Restriction Enzyme Digest of Genomic DNA - Problem with RE-digest + PCR in CpG island assay (Jul/18/2009 )
So my problem is as follows;
I want to screen the promoter region of a gene that I suspect is silenced by methylation in cancer.
Now what most of you think now is probably BiS-sequencing, MSP or something like that.
But nope, in my lab we do it the old-school way (Restriction enzyme digest followed by PCR, UnMe--> no band. Methylated--> band).
So I dezigned 4 primer pairs for my promoter and picked out 6 methylation sensitive RE's that (theoretically) allow me to probe the methylation status of 10 CpG's in the CpGisland.
Now to the problem part.
I'm trying to optimize the system now so what I do is I incubate 20ng of:
1) Fully methylated DNA
2) 5-AzaC treated (UnMe)
3) a random cancer cell line DNA (for the heck of it)
(also about 1ug of a PCR-product that I know the enzyme cuts and that I can image on a gel right after digest, just to see if the enzyme has worked)
I do this with and without enzyme (25 fold excess now, started at 10) over night in a 20ul volume.
I then use 5ul of my digest mix (5ng) as template for a PCR to hopefully get bands from both CpG-methylated templates (+/- enz), a band from the no RE 5AzaC template and something in between in the cell line version...
I run a Touchdown PCR protocol that has worked well for getting strong bands in the PCR-optimization stage but now my problem is that it works too well!
I get bands from all templates, I've adjusted the cycle count downwards to get it into a semi-quantitative range (want to roughly be able to tell the difference between fully methylated, unmethylated and 50%methylated) but my problem is that the 5-AzaC treated DNA outperforms the CpG methylated DNA every single time!!!
I'm soon at the detection limit for my other templates but the "unmethylated" DNA gives a stronger band than the others.
I read that the 5AzaC DNA is only 70% unmethylated but still I feel I should be able to digest away enough of it so that I can keep it from being detected (by adjusting the cycle count) while the CpG methylated would give bands.
But when looking at my post PCR bands I can hardly see any effect of RE-treatment on the 5AzaC DNA (Maby a little if I try hard), but still the RE-easily chews up 1 freaking ug of control DNA (incubation of all in the same PCR-strip using the same RE-mastermix so no variance there).
Should I throw away the 5AzaC DNA and just go with GenomiPhi or BioScore amplified template in the unmethylated control (or atleast try them to see if the 5AzaC thing is screwing with me)?
What can I do to fix this, is it fixable?
Im really running out of ideas but I have invested a lot of time in this so scrapping the setup would really not be fun.
The reason for why I'm using so little template DNA is that once the system is optimized I will be probing primary tumor DNA (which is a limited resource).
The concentrations for the templates have been determined using a NanoDrop and are within 1ng/ul of each other.
If anyone can come upp with suggestions for fixing my problem I would be gratefull (And sorry for writing such an essay).
I would suggest that give it up and switch to bisulfite based PCR methods. If restriction digestion is feasible for methylation assay, it should have become the gold strandard long time ago. The problem with restriction digestion is incomplete DNA digestion. Any residue will be amplified and shows up as false positive or negative.
In addition, Aza treated DNA is not a good unmethylated DNA control unless you have bisulfite sequenced the DNA that the treatment has removed all methyl-cytosines.