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High concentration of primer in PCR

High concentration of primer in PCR


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Why is it that higher concentration of primers than the DNA template in PCR will favor the annealing of a template to a primer?


To simply put it, higher concentration of primers means more primers in the mix, therefore more chances of it annealing to your DNA template, however more is not always good. The optimum concentration of primers in a PCR reaction is between 0.1 and 0.5 µM. For most applications 0.2 µM suffices. Using very high concentration of primers can be troublesome as it can cause your primers to bind to partially complementary sites resulting in non-specific PCR products.

https://www.embl.de/pepcore/pepcore_services/cloning/pcr_strategy/optimising_pcr/


Using high concentration of primers ensures that after denaturation template DNA binds to primers instead of binding to each other. Also higher primer concentration enhanced product yield, as primers act as limiting factor for annealing.

(Reference: https://academic.oup.com/nar/article/24/5/985/1047654)


Calculating Concentrations for PCR

i) Oligonucleotide primers are generally supplied as "so many OD units/ml" - but what does this mean, in terms of mg/ml, or mmol/ml, etc?

Given: a primer is Y nucleotides (nt) long

Given: the MW of ssDNA is (330 daltons per nt) x (length in nt) (Sambrook et al., 1989 p. C.1)

Given: the concentration of primer (=ssDNA) producing an OD of 1 at 254 nm in a 1 cm cuvette, is 37 ug/ml

Then: the MW of the primer is 330.Y daltons

And: X OD/ml = 37.X ug/ml = 37.X mg/l = 37.X /330.Y mM = 37.X.1000/330.Y uM

For example:

B 88/77 primer - a 17-mer oligodeoxynucleotide - as supplied is 12.6 OD units/ml. We need to make a 5 uM stock solution for PCR.

MW: 17 x 330 = 5610

Concentration: 12.6 OD x 37 ug/ml = 466 ug/ml = 466 mg/l = 0.466 g/l

Molarity: 0.466/5610 = 0.000083 Molar = 83 uM

Therefore: we need 5 ul of oligo stock solution in 83 ul (+78 ul water) to make a 5 uM solution (if 1 ul in 83 ul gives a 1 uM soln. )

ii) Calculation of amounts for PCR reactions: if we need a final concentration of 0.5 uM oligo in the PCR reaction mix (final volume 50 ul), we add 5 ul of 5 uM stock to the reaction mix (1/10 final dilution).

B) Nucleotides:

Stocks of nucleotides for PCR (or other procedure) are NEARLY ALWAYS dNTP s (deoxynucleotides), and concentrations is almost always given in EACH dNTP: that is, the given concentration is EACH nucleotide in the mix, NOT the total concentration. This means that a 2.5 mM dNTP mix for PCR contains 2.5 mM of EACH dNTP, and 10 mM TOTAL dNTPs.

Example:

i) Make up a 2.5 mM stock solution of dNTPs from stock 100 mM individual dNTPs, supplied by Promega:

  • FIRST mix equal volumes of each nucleotide (eg: 50 ul): this gives you 200 ul of 25 mM mixed dNTPs (Remember: concn. expressed in EACH dNTP).
  • THEN dilute this (or aliquot) 1/10 with WATER - aliquot into 100 ul amounts and freeze.

ii) Prepare a 1 mM stock of dNTPs with dTTP substituted to 10% (w/w) by digoxigenin-11-dUTP (DIG-dUTP) for use as a labelling mix for PCR labelling of PCR products:

  • DIG-dUTP supplied (by Boehringer Mannheim) at 25 nmol/25ul = 1 umol/ml = 1mM final concentration of DIG-dUTP must be 1/10th that of other nucleotides, and [DIG-dUTP] + [dTTP] must = [any other dNTP]. Therefore to get a 1 mM dNTP stock one must dilute DIG-dUTP stock 1/10.
  • FIRST dilute separate 100 mM dNTP stocks to 10 mM (eg. 5 ul to 50 ul, in water).
  • THEN mix equal volumes (eg. 10 ul) of 10 mM dCTP, dGTP and dATP stock, and 9/10ths volume of dTTP (9 ul). Add equal volume (eg. 10 ul) of of 1 mM DIG-dUTP.
  • THEN add water to 10 vol (=100 ul add 51 ul): final concentration each dNTP = 1 mM final concn DIG-dUTP = 0.1 mM, and of dTTP = 0.9 mM.

iii) USE mix made above at 50 uM each dNTP in a PCR reaction mix, final volume 25ul:


Protocol

Reaction setup:

We recommend assembling all reaction components on ice and quickly transferring the reactions to a thermocycler preheated to the denaturation temperature (95°C).

Component 25 &mul reaction 50 &mul reaction Final Concentration
10X Standard Taq Reaction Buffer 2.5 &mul 5 &mul 1X
10 mM dNTPs 0.5 µl 1 &mul 200 µM
10 µM Forward Primer 0.5 µl 1 &mul 0.2 µM (0.05&ndash1 µM)
10 µM Reverse Primer 0.5 µl 1 &mul 0.2 µM (0.05&ndash1 µM)
Template DNA variable variable <1,000 ng
Taq DNA Polymerase 0.125 µl 0.25 µl 1.25 units/50 µl PCR
Nuclease-free water to 25 µl to 50 µl

Notes: Gently mix the reaction. Collect all liquid to the bottom of the tube by a quick spin if necessary. Overlay the sample with mineral oil if using a PCR machine without a heated lid.

Transfer PCR tubes from ice to a PCR machine with the block preheated to 95°C and begin thermocycling.

