PCR Primer optimization
Written by Steve Doyle (s.doyle@latrobe.edu.au) - March 2014
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PCR is a simple molecular technique in theory, however, it can be difficult in practice,
sometimes without any logical reason as to why.
It is important therefore to optimize and control the parameters that you can to
ensure you can have the best chance of it working correctly.
To do this, the main steps that we take every time we get a new primer set are to:
o Correctly determine the primer concentrations (PCR/HRM/qPCR)
 You should not believe the values that the primer synthesis company
gives you – in most cases, it is more that what is stated, and having
too much primer can lead to non-specific amplification and primer
dimers
o Perform a gradient PCR to determine the actual optimal annealing
temperature (PCR/HRM/qPCR)
 Again, you should not trust the providers’ estimate of Tm. There are
many algorithms out there to predict the Tm, however, in your hands,
it will ultimately depend on the polymerase and buffer that you use.
The old general rule of annealing temp = Tm-5C is fine, however, if
you have access to a gradient PCR machine, it is best to test it out for
yourself.
o Determine the primer amplification efficiency (HRM/qPCR)
 Primer efficiency tests the doubling rate of the PCR. A perfect PCR will
have an efficiency of 100%, which means that the concentration of
PCR product will double with each cycle (ie. 2n , where n= # PCR
cycles). However this is not always the case, and primer efficiencies
will deviate from 100% due to factors such as non-specific
amplification or PCR inhibitors. Knowing the efficiency of the reaction
is especially important for qPCR, as it is often used in the analysis
calculations
o Perform a melt curve analysis at the end of the PCR (HRM/qPCR)
 This is only relevant if you are using the qPCR machine. You can add a
melt curve to the end of the PCR cycle program, which allows to
determine if the PCR product you have generated is a single product,
or if there has been non-specific amplification such as primer dimers.
Both HRM and qPCR require a single specific product, which will
translate into a single melt peak.
The above recommendations are probably the most critical for success, however,
other relevant parameters that could be optimized are:
o Primer concentration
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o DNA template concentration (we will address this in the qPCR section)
o Magnesium concentration
o Cycling times
Relevant reading/links associated with the section
o IDTDNA website (www.idtdna.com) and oligo analyser tools
(https://sg.idtdna.com/analyzer/Applications/OligoAnalyzer/)
 My preferred website for predicting primer properties.
o MIQE guidelines (see Bustin 2009 paper; Biorad tech note - a practical
approach to RT-qPCR)
 These are quite specific for qPCR, but have very relevant info for
standard PCR and HRM
o Biorad notes
 qPCR assay design and optimization (http://www.bio-rad.com/enau/applications-technologies/qpcr-assay-design-optimization)
 SsoAdvanced Universal SYBR Green Supermix (http://www.biorad.com/en-au/product/ssoadvanced-universalsupermixes/ssoadvanced-universal-sybr-green-supermix)
STEP 1: Determine Primer concentrations
- primers are typically delivered to you as a lyophilised pellet, which need to be
resuspended in either TE buffer or H2O.
- we typically resuspend in H2O to make a stock concentration of 100 μM
- once the concentration is confirmed, we make aliquots at 10 μM working
concentrations
- to determine the concentration of the dissolved primer stock, we need to measure
the absorbance of the primers using a spectrophotometer
Equipment and consumables needed
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vortex
centrifuge
spectrophotometer
Protocol
1. before you start, calculate the volume of liquid required to achieve a concentration
of 100 μM
2. centrifuge the primer tube at max speed for 30 seconds
a.
some primer may be stuck to the sides/top of the tube, and you don’t want it to escape
when you open the tube
3. add the required volume of liquid to the primer tube
4. vortex the tube for 5 seconds
5. centrifuge the tube for 10 seconds
6. repeat steps 4 and 5 for a total of 2X vortex/spins
7. per primer stock, make three 1/100 dilutions, ie. triplicates
a.
add 2 μl of primer to 198 μl of H20 in a new microcentrifuge tube
8. determine the absorbance at 260 nm (A260) for each of the triplicates for each
primer stock using the spectrophotometer
9. calculate the mean absorbance (μA260) value of your triplicate tubes
10. calculate the concentration using the following formula (Beer-Lambert Law)
a. Concentration (M) = (μA260 x Dilution Factor) / extinction coefficient
i. Dilution Factor = 100 ; you made 1/100 dilutions above
ii. Extinction coefficient = specific for each primer – the primer synthesis provider
should give you this, but if not, you can use the Oligo Analyser tools at
www.idtdna.com
11. Adjust the concentration of the stock tube to 100 μM using C1V1=C2V2
12. Make 1/10 dilutions of the primer stocks to achieve the final 10 μM working stock
13. Primer stocks can now be stored in the freezer until needed
STEP 2: Gradient PCR
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gradient PCRs are performed using PCR machines that are capable of generating
thermal gradients across the 96-well block
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depending on the machine, thermal gradients may run from the bottom (cool) to the
top (hot), OR, from left (cool) to right (hot). Make sure you know which gradient
direction your machine uses! The Biorad CFX machines have gradients running from
the bottom to the top.
