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Research article
A comprehensive collection of experimentally validated primers for Polymerase Chain Reaction quantitation of murine transcript abundance
Athanasia Spandidos, Xiaowei Wang, Huajun Wang, Stefan Dragnev, Tara Thurber and Brian Seed*
Corresponding author:
Center for Computational and Integrative Biology, Massachusetts General Hospital. MA, USA
Department of Genetics, Harvard Medical School, 185 Cambridge Street, Boston, MA , USA
Division of Bioinformatics and Outcomes Research, Department of Radiation Oncology, Washington University School of Medicine, 4921 Parkview Place, St. Louis, MO 63110, USA
Idearc Media Corp, 1601 Trapelo Road, Waltham, MA 02451, USA
For all author emails, please .
BMC Genomics 2008, 9:633&
doi:10.64-9-633
The electronic version of this article is the complete one and can be found online at:
Received:19 September 2008
Accepted:24 December 2008
Published:24 December 2008
& 2008 S licensee BioMed Central Ltd.
This is an Open Access article distributed under the terms of the Creative Commons Attribution License (), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.
Background
Quantitative polymerase chain reaction (QPCR) is a widely applied analytical method
for the accurate determination of transcript abundance. Primers for QPCR have been
designed on a genomic scale but non-specific amplification of non-target genes has
frequently been a problem. Although several online databases have been created for
the storage and retrieval of experimentally validated primers, only a few thousand
primer pairs are currently present in existing databases and the primers are not designed
for use under a common PCR thermal profile.
We previously reported the implementation of an algorithm to predict PCR primers for
most known human and mouse genes. We now report the use of that resource to identify
17483 pairs of primers that have been experimentally verified to amplify unique sequences
corresponding to distinct murine transcripts. The primer pairs have been validated
by gel electrophoresis, DNA sequence analysis and thermal denaturation profile. In
addition to the validation studies, we have determined the uniformity of amplification
using the primers and the technical reproducibility of the QPCR reaction using the
popular and inexpensive SYBR Green I detection method.
Conclusion
We have identified an experimentally validated collection of murine primer pairs for
PCR and QPCR which can be used under a common PCR thermal profile, allowing the evaluation
of transcript abundance of a large number of genes in parallel. This feature is increasingly
attractive for confirming and/or making more precise data trends observed from experiments
performed with DNA microarrays.
Background
Quantitative polymerase chain reaction (QPCR) has become a widely applied technique
for quantitative gene expression analysis [,]. The technique is frequently used to validate and improve the precision of measurement
of differences in transcript abundance detected by DNA microarray experiments []. In QPCR, product formation is monitored at the end of each thermal cycle by determining
the strength of a fluorescent signal that is proportional to the amount of product
[,]; QPCR thus provides more information than can be inferred from signal detected at
the end of multiple cycles of reaction, as in conventional PCR analysis [-]. Because data can be collected from the exponential phase of the reaction a generally
reliable quantitation of target DNA concentration can be achieved []. Detection of QPCR product concentration is usually accomplished by one of two general
fluorescence-based approaches: the measurement of a target sequence-selective signal
arising from a conformational change in a labeled primer, or the measurement of total
DNA formed during the reaction. In the former method, target-specific probes containing
fluorophores, such as hydrolysis probes [-], dual hybridization probes [], molecular beacons [] or scorpions [,] are designed. These detection systems provide partial protection against the risk
of generation of signals from off-target amplicons but the primers are considerably
more expensive to generate than conventional unlabeled primers. In a more widely practiced
variant of QPCR, sequence non-selective fluorescent dyes that bind to double-stranded
DNA, such as SYBR Green I, are used [,]. The quantum yield of SYBR Green I dye intercalated into double-stranded DNA is much
greater than the quantum yield of free dye, leading to an increase in fluorescence
intensity that, at saturating dye concentration, is proportional to DNA concentration
[]. This yields a simple inexpensive way to measure product amplicon formation. However,
the contribution of fluorescence from DNA arising by amplification of undesired sequences
cannot be determined without some additional measure, such as thermal dissociation
analysis [].
Several online resources have been described that can be used to design primers for
PCR and QPCR [-] and are useful for gene expression analysis, when a small number of genes are of
interest. We have previously described a resource of designed primers that can be
used for real-time PCR with sequence independent detection methods, such as SYBR Green
I detection, and that can work under a common PCR thermal profile []. Amplification of undesired sequences is a common problem in QPCR, and poses greater
difficulties when the amplification conditions cannot be tailored to the primer pair
of interest, as for example would be the case for massively parallel QPCR. The primer
design algorithm used for the selection of primers for this study was based on a previous
approach to the prediction of oligonucleotides for the study of protein coding regions
by microarrays [], but differed by the addition of filters thought to be important for PCR primer specificity.
Primers were designed from cDNA sequence information and the principal filter for
cross-reactivity was the rejection of primers containing contiguous residues (15 bases
or longer) present in other sequences []. Additionally, the selected primer pairs had no self-complementarity, low 3' end
stability and high complexity. Low complexity regions may contribute to primer cross-reactivity
[], so they were excluded using the DUST program []. The primer Tms were in the same range, as well as their GC contents. Short amplicons (60–350 bp)
were favored during primer selection, but in some cases 100–800 bp amplicons were
also considered when the design criteria could not be met for shorter amplicons.