Thermocycling conditions for a routine PCR:

STEP
TEMP
TIME
Initial Denaturation
95°C
30 seconds
30 Cycles 95°C
45-68°C
68°C
15-30 seconds
15-60 seconds
1 minute/kb
Final Extension 68°C 5 minutes
Hold 4-10°C


General Guidelines:

Oligonucleotide primers are generally 20&ndash40 nucleotides in length and ideally have a GC content of 40&ndash60%. Computer programs such as Primer3 (https://bioinfo.ut.ee/primer3) can be used to design or analyze primers. The final concentration of each primer in a reaction may be 0.05&ndash1 &muM, typically 0.1&ndash0.5 &muM.

Mg ++ concentration of 1.5&ndash2.0 mM is optimal for most PCR products generated with Taq DNA Polymerase. The final Mg ++ concentration in 1X Standard Taq Reaction Buffer is 1.5 mM. This supports satisfactory amplification of most amplicons. However, Mg ++ can be further optimized in 0.5 or 1.0 mM increments using MgCl2.

Amplification of some difficult targets, like GC-rich sequences, may be improved with additives, such as DMSO (3) or formamide (4).

The final concentration of dNTPs is typically 200 &muM of each deoxynucleotide.

We generally recommend using Taq DNA Polymerase at a concentration of 25 units/ml (1.25 units/50 &mul reaction). However, the optimal concentration of Taq DNA Polymerase may range from 5&ndash50 units/ml (0.25&ndash2.5 units/50 &mul reaction) in specialized applications.

An initial denaturation of 30 seconds at 95°C is sufficient for most amplicons from pure DNA templates. For difficult templates such as GC-rich sequences, a longer initial denaturation of 2&ndash4 minutes at 95°C is recommended prior to PCR cycling to fully denature the template. With colony PCR, an initial 5 minute denaturation at 95°C is recommended.

During thermocycling a 15&ndash30 second denaturation at 95°C is recommended.

The annealing step is typically 15&ndash60 seconds. Annealing temperature is based on the Tm of the primer pair and is typically 45&ndash68°C. Annealing temperatures can be optimized by doing a temperature gradient PCR starting 5°C below the calculated Tm. The NEB Tm Calculator is recommended to calculate an appropriate annealing temperature.

When primers with annealing temperatures above 65°C are used, a 2-step PCR protocol is possible (see #10).

The recommended extension temperature is 68°C. Extension times are generally 1 minute per kb. A final extension of 5 minutes at 68°C is recommended.

Generally, 25&ndash35 cycles yields sufficient product. Up to 45 cycles may be required to detect low-copy-number targets.

When primers with annealing temperatures above 65°C are used, a 2-step thermocycling protocol is possible.


Protocol

Reaction setup:

We recommend assembling all reaction components on ice and quickly transferring the reactions to a thermocycler preheated to the denaturation temperature (95°C).

Component 25 &mul reaction 50 &mul reaction Final Concentration
10X Standard Taq Reaction Buffer 2.5 &mul 5 &mul 1X
10 mM dNTPs 0.5 µl 1 &mul 200 µM
10 µM Forward Primer 0.5 µl 1 &mul 0.2 µM (0.05&ndash1 µM)
10 µM Reverse Primer 0.5 µl 1 &mul 0.2 µM (0.05&ndash1 µM)
Template DNA variable variable <1,000 ng
Taq DNA Polymerase 0.125 µl 0.25 µl 1.25 units/50 µl PCR
Nuclease-free water to 25 µl to 50 µl

Notes: Gently mix the reaction. Collect all liquid to the bottom of the tube by a quick spin if necessary. Overlay the sample with mineral oil if using a PCR machine without a heated lid.

Transfer PCR tubes from ice to a PCR machine with the block preheated to 95°C and begin thermocycling.

Thermocycling conditions for a routine PCR:

STEP
TEMP
TIME
Initial Denaturation
95°C
30 seconds
30 Cycles 95°C
45-68°C
68°C
15-30 seconds
15-60 seconds
1 minute/kb
Final Extension 68°C 5 minutes
Hold 4-10°C


General Guidelines:

Oligonucleotide primers are generally 20&ndash40 nucleotides in length and ideally have a GC content of 40&ndash60%. Computer programs such as Primer3 (https://bioinfo.ut.ee/primer3) can be used to design or analyze primers. The final concentration of each primer in a reaction may be 0.05&ndash1 &muM, typically 0.1&ndash0.5 &muM.

Mg ++ concentration of 1.5&ndash2.0 mM is optimal for most PCR products generated with Taq DNA Polymerase. The final Mg ++ concentration in 1X Standard Taq Reaction Buffer is 1.5 mM. This supports satisfactory amplification of most amplicons. However, Mg ++ can be further optimized in 0.5 or 1.0 mM increments using MgCl2.

Amplification of some difficult targets, like GC-rich sequences, may be improved with additives, such as DMSO (3) or formamide (4).

The final concentration of dNTPs is typically 200 &muM of each deoxynucleotide.

We generally recommend using Taq DNA Polymerase at a concentration of 25 units/ml (1.25 units/50 &mul reaction). However, the optimal concentration of Taq DNA Polymerase may range from 5&ndash50 units/ml (0.25&ndash2.5 units/50 &mul reaction) in specialized applications.

An initial denaturation of 30 seconds at 95°C is sufficient for most amplicons from pure DNA templates. For difficult templates such as GC-rich sequences, a longer initial denaturation of 2&ndash4 minutes at 95°C is recommended prior to PCR cycling to fully denature the template. With colony PCR, an initial 5 minute denaturation at 95°C is recommended.

During thermocycling a 15&ndash30 second denaturation at 95°C is recommended.

The annealing step is typically 15&ndash60 seconds. Annealing temperature is based on the Tm of the primer pair and is typically 45&ndash68°C. Annealing temperatures can be optimized by doing a temperature gradient PCR starting 5°C below the calculated Tm. The NEB Tm Calculator is recommended to calculate an appropriate annealing temperature.

