Hi all,
I've been attempting to ligate illumina adapters to a PCR amplified dsDNA fragment (sticky ends, 4bp overhang) but on all of my gels I can see the adapter and the insert separately, not ligated together.
I think part of this has to do with my insert (59bp) being so close in size to my adapters (79bp and 69bp). For this reason I have been using a 1:1 ratio of insert:adapter. I have tried 1ng, 100ng, 200ng, and 500ng. I have tried using the Blunt/TA Master Mix (NEB#M0367) with equal volumes of master mix and DNA/Adapter. I have also tried regular T4 Ligase (NEB#M0202), in a 50µL rxn I use 5µL T4 buffer and 100 Units T4 Ligase (I just scaled it from the protocol online-- is that too much?)
When I amplify the ligations (using primers that bind to the restriction enzyme cut site, I call them "bridge primers" because they bridge the insert to adapters) I get a product of the desired size, but when I send the sample for sequencing the only sequences that align are where the bridge primers bind.
I would love any advice on this, and I also have one specific question: I heard that spin column cleanup is not an ideal method for downstream ligations. Since my insert is PCR amplified, is there any other cleanup I can use to remove the excess salts and EDTA that might be an issue? I am doing SPRI bead cleanups between ligations, but what about cleaning up the PCR product before ligation?
qPCR is much more accurate than a microarray and much more specific than RNAseq
You can do HPLC to purify your pcr product. Also I can say that chelating agents may prevent the enzyme activity because they tend to keep metal ions. As a summary, during T4 ligase protocol, addition of Mg+2 keeps 2 phosphates of atp to make Atp sterically hindered. With the help of Magnesium, phosphates of Atp become reachable and than these phosphates are removed . Therefore, removing chelating agents and salts from your pcr products will fix the problem I think. HPLC is a nice application for your purpose.
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