Bisulfite sequencing of very small samples

Introduction

(based on Olek et al., 1996, Schoenherr et al., 2003)

DNA methylation is a stable epigenetic mark, which can mediate gene silencing. Bisulfite sequencing allows for precise identification of methylated cytosines within DNA (Frommer et al. 1992). This method is based on different rate of chemical conversion of methylated and non-methylated cytosines to uracil (Figure 1) where non-methylated cytosines are converted efficiently while methylated cytosines remain non-reactive. This method was further developed by embedding analyzed DNA into an agarose bead (Olek et al. 1996). The protocol presented here was further optimized for bisulfite sequencing of small samples where the starting material was a small number of cells (Fedoriw et al. 2004; Svoboda et al. 2004). The smallest amount of material from which several unique clones were recovered was 25 oocytes, which corresponds to 100 DNA molecules in the initial material (Svoboda et al. 2004). This protocol can be also used for analyzing up to 200ng purified genomic DNA in one sample

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Procedure

Primer design

Good primers are absolutely critical for successful bisulfite sequencing. Use genomic DNA sequence from the region you want to amplify (It sounds obvious but it is not uncommon that people accidentally design primers for cDNA sequence).

Convert genomic DNA in silico:

1. Open sequence file (FASTA format) in Word
2. Edit/Replace - replace CG with XY
3. Edit/Replace - replace C with T
4. Edit/Replace - replace XY with CG
5. Use this sequence to design primers

Design primers � We manually select primer sequences to amplify a region of interest and then we check the Tm and other parameters in the Vector NTI. Primers are usually ~ 28-30 nt long and they have Tm approx. 55 °C (comment 1). Choose primers such that they contain 5-8 G’s and try to choose primers that have higher sequence complexity. Try to avoid CpG’s in primer sequences � if you have no choice, use degenerated nucleotide in that position. Also, try to avoid having very T-rich sense and A-rich antisense primers, which would pair too much. If there is no other choice, you can accept -ΔG up to approx. �2.0 if it is a single strong pairing in the middle or at the 5’ end of the primer and it does not provide a good template for Taq pol (which could inactivate this primer pair). Keep the PCR product under 600 bp (ideally 300-350 bp).

Day 1 (if your starting material is DNA go to day 2)

1. Prepare 2% LMP Agarose in pure water. Melt it in the microwave, mix on magnetic stirrer (note 1).
2. Add 20 µL of melted agarose to cells placed at the bottom of a 2 mL Eppendorf tube, spin briefly down (note 2).
3. Overlay with 300 µL of mineral oil and incubate for a few minutes at 65 °C (note 3).
4. Chill on ice for 2 minutes to solidify
5. Add 800 µL of Lysis buffer to the bead.
6. Add 2 µL od Proteinase K per sample (40 µg per reaction � stock 20 mg/ml)
7. Incubate overnight at 50 °C for 12 to 14 hours, with gentle shaking if possible (note 4).

Day 2

1. Wash the bead 3 times with 1 mL of TE (10 mM Tris-HCL, pH 7.6, 1 mM EDTA) for 15 minutes with gentle mixing (note 5).

If you are using genomic DNA start here:

1. • Add 30 µL of melted 2% LMP Agarose to DNA (up to 200 ng in up to 4 µL)
   • Overlay with 300 µL of paraffine oil and incubate for a few minutes at 65 °C
   • Chill on ice for 2 minutes to solidify


