Hi all,

We have been trying to figure out why we see such differences in our base pair (bp) fragment size between doing a 1-step PCR reaction vs a 2-step PCR reaction to prepare metabarcoding libraries for Illumina HTS. The PCR recipe is the exact same. The PCR reaction conditions are the almost the exact same between the 1-step and 2-step, except 1-step is 25 cycles and 2-step is 25 cycles for amplicon (1st PCR) and 7 cycles for index PCR (adding the barcodes, 2nd PCR). The primers are much larger in the 1-step PCR with everything in the primers needed for sequencing. Annealing temperature is 62C and based on the primers melting temperature (515F, 939R) without any of the overhangs and the use of Kapa HiFi hotstart ready mix recommendation to be near primer melting temperature for annealing.

The big change between the two is a double edged peak in 1-step versus 2-step PCR. 1-step has a double peak - one at 490 and one at 550. 2-step has a standard peak at 570 bp that is what we anticipate.

In both, we also have a tail of additional products that we have been trying to figure out what they are for years. The guess is heteroduplexes or daisy chains. If anyone has any insight would love any thoughts on that also. We size select (using an e-Gel) to get rid of them in the 2-step library prep. We have had issues with size selecting with the 1-step because of the larger smear from 400 to 700.

We are having challenges with consistent runs on the MiSeq since we transitioned to 1-step library prep, and trying to sort out the 400-700 bp smear - why double peaks? - and planning to try to figure out how best to size select again.

Any insight on why double peaks happen in PCR, why there are larger fragments present after PCR and how best to size select when you have a larger smear would be greatly appreciated on any of those items.

Thank you.