POLYMERASE CHAIN REACTION
Objectives
1) To understand factors governing choice of PCR primers.
2) To understand factors limiting PCR.
3) To understand real time PCR.
Basic mechanism of PCR
The polymerase chain reaction can be used to radically amplify a short region of the genome. The
genomic DNA is denatured, primers are annealed bracketing the region, and DNA synthesis is conducted
from the primers doubling the number of molecules in this region. The cycle is repeated many times, until
most of the DNA in the tube is from between the primers. The process is automated by a machine that
cycles between polymerization temperature and a denaturing temperature, and the use of a thermostable
polymerase (AmpliTaq) that survives the denaturing cycle.
Ref: Saiki, et al., Science 439,487 (1988).
A typical PCR profile is shown above. Denaturation time is usually about 90 sec., and mainly
represent the time your particular machine needs to stabilize at the new temperature. Annealing time
reflects the kinetics of the annealing reaction, and mainly depends on the primer concentrations. (1uM
primer anneals to completion in about 45 sec; half that concentration takes twice as long). The extension
time depends on the length of the products. (The polymerase only goes about 2000 bases/min.) The
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annealing temperature is usually about 5oC below the TM of the primers. The extension and denaturing
temperatures depend on the polymerase being used. The annealing temperature depends on the TM of the
primers being used.
The use of the term TM in PCR hides a somewhat complex situation. TM of a duplex is the
temperature at which it is half duplexed at equilibrium. That value is dependent on salt concentration and
is usually indexed to 1 M NaCl. The TM of an oligo against a template is the temperature at which the
template is half duplexed at equilibrium. That value is also dependent on oligo concentration (about 1oC
per factor of 2 around the primer concentration used for PCR). For PCR you want the template to be
mostly all duplexed (primed) so that each cycle will double the amount of product. You also have a
kinetic limitation in that the hybridization must approach completion in the short time period set aside for
annealing. The primer concentration and salt conditions in a PCR reaction have been chosen to achieve
approach to equilibrium in this short time, and also to make the effective TM of the experiment come out
close to the thermodynamic TM of a duplex. Hence you will see formulas applied to calculating PCR
annealing temperatures that would not immediately seem to be relevant.
A consequence of this is that if the PCR primer concentration drops, the efficiency of reaction will fall off
dramatically. The main causes of low primer concentration are hairpin formation or primer dimer
formation (see below). It is also true that the discrimination for priming with temperature is not as sharp
as TM curves you may be familiar with. Having two PCR primers discriminating is very important to
achieving discrimination. Some procedures intentionally take advantage of priming that would seem to be
prohibited by the Tm. Remember that once a primer primes in spite of mismatches, the product it makes is
now going to be perfectly matched in future cycles.
Primer Design (ref: Wetmur, J. (1991) Crit. Rev. in Bioch. and Mol. Biol. 26:227-259.)
In a large target (ie., the human genome), one can expect any primer to adventitiously prime at
some incorrect sequences. Specificity is only possible, because it is unlikely for two such false priming
sites to be close enough together to make a PCR product. However, if you do something to increase the
potential for non-specific priming, you're going to see a lot of strange and unrelated PCR products. You
can save a lot of trouble by designing the primers according to the criteria below. Computer programs are
available that do a good job of identifying primers that will work cleanly.
1) Balance the TM's of the two primers.
2) Avoid self-complementarity and complementarity between the two primers.
3) Avoid complementarity with repetitive sequences in the target DNA.
Balancing the TM's
The reason for this is obvious. If the primers have radically different TM's, then one of them must
be working at low stringency. The original protocols call for making the final adjustment of stringency by
altering the Mg+2 concentration. (This is simpler than changing the annealing temperature itself, because
you can put tubes with different Mg+2 in the PCR machine at the same time). However, that won't make
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the primers better balanced; it will just find the best compromise stringency. The better approach is to get
the TM's right in the first place.
The effect of different Mg+2 concentrations on TM.
total Mg+2
mM
--------0.8
1.3 (std.)
2.0
2.5
3.0
3.5
4.0
free Mg+2
mM
---------0.175
0.575
1.375
1.875
2.375
2.875
3.375
T elev.
