One of the very first things you need to do when getting set up for quantitative PCR (qPCR) is to determine the efficiency of the assay because knowing the assay efficiency is critical to accurate data interpretation. And you have to do this every time you design and purchase a new primer pair.
Ideally, the efficiency of the assay should be 100%. This means that during the logarithmic phase of the reaction, the PCR product of interest is doubling with each cycle. Perfect PCR efficiency will demonstrate a change of 3.3 cycles between 10 fold dilutions of template.
Sometimes the efficiency is below 100% and sometimes you can get readings of PCR efficiency above 100%. In a future article, we’ll discuss what causes high and low PCR efficiencies and how to fix it.
Today’s article is about the important considerations when setting up a PCR efficiency test for your new assay. The three key ingredients to focus on when getting started are the type of template, the primers, and the chemistry.
Your PCR template can be one of several types of nucleic acids. It can be a plasmid containing the gene of interest (in which case, linearize it), a DNA oligo (most people order a 60-70mer), genomic DNA (works best only if the target exists in multiple copies, such as actin, gapdh, or rRNA gene), and it can also be the PCR product of interest purified and quantified using a spec or picogreen.
Your template for the standard curve needs to be quantified and then diluted serially. I recommend at least 5 dilutions and 1:10 dilutions for the widest linear range. Many people use 1:4 dilutions which is fine, but limits the range in which you can be sure the primer pair is accurate.
Duplicates of each template dilution are required and triplicates will give you greater confidence in the data, especially if one data point drops out for an unknown reason.
I recommend diluting the templates so that the first sample (the most concentrated) comes up around cycle 16-18. If your first sample is coming up too early- like around cycle 8-10, you may run into problems with the subtraction of baseline fluorescence from the samples. This causes shifting curves and loss of sensitivity for detection of low copies of template.
Every time you have a new pair of primers for qPCR, check the efficiency. Even if they are primers you know work well and you just received a new batch. You cannot assume that the primers from one lot to another work the same. Better to catch the problem of a poor synthesis early on than to perform a lot of work on irreplaceable samples only to realize later that the new primers were only working at 80% efficiency.
The concentration of primers to use depends on the enzyme chemistry. It’s a good idea to start with a concentration recommended by the kit supplier as a start and then modify if the efficiency is low. Using too much primer can lead to dimer formation, especially at the low dilutions. This will have a greater impact on SYBR Green data but will also impact hydrolysis probe results. Too little primers and the efficiency will be negatively affected.
Sometimes no matter how much tweaking and optimizing (and enzyme kit changes) you do, you cannot get the assay efficiency to a level that allows for the sensitivity you need. In this case, re-design the primers. If you are trying to make accurate quantifications out past cycle 35, you need the most efficient assay possible. If all of your data is between cycles 18-30, maybe you can tolerate an assay that is only 80% efficient. You can decide what works for you.
The kit enzymes and buffer systems are going to play a big role in the results as well. So when buying a new lot of kit, re-check the assay efficiency as well. Chances are that the kits will not vary, but it is better to check and make sure.
When using SYBR Green assays, the PCR efficiency will be affected by the presence of primer dimers so make sure to always include the melt curve analysis. This is a great benefit of performing SYBR Green qPCR. Primer dimers will affect a hydrolysis probe (such as TaqMan) as well. If reaction components are used up amplifying dimers instead of real product, efficiency will be low. For hydrolysis probe assays, you’ll want to save the reactions and run them on a gel if the efficiency is low so you can analyze what else may be amplifying in the reaction.
With Probe assays, there is also the consideration of the Probe design and the compatibility of three primers in the mix. The probe needs to have a higher Tm so that it lays down on the DNA first. With Probe assays, low efficiency can also be caused by the Probe design or Probe labeling. To determine if efficiency problems are the primers or the probe, run your primers with a SYBR Green kit. If they work well, then the issue is the Probe. If they do not, you know you need to optimize the primers.
Set up a qPCR experiment to test PCR efficiency:
Dilute the template used for the assay with clean water. Make your 10 fold dilutions and make enough for the number of reactions plus one to account for pipetting error. Use a P10 pipettor (or similar pipettor accurate to very low volumes) to achieve low standard deviations in the Cq values between replicates.
To make it easy for yourself, prepare the template DNA so that the same volume is added per reaction (2 ul) in all wells.
In a PCR hood and using designated pipettors, prepare your mastermix (buffer, water, and enzyme) or aliquot your purchased mastermix into the wells of the plate or the special tube or cuvette you are using for your instrument. Make sure to mix the mastermix well before use, especially for commercial mastermixes that may have been sitting. In the bottle, the SYBR Green can settle out and then be distributed unequally between the wells.
If you are running unknowns in the same plate, make sure to have room for 10 wells (duplicates) or 15 wells (triplicates) for the standard curve and then you’ll want to leave 2 or 3 wells for negative control (water alone). If you are doing RT-qPCR, you will need to have a -RT control as well.
Add the template DNA to the wells in a separate location. In our lab, we use different pipettors for this.
Seal your plate with tape or close your cuvettes, and for plates, we centrifuge in a 96 well plate centrifuge briefly. You can also try the method described by Shoba using a Salad Spinner. We never allow the plate to touch any counter surface to protect the outside wells from attracting dust that might affect the path of light for emission or detection.
Put the samples or plate into the instrument and run your pre-set program.
That’s about it for getting started.
The main decisions you need to determine before getting started are the template type, primer sequences, and the chemistry (SYBR vs. probes).
Getting good results begins with a lot of fore-thought and planning in the early steps. The combination of primers, chemistry, and template are the three essential ingredients for high quality data and accurate results. Work this out systematically up front and save yourself a lot of time and pain later.
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