The collection of designed primer pairs has been deposited in a public resource called
PrimerBank []. PrimerBank
contains primers for most known human and mouse genes (Table ). The primers designed for the mouse genome cover 27684 genes, but because of some
redundancy – one primer pair can represent multiple genes, in most cases isoforms
– only 26855 primer pairs were synthesized to represent once each of these 27684 genes
(Table ). For another 1165 mouse genes, it was not possible to design primers, mainly due
to low sequence quality. The average sequence length for these genes, the majority
of which are 'unknown' or RIKEN sequences, is 435 bp while the average mouse gene
has 1293 bp. All primers have been designed to have uniform properties and work using
the same PCR conditions which simplifies analyzing the expression of many genes in
parallel by QPCR.
Statistics of primers contained in the PrimerBank database.
PrimerBank primer design and gene representation.
Previously we tested by conventional and QPCR 112 primer pairs from PrimerBank representing
108 genes []. These primers amplified successfully and specifically the genes for which they had
been designed, even though some genes were from closely related gene families. As
a second step, we tested by QPCR 26855 PrimerBank mouse primer pairs, representing
most known mouse genes, in order to determine if they can successfully amplify the
genes for which they had been designed. From the experimental validation procedure,
we identified 17483 pairs of primers that amplify unique sequences corresponding to
distinct murine transcripts. We also validated on genomic DNA some of the primer pairs
that initially failed by QPCR, to provide explanations for these failures. We determined
the uniformity of amplification using 96 PrimerBank primer pairs, and the technical
reproducibility of the QPCRs, using the same primer pairs. In addition, SYBR Green
I sequence specificity was investigated, using a set of sequences differing in length
and base composition. Successful primer pair information is now freely available from
the PrimerBank database together with the experimental validation data (Figure ). The mouse serves as an excellent model for studying the function of human genes
in vivo [] and currently more genomic resources exist for mouse compared to human. The experimental
validation of PrimerBank mouse primers can be applied to functional analysis of human
A screenshot of the web interface for PrimerBank. Several primer search terms can be used, such as: GenBank accession number, NCBI
protein accession number, NCBI gene ID, PrimerBank ID, NCBI gene symbol or gene description
(keyword). Website:
High-throughput primer validation procedure
A collection of primer pairs from PrimerBank covering most known mouse genes was tested
by QPCR, agarose gel electrophoresis, sequencing and BLAST. An overview of the procedure
used for primer validation can be seen in Figure . Universal mouse total RNA was reverse transcribed using random hexamers and the
cDNA was used as a template. 26855 primer pairs, corresponding to 27684 transcripts,
were tested by QPCR and the amplification plots and dissociation curves were analyzed.
The same PCR conditions were used for all reactions. PCR amplification plots indicate
SYBR Green I fluorescence which is proportional to PCR product formation. Dissociation
curves indicate the loss of SYBR Green I fluorescence as the PCR product duplex dissociates.
Tm and the shape of the dissociation curve are a function of GC content, sequence and
length [,]. From the amplification plots, PCR products appeared typically between 19 and 27
cycles of PCR, with a small variation of 1 or 2 cycles depending on the length of
the PCR product and thus the amount of SYBR Green I bound to it. As a general observation,
most shorter length products (from 60 bp) appeared between 20 and 27 cycles and their
Tms were between 75°C and 85°C, and most longer length products (&200 bp) appeared between
17 and 27 cycles and their Tms were between 80°C and 90°C.
Summary of procedure for experimental validation of PrimerBank mouse primers.
Agarose gel electrophoresis was used to confirm the correct size of the PCR product,
and sequencing and BLAST were used to confirm that the expected transcript had been
amplified. All successfully sequenced samples (24476) were BLAST analyzed. From the
primer validation procedure, primer pairs were grouped into successful or failed,
according to the analysis criteria. From 26855 primer pairs tested 1%) primer
pairs, corresponding to 18324 transcripts, were found to be successful by QPCR, agarose
gel, sequencing and BLAST analysis. 2%) primer pairs were successful based
on agarose gel electrophoresis analysis and 1%) primer pairs were successful
based on BLAST analysis. Primer pairs which failed based on the experimental validation
procedure can be grouped into various types. Table
presents a classification of the types of failures. In a few cases (less than 0.8%),
primer pairs were found to be successful based on the gel or BLAST analysis criteria,
but no amplification could be detected with SYBR Green I. Sequencing can be very sensitive
and a low abundance amplicon can thus be sequenced successfully despite low amounts.
Also, in many cases where PCR products were short (~60–80 bp) it was not possible
to obtain sequencing information for these samples.
Classification of failed PrimerBank primer pairs.
A few representative examples of primer pairs are described [see Additional files
, , , , ], to demonstrate in detail the analysis of the results generated from the high-throughput
primer validation procedure. Data are shown for five successful primer pairs, five
primer pairs that failed based on agarose gel electrophoresis analysis and five primer
pairs that failed based on BLAST analysis. Information on these primer pairs, such
as PrimerBank IDs, primer sequences and amplicon lengths, is shown here [see Additional
file ]. More information on these primers, such as their Tm and location on the gene, can be found in PrimerBank, as well as alternative primer
pairs designed for these transcripts.
