It can also occur from inaccurate standards (e.g. old, improperly diluted, etc).

Do you get much more than 100%?

Also, check in your melting curve if you have any kind of unspecific amplification.

I can get 123%. And I already checked the melting curve, everything is fine.

So what should I do? Consider 100% efficiency? change the primers? Do an efficiency curve with lower cDNA concentration?

I don't think the issue is with your primers, as they seem to be producing specific amplification. I think the issue is with your reaction set-up or standards dilution series. Johan is probably correct that a reduction in the amount of DNA used for your standard curve will fix this issue. If you've already done a relatively wide dilution series in the first instance, you can check the slope of your standard curve at the most diluted standards. If the slope at these points is closer to 100% (ignoring the least diluted standards) than you will have your answer. Good luck!

Johan and Christopher are right. Efficiency over 100% occurs when you have inhibitors in your standards. To diagnose this, check the slopes of your amplification curves to see if they look "slept" and if there are differences in the shape and in the number of cycles between the more concentrated and the more diluted standards. Try diluting your cDNA without loosing a log point, or doing instead a PCR product curve. Good luck!

Dear friend,

It is quite common in case of real-time PCR. And, most of the researchers face this problem during the start of the reaction. There are numbers of reason for this. For example, one is the standard curve (improper dilution, or pipetting error) -- usually, this happens because of your most diluted standard dilution. Try to see the efficiency and R square by selecting the first few dilution by omitting the later dilution. See whether or not that works.

I don't know which PCR machine you are working with. Normally, every PCR machine has their own acceptable efficiency range. So, make sure to check the specification of the machine you are using. You can find that information in ther user manual or in the official site of the company of manufactor. If I am not wrong, for example, Stratagene MxPro 3005 have efficiency range of 85% to 115%. Whatever, efficiency you get within this range will be acceptable. However, still, the efficiency close to 100% if of course most acceptable and kind of ideal.

Sometimes it might be because of the contamination in your sample, like phenol, ethanol or some chemical that might not have completely churned out during your purification step or something that might have still stick to your sample during the DNA or mRNA extraction process.

Sometimes even higher concentration of primer might inhibit the reaction. Try to follow some paper related to your work, and try to check the ratio of sample to primer they have used. That might help, I guess in few instances.

Wish you all the best.

PS. Usually machine itself calculates the efficiency and R square from the equation it generates from the standard curve.

I also suggest to use the method described by Ramakers et al (2003) to estimate the PCR efficiency of single wells based on the linear regression of Log(fluorescence) versus cycle number. This method is alternative to standard curve of serial dilutions for calculation of PCR efficiency. Even if you apply the standard curve method, the analysis of single runs can help you to spot the problem.

Bye

Isacco

link to the article

http://www.gene-quantification.biz/ramakers-2003.pdf

LinReg Software can do this calculation from raw fluorescent data.

link to the software

http://www.hartfaalcentrum.nl/index.php?main=files&fileName=LinRegPCR.zip&description=LinRegPCR:%20qPCR%20data%20analysis&sub=LinRegPCR

Hey

Just try the following, I am pretty sure that you will get the efficiency close to 100%

1. use BSA @ 0.3 uM in each reaction

2. Increase the volume of water when you make the dilutions;

1+19 or 1+39 in place of 1+9 and add more volume of dilutions in reaction like in case of 1+19 add 2 ul of it and in case of 1+39 add 4 ul of it. I always do this to get rid of the inhibitors

3. use low retention tips when you make the dilutions.

Let me know if you need a detail protocol for dilutions

Dear Virginie

I suspect that you have constant forming and dissociating primer dimmers and because you have more primer dimer molecules then your specific DNA, it results in higher efficiency. You can check that in melting curve analysis graph representing – (d/dt)fluorescence (483-533) as fluorescence fluctuation or 0.3 higher is than initial fluorescence from 60 °C. You can also try to measure fluorescence not at 72 C but instead at higher temp ( primer dimers dissociate normally around 80 C if the primers are not GC rich). So it can be safe to measure fluorescence 5 degrees lower than the melting temp of specific product

Good Luck

I would suspect that you either have primer dimers or as mentioned above any by-product from RNA purification. Since I suppose that you have checked your primers for non-specific amplification on the melt curve and on an agaroze gel, I would suggest you reduced the cDNA amount you use for the standard curve or the primer concentration.

Efficiency should vary between 95-115% but even so it is relative; what you really need to make sure is that the amplification efficiency of your GOI and reference gene will not differ more than 5%. Otherwise, the rate will be different and you will not be able to deduce safe conclusions.

Furthermore, it is also important what your fluorescence threshold will be. Although the machine usually calculates this value, it is also possible to interefere manually and set an arbitrary threshold so that you in the end get a good amplification efficiency. By experience, this should lie around 2/3 of the maximum of fluorescence signal, as more than that would mean that the cycle values are higher and so many undesired by-products (e.g. primer dimers) will be enough to increase amplification efficiency. The opposite applies to less than 2/3, where no more specific product is already produced and thus you would have an efficiency

Ideally the efficiency should be close to 100%, meaning that your PCR product doubles during each cycle. Efficiencies below 100% normally mean poor reaction cconditions (bad primers, bad chemistry …). When you see efficiencies above 105-110% it may be due to amplification of nonspecific products, primer-dimers, or also the presence of inhibitors carried from your RT reaction. Did you try to analyze your dilution series omitting early and late points? Does the efficiency change after you do that? How may points do you have in your dilution series and how early does amplification starts (at what cycle?). For the first point of the dilution series, do you use diluted cDNA or cDNA straight from the RT reaction?

I was asking the same question but here what a rich topic and answers already available! tnx everybody

It is probably common problem for SYBR based qPCR as you are measuring total fluorescence. Presence of unspecific products of all kinds could build up florescence making it looks like >100% efficiency which is theoretically impossible. If the efficiency is < 110% then I would not worry and just proceed with deltatdelta Ct method. Off course as mentioned already above it is important to have efficiencyyou’re your gene of interest and reference gene as close as possible. 90% for one and 110% for another it is too big of a difference from my point of view. Check you melting curve again and play around with primer concentration and annealing temperature – it could help sometimes. Or just order completely different primers for another region of the same gene.

Simpler way to solve the problem – switch to TaqMan probes from Life technologies or IDT. We always use them and never seen a problem.

Hi Mohammad,

I do have the same problem would you please send the detail protocol for dilutions?

Thanks

Sometimes when you work with very low concentrations of cDNA (in your serial dilution) there can be some pippeting problems that can increase your error and give you an efficiency greater than 100%. It's not the same pippeting 100 more copies of cDNA in a mixture that has 1 million of copies of cDNA (that would be a 0.01% of error) that pippeting 1 more copy of cDNA in a mixture of 100 copies of cDNA (that would be a 1% of error). Therefore, in this type of experiments if you work with a very low concentration of cDNA, the efficiency can be usually greater than 100%.

One additional suggestion that hasn't been brought up here: check your pipette. Try using a different pipette and see if the result is the same. Since low volumes are sensitive, iIf you add an extra 5-10% cDNA to each well, your true dilution factor could be quite different from your theoretical dilution series. You might just need to re-calibrate.

I had this problem recently with a set of primers where my efficiency was 160%, but all my sanity checks were good. Might have been a primer hairpin that was supposed to melt at 14'C below the temp where fluorescence was taken. So a second reminder is that primer Tm is only the median where ~50% of dimers are dissociated. Some fraction of primer dimers might remain stable at temperatures much higher than you'd think.

Thanks to Behrouz for the recommendation of measuring fluorescence at a higher Tm to try and solve this issue!