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Polymerase

Recombinant Taq and native Taq polymerase are identical in terms of their activity, specificity, thermostability, and performance in PCR. Recombinant Taq has been expressed in a bacterial system and purified, whereas native Taq has been purified from the host.

We offer Pfx50™ DNA polymerase that displays 50 times higher fidelity than Taq.

Please see the following comparison table:

Enzyme Relative fidelity Amplicon length 3’ A-overhang
Taq 1 <5 kb +
Platinum® Taq 1 <10 kb +
AccuPrime™ Taq 2 <5 kb +
Platinum® Pfx 26 <12 kb -
AccuPrime™ Pfx 26 <12 kb -
Pfx50™ 50 <4 kb -
Platinum® Taq HiFi 6 <20 kb +/-
AccuPrime™ Taq HiFi 9 <20 kb +/-
AmpliTaq® 1 <5 kb +
AmpliTaq Gold® 1 <5 kb +
AmpliTaq Gold® 360 1 <5 kb +

Taq error rate: 1 x 10-4 to 2 x 10-5 base/duplication

Yes, you can use a proofreading polymerase for PCR. However, you will need to add 3′ A-overhangs to your PCR product prior to TA cloning.

We would recommend using our AccuPrime™ GC-Rich DNA Polymerase which has been formulated for use with difficult GC-rich templates (>65% GC content). Alternatively, our PCRx Enhancer System can be used in conjunction with DNA polymerases, including native/recombinant Taq, Platinum® Taq, and Platinum® Taq High Fidelity to optimize PCR of problematic and/or GC-rich templates. Finally, AmpliTaq Gold® 360 with the 360 GC Enhancer buffer has been shown to work with templates containing up to 80% GC.

We strongly suggest using MgSO4. While MgCl2 may work in some cases, MgSO4 usually produces more robust and reproducible products, as sulfate is the best anion found for the Platinum® Taq High Fidelity enzyme.

With Platinum® technology, anti-DNA polymerase antibodies bind to the enzyme until the denaturing step at 94°C, when the antibodies degrade. The polymerase is now active and primer extension can occur. AccuPrime™ Taq combines Platinum® Taq (Taq + Platinum® antibodies) with proprietary thermostable AccuPrime™ accessory proteins. The 10X reaction buffer contains the accessory proteins which enhance specific primer-template hybridization during each cycle of PCR.

Both AmpliTaq Gold® and Platinum® Taq are hot-start enzymes that allow you to set up your reactions on the benchtop without the need for ice. AmpliTaq Gold® is a chemically-modified hot-start enzyme, provided in an inactive state. Heat activates the enzyme, with full activity after 10 min at 95°C. Platinum® Taq is an antibody-mediated hot-start enzyme. The anti-Taq antibodies bind and inactivate the enzyme, until the heat denaturation step of the PCR reaction (30 sec to 2 min), which activates the enzyme.

Yes, the enzyme mix leaves 3′ A-overhangs on a portion of the PCR products. However, the cloning efficiency is greatly decreased compared to that obtained with Taq polymerase alone. It is recommended to add 3′ A-overhangs to the product for TA cloning.

In our experience, the Gold Buffer gives the best performance. Here are the formulations:
GeneAmp® 10X PCR Buffer: 100 mM Tris-HCl, 500 mM KCl, 15 mM MgCl2, pH 8.3, 0.01% (w/v) gelatin
GeneAmp® 10X PCR Buffer II*: 100 mM Tris-HCl, 500 mM KCl, pH 8.3
GeneAmp® 10X PCR Buffer Gold*: 150 mM Tris-HCl, 500 mM KCl, pH 8.0
*Buffer comes with separate MgCl2 Solution (25 mM).

PCR Reaction

The main steps are: denaturation, annealing, and extension. The template is typically heated to a high temperature (around 94–95°C) allowing for the double-stranded DNA to denature into single strands. Next, the temperature is lowered to 50–65°C, allowing primers to anneal to their complementary base-pair regions. The temperature is then increased to 72°C, allowing for the polymerase to bind and synthesize a new strand of DNA.