When primers with annealing temperatures above 65°C are used, a 2-step PCR protocol is possible (see #10).

The recommended extension temperature is 68°C. Extension times are generally 1 minute per kb. A final extension of 5 minutes at 68°C is recommended.

Generally, 25&ndash35 cycles yields sufficient product. Up to 45 cycles may be required to detect low-copy-number targets.

When primers with annealing temperatures above 65°C are used, a 2-step thermocycling protocol is possible.


Results

Blocking primer performance

Adding a blocking primer to the PCR mixture clearly decreased the number of predator fragments amplified and enriched the number of rarer prey fragments. In rDNA mixtures containing 1000 times as many "predator" (krill) rDNA fragments compared to "prey" (algae) rDNA fragments, only peaks corresponding to krill rDNA could be detected by fragment analysis when no blocking primer was added (Table 3). Adding an annealing inhibiting blocking primer in a ratio 4:1 compared to the corresponding universal primer (1.0 μL blocking primer and 0.25 μL universal primer, both 10 μM) led to reduced amplification of krill rDNAs but not to complete amplification arrest (Table 3). And by adding 10 times as much blocking primer as universal primers (2.5 μL blocking primer and 0.25 μL each universal primer), algal rDNA was almost exclusively amplified (Table 3). Adding 20 times as much blocking primer (5.0 μL blocking primer and 0.25 μL each universal primer) decreased the krill peak even further (Table 3).

The "normal" primer Short28SR-blkKrill3'c3 seemed to be somewhat more efficient in blocking predator DNA compared to the DPO primer Short28SF-DPO-blkKrill. Peaks corresponding to krill rDNA were lower in the 4:1 blocking primer: universal primer mixtures for both the 1:100 and the 1:1000 prey: predator rDNA samples with the normal primer compared to the mixtures with the DPO primer (Table 3). However, the DPO primer largely blocked krill DNA amplification when added in a ratio of 10:1 to the unmodified primer (Table 3).

The elongation arrest primer 28S-ElArKrill-3'c3 did not work. No PCR product at all was generated when it was added to the PCR mixture (data not shown).

Krill stomach analysis

In total 111 clones from 13 different krill stomachs were sequenced revealing 36 different sequences falling into 6 major groups. Among them there were no sequences corresponding to the wild type of krill 28s rDNA. 10 different (15 in total) sequences were however clearly of krill origin (Figure 3) and probably pseudogenes or low copy versions normally not detected by direct sequencing.

Sequence similarity tree. Sequence similarity tree of sequences amplified from krill stomachs and related sequences. The krill stomach sequences are named so as the first three letters means sampling date, i.e. M24 = March 24. 2007, S17 and S20 = September 17. and September 20. 2007, St. means Stomach number and the number in parenthesis equals the number of identical sequences of each different sequence found in the different stomachs. The tree is unrooted.

Four different groups of algal sequences were found. Two of these groups, only detected in the March samples, had their closest match to species belonging to Bacillariophyta, respectively the genus Phaeodactylum and the genus Skeletonema (Figure 3). A third group, found in stomachs from all three sampling dates seemed to be somewhat similar to the Bacillariophyta, but did not match anything in GenBank (Figure 3). The 200 bp area sequenced was too uninformative to reveal where the group belonged by phylogentic analysis, besides that the group most likely was of algal origin. The last algal group was one sequence from September belonging to Chlorophyta (Figure 3).

One large group of sequences, which was prevalent in the September sample, could not be unequivocally identified by reference to current sequences in GenBank. The closest matches was a 96% match to Stauroteuthis (Mollusca). It is possible that this may represent mollusc food items such as larval squid. However, it is also possible that the Stauroteuthis sequence has been misreported and is actually derived from a contaminant of the original Stauroteuthis sample.

The sequences have been deposited to GenBank under the accession numbers EU378965 – EU379000.


Taq Polymerase


Unit Definition: One unit is defined as the amount of the enzyme required to catalyze the incorporation of 10 nmoles of dNTP's into an acid-insoluble form in 30 minutes at 70 °C using hering sperm DNA as substrate.

Shipping: shipped on blue ice

Storage Conditions: store at -20 °C
avoid freeze/thaw cycles

Shelf Life: 12 months

Concentration: 5 units/&mul

Description:
Taq Polymerase is recommended for routine PCR applications (up to 4 kb fragment length), high throughput PCR or genotyping. The buffer system guarantees robust and reliable amplification results in almost all PCR applications. The Crystal Buffer contains a well-balanced ratio of potassium-, ammonium- and magnesium-ions to ensure high specificity and minimal by-product formation without the need of additional optimization steps.
Ruby Buffer additionally contains gel loading buffer and an inherent red dye. The red dye allows an easy visual control during PCR set-up and in combination with the density reagent the direct loading of the reaction product into the gel.
The enzyme replicates DNA at 72 °C. It catalyzes the polymerization of nucleotides into duplex DNA in 5'→3' direction in the presence of magnesium. It also possesses a 5'→3' polymerization-dependent exonuclease replacement activity but lacks a 3'→5' exonuclease (proof-reading) activity.