Additional file 1. Five representative examples of primer pairs that were successful throughout the validation
procedure.
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Additional file 2. Five representative examples of primer pairs that failed based on agarose gel analysis.
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Additional file 3. Five representative examples of primer pairs that failed based on BLAST analysis.
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Additional file 4. Information for mouse primer pairs from PrimerBank tested using QPCR.
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Additional file 5. NCBI BLAST analysis of successfully sequenced PCR products.
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PrimerBank user interface
All data generated from the high-throughput primer validation procedure can be freely
accessed from PrimerBank
. See Figure
for the PrimerBank homepage. Users can search the PrimerBank database for primers
for their gene of interest using several search terms such as: GenBank accession number,
NCBI protein accession number, NCBI gene ID, PrimerBank ID, NCBI gene symbol or gene
description (keyword). Search results include primer sequences together with some
information about the primers, such as expected amplicon size and Tm. cDNA and amplicon sequences, and validation data can be viewed by clicking on the
appropriate links. All validation data can be accessed from PrimerBank, since the
validation criteria may be different from the criteria of the users. Also, users can
use a BLAST tool found on the PrimerBank homepage (see Figure ), to find any primers contained in the PrimerBank database that would amplify their
sequence of interest. A BLAST tool for the PCR product sequence obtained from the
validation procedure can be used to query the NCBI database and this can be found
on the validation data webpage. The QPCR and reverse transcription protocols can be
found on PrimerBank, as well as a troubleshooting guide.
Analysis of failed primer pairs
A schematic representation of the agarose gel fail distribution can be seen in Figure
. This analysis was based on determining whether one PCR product of the correct size
could be visualized from agarose gel electrophoresis data. Most primer pairs were
successful based on at least one step of the primer validation procedure. Two major
types of failed primer pairs that comprise most of the failures are primer pairs that
failed on agarose gels but were successful by BLAST and primer pairs that failed on
BLAST but were successful on agarose gels. 3695 primer pairs failed based on BLAST
analysis alone and another 1864 primer pairs failed based on agarose gel analysis
alone. In most cases a primer pair failed in one of the analysis steps based on the
criteria, but was successful in other analysis steps. The failed samples did not overlap
in many cases and this could have been in some cases due to strict BLAST analysis
criteria and new splice isoforms seen on the agarose gels. Also, some primer pairs
failed by both BLAST and agarose gel analysis, although these are numerically minor.
For a detailed description of the analysis criteria see Table . The criteria for success or fail may be different from the criteria users might
apply and for this reason all validation data can be accessed from PrimerBank.
Distribution of agarose gel failures. Multiple amplification visualized as two or more bands on the gel accounted for
46.7% of the failed samples. Undesired amplification visualized as the wrong size
bands on the gel accounted for 13.8% of the failed samples. Poor amplification visualized
as a faint band on the gel was observed in 4.8% of the failed samples and no amplification
took place in 34.7% of the failed samples.
From the total agarose gel failed reactions, 46.7% were due to multiple amplification
products apparent by gel electrophoresis. 13.8% of the total failed reactions were
due to undesired amplification, seen as the wrong size band on the gel. 4.8% of the
total failed reactions were due to poor amplification, and 34.7% of the total failed
reactions were due to no amplification taking place. Multiple or undesired amplifications
accounted for the majority (60.5%) of the agarose gel failed reactions. These may
represent undocumented transcripts or splice isoforms that could have been amplified
in addition to or instead of the expected transcripts. For the reactions that failed
because no amplification had taken place, the template sequences may not have been
present or present in very low copy number.
Validation of primer pairs that failed amplification using genomic DNA
From the high-throughput PrimerBank mouse primer pair validation, 1745 samples (6.5%)
failed because of no amplification, as seen from the QPCR amplification plots. From
the gene description information we found several to belong to olfactory receptors,
vomeronasal receptors, transcription factors and low abundance transcripts while others
were of unknown function or RIKEN sequences (data not shown). In order to investigate
the possibility that the templates for the failed amplification primer pairs were
not expressed in the cDNA sample used, we repeated these reactions using genomic DNA
as a template. It can be difficult to achieve amplification using genomic DNA as template
in general, due to its complexity. However, it can be used successfully if technical
difficulties are overcome and can be useful as a universal template as it contains
a copy of all genes, and the same amount of template is present for all single-copy
genes []. We have found that enzymatic digestion (such as EcoRI/BamHI digestion used here) can be used for reduction of the complexity of the DNA and
thus higher amplification rates. We matched 864 primer pairs to mouse genome sequences
obtained from the UCSC genome browser. The remainder of the sequences could not be
matched, probably because they were located on exon junctions. 640 of these primer
pairs have no EcoRI/BamHI restriction sites in their expected PCR amplicons, and were used with EcoRI/BamHI digested DNA template to prepare the validation reactions. We tested 192 representative
samples, from the 1745 total number of failed primer pair samples, whose expected
PCR amplicon lengths range from 60 bp to 123 bp and whose amplicons have no EcoRI/BamHI restriction sites. 50 ng EcoRI/BamHI digested 129 mouse ES cell genomic DNA was used per 25 μl PCR reaction.
