Hello everyone,
I am having some trouble with the sample prep I am doing pre-Illumina sequencing. I have modified the standard protocol, which I will attempt to explain as clearly as possible. I begin by fragmenting genomic DNA to about 2kb in size. I then do what essentially amounts to T-linker ligation. First, I do a primer extension reaction using a primer that is specific to a fairly abundant region of DNA in the genome (eg, Alu or LINE-1). This selectively adds 3' Adenine overhangs to those fragments that contain the sequence I want to select for, allowing the T-linker ligation to occur in a selective manner. I ligate a duplex T-linker to my fragments. Then I perform a nested PCR, first using the same primer used for the primer extension (and a linker-specific primer). Then, in the second (nested) round of PCR, I add the necessary sequences required for PE cluster generation and sequencing by adding them to the 5' end of my internal (nested) primers.
This second PCR reaction is what goes onto an agarose gel to be size-selected. I cut bands at about 500bp and 750bp and then amplify using the PE Illumina Primers and Phusion Mastermix (as per the Illumina protocol) for 10 cycles to enrich for the DNA. However when I run the samples on both a bioanalyzer and a gel, the fragments that are created by the PCR are not the expected 500 and 750 bp in size, but rather are only about 250 bp (and both are the same). I am at a loss for what might be happening here, and would appreciate any and all suggestions.
Hope I haven't made the explanation of my modifications too complicated.
Thanks!
I am having some trouble with the sample prep I am doing pre-Illumina sequencing. I have modified the standard protocol, which I will attempt to explain as clearly as possible. I begin by fragmenting genomic DNA to about 2kb in size. I then do what essentially amounts to T-linker ligation. First, I do a primer extension reaction using a primer that is specific to a fairly abundant region of DNA in the genome (eg, Alu or LINE-1). This selectively adds 3' Adenine overhangs to those fragments that contain the sequence I want to select for, allowing the T-linker ligation to occur in a selective manner. I ligate a duplex T-linker to my fragments. Then I perform a nested PCR, first using the same primer used for the primer extension (and a linker-specific primer). Then, in the second (nested) round of PCR, I add the necessary sequences required for PE cluster generation and sequencing by adding them to the 5' end of my internal (nested) primers.
This second PCR reaction is what goes onto an agarose gel to be size-selected. I cut bands at about 500bp and 750bp and then amplify using the PE Illumina Primers and Phusion Mastermix (as per the Illumina protocol) for 10 cycles to enrich for the DNA. However when I run the samples on both a bioanalyzer and a gel, the fragments that are created by the PCR are not the expected 500 and 750 bp in size, but rather are only about 250 bp (and both are the same). I am at a loss for what might be happening here, and would appreciate any and all suggestions.
Hope I haven't made the explanation of my modifications too complicated.
Thanks!
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