Museum DNA extractions/PCRing

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Museum DNA extractions/PCRing

Post by LMOE » Thu Jun 23, 2005 4:48 pm

I am having problems extracting/PCRing my museum DNA samples. I am using rodent skins.

For the extraction, I have been using a Qiagen Kit with a special (homemade) lysis buffer which contains 10mM Tris-Cl (pH 8.0), 10mM EDTA, 100 mM NaCl, 40mM DTT, 2% SDS, and 250 micro-grams/milli-liter Prot. K. I have been doing my initial digestion with pro-k (and the rest of the reagents) for 24 hours.

For the PCR, I am using microsatellite (nuclear) primers. I have actually successfully amplified my museum samples using mitochondrial, it does appear that I am successfully extracting mitochondrial DNA. When I PCR using my microsat. primers, I do not get any bands on an agarose gel nor do I get any bands on a polyacrylimide capillary gel. For my PCR, I use Taq beads that contain the following: 2.5 units of PuReTaq DNA polymerase, 10 mM Tris-HCl, (pH 9.0 at room temperature), 50 mM KCl, 1.5 mM MgCl2, 200 µM of each dNTP, stabilizers, and BSA. All of my PCR reactions are run with a positive control (high quality, non-museum DNA), which successfully amplifies.

I am not sure if my problem is with my extraction or with my PCR. I have already tried concentrating (pelleting) down the extracted DNA in a vacuum centrifuge and rehydrating it with 10 micro-liters of water, but the PCR still did not work.

I have successfully extracted/amplified DNA using my microsatellite primers from fresh muscle tissue.

If anyone has any ideas, I would be very, VERY greatful.


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