C
-----5.2
8.1
9.9
10.7
11.4
12.0
12.5
% G+C tolerated
in template
--------------70.5
63.4
58.9
56.9
55.2
53.9
52.7
A crude estimate for the TM of an oligonucleotide is: 2oC x (A+T residues) + 4oC x (G+C residues). This
equation is indexed to 1 M NaCl, 0 mM MgCl2, and 100 pM concentration. As it turns out, the changes to
PCR conditions (50 mM NaCl, 1.3 mM MgCl2, 1 uM primer concentration) nearly cancel out. This
equation tends to predict a temperature about 5oC under the true TM and so the temperature predicted is
often used directly as the annealing temp.
Mispriming on slightly mismatched sequences is difficult to exclude during PCR. As a rule of
thumb, the template will be about 10x less primed for each 4-5oC above the TM of the primer. The same
oligo with a 1 bp mismatch is less stable by 1oC/percent mismatch or about 5C. However efficiency of
PCR priming doesn't fall off as sharply. An oligo used 10oC over its TM (say because there are 2
mismatches) will only be duplexed to 1/100 of the template at any time. But the polymerase will extend
transiently bound primer, possibly yielding nearly complete priming by the end of the annealing period.
Suppose enough mismatches are present to exclude priming to 1/1000 of the perfectly matched sequence,
even given the polymerase effect. On subsequent rounds the misprimed material now perfectly matches
the primer and amplifies normally. The misprimed product will appear about 10 cycles after the fully
matched product. The efficiency of this low stringency priming mode tends to be more affected by
mismatches near the 3' end of the primer than by the overall primer stability. The outcome of these
considerations are:
 It is not practical to exclude false priming based on one or two mismatches.
 If you amplify enough cycles trying to save a failed experiment, eventually all sorts of strange
products will appear.
 False priming is reduced if the 3' end of the primer is AT rich.
Mg+2 concentration also affects the TM of the PCR product with respect to denaturation on each
cycle. At %GC in the product of > 65%, the product may not be able to denature at all in the presence of
the reaction buffer. In this case you can add some DMSO (or other denaturants), reduce the [Mg+2], and/or
use an alternate thermostable polymerase that survives a few degrees higher denaturing temperature. You
do get some help with this problem if your PCR product is short (<500 bp), since that destabilizes it a
degree or two. To realize this benefit, you would have to do "hot start" (see below) and hold the Mg out
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until after the original template had a chance to denature. An ultimate solution would be to substitute a
base analogue for G (McConlogue, L., Brow, M.A.D., and Innis, M.A. (1988) Nucl. Acids Res. 16: 9869.
There are computer programs that do a more sophisticated calculation of the oligo's Tm taking into
account the dinucleotide composition, the anticipated ionic conditions, and the anticipated primer
concentration. Most people try for 20 base oligos with about 50% G+C and a TM of 60oC.
Another issue that requires a good understanding is whether or not to choose primers with G+C rich 3'
ends. The idea behind G+C rich 3' ends is to clamp down the end recognized by the polymerase in the
case where there are some mismatches between the oligo and the template. If you're trying to prime in the
presence of mismatches, then a G+C rich 3' end is good. On the other hand, if your primer is known to
perfectly match its template, then the G+C rich 3' end is only going to promote false priming at the wrong
sites. In this case make the 3' end of the primer A+T rich. This principle also applies to the design of
DNA sequencing primers.
On a 20 base primer, it costs about 5oC to tolerate each mismatch to the template. So if you are raising
nonspecific products, move the annealing temperature up by 5oC. It's unclear if any of the predictive
methods are better than +- 5oC.
Software
There are many programs available that profess to design PCR primers. UTHSCSA has a site licensed
copy of Oligo 6.0. This is a full capability program with excellent documentation concerning the
scientific principles. I recommend using this program, or benchmarking other programs against it. To
successfully handle oligos with pathological sequences, a program should use a nearest neighbor
algorithm. A well respected web-based program is primer3 (at MIT, http://frodo.wi.mit.edu/). For jobs
that require batch processing of lots of sequence for automatic PCR primer picking, request the
bioinformatics center to import and install primer3. The prime program in the GCG package also uses a
nearest neighbor algorithm.
