22 replies to this topic

[labtechie]

Hi-- I am a just-out-of-college lab tech with the assignment of determining methylation status of two extensive CpG islands in promoters of candidate tumor suppressor genes.  I have tried methylation-sensitive restriction then PCR (got it to work pretty well but had problems with false positives from incomplete digestion), bisulfite then sequencing (abandoned because my primers didn't work very well and I also am not in charge of the sequencing so cannot control special conditions)and am now optimistic about MSP.  My W primers work beautifully in untreated DNA using Roche Faststart Taq (some nonspecific bands due to low annealing temps but I can deal with that) so I am hoping my M and U primers will work as well, as they cover basically the same amplicon.  The problem is that I seem to have lost my DNA during the bisulfite treatment.  Now I have read every paper I can find with the words "bisulfite treatment" and "optimization" in it, and everybody uses slightly different methods.  I followed the protocol found here: http://www.protocol-...ailed/3160.html very closely except that I used Promega Wizard columns.  I think I lost the DNA during the Wizard clean-up column step, as I saw no pellets during the final EtOH precipitation.  Thus I was already expecting my sad result.  The other scary thing is that the only faint band I see in a "treated" lane is in the W lane, so it seems like not only did I lose most DNA but the bisulfite didn't work in the first place.

Does anyone have a "favorite" protocol they'd like to share?  My lab has suggested always using TE instead of water (to protect the DNA) and to find out better ways of cleaning up the DNA from the column.  I am not in favor of a kit as I see no reason to use "reagent 1" that is really NaOH and "reagent 2" that is just bisulfite at a 400% markup.  

Any advice you can offer would be MUCH appreciated.  I have been spinning my wheels for four months now and I only have this job for a year!  I need results!

[pcrman]

Hello,

First, loss of DNA during purification is one of the many possibilities for failed PCR, especially with Promega Wizard kit. I think using some bisulfite modification kit will help such as the kit from chemicon because of the built-in purification method.

Second, cycle your PCR up to 40 times if you have not done so.

[labtechie]

Hi all-- I have very good advice that comes from trying many different protocols for bisulfite treatment.  Use the one that involves embedding your DNA in agarose beads and using a higher concentration of bisulfite for a shorter amount of time.  It works SO WELL.  It is Olek et. al., 1996 or send me a message if you want me to forward you my slight modifications to the protocol.

[paulina]

Hi labtechie, could you share your method with us by posting it here?

Thanks.

[labtechie]

Here it is:
1. Set 300 ul mineral oil (I use Sigma M5904) in 2 ml tubes on ice for 30 minutes.
2. Prepare less than 1ug DNA (you may digest it with a restriction enzyme that has no site within your region of interest) in a volume of 21 ul TE.
3. Add 4  ul of 2M freshly prepared NaOH and incubate 15 minutes in a 50C water bath
4. Make a 2% LMP agarose solution (seaplaque agarose from cambrex) and add 50 ul of it to the 25 ul of DNA solution
5. Before it cools, form beads by gently pipetting 10 ul into the middle of the mineral oil layer (only one bead per tube—therefore up to six tubes per sample)
6. Leave for at least 30 minutes
7. Prepare bisulfite solution (avoid light by wrapping tube in foil): 3.8 g sodium betabisulfite in 5 ml H20 (this is 5 M) and also prepare 110 mg hydroquinone in 1 ml H20.  Mix these two solutions, check pH, and adjust pH to 5 with 2 M NaOH.
8. Add 500 ul of this solution to tubes—this is the important part. Pipet very gently to get the layers to separate correctly and check that your bead is in the aqueous layer. Other protocols say “invert”—that doesn’t work.
9. Cover tubes with foil (light exclusion) and incubate in hybaid for 4 hours at 50C.
10. The following steps require both p1000 and p200 pipets (avoid coming near the bead with the tip of the p1000, it may break it): remove oil and solution and rinse with 1 ml TE ph 8, 4 x 15 minutes (I use a rotating tray).
11. Desulfonate with 2 x 15 minutes in 500 ul 0.2 M NaOH.
12. Stop treatment in 1 ml TE 2 x 10 minutes, store overnight (or a little longer) in 1 ml TE.
13. Before PCR, rinse with 1 ml H20, 2 x 15 minutes.  Use one bead (something less than 100 ng) per PCR reaction
14. PCR hints:  Use a hotstart taq with NO PROOFREADING ABILITY—I use Faststart from Roche, it’s cheap. Also do a nested PCR and do not expecct any products from your large primers.


I am now working on cloning and sequencing these products, so if anyone has any hints.... they'd be much appreciated.  I'm using the PCR-script cloning vector from Stratagene.

-labtechie

[pcrman]

Thank you, labtechie, for sharing the protocol.

