CleanAmp™ dUTP is a dUTP analog which contains the CleanAmp™ thermolabile protecting group for improved PCR specificity.
How do I use CleanAmp™ dUTP?
Optimal PCR results are achieved when CleanAmp™ dUTP is in a three-fold excess over dATP, dCTP and dGTP. Use CleanAmp™ dUTP with standard dNTPs for well-behaved targets and with CleanAmp™ dNTPs for problematic targets.
Is CleanAmp™ dUTP compatible with carryover decontamination schemes?
CleanAmp™ dUTP can be used with uracil-N-glycosylase (UNG) in PCR-based carryover decontamination schemes. Please refer to the enzyme manufacturer for the recommended conditions for the UNG step of the reaction.
What is the DADE linker and what are its advantages over other carboxyl linkers?
The DADE linker is a pre-activated carboxyl linker for the 5’ terminus of an oligonucleotide. It was designed for the solid phase conjugation of amine bearing compounds directly to an oligonucleotide, although it has other applications. If it is left in its activated form when the oligonucleotide is deprotected, it will react with the prevalent nucleophile in the solution (ammonia in ammonium hydroxide forming the amide, the hydroxyl in sodium hydroxide forming carboxylic acid, etc.). Its major advantage is that it can be used for solid phase conjugation to amine bearing compounds, negating the need for costly succinimidyl esters. This allows high throughput screening, and the reuse of compound that did not conjugate in the first reaction. It also allows for the use of large excesses that will enhance conjugation efficiencies, and reduce cost. It has many other applications.
Can I buy linker reagents directly from TriLink?
Yes, click here to view the reagents still sold through TriLink. All of our other DNA synthesis reagents are now sold through Glen Research.
What is the difference between the TFA C6-Amino Linker and the MMT C6-Amino Linker?
The main difference is the protecting group. The MMT linker is protected with a monomethoxytrityl group and the TFA linker is protected with a trifluoroacetate group. The MMT linker is acid labile, whereas the TFA group is base labile. MMT is the best choice when you want a pure amino labeled oligonucleotide. Since the MMT group is more nonpolar, it acts as a purification handle on reverse phase HPLC just like the DMT group. The TFA protected linker is better for crude conjugations where the oligonucleotide is used after deprotection. As a word of caution, remember to exchange the ammonia for another salt prior to conjugation since excess ammonia will severely reduce the efficiency of many conjugations.
Should I still use additives such as DMSO, Betaine or Glycerol?
CleanAmp™ 7-deaza-dGTP and CleanAmp™ 7-deaza-dGTP Mix function well on their own for most sequences with <80% GC-content. However, should you have a problematic target or one that is >80% GC-content, addition of any one of these additives may increase specificity. We have found 0.5M Betaine or 10% Glycerol give the best results.
Q. How versatile is the CleanAmp™ 7-deaza-dGTP Mix?
Preliminary results show CleanAmp™ 7-deaza-dGTP Mix functions with a variety of enzymes. However, we recommend Invitrogen™ Taq DNA polymerase for best results. For questions regarding your specific enzyme, please contact us.
What components are in the CleanAmp™ 7-deaza-dGTP Mix?
CleanAmp™ 7-deaza-dGTP Mix contains the CleanAmp™ modified nucleoside triphosphates of dA, dC, (dG:7-Deaza-dG) and dT at 10 mM. The dG mix is a 1:3 ratio of CleanAmp™ dGTP:CleanAmp™ 7-deaza-dGTP.
Should I use the CleanAmp™ 7-deaza-dGTP Mix or just CleanAmp™ 7-deaza-dGTP alone?
For targets with <70% GC-content, addition of theCleanAmp™ 7-deaza-dGTP with standard dNTPs should amplify the target well. For targets with >70% GC-content, we recommend theCleanAmp™ 7-deaza-dGTP Mix for a robust amplification.
Do CleanAmp™ dNTPs form a modified amplicon?
No, CleanAmp™ dNTPs are reverted back to the corresponding standard dNTP prior to being incorporated into the amplicon. The 3’-hydroxyl modification blocks enzyme incorporation until it is removed during heat activation. After heat activation, the enzyme efficiently incorporates the corresponding standard dNTP.
Will CleanAmp™ dNTPs activate at 60°C?
We are currently testing the kinetics of the deprotection of CleanAmp™ dNTPs at lower temperatures such as 60°C. Both the temperature and the pH of the PCR reaction affect the deprotection rate of the CleanAmp™ dNTPs where increased temperature and acidic conditions accelerate deprotection.
Our studies of deprotection at 55°C in 1X PCR buffer (pH 8.3) indicate that 40% of the dNTPs are deprotected after 2 hrs. Although deprotection is slower than at the elevated temperatures of PCR (95°C), adjustments to your reaction conditions, such as higher dNTP concentrations will likely allow for success.