Primer dimer
If the primers can prime on themselves or each other, then they will be used up making a small
dimeric product. Since the primer concentration is a million or more times higher than the template, both
its TM is pushed up and it only has to get within about 30oC of the true temperature to make enough
primer to subsequently amplify as well as the perfectly matched template. As little as a 3 base perfect
complementarity of a 3' end can cause detectable primer dimer synthesis.
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Rather than quantitatively trying to calculate primer dimer efficiencies, most people use these rules of
thumb:
 Avoid 3 base match of 3' end of a primer to itself or its partner.
 If 3 base match can not be avoided, make them be A's and T's as much as possible.
 If a primer dimer problem seems unavoidable, compose and/or use the primers for annealing at
the maximum temperature allowable (the extension temperature).
Length effect, and the effect of over-amplification.
The PCR product doubles nearly quantitatively through early cycles, but then the accumulation
levels off. One limiting factor is that the polymerase misincorporates bases and then fails to extend the
unpaired 3' end. Another is that as the concentration of product builds up, it begins to anneal with itself
rather than the PCR primers. For reasons that are not totally clear, if you continue cycling long enough,
the product is converted to a smear of lower gel mobility, and eventually sticks entirely in the slot. I
suspect that this happens because the final PCR products prime illegitimately on one another making
mosaic structures that denature and reanneal to form networks. This most commonly happens to people
who amplify from a plasmid or a previous PCR product and put in far too much template. The reaction
plateaus in the first few cycles and then spends the rest of the time getting into this aberrant mode. If you
see this, try using 1/1000 as much template, then 1/1,000,000 as much.
The larger fragments reanneal more effectively, and therefore are product inhibited earlier in the
reaction. There are practical size limits of several kilobases on the size of the products that can be
amplified. 75 to 500 bp is optimal.
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The PCR primers can be expected to prime spuriously at a low level; and eventually to generate a
small spurious product with a primer on both ends. Once such a product is produced, it will amplify at a
normal rate and eventually overtake the expected products. Consequently, it doesn't pay to over-amplify.
The greatest risk of spurious priming is when the reaction is assembled and before it is heated up
for the first cycle. This problem can be suppressed by leaving out one of the components of the reaction
until it reaches the first denaturation cycle. This is sometimes called "hot start" PCR.
Hot start PCR is implemented in several ways. The simplest is to leave out the Mg+2 until the tubes are
hot, then open them and add the Mg+2. Another way is to solidify a wax on the top of the reaction volume
and then add the Mg+2 over the wax. When the wax melts, the Mg+2 mixes into the reaction. A third way
is to add single stranded binding protein to the reaction. It ties up the primers until the temperature goes
up and denatures the protein. Finally, the polymerase can be tied up with antibody until the denaturing
temperature denatures the antibody.
Ref: D'Aquilla, et al. 1991. Maximizing sensitivity and specificity of PCR by preamplification heating.
Nuc. Acids Res. 19:3749.
Long-accurate PCR
The maximum length of a PCR product can be increased up to 50 kb by using a mix of a
thermostable polymerase that cannot proofread with a little bit of thermal stable polymerase that can
proofread. Apparently the proofreading enzyme resolves mismatches caused by misincorporation. This
method is still finicky at the time of this writing. To attempt this, definitely look for a kit that comes with
a positive control.
Ref: Wayne M. Barnes. 1994. PCR amplification of up to 35-kb DNA with high fidelity and high yield
from lambda bacteriophage templates. Proc. Natl. Acad. Sci. USA 91: 2216-2220.
Uses of PCR
Diagnostics
The simplest use of PCR is the determination of whether the target sequence is present or not.
PCR is sufficiently sensitive to detect the presence of one copy of HIV genome per million blood cells.
The theoretical amplification is 2 to the power of the number of cycles.
Number of cycles
20
40
Amplification
106
1012
The true amplification can be expected to be lower than the theoretical amplification. The usual
number of cycles used is 30-40 cycles for amplification of single copy mammalian genes, and up to 60
cycles for HIV detection.