Content:
Taq Polymerase (red cap)
5 units/&mul Taq DNA Polymerase in 20 mM Tris-HCl, 100 mM KCl, 0.1 mM EDTA, 1 mM DTT, 0.5 % Tween-20, 0.5 % Nonidet P-40, 50 % (v/v) Glycerol, pH 8.0 (25°C)

Ruby Buffer (black cap)
10 x conc. complete PCR buffer containing 200 mM Tris-HCl, KCl, (NH4)2SO4 and 20 mM MgCl2, red tracking dye and density reagent for gel loading

Crystal Buffer (green cap)
10 x conc. complete PCR buffer containing 200 mM Tris-HCl, KCl, (NH4)2SO4 and 20 mM MgCl2

componentPCR-211SPCR-211LPCR-211XL
Taq
Polymerase
200 units
/ 40 &mul
1000 units
/ 200 &mul
5000 units
/ 1 ml
Ruby Buffer1.2 ml5 x 1.2 ml25 ml
Crystal
Buffer
1.2 ml5 x 1.2 ml25 ml


Assay Set-Up:
Before starting, vortex all components thoroughly to ensure homogeneity.
Prepare a premix for the number of assays you need according to the following protocol:

comp.capstock conc.final conc.1 assay @ 20 &mul1 assay @ 50 &mul
PCR-grade Waterwhite fill up to 20 &mulfill up to 50 &mul
Ruby Buffer or Crystal Bufferblack or green10x1x2 &mul5 &mul
dNTP Mix / 10 mM #NU-1006white10 mM200 &muM0.4 &mul1 &mul
Taq Polymerasered5 units/&mul0.025 units/&mul0.1 &mul0.25 &mul
primer mix or each primer 10 &muM each primer200 - 400 nM each primer0.4-0.8 &mul1 - 2 &mul
template
/sample DNA
10 &mul 1)
elongation 2)
95 °C
50 - 68 °C
72 °C
10 - 20 sec
10 - 20 sec
20 sec - 4 min

25 - 35x

Gel Loading and Down-Stream Applications:
Ruby Buffer (#PCR-272) includes a density reagent + tracking dye and allows the direct loading of the PCR products into a electrophoresis gel. For DNA detection / fluorescent DNA staining we recommend to use new generations dyes (i.g. SYBR DNA Stain, #PCR-273) instead of the classical but highly mutagenic ethidium bromide.
Crystal Buffer(#PCR-271) is recommended for down-stream applications such as DNA sequencing, ligation, restriction digestion or where an analysis of the PCR product by absorbance or fluorescence excitation is required. For gel electrophoresis add gel loading buffer and fluorescent DNA stain (i.g. Gel Loading Buffer with DNA Stain, #PCR-274 - #PCR-276) before loading the PCR into the gel. Using pre-stained gels or post-run staining protocols is also possible.

Additional Buffer Systems:
Labeling Buffer (#PCR-263) is recommended for DNA labeling or mutagenesis applications. The buffer is specially optimized for incorporation of labeled or modified nucleotides into DNA. It gives superior results in a broad range of reaction conditions with most primer-template pairs but amplification may also tend to an increased unspecifity.
KCl Buffer (#PCR-262) is recommended for use in routine PCR reactions. The buffer is optimized for highest specificity but may require additional fine-tuning of assay parameters like MgCl2 concentration and annealing temperature.

Optimization of MgCl2 concentration:
A final Mg 2+ concentration of 2.0 mM is recommended in combination with Labeling Buffer. However, if an individual Mg 2+ optimization is essential add 25 mM MgCl2 stock solution (#PCR-266) as shown in the table below.

final MgCl2 conc.20 &mul final assay volume50 &mul final assay volume
2 mM--
3 mM0.8 &mul2.0 &mul
4 mM1.6 &mul4.0 &mul
5 mM2.4 &mul6.0 &mul

Product Citations:
Please click the black arrow on the right to expand the citation list. Click publication title for the full text.


Discussion

We describe here a novel technique for amplifying DNA to reduce the negative effects of mismatches between primer and template on the efficiency of amplification of target templates. This method is an extremely general and simple modification to the PCR reaction, and involves three enzymatic steps, including a polymerase-mediated linear copying of genomic DNA templates, an exonuclease digestion of primers, and a final PCR amplification using primers targeting target-independent linker sequences. As shown in this manuscript, this approach allows the exponential amplification of the target of interest to be performed with primers that have no mismatches with any templates in the reaction. This significantly reduces bias associated with mismatches and degenerate primers that can accumulate during PCR by limiting primer-template interactions to two cycles only.

The objectives of this study were to: (i) demonstrate that the first and second stages of PCR could be separated, and still generate reliable amplification (ii) determine if degeneracies in the primer pools led to obvious distortion of the observed microbial community, and if the PEX PCR technique could be used to circumvent or reduce such distortion, (iii) develop a robust workflow for this approach to be implemented for any primer set or sets, and (iv) identify applications to which this approach is best suited. The results herein demonstrate that indeed the two defined types of interactions within PCR (i.e. natural and artificial interactions) can and should be separated when degenerate primers are used or when mismatches with the template are anticipated. The stages should be separated because genomic DNA template-primer interactions have the greatest potential for bias due to mismatches derived from true mismatches in the gDNA and from degeneracies in the primer pool. Our analysis of an artificially synthesized mock community demonstrates the strong potential for a degenerate primer pool of oligonucleotides of varying melting temperatures to preferentially select templates based on sequence variations in the primer site. Our strategy limits the gDNA template-primer interaction to two cycles, with all subsequent amplification cycles employing non-degenerate, non-template interactions. Furthermore, because only two cycles are utilized during the first stage of gDNA-primer interactions, unusual annealing and elongation conditions can be utilized. In this study, we employed long annealing times (20 minutes) at low annealing temperatures to allow for adequate time for the polymerase to bind target sites and elongate without raising the reaction temperature. Such reaction conditions are likely more tolerant of low primer annealing efficiency of primer-template pairings with mismatches. In a systematic study of primer-template interactions, Wu et al. [31] reported that single mismatches occurring in the last 3–4 positions from the 3’ end of the primer yielded minimal or no primer extension. We observe here that 3’ mismatches can be overcome using the PEX PCR method, and that primers with 3 or 4 mismatches can still anneal with genomic DNA targets and yield polymerase extension. This was revealed through an analysis of primer utilization patterns in mock community analyses. We note that such an analysis cannot be performed using standard PCR approaches, and requires the PEX PCR method.

