Bio 181
Spring, 2009
qPCR
QUANTITATIVE REAL-TIME PCR I:
Determining Primer Efficiencies
Introduction
We have conducted PCR in several of our labs. In these experiments, we monitored the reaction by
examining the products after the reaction was complete using agarose gel electrophoresis. For this
reason, this approach is often called end-point PCR. A relatively recent advance in PCR has been the
development of techniques that improve our ability to monitor PCR product accumulation. In these
techniques, the amount of product resulting from PCR is monitored during the PCR rather than just at the
end-point of the reaction. Since the product is monitored as it is being produced, these techniques have
been named real-time PCR.
All of the real-time PCR techniques that have been developed monitor the amount of product during the
reaction using some method to fluorescently label the product and then record the resulting fluorescence.
Several techniques have been developed to accomplish this. We will utilize a strategy that uses the dye
SYBR Green I to fluorescently label the PCR products. SYBR Green I, like EtBr, binds double-stranded
and becomes fluorescent once bound. Several companies have developed thermocyclers (PCR units) that
allow fluorescence to be monitored. We will utilize a real-time unit that allows up to 96 reactions to be
monitored at one time by carrying out reactions in a 96 well plate. This unit includes a typical PCR heat
block to control reaction temperatures as well as fluorescence unit that will deliver light to the sample
plated at the correct excitation wavelength and then collect the fluorescent signal from each well during
the course of the reaction. An example of the results of PCR of 96 identical reactions is shown below
(Fig. 1). Note that at the point at which these reactions might be analyzed in end-point PCR (likely 30 or
more cycles) significant differences begin to become apparent between samples.
Figure 1. Real-time detection of the amplification of 96 identical reactions. (From: iQ5 Real-Time
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Spring, 2009
PCR Detection System. Kindly provided by Dan Konet, Bio-Rad, Inc.)
While “identical” reactions begin to show differences as the number of cycles increases, early in the
exponential amplification phase carefully prepared “identical” reactions are very similar to one another.
Real-time PCR analyses take advantage of this to increase the accuracy of comparisons between samples
by comparing reactions early in the exponential amplification phase. To make this comparison, an
arbitrary amount of product is set as the “threshold” amount of product and the cycle at that point is the
“threshold cycle”—CT. This is illustrated in Figure 2 below. The horizontal bar is the threshold amount
of product and the vertical arrow is the threshold cycle CT. Note that the larger the amount of starting
template, the lower the CT value will be because fluoresence will accumulate to the threshold level more
quickly.
Figure 2. Determination of CT in a typical PCR. (From: iQ5 Real-Time PCR Detection System.
Kindly provided by Dan Konet, Bio-Rad, Inc.)
Now, since the amount of product yielded in a reaction is proportional to the amount of template in the
reaction initially, we can compare reactions by comparing CT’s. Figure 3 (below) illustrates this by
comparing of a series of 10-fold dilutions. The formula at the left demonstrates that the CT’s of 10-fold
dilutions should differ by a factor of 3.32. Figure 3 also demonstrates that we can relate CT’s to the
amount of starting material. In the same way, we can compare reactions using different templates as long
as the efficiencies of the two PCRs are very close to 100%. A dilution series such as that shown in Figure
3 below is used to test the efficiency of an individual template/primer combination.
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Figure 3. Real-time measurement of PCR yields in reactions with serial dilution of a single
template.
In this lab we will carry out PCRs of a serial dilution of template DNA testing two different primer pairs.
If these primer pairs have PCR efficiencies roughly equal and near 100% we will then have determined
that we can use these primer pairs in a real-time assay. This assay is being developed to examine the
amount of tomato LHA2 gene expression and copy number in Arabidopsis plants that have been
transformed with a construct that includes LHA2.
In order to quantify LHA2 levels we will compare its level to an Arabidopsis gene—GAPDH—which is
normally expressed at moderately high, unchanging levels throughout the life of the plant. To accomplish
this, we will first generate a standard curve across a series of dilutions of Arabidopsis genomic DNA
containing GAPDH and the LHA2 transgene—for which we can use the PCR efficiency test that we are
conducting today.
Standard curves give us an idea of the dynamic range of a particular primer pair for real time PCR. The
dynamic range refers to the concentrations of starting template from which accurate CT values can be
obtained. They also provide an estimate of the average PCR amplification efficiency for the primer pair.
PCR efficiency is a value between 1 and 2 that describes how much of the target amplicon is present after
one PCR cycle compared to the amount present beforehand. A PCR efficiency of 2 indicates 100%
efficiency (after one cycle there are now 2 copies of the target sequence compared to the one copy present
before if we have a perfect doubling). Likewise, a PCR efficiency of 1.9 is a 90% increase in the amount
of target sequence.
Once we obtain CT values for each dilution of the samples, we can create a standard curve by plotting the
log of the starting template amount versus the average CT value. Using the log of the starting quantity is
important because PCR amplification is exponential, so we must adjust the parameters to get a linear
relationship between a function of our starting quantity and the CT value. This plot is our standard curve.
Note that it is not necessary to always know the exact concentration or copy number of the target
sequence because we will still know the relationship between the starting amounts in each case (an order
of magnitude difference between each 10-fold dilution, assuming the dilutions were performed
accurately). If we do not know the exact amount, we can arbitrarily set the value of one dilution as 1 (so
that value used on the graph would be log (1) = 0) and express the starting amounts of the other samples
relative to that.