Hot start is a way to prevent DNA amplification from occurring before you want it to. One way to do this is to set up the PCR reaction on ice, which prevents the DNA polymerase from being active. An easier method is a use a ‘hot-start’ enzyme, in which the DNA polymerase is provided in an inactive state until it undergoes a high-heat step.

The MgCl2 should be optimized for each template and primer pair. In general, the final concentration varies between 0.5–2.5 mM (when using 0.2 mM dNTPs). Note: EDTA or excess dNTPs can inhibit amplification by chelating the magnesium ions necessary for Taq DNA polymerase activity.

While the volume is dependent on the starting amount of RNA used for the first-strand synthesis and the abundance of the target gene, we’d recommend starting with 10% of the first-strand reaction for your PCR reaction.

Touchdown PCR involves decreasing the annealing temperature by 1°C every second cycle to a ‘touchdown’ annealing temperature which is then used for the remaining cycles. Touchdown PCR increases specificity and reduces background amplification. By starting at a high annealing temperature, only your gene of interest is amplified, allowing the target product to accumulate. Decreasing the annealing temperature through the remaining PCR cycles permits efficient amplification of the specific template.

Nested PCR requires two separate amplifications—the first one using one set of PCR primers and the second one using internal "nested" primers plus 1% or less of the first PCR reaction as a template. Nested PCR is used when the target is present in low abundance or when nonspecific PCR products are being produced along with the specific product. Semi-nested PCR is used when there is only enough sequence information to make a primer internal to one end of the primary PCR product such as in RACE (rapid amplification of cDNA ends).

Typically, primer annealing temperature is 3–5°C lower than the lowest primer melting temperature. If the temperature is too high, primers anneal poorly. If the temperature is too low, nonspecific annealing can occur.

A GC-rich template often has a higher melting temperature and may not denature completely under the normal reaction conditions.

You can try adding 5–10% DMSO, up to 10% glycerol, or 1–2% formamide or a combination of these to facilitate difficult templates. Note: the use of cosolvents will lower the optimal annealing temperatures of your primers.

You may choose to do a two-temperature protocol when the annealing temperature is relatively high. In this case, you would combine the annealing and the elongation steps, i.e., both can occur together at a temperature >62°C. The advantage of a two-temperature protocol is that it is considerably quicker in comparison to the conventional three-temperature protocol.

Primers and Oligos

These guidelines may be useful as you design your PCR primers:

  • In general, a length of 18-30 nucleotides for primers is good.
  • Try to make the melting temperature (Tm) of the primers between 65°C and 75°C, and within 5°C of each other.
  • If the Tm of your primer is very low, try to find a sequence with more GC content, or extend the length of the primer a little.
  • Aim for the GC content to be between 40 and 60%, with the 3′ of a primer ending in C or G to promote binding.
  • Typically, 3 to 4 nucleotides are added 5′ of the restriction enzyme site in the primer to allow for efficient cutting.
  • Try to avoid regions of secondary structure, and have a balanced distribution of GC-rich and AT-rich domains.
  • Try to avoid runs of 4 or more of one base, or dinucleotide repeats (for example, ACCCC or ATATATAT).
  • Avoid intra-primer homology (more than 3 bases that complement within the primer) or inter-primer homology (forward and reverse primers having complementary sequences). These circumstances can lead to self-dimers or primer-dimers instead of annealing to the desired DNA sequences.
  • If you are using the primers for cloning, we recommend cartridge purification as a minimum level of purification.
  • If you are using the primers for mutagenesis, try to have the mismatched bases towards the middle of the primer.
  • If you are using the primers for a PCR reaction to be used in TOPO® cloning, the primers should not have a phosphate modification.

Read more about primer design tips and tools here.

Yes. OligoPerfect™ Designer can be used to design primers for sequencing, cloning, or detection.

Centrifuge the tube for a few seconds to collect the oligonucleotide at the bottom of the tube. Carefully open the tube, and dissolve the oligonucleotide in the appropriate volume of TE (10 mM Tris-HCl, pH 8.0, 1 mM EDTA). TE is recommended over deionized water since the pH of water is often slightly acidic and can cause hydrolysis of the oligonucleotide. It is also best to pipette the solution up and down at least 10 times. Please visit this webpage for more information on how to calculate primer concentration and resuspension volume.