The amplification plots of all 192 samples (2 × 96 well plates) are shown here [see
Additional files , ]. The success rate of QPCR based on the amplification plots was high: 88.5% for the
first plate [see Additional file ] and 90.6% for the second plate [see Additional file ]. However, Ct values differed significantly, from roughly 23 to 40 [see Additional
files , ]. The location of the reactions on the plate did not explain this variation. The
samples were also analyzed by agarose gel electrophoresis and sequenced (data not
shown). Sequences obtained were BLAST analyzed and matched to the expected sequences,
confirming that the correct templates had been amplified (data not shown). Therefore,
these primer pairs had originally failed because their respective templates were not
present in the cDNA sample used and not because of poor primer design, in general.
Additional file 6. Validation of 96 PrimerBank primer pairs which had failed QPCR during the high-throughput
validation procedure.
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Additional file 7. Validation of 96 PrimerBank primer pairs which had failed QPCR during the high-throughput
validation procedure.
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Uniformity of amplification and technical replicate tests
We next set out to determine the uniformity of amplification using fully validated
PrimerBank primer pairs ie. primer pairs that had been successful in all steps of
the validation procedure. 96 primer pairs were chosen with expected PCR amplicon length
ranging from 80 bp to 120 bp and containing no EcoRI/BamHI restriction sites in their sequences. Both forward and reverse primers were chosen
to be on the same exon in order to amplify the same template on genomic DNA. EcoRI/BamHI digested 129 mouse ES cell genomic DNA was used as template. After digestion the
DNA was purified for PCR by phenol extraction and ethanol/salt precipitation. 50 ng
of DNA template was used per 25 μl PCR reaction, which was found by optimization experiments
to give a reasonable Ct value.
See Figure
for the amplification plots and dissociation curves. As can be seen from Figure , the Ct values for each sample are not exactly the same. This is expected since there
will be some stochastic variation. Also, different primer pairs were used for each
sample. However, the Ct values are similar, so amplification using PrimerBank primers
appears to be relatively uniform. The statistical significance of the difference in
Cts observed was determined by plotting a frequency distribution of the number of
samples versus the Ct (Figure ). A statistical normality test was also used for the analysis of these Ct values,
but the data did not pass this test. The effect of primer length and primer GC% on
the Ct was studied, by plotting these values against the Ct, and no correlation between
these parameters was found (see Figure ). The effect of the PCR product Tm on the Ct was also studied, by plotting the Tm values against the Ct, and again no correlation was found (see Figure ). Since the expected PCR product size varies from 80 bp to 120 bp, some small variation
in Tm is expected, and this can be seen from the dissociation curve data (see Figure ). The Tm data (obtained from the dissociation curves) was also plotted as a frequency distribution
and did not pass the statistical normality test (data not shown).
Uniformity of amplification test using 96 PrimerBank primer pairs. A. PCR amplification plots. B. Dissociation curves plotted as the raw fluorescence
with respect to temperature. Expected PCR product lengths range from 80–120 bp.
Analysis of uniformity of amplification test. A. Ct frequency distribution. B. Correlation of Ct to total primer length, R: 0.08.
C. Correlation of Ct to GC%, R: -0.12. D. Correlation of Ct to Tm, R: -0.29.
In order to determine the technical reproducibility of the QPCRs, five 96 well plate
assays were prepared using the same technical procedure. Reactions were set up using
the same 96 primer pairs and DNA template (129 mouse ES cell EcoRI/BamHI digested genomic DNA) that were used for the uniformity of amplification test.
The coefficients of variation for each 96 well plate assay are all & 0.1 and the average
coefficient of variation for all assays is 0.07 [see Additional file ]. The individual primer pair Cts for each 96 well plate assay and coefficients of
variation are shown here [see Additional file ]. Ct data from each assay initially did not pass the statistical normality test.
The Ct values were normalized, using the formula:
Additional file 8. Analysis of technical replicate experiments.
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Additional file 9. Analysis of individual primer pairs from technical replicate experiments.
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(LnCt - LnCtav)/SD,
where LnCt is the natural logarithm of the Ct value used, LnCtav is the natural logarithm of the average Ct value of the assay and SD is the standard
deviation of the LnCt values for each assay, and outliers were removed. The normalized
data passed the normality test, so the data appear to be log normal. The plots of
the frequency distributions of the log normal data are shown here [see Additional
Additional file 10. Frequency distributions of log normal data from five technical replicate tests.
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Analysis of pipetting variation during liquid transfer of the fluid handling system
was carried out and the transfer efficiency of the robot was found to be 97.3% [see
Additional file ]. The data from the liquid transfer test passed the statistical normality test only
after the 9 lowest value outliers were removed (data not shown), but the coefficients
of variation are low (less than 0.03) [see Additional file ]. Variation in liquid transfer can only account for a small amount of the variation
observed in QPCR reactions, and hence other factors must be responsible for the differences
observed in Ct values.
Additional file 11. Comparison of pipetting variation between manual and robotic liquid transfer.