In order to estimate the average PCR efficiency from this graph, a regression line must be fit to the data.
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Real-time PCR thermocyclers are connected to computers and often have software that can graph the data
and perform these calculations for you, but it is straightforward to do this yourself using a spreadsheet
such as Excel. Once a regression line has been fit to your data, the slope of the line is related to the
average efficiency of the primer pair by the following equation:
PCR efficiency = 10-1/slope – 1
An ideal PCR efficiency will give a slope of -3.32, but typically the acceptable range is -3.6 to -3.1. Note
that this is related to the theoretical 3.32 difference in CT value for each 10-fold dilution mentioned
earlier.
A common way to quantify mRNA expression using real time PCR is to compare the amount of the target
mRNA to another mRNA whose expression level does not vary significantly (often called a housekeeping
gene). GAPDH will serve this function in the LHA2 experiments in Dr. Ewing's lab. This is accomplished
by dividing the starting amount of LHA2 in a particular sample by the starting amount of GAPDH in that
same sample. This produces a normalized value that can be directly compared between different samples,
such as different time points or developmental stages, However, calculating these values requires
plugging the CT value obtained from each primer pair into the regression equation for the standard curve
and calculating a starting quantity which can then be used in subsequent calculations. As a result, being
able to express this ratio in terms of CT values themselves would be advantageous.
This is possible by simply subtracting the CT value of the housekeeping gene from the CT value of the
target gene to produce the ∆CT value ( CT of target - CT of reference) . The ∆CT values of two different
samples can be compared by calculating the ∆∆CT value (∆CT of experimental sample - ∆CT of reference
sample). The ∆∆CT value can then be used in the following equation to extrapolate the difference in
starting quantity of the target sequence between the two samples:
Comparative expression level = 2-∆∆Ct
However, in order for this to provide accurate estimates of relative gene expression, some conditions must
be met. In order to compare CT values directly, the average amplification efficiencies of both the target
and the housekeeping gene must be approximately equal. This can be demonstrated by construction of
standard curves for each amplicon and showing that the amplification efficiencies are approximately
equal (i.e. comparing the slopes of the regression lines). The best way to do this is to calculate CT
values for each starting template amount and graph the log (starting quantity) versus ∆CT. If the slope of
the regression line is less than 0.1, the primer pairs can be used together for the ∆∆CT method.
There is one further complication: the ∆∆CT method assumes that the PCR efficiency approaches 100%,
which is not always possible. However, it is easy to get around this since we already had to estimate the
average PCR efficiency. We can simply substitute the calculated efficiency for 2 in the above equation to
represent the actual efficiencies observed. Recall that 2 represented the maximum PCR efficiency of
100%, so if our actual efficiencies are 90% we can simply plug in 1.9.
A comparison of the CT of the amplification of an unknown to a standard curve:
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Procedures
____________Reagents__________
1.
2.
3.
4.
Arabidopsis genomic DNA template stock (10 ng/µl stock)
Internal standard primer pair 1: GAPDH for / GAPDH rev
LHA2-specific primer pair: LHA2<250 f / LHA2<250 r
iQ SYBR Green Supermix
Template dilution series:
1. Obtain 6 tubes and label them 1-6.
2. Prepare serial dilutions of the plasmid template as follows:
a. Aliquot 90 µl of sterile H20 into each of the six tubes.
b. Pipet 10 µl of plasmid template (tube T in your ice bucket) into tube 1 (a ten-fold
dilution).
c. Mix thoroughly and transfer 10 µl of this into tube 2 (another ten-fold dilution)
d. Repeat for tubes 3-5. Leave tube 6 as a no template control (i.e. do not add template to
tube 6).
PCR reactions:
Each group will prepare reactions in duplicate for each of the five dilutions along with duplicate no
template controls—for a total of 12 reactions per group.
We will run all of our reactions on a single 96 well plate and so will need to rotate through the
loading station.
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1. Prepare Master Mix in a 1.5 ml tube according to the following table:
Reagent
2x iQ Supermix*
Primer 1 (10 µM)
Primer 2 (10 µM)
dH20
DNA
TOTAL
Single
reaction
µl
12.5
0.5
0.5
6.5
5
25
Master Mix (for 13
rxns)
µl
162.5
6.5
6.5
84.5
none
260
* 2x Supermix includes dNTPS, Taq (heat activated), SYBR green, Mg 2+ and buffer.
2. Lay-out reactions on the 96 well plated. We will decide this in class and assign each group 12
wells. With 8 groups we will fill the entire plate.
3. We will set up each dilution series in duplicate. To do this, pipet 5 µl into adjacent wells (5 µl
/well) for each of the 5 dilutions (tubes 1-5).
4. Repeat 5 µl into each of two adjacent wells for tube 6 also (as a no template control).
5. Next pipet 20 µl of Master Mix into each well that contains template.
6. Once all groups have loaded their reactions, the plate will be sealed, given a quick spin to collect
all reactions in the bottom of the wells and placed in the real-time unit.
7. The PCR program is as follows…we’ll watch it as it goes!
PCR Program:
The small volume and short amplicon allow short incubation times at each step. In addition, since the
short amplicon is likely completed at the end of annealing we can shorten our cycle to just two steps!
The program is:
1. 94 oC for 2 min. (hot start)
2. 95 oC for 15 sec. (denature)
3. 55 oC for 30 sec (anneal)
4. return to step two 39 times
5. end
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