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Contamination problem
Due to the extreme sensitivity of the PCR assay, it is vulnerable to contamination. The amount of
the target DNA fragment in the sample might be 10-6 to 10-13 of the amount of PCR product produced by a
positive assay. Consequently, even a minute contamination with a prior PCR product into a DNA sample
will produce a false positive signal. The amount of contamination required can spread as an aerosol. The
following precautions are recommended:
1. Use of a separate set of micropipets and other implements to prepare templates from those used to
manipulate PCR products. Some labs use an entirely separate template preparation room. Note: a
common oversight is to prepare a control reaction by diluting a control plasmid with the "clean"
pipets. The control plasmid, having a high concentration of positive DNA, should be kept away
from the "clean" pipets and template area for the same reason the PCR products are kept away.
2. Amplification of a negative control with all of the components except the template. Even experienced
PCR practitioners suffer from contamination problems on a regular basis.
3. Use of dUTP instead of dTTP in PCR amplification coupled to pretreatment of the reaction with uracilN-glycosylase. The idea is that a major source of contaminants is previously produced PCR
products. If the previously made PCR products were made with dUTP, then they can be destroyed
with the enzyme uracil-N-glycosylase (ung). The uracil-N-glycosylase is mostly destroyed during
the high temperature initial DNA denaturation, and remains inactive above 55oC. However, it can
interfere with the amplification if annealing is done below 55oC. It can also interfere with post
amplification analysis of the PCR product, and will preclude cloning of the PCR product unless the
host strain is ung-.
Mutation Screening.
PCR products can be directly sequenced. Therefore, it is possible to directly amplify and
sequence DNA from any genome without the intermediate cloning step. The preferred sequencing method
is cycle sequencing (because it works well on double stranded linear molecules).
The PCR product can be run on a gel system for which any base change within the fragment alters
the mobility. To accomplish this, the PCR product is denatured and then run in a native gel system
without reannealing. Each strand then forms hairpins as available from its sequence. Virtually any base
change alters the potential to form some hairpins and causes a mobility shift on the gel. This is called a
Single Strand Conformational Polymorphism (SSCP). In addition to screening exons for novel mutations,
this method can also be used to create a genetic marker where no more convenient method exists. The
fragments analyzed for SSCP have to be relatively small (~100 bases), because larger fragments form
complex patterns by falling into multiple secondary struture patterns.
Ref: Orita, M., Iwahana, H., Hayashi, K., and Sekiya, T. 1989. Detection of polymorphisms of human
DNA by gel electrophoresis as single-strand conformation polymorphisms. PNAS 86: 2766-2770.
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To hunt for a specific mutation, PCR products can be hybridized to Allele-Specific Oligonucleotides
(ASOs). Note below that allele-specific probes have been introduced into Real Time PCR.
Fidelity Problem
The original thermostable polymerase (Taq polymerase) lacked a 3' exonuclease activity (for proof
reading) and therefore has a poor fidelity of copying. This lack of fidelity is compounded by the multiple
rounds of synthesis. The error rate can exceed 1/100 bp, but more typically it should be around 1/3000 bp
(after 20 cycles).
Several precautions are available to avoid fidelity problems:
1) New thermostable DNA polymerases are now on the market that do proofread. However, their fidelity
turns out to only be about 10 x better than Taq polymerase. Also, they tend to nibble at the primers,
so longer primers are needed to compensate.
2) The PCR product can be sequenced without cloning. That way the individual errors in the different
molecules average out.
3) If the PCR product is cloned, then two different clones from different amplifications should be
sequenced.
Physical mapping with PCR
Ref. Olson et al., Science 245, 1434 (1989)
Any pair of PCR primers that amplify a unique sequence out of a genomic clone serve to identify that
sequence in any other genomic clone. Such a sequence is called a sequence tagged site (STS). If one
investigator publishes the sequence of a STS from his clone, another investigator can synthesize the
primers and tell if his clone overlaps without exchanging any materials. YAC contigs are primarily put
together by determining the overlap of STS's. A YAC contig covering the complete human Y
chromosome has been put together in this way (Foote et al, Science 258:60-66 (1992). A very small
sample appears below:
The same STSs were screened against the DNA from people with naturally occurring deletions of
part of the Y chromosome. This allows correlation between the genetic map defined by these deletions
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and the physical map defined by the YAC contig. STSs are similarly used to correlate YAC contigs with
deletion maps originating in interspecific somatic cell hybrids.
