, we observed different electrochemical sensitivities for oligonucleotides with different lengths. Figure 1 shows background-subtracted square wave voltammograms (SWVs) for 20 -deoxyribonucleoside 50 monophosphates (50 -dNMPs) mixtures and two kinds of synthetic CpG repeated sequences with different lengths (20mer; CpG20 = 50 -CGCGCGCGCG CGCGCGCGCG-30 , 60mer; CpG60 = 50 -CGCGCGCGCG CGCGCGCGCG CGCGCGCGCG CGCGCGCGCG CGCGCGCGCG CGCGCGCGCG-30 , all CpG samples were purified using HPLC). Note that all the samples show the equivalent concentration of each base. When we look at the responses of G, its electrochemical responses from

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Figure 1. Background subtracted SWVs of 60 μM dGMP + 60 μM dCMP, 6 μM CpG oligonucleotide (20mer, CpG20), and 2 μM CpG oligonucleotide (60mer, CpG60) at the nanocarbon film electrode in 50 mM, pH 4.4 acetate buffer containing 2 M NaNO3. Amplitude = 25 mV; ΔE = 5 mV; f = 10 Hz.

the inherent CpG oligonucleotides were much smaller than that of dGMP in the mixtures, despite the fact that these samples have the same base concentrations. With CpG20, the G oxidation current was 24% of that of the dNMP mixtures. The current decrease became more pronounced during the measurement of the longer CpG60 (13% of that of the mixtures). In general, larger molecules show smaller electrochemical signals because of their lower diffusion coefficient.44,45 Indeed, we calculated the oxidation current46 of G in CpG60 theoretically by considering the diffusion coefficient decrease and found that the current fell to approximately 50% of that of dGMP. However, this decrease is much larger than the experimental data shown above, even taking account of the reduction in the diffusion coefficient of the longer oligonucleotide. This is probably because the CpG oligonucleotides used in this study, which consist of only a CG rich structure, are also capable of forming double strand oligonucleotide conformations via an inter/intra interaction. In fact, such structures were observed using certain kinds of electrophoresis measurement (polyacrylamide gel electrophoresis (PAGE) or capillary electrophoresis (CE)). This formation makes it more difficult to achieve the direct oxidation of the DNA bases because DNA bases of dsDNA are distributed inside the helix as other groups have also suggested.40,41,45 As a result, this causes a decrease in the quantitativity of the mC measurement in the oligonucleotide. To improve the quantitativity for the base content, especially the mC of the CpG oligonucleotide samples, we used nuclease P1 to digest the CpG60 before direct electrochemical oxidation in this study. nuclease P1 is well-known as an endonuclease, which cleaves the 30 -phosphodiester bond of single-stranded DNA/ RNA samples yielding an identical 50 -dNMP.4749 The reactivity of nuclease P1 depended on the concentration and length of the target DNA. Although in some cases nuclease P1 does not recognize or react on double strand DNA when there is a high salt concentration, the enzyme reacted completely on the duplexed oligonucleotide in our sample.47 We investigated the nuclease P1 activity against CpG60 samples, especially the methylated CpG60, using a CE based on a microfluid chip (Bioanalyzer Agilent 2100) since it is unclear whether or not P1 activity against methylated oligonucleotide is equivalent to that of an unmethylated sample. 7596

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Analytical Chemistry Therefore, it is important to study how different P1 nucleases catalyzed the digestion of the methylated CpG sample compared with the unmethylated CpG sample. Figure 2 outlines our experimental procedures when nuclease P1 is used. Briefly, 1 μL of a 100 μM CpG60 sample was treated with 10 μL of 2 mU/μL nuclease P1 (Yamasa Shoyu, Co., Tokyo) dissolved in 40 mM sodium acetate buffer (containing 0.2 mM ZnCl2, pH 5.3) and 9 μL of water. The solution (20 μL) was incubated at 37 °C (0120 min) and then held at 95 °C for 20 min. The digested sample solution was centrifuged at 15 000 rpm (15 min) using a spincolumn to remove the enzyme. Figure 3 shows an electrophoregram of unmethylated and methylated CpG60 (methylation ratio = 100%) enzymatically digested using nuclease P1. In both CpG60 samples, we observed some peaks at 50100 bp. These results

Figure 2. Schematic diagram of the experiment performed in this study.