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SYBR Green I sequence specificity
The SYBR Green I dye has been widely used as a non-sequence specific dye for fluorescence
detection of QPCR products []. Studies of SYBR Green I-DNA binding showing some sequence specificity of the dye
have been reported but these have not been conclusive [,,]. We investigated whether SYBR Green I is sequence specific by adding the dye to a
series of amplicons and taking fluorescence readings. 8 amplicons of increasing length
and 7 amplicons of increasing AT% [see Additional file ] were used, whose concentrations were accurately determined (see methods). From these
experiments, we did not observe any length dependent or AT/GC dependent sequence specificity
of SYBR Green I [see Additional file ]. However, we cannot exclude the possibility that SYBR Green I can show specificity
to sequences such as homopolymer regions of DNA [] or specific sequences. We also investigated whether SYBR Green I dye binding is sequence
specific by estimating the number of PCR product molecules at threshold using the
ABI PRISM 7000 Sequence Detection System (Applied Biosystems) [,]. For this, the same 14 amplicons as above were used and a template titration series
of reactions was prepared for each amplicon. SYBR Green I threshold cycle (Ct) fluorescence
will be the same for all amplicons (and all reactions), since the same threshold was
used to compare all reactions. However, if SYBR Green I is sequence specific, this
fluorescence will correspond to a different number of molecules at threshold for each
amplicon. These experiments were inconclusive, as the stochastic error was too large
to be able to accurately determine the molecules detected at the threshold (data not
Additional file 12. Amplicons used for SYBR Green I sequence specificity experiments.
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Additional file 13. SYBR Green I binding to dsDNA of increasing length and AT%.
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Estimation of QPCR amplification efficiency
The most common method for the calculation of the amplification efficiency of a QPCR
reaction requires preparation of a series of serial dilutions of the sample and creation
of a standard curve, whereby efficiency is estimated from the slope of the standard
curve [,]. However, this method does not provide an accurate value of the efficiency, as the
efficiency can vary between different reactions and as input concentration changes.
A number of analytical methods have been described for the calculation of the amplification
efficiency of a reaction from single reaction kinetics [] (for a correction in equation 3 of this paper see: []), [-]. These methods can be more accurate and, when automated, less laborious compared
to the standard curve method []. Using the following analytical method, we estimated the amplification efficiency
values for 13 QPCRs using PrimerBank primer pairs that had been previously used. The
log2 fluorescence data was plotted versus the Ct number and the slope of the linear
regression was taken to be equal to the efficiency of each reaction [see Additional
file ]. Cycle values closest to the Ct were used, as this region will be the most accurate.
The efficiency values ranged from 79% to 96% [see Additional file ]. Replicates can be used to improve accuracy when using either the standard curve
or analytical single reaction kinetics methods [,].
Additional file 14. Amplification efficiency estimation from single reaction kinetics data.
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We compared amplification efficiency estimation using the standard curve and analytical
methods in order to determine the accuracy of each method using the same 13 PrimerBank
primer pairs as above [see Additional file ]. Either the log2 of pg of input template DNA data, for the standard curve method,
or the log2 fluorescence data, for the analytical method, was plotted versus the Ct
number. Ct was the independent variable and log2 of pg of input template DNA/fluorescence
was the dependent variable. The slope of the linear regression was taken to be equal
to the efficiency of each reaction. From these results the analytical method shows
a smaller variance of efficiency values and the range is smaller compared to the standard
curve method [see Additional file ]. One-way ANOVA analysis was done to determine if amplification efficiency varied
significantly between different PrimerBank primer pairs, using each primer pair in
a series of titration reactions of template DNA [see Additional file ]. The average efficiency, standard deviation and coefficient of variation for each
group of primer pairs are shown here [see Additional file ]. The P value is & 0.05 (0.7338) therefore the amplification efficiency is similar
between these groups.
Additional file 15. Amplification efficiency estimation using analytical and standard curve methods.
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Additional file 16. One-way ANOVA test to determine if amplification efficiency varies significantly between
different PrimerBank primer pairs.
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Additional file 17. PrimerBank primer pair groups used for one-way ANOVA analysis.
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In order to account for sample effects, it is useful to provide a model of the experimental
measurement of fluorescent PCR product accumulation [-]. The following equations can be used:
Log2pgDNA = β0 + βCtxCt + ε,(1)
where Log2pgDNA is the dependent variable, β0 is the intercept, βCt is the regression coefficient for the x independent variable, and ε is the error.
Equation 1 can be used for the standard curve method.
Log2Fluorescence = β0 + βxxc + ε,(2)
where Log2Fluorescence is the dependent variable, β0 is the intercept, βx is the regression coefficient for the x independent variable of cycle c, and ε is
the error. If βx = 1, amplification efficiency is 100%. Equation 2 can be used for the analytical methods.
PrimerBank primer pair gene location
PrimerBank primer pairs have been designed irrespective of their location on exons.
Data from the UCSC genome browser were downloaded and used to find the location of
26854 mouse primer pairs with respect to exons (see Table ). 19668 primer pairs matched to sequences from the genome browser. Most of the matched
primer pairs (16356) are located within exons and at least one primer from the rest
of the primer pairs is located on an exon boundary. Primers can be designed to be
located on exon boundaries, in order to avoid non-specific amplification of genomic
DNA during PCR, but in many cases it was not possible to design primers located on
exon boundaries that fulfilled all of the criteria for primer design, most trivially
because some transcripts consist of a single exon.
