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View additional product information for Custom DNA Tube - FAQs (10336022)
21 product FAQs found
Yes, we can synthesize primers with degenerate bases. Please see here for the electronic ordering code to use (https://www.thermofisher.com/us/en/home/products-and-services/product-types/primers-oligos-nucleotides/invitrogen-custom-dna-oligos/oligo-ordering-details/oligo-modification-options.html#2).
We offer TE Buffer, pH 8.0 (Cat. Nos. AM9849 and AM9858), that consists of 10 mM Tris (adjusted to pH 8.0 with HCl) and 1 mM EDTA.
Unless you specifically request that primers be supplied in solution, they are shipped lyophilized. We recommend reconstituting them in the appropriate volume of TE (10 mM Tris-HCl, 1 mM EDTA, pH 8.0).
No, the primer will not have this modification. You must specify that it should have a 5‘ phosphate modification.
It is important to differentiate naturally occurring mutations linked to the chemical nature of the oligo manufacturing process from the perceived mutations that occur when desalted oligos are used in certain applications.
If the mutation you find is not consistent in all of your clones, this probably falls into the category of a "perceived" mutation that is actually an expected by-product of an oligo reaction. This has a few causes:
-Following DNA synthesis, the completed DNA chain is released from the solid support by incubation in basic solutions such as ammonium hydroxide. This solution contains the required full-length oligo but also contains all of the DNA chains that were aborted during synthesis (failure sequences). If a 30-mer was synthesized, the solution would also contain 29-mer failures, 28-mer failures, 27-mer failures etc. The amount of failure sequences present is influenced by the coupling efficiency. For an oligo of this type, the percentage of full-length oligo would be between 74 and 54%, assuming a 99 or 98% coupling efficiency (the percentage of full-length oligos produced from a given synthesis declines as the length of the oligo increases). The truncated oligos usually result because oligos are synthesized from 3' to 5' end. Primers that are desalted and not purified for length will have truncated product missing one or more bases at the 5' end. Hence, oligos that are desalted are only recommended for diagnostic PCR, microarrays, or sequencing. We recommend purification of the oligos if they will be used in certain demanding applications such as mutagenesis or cloning, especially if restriction sites are added to the 5' end of them.
-Other sources of perceived mutations for both desalted and purified oligos are sequencing artifacts, point mutation introduced during PCR, unstable stem loop structures in the primers, propagation of the plasmid DNA after cloning in an E. coli strain that is muS, mutD or mutT or a silent mutation selected by the bacterial strain because of codon usage in that strain.
Naturally occurring mutations are a rare but inherent event in the chemical synthesis of the oligos. The chances of having one single insertion or deletion in a given oligo of about 30 bases is about 2%.
For a given oligonucleotide, the extinction coefficient and A260/280 ratio depend on the sequence (each nucleotide has its own absorbance characteristics). For accurate determination of oligonucleotide concentration, calculate the extinction coefficient using an equation that incorporates the contribution of each base and the effect of base stacking [Reference: Newton, P.R. (1995) PCR Essential Data, John Wiley and Sons, New York, New York. p.55.]
Also, because the A260/280 ratio of a given oligo depends on the sequence, that measurement is not a reliable indicator of purity.
While not the case for large DNA molecules, the migration of oligonucleotides in polyacrylamide gels is affected by the sequence and composition. Comparing an oligo to a molecular size standard or another oligonucleotide of known size is not a reliable way to determine its size.
The addition of 40% formamide (in addition to 8 M urea) to the acrylamide gel solution removes most sequence-dependent migration effects. [Reference: Ausubel, F.M., Brent, R., Kingston, R.E. Moore, D.D., Seidman, J.G., Smith, J. A., and Struhl, K. (1994) Current Protocols in Molecular Biology, John Wiley and Sons, Inc., New York, NewYork.]
In addition, the use of ethidium bromide for detection and gel densitometry-based quantitation of an oligonucleotide is not reliable. The ability of ethidium bromide to stain oligonucleotides is poor and varies depending on sequence and composition.