STS content can detect chimeric inserts. If an STS appears in a clone that can't possibly be there
based on clones that it overlaps, then that segment is probably chimeric. The best way to detect multiple
inserts in a large insert library is to make an STS from the sequence at the extreme end of each clone and
then see if that STS is detected in all overlapping clones.
Recovering DNA next to a known sequence (anchored PCR).
There are a variety of methods to do anchored PCR from double stranded DNA. The object is to
recover DNA next to the sequence that you know, without knowledge of any sequence on the other side.
Most methods involve ligating something to the end of a fragment to act as the second priming site. The
key problem is to avoid end-to-end amplification of all of the fragments.
Bubble PCR
One can ligate on a pair of oligonucleotides with a noncomplementary bubble, which is designed
to support amplification only on fragments where the gene specific internal primer has primed.
Inverted PCR
In inverted PCR, the fragments are first circularized and then recleaved at a different site to move
the unknown region between two primers.
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Using the vector.
One can ligate the fragments into a vector and then use an insert specific primer with vector
primer.
Recombination Mapping
Arbitrary Primed PCR (AP-PCR) also known as Random Amplified Polymorphic DNA (RAPD).
Ref: Welsh, J., & McClelland, M. 1990. Fingerprinting genomes using PCR with arbitrary primers. Nucl.
Acids Res. 18:7213-7218.
When genomic DNA is amplified with a PCR primer of 9 or 10 bases and an arbitrary sequence, a
pattern of products is amplified by chance complementarity. Some of these products are polymorphic. By
working through a collection of such arbitrary primers, a recombination map can be constructed with these
markers. This method is often used to gain a preliminary genetic map of an organism for which there is
no prior characterization.
Simple Sequence Length Polymorphisms (SSLPs)
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If the PCR product for a STS contains an internal tandem repetitive sequence, the length of the
PCR product will be polymorphic. In this case the PCR product can be used as a genetic marker, and
placed on the genetic map along with polymorphic genes, RFLPs, and disease loci. These genetic markers
are favored by human geneticists because they have a large number of alleles. They are variously called
Simple Sequence Length Polymorphisms (SSLPs), minisatelite repeats, microsatelite repeats, VNTRs,
dinucleotide repeats, & CA repeats. A typical case is shown below:
Stuttering problem
Each individual allele in the figure above appears not as a single size, as you would expect, but as
a small cluster of bands separated by 2 bp in size. This is because the polymerase is able to allow the
polymerizing strand to slip on the template during polymerization through the tandem repeat. This
problem is sometimes referred to as "stuttering". Stuttering is tolerated in the above experiment as long as
one can tell the alleles apart.
If one attempted to sequence a PCR product of the kind illustrated above, the sequence would
become unreadable as one passed through the tandem repeat. Given the option, one should try not to put
simple tandem sequences between the PCR primers (eg. poly(A) stretches). Alternatively, one could
clone the amplified fragment, and thus purify a homogeneous sequence.
Recovery of homologous sequences
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PCR can be used to amplify any sequences straight out of the genome for which enough
information is available to make the primers. This might be done to recover different alleles of the same
gene, different gene family members, or the homologous genes from a different organism.
Since the PCR primers may encounter mismatches on the intended priming sites, it is wise to use a
longer primer and to prime at a reduced stringency. This, however, promotes the production of extraneous
products. Consequently, one needs to be prepared to identify the correct product among a variety of
different products amplified. This can be done by predicting the size of the correct product, predicting a
restriction site found within it, or probing the products with an oligonucleotide probe designed to
hybridize between the two primers.
Differential Display PCR
Ref: Liang, P., Averboukh, L., and Pardee, A.B. 1993. Distribution and cloning of eukaryotic mRNAs by
means of differential display: refinements and optimization. Nucl. Acids Res. 21(14)3269-3275.
This method is now often used instead of subtraction libraries to find cDNAs that are differentially
induced in a tissue or cell line. It is an example of arbitrarily primed PCR. If RT PCR is conducted on an
mRNA sample using an arbitrary 10 base primer paired with oligo dT, chance matching of the arbitrary
primer to some RNAs will produce a pattern of bands on a gel. If several reactions are conducted with
different arbitrary primers, one can hope that all of the mRNAs are represented by a band somewhere on
the gel.