Primer pair location with respect to exons.
Discussion
Source of DNA template
A commercial composite mouse RNA preparation was chosen as the source of DNA template
for QPCRs, which contains RNA from a panel of eleven different mouse cell types for
a good representation of the majority of mouse genes. The composite mouse RNA is composed
of total RNA from: whole embryo, embryonic fibroblasts, kidney, liver, lung, B-lymphocyte,
T-lymphocyte, mammary gland, muscle, skin and testis. The success rate of the high-throughput
PrimerBank primer validation experiments was high as seen both from agarose gel and
BLAST analysis. We validated some of the failed reactions using genomic DNA as template,
and found that most of the failures in which no PCR product had formed could be due
to very little or no cDNA present in the source of DNA template. In order to increase
amplification success, specific tissues may be used as sources of cDNA templates where
expression of the genes of interest is known.
Primer specificity
The PrimerBank primer design was based on a successful approach for the prediction
of oligonucleotides for the interrogation of protein coding regions by microarrays
[]. However the primer design differs by the addition of filters that are thought to
be important for primer specificity []. All primers have been designed to work using a relatively high annealing temperature
of 60°C and this temperature was used throughout the primer validation experiments
described here. High annealing temperatures help reduce non-specific amplification.
A high percentage of the total failed samples were due to undesired or multiple amplification,
however this may have been for other reasons such as new unidentified genes or splice
isoforms. In 3.9% of the cases where multiple bands could be seen on the agarose gel
and in 14.6% of the cases where bands of other than the expected size could be seen
on the agarose gel, no sequencing information was obtained. Also, 29.7% and 55.2%
respectively, did not match to the expected sequences by BLAST. So, sequence homology
existed in most cases of undesired or multiple amplification. From the genome-wide
primer validation experiments presented here, we have found a high success rate of
primer pairs that amplify the transcripts for which they had been designed. For primer
pairs that failed because no amplification could be detected, we found that the reason
for which they had initially failed was because their target sequences were not present
in the target cDNA used. Another reason for failure in the high-throughput validation
procedure, may be that protein coding genes in the human genome are fewer than previously
thought, and the same may apply to the mouse genome [].
A collection of potential new splice isoforms
As mentioned previously, larger than expected or multiple bands were visible on the
agarose gel for some samples, however, sequences for these matched confidently by
BLAST to the expected sequences. Therefore, the template sequences amplified in these
cases could be new genes or splice isoforms. These unrecognized genes or splice isoforms
may contribute to primer cross reactivity which results in a lower success rate on
the agarose gels. Good primer design depends on accurate genomic information about
genes and splice isoforms and it is suggested that many unidentified genes and splice
isoforms could exist. All primer pairs that failed because of non-specific amplification,
but when BLAST analyzed matched to the expected sequence, could have amplified new
non-identified isoforms. This information would be very useful for other researchers,
in addition to other strategies for identifying new genes and splice isoforms [,]. PrimerBank primers could also be used for determining copy-number variation of a
gene or splice isoform [,].
The PrimerBank database
Several online databases exist containing experimentally validated primers, however,
only a few thousand primer pairs are currently present in these databases [-]. We have previously designed PCR primers for the human and mouse genomes, which are
available from PrimerBank []. The PrimerBank database currently contains 306800 primers for the mouse and human
genomes and is tightly integrated with information from the NCBI databases. PrimerBank
has been designed so that researchers can search for primers for their gene of interest
using several search terms such as: GenBank accession number, NCBI protein accession
number, NCBI gene ID, PrimerBank ID, NCBI gene symbol or gene description (keyword).
Currently, all validated primers can be retrieved by searching PrimerBank. In many
cases, alternative primer pairs for genes also exist in PrimerBank. NCBI sequences
have been attached to the primer information page and NCBI LocusLink indices have
been used internally for gene locus mapping. All primers have uniform properties such
as Tm, length and GC content and can work using the same PCR conditions.
Conclusion
We tested by QPCR 26855 PrimerBank mouse primer pairs in order to determine if they
can successfully amplify the genes for which they had been designed. We identified
17483 primer pairs that amplify unique sequences that correspond to distinct murine
transcripts. All primers have been used under a common PCR thermal profile, allowing
the experimentally validated primer collection to be used to evaluate the transcript
abundance of a large number of genes in parallel. We used genomic DNA as a template
to validate primer pairs that had initially failed by QPCR and provided explanations
for the various modes of failure. We determined the uniformity of amplification of
the QPCRs using 96 PrimerBank primer pairs. From the uniformity experiments, we found
a small variation in Cts which could be due to differences in PCR product length and/or
stochastic variation. However, overall amplification appears to be uniform using PrimerBank
primers. We investigated the reproducibility of the QPCRs, using the same 96 primer
pairs that were used for the uniformity experiments, by comparing Ct values between
five technical replicate plates and found coefficients of variation to be low. In
addition, SYBR Green I sequence specificity was investigated, using a set of sequences
differing in length and base composition. We found no SYBR Green I specificity for
the sequences used, but cannot exclude SYBR Green I specificity towards specific sequence
motifs. Furthermore, we calculated the efficiency of the reactions from single reaction
kinetics data and found the estimated efficiencies to be within a reasonable range,
and also that the efficiency can vary between different templates. PrimerBank provides
a useful tool for quantitative gene expression analysis by QPCR and facilitates high-throughput
High-throughput primer validation procedure
Oligonucleotide synthesis
Oligonucleotides for QPCR were synthesized at Synthesis Core lab of Center for Computational
and Integrative Biology at Massachusetts General Hospital. The quality and quantity
of the synthesized oligonucleotides were determined by capillary elecrophoresis using
the MCE 2000 (CombiSep) instrument and by OD260 reading using the Spectra Max Plus
Spectrophotometer (Molecular Devices). Forward and reverse primer mixtures were normalized
to 2 μM of each primer for use in QPCR.
