Quantitative PCR analysis
Written by Steve Doyle (s.doyle@latrobe.edu.au) - March 2014
-
-
-
Quantitative PCR (qPCR) (or “real-time” PCR) is a technique used to detect the
change in DNA template concentration during a PCR reaction
Its basis for detection is the measurement of fluorescence after each PCR cycle,
using either dsDNA-specific fluorescent dyes (eg. SYBR green) or probes (eg. TaqMan
hydrolysis probes)
There are two main types of qPCR: relative and absolute
o Relative qPCR is mainly used to detect changes in gene expression in a cell or
tissue by using RNA as a template for the PCR. It does this by measuring the
“relative” amount of PCR amplification of a target gene compared to a
reference gene.
o Absolute qPCR can also be used to measure gene expression changes,
however, it aims to determine the exact copy number of DNA/RNA
molecules. It is possibly to do this by comparing a unknown sample of
interest against a standard curve of known concentrations
This workshop will focus on using absolute qPCR to analyse our O. volvulus DNA
samples, and we will do so for a number of reasons:
o Determine DNA concentration: samples with low DNA concentrations that
cannot be detected using conventional spectrophotometric methods may still
be quantified by qPCR.
o Determine optimal DNA concentration for assay: by generating standard
curves, we can determine the necessary and sufficient DNA concentration
that will allow robust amplification. This is particularly important as too much
or too little DNA can influence how well a PCR performs. As qPCR is very
sensitive in its ability to detect DNA, it also will allow you to use much less
DNA than you think, and therefore, increase the number of assays that can
be performed with a particular sample
o Determine assay efficiency: in theory, PCR is an exponential reaction that
doubles the DNA concentration after each cycle. However, in reality, this is
not always the case: non-specific amplification or inhibitors present in a
sample can change the reaction efficiency, and therefore influences the
interpretation of the fluorescence signal. We therefore measure the reaction
efficiency across a standard curve to ensure that we have confidence in
comparing different samples and different assays.
o DNA standardization: in preparing samples for HRM, we need to ensure that
the DNA concentrations of all our samples are (1) within an approximate 10fold concentration range ( ~3.3 PCR cycles = 10-fold concentration difference)
of each other, and (2) that the cycle threshold values (Cq; this will be
explained later) for amplification are < 30 cycles. We can use the
-
-
-
amplification plots to adjust the DNA concentrations of our samples to be
within an acceptable range
The DNA used to generate a standard curve needs to be from either good quality
genomic DNA of a known concentration, cDNA (if measuring gene expression), or a
cloned PCR product in a plasmid.
o Often cloned PCR products are convenient to use as it is easy to get sufficient
high quality DNA without having to waste precious genomic DNA samples.
We will use both plasmid DNA and genomic DNA to illustrate both.
o It is important to accurately quantitate the DNA used for generating the
standard curve as it will influence the analysis of all samples with unknown
DNA concentration.
 We use a Qubit (Life Technologies) and will show you how to use it,
however, a combination of spectrophotometric and gel
electrophoresis to determine DNA concentration can be sufficient
 NOTE:
The qPCR enzyme that we will be using is called SsoAdvanced Universal SYBR Green
supermix (Biorad), which we have shown to work quite well with poor quality and
low concentrated O. volvulus samples.
Relevant reading
o Ssoadvanced universal SYBR green supermix:
 See “Biorad - SsoAdvanced Universal SYBR Green Supermix Instruction Manual.pdf”;
 http://www.bio-rad.com/en-us/product/ssoadvanced-universalsupermixes/ssoadvanced-universal-sybr-green-supermix
o Qubit fluorometer
 http://en.wikipedia.org/wiki/Qubit_fluorometer
 http://www.lifetechnologies.com/order/catalog/product/Q32851
o qPCR
 http://www.bio-rad.com/en-au/applications-technologies/real-timepcr-data-analysis
 MIQE guidelines – an attempt to improve qPCR practice
 NOTE: many aspects are specific to using RNA for gene
expression experiments, however, there is still a lot of good
information about experimental design and sample quality
that is relevant.
