I have plasmid containing my gene of interest. It has been linearized, gel purified, and is at known concentration.
I will use this standard to generate a standard curve for my qPCR runs, and test them against my mammalian cell culture cDNA samples. (FYI: My cell samples are 1)WT Control 2)GFP Control 3) GOI+GFP tag.)
But I'm wondering if the DNA standard is valid to use to calculate qPCR efficiency.
The DNA standard is a cleaner product. My cDNA samples were extracted from a mammalian cell line, gone through RNA extraction, post-DNAse treatment (AMPD1), and reverse transcription. All of these procedures could have included in the mix compounds that may reduce efficiency.
I can see how the DNA standards can be used to calculate copy number, but should I be using a cell sample control to calculate efficiency? Should efficiency be calculated for each of the cell samples anyway, or just a control?
One would hope that your final cDNAs are not so dirty that they impair PCR reactions (this would render your assay semi-quantitative, essentially), but if you are worried, why not try the following: simply mix your plasmid standards with an extract that has been treated identically to all your other samples, with the simple omission of reverse transcriptase.
You could always do a final cleanup step on your cDNA, but I've never had a problem with "dirty" cDNA. It sounds like you've made your plasmid standard correctly. Best of luck!
Dear Martin
I cannot add much to the comments above except to say
- It is valid to use plasmids to verify primer efficincy but more accurate to use real samples or a simulacrum of a real sample as suggested by John
- People only tend to use high copy number fragments to measure primer efficincy when they have no alternative, i.e. the gene of interst is of such low copy number that the undiluted cDNA yields Ct values > 30 and thus a titration is not possible
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