I am using a 2-step RT-PCR assay for real time quantification (absolute). The primers are optimized and produce a 100bp product. I know the primers are optimized since I consistently only observe a single peak when doing the melt curve analysis. The assay is SYBR green based. I have generated RNA standards by in vitro transcription and have diluted them from 10^10 down to 10^1 in 10-fold serial dilutions. I used these RNA samples for cDNA synthesis (NEB Protoscript II) and then qPCR.
I have consistently been observing PCR efficiencies of 145-155%! I don't think I should use this data, but everything else looks great. The R^2 value is always around 0.99 and my first Ct is reached around cycle 20 (10^7 copies). It also seems to be very sensitive (10^2 copies detected around cycle 34). I only observe about 2.5 cycles between my Ct values (it should be 3.3) for 10-fold dilutions.
What are some explanations for such high PCR efficiency values? I have diluted my cDNA samples, thinking it could inhibit, but the slope remains the same with diluted samples. I find it highly unlikely that I have inhibitors present from the in vitro transcription. These samples have been diluted to nearly 10^10 and I cant imagine EtOH or any other inhibitor would be effective at that dilution.
I have attached some figures (amplification, melt curve, standard curve).