Preparation of cDNA sample
Universal Mouse Reference total RNA (Stratagene) was used for the preparation of the
cDNA sample. Reverse transcription using random hexamers was performed using the Superscript
First-Strand Synthesis System for RT-PCR (Invitrogen). Based on the recommended protocol,
20 μg of total RNA was used for each reaction and cDNA samples prepared were in a
final volume of 84 μl. The quality of the individual first strand cDNA preparations
was tested in a QPCR reaction using mouse actin primers (PrimerBank ID: ,
F: GGCTGTATTCCCCTCCATCG, R: CCAGTTGGTAACAATGCCATGT).
QPCRs were performed in polypropylene 96 well plates on the ABI PRISM 7000 Sequence
Detection System and ABI 7300 Real-Time PCR System (both from Applied Biosystems).
SYBR Green PCR Master mix (Applied Biosystems) or Absolute Q-PCR SYBR Green ROX mix
(ABgene) were used. For each reaction, 12.5 μl of the 2× SYBR Green PCR mix were added
to 2.5 μl of 2 μM forward and reverse primer mix (final concentration of each primer
is 200 nM), 1 μl of cDNA and made to 25 μl with water. The Biomek FX Laboratory Automation
Workstation (Beckman Coulter), as well as manual pipetting, was used to prepare the
reactions. PCR conditions used were the following: 50°C for 2 minutes (step 1), 95°C
for 10 minutes (for Applied Biosystems PCR mix) or for 15 minutes (for ABgene PCR
mix) (step 2), 95°C for 15 seconds, 60°C for 30 seconds, 72°C for 30 seconds (step
3 – repeated another 39 times ie. 40 cycles in total). In some QPCRs an additional
elongation step was added at 72°C for 10 minutes (step 4). Dissociation curves were
obtained by heating and cooling the samples at: 95°C for 15 seconds, 60°C for 30 seconds,
95°C for 15 seconds. DNA was renatured for agarose gel electrophoresis using the following
conditions: 50°C for 2 minutes (step 1), 95°C for 15 seconds, 60°C for 30 seconds
(step 2 – repeated one more time) and 72°C for 5 minutes (ABI PRISM 7000 Sequence
Detection System) or 95°C for 30 seconds, 60°C for 2 minutes (ABI 7300 Real-Time PCR
Preparation of samples for agarose gel electrophoresis and sequencing
PCR products were purified using Standard Performa 96 well plates and QuickStep 2
SOPE resin (both from EDGE BioSystems), following the recommended procedure.
Agarose gel electrophoresis of purified QPCR products
For each sample 10 μl of 2× Orange G loading buffer (composition shown below) was
added to 5 μl of the purified PCR product and made to 20 μl with water. Samples were
prepared in 96 well plates using the Biomek FX Laboratory Automation Workstation (Beckman
Coulter) and using the same instrument applied to 2% agarose 96 well E-gels (Invitrogen).
For 10× Orange G loading buffer, a solution of 30% Ficoll 400 (AlfaAesar), 10 mM EDTA
(Sigma) was prepared and Orange G dye (Fisher Scientific) was added for color. E-Gel
Low Range Quantitative DNA Ladder (Invitrogen) was used as a marker for PCR product
size. The gels were run for 12 minutes on the E-Gel 96 Base (Invitrogen) and analyzed
using the E-Editor Software (Invitrogen).
Sequencing of purified QPCR products
Purified QPCR products were sequenced at Sequencing Core lab of Center for Computational
and Integrative Biology at Massachusetts General Hospital.
NCBI BLAST analysis
Sequences obtained were BLAST analyzed as batch sets against the NCBI database []. In order to identify successful samples, the main parameters considered were the
alignment length, the expected sequence match position to the sequence returned by
NCBI BLASTn and the percent identity of the two sequences. If more than 50% of the
length of the expected PCR product sequence aligned with the expected sequence as
first match and there was more than 92% identity between the sequences, this was considered
to be a successful sample. In cases where a primer pair had been designed to also
amplify a redundant gene and the redundant gene matched first to the sample, the reaction
was still considered successful. In these cases the primers have been designed to
amplify the same region of the two sequences, so it is not possible to determine by
agarose gel or BLAST analysis if one or the other species was amplified during PCR.