Coupling efficiency is the major factor dictating the length of DNA that can be synthesized. Base composition and synthesis scales are also contributing factors. For example, at 99% coupling efficiency, a crude solution of synthesized 95-mers would contain 38% full-length product and 62% failure sequences. This calculation doesn't take into consideration other chemical side reactions that lead to failed sequences such as depurination. The frequency of depurination is small but the effects of depurination have an increasing impact as the length of the primer increases. 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.
Following DNA synthesis, the completed DNA chain is released from the solid support by incubation in basic solutions such as ammonium hydroxide. This solution contains the required full-length oligo but also contains all of the DNA chains that were aborted during synthesis (failure sequences). If a 20-mer was synthesized, the solution would also contain 19-mer failures, 18-mer failures, 17-mer failures etc. The amount of failure sequences present is influenced by the coupling efficiency. These failure sequences can compete with the full-length product in applications such as PCR, and may need to be removed before the oligo can be used successfully.
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 the coupling efficiency.
Every DNA base added during DNA synthesis has a dimethoxy-trityl (Trityl) protecting group attached. This Trityl group protects the base from undergoing unwanted chemical reactions during the synthesis cycle and is only removed immediately before a new base is added.
Coupling efficiency is a measurement of 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 effiency raised to the power of its length (i.e. number of cycles), e.g. 0.99^22 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 0.99^64 x 100 = 53% full-length primer (and keep in mind that 53% full-length primer is, and this result is based on a 99% efficiency at every cycle.
DNA synthesis is a complicated process that has improved significantly over the last 10 years. Despite these improvements, all manufacturers have an inherent failure rate in their oligo synthesis runs. We are constantly developing our processes and systems to minimize these losses, however it is inevitable that we will occasionally have to re-make some oligos. When ordering, you can choose whether you would like to receive partial orders or not, on the ordering form.
The phosphorothioate modification is actually a modification to the normal phosphodiester bond between the bases. The codes used to indicate phosphorothiate bases are F, O, E, and Z, representing A,C,G, and T respectively. For instance, "E" is used to specify a G followed by a downstream phosphorothioate modification linkage that links the G to the downstream (next) base. Therefore, chemically it is not possible to have a "phosphorothioate" as the 3' most base, since there is no base downstream of it. The F, O, E, or Z codes should therefore not be used in the 3' most position when placing an order for S-oligos. Use the normal A, C, G, or T codes in this position.
There are two options after deprotection with ammonium hydroxide, and both work well.
(1) The primers can be lyophilized. This will take around 6-8 hours. Then solubilize them in TE or water.
(2) Alternatively the primers can be desalted directly using Sephadex G-25 gel filtration medium.
Primers last a long time if stored in buffer at -20 degrees C. They last indefinitely when lyophilized.
No. Longer primers work fine. Generally, 50 nmol of desalted-purity oligos are adequate for most applications. Oligos should be adjusted to a concentration of 20-50 µM and the concentration should be verified by spectrophotometry. For cloning large PCR products (>5 kb), colony output can be increased if long primers (>65 bases) are further purified (i.e., cartridge purification or PAGE). Go to https://www.thermofisher.com/us/en/home/products-and-services/product-types/primers-oligos-nucleotides/invitrogen-custom-dna-oligos.html for more information.
Example 1: the COA specifies we have 24 nmole of oligo. If we resuspended oligo in 1 mL:
1 ml = 0.001 L
24 nmole/0.001 L = 24000 nmole/L or 24000 nM
24000 nmole/L X 1 µmole/1000 nmole = 24 µmole/L or 24 µM
Explanation: We convert the volume in which the oligo was resuspended into liters. Then the total nmole amount of oligo is divided by the volume to get nM concentration. The nmoles are converted to µmole to get the µM concentration.
Example 2: making 100 µM primer stock.
If the COA specifies we have 24 nmole of oligo:
24 nmole X 1 µmole/1000 nmole = 0.024 µmole
0.024 µmole/100 µmole/liter = 0.00024 L
0.00024 L X 1000 mL/L = 0.24 mL or 240 µL
Explanation: We convert from nmole to µmole then divide by the desired concentration in µmole/L. The µmoles cancel out giving the needed volume in liters. We then convert liters to mL. So in this example the oligo should be suspended in 0.24 mL to get a 100 µM solution.