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Gradient PCRs therefore aim to vary the temperature within each well at a defined
step of the PCR, which is typically the annealing step
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This allows you to find the optimal annealing temperature of your PCR, which is the
highest temperature that shows good amplification, with hopefully no non-specific
amplification
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DNA: it is best to begin with a sample that you know is good quality and has
sufficient concentration to allow you to do a few assays, ie. not a precious sample.
As a starting point, I would make an aliquot of DNA at a concentration of 0.1-1 ng/ul.
We can work out what the lower detection limits are later, but for now, use a
concentration that should work easily.
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It is good practice to keep all of your PCR reagents and set up your master-mixes in a
dedicated DNA-free area that is separate to where you keep and aliquot your DNA,
ie. your workbench. At LTU, we have a DNA-free room dedicated to this. This will
help prevent contamination.
Equipment and consumables needed
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Gradient PCR machine
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8-well PCR strips and caps
Protocol
In your DNA-free area:
1. For each primer set, prepare a PCR master-mix in a single microcentrifuge tube
Reagent
Volume (μ) – 1X
Master-mix Volume
(# samples +1 extra) – 10
rxns
SsoAdvanced Universal
5
50
Sense primer (10 μM)
0.5*
5
Antisense primer (10 μM)
0.5*
5
H2O
2
20
SYBR Green master mix
* primer concentration is at 500 nM to start with. This can be optimized later if
desired
In your DNA work area:
2. aliquot 8 μl of master-mix into each well of an 8-well strip, and an extra single PCR
tube for a negative (no template) control (NTC)
3. add 2 μl of H2O to the NTC and close the lid – do this before opening any DNA
4. prepare your standardized DNA sample if you have not done so already – you will
need at least 16 μl of a 0.1-1 ng/μl solution
5. add 2 μl of the DNA stock to each well in the 8-well strip, and close the strip with
strip caps
a.
each well contains exactly the same reaction – what will differ is the temperature applied to
each well
6. ideally – you should centrifuge the 8-well strip for at least 30 secs at max speed to
ensure all liquid is at the bottom of the well and that bubbles are minimized
a.
bubbles can cause two problems: (1) they cause uneven temperature in the well, and
therefore you can have problems with amplification, (2) they can cause problems with
fluorescence detection, and may not be detected properly
7. load the 8-well strip in the PCR machine, top to bottom, and load the single NTC at
the bottom
a.
the NTC should be at the coolest temperature of the gradient, as this is where you are most
likely to see non-specific amplification. If it is clear, you can be pretty confident that at high
temps it will also be clear. If it is not clear, but your optimal temperature in your DNA wells is
higher, subsequent PCRs at the optimal temperature will also have an NTC, and you can
check it again then.
Setting up the PCR conditions
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to start with, a good place to set the gradient is from 50-70C. This can be refined
later if needed, but for the most instances, it will give you all the information you
need.
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The SsoAdvanced Universal manual will give you basic cycling parameters, which
includes using a 2-step PCR (denaturation, followed by annealing and extension in a
single step). As we are trying to find the optimal annealing step conditions, we will
use a standard 3 step PCR (denaturation, annealing and extension as separate steps).
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qPCR and HRM typically runs for 45 cycles. This is obviously longer than a
conventional PCR, however, it is important that the amplification curves of each
product all reach a plateau so that they can be compared properly. Therefore, we
need to cycle for longer.
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We will include a melt curve here (best to do it on all PCRs done on the real-time
machine), so we can compare the amplification curve and melt curve to find the
optimal conditions.
8. setup the following protocol on the PCR machine:
98C
2 min
Pre-amp.
denature
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98C
5 sec
Gradient
15 sec
72C
15 sec
PCR amplifica on
(45 cycles)
95C
30 sec
60C
2 min
Post amplifica on
melt and anneal
65C
15 sec
95C
0.5C/read
Melt curve
the run consists of a conventional PCR that is run for 45 cycles, followed by a fast
melt curve analysis
o the PCR is run for 45 cycles to ensure that the amplification reaction is
finished and the reaction rate has reacted a plateau
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o this is important to enable comparison between samples
it is important to ensure the sample volume is set correctly to 10 ul on the machine.
o The volume affects the rate and time it takes to cycle between temperatures,
and if set incorrectly, will result in different temperatures in each well than
those programmed
NOTE: the gradient step should be set from 50-70C