Furthermore, we demonstrate through the comparison of PEX PCR method with and without exonuclease that the primers targeting the source gDNA (or mock DNA) do contribute to the distortion during later cycles of PCR even in the presence of high concentrations of the second stage PCR primers. The complete removal of unincorporated primers from the first stage reaction, although desirable, was found to be difficult. Even after dilution, exonuclease digestion, and lowering of the primer concentration during the first two cycles, some limited amount of first stage primer may be propagated to the second stage PCR. Despite this, treatment with exonuclease significantly reduces the impact of primer carry-over, and is an essential part of the PEX PCR methodology. In analyses of both mock community and environmental gDNA, exonuclease treatment significantly alters the observed microbial community. It is possible that in place of exonuclease treatment, blocking oligonucleotides of the inverse complement of the forward and reverse template-specific primers (e.g. inverse complement of 515F and 806R primers without CS1 and CS2 linkers) could be added to the second PCR stage of the PEX PCR reaction to prevent gDNA template-primer interactions.

The PEX PCR method also provides a novel and robust mechanism to explore primer-template interactions in analyses of complex gDNA samples. The PEX PCR method preserves the sequence of the primer annealing to the gDNA template during the first two cycles of the reaction, and these can be bioinformatically interrogated to determine which primers within a degenerate pool are truly involved in annealing and extension. We observed that at high annealing temperatures, perfect match annealing is favored, and a lower diversity of the primers in the pool were utilized. This appears to be detrimental for the amplification reaction, as perfectly matching primers are present at a low overall abundance in a heavily degenerate primer pool. At lower annealing temperatures, a broader spectrum of primers, containing mismatches with the template, are involved in annealing and elongation. This appears to be beneficial, as analyses of the mock community under lower annealing conditions, generated better representations of the true underlying distribution of mock DNAs. Primers with 0–4 mismatches with various templates were observed to anneal and allow for polymerase extension. When 3’ mismatches were introduced into mock DNA templates, the primer utilization distribution shifted towards primers with 1 or 2 total mismatches to the template. Therefore, the heavy degeneracy of primer sets may not be beneficial when using the PEX PCR method. Instead, it may be appropriate to select “intermediate” primers that have at most 1 or 2 mismatches to all potential priming sites, with the assumption that every variant does not require a unique primer to be targeted. Further research is needed to determine the best combination of primer degeneracy and annealing temperatures for other, more degenerate targets such as microbial functional genes. Such a strategy may enable a direct PCR-based method for analysis of a single copy gene present in all microorganisms for the purpose of community structure analyses. We further note that additional strategies may be required to allow the PEX PCR method to work effectively at lower annealing temperatures (e.g. <45°C), such as the introduction of single-stranded DNA binding protein into the amplification master mixes.

The PEX PCR method is recommended for: (i) any PCR reaction in which a degenerate primer pool is used (ii) any PCR reaction in which a non-degenerate primer is used but where DNA template variability at the priming site is possible (iii) reactions in which high-level degeneracy may be utilized to target all known variants of a gene and (iv) when multiple primer pairs are to be utilized simultaneously. We show here that the method can be used to amplify and sequence templates with mismatches at the 3’ end of the primer site, which have been shown to be highly destabilizing in PCR [31,32]. Crosby and Criddle [33] previously employed a strategy using hybridization capture followed by random-primed amplification and sequencing to target DNA-directed RNA polymerase genes (rpoC). The PEX PCR method may be adaptable to a direct PCR amplification of rpoC genes from all microorganisms within a single amplification reaction. This would preserve the original relative abundance found in the template DNA, and provide a direct proxy for relative abundance of organisms in the sample. This is unlike amplification and sequencing of rRNA genes, as performed in this study, since a wide range of gene copies of rRNA operons are found across the domains Bacteria and Archaea [34]. We do not yet know if the level of degeneracy at conserved regions of the rpoC (or other similar) gene is likely to be a major impediment during the first two cycles of stage “A” of the PEX PCR method, but this is a clear next step in the development of this technique. If primer dimerization in reactions with extremely degenerate primer pools is observed, purification of stage “A” components using a size-selection protocol (e.g. AMPure beads) instead of exonuclease will reduce the transfer of primer dimers which are insensitive to single-strand exonuclease activity.

The PEX PCR method may also find wide-spread application for quantitative PCRs in which degenerate primers are employed. Quantitative PCRs could be performed by initially processing DNA samples prepared using stage “A” of the PEX PCR method. Subsequently, qPCR would be performed using primers targeting linker sequences instead of template-specific primers. This could potentially greatly increase qPCR efficiency and target range for broad-target degenerate primers common to environmental microbiology, and avoid problems deriving from differential amplification efficiencies for different targets. A similar approach has in fact been developed, with the aim to reduce the impact of bacterial DNA contamination in qPCR reactions [35].

Finally, we note that this method has conceptual similarities to a study previously performed by Crosby and Criddle [33], in which linker sequences connected to random primers were used to amplify functional genes that were captured using hybridization probes. In that study, two cycles of annealing and elongation were used for labeling, with subsequent amplification. In addition, Illumina has developed a target-capture approach in which two primers with linkers either at the 5’ or 3’ ends are allowed to anneal to a single strand of template DNA (i.e. TruSeq Amplicon). After polymerase extension, ligation is used to link the elongated fragment to the 3’ terminal primer with a 3’ flanking linker. Subsequently, PCR amplification using the linker sequences is used to prepare fragments for sequencing.


