How To Calculate PCR Primer Efficiencies

Why bother with PCR primer efficiencies?

Every time you receive a new set of primers, especially when using SYBR Green chemistry during quantitative polymerase chain reaction (qPCR), you should always run a standard curve to calculate the efficiency of your PCR primers.

The reason we bother calculating PCR primer efficiencies is to be able to correctly analyse the results. For the calculation of gene expression, such as the delta-delta Ct method, it is assumed that the PCR primer efficiencies are comparable for the gene of interest and for the housekeeping gene. Therefore, dissimilar PCR primer efficiencies within your experiment can impact your final result.

Mastering qPCR

A video tutorial on qPCR primer efficiency calculation can be found in our Mastering qPCR course.
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What is the correct PCR primer efficiency value?

Obviously, a perfect primer set will have a primer efficiency of 100%. In other words, for every PCR cycle, the number of copies of the PCR product will double in size during the logarithmic phase of the PCR reaction.

To get a 100% primer efficiency for all of your primer sets is highly unlikely. Therefore, it is recommended that all the primer sets used in your experiment lie between 90 – 110% efficient. If so, they are deemed comparable.

How to perform a standard curve

So you have designed and received your new primers. Now what? Well, the first thing you will need is a template to use for standard curve generation. Ideally, this should be from the same source as what will be used during your experiment. So, if you have generated complementary DNA (cDNA) from RNA extracted from a cell culture experiment, for example, then use one of these samples as your template.

To create a standard curve, it is recommended to start with the undiluted cDNA sample as your first point. From this, you need to create a serial dilution series. A 1:10 dilution is commonly used to create a standard curve with at least 5-points. If you can include more points in your standard curve, then this would be better. So long as the standard curve covers the Ct values of your experimental samples then this is fine.

Here is an example of a 1:10 serial dilution standard curve containing 5 points:

Perform a qPCR reaction using your standard curve containing the recommended reagents and concentrations for the qPCR master mix of your choice, as a starting point. Make sure to perform each sample in duplicate at the very least, or even better, triplicate. Also, don’t forget to include no template controls (NTCs), i.e. PCR-grade water instead of the sample, on your plate to identify any contamination.

How to calculate primer efficiencies

Some qPCR machines will be able to calculate this for you, but I prefer to export my raw results and calculate the PCR primer efficiencies manually.

Here is how to calculate a primer efficiency using Microsoft Excel. The Excel formula used in each section is highlighted in grey.

1. Calculate your average Ct values from each of your replicates/triplicates

The first step is to average the technical replicate Ct values.

The function in Excel is found below, where the Ct1 and Ct2 values represent the cells for each technical replicate.

=AVERAGE(Ct1,Ct2)

2. Calculate the log of each sample dilution

The starting quantity is based on your dilutions. So, for example, I like to call the first value ‘1‘ since this is the stock, undiluted cDNA. Then do 1:10 dilutions of this value.

Then, the log value of these should be determined. Simply use the LOG function in Excel to do this.

=LOG(Starting quantity)

3. Get the slope of the regression between the log values and the average Ct values

To quickly calculate the slope of the line, use the SLOPE function in Excel. Specifically, it is the slope between the log values just created and the average Ct values.

=SLOPE(Average Ct value range, log quantity range)

Alternatively, you can plot the log values against the average Ct values as a scatter plot. To do this, first, create a scatter plot between the average Ct values and the log values. Then select ‘Add Chart Element > Trendline > More Trendline Options …’. From here, select the ‘Display Equation on chart’ option to view the regression equation. The slope value will be the value at the start (just before the x). In this example, the slope is -3.359.

4. Calculate the primer efficiency by using the slope value

Primer efficiency values are presented as a percentage. To calculate primer efficiency values, use the following equation.

Primer efficiency PCR formulaThe formula to do this in Excel can be found below.

=(10^(-1/The Slope Value)-1)*100

This will give you a primer efficiency score as a percentage. Hopefully, this is between 90 – 110%. By using the above dataset, the efficiency comes to 98%.

If your standard curve and primer efficiency is not within the desired range, don’t worry. There are a few things you can do to improve your PCR primers efficiencies, such as adjusting the primer concentrations and the annealing temperature of your reaction. If you are really struggling after that, then I suggest designing new primers.

PCR primer efficiency calculator

If you prefer, I have created a PCR primer efficiency online calculator. To use this, simply enter the slope of the line, as determined above, and the calculator will return the primer efficiency value and the amplification factor (E).

Free PCR primer efficiency Microsoft Excel template

Are you still struggling to calculate your PCR primer efficiencies? Well, I have made an Excel worksheet to hopefully help you out.

All you have to do is to fill in the Ct values from your replicates and the dilution factor used when making the standard curve, e.g. 10, and the sheet will (hopefully) work out the rest for you. This will also work out the slope, R2 value and the PCR primer efficiency value as a percentage.

Click here to download the template.