Our CleanAmp™ Primers contain a different modification, but provide Hot Start activation using the same principle. At lower temperatures the rate of deprotection is slower, with a lower concentration of activated primers. To counter act the slower rate, we recommend using a higher concentration of primers.
How versatile are CleanAmp™ dNTPs?
CleanAmp™ dNTPs can be used with a broad selection of DNA polymerases including Taq, Pfu, Pfu (exo-),Dynazyme™, Deep VentR™, Tth and Tfi. When using CleanAmp™ dNTPs with these polymerases, all were able to produce the desired amplicon.
Can you provide protocols for CleanAmp™ dNTPs?
Yes, please see the CleanAmp™ dNTPs Product Manual.
How long is the initial denaturation time for CleanAmp™ dNTPs?
When standard cycling protocols are employed, a 2-5 minute initial denaturation step at 95°C allows for robust amplification. Recently we have found that initial denaturation times as short as 30 seconds can be used without any effect on PCR efficiency.
How should I handle CleanAmp™ dNTPs?
For handling stock solution, thaw for 5-15 minutes at room temperature. Do not thaw by heating. The unused portion should be returned to the freezer as quickly as possible. We recommend aliquotting stock solutions into several tubes to prevent extended room temperature exposure over many uses.
How should I store CleanAmp™ dNTPs?
For best performance, we recommend distributing your stock solution into smaller aliquots that are sufficient for one week of work. To avoid prolonged exposure of the CleanAmp™ dNTP Mix stock solution to room temperature, store stock nucleotide solutions in the freezer at -20°C. We recommend not subjecting the CleanAmp™ dNTPs to more than 20 freeze-thaw cycles.
What does the CleanAmp™ dNTP Mix consist of?
It contains the modified nucleoside triphosphates of dA, dC, dG, and dT. We currently offer CleanAmp™ dNTPs in a mix but the CleanAmp™ dNTPs are also available individually upon request. We have found that sometimes the replacement of just one or two of the natural nucleotides with CleanAmp™ dNTPs is enough to have the desired effect.
What are the benefits of CleanAmp™ dNTPs?
CleanAmp™ dNTPs improve PCR specificity, make PCR reactions cleaner, eliminate or reduce off-target amplicon formation, and are compatible with both Hot Start and non-Hot Start DNA polymerases that employ different buffer compositions (pH 7.5-9, 25°C). CleanAmp™ dNTPs are water soluble, stable for at least 1 year when frozen at –20°C and are inexpensive compared to other Hot Start technologies.
How stable are CleanAmp™ dNTPs?
CleanAmp™ dNTPs are most stable in the shipped mix, diluted in PCR buffer (8-9 pH). In these conditions, they are stable for at least one year at –20°C. Exposure to higher temperatures during shipment does not pose performance risks. Avoid repeated freeze/thaw cycles and exposure to room temperature for more that 24 total hours. Upon first use, it is recommended to aliquot samples into single use portions.
Why use CleanAmp™ dNTPs versus unmodified dNTPs?
CleanAmp™ dNTPs are able to decrease the formation of primer dimer products while increasing the specificity of the desired product better than unmodified dNTPs. Use of CleanAmp™ dNTPs also reduces data fluctuation, giving a more consistent PCR result.
What are CleanAmp™ dNTPs and how do they work?
CleanAmp™ dNTPs contain thermolabile modification groups that allow for a dNTP-mediated Hot Start activation approach in PCR. The introduction of temperature sensitive protecting groups onto the 3’-hydroxyl of a dNTP blocks primer extension at the less stringent, lower temperatures of PCR reaction preparation. When the reaction is heated to the elevated temperatures of PCR, the protecting group is removed to form the corresponding standard dNTP, which is now a suitable DNA polymerase substrate.
What is the difference between stock and working solutions?
The stock solution is the DMSO solution in which your primers are shipped. Working solutions are any dilutions that are made using the stock, generally into aqueous solution.
How long is the initial denaturation time for CleanAmp™ Primers?
The initial denaturation time is 2-10 minutes at 94°C for standard thermal cycling protocols. If used in combination with a chemically modified DNA polymerase, such as AmpliTaq Gold DNA polymerase, an initial denaturation of 10-20 minutes may be required. For fast cycling, shorter denaturation times at 94°C can be employed.
How should I handle CleanAmp™ Primers?
For handling stock solution: Thaw for 5-15 minutes at room temperature. Do not thaw by heating. The unused portion should be returned to the freezer as quickly as possible. We recommend aliquotting stock solutions into several tubes to prevent extended room temperature exposure over many uses. For handling working solutions: Dilutions of the stock solution should be made as needed, using water or aqueous buffer (pH 7-9) as the diluents, and then stored on ice. Working solutions should be used immediately.