Real-Time qRT-PCR standard curves. efficiency is too high! - (Jun/29/2005 )

In my Real-Time qRT-PCR experiments, I employ the standard curve method for quantification of gene expression. However, standard curves seem to be a huge hit-or-miss procedure for me, even with genes that are well-established to work well with Real-Time such as GAPDH.

At times, I am able to produce excellent standard curves with slopes at approximately -3.3 (

100% efficiency) the associated melting curves for each dilution are excellent as well.

However, most of the time I may get excellent melting curves for each dilution, but the slope for my standard curve may fall at around -2.6 (

Can anyone offer some explanations for this strange dilemma? Consequently, can anyone offer some suggestions to solve this problem?

One thing I can think of is that I'm getting disuniform amplification efficiencies at different RNA concentrations. FYI, I typically use 100 ng/uL RNA for "1x" and make serial dilutions up to 100x. I understand I really should be using a larger range of dilutions, but greater than 100x dilutions just don't work out for my genes/primers.

Thanks for any help I can get.

What is the range of the Cts? Did you see any contamination in your negative control?

My Ct values for the standard curves usually range from around 14-20, and the negative controls are showing no DNA contamination.

What instrument are you using?

Dear YuJ,
Are you using SYBR Green as reporter dye and you are using iCycler?
What is your lowest dilution in your standard?

This is what always happen to me. If i run a SYBR Green assay using standard ranging from 10e6 to 1, i will get > 100% efficiency.
So what i did was I unselect the last two dilution (10 and 1) and the efficiency become

3.3.
I think this is due to the primer dimer formation that contribute to the false signal in you reaction. And the effect of the primer dimer is great enough to afact you data especially in the low concentration standard.

I am indeed using SYBR Green chemistry (ABI SYBR Green PCR Mix), but the instrument I use is the Applied Biosystems 7900HT.

I have tried removing certain dilutions in all possible permutations before plotting the trendlines, but the outcomes are inconsistent between experiments. I also don't believe I have any primer-dimers because of the very pronounced product peaks and complete absence of primer-dimer peaks from the dissociation/melting data.

Hi YuJ,
Could you please describe how you preparing your quantitative standard?

My quantitative standards are prepared as such.

1. RNA isolation using TRIZOL reagent following manufacturer instructions plus an additional overnight purification step with ethanol (100%) and sodium acetate (3 M).

2. Spectrophotometric determination of [RNA]. A260/A280 ratio is usually at around 1.8.

3. Serial dilution of RNA at 1x, 10x, 25x, 75x, 100x, and sometimes 1000x (with 1x being 100 ng/uL diluted from stock, 10x being 10 ng/uL, etc.) using DEPC-treated water. I make sure to vortex each tube very well before pipetting for the next dilution.

4. Reagents in RT-PCR reactions include: Multiscribe Reverse Transcriptase (ABI), RNaseOUT RNase Inhibitor (Invitrogen), SYBR Green 2x PCR Mix (ABI). These, plus DEPC-treated water, are combined as a master mix.

5. Primer concentrations have been previously optimized for each primer. In the case of GAPDH, I determined the ideal concentration to use would be 0.06 uM per reaction for both the forward and reverse primer.

6. In each reaction tube, I first add the water and master mix, then the primers, and then finally the appropriate RNA template.

7. Each tube is vortexed and loaded in triplicate into the appropriate optical reaction plate. Plate is centrifuged to get rid of air bubbles and placed into the ABI 7900HT.

8. Thermocycler is set for a 45C RT phase (30min), 95C melting + 60C annealing phase (40 cycles), and then an additional dissociation phase at the end for generating the dissociation curves.

I probably included a lot of irrelevant facts, but hopefully this is what you meant by how I prepare my quantiative standards.

Dear YuJ,
Thanks for your description. I would strongly suggest you to use invitro transcription to prepare your GAPDH mRNA rather than using TRIzol extracted total RNA.

The reason is when you use TRIzol to extraction, you will get total RNA not GAPDH specific mRNA. So when you quantitate using spetrophotometrically, the reading will be out. Thus it would not give you a

You may try to clone the GAPDH gene into a plasmid and invitro trascripted(IVT) into GAPDH mRNA --> treat with DNase I to completly remove DNA conteminant--> stop DNase reaction --> purify the mRNA --> quantitate you mRNA*-->converte xx ug/ul into xx RNA copy/ul--> perform 10 fold serial dilution --> run RT-PCR.

This should give you a batter result.

*optional:
If you want to make sure that you IVT GAPDH mRNA is free from DNA conteminat, you may run a RT-PCR and a PCR (no RT) simultaneously. Your (no RT) PCR should not give you any band. If it does, treat you standard again with DNase I.

Note:
when you do a cloning please in clude 10-20 bp extra flanking at the both 3' and 5' end of your acture mRNA target. This will provide bater stability against exonuclease and yield prolong shelf life to your standard.

I would like to recommend you to use Ambion produst for IVT purpose.

Thank you for your advice, Hadrian, but I do not believe that method is suitable for me.

Although I've only mentioned GAPDH serial dilution as an example, I am actually conducting a gene expression study for some other genes, merely using GAPDH as an internal control.

Also, when I made the spectrophotometric measurements, I was just interested in the A260/A280 ratio as a measure of RNA purity and a rough estimate of the total RNA concentration in the stock tube. Indeed I am measuring total RNA, but gene-specific primers are used in the RT-PCR for actual quantification.

Sorry, I should have first mentioned what I'm actually doing instead of assuming that others can read my mind.


