I have had some preliminary evidences showing different q-PCR efficiencies (HKG and target genes) for different cDNA samples, using the same pair of primers. I reckon that, theoretically, efficiency of the HKG and target genes must be constant in every sample, but in my preliminary results, E encompasses from 85 to 95%. I'd like to know which could be the reason of the differences. RNA samples come from woody plant leaves. Thank you!
The sample can certainly affect the apparent amplification efficiency of your PCR reactions, as they may contain inhibitors that reduce the efficiency. This is most usually apparent as an increased Cq and/or an abnormal slope of your amplification plot.
At the same time we know that RT efficiencies vary widely, dependent on the supplier, but also on the sample and target being investigated. You also say you are working with plans material, so there is a well known background of co-extraction of RT-qPCR inhibitors. I would advise you, if possible, to see whether you can use dilutions of your RNA and so avoid any inhibition or, if not, run dilution curves with your samples and use each dilution curve to calculate target copy numbers.
I think that as far as the HKG and the gene of interest have the same efficiency in each sample you don´t need to worry. You are using the HKG to normalize for differences between the different samples. However, it is important that your RNA has good 260/280 and 260/230 ratios
Dear Rafael,
A difference from 85 to 95 % is rather small and therefore neglectable IMO. What you see could be a "natural" pipetting error or the normal degradation process of your cDNA rather than a real inhibitor.
Keep in mind that cDNA degrades over time even when stored at -80°C. If you planned wisely you won' t freeze and thaw your samples but use a fresh aliqout for each qPCR. Using the same aliquot again for several PCRs and thawing it more than three times can definitely lower your PCR efficiency.
I experienced that when I did semi-quantitative RT-PCR - samples that had been reused showed a weaker signal on the gel in each following PCR. Using a new aliquot for each PCR eliminated the problem.
As mentioned above of course there still could be a contamination in your samples. It is very likely that you have co-isolated non-plant RNA as well and that this alien RNA/cDNA interferes with your qPCR. This problem is well known from samples out of soil like roots - a tissue I am working on. Here, humins can cause trouble in the PCR. Of course secondary plant metabolites can be diadvantageous as well but I would start to think int that direction only if your RNA or efficiencies are really bad or you cannot detect any transcription at all - which is not the case for you.
There definitely are situations where PCR inhibitors are present in some cDNA samples, and not others. I've observed this consistently across technical and biological replicates for a treatment that induces a range of phenolic compounds. When phenolics are induced, the PCR efficiency of individual reactions is reduced (determined based on amplification plots). The steepness of the amplification plots were very clearly different, visible by eye. You can account for these differences using efficiency-corrected calculations (e.g. eGOI^(-Ct GOI)/eHK^(-Ct HK).
Typical standard curve estimates of PCR efficiency do not account for this source of variation - and often are performed on perfectly clean, diluted plasmids. At best, they provide the reaction efficiency for a primer pair under optimal conditions. Many qPCR experts advocate the use of estimating PCR efficiency from amplification plots - e.g. using LinReg. I'd highly recommend reading the commentary '11 golden rules for qRT-PCR' published in the Plant Cell is a great resource.
Hope this helps!
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