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Frequently asked questions

Can you please enlighten me on the base degenaracy codes that you use, eg. R, W, K, V, S

The standard IUB codes for degenerate bases are:


A = Adenosine;

C = Cytosine;

G = Guanosine;

T = Thymidine;

B = C, G, or T;

D = A, G, or T;

H = A, C, or T;

V = A, C, or G;

R = A or G (puRine);

Y = C or T (pYrimidine);

K = G or T (Keto);

M = A or C (aMino);

S = G or C (Strong -3H bonds);

W = A or T (Weak - 2H bonds);

N = aNy base.


I used oligos for cloning, sequenced a clone and found a mutation within the oligo sequence. Can mutations happen in synthetic oligos?


Mutations or alternations in sequence are rare in synthetic oligos but they can happen. Oligos may become depurinated in some positions during the synthesis process. The longer the oligonucleotide, the more depurinated sites may occur. These depurinations may become visible in applications like cloning where single molecules are selected and propagated. Typically the depurinated sites are replaced by any base. Usually only some oligonucleotide molecules are affected. If you have sequenced only one clone and found a mutation in it, sequencing another independent clone will in most cases result in the correct sequence. When choosing clones, it is most important to select independent ones: To get independent clones, the pre-incubation time before plating should be based on the manufacturer instructions of your cloning system. Optimal density of the clones on the plate ensures that well separated clones can be selected.


Longer oligos used for cloning should be purified to remove any truncated sequences. As the risk of depurinations is increasing with the length of the oligonucleotide, we recommend selecting sequences for cloning that are as short as possible. If long oligos (>40 mer) have been used for cloning, consider increasing the number of clones that need to be sequenced.


Please note that synthetic oligos do not contain a phosphate at the 5' end which is necessary for enzymatic ligation reactions. The phosphate can be added as a modification to the 5' end of the oligo.


Do fluorescent dye-labeled oligos require special storage and handling?


Yes. Fluorescent dye-labeled oligos are more fragile than unmodified oligos. Their ability to fluoresce will decrease over time. Before opening, spin the tube at 1000 rpm for 5 minutes to ensure the oligos are at the bottom of the tube. To ensure high quality, store the oligo dry at -20℃ in small aliquots. Note also that fluorescent dye-labeled oligos should be stored in the dark as light can slowly degrade the fluorescent moieties.

For optimal long-term storage, it is recommended that the labeled oligonucleotides should be stored dry at -20℃ in the dark. If numerous experiments are planned using the same oligonucleotide, prepare aliquots, dry all aliquots, and store the aliquots at -20℃. Use clean, sterile labware for all transfers.


How long will my fluorescent dye-labeled oligos last?


Fluorescent dye-labeled oligos can be used up to 6 months from purchase when stored dried at -20℃ in the dark.


How should I resuspend my fluorescent dye-labeled oligos?


Oligonucleotides labeled with fluorescent dyes, such as FAM, HEX, TET, JOE, and TAMRA should be resuspended in a slightly basic solution (e.g., TE at pH 8.0). Cy3 and Cy5 labeled oligonucleotides should be resuspended at pH 7.0, and they begin to degrade at pH <7.0. If brought to a pH below 7, it has been shown that the oligo can begin to degrade and may be unusable within a few weeks.


Why are my oligos colored?


Synthetic oligos vary in appearance depending on a large number of factors. The lyophilized oligo may appear dry or oily with a clear, white, or brownish color. A colored product is not a sign of impurities and should not be a cause for concern. All of SBS’s oligo products meet strict quality control standards before being shipped to our customers.


How long will my oligos last and how should I store them?


Unmodified oligos are stable molecules and can be used for at least 12 months after purchase when stored at -20℃. For long-term storage, oligos should be stored dry at -20℃. If numerous experiments are planned using the same sequence, aliquot the sample, dry all aliquots, and store at -20℃. If the oligos are stored wet, avoid repeated freeze-thaw cycles as this process can lead to physical degradation of the oligo. Oligos generally last longer in TE than in water. Careful handling is recommended to avoid the possibility contamination with nucleases or bacteria.


How should I resuspend my oligos?