 Biorad tech note - a practical approach to RT-qPCR - publishing
data that conform to the MIQE guidelines
 Bustin 2009 - The MIQE guidelines - Minimum Information for
Publication of Quantitative Real-time PCR Experiments
 Taylor 2013 - The state of RT-qPCR - firsthand observations of
implementation of MIQE
STEP 1: generating dilution series
1. as discussed above, it is important to accurately determine the DNA concentration of
the sample you will use to generate the standard curve
a.
follow the Qubit protocol
2. make a aliquot of DNA at 0.5 ng/ul concentration
a.
the volume you will need will depend on how many standard curves you plan to generate –
you will be 2x 2ul for each primer set you are testing
b.
minimum 20 ul, ie. 10 ng in 20 ul
c.
this will be the first dilution (10-1)
3. vortex for 5 seconds, pulse spin, vortex for another 5 seconds and pulse spin for a
second time
a.
this is important to ensure the dilution is well mixed prior to making the next dilution
4. aliquot 90 ul of pure water (MilliQ; HPLC grade; molecular biology grade H2O) into
each of a series of 7 microcentrifuge tubes
a.
this will generate the remaining dilution series ( 10-2, 10-3, 10-4, 10-5, 10-6, 10-7, 10-8 )
5. Transfer 10 ul of DNA from the 0.5 ng/ul tube (10-1) into the second dilution tube
(10-2)
6. vortex for 5 seconds, pulse spin, vortex for another 5 seconds and pulse spin for a
second time
7. repeat steps 5 and 6, transferring 10 ul of the previous dilution into the next tube,
until all dilutions are made
8. 2 ul of each dilution will be used per well in the qPCR assay. This means that the
amount of DNA in the standard curve will be 1 ng, 100 pg, 10 pg, 1 pg, 0.1 pg, 0.01
pg, 0.001 pg, 0.0001 pg
a.
the lower concentrations are likely to be too dilute for genomic DNA samples to amplify,
however, they will likely still be relevant for plasmid DNA templates
STEP 2: Master Mix Setup
- The basic master mix setup for SsoAdvanced Universal SYBR Green super mix is
outlined in the product manual. We have modified this protocol from a 20 ul
reaction to a 10 ul reaction.
- As discussed in the PCR optimization protocol, 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.
1. Determine the number of reactions needed per primer set – all reactions will be
performed in duplicate, including a no template control (NTC)
a.
Use the 96-well plate map to design your experiment and placement of sample wells – this
will keep track of your samples while pipetting, and will help to input the data into the CFX
manager for data analysis
1
A
B
C
D
E
F
G
H
2
3
4
5
6
7
8
9
10
11
12
In your DNA-free area:
2. Combine the following per primer set:
Reagent
Per reaction
Mastermix volume
(#rxns + 1)
SsoAdvanced Universal
5
buffer/enzyme mix (2X)
Sense primer (10 uM
0.5
stock, 0.5 uM final)
Antisense primer (10 uM
0.5
stock, 0.5 uM final)
H2O
2
TOTAL Volume
8 ul
In your DNA working area:
3. Aliquot 8 ul of master mix into an 8-well strip / 96-well plate (depending on how
many reactions you are performing
4. add 2 μl of H2O to the NTC
5. add 2 ul of each DNA dilution to the relevant wells
6. once all DNA is distributed in the 8-well strip / 96-well plate, seal with a cap-strip or
plate seal well, and centrifuge for 2 minutes at 2000 RPM
7. proceed with loading onto the PCR machine
a.
NOTE: if you are not planning on doing the run straight away, you can store the strip/plate at
-20C in the dark. I would not recommend leaving it for more than a few hours.
Step 3: PCR machine setup
- the basic program that we will use is outlined in the diagram below
98C
2 min
Pre-amp.
denature
-
-
-
98C
5 sec
Op -Tm
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
it 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
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
The “Opti-Tm” step refers to the optimized annealing temp for the particular primer
set that you are using. This should be been already determined using a gradient PCR.