How should I store CleanAmp™ Primers?
Store at or below -20°C to avoid loss of hot start activity.
Can you provide protocols for CleanAmp™ Primers?
Yes, please see the CleanAmp™ Primer Product Manual.
Which DNA Polymerase should I use with CleanAmp™ Primers?
You do not need to use a DNA polymerase with CleanAmp™ Primers. If you would like to include a DNA polymerase, some validated sources of Taq DNA polymerase are as follows: Taq DNA polymerase (native and recombinant) from Invitrogen; Taq DNA polymerase (recombinant) from New England Biolabs; and EconoTaq™ DNA polymerase from Lucigen. Other validated DNA polymerases with comparable performance to Taq are: Pfu DNA polymerase (exo + and exo -) from Strategene; Dynazyme™ DNA polymerase from Finnzymes; Tth DNA polymerase from USB; Tfi DNA polymerase from Invitrogen; and Deep Vent™ DNA polymerase from New England Biolabs.
Can I use DMSO with CleanAmp™ Primers in my reaction?
DMSO improves PCR efficiency by disruption of GC-rich sequences. Typically, 0-10% DMSO is introduced into the reaction. DMSO can also inhibit PCR performance, so be sure that the percent DMSO introduced by CleanAmp™ Primers is not inhibitory to the reaction. At high DMSO percentages, the performance of CleanAmp™ Primers may be reduced. Therefore we recommend the addition of DMSO into your reaction only if absolutely necessary and to a maximum of 2% volume per volume.
What concentration should I use for CleanAmp™ Primers?
CleanAmp™ Primers can be employed over a much wider concentration range than unmodified primers. It is recommended that you perform a titration of your primers to determine the optimal concentration for improved Ct, with reduced off-target amplicon formation. Generally, primer template systems prone to mis-priming require a much lower primer concentration than those prone to primer dimer formation.
How do I select the CleanAmp™ Primer type that I need?
There are two forms, which differ in the rate of release of the protecting group. CleanAmp™ Turbo Primers are for fast cycling (<45 minutes) and multiplexed PCR. You will see improved amplicon yield and reduced mis-priming and primer dimer formation. CleanAmp™ Precision Primers are for standard cycling (>45 minutes). You will see improved specificity of amplification, improved limit of detection and the greatest reduction in mis-priming and primer dimer formation.
How stable are CleanAmp™ Primers?
When in stock solution, they can be stored at room temperature for 48 hours, at 4°C for 60 days and at -20°C for 1 year. When in working solution, they can be stored at room temperature for 3 hours, at 4°C for 3 days and at -20°C for 20 days.
How much material will I receive?
You will receive the typical synthesis yield of 10-15 ODs.
How will my CleanAmp™ Primers be delivered?
In DMSO at ~200 uM.
How much triphosphate will I receive if I send you 10 mg of a modified nucleoside?
That depends on the modification, but in general, we obtain a 20-40% isolated yield of triphosphate >90% pure (and often >95%) if the nucleoside is clean and behaves normally. That translates very roughly into 10-15 μmoles of nucleotide. The yield can be much lower or higher in any individual case.
What is your maximum scale for custom chemistry products?
The answer depends on many factors, beginning with how easy it is to prepare your particular compound. In general, we are willing to consider quoting on kilogram or multi-kilogram quantities of straightforward phosphoramidites, such as our linkers. We are able to prepare modified nucleotides in 10 grams or more per single synthetic prep if the purification is not problematic. Before we consider a large scale synthesis, we will try it on a much smaller scale first. We can also prepare multiple smaller synthetic batches (this reduces risk to both parties) that can be recombined prior to analysis to prepare single large lots. Please contact us to discuss your particular project.
What is a phosphoramidite?
A phosphoramidite is the actual building block we use to chemically synthesize oligonucleotides. It usually consists of a protected nucleoside with a 3’ phosphitylating reagent, where phosphorus is in the trivalent state and protected with a stable amine and ester. The amine is activated during synthesis with acid, producing a good leaving group. The phosphorus is then attacked by the 5’ hydroxyl of the growing oligonucleotide chain. After coupling, the phosphorus is oxidized to the pentavalent phosphorus state using iodine and water.
Do you provide a “money-back” guarantee if it turns out that you are unable to convert a nucleoside into a triphosphate due to the nature of the nucleoside modification?
Before we agree to do a triphosphate conversion, we need to look at the structure of your specific modified nucleoside. Based on the structure, we can determine whether or not we will be able to convert it to a triphosphate. You will only be charged if we are successful in the conversion.
Which RNA polymerase do you suggest for incorporation of 2’ OMe RNA in transcription?