The Degenerate Primer Design Problem: Theory and Applications

A PCR primer sequence is called degenerate if some of its positions have several possible bases. The degeneracy of the primer is the number of unique sequence combinations it contains. We study the problem of designing a pair of primers with prescribed degeneracy that match a maximum number of given input sequences. Such problems occur when studying a family of genes that is known only in part, or is known in a related species. We prove that various simplified versions of the problem are hard, show the polynomiality of some restricted cases, and develop approximation algorithms for one variant. Based on these algorithms, we implemented a program called HYDEN for designing highly degenerate primers for a set of genomic sequences. We report on the success of the program in several applications, one of which is an experimental scheme for identifying all human olfactory receptor (OR) genes. In that project, HYDEN was used to design primers with degeneracies up to 10 10 that amplified with high specificity many novel genes of that family, tripling the number of OR genes known at the time.


High concentration of primer in PCR - Biology

The 64-bit Mac version should work on most modern Macs (OS X 10.7 or newer).

FreeBSD users may simply type pkg install pooler

For all other systems (GNU/Linux, older Mac, Solaris. ) please compile from source (below).

Example primers file

Source code

  • a C compiler (GCC or Clang) and basic Unix tools,
  • MingW compiler(s) if you want to cross-compile for Windows.

Usage

  • If you opt to use Score when your primers and/or tags are very long, you will be asked if you are really sure you don't want to use deltaG instead.
  • If you opt for deltaG, the following questions will be asked: Temperature: Enter a number (decimal fractions are allowed). You can enter it in Celsius, Kelvin, Fahrenheit or Rankine. Do not enter the suffix C or K or F or R---Primer Pooler will determine for itself which unit was meant, and ask you to confirm. (Recent versions of Primer Pooler offer 5 additional obscure temperature scales if you decline all of the more probable ones.) Magnesium concentration in mM (0 for no correction): Enter your concentration of magnesium in nanomoles per cubic metre (decimal fractions are allowed). Enter 0 if you don't mind the deltaG figures not being corrected for magnesium concentration. Monovalent cation (e.g. sodium) concentration in mM: Enter your concentration of sodium etc in nanomoles per cubic metre (decimal fractions are allowed). If in doubt, try 50. dNTP concentration in mM (0 for no correction): Enter your concentration of deoxynucleotide (dNTP) in nanomoles per cubic metre (decimal fractions are allowed). Enter 0 if you don't mind the deltaG figures not being corrected for dNTP concentration.
  • If you answered yes to this question, the summary will be displayed on screen, and you will be asked if you also want to save it to a file. If you answer yes to this, you will be asked for a filename.
  • These up-front counts will include self-interactions (a primer interacting with itself), and interactions between the pair of primers in any given set. Self-interactions and in-set interactions are not counted when summarizing the counts of each pool (below).
  1. Go to http:// hgdownload. cse. ucsc. edu/ downloads. html
  2. Choose a species (e.g. Human)
  3. Choose "Genome sequence files"
  4. If you're under hg38, choose "Standard genome sequence files"
  5. Scroll down to the links, and choose the one that ends .2bit (e.g. hg38.2bit)
  • After the overlap scan is complete, Primer Pooler will then have enough data to write an input file for MultiPLX if you wish to run that software as well for comparison. If you decline this, it will ask if you want it to write a simple text file with the locations of all amplicons, which you may accept or decline.
  • If you do not opt to check for overlaps in the genome, then Primer Pooler will not take overlaps into account when generating its pools. This is rarely useful unless you have already ensured there are no overlaps in the set of amplicons under consideration. Even then, I would recommend performing a scan anyway, just to double-check: an early version found 11 overlaps in a supposedly overlap-free batch drawn up by an experienced academic---we all make mistakes. But bypassing the overlap check might be useful if you are sure there are no overlaps and you don't want to download a very large genome file to the workstation you're using.

You will not be allowed to set the maximum size of each pool lower than the average size of each pool, since that would make it logically impossible to fit all primer-sets into all pools. It is not advisable to set it just above the average either, since being overly strict about the evenness of the pools could hinder Primer Pooler from finding a solution with lower dimer formation. You might want to experiment with different maxima---you will be able to come back to this question and try again. Do you want to give me a time limit? (y/n): If you answer y, you will be asked to set a time limit in minutes. Normally 1 or 2 is enough, although you may wish to let it run a long time to see if it can find better solutions. You don't have to set a time limit: you may manually interrupt the pooling process at any time and have it give the best solution it has found so far, whether a time limit is in place or not. Additionally, Primer Pooler will stop automatically when it detects better solutions are unlikely to be found. Do you want my "random" choices to be 100% reproducible for demonstrations? (y/n): If you answer y, Primer Pooler's random choices will be generated in a way that merely look random but are in fact completely reproducible. This is useful for demonstration purposes---you'll know how long it will take to find the solution you want. Otherwise, the random choices will be less predictable, as a different sequence will be chosen depending on the exact time at which the pooling was started. Pooling display While pooling is in progress, Primer Pooler will periodically display a brief summary of the best solution found so far, showing the pool sizes, and the counts of interactions (by deltaG range or score) within each pool. As instructed on screen, you may press Ctrl-C (i.e. hold down Ctrl while pressing and releasing C, then release Ctrl) to cancel further exploration and use the best solution found so far. Do you want to see the statistics of each pool? (y/n): After the pooling is complete, or after you have interrupted it (by pressing Ctrl-C as instructed on screen), you will be asked if you wish to see the interaction counts of each pool (rather than a simple summary of all pools as appeared during pooling). If you want this, you will also be asked if you wish to save them to a file, and, if so, what file name. Do you want to see the highest bonds of these pools? (y/n): If you answer Yes, you will be asked for a deltaG or score threshold, and all interactions worse than that threshold will be displayed on-screen with bonds diagrams such as:and you will then be asked if you wish to save it to a file, and, if so, what file name. You will then be asked if you would like to try another threshold. Shall I write each pool to a different result file? (y/n): If you answer y to this, you will be asked for a prefix, which will be used to name the individual results files. Otherwise, you will be asked if you wish to save all results to a single file. If you decline saving all results to a single file, the results will not be saved at all---this is for when you weren't happy with the solution and want to go back to try a different number of pools or a different maximum pool size. Do you want to try a different number of pools? (y/n): This question is self-explanatory. You can go back as many times as you like, trying different numbers of pools. But many researchers have a pretty good idea of how many pools they want to use, or else are happy with the computer's initial suggestion. Would you like another go? (y/n): If you answered No to trying a different number of pools, or if you didn't want the program to do pooling at all, then you will be asked if you want to start the program again. Answering No to this question will exit.