Before opening, spin the tube at 1000 rpm for 3 min to ensure that the oligonucleotides are at the bottom of the tube. It is recommended that the oligonucleotides should be resuspended in a sterile buffered solution (e.g., TE at pH 7.0). Please vortex your oligonucleotides thoroughly after resuspension. The oligonucleotides may not readily dissolve in sterile, distilled water. If NaOH is added to the water, the pH will rise to 7.0 and this should help. If the oligonucleotides are resuspended at pH <7.0 (deionized water may have a pH as low as 5.0), the oligonucleotide could begin to degrade and may lose functionality within a couple of weeks.


For optimal long-term storage, it is recommended that the oligonucleotides should be stored dry at -20℃ in the dark. If numerous experiments are planned using the same oligonucleotide, prepare aliquots, dry them and store the aliquots at -20℃. Use clean, sterile labware for all transfers.


How are oligonucleotides quantified?


Oligonucleotides are most commonly quantified by measuring their UV absorbance at 260 nm. A single-stranded oligonucleotide dissolved in a neutral aqueous solution at a concentration of 33 µg/ml will have an absorbance at 260 nm of approximately 1.0 A.U. The optical density (OD) unit is defined as the absorbance (at 260 nm in a 1 cm cuvette) multiplied by the sample volume (in milliliters), so that 1 OD unit is roughly 33 µg of ssDNA.


Why are my measured yields lower than the ones provided by the company?


The most common reason for this is that the oligos are not fully dissolved. If you resuspended in water, you may need to add NaOH to raise the pH to between 7 and 8. Oligos resuspended in water tend to have a pH of around 5, and this is often too low for them to completely dissolve. We recommend that most oligos be resuspended in TE at pH 7. We find that 10 mM Tris-HCl, 1 mM EDTA works well, but if your experiment cannot tolerate EDTA, you may use just Tris-HCl.


Why do I see extra bands in my PCR product?


Complementary primers can cause PCR artifacts, especially in control lanes (without template). The primers anneal to each other and extend, sometimes creating large products. The extra bands that appear in the control lane often disappear in the presence of competing template. Depending on the degree of complementarity, raising the annealing temperature may help if there are extra bands in other lanes.


Why did my PCR reaction stop working?


If your PCR reaction used to work and then stopped, the primers or template may have become degraded. The primers can last up to 1 year if stored lyophilized and aliquoted at -20℃. Freezing and thawing causes the primers to degrade. Primers or template can also degrade due to contamination. Millipore filtration systems, which require the filter to be changed periodically, can supply water of questionable quality at the end of a cycle. If you are using a filter system, check to see if your filter was changed soon after resuspending your oligos. You may want to consider changing the filters more often if this is the case.


What are some design guidelines for PCR Primers and is there a tool to help me with this?


In PCR, it is important to design the two primers with similar melting temperatures to ensure efficient annealing. There is a trade-off in PCR primer design between sequence-specific primers and primer length. A primer needs to be long enough to be specific for its target region, yet not too long since longer primers are less efficient at annealing. It is also important to ensure that each primer does not form hairpins or dimers with itself or the other PCR primer. Dimerization and hairpin formation prevent the primers from participating in the amplification reaction and may contribute to nonspecific products if a 3' overhang is formed. Careful consideration should be given to avoid mis-priming. If the selected primer shares homology with other areas of the target region, mis-priming can generate multiple undesired PCR products. Note: The rules of primer design are based on theoretical considerations. The true test of a primer’s function is its use in a functional assay. Usually, a primer length of 18~30 bases is optimal for most PCR applications. This is based on the complexity of the target template DNA. Theoretically, a primer of 18 bases represents a unique DNA sequence amongst 418 = 7×1010 nucleotides and should hybridize at only one position in most eukaryotic genomes which consist of approximately 109~1010 base pairs. A shorter primer such as a 15mer would have a greater chance of annealing at more than one complementary site within the genome. This may lead to amplification of nonspecific PCR products. In contrast, when using the same shorter primer with a less complex DNA template such as plasmid DNA, PCR will only generate the specific PCR product.


Design Guidelines for Sequencing Primers.


SBS strongly recommends the use of software to design DNA sequencing primers. The Oligo Toolkit and Plotter display secondary structure as well as other useful information such as melting temperature, GC content, etc. For cycle sequencing, primer length should be between18 and 28 bases with a GC content of at least 50%. The melting temperature of the primer should be between 50℃ and 70℃. It is important to avoid selecting primers that form dimers or hairpins as molecules in this conformation will not participate in primer template hybridization. Additionally, if a 3' overhang is formed by a dimer or hairpin, the polymerase will extend and form unwanted products.


