consistent real-time PCR

10 Tips for Consistent Real-Time PCR

Real-time PCR is a specialized technique that delivers far more information about your DNA or RNA than end-point PCR.

Essentially, real-time PCR is a way to visualize the amplification of specific DNA fragments as it is happening (in real time) and allows for the ability to quantify exactly how much DNA (or RNA) was in the original sample.

Fluorescent dyes (either SYBR Green or dye labeled DNA oligos) are mixed in the amplification reaction, fluoresce when they bind to DNA, and are measured by the machine during replication.

The amount of fluorescent signal measured directly correlates to the amount of product in the tube at that moment in time. Because of the sensitivity of fluorescent dyes, concentrations as low as picogram quantities of DNA (or as low as a single cell) can be accurately detected.

While it has become second nature in some labs, the technique does require a certain amount of technical finesse to get consistent real-time PCR data every time. Because the cost of real-time PCR kits is so much higher than standard PCR, getting every experiment right is critical.

The main bottlenecks people encounter when getting started with real-time PCR are contamination issues or inconsistency between replicates. Here are some simple but important pointers to help make sure you prevent the cause of many grey hairs and frustration at lab meetings.

1. Always Mix the Reagents Well Before Use

The reagents contain dyes, nucleotides, and enzymes that may have settled while sitting in the freezer or refrigerator. Give the master mix a good mix before your start aliquoting into your plate or tubes to avoid uneven distribution of reagents between samples.

2. Store Primers in a Buffer to Protect their Stability

When your primers arrive, avoid resuspending the master stock in water. The pH of water can be low (especially if it is DEPC treated) and this will be damaging to DNA over time.

Use a buffered solution at neutral pH to protect from acid hydrolysis. EDTA (1 mM) in the master stock is also a good idea to protect against DNases and when you dilute the primers for working stocks, the EDTA will be sufficiently dilute that it will not interfere with Taq activity.

3. Aliquot the Primers

Once you have a master stock (usually 100-200 mM), you will want to make some working stocks so that you are not continually freeze/thawing your primary source. Prepare 10-20 mM working stocks in neutral pH buffer and prepare aliquots that allow you to freeze/thaw the working stock 3-5 times at the most.

Continual freeze/thawing of the primers can cause some break down and this will lead to drift in results such as worsening PCR efficiency and sensitivity. Preparing aliquots will also help avoid contamination problems.

If you accidentally contaminate one of the tubes of primer, you can throw it away and take a fresh one without worrying about contaminating the main stock.

4. Use Pipettors Accurately for Low Volumes

If you require absolute accuracy in quantification and want spot on standard curves, use a pipettor calibrated for low volume pipetting (such as a P2 or P10).

This will ensure reproducibility between replicates and make sure that when you are measuring efficiency based on the standard curve, that you are truly measuring efficiency of the reaction and not your pipetting skill. Read more to learn more about pipetting accuracy.

5. Perform a Standard Curve for Every New Primer Pair

Don’t assume that every set of primers ordered is going to work as well as the last. PCR efficiency can be impacted by a number of factors. The best practice is to run a 5 point standard curve with 10 fold dilutions for every new primer pair and make sure you can get at least 90% PCR efficiency with control DNA.

7. Follow the Three Room Rule

One of the biggest causes of contamination is from using the same pipettors for extraction or handling PCR products post-run for reaction set up. Even if aerosol resistant tips are used all the time, this is a big no-no. Buy a complete set of pipettors that are used for PCR set up and nothing else.

In addition to new pipettors, you will want to keep them in a different location, and preferably a different room than the room used for extractions. The ideal set up is to have three rooms; one for RNA or DNA extractions, one for reaction set up (and using a hood with a UV lamp to pre-treat the pipettors and plastics between users), and one for the real-time cycler.

This is the most assured way to make sure you never have amplification in your negative controls.

8. Double Check the Cycling Conditions

This is important if you are using a shared instrument. Even if you have your own template file set up, before hitting start, make sure the machine has the correct run cycle for your experiment. Someone may have used your template and made changes to the annealing temperature or the hot start activation time without your knowledge.

Some instruments default back to standard settings and if you are using an instrument for the first time, you may find that your settings didn’t save. It never hurts to double check the run settings.

9. Dilute the Template (Less is More)

Depending on the gene of interest, you might actually be starting with too much template. Real-time PCR is sensitive enough that sometimes less template gives a more accurate measurement.

You will want samples to cross the threshold between cycles 20-30. Samples that cross the threshold below cycle 15 will fall into most instruments default baseline setting and this will cause a subtraction of fluorescence from the rest of the data.

This can be remedied by adjusting the baseline setting, but if you are unfamiliar with your instrument, it may require a call to technical service to figure it out. Also, if there were any inhibitors in the sample from the purification step (guanidine salts or ethanol, for example) diluting the sample will eliminate their impact on the results and give you an accurate quantitation of the sample.

The best approach for a new sample is to perform a standard curve- even just a 3 point dilution series- to see what concentration will give you a Ct that falls in your standard curve and is most accurate.

10. Make Dilutions Fresh

Nucleic acids stick to plastic so if you make a dilution series and want to store it for future runs, you will need to protect the samples from absorbing to the tubes walls and becoming diluted out over time.

This can be done by using a carrier nucleic acid, such as tRNA, or by using specially treated plasticware that does not bind nucleic acids. Several manufacturers offer low retention tubes (Axygen is one) or silicon treated tubes to help prevent this occurrence.

If you do store dilutions in non-treated tubes, you may want to re-quantify the most concentrated dilutions on a Nanodrop before using to make sure they still match the expected concentration.

Some of these tips may seem like common sense- and they are- to people who have been doing this for a long time. But for many people just starting to use this technology, a lot of time and money can be saved with these simple steps that can make a big impact on results.