Command-line usage

The only mandatory argument (if not running interactively) is a filename for the primers file. This should be a text file in multiple-sequence FASTA format, such as:(this example does not represent real primers). Degenerate bases are allowed using the normal letters, and both upper and lower case is allowed. Names of amplicons' primers should end with F or R, and otherwise match. Optionally include tags (tails, barcoding) to apply to all primers: >tagF and >tagR (tags can also be changed part-way through the file).

Processing options should be placed before this filename. Options are as follows: --help or /help or /? Show a brief help message and exit. --counts Show score or deltaG-range pair counts for the whole input. deltaG will be used if the --dg option is set (see below). This option produces a fast summary of how many primer pairs (in the entire collection, before pooling) have what range of interaction strengths. This could be used for example to check a pool that you have already chosen manually, or if you want a rough idea of the worst-case scenario that pooling aims to avoid. --self-omit Causes the --counts option to avoid counting self-interactions(a primer interacting with itself), and interactions between the pair of primers in any given set. --print-bonds=THRESHOLD Similar to --counts , this can be useful for checking a manual selection or for a rough idea. All interactions worse than the given threshold (deltaG if --dg is in use, otherwise score) will be written to standard output, with bonds diagrams. --dg[= temperature[, mg[, cation[, dNTP]]]] Set this option to use deltaG instead of score. Optional parameters are the temperature (default is human blood heat), the concentration of magnesium (default 0), the concentration of monovalent cation (e.g. sodium, default 50), and the concentration of deoxynucleotide (dNTP, default 0). Decimal fractions are allowed in all of these. Temperature is specified in kelvin, and all concentrations are specified in nanomoles per cubic metre. --suggest-pools Outputs a suggested number of pools. This is the approximate lowest number of pools needed to achieve no worse than a deltaG of -7 (or a score of 7) in each. --pools[= NUM[, MINS[, PREFIX]]] Splits the primers into pools. Optional parameters are the number of pools (if omitted or set to ? then the suggested number will be calculated and used), a time limit in minutes, and a prefix for the filenames of each pool (set this to - to write all to standard output). --max-count=NUM Set the maximum number of pairs per pool. This is optional but can make the pools more even. A maximum lower than the average is not allowed, and it's usually best to allow a generous margin above the average. --genome=PATH Check the amplicons for overlaps in the genome, and avoid these overlaps during pooling. The genome file may be in .2bit format as supplied by UCSC, or in .fa (FASTA) format. --scan-variants When searching for amplicons in a genome file, scan variant sequences in that file too, i.e. sequences with _ and - in their names. By default such sequences are omitted as they're not normally needed if using hg38. --amp-max=LENGTH Sets maximum amplicon length for the overlap check. The default is 220. --multiplx=FILE Write a MultiPLX input file after the --genome stage, to assist comparisons with MultiPLX's pooling etc. --seedless Don't seed the random number generator --version Just show the program version number and exit.

Changes

Defects fixed

  1. an error in incremental-update logic sometimes had the effect of generating suboptimal solutions (in particular, pools could be unnecessarily empty, and/or full beyond any limit that was set)
  2. an error in the user-interface loop meant that if you use tags, run interactively, and answer "yes" to the question "Do you want to try a different number of pools", the second run will have been done without the tags, and its results will have been de-tagged twice, removing some bases from the output moreover, the resulting truncated versions of your primers will have made it into the interaction calculations for any third run.

Versions prior to 1.17 also had a display bug: the concentrations for the deltaG calculation are in millimoles per litre, not nanomoles as stated on-screen in interactive mode (please ignore the on-screen instruction and enter millimoles, or upgrade to the latest version which fixes that instruction).

Versions prior to 1.34 would round down any decimal fraction you type when in interactive mode (for deltaG temperature, concentration and threshold settings). Internal calculation and command-line use was not affected by this bug.

Versions prior to 1.37 did not ignore whitespace characters after FASTA labels and the label) -->.

Notable additions

Version 1.2 added the MultiPLX output option, and Version 1.33 fixed a bug when MultiPLX output was used with tags and multiple chromosomes. Version 1.3 added genome reading from FASTA (not just 2bit), auto-open browser, and suggest number of pools.

Version 1.36 clarified the use of Taq probes, and allowed these to be in the input file during the overlap check. It's consequently stricter about the requirement that reverse primers must end with R or B : previous versions would accept any letter other than F for these.

Version 1.4 allows tags to be changed part-way through a FASTA file. For example, if there are two >tagF sequences, the first >tagF will set the tags for all F primers between the beginning of the file and the point at which the second >tagF is given the second >tagF will set the tags for all F primers from that point forward. You can change tags as often as you like.

Version 1.5 allows primer sets to be "fixed" to predetermined pools by specifying these as primer name prefixes , e.g. [email protected]:myPrimer-F fixes myPrimer-F to pool 2.

Version 1.6 detects and warns about alternative products of non-unique PCR. It was followed within hours by Version 1.61 which fixed a regression in the amplicon overlap check.

Version 1.7 makes the ignoring of variant sequences in the genome optional, and warns if primers not being found might be due to variant sequences having been ignored.