We recommend the enzyme mixture T7 SP6 from Epicentre Biotechnologies.
Which of your modified nucleoside triphosphates can be incorporated into DNA molecules through PCR?
Aminoallyl-dNTPs, biotin-AA-dNTPs, 2-amino-dATP, 7-deaza-dGTP, and 7-deaza-dATP, 5-Methyl-dCTP, 5-Iodo-dUTP, 5-Bromo-dUTP, 5-Fluoro-dUTP, N4-methyl-dCTP, 5-propynyl-dUTP and 5-propynyl-dCTP, 2-thio-dTTP, 4-thio-dTTP and alpha-thio-dNTPs can be incorporated through PCR. As not all of these analogs will incorporate with similar efficiencies in PCR; some optimization may be needed. We recommend performing some initial PCR experiments using natural:modified dNTPs in ratios such as 1:0, 3:1, 1:1, 1:3 and 0:1 to identify the best conditions for modified nucleotide incorporation with robust amplicon yield.
Which enzymes work with which modified nucleotides?
There is very little known about many of the triphosphates we sell. Please see our table of Enzymatic Activity of Selective NTPs, or our NTP bibliography in our website and catalog to see if there is a publication regarding your question. If not, our stock answer is often tongue in cheek: buy the compound, be the first to do the research, publish the paper and then we will both know. Seriously, we are attempting through both in-house and collaborative efforts to learn more about potential applications of our compounds, but this is a long term effort. Please contact us to see if there is any news about your particular application.
What type of mass spectroscopy do you use to verify the mass of the triphosphate?
Electrospray Ionization (ESI).
How do you make sure that no salts are left in your preparations after purification and precipitation?
We QC our products by 31P NMR which reveals inorganic phosphate salt. This is usually the salt of most concern for enzymologists since it is an inhibitor of many polymerases. For other salts, the lambda max and epsilon values of the modified nucleoside are used to make sure the optical density matches the actual dry weight. The concentration of NTP in the final solution is determined spectrally. This spectra is not affected by the salt form of the triphosphate, assuring we deliver accurate quantities of our NTPs, based on the molecular weight of the free acid form.
What is the expiration date of my modified nucleotide?
All of our modified nucleotides are submitted to quality control frequently to ensure that they still meet our specifications, therefore we do not assign expiration dates to our modified nucleotides other than our CleanAmpTM dNTPs.
What kind of nucleotides can you prepare under GMP conditions?
We can prepare any kind of nucleotide in our GMP facility. We first develop, if necessary, the process in our R&D lab, then validate it in our GMP facility with our equipment and reagents. All TriLink products are GMP and manufactured in compliance with ISO 9001.
What are the likely contaminants when preparing a triphosphate?
The most common contaminants are the mono- and diphosphate forms of the nucleotide. Our purification procedures are designed to minimize the amount of other phosphates present, but since they result from hydrolysis that occurs at some finite rate with all nucleotides, they are generally present in the 0.5-3% range as detected by anion exchange HPLC. We routinely analyze older lots to identify these species and repurify the NTP if necessary to ensure it meets our purity specification (>90% for catalog products) before shipping. Two other potential contaminants are salts and inorganic phosphate. Salts are removed during the reverse phase HPLC and precipitation steps. Inorganic phosphate can be particularly problematic in some applications, therefore we take special measures to remove as much as possible. Every lot of NTP we manufacture is analyzed for the presence of inorganic phosphate using phosphorous NMR.
How do you prepare your nucleotides?
All of our nucleotides are prepared chemically using a variety of methods, several of which are proprietary to TriLink. We use a multi-step purification process that includes two DEAE columns and a reverse phase HPLC. Finally, the compounds are precipitated to remove any remaining salts and analyzed for quality. They are stored as an aqueous solution at -70°C.
What triphosphates do you sell that have nucleosides with fluorescent properties?
Triphosphates of fluorescent nucleosides can be found in the Isomorphic-Intrinsically Fluorescent Analogs category. Their quantum yields and uses have been described in the literature. If you don’t see the triphosphate you need, request a quote.
Do you sell dye-labeled NTPs?
Yes. Currently, we sell biotin and dabcyl labeled NTPs. Other labeled NTPs will be added to our product offering in the future.
My NTP arrived thawed. Is it ok?
Although NTPs are stable at room temperature for over a week, we ship them on ice to ensure quality in the event of a delay in transportation. We are still confident in the integrity of our NTPs if they have arrived thawed. However, if your ice has disappeared when your NTP arrives and you have concerns, please contact us immediately.
How should I store my NTP and how long will it last?
