Defining standards in qPCR softwares - (Dec/06/2013 )
I have a question regarding standard curves and the way to enter their values into the software.
Now, lets suppose that you have a series of standards with concentrations that is given in copies/ul. When you setup the plate layout, do you enter the standard concentration as it is or you enter the absolute quantity you're introducing in the reaction, for example if the standard concentration is 1E3 copies/ul, and you adding 5ul of that standard into the reaction, do you enter that value as it is (1000), or you calculate the absolute quantity you which is 5000 copies.
In the above example, I am assuming the software does not give you the option to choose the standard unit. On another note, if the software gives you the chance to chose the unit, does it really matter, does the software use that for calculations? or is it just a place holder, and something to appear in the results/reports?
The main platform I am asking about is the Biorad IQ5 optical software, but any information about the whole issue will be extremely helpful.
The software doesn't actually care at all what you put there as units. It can be copies, copies, nanograms,copies/ul, ng/ul, gummy bears or even just dilution factor if you don't have absolute amounts.
You should care how you interpret them.
If you have a concentration of 1000 copies/ul and you put 5 ul into reaction of both standard and sample, then putting a value "1000" in there (as per ul) and having the unknown sample concentration extrapolated as "300" would mean the concentration of your sample with identical units = 300 copies/ul.
If you put there "5000" as absolute quantity in reaction, the result would be calculated as 1500 for the same sample, which you could then also reformulate as 300 copies/ul (given that you still have same 5 ul of both standard and sample).
Both ways are possible and mathematically correct if you use a common sense while interpreting results. Though I would personally probably rather enter the overal amount of copies (5000), because it is not dependent on the same volume of standard and sample (you can have only 2 ul of sample and in that case the 1500 value of the sotware calculation would make the sample concentration 750 copies/ul). But I can't say that the other way is bad. It's not if you don't use it incorrectly. But probably overal amount is less likely to cause confusion, if you share the results.
Thank you Trof
On another note Trof, let us assume that we have a kit with standards with 7 logs dynamic range, and a good sensitivity. Now, lets assume I ran an expirement without using the lowest 2 standards. When analysing the results, assuming a good efficiency, shouldn't most of the software calculate the unknown even if it is lower than the lowest used-standard in that expirement?
The golden rule is to have at least 3 points of standard curve (I've seen variations like seven points without replicates or - what I usually do - three points only in triplicates) and have the samples within dinamic range of the standards. I use only three because I thing every serial dilution adds up errors and mostly because I like 10-fold spans and usually don't have the original standard such concentrated, to be dilluted more than 1000 times and still amplify in the usefull range.
But if you have kit standards, then you should assume they be more accurate.
Anyway, having samples outside standard curve is not recommended. If you only want to use some of them, then just omitt some in the middle, the standards don't need to be evenly distributed. If you don't want to use the lowest standards, because they amplify badly, having a sample at that range would be probably not very accurate anyway.
But, you can still use all standards and those that don't have close replicates or other problems you can discard from analysis, that should be possible in your machine (I would guess).
Reasons why extrapolation is not so good is that actually you can have a non-linear standard curve also (if the machine can calculate it, LightCycler 480 that I have, can) and then the would be a shift.
It depends why you don't want to use them and how big probability is you will have such low concentrated sample and how important it is for you to measure it accuratelly.