Hello
I've performed qPCR experiments for many years but I could not really figure out the following.
First question:
I've prepared standard curves from a DNA template (concentration determined by Nanodrop several times) and then run the qPCR. So I get a set of Ct values for this standard curve.
In a subsequent attempt, I prepare a new set of standard curves from the same DNA template (from the same tube) and run the qPCR. I get a different set of Ct values sometimes differing by Ct value of 2 from the first attempt for the highest concentration. In both cases I've obtained R^2 values of >0.99 and efficiency 90-110%.
So my question is should I expect the Ct values to be the same every time I run the qPCR assuming the DNA template, primers are always the same?
Second question:
If I use the same primers for different DNA templates but they all contain the same region that the primers target, should I prepare a different standard curve per different unknown or can I just use this standard curve across different unknowns as long as the DNA templates contain the same target region?
E.g. mCherry-WPRE and EGFP-WPRE, both contains WPRE and my primers are designed to target the WPRE region. Should I prepare standards based on both or only one is needed to determine unknowns for both? Their total size and sequence are not the same but the WPRE sequence is the same.
Hi,
I will try to answer your questions.
1. In the ideal case the Ct values from the same DNA should be "identical" (SD approx. 0.2 Ct but depending on the target amount). So you may think about possible errors in one of the experiments. Thawed DNA not mixed? The same threshold in both experiments? Any problem with the optical system of the cycler?
2. I think you can use one of the plasmids as standard for all samples. But there may be special cases were you have to think in detail about the use.
Best
Norman
I respectfully disagree with the answer from Norman Hafner .
He's correct saying that Ct values should be within an SD of about 0.2 - but this refers to technical replicated within the same plate/run. You were asking for Ct variation of the same sample/standard between different plates/runs. Here it may well be that you observe differences of 1-2 cycles (perhaps even more), as it depends on many things such as on how the software interprets the background signal, the measured signal noise in the first cycles and how the threshols is chosen. Most of it is not under your control. So you should never expect that Ct values from differentplates/runs are comparable.
When you run a new late and you want the Ct values being comparable with that of other plates/runs, you need to include a plate-standard. This can be one, two,... or all of the standard dilutions you have prepared. You can use this to either simply correct the Ct values (by subtracting the observed difference between the Ct values for the standard on the different plates) or by expressing the sample concentration in units of the standard (e.g. in copies/µl), if this is known for the standard you are using.
To your 2nd qustion:
you can/should use the same primers, if you want to compare the concentrations of mCherry-WPRE and EGFP-WPRE.
This is likely not relevant for you, but if the DNA tempates are structually different (e.g. onesample is bacterial DNA, the other is plant DNA, and yet another is a plasmid - all containing the target sequence), the structual context can cause considerable differences in the amplification efficiencyin the first few cycles,where the sequence context of the target sequence plays an important role (e.g. adjacent GC-clamps in gDNA can hinder efficient melting and amplification, where as a plasmid maynot contain such GC-clams; this would lead to systematically higher Ct values for gDNA). In these cases it's a good idea to fragment high-molecular templates. It's also good to nick or linearize plasmids.
...Jochen Wilhelm is correct. The interassay reproducibility is lower than the intraassay reproducibility and you cannot influence it completely. However, in our experience, using identical experimental conditions we have interassay SD of 0.1-0.6 Ct for a large range of copy numbers and different amplicons. This may be higher for the normal real life ;-)
To be sure that your assay is valid in different experiments you may use a calibrator sample as suggested by Jochen Wilhelm. However, I would use an additional sample which can be quantified by the standard curve. If your standard is ok, than the calibrator sample should give the same results in different runs if quantified by the standard curve.
@jochen wilhelm Thank you so much. Now I understand it better.
For the second question;
I wanted to measure the concentration of both but not really to compare each other. Should I prepare two different standards for both plasmids and determine the unknowns for each one of them accordingly? Or is it sufficient to prepare only one standard curve for all? Because 99% of the time I performed qPCR the different plasmids I have contain the same region WPRE and I've used the same primers for all of them. These are used to quantify the viral titer of each viral construct and I've been making separate standard curves for each viral construct. Even if the primers target the same region in all of them the Ct values do differ from 1 plasmid to another.
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