There are a lot of resources for real-time PCR help as well, including a very active Yahoo List group and the BioTechniques® Molecular Biology Forums. Fortunately, there are many experts who enjoy helping others master the art of real-time PCR.

And if any experts out there reading this want to list some common mistakes you see in your labs or best practices tips, please let us know in the comments field.

Originally published on January 20, 2009.  Revised and updated April 4, 2016.


  1. Catherine on November 8, 2016 at 4:42 pm

    What is an appropriate concentration of tRNA. 1ng/ml or 10ng/ml as a diluent? I have conflicting notes.

    Thank You.

  2. Adriano Alcantara on May 2, 2016 at 3:28 am

    Hi, Yunes.

    I hope I can help you with your questions.

    1) It seems that a CT difference of more than 0.3 between replicates and/or a calibration curve R2 of less than 0.99 are unacceptable. How do we overcome the problem if that is the case?
    I don’t think they are unacceptable but they are not the best results you want to have. So, you need to try to decrease the differences about your replicates to the minimal. If I did not get it wrong, Suzanne said up here in the text, o.9 at least.

    2) People use the blue and the yellow solution mixes. Blue is the Syber mix and yellow is the cDNA. Which one to go first and how to avoid contamination?

    Usually, the normal standard, was to distribute the SYBR mix first and then the cDNA. In this order, the chance to make air disturbance able to take the cDNA from one well to the other is less probable.

    Do we need to change pipette tips for every well? or keep it for every set of replicates?
    In my humble opinion, You should avoid to pipette anything with the same tip to more than one well. I do change tips everytime.

    Finally, There is no such a thing as elementary questions. Your questions are the most important for the right flow of your work. So, keep on doing all questions you have in order to eliminate them. Ok?

    Good luck!

  3. Yulduz on April 14, 2016 at 1:10 pm

    Hi everyone, I wanted ask about data analysis. I`ve read about it a lot but still couldn`t understand. I have done many qPCR and I’m doing data analysis in Excel, but I couldn`t understand what is it fold changes?
    Thank you!

  4. Carin on April 7, 2016 at 2:59 pm

    Master mixes are your friend. I find putting my template in the master mix and adding the primers to the plate give the best results. Make the biggest master mix possible at each stage (without template/primers; with template/primer). This is for two-step qPCR where I’ve made and stored my cDNA and use it for lots (and lots) of templates.

    My own personal method to reduce the number of tips without cross-contaminating is to pipette the primers into the wells with the plate back-to-front (top row facing you) and then swivel it around to add the master mix. Seal with adhesive film and centrifuge at 3500 rpm for 30s to collect everything at the bottom.

    I use an external standard (lambda phage gDNA) to quantify by templates because my samples don’t have a stable internal reference to normalise to (the treatment affects primary, secondary and “housekeeping” metabolism). See papers by Rutledge on LRE for details.

  5. Fatma jumapili on March 14, 2016 at 8:34 am

    I will be able to understand my difficult situations in this new career.

  6. livid11 on September 7, 2011 at 8:32 am

    Hi, I was wondering do you do your standard curve dilutions in TE, water, or some other buffer? Do you think it matters?

    • ofira on September 5, 2017 at 6:56 am

      did you get a reply? because i was wondering about it as well

      • Dr Amanda Welch on September 5, 2017 at 6:50 pm

        You should do your dilutions in whatever you have your DNA dissolved in. So, if it’s in pure water, then use pure water and so on.

  7. Yunes on June 22, 2011 at 10:06 pm

    Hello Suzzane,

    I am a beginner on RealTime PCR and am using Applied Biosystem SyberGreen mix for my PCR and I have the following questions:
    1) It seems that a CT difference of more than 0.3 between replicates and/or a calibration curve R2 of less than 0.99 are unacceptable. How do we overcome the problem if that is the case?
    2) People use the blue and the yellow solution mixes. Blue is the Syber mix and yellow is the cDNA. Which one to go first and how to avoid contamination?
    Do we need to change pipette tips for every well? or keep it for every set of replicates?

    Sorry for asking elementary questions.

  8. Marisa on April 29, 2010 at 10:37 am

    Great tips!

    But just one question…where and at which temperature you store your primers? Should we avoid low temperatures like -80?

    • Suzanne on April 29, 2010 at 1:37 pm

      Hi Marisa,
      You can store your primers at -80C but aliquot them to avoid multiple freeze thaws. I would recommend resuspending the concentrated stock of primer in a TE buffer so the little bit of EDTA will protect from DNase degradation and the buffer will protect from the acidity of water causing hydrolysis. You can aliquot this stock and put them at -20C. Then you make your dilutions of a working stock. You can make the working stocks in just Tris buffer so the EDTA is diluted out (which is even more diluted once the primer is added to the PCR).
      The working stocks should also be aliquoted so that you do not freeze/thaw them more than a few times and also, in case they become contaminated with PCR product, you can easily solve the problem and not have to throw away the entire stock of primers.

      Storage at -20C is ok too and -80C is fine if you want to put them away for a long time without needing to get to them frequently. If you are in the middle of a project and need easy access to the primers, use -20C.

      • Ian Mackay on November 4, 2015 at 9:57 pm

        -20 works well. I recently went back to a 7 or so year old “TaqMan” oligoprobe and it work as well as ever-in fact better than what we were using leading to some questions for the current company I uses!

        • Mordechai Applebaum on March 15, 2016 at 6:39 am

          Surprises happen, quite often inexplicable. I used a >10-year old restriction enzyme stored at -20 (in glycerol) and it worked fine. But I would never count on it or plan that it would work. Good practices in lab work ultimately saves time, money and most importantly frustration.

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