17 COMMENTS

  1. Hello, Dr. Steven!

    This is such a great site and the excel template you provided is really useful. One of my primers have an efficiency of 114%, and since you mentioned the ideal range to be 90 – 110%, I was wondering if you would consider 114% as acceptable or whether I should start thinking about the effect of contaminants/inhibitors in my run.

    Also, I want to know how you would go about troubleshooting low primer efficiencies (60 – 75%), will you change the primer concentration first or the annealing temperature? Both AGE and melting curves suggest single products in my intended product size, so I’m leaning more towards optimizing the conditions rather than redesigning primers.

    Thank you again for your help!

    • Hi Marvin,
      Many thanks for your message and I am sorry this is late getting back to you.
      So in your case, I would say 114% is acceptable. I would, however, use the Pfaffl method to account for this difference to your other primer sets.
      With regards to primers that have low efficiencies, this can be due to suboptimal primer design. So you may want to look at your primer sequences and possible redesign them. Or, incorrect primer conditions. This could be due to the wrong annealing temperature being used. So ideally you want to run a gradient PCR to determine the optimal annealing temperature. Also, yes you are correct, changing the primer concentrations can also help to imprive the efficiency of the primers.
      Best wishes,
      Steven

  2. So thankful to have found this site. I’m learning qPCR on my own here having only had experience with good ol’ PCR many years ago and your write ups are easy enough to understand. Thanks Steven.

    • Many thanks for the comment JC, I really appreciate it.
      Good luck with your qPCR and feel free to reach out if you get stuck.
      Thanks,
      Steven

  3. Hi Steven,

    I found primer efficiencies vary due to SybGreen dye on the qPCR. If I use UPL way, and it is probe, do I still need to calculate the primer efficiencies?

    Best,
    Lucas

    • Hi Lucas,
      So if the probe-based assay was created by a company, usually they would optimise the primers for you. So there is generally no need to do a primer efficiency calculation. Personally, I would do one anyway just for your record.
      Good luck!
      Steven

    • Hi Ann,
      Good question! There are a few reasons why this happens. Mainly:
      1. There may be primer-dimer formation. Check there are no primer dimers by running some product on an agarose gel or by checking the melt curve (if using intercalating dyes).
      2. There could be PCR inhibitors in your reaction, such as ethanol. Ensure your starting material is free from potential PCR inhibitors. You can also dilute your starting material, which will dilute the inhibitors in the reaction.
      3. There could be too much template added to the reaction, especially at the start of your standard curve. Try diluting the starting template further.
      I hope that helps.
      Best wishes,
      Steven

  4. Hi Dr.Bradburn,
    first of all thank you so much for your excellent website. actually, I think the slope formula which has been shown in grey is wrong. the right one is it:
    =slope(average Ct value range, Log starting quantity range).
    best wishes.

    • Hi Mohsen,
      Many thanks for your comment.
      You are right! Apologies, I seem to have placed the x and y values for the SLOPE function the wrong way around. I have since amended this.
      Thanks
      Steven

  5. Hi Steven :
    Many thanks for the useful information. We use PCR in determination of presence/ absence of foodborne microorganisms by a kits validated by AOAC like ( foodproof® Listeria monocytogenes ,Detection LyoKit-5”Nuclease-LP. Protocol , foodproof® E.coli 0157 Detection Kit -5”Nuclease – Protocol, foodproof ®Campylobacter Quantification Kit-5”Nuclease- Protocol). Are we still needs to determine the amplification efficiency by method mentioned above? taking into account the fixed volume of sample and mix determined by kits .

    • Hi Safa,
      Many thanks for your message. No, you should not need to perform the primer efficiencies for this assay. Mainly, this is a detection kit rather than gene expression analysis. Further, the company will have optimised the assay to work. Therefore, I would not worry about primer efficiencies for this assay.
      Best wishes,
      Steven

  6. Thanks alot for that information, it is very helpful and useful. But I have a question. If my target tissue is liver and I have liver with cancer. Can I use healthy liver tissue as a standrad?

    • Hi Soha,
      Many thanks for your message. Yes, you can use a healthy liver tissue as your standard in your case. Or, you could also pool a few samples together if you don’t want to waste all of one sample.
      The main point is that the Ct values of your standards should cover the range of Ct values in your experimental samples. So if your highest standard Ct value is 16 and your lowest standard Ct value is 33, then your experimental sample Ct values need to be within Ct values of 16 – 33.
      Hope that helps.
      Best wishes,
      Steven

  7. Thanks for your useful information. Can we make a 1:2 dilution series for primer efficiency calculation? sometimes the PCR cannot be accomplished in a wide range of dilutions (1:10)

    • Hi Mehdi,
      Many thanks for your comment.
      Of course. Sometimes certain genes are expressed at low levels so a smaller dilution is required. Try to do at least 5 points in your dilution series. Also, ensure that when it comes to do your experiment, your Ct values are within the range of Ct values from your dilution series when you calculated the primer efficiencies.
      Best of luck,
      Steven

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