Preparation of digested genomic DNA for QPCR
129 Embryonic Stem cell mouse genomic DNA (isolated by ethanol precipitation) was
used. The DNA was digested completely using EcoRI and BamHI restriction enzymes. Digests were made by adding 20 μl EcoRI buffer (10×) (New England Biolabs), 20 μl 10× BSA, 4 μg DNA, 40 U BamHI (New England Biolabs), 40 U EcoRI (New England Biolabs) and water to 200 μl total volume. Digests were incubated
at 37°C for 4 hours and 30 minutes and heat inactivated at 75°C for 10 minutes. The
digested DNA was phenol extracted and ethanol/salt precipitated. DNA pellets were
resuspended in TE pH 8.0.
QPCRs for uniformity, technical replicate and primer validation tests
QPCRs were performed in polypropylene 96 well plates on the ABI 7300 Real-Time PCR
System (Applied Biosystems). For each reaction, 12.5 μl of Absolute Q-PCR SYBR Green
ROX mix (ABgene) were added to 2.5 μl of 2 μM forward and reverse primer mix (final
concentration of each primer is 200 nM), 1 μl of 50 ng/μl BamHI/EcoRI digested genomic DNA and made to 25 μl with water. The following PCR conditions
were used: 50°C for 2 minutes (step 1), 95°C for 15 minutes (step 2), 95°C for 15
seconds, 60°C for 30 seconds, 72°C for 30 seconds (step 3 – repeated another 39 times
ie. 40 cycles in total), 72°C for 10 minutes (step 4). Dissociation curves were obtained
by heating and cooling the samples at: 95°C for 15 seconds, 60°C for 30 seconds, 95°C
for 15 seconds.
Large-scale amplicon preparation for SYBR Green I sequence specificity experiments
Amplicons were prepared large-scale by PCR, in two steps. For the first step PCR,
75 μl PCR reactions were prepared for each sample. For each reaction, 37.5 μl Absolute
Q-PCR SYBR Green ROX mix (ABgene) were added to 3 μl of 5 μM primer pair mix (final
concentration of each primer is 200 nM), 3 μl universal mouse cDNA (see: 'preparation
of cDNA sample' section in methods) and made up to 75 μl with water. The following
PCR conditions were used: 95°C for 15 minutes (step1), 95°C for 15 seconds, 60°C for
30 seconds, 72°C for 30 seconds (step 2 – repeated another 39 times ie. 40 cycles
in total), 72°C for 10 minutes (step 3). The PCR products were purified using the
MinElute PCR purification kit (Qiagen). Purified amplicons were used as templates
in large-scale 40× 100 μl PCRs, each reaction containing 50 μl 2× LC1v3 buffer (40
mM Tris-HCl pH8.8, 40 mM KCl, 40 mM ammonium sulfate, 4 mM MgCl2, 200 μg/ml BSA, 0.2% Triton X-100, 400 μM dNTP mix, 2.5 M betaine), 4 μl of 5 μM
forward and reverse primer mix, DNA template, 1 μl Taq polymerase and water to 100
μl. The PCR conditions used were the following: 95°C for 3 minutes (step 1), 95°C
for 15 seconds, 60°C for 30 seconds, 72°C for 30 seconds (step 2 – repeated another
39 times ie. 40 cycles in total), 72°C for 10 minutes (step 3).
PCR reactions were phenol extracted and isopropanol precipitated. DNA pellets were
resuspended in TE pH8.0. DNA was purified using Performa DTR Gel Filtration Cartridges
(EDGE BioSystems), following the recommended procedure. Amplicon concentrations were
determined by taking OD260 readings of each preparation using the ND-1000 Spectrophotometer
(Nanodrop). The average value was taken and the OD260 reading from a no DNA template
control was subtracted, in order to remove the contribution from primers and buffer
components to the spectrophotometric absorption.
SYBR Green I sequence specificity experiments
DNA samples in 1× Absolute Q-PCR SYBR Green ROX mix (ABgene) were pipetted into OptiPlate-96F
black 96 well plates (Perkin Elmer). SYBR Green I fluorescence was detected using
the Analyst AD fluorescence plate reader (Molecular Devices) by excitation at 485
nm and emission at 530 nm (505 nm dichroic mirror).
Robotic and manual liquid transfer test
5 μl of 10 mM dNTP solution were added to 95 μl water and the OD260 readings were
taken using the Spectra Max Plus Spectrophotometer (Molecular Devices).
Primer genome location analysis
Mouse genome sequences were downloaded from the UCSC genome browser [] and the primer pair sequences were matched by BLASTn to the genome sequences, to
identify the primer locations with respect to exons.
Authors' contributions
AS designed and performed the experiments, analyzed experimental data and prepared
the manuscript. XW designed the primer algorithm. HW and SD provided bioinformatics
support. TT performed the automation experiments. BS designed and directed the experiments
and prepared the manuscript. All authors have read and approved the final manuscript.
Acknowledgements
We thank our colleagues Chen Liu and Don Dwoske for their technical help, the Automation,
Synthesis and Sequencing Core labs of Center for Computational and Integrative Biology
at Massachusetts General Hospital for their contributions and Naifang Lu for the 129
ES cell mouse genomic DNA. This work was supported by the National Institutes of Health
Program for Genomic Applications, grant U01 HL66678.
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