Oligonucleotides from Life Technologies™ come lyophilized. It is recommended to store lyophilized (and reconstituted) oligos at -20°C. Lyophilized oligonucleotides are stable at -20°C for at least 1 year. Oligonucleotides dissolved in TE are stable for at least 6 months at -20°C or 4°C. Dissolved in water they are stable for at least 6 months at -20°C in the absence of nucleases. Lyophilized oligos should be stable for at least a few months at RT. Oligos in solution are stable for at least 6 months if they are stored at -20°C at a concentration greater than 10 μM. If the oligos are stored at 4°C, it is important that they are resuspended in TE, NOT water. (Note: at 4°C in water, the oligo will hydrolyze over time). AP and HRP conjugates are shipped in their respective storage buffers. Please store them at 4°C.

Depending on your specific application, a different level of purification may be required. Please refer to this table.

Oligos are made using a DNA synthesizer which is basically a computer-controlled reagent delivery system. The first base is attached to a solid support, usually a glass or polystyrene bead, which is designed to anchor the growing DNA chain in the reaction column. DNA synthesis consists of a series of chemical reactions.

  1. Deblocking: the first base, attached to the solid support via a chemical linker arm, is deprotected by removing the trityl protecting group. This produces a free 5′ OH group to react with the next base.
  2. Coupling: the next base is added, which couples to the first base.
  3. Capping: any of the first bases, which fail to react, are capped. These failed bases will play no further part in the synthesis cycle.
  4. Oxidation: the bond between the first base and successfully coupled second base is oxidized to stabilize the growing chain.
  5. Deblocking: the 5′ trityl group is removed from the base, which has been added.

The scale of synthesis is the starting point for synthesis, not the guaranteed final amount. We guarantee the total yield of oligonucleotide as a minimum number of OD units. Use this link for the minimum yield guarantees we offer for our oligos.

Coupling efficiency is the major factor affecting the length of DNA that can be synthesized. Base composition and synthesis scales will also be contributing factors. At 99% coupling efficiency, a crude solution of synthesized 95-mers would contain 38% full-length product and 62% (nx) failure sequences. This is before other chemical effects have been taken into account such as depurination. Depurination mainly affects the base A. The frequency of depurination is small but will increase significantly with primer length. For these reasons, we specify a maximum length of 100 bases, which we believe is the maximum length that can be synthesized routinely and economically.

Coupling efficiency is a way of measuring how efficiently the DNA synthesizer is adding new bases to the growing DNA chain. If every available base on the DNA chain reacted successfully with the new base, the coupling efficiency would be 100%. Few chemical reactions are 100% efficient. During DNA synthesis, the maximum coupling efficiency obtainable is normally around 98–99% (99% is typical). This means that at every coupling step, approximately 1–2% of the available bases in the chain fail to react with the new base being added. An approximation of the percentage of full-length oligonucleotide is obtained by the coupling efficiency raised to the power of its length (i.e., number of cycles), e.g., 0.9922 x 100 = 80% full-length primer. You can have your primers further purified to 95% full-length. Purification is highly recommended for long oligos. For example, a 64-mer synthesis will yield 0.9964 x 100 = 53% full-length. This will only be obtained if the efficiency is 99% at every cycle.

The trityl group is colorless when attached to a DNA base but gives a characteristic orange color once removed. The intensity of this color can be measured by UV spectrophotometry and is directly related to the number of trityl molecules present. By comparing the absorbance of trityl releases throughout synthesis, it is possible to calculate the percentage of bases coupling successfully and hence the coupling efficiency.

Coupling efficiency is important as the effects are cumulative during DNA synthesis. The table below shows the effect of a 1% difference in coupling efficiency and how this influences the amount of full-length product available following synthesis of different length oligos. Even with a relatively short oligo of 20 bases, a 1% difference in coupling efficiency can mean 15% more of the DNA present following synthesis is full-length product.

