I am running standard dilutions (1:5) of some cDNA template to check the efficiency of primers. I found to have apparently PCR inhibition, as the efficiencies that I am getting for all the targets are over 110 % (all are around 130-140 %), with R2 >0.98. However, I can't understand where I can be getting PCR inhibitors. I measured the purity ratios of the RNA after isolation, and they are 260/280=2.09 , 260/230=2.23. I also measured the purity of the cDNA after the reverse transcription , and the values are 260/280=1.77 , 260/230=2.22. Although the 260/280 ratio drops ( I guess because of the reverse transcriptase), I don't think is too dramatic. I even dilute the template for the first dilution point, to put no more than 50 ng per reaction. If I am introducing inhibitors I think it can only be during the real time qPCR. I am using the LightCycler 480 SYBR Green I Master kit from Roche, but it is brand new. I analyzed the data with the ThermoFisher Cloud, and I am getting Amp scores over 2 for all reactions, so the quality of amplification is good. I only see one peak of melting curve for each reaction. For all these reasons I don't understand how I have PCR inhibition.
Can it be an issue of the set up? I thought that working out of optimal temperatures and cycle times will decrease the efficiency below 90%, not increase it. over 110%
Does someone has any clue of what can be happening?
Edited by Lord Apoptosis, 19 March 2019 - 06:25 AM.