Most NTPs should be stable for several years when stored properly at -20oC. We recommend aliquotting your NTP into multiple, single-use tubes upon first thawing to avoid repeated freeze/thaw cycles, which can lead to significant physical degradation. This is especially important for NTPs with amino modifications, which are less stable in solution than other NTPs.
Your NTPs are delivered in solution. What is the pH?
Our triphosphates are resuspended in water so the pH of the solution will be very close to neutral.
What are the NTPs dissolved in?
Water.
How many μL are in a 1 μmole vial of an NTP at 10 mM concentration? At 100 mM?
The 1 μmole size of an NTP at a 10 mM concentration will have 100 μL of solution and a 100 mM concentration will have 10 μL (0.01 mL) of your NTP solution.
What type of QC do you run on your triphosphates?
After purifying an NTP by anion exchange and reverse phase HPLC, the compound is precipitated and dissolved in water, an analytical scale anion exchange HPLC is run to verify each compound is >90% triphosphate. In addition, 31P NMR, Mass Spec and UV analyses are done to ensure the triphosphate has little or no inorganic phosphate. We then match the calculated molecular weight and confirm the concentration of the final solution.
What analytical data do you provide for NTPs?
Each NTP is accompanied by a certificate of analysis that lists the methods of analysis, concentration, purity, MW and extinction coefficient. A copy of the HPLC is also included.
What’s the difference between a nucleoside and a nucleotide?
A nucleoside is only the base and the sugar (5’-OH) while a nucleotide refers to a nucleoside with a phosphorylated sugar of any kind. This includes the phosphate groups on the 2’, 3’ or 5’ ends.
What is FRET?
FRET stands for fluorescence resonance electron transfer. This phenomenon is observed when two dyes are placed within proper distance of each other (one full turn of DNA is an excellent length). If the first dye is excited and emits photons, and the second dye absorbs that wavelength, the only observed emitted wavelength will be that of the second dye. This ability to see another wavelength when two dyes come into proximity of each other has a number of useful applications for diagnostic assays.
How many dyes of the same type can you place on an oligonucleotide?
At most, we recommend one conjugate every 5-10 bases. Theoretically, we can place a conjugate at every location and at the 3’ and 5’ ends. Unfortunately, it is very difficult to synthesize these types of compounds and just as difficult to analyze for labeling efficiency. In many cases, the resulting products are essentially insoluble. In other cases, such as fluorescein, the dye actually quenches itself if too many are present, becoming almost non-fluorescent. The length is also limited.
How many different dyes can you place on a single oligonucleotide?
It is possible to place up to six different dyes on one oligonucleotide, but that can only be done with a very specific sequence of dyes and is of no practical use. We routinely place two dyes on an oligonucleotide, and can place up to four with some degree of flexibility in dye choice using a combination of synthesis reagents and an orthogonal linker scheme. However, any combination above two gets more problematic to synthesize and purify, which will affect purity and yield. Please contact us to discuss your specific needs.
Can you make an oligonucleotide with two different dyes or other conjugates?
The answer is complicated because there are many ways to place different conjugations on an oligonucleotide, but we are limited by what is available for conjugation and by compatibility issues. The easiest way is if one or both of the conjugates are available as a phosphoramidite and/or support bound reagents. If at least one reagent is available, the other conjugate can often be attached through a primary amine at the other end of the oligonucleotide. Chemical compatibility is avoided in this case. If neither conjugate is available as a phosphoramidite or support bound reagent, then we often will use an orthogonal linker scheme incorporating both a thiol linker and an amino linker. This requires that one of the conjugates is available as a maleimide, which is thiol specific under neutral pH conditions. Most of all the commonly used dyes are available in both the succinimidyl and maleimidyl ester forms. The most common, such as fluorescein, TET, HEX, TAMRA, Cyanine 5 and Cyanine 3 are also available as phosphoramidites or support bound reagents. We will help you design your specific oligonucleotide when you place your order.
What are the absorption max and emission max values for my fluorescent labeled oligonucleotide?
Spectra data is listed for each dye on our website and in our catalog. These values are provided by the manufacturer of the fluorescent dye and are generally calculated from the free dye not attached to an oligonucleotide. As a general rule, these values work fine for the common oligonucleotide user as there is little change, if any, when the dye is attached to an oligonucleotide. The particular base composition of an oligonucleotide can play a role as can pH in many cases, particularly with fluorescein and related dyes.
Can I use a dye or modification that is not listed here?
Yes. Tell us the dye or modification you are looking to use; we will find out if it is commercially available. If it is not, there is a possibility we can make it in-house. If neither of these is an option we will do our best to see if there is another dye or modification that can give you similar results. If any of these options work for you, we will then send you a quotation.
Why do you no longer offer pteridine oligonucleotides?
The original supply of reagent was finally exhausted, and the cost of replacement far outweighed the potential sales. We had to make the unfortunate decision to relinquish our license.
