I am trying to define a standard curve for the primers that I have designed for qPCR and I ordered to sythetize the correpondent oligo for test the primers in different concentrations, but I don't know how much pure oligo I should use in the dilutions and I can't find some work that explain this issue. Somebody can help me?
I usually use 2500ng for the first dilution and 0,25ng for the last one in 50ul per reaction, but I have used total RNA
It is said that linear range of quantitative PCR has 9 orders of magnitude. Therefore, theoretically qPCR can measure DNA starting from 1*10^9 to 1*10^1 (actually more than 3) copies of template per reaction; in practice linear range usually narrower. I think, 1*10^6-1*10^5 of copies per reaction is a good starting point for calibration curve. It is also recommended to do at least five 5-fold dilutions of the starting point to check linearity of qPCR; dilutions should be prepared in the presence of carrier (like Lambda DNA) to prevent sedimentation of the template at tubes walls.
You don't use your primer oligos to make the standard curve. Do you means "how many copies of my target gene should I use for my standard curve"? Most folks use a CLONED target gene because they can calculate exact copy numbers and do not use a diluted PCR product.
BioRad has some good information for folks who are new to qPCR
http://www.bio-rad.com/en-us/applications-technologies/qpcr-assay-design-optimization
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