If the same set of reactions is conducted with RNA from two tissues, or from cells in a different
state of induction, one can hope that those RNAs that are differentially expressed will appear as a band
that is present on one pattern and absent from the other. This band can be rescued by PCR and cloned for
subsequent analysis. In practice, about 50% of rescued clones turn out to really represent differentially
expressed RNA.
PCR to Recover cDNA after Hybridization to Genomic DNA
Ref: Parimoo et al., PNAS 88, 9623-9627 (1991)
The following procedure uses PCR to rescue the small amount of cDNA that is bound to a filter as
a result of having used it as a hybridization probe.
The purpose of the procedure is to start with a mixture of total cDNA from some tissue, and then
to select those molecules that hybridize to a given YAC. The YAC DNA is fixed to a filter, and the
cDNA will be physically selected by hybridization to the filter, then melted back off. Before hybridization
all of the cDNA molecules will have PCR priming sites added to their ends so that any molecules
recovered can be PCR amplified. The priming sites could be added by ligating linkers onto a freshly made
cDNA; however, more commonly the cDNA has already been cloned and one uses the vector priming
sites.
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This is usually done with a library of small cDNAs (so that they will amplify well). One has to expect to
resort to the methods described above to recover extended cDNA sequences.
Quantitative RT (Reverse Transcription) PCR
The intensity of a PCR product can be used to estimate the amount of the original template. One must
measure the amount while it is still in the log-linear portion of the reaction. Originally one measured
incorporation of a radioactive primer in a series of amplifications differing in the number of cycles. The
advent of Real Time fluorescence PCR (see below) has made quantitative PCR much more accessible.
Often this method is used to find the ratio of an inducible RNA to a constitutive RNA in a sample. There
are a variety of strategies to conduct the reverse transcription reaction followed by PCR in a single
reaction tube. One of these is to use an enzyme (rTth) that is both a thermostable reverse transcriptase and
a thermostable DNA polymerase.
Real Time PCR.
In Real Time PCR, the thermocycler is equipped with a fluorescence detector to monitor the buildup of
product. Our department has purchased an ABI 7500 Real Time PCR machine that can simultaneously
amplify and monitor 96 samples. Training is required before you use this instrument. See Dr. Kathy
Howard.
Detection methods.
A method that is straight forward, but lacks specificity for the target sequence, is to incorporating a
fluorescent dye into the reaction. Although some initial real time experiments employed ethidium
bromide for detection, now a more sensitive fluorophore named SYBR Green is used for this purpose.
The dye fluoresces after associating with double stranded DNA, so it directly detects the build up of
product. Since this detection method is not sequence specific, one is vulnerable to being mislead by
amplifcation of spurious products. The instrument can be programmed to do a denaturation curve on the
sample after amplification, which may provide a warning that the amplified product is heterogeneous.
The available sequence specific detection methods involve an oligo probe with a fluorophore on one end
and a quencher on the other. These probes are added in addition to the two PCR primers, and are
designed to fluoresce if they can hybridize to the accumulating PCR product. In order to prevent the
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probe from acting as a primer, they are phosphorylated at the 3' end. There are several variations on two
general formats:
One type of probe fluoresces directly upon hybridizaiton. The molecular beacon web site has extensive
documentation: ( http://www.molecular-beacons.org/). The oligos with a fluophore on one end and a
quencher on the other has the ends in a hairpin so that usually the quencher is brought close to the
fluorophore and quenches it. If the oligo is hybridized to another nucleic acid, the quencher is separated
from the fluorophore and fluorescence increases. Hybridization is therefore detected directly by
fluorescence without having to remove the unhybridized oligo. This kind of probe can also be used for
quantitative in-solution hybridization or as a substrate on a microchip or microarray.
Another type of probe is called a hydrolysis probe (TaqMan). These are also called "fluorogenic
probes". The fluorophore is at the 5' end and the quencher at the 3' end. In this case the probe is fully
complementary to the intended product and is short enough that fluorescence is quenched as long as the
probe is intact. This probe hybridizes to the template in the path of the polymerase. Upon colliding with
the probe, the polymerase hydrolyzes the 5' nucleotides releasing the fluorophore which is now
unquenched. When designing hydrolysis probes, one is advised to give it an extra 5oC over the TM of the
primers to account for any destabilizing effect of the dyes. Some of the available fluorophores are
quenched by guanine, so one is advised to not put a G at the 3' end of the probe. Also one is advised to
choose the strand that puts more C's than G's in the probe. A common strategy to avoid detection of
genomic DNA during a reverse-transcription PCR assay is to make the probe cross an intron exon
boundary. Finally, more than one probe with different fluorophores may be used to detect the same
amplicon.