Example 3: Calculate from OD.
If the primer OD is 0.14 and the µg/OD reported on the COA is 36.6:
0.14 OD/mL X 1000 µL/10 µL = 14 OD/mL stock
14 OD/mL X 36.6 µg/OD = 512.4 µg/mL
Explanation: The OD/mL is multiplied by the dilution factor to get the stock OD/mL. The OD is converted to µg/mL by multiplying the OD/mL of the stock by the µg/OD conversion factor listed on the COA. The µg cancel out giving µg/mL.
If the primer OD is 0.14 and the nmole/OD reported on the COA is 4.9:
0.14 OD/mL X 1000 µL/10µL = 14 OD/mL stock
14 OD/mL X 4.9 nmole/OD = 68.6 nmole/mL
68.6 nmole/mL X 1000 mL/L = 68600 nmole/L or 68600 nM
68600 nmole/L X 1 µmole/1000 nmole = 68.6 µmole/L or 68.6 µM
Explanation: The OD/mL is multiplied by the dilution factor to get the stock OD/mL. The OD is converted to nmole using the conversion factor on the COA. Then mL are converted to liters and nmole are converted to µmole to get the µM concentration.
Example 4: Calculate from MW. The COA specifies a molecular weight for the oligo of 7440.0:
7440.9 g/mole = 7440.9 µg/µmole
7440.9 µg/µmole X 68.6 µmole/L = 510445.74 µg/L
510445.74 µg/L X 1 L/ 1000 mL = 510.4 µg/mL
Explanation: g/mole is the same as µg/umole. The molecular weight expressed in µg/umole is multiplied by the µM concentration determined in example 3. The µmole cancel out leaving µg/L. Liters are converted to mL to give the µg/mL concentration.
The MgCl2 should be optimized for each template and primer pair. In general the final concentration varies between 0.5 and 2.5 mM (when using 0.2 mM dNTPs). EDTA or excess dNTPs can inhibit amplification by chelating the magnesium ions necessary for Taq DNA polymerase activity.
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. Typical annealing temperatures range from 55-70 degrees C. Annealing temperatures are generally about 5 degrees C below the Tm of the primers. Since most formulas provide an estimated Tm value, the calculated annealing temperature is only a starting point. Troubleshoot PCR specificity problems by performing several PCR reactions with increasingly higher annealing temperatures and selecting conditions that amplify only the product you're interested in.
The annealing temperature should be 5 degrees C below the melting temperature (Tm) of the amplification primers. Temperatures in the range of 55-72 degrees C generally yield the best results. The Tm can be determined by most computer programs used to design primers. The general formula Tm = 4(G+C) + 2(A+T) may be used to determine Tm of oligonucleotides less than 10 bases in length. For oligonucleotides greater than 10 bases, use the following formula: [67.5 + 0.34(%GC as a whole number) - 395/length of the oligonucleotide]
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 (https://www.thermofisher.com/us/en/home/life-science/oligonucleotides-primers-probes-genes/custom-dna-oligos/oligo-technical-resources/oligo-protocols.html) for more information on how to calculate primer concentration and resuspension volume.
The lyophilized oligonucleotide is stable at -20 degrees C for 1 year or more. The oligonucleotide dissolved in TE is stable for at least 6 months at -20 degrees C or 4 degrees C. Oligonucleotides dissolved in water are stable for at least 6 months at -20 degrees C. Do not store oligonucleotides in water at 4 degrees C.
Primers are lyophilized and shipped at room temperature in a 2 mL screw cap, clear, polypropylene tubes. Large orders (over 48 primers) may be arrayed in 1-mL 96-well polypropylene plates. Fluorescent-labeled primers are supplied in amber screw cap polypropylene tubes. The alkaline phosphatase- and horseradish peroxidase-labeled primers are supplied in solution in a storage buffer unique to the enzyme label.