I would like to design a very simple, idiot-proof protocol for measuring the relative amount of methylation in the genome as a whole (working with non-model organisms). That is, I would like to be able to show that the genome of organism A has about 3% cytosine methylation and that of organism B has 3.5% (ideally with high accuracy, as my organisms are insects and thus have less methylation than mice/humans). I would also settle for having methylation in arbitrary units relative to some sort of reference. I don't care where the methylation is in the genome (yet) - I only want to know the genome-wide rate of methylation.
Here is my current plan - is there anything wrong with it? I mostly don't do molecular work, so it might be way off:
1. Extract the DNA and sonicate it into smallish pieces (150bp?).
2. Denature the DNA into single-stranded DNA at 95oC - this is needed because the antibody only binds ssDNA.
3. Add primary antibody that binds to 5-meC, and incubate for 2 hours at 4oC.
4. Add secondary antibody, and incubate - this will stick to the primary-labelled DNA, and presumably also to the unbound antibody in the supernatent.
5. Add AMPure beads - all the DNA sticks to them, both antibody-labelled DNA and the non-labelled DNA pieces.
6. Discard supernatent and wash the beads a few times. This removes the excess primary and secondary antibody.
7. Remove the DNA from the beads with elution buffer, and measure the amount of (methylated) DNA using a plate reader. Then, measure the total amount of DNA using Qubit ssDNA kit.
8. Take the ratio of the two readings in step 7 to get a number expressing the relative amount of genome-wide DNA methylation.
9. Repeat steps 1-8 with some reference samples which contain 100% methylated DNA and 100% non-methylated DNA, mixed in various ratios like 100:1, 50:1, 10:1 etc. These will allow the values in step 8 to be converted into estimates of the % methylation rate.
Any tips are much appreciated!