There is no one value that works for everyone. You usually can determine this from your standard curve set up if you have included very high template concentrations. As you keep increasing template concentration there will be a point at which a doubling of template concentration does not result in one less Ct. That concentration and corresponding Ct is your limit. In practice I have found reactions can amplify efficiently down to a Ct as low as 6, and that limit was determined by the quantitation algorithm, not the reaction per se.

There is no one value that works for everyone. You usually can determine this from your standard curve set up if you have included very high template concentrations. As you keep increasing template concentration there will be a point at which a doubling of template concentration does not result in one less Ct. That concentration and corresponding Ct is your limit. In practice I have found reactions can amplify efficiently down to a Ct as low as 6, and that limit was determined by the quantitation algorithm, not the reaction per se.

Roger is right. Run a standard curve. That will tell you the saturation level, the limit of detection and te efficiency. Remember, if you are using the delta Ct method you target gene and normaliser should have similar efficiencies as well. I have had reactions in the 12 Ct region before with no problem.

Hi, thanks for your answers that's a lot clearer now.

Just for a bit of follow up. The reason I ask is because I'm using Roche designed PCR assays and I don't have access to the standard curve calibration experiments.

I guess the reason that people generally say 12-15 is because it's a safe bet that most assays are well within the quantifiable range of the assay at these concentrations.

In that case I suggest you do your own standard curves - and make sure it spans a very large concentration range so that it covers very high and very low Cts. The primer/probe sets worked for Roche with an efficiency of 100% - but you cannot know they are working the same in your set up. Roche implies the same in the last sentence of this document: http://www.roche-applied-science.com/wcsstore/RASCatalogAssetStore/Articles/RTR_04.11.pdf

I have tried some published primer sets and found they do not work as described. Assumption is the mother of ...

Always design your own primers. If you run and analyse qPCR experiments you should have the know how to design primers (in 5 minutes) as good if not better than the commercially available pre-designed primers. You will also get 1000 times more for about 1/10 the price. Don't be fooled by sales info from the likes of Qiagen saying their primers are validated. What they mean is they are bio-informatically validated which essentially means running them through BLAST, which you should do anyway.

Really you should always run your own standard curves to at least calculate your reaction efficiency. It is easy.

Hi Mark, Hi Roger,

Thanks for your feedback once again.

I would normally agree with you Mark on designing your own primers because of the advantages you list. However, in this case I am using the Roche PCR arrays in which the primers and probes are dried directly into 384 well plates. Therefore I do not have the ability to run the assay outside of this format and I do not know what the optimised primer concentration used in these assays is.

I am using these arrays in conjunction with a cDNA pre-amp step as the samples are archival clinical samples.

Because the assays are in this format I cannot do the standard curve experiment which would solve this issue of what dilution factor of my pre-amped cDNA to use.

I got around the issue by ordering in the primers used in the assay and running a number of dilutions of a pre-amp cDNA library on a standard SYBR green QPCR plate. Since I don't have the exact sensitivity range information for the assays I'm going to have to assume that a CT of ~15-17 for my highest expressing genes on the array will suffice. *Fingers crossed