How does a quencher work?
A quencher is very efficient at absorbing certain wavelengths. When near a dye that emits at the absorbed wavelength, the light is “quenched”, and no longer visible. Quenchers are very similar in structure to dyes. The difference is they emit in undetectable ranges, or in undetectable amounts. The ability to quench is a function of distance from the dye in most cases. Molecular beacons are effective in that the quencher actually comes in contact with the dye. Different quenchers are best for different dyes.
What is the best design for a molecular beacon?
This is a difficult question since so much depends on your specific target sequence and application. A good set of rules for the design of molecular beacons can be found on www.molecular-beacons.org. In general, the most successful compounds are 30-40 nucleotides in length, with the stem comprising the last 6 bases on either terminus. Another general rule is that the melting temperature of the stem should be about 10°C lower than that of the target/probe section of the molecular beacon. This is where the trial and error comes in as the design tools are only predictive of nature.
What is the difference between a 6-FAM and FITC labeled oligonucleotide?
Both 6-FAM (6-carboxyfluorescein, single isomer) and FITC (fluorescein-5,6-isothiocyanate, mixed isomers) are forms of the fluorescent dye fluorescein. FITC is a particular form of reactive species, the isothiocyanate, of the dye. It yields a urea linkage upon reaction with a primary amine. 6-FAM is the preferred reagent for labeling an oligonucleotide. It results in an amide bond when reacted with a primary amine. The chemistry is more robust and better yielding. Furthermore, it has been shown that 6-FAM is less susceptible to photobleaching. 6-FAM and FITC conjugated oligonucleotides have similar spectral properties otherwise.
Can I have a dye conjugated to my oligonucleotide internally?
Yes. We use the succinimidyl ester version of the dye and conjugate it to a selective placement linker. We then post-synthetically conjugate the dye to the oligonucleotide. Other options include using a dT-FAM or dT-TAMRA. These dyes are already conjugated to the dT-C6 amino linker.
How do I deprotect my 5’ maleimide oligonucleotide?
The protected maleimide modification is deprotected using a retro Diels-Alder reaction. This can be done with microwave irradiation or by heating in toluene. The second approach is described below. Retro Diels-Alder reactions should be conducted in anhydrous conditions as exposure to significant levels of moisture can cause incomplete deprotection, hydrolysis and/or addition of water to the maleimide.
1. Suspend lyophilized oligo in anhydrous toluene.
2. Let sit for 4 hrs at 90°C.
3. Cool to room temperature.
4. Evaporate toluene.
The oligonucleotide is now ready for conjugation to a thiol compound.
What is the machine validation charge for trimer modified oligonucleotides?
When trimer modified oligonucleotides require machine synthesis, a machine validation is performed prior to starting the job. This validation ensures that all aspects of the synthesizer are functioning properly and decreases the chance of synthesis failure. Since the trimer mix amidite is an expensive modification we take all necessary precautions to limit the chance of a synthesizer malfunction which could waste trimer mix amidite. A machine validation charge is required for each new synthesis run so it is beneficial to order all of your trimer modified oligonucleotides at once. This allows us to split the validation charge across all oligonucleotides in the order, reducing cost per oligonucleotide.
Why does the price of trimer modified oligonucleotides vary?
Trimer modification requires a highly custom synthesis protocol in which multiple additional factors are taken into account. Each trimer mix job is individually analyzed in order to provide the most cost effective pricing in terms of synthesis cost and trimer amidite usage. If a small number of trimer mix incorporations are needed then it is more cost effective to manually couple the trimer mix rather than machine couple. While manual coupling adds increased labor to the quote, the amidite usage is decreased since there is no need to prime the machine. Also, manual coupling eliminates the need for a machine validation requiring an additional charge.
Does TriLink provide LNA modified oligonucleotides?
Yes, we are able to synthesize LNA modified oligonucleotides, however there are a few stipulations. Since this material is under license by Exiqon we require that our customers obtain a license from Exiqon, purchase the LNA amidites and have the material shipped to our facility for synthesis.
Can you provide isomerically pure phosphorothioate modified oligonucleotides?
It is very challenging to distinguish between the two species. We, therefore do not offer purification of phosphorothioate oligonucleotides at this time.
The analysis of my purified thiol modified oligonucleotide shows three different species. Are these impurities?
No, this is generally not an indication of impurity. A thiol modified oligonucleotide which is not fully reduced will be in three states: free thiol (oligonucleotide-SH), protected thiol (oligonucleotide-S-S-C6-O-DMT) and dimerized oligonucleotide (oligonucleotide-S-S-oligonucleotide). We recommend that you reduce these oligonucleotides just prior to use to ensure that only the free thiol is present. DTT or TCEP are the preferred reductants.
