I have been designing qPCR primers for use in assays with Promega GoTaq qPCR mastermix. This uses BRYT Green, which is supposed to be comparable to SYBR chemistry. Using a non-precious positive control cDNA sample to optimise primer concentration and temperature initially, I select conditions that give single melt curve peaks/gel electrophoresis bands, blank NTCs and high efficiency.  These are inducible genes with quite late Cq values. However, when I run the gene assay on my experimental samples, I get non-specific bumps/peaks in the melt curves, sometimes even in addition to the gene-specific peak in a given sample. I guess primers can act in unexpected ways in the presence of DNA with low amounts of template, that can’t be predicted from template-abundant positive control and cDNA-free reactions. I always run my RNA samples on an agarose gel so I know my experimental samples aren’t degraded.

Would anyone have any suggestions for how to tackle this? I am aware that TaqMan is probably more appropriate for my situation but we don’t have the money or time in the lab to switch at this point. I try to keep primer concentrations to 200-300nM, as I’m afraid going lower would mean I wouldn’t be able to detect anything in my experimental samples at all. I aim to reverse transcribe 1ug of RNA per cDNA synthesis reaction, though for some sets of experiments I have had to reduce this to 300ng. I dilute this 1:10 and use 2ul cDNA in a 20ul qPCR reaction volume- perhaps I should be adding more cDNA?

Any suggestions or comments on people’s own experiences welcome. Thanks!

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