Mutation detection: In the hydrolysis probe method, the probe oligo can be designed such that a single
nucleotide mismatch will prevent hybridization. Hence the probe could distinguish a one base difference
in a sequence.
Multiple fluorophore detection: The instrument simultaneously monitors 5 different wavelengths
corresponding to 5 commonly used fluorophores. In principle, one could have 5 differently colored
probes in each reaction. In practice, there should probably not be more than two differently colored
probes in a reaction.
Quantification: For quantification, typically quadruplicate samples are amplified for each unknown.
Reactions that fail to follow a log-linear accumulation are thrown out, and the rest are averaged to find the
fractional cycle number (Ct) that the reaction crosses a threshold within the log linear part of the
accumulation. Ct will have a log-linear relationship with the initial concentration of the template in each
sample. Consider a standard curve created by amplifying a series of samples composed by serial dilution
from a stock solution of known concentration. Such a stock can be created by quantifying the product of a
prior PCR amplification. The instrument will plot the standard curve as Ct versus log concentration. This
should ideally produce a straight line with a slope of -3.3 (= -1/log10(2)). Amplifying an unknown and
finding its initial concentration from this curve would be a very tight way to calculate the intial
concentration. The standard curve would automatically compensate for any inefficiency in preparing the
sample (eg. the efficiency of a reverse transcription step), and it would provide a true slope to use if the
amplification was proceeding with less than an 2 fold increase on each cycle. The software also reports
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deviation from ideal amplification as the efficiency of amplification. An efficiency of less than 0.9 means
that the assay is significantly suboptimal in some parameter such as annealing temperature, primer selfannealing, or primer dimer formation.
When comparing the RNA level between two different preparations, a major risk is that the samples might
not have been prepared comparably. For example, a different number of cells may have been included. A
notorious problem for RNA is that a differential amount of RNA degradation may have occurred. So,
particularly for measuring levels of RNA, it is customary to also measure the Ct for another molecule that
is believed to be expressed the same in both samples. The signal of interest can then be normalized by
this reference signal. To be especially careful, you might do a dilution series on one of the samples and
plot the Ct vs. the dilution factor to reveal if the true slope for either the target or the reference RNA
deviated from the ideal -3.3 and take that into account.
The Real Time PCR machine has software to assist in these analyses and is accompanied by substantial
documentation. Hydrolysis probes are designed similarly to PCR primers with respect to the Tm, but
have the additional constraints that they should not have a C at the 3' end, and should have more C's than
G's.
Some resources for real time PCR:


Tevfik Dorak's Real Time PCR page: http://dorakmt.tripod.com/genetics/realtime.html
Lunge VR, Miller BJ, Livak KJ, Batt CA. 2002. Factors affecting the performance of 5' nuclease PCR
assays for Listeria monocytogenes detection. J. Microbiol. Methods. 51:361-8.
Study Questions
1) For perfectly matched PCR primers, it generally isn't considered of any benefit to make them longer
than about 30 bases. Why?
2) Does the amount of PCR product at the end of the amplification reflect the amount of original template
in the sample? What conditions must be met for the amount of PCR product to reflect the amount
of the original template?
3) If you knew exactly 30 bases of sequence with which to recover a cDNA, would you make the PCR
primer 30 bases long, or only 20?
4) Overly stable PCR primers often raise a background of nonspecific products, but over stable
fluorogenic probes don't cause any particular problem. Why?
5) A real time reverse transcriptase/PCR assay for a message is designed with one gene-specific PCR
primer paired with oligo dT as the other PCR primer, and detection by SYBR Green. What is
wrong with this design?
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6) One PCR primer in a particular set is diagnosed as having a hairpin of marginal significance. What
symptoms might you expect if this hairpin causes a problem? How might conditions be adjusted to
overcome the problem?
Last update 3/01/05 - Steve Hardies
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