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suggest that both samples formed high-order structure conformations including double strand oligonucleotide conformations via an inter/intra interaction as described above. With the unmethylated CpG oligonucleotide, P1 exhibited high enzyme activity that resulted in almost 100% digestion within 10 min. On the other hand, the reaction with the methylated CpG oligonucleotide was much slower than that obtained for the unmethylated CpG oligonucleotide. In fact, we observed a small broad peak at 1525 bp in the electrophoregram even after 90 min of the P1 reaction (Figure 3b). This suggests that C methylation in the oligonucleotide affects the P1 reactivity. Therefore, we optimized the experimental conditions for the digestion of the methylated CpG samples and found that the optimized condition for almost 100% digestion of the methylated sample was 120 min as shown in Figure 3b. Subsequent experiments were therefore performed under the above conditions. We measured the digested CpG60s with different numbers of mC using a nanocarbon film electrode. The CpG60 (methylation ratio = 0, 30, 50, 66, 70, and 100%) consisted of 0, 9, 15, 20, 21, and 30 of mC from 50 -end in the oligonucleotide, respectively. Figure 4 shows background-subtracted SWVs of the digested samples from the unmethylated CpG oligonucleotide and its methylated CpG oligonucleotides. This P1 treatment works well, and the current responses coincide precisely with the base content of each inherent oligonucleotide at the nanocarbon film electrode. At the same time, these voltammograms exhibited almost the same responses produced for dGMP oxidation. These current responses for dGMP in the CpG60 sample obtained after P1 treatment agreed well with that of the equivalent dGMP as a standard sample, which is unlike the result in Figure 1. These results also reveal that we succeeded in realizing an almost 100% enzyme reaction as estimated in Figure 3. Moreover, the treatment effectively provided clearer peak separation between mC and C. This is presumably due to the decrease in the current obtained for the digested samples in the higher potential region.

Figure 3. Typical electrophoregrams of (a) unmethylated CpG60 and (b) methylated CpG60 (methylation ratio = 100%). In the electrophoregrams in (a), the P1 enzyme reactions were 0 and 10 min in the left and right panels, respectively. In the electrophoregrams in (b) the reaction times were 0, 90, and 120 min in the left, middle, and right panels, respectively. The two peaks at 15 and 1500 bp indicate the reference markers. 7597

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Analytical Chemistry

Figure 4. Background subtracted SWVs of 2 μM of digested CpG60s with different methylation ratios at the nanocarbon film electrode in 50 mM, pH 4.4 acetate buffer containing 2 M NaNO3. The SWV conditions are the same as those in Figure 2.

Therefore, the oxidation currents of both mC and C in the digested samples could be superior to those obtained by direct detection of the inherent CpG oligonucleotide, revealing that the C methylation ratio in oligonucleotides is quantified from the heights of the two peak currents. We estimated the resolution of our method regarding DNA methylation detection for a P1 digested CpG60 sample. Figure 5 shows the relationship between the methylation ratio in the oligonucleotides and the current response assigned to the mC of the digested CpG oligonucleotides with different methylation ratios. The slope for the calibration of the P1 treated sample was more than 4.4 times greater than that obtained for a direct measurement without P1 treatment, indicating that this method enabled us to quantify the methylation ratio with higher sensitivity. It is noteworthy that we could achieve excellent discrimination of C methylation in oligonucleotides without any separation process. For example, the difference in the current response of the mC content obtained from 66% and 70% methylated CpG oligonucleotides is attributed to single C methylation for these CpG oligonucleotides. Furthermore, we found that the current responses formed a straight line that passes very close to the origin with a correlation coefficient of 0.996 (y = 0.0264x + 0.0323). On the other hand, although direct measurement (without P1 treatment) gave a straight line, it did not pass through the point of origin (y = 0.006x + 0.2056, r = 0.994). Therefore, the P1 treatment of the CpG samples played an important role in improving the sensitivity and the linear range of the calibration curve. Several groups have reported the quantification of mC in DNA samples using nuclease coupled with HPLC (UV or MS detection) to separate mC and C nucleotides.5053 Although the digestion/HPLC technique provides no information about the specific loci of methylated DNA samples, it is one of the most reliable and sensitive methods for detecting total DNA methylation.5456 More recently, HPLC with electrochemical detection (HPLC-ECD), which generally provides superior sensitivity to the conventional HPLC-UV method, was also employed for a DNA methylation assay using a BDD electrode after chemical