Quote

I am now working on cloning and sequencing these products, so if anyone has any hints.... they'd be much appreciated. I'm using the PCR-script cloning vector from Stratagene.

If you have got good PCR products, cloning and sequencing is not a big deal. Just purify your PCR products using Qiagen PCR or Gel purification kit and do a 5-min TA cloning. Pick 10 clones for sequencing (I have found ten represents well the actual profile and no more is needed).

[labtechie]

Hi pcrman,
Thank you SO much for all your help.  I think I'm confused as to how I get from the ten clones I'll pick to the sequence-- is it a miniprep followed by dideoxy sequencing?  Also, how do I get the clone off the plate?  Any advice would be much appreciated, as my boss is out of town and I have no idea what I am doing.
Thanks,
labtechie

Edited by labtechie, 11 January 2005 - 09:01 AM.

[pcrman]

After you have done a sucssessful PCR,  purify your PCR products and use the fresh products to do a TA cloning (ligation->transformation->plating). You will see white and blue colonies on the plates. For each PCR reaction (transformation) or DNA sample, randomly pick ten white, big, shiny and well isolated colonies using a yellow pipet tip and drop the tip into a 14-ml or 50-ml tube containing 2-5 ml LB medium with antibiotics added. Grow the bacteria overnight at 37C with virgorous shaking.  The next morning, take out the tubes and take 1 ml overnight growth for miniprep.  You can pick more than 10 colonies (such as 12) just in case some minipreps fail. If cost and labor is not a problem, screen 20 colonies. After you got the plasmid DNA, send it out for sequencing.

Hope that helps.

[labtechie]

Thank you so much!  That's exactly what I needed.

[methylnick]

labtechie, on Jan 7 2005, 07:19 AM, said:

11. Desulfonate with 2 x 15 minutes in 500 ul 0.2 M NaOH.

Labtechie,

I am not exactly sure about the chemistry of the bisulfite reaction and if NaOH is enough to desulfonate the adducts.

From my understanding, NaOH helps denature the DNA strands to expose the sulfonated adducts to another chemical such as ammonium acetate.

That could be a reason why you are not yielding converted DNA.

Please correct me if my understanding of the reaction is wrong anyone!

Nick

[labtechie]

With this new (agarose bead) protocol, I get great PCR and great sequence, so.... I think it's ok, but thanks for your concern  Anyone have any input?  I'm not seeing much methylation, but that wouldn't be a result of things not getting desulfonated properly, since in my understanding the 5mC doesn't react with bisulfite in the first place, right?

[methylnick]

labtechie, on Jan 26 2005, 06:54 AM, said:

I'm not seeing much methylation, but that wouldn't be a result of things not getting desulfonated properly, since in my understanding the 5mC doesn't react with bisulfite in the first place, right?

Yes you are absolutely correct, 5mC is unreactive to the bisulfite reaction.

[spjgjsm]

Thanks for that protocol labtechie! It worked great - it even successfully converted some really dodgy samples that the chemicon kit had failed to work on. Just to add, I've cut out the number of washes at the end and it still seems to work fine.

Jon

[labtechie]

Hi Jon!  That makes me so happy.  Questions:  How many washes did you cut it down to?  You still have to wash initially in TE and then in water before PCR, right?  Are you seeing any CpG methylation?  I see none, I assume because that's a negative result but I'm worried that perhaps my protocol works TOO well, I need a positive control   Do most people reading this buy fully methylated DNA or is there a region I can amplify that is known to always be methylated?  Also, the only artifacts I get are sometimes a non-converted nonCpG C a few bp away from a CpG.  Do you get that?  Interesting artifact, a result of steric interactions or secondary DNA structure maybe?

Just curious about all these things... so glad your protocol works!

Ellen

P.S.  All:  I do NOT pre-digest my DNA with a restriction enzyme and this protocol still works FINE.

[methylnick]

labtechie, on Feb 23 2005, 11:10 AM, said:

I need a positive control   Do most people reading this buy fully methylated DNA or is there a region I can amplify that is known to always be methylated?  Also, the only artifacts I get are sometimes a non-converted nonCpG C a few bp away from a CpG.  Do you get that?  Interesting artifact, a result of steric interactions or secondary DNA structure maybe?

Hi Ellen,

as a positive negative control, instead of buying methylated DNA it might be worth your while to methylate some DNA yourself. You can use SssI methylase (NEB #M0226) and this enzyme methylates every CG residue available. As a control I would methylate a plasmid such as pGEM and then "spike" my sample with the methylated plasmid prior to bisulfite treatment. You can assume that all CG's on the plasmid are methylated after SssI methylase and therefore should all come up as CG after bisulfite sequencing.

As for non-CG non conversion close to CG residues, this is due to incomplete denaturation of the strands and thus inefficent bisulfite conversion.  

Hope this helps

Nick