Is it best to have my 5’ thiol linker modified oligonucleotide delivered with or without the thiol protected?
There is simple answer to this question, as it depends on your experience and what you plan to do with the molecule. In most applications, the thiol will be reduced just prior to use to ensure that unwanted dimers are cleaved. If the conjugation is to a maleimide, the reduction can actually be done in situ during conjugation. In that case, there is very little difference. Technically, it is easier to purify a protected thiol, but it is very difficult to keep it on completely intact.
What other linkers can you offer beyond those listed already?
We can prepare almost any linker not covered by a patent. Please contact us for more information.
What amino linker lengths do you have available?
Click here to view the full list of linkers currently available. For more information, please see our article DNA Linker and Spacer Reagents and Their Utility.
How do I reduce my thiol modified oligo?
1. Make a solution of 60 mM TCEP. (18 mgs TCEP in 1 mL H2O)
2. Add 125 uL 0.1M NaHPO4, 0.15M NaCl, pH 7.26, vortex & make certain oligo is in solution.
3. Add 75 uL 60 mM TCEP to oligo solution.
4. Vortex.
5. Let sit for 2 hrs at room temperature.
6. Run size exclusion column, RP-HPLC.
Do you still make 5-propynyl oligonucleotides?
Yes. Formerly, we were directly licensed from Isis and all potential customers had to first fill out a questionnaire regarding use for Isis to approve. Now we can buy the reagent from the licensed supplier (Glen Research) and make oligonucleotides for research use.
What are the advantages of using 2’OMe RNA bases in an oligonucleotide?
2′ O-Methyl RNA is a more stable form of RNA with similar properties. It has the same low rate of depurination, but it is protected against base hydrolysis. 2′ OMe RNA is chemically more stable than RNA as well, and allows for greater flexibility in conjugation schemes.
2′ O-Methyl RNA:RNA duplexes are the most stable. RNA:RNA and DNA duplexes have lower melting temperatures. 2′ OMe RNA is most commonly used in therapeutic applications due to its nuclease resistance.
What’s the difference between a phosphodiester and a phosphorothioate backbone?
A phosphorothioate linkage has a sulfur in a non-bridging location on the phosphate backbone. This modification is known to greatly retard nuclease degradation of oligonucleotides. Phosphorothioates also have a lower melting temperature, which is a measure of the association and disassociation rates. A phosphodiester linkage contains all oxygens on the phosphate backbone.
How do you purify your oligonucleotides?
We offer several different purification schemes. They offer a range of capabilities and applications. Our most common method is reverse phase (RP) HPLC. We use C-18 columns for most of our work. The vast majority of our compounds are at least double purified, first undergoing a trityl-on prep, followed by another full HPLC purification after removal of the trityl group. This ensures a high level of both length and chemical purity, which is comparable or even superior in many cases to more sophisticated and costly anion exchange (AX) HPLC methods. We have demonstrated this to many customers and would be happy to demonstrate the same to you with a sample synthesis. We do offer AX-HPLC, but in a more limited capacity. Another popular method is polyacrylamide gel electrophoresis (PAGE). This method is limited to just a few tens of milligrams of material (15 μmole starting scale) and becomes expensive very quickly. It is a good method to enhance length purity, but is limited in its ability to remove chemical contaminants.
Is there excess salt in my oligonucleotide?
It is very difficult to remove all excess salt. However, we do routinely obtain less than 5% excess salt, and our diafiltration process for larger samples can reduce the salt content to even lower amounts.
What is the “extra purification step” that is needed for many of the fluorescent dye oligonucleotide conjugates?
Dyes listed with the “extra purification step” requirement are made in a two-step process. First, the unlabeled oligonucleotide with the appropriate amino or thiol linker is synthesized and then purified to isolate only the full-length oligonucleotide. The oligonucleotide is then conjugated to the dye and purified again to remove any unlabeled material. Some fluorescent dyes, such as 6-FAM, TET and HEX, do not require this extra purification step. These dyes are available as phosphoramidite reagents, allowing us to label the oligonucleotide during the synthesis procedure and use a single purification step.
Do I need to have my oligonucleotide purified? If so, what method is best?
The need for oligo purification is dependent on a number of factors, including your application and oligo complexity. HPLC and PAGE are the most common purification methods. Reverse phase HPLC (RP-HPLC) is by far the most common, and is very useful in separating dye-labeled from unlabeled oligos, and in separating full-length oligos from truncated species (n-1, n-2, etc.). A crude/desalted preparation of a primer is usually sufficient for a PCR application. Long oligos (45+ bp) do not resolve well by HPLC, therefore PAGE purification is recommended. RNA requires purification under sterile conditions and sterility is most easily controlled during a PAGE purification procedure. Our technical support team can assist you in determining the best purification method for your construct and requirements.