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Figure 5. Calibration of methylation ratio obtained from SWVs of some CpG60 samples with (a) and without (b) nuclease P1 treatment.

hydrolysis.57 The HPLC-based technique is time-consuming in regard to the separation steps, whereas the DNA methylation detection method described in this study is based on the direct electrochemical oxidation of both bases, meaning there is no need for a separation system. These factors are highly advantageous with a view to achieving the rapid and separative measurement of mC and C in oligonucleotides with the high sensitivity needed to detect a single C methylation difference. Of course, our nanocarbon film electrode can also be employed as an HPLC detector since its electrochemical activity for DNA bases is superior to that of a BDD electrode as a result of its sufficient potential window and low biomolecule adsorption compared with conventional GC electrodes. Our approach can achieve more sensitive and quantitative results in a DNA methylation assay. For this, we must evaluate our method in terms of reliability and versatility for detecting a single C methylation from various types and lengths of sequences including non-CpG samples. In conclusion, we successfully used nuclease P1 to digest methylated CpG oligonucleotides and thus realized more quantitative and sensitive detection of the DNA methylation ratio by direct electrochemical oxidation using a nanocarbon film, without the need for bisulfite treatment or separation processes. With nuclease P1, there was an approximately 4.4-fold increase in the sensitivity for both C- and mC-positive assays compared with an assay without the enzyme. As a result, we could detect a subtle C methylation difference in 60mer CpG oligonucleotides without any separation process. We will use this method to detect DNA methylation in much longer oligonucleotides including non-CpG sequences, where it would be hard to use direct electrochemical oxidation without an enzyme. This technique will be necessary if we are to detect DNA methylation in real genomic samples.

’ AUTHOR INFORMATION Corresponding Author

*Address: National Institute of Advanced Industrial Science and Technology (AIST), Tsukuba Central 6, 1-1-1 Higashi, Tsukuba, 7598

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Analytical Chemistry Ibaraki, Japan. Tel: +81-29-861-6158. Fax: +81-29-861-6177. E-mail: [email protected].