Number of
bases added
99% coupling
full-length
Failures 98% coupling
full-length
Failures
1 99 1 98 2
2 98.01 1.99 96.04 2.96
3 97.03 2.97 94.12 5.88
10 90.44 9.56 81.71 18.29
20 81.79 18.21 66.76 33.24
30 73.79 26.03 54.55 63.58
50 60.5 39.5 36.42 63.58
95 38.49 61.51 14.67 85.33

The percentage of full-length oligonucleotide depends on the coupling efficiency of the chemical synthesis. The average efficiency is close to 99%. To calculate the percentage of full-length oligonucleotide, use the formula: 0.99n-1. Therefore, 79% of the oligonucleotide molecules in the tube are 25-bases long; the rest are <25 bases. If you are concerned about starting with a preparation of oligonucleotide that is full-length you may want to consider cartridge, PAGE, or HPLC purification.

Please take a look at this list of standard modification options that we offer. If you do not see the modification option you would like, please email our Technical Support team at techsupport@lifetech.com to see if we can accommodate your request.

A common equation used to calculate primer Tm is as follows:

Tm (in °C) = 2 (A+ T) + 4 (G + C)

An important parameter for primers is the melting temperature Tm. This is the temperature at which 50% of the primer and its complementary sequence are present in a duplex DNA molecule. The Tm is necessary to establish an annealing temperature for PCR. Reasonable annealing temperatures range from 55°C to 70°C. Annealing temperatures are generally about 5°C below the Tm of the primers. Since most formulas provide an estimated Tm value, the annealing temperature is only a starting point. Specificity for PCR can be increased by analyzing several reactions with increasingly higher annealing temperatures.

A GC-rich template often has a higher melting temperature and may not denature completely under the normal reaction conditions.

You can try adding 5–10% DMSO, up to 10% glycerol, or 1–2% formamide or a combination of these to facilitate difficult templates. Note: the use of cosolvents will lower the optimal annealing temperatures of your primers.

You may choose to do a two-temperature protocol when the annealing temperature is relatively high. In this case you would combine the annealing and the elongation steps, i.e., both can occur together at a temperature >62°C. The advantage of a two-temperature protocol is that it is considerably quicker in comparison to the conventional three-temperature protocol.

Value Oligos are the most cost-effective and fastest way to order oligos. They are available for 5–40-mers, at a 25 or 50 nanomole scale, with a range of purification options to suit your needs, and are eligible for next-day delivery. The cost is calculated per oligo as opposed to per base. Value Oligos are not available with modifications. Value Oligos undergo the same QC standards as our standard oligos with the same manufacturing process.

As oligos increase in length, the column purification is less effective in separating the failure oligos from the correct products. PAGE purification would be the method of choice in this case.

Tm values are not absolute—they are an approximation of the melting temperature range which exists. A thermal profile for a given oligo shows a 10–15 degree range of melting depending on the amount of salt but also on the base composition and concentration of primer in the reaction which are not precisely defined. One should not rely solely on the given Tm value as the only one that will work. Tm is the temperature at which 50% of the primer and its complementary sequence are present in a duplex DNA molecule. The Tm is necessary to establish an annealing temperature for PCR. Reasonable annealing temperatures range from 55°C to 70°C. Annealing temperatures are generally about 5°C below the Tm of the primers. Since most formulas provide an estimated Tm value, the annealing temperature is only a starting point. Specificity for PCR can be increased by analyzing several reactions with increasingly higher annealing temperatures.

The plate orders must contain an average of 24 or more oligos per plate for 96-well plates or 192 or more oligos per plate for 384-well plates across the entire order.

For 25, 50, and 200 nmol desalted and cartridge-purified DNA oligos, there is 100% A260 analysis. Random samples of 25% of the oligos produced are tested by either capillary electrophoresis or mass spectrometry. DNA oligos that are desalted and ordered at 25 and 50 nmol scales also have 100% real-time digital trityl monitoring during analysis. Desalted DNA oligos ordered at 1 and 10 μmols, DNA oligos at any scale that are purified by HPLC and PAGE, the majority of the DNA oligos with 3′ and/or 5′ modifications, and RNA oligos have 100% A260 analysis and capillary electrophoresis or mass spectrometry.

No, we do not guarantee 50/50 of mixed bases. If a mix of GC bases is requested, for example, the synthesizer would deliver half the normal amount of G and half the normal amount of C. Coupling efficiency is not taken into account. Therefore, it is possible that a mix, such as 30/70, will be delivered.

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