2. Add 500 µL of 0.3M NaOH and incubate for 15 min, remove NaOH solution.
   (denaturation of DNA strands)
3. Repeat incubation with 500 mµL of 0.3M NaOH for 15 min, remove NaOH solution.
4. Incubate with 500 µL of 0.1M NaOH for 10 min, remove NaOH solution.
   (to solidify agarose beads)
5. In the meantime start preparing fresh 10 mM hydroquinone and 40.5% Sodium bisulfite in pure water (note 6). Both solutions are light sensitive, so try to avoid any unnecessary exposure to light (wrap Falcon tubes with aluminum foil, cover beaker with dissolving sodium bisulphate etc.).
6. Transfer the bead into a new 2 mL Eppendorf tube:
   • Cap the tube containing the bead with a cap of a new tube, invert and tap on the tube with the bead � the bead drops on the cap.
   • Then release carefully the cap with the bead, close the new tube and tap on it to place the bead at the bottom. This step prevents diluting LMP Agarose and increasing bead size in subsequent steps.
7. Overlay the bead with 300 µL of mineral oil, heat to 80 °C for 15 minutes.
   (full separation of individual DNA strands)
8. Resolidify the agarose bead by chilling on ice (at least 2 minutes).
9. Set up the bisulfite reaction. Add 15 µL of 10 mM hydroquionone and 255 µL of 40.5% Sodium bisulfite
10. Incubate at 50 °C for 4-6 hours with gentle mixing (e.g. Eppendorf Thermomixer at 400 rpm), cover with aluminum foil to protect samples against light (note 7).
11. Stop the bisulfite reaction by removing solutions and mineral oil and wash the bead with 1 mL of TE for 15 minutes.
12. Repeat wash 5 times with TE. If necessary, you can store the bead in TE after the second wash at 4 °C overnight and continue the next day
13. Incubate the bead with 500 µL of 0.2 M NaOH for 15 minutes at room temperature, remove NaOH.
14. Incubate the bead with 500 µL of 0.2 M NaOH for 15 minutes at 37 °C. (desulfonation step)
15. Neutralize NaOH with 100 µL of 1M hydrochloric acid.
16. Briefly wash the bead with 1 mL of TE.
    (pH is often not neutral after HCl neutralization, this step assures proper neutralization)
17. Wash twice with 1 mL of pure water, 15 minutes each.
18. After removing the last wash, transfer the bead into a new Eppendorf tube.
19. Add pure water to the final volume of approx. 100 µL. Heat the tube at 80 °C for 5 min, mix briefly by vortexing and use aliquots of diluted melted agarose for PCR reaction (note 8).

Suggestions for PCR, cloning and sequencing of PCR products

PCR

For very small samples: to assure detection of as many individual strands as possible we recommend to run more PCR reactions for one locus (four to ten) and then clone individual amplicons separately. If one uses a single PCR reaction for a very small sample, there is a high risk of a strong clonal effect � i.e. a large fraction of sequenced clones will be derived from a single converted template molecule. If the methylation pattern is homogeneous, one cannot distinguish between identical sequences stemming from the same single DNA molecule and sequences stemming from different molecules with the same pattern. This problem can be addressed if one generates independent amplicons and clones them separately. Identical sequences cloned from different amplicons give a better idea about the distribution of DNA methylation.

When starting with 200 ng of genomic DNA, clonal effects are relatively small, so one PCR reaction for each locus is typically sufficient. We usually use 1/10 of a melted bead in 50 µL PCR reaction. If there is a suspicion of clonal effects we suggest to run more individual PCRs on the rest of the sample.

When setting up PCR, we melt the sample in the Eppendorf Thermomixer (800 rpm) while preparing PCR mastermix. Then we divide diluted melted agarose into PCR tubes containing the rest of the PCR reaction mixture. It is recommended to use a hot-start PCR to avoid nonspecific primer amplification (comment 2). We use Amplitaq GOLD polymerase, which requires heat activation (15 minutes at 94 °C).

We typically use 20-25 µL of diluted melted agarose in 100 µL PCR for very small samples (20-50 oocytes) and 10 µL of diluted melted agarose in 50 µL PCR for genomic DNA (comment 3). To further improve specificity of amplification, we use touch-down PCR. The following program works well for primers with Tm ~ 55 °C

94 °C for 15 min

94 °C for 30 sec
62->55 °C for 30 sec - 14 cycles with a gradual decrease of Tm 0.5 °C per cycle
72 °C for 1 min

94 °C for 30 sec
55 °C for 30 sec - 36 cycles
72 °C for 1 min

72 °C for 15-20 min - 1 cycle

It is possible to sequence the PCR product directly but we do not recommend it as it yields misleading results. The main reason is that the height of peaks in the sequencing chromatogram is extremely biased because of the biased nucleotide content of the sequence. In some experiments direct PCR product sequencing can produce a completely different results compared to the analysis of individual clones (figure 2). Therefore, for direct analysis of PCR products one should use an appropriate quantitative approach such as, for example, pyrosequencing (comment 4).

PCR cloning:

It is possible to use completed PCR reaction directly for cloning but we prefer to purify amplicons via gel extraction. We run 50 µL of PCR reaction in 1.2% agarose gel (regular agarose, in 1x TAE or 0.5 x TBE), gel extract amplicons (Qiagen Gel extraction kit, final elution volume 30 µL) and use 4 µL for TOPO TA cloning (Invitrogen, TOPO TA cloning kit � dual promoter). As noted above, if you work with small samples, it is better to clone each amplicon separately. Use blue-white selection for TOPO TA cloning.

Pick at 5-10 colonies from each plate for miniprep. Submit for sequencing (SP6 primer is the best for pCR II plasmid � for some reason the PCR product tends to be cloned into the vector in one orientation (antisense with respect to the pCR II sequence). Sequencing of GT rich regions is often very difficult and because of the bias in cloning orientation, the SP6 primer usually sequences the easier-to-sequence CA-rich strand).