’ ACKNOWLEDGMENT This work was supported by a Grand-in-Aid for Scientific Research (O.N. No. 23350038 and No. 20350042) from the Ministry of Education, Culture, Science, Sports and Technology of Japan. We thank Shiori Yamazaki for help with the CE experiments. We also thank Kohei Nakamoto (University of Tsukuba) for his creative support in the preparation of the TOC graphic. ’ REFERENCES (1) Laird, P. W. Nat. Rev. Cancer 2003, 3, 253–266. (2) Zhang, Z. J.; Chen, C. Q.; Manev, H. Anal. Chem. 2004, 76, 6829–6832. (3) Ushijima, T. Nat. Rev. Cancer 2005, 5, 223–231. (4) Frigola, J.; Song, J.; Stirzaker, C.; Hinshelwood, R. A.; Peinado, M. A.; Clark, S. J. Nat. Genet. 2006, 38, 540–549. (5) Robert, S.; Robert, S.; Marvin, W. J. Am. Chem. Soc. 1970, 92, 422–424. (6) Frommer, M.; Mcdonald, L. E.; Millar, D. S.; Collis, C. M.; Watt, F.; Grigg, G. W.; Molloy, P. L.; Paul, C. L. Proc. Natl. Acad. Sci. U.S.A. 1992, 89, 1827–1831. (7) Herman, J. G.; Graff, J. R.; Myohanen, S.; Nelkin, B. D.; Baylin, S. B. Proc. Natl. Acad. Sci. U.S.A. 1996, 93, 9821–9826. (8) Chen, R. Z.; Pettersson, U.; Beard, C.; Jackson-Grusby, L.; Jaenisch, R. Nature 1998, 395, 89–93. (9) Dupont, J. M.; Tost, J.; Jammes, H.; Gut, N. G. Anal. Biochem. 2004, 333, 119–127. (10) Chhibber, A.; Schroeder, B. G. Anal. Biochem. 2008, 377, 46–54. (11) Toyota, M.; Ho, C.; Ahuja, N.; Jair, K. W.; Li, Q.; Ohe-Toyota, M.; Baylin, S. B.; Issa, J. P. J. Cancer Res. 1999, 59, 2307–2312. (12) Yamada, H.; Tanabe, K.; Nishimoto, S. Bioorg. Med. Chem. Lett. 2005, 15, 665–668. (13) Weber, M.; Davies, J. J.; Wittig, D.; Oakeley, E. J.; Haase, M.; Lam, W. L.; Schubeler, D. Nat. Genet. 2005, 37, 853–862. (14) Rauch, T.; Li, H. W.; Wu, X. W.; Pfeifer, G. P. Cancer Res. 2006, 66, 7939–7947. (15) Blair, S.; Yu, Y. N.; Gillespie, D.; Jensen, R.; Myszka, D.; Badran, A. H.; Ghosh, I.; Chagovetz, A. Anal. Chem. 2010, 82, 5012–5019. (16) Craighead, H. G.; Cipriany, B. R.; Zhao, R. Q.; Murphy, P. J.; Levy, S. L.; Tan, C. P.; Soloway, P. D. Anal. Chem. 2010, 82, 2480–2487. (17) Tanaka, K.; Tainaka, K.; Kamei, T.; Okamoto, A. J. Am. Chem. Soc. 2007, 129, 5612–5620. (18) Tanaka, K.; Tainaka, K.; Umemoto, T.; Nomura, A.; Okamoto, A. J. Am. Chem. Soc. 2007, 129, 14511–14517. (19) Bareyt, S.; Carell, T. Angew. Chem., Int. Ed. 2008, 47, 181–184. (20) Flusberg, B. A.; Webster, D. R.; Lee, J. H.; Travers, K. J.; Olivares, E. C.; Clark, T. A.; Korlach, J.; Turner, S. W. Nat. Methods 2010, 7, 461–U72. (21) Kelley, S. O.; Boon, E. M.; Barton, J. K.; Jackson, N. M.; Hill, M. G. Nucleic Acids Res. 1999, 27, 4830–4837. (22) Drummond, T. G.; Hill, M. G.; Barton, J. K. Nat. Biotechnol. 2003, 21, 1192–1199. (23) Fang, Z. C.; Kelley, S. O. Anal. Chem. 2009, 81, 612–617. (24) Slinker, J. D.; Muren, N. B.; Gorodetsky, A. A.; Barton, J. K. J. Am. Chem. Soc. 2010, 132, 2769–2774. (25) Hashimoto, K.; Ito, K.; Ishimori, Y. Anal. Chem. 1994, 66, 3830–3833. (26) Ihara, T.; Nakayama, M.; Murata, M.; Nakano, K.; Maeda, M. Chem. Commun. 1997, 1609–1610. (27) Inouye, M.; Ikeda, R.; Takase, M.; Tsuri, T.; Chiba, J. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 11606–11610.

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