I am currently developing a protocol to process approximately 400 samples using the Illumina MiSeq platform. We are using the two-step PCR approach and have designed our own primers for this.
The samples I am processing have already undergone a 35 cycle PCR once and as these samples are in a limited supply now, I am using a 1ul of a 1:10 dilution of the PCR product for each sample as the DNA template in the first PCR of the two required for the sequencing approach.
I was just looking for some further advice on protocol development. So far, I have managed to reduce the first PCR of the two steps to a 10 cycle PCR. I then planned to use a 96-well format PCR purification kit to clean the product up, quantify it and then use 50ng of DNA for each sample as the template in the second PCR. I have had a look through the Nextera XT sample preparation guide which has a PCR program for the second PCR, but this is 12 cycles as the Nextera kit is designed to amplify sequences from transposon tagged genomic DNA , so I was thinking of trying to reduce this to 5 cycles as I want maximum amplification for the minimum number of cycles and then clean up again before quantifying and pooling to get ready for sequencing.
Does anyone have any opinions/comments/recommendations?
The protocol depends what kind of samples you are using and what is your aim. By doing two step PCR , you are reducing the complexity of your library. The two step PCR will increase PCR-based bias amplification. In my opinion, you should do one step PCR amplification of 20-24 cycle and then purify.
I agree with Mr. Singh, but I would recommend only 10 cycles PCR instead 20-24. Again, as he said, it always depends on your aim. The Illumina protocols are quite detailed in terms of available kits and your needs.
A two-step protocol with dual indexing would reduce the costs of ordering primers, specially if you work with several amplicons.
Could you optimize the protocol Emma?
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