Design Guidelines for Dual-Labeled Probes and Primers for 5' Nuclease Assay.


Dual-labeled probes (fluorescence quencher or FQ probes) have a fluorescent reporter and a quencher at their 5' and 3' ends, respectively. These probes can be used in quantitative PCR systems that take advantage of the 5'→3' exonuclease activity of Taq DNA polymerase). A probe specific for the sequence of interest is used together with specific PCR primers. This probe is designed to anneal between the PCR primers. During the extension phase of PCR, the 5'→3' exonuclease activity of Taq DNA polymerase cleaves the fluorescent reporter from the probe. The amount of free reporter accumulates as the number of PCR cycles increases. The fluorescent signal from the free reporter is measured in real-time and allows quantification of the amount of target sequence. There are several design considerations to keep in mind when designing dual-labeled probes. Placement of the probe is important; the probe should be designed first, followed by the design of the PCR primers. The probe should anneal near the center of the amplicon, and the amplicon should be 50 to 150 bases long. The probe's melting temperature should be 68℃ to 70℃. The probe should be at least 20 bases long to prevent nonspecific annealing. Finally, avoid designing the probe with a dG at the 5' terminus as dG is a weak quencher. Any dG in the primer should be at least 2 residues away from the 5' terminus.



Probe:

G-C Content: 30~80%

polyNNN: Avoid stretches same nucleotide, especially 4 or more Gs

End Composition: No Gs at 5' end

Tm: 68-70℃

Strand: Select strand with more Cs than Gs. If the compliment is used, make sure there are no Cs at the 3' end.

Length: Less than 30 bp

Design: Quencher on 3' end dye on 5' end

Primer:

G-C Content: 30-80%

polyNNN: Avoid stretches of same nucleotide, especially 4 or more Gs

End Composition: 5 bases at 3' end should have no more than 2 G/Cs. Try to use As at 3' end so primer dimers will be degraded more efficiently.

Tm: 58-60℃

Product: 50 to 150 bases

Design: Choose after probe. Design as close to probe as possible without overlap. Have one primer cross one exon junction to amplify mRNA.


Design guidelines for Molecular Beacons.


Molecular Beacons also have a fluorescent reporter and a quencher at their 5' and 3' ends, respectively. The sequence of these probes is designed so that they form a hairpin structure in which the fluorescent dye and the quencher are in close proximity. A Molecular Beacon specific for the sequence of interest is used in PCR. The probe is designed to anneal between the PCR primers. When the probe hybridizes to its target sequence in the PCR annealing step, the loop opens and the fluorescent reporter and quencher are separated, resulting in a fluorescent signal. The amount of signal is proportional to the amount of target sequence, and is measured in real-time to allow quantification of the amount of target sequence. In order to successfully monitor PCR, Molecular Beacons should be designed so that they are able to hybridize to their targets at the annealing temperature of the PCR, whereas the free Molecular Beacons should remain closed and be non-fluorescent at these temperatures. This can be ensured by choosing the length of the probe sequence and the length of the arm sequences appropriately. The length of the probe sequence should be such that it will dissociate from its target at temperatures 7~10℃ higher than the annealing temperature of the PCR. The melting temperature of the probe target hybrid can be predicted using the "percent-GC" rule, which is available in most probe design software packages. The prediction should be made for the probe sequence alone before adding the stem sequences. In practice, the length of the probe sequence is usually in the range from 15~30 nucleotides.



The schematic diagram above represents the principle of Molecular Beacons in quantitative, realtime PCR. A: When not bound to its target sequence, the Molecular Beacon forms a hairpin structure. The proximity of the fluorescent reporter with the quencher prevents the reporter from fluorescing. B:During the PCR annealing step, the Molecular Beacon probe hybridizes to its target sequence. This separates the fluorescent dye and reporter, resulting in a fluorescent signal. The amount of signal is proportional to the amount of target sequence, and is measured in real-time to allow quantification of the amount of target sequence.


·         Tm of probe + target hybrid (loop only) has to be 7~10℃ above Tanneal

·         Tm of stem is 7~10℃ higher than the detection temperature (=annealing temperature of the PCR)

·         The loop of the probe should be 15~30 bases

·         The stem of the probe should be 5~7 bases on each side


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