Do you offer small-scale oligonucleotides under GMP conditions?
Yes. We offer full GMP quality material at each scale we offer. However, we do not have a small scale synthesizer in the clean rooms, which are reserved for programs requiring each oligonucleotide to be synthesized one at a time in an exacting environment. Most of our small-scale oligonucleotide programs include multiple oligos being prepared simultaneously, albeit under very controlled circumstances with validated protocols and instruments. The cost of doing a small-scale synthesis one compound at a time in a clean room is very expensive and normally unnecessary except in the most unusual cases.
I received less/more of this oligonucleotide the last time I ordered it. Why the difference?
The synthesis and purification of an oligonucleotide is a very complicated process with hundreds of individual steps. The overall success of a synthesis can depend on changes in minor amounts of contaminates from synthesis to synthesis, or even on uncontrollable variables such as the humidity level on the day of your particular synthesis, despite our environmental control systems. All of these (and other factors) can affect the amount that can be isolated of a particular compound with the proper purity.
I ordered a 1.0 µmole scale synthesis. Why did I not receive 1.0 µmole of final product?
Research scale oligonucleotides are sold (by convention) by the starting scale, not the delivered amount. The reason is that each oligonucleotide is different, and will result in different yields, although the cost of material and labor is equivalent for similar constructs with the same starting scale. Therefore, the price is based on actual cost. If you need a defined amount of an oligonucleotide, please note clearly on your order that you are ordering based on the final quantity. We will then determine the scale needed to deliver that amount and price your oligo accordingly.
How much material will I get from a 1.0 μmole scale, HPLC purified?
Typically our yields for a 1.0 µmole starting synthesis scale, purified are about 30 -60 OD260 units, or ~1-2 mg. Yields vary greatly depending on modifications and even on the sequence itself.
What is the longest oligonucleotide TriLink can synthesize?
TriLink has successfully synthesized 220mer unmodified DNA and 110mer unmodified RNA. However, TriLink is willing to try longer sequences. Please contact us discuss your needs.
How do you make an oligonucleotide?
We manufacture oligonucleotides on synthesizers (an automated reagent delivery machine) and/or manually, depending on the type and complexity of the oligo. Synthesis occurs 3’ to 5’, with the 3’ base anchored to a solid support within the synthesis column. The synthesis cycle involves five chemical reactions and repeats as it adds each base to the growing oligonucleotide chain. For further detail, refer to our article A Short History of Oligonucleotide Synthesis.
Why is the OD reading for my duplex lower than what is stated on my Certificate of Analysis?
The extinction coefficients of the single strands are added together to provide the final data on our Certificates of Analysis. When oligonucleotides are annealed; however, these constants are not entirely additive which can cause confusion when calculating the amount of product present in a sample. Duplex extinction coefficients can be 15-25% lower than expected due to the hyperchromicity of the duplex. Therefore, we provide a calculation on our duplex Certificates of Analysis to help better determine the actual amount of duplex present in a sample:
(Measured ODs/Total ODs of single strands) x Total Extinction Coefficient = Duplex Extinction Coefficent
How do I calculate the number of μmoles of oligonucleotides I have?
On the Certificate of Analysis you received with your product, you are supplied with two numbers needed for this calculation: the number of OD260 units of product (the absorbance reading you would obtain from a 1 cm cell path if you dissolved your sample in 1 ml of water, and usually serially diluted for the reading) and its calculated extinction coefficient (ε). The ε is given in units that are readily used for our application, OD260 units per μmole. Divide the number of OD260 units by ε to calculate the number of μmoles directly. For example, if you obtain 60 OD260 units of an oligonucleotide with an ε of 300 OD260units/μmole, you have received 0.2 μmoles of product.
What are OD260 units?
The optical density unit, or more commonly the OD260 unit, is a spectrophotometric measurement of an oligonucleotide. It is a normalized unit of measurement that is defined as the amount of oligonucleotide required to give an absorbance reading of 1.0 at 260 nm in 1.0 mL of solution using a 1 cm light path. Each of the bases in a nucleic acid strand has an absorbance at or near 260 nanometers, due to their conjugated double bond systems. Because the exact base sequence and composition is known, the OD260 unit is a precise method to quantify an oligonucleotide. Utilizing absorbance measurements is the recommended method for quantitating or aliquotting an oligonucleotide.
How do I determine the mass of my oligonucleotide?
The molecular weight of your oligonucleotide is provided on the Certificate of Analysis. Multiply that number by the number of μmoles. For example, if the molecular weight of your oligonucleotide is 6950 au, and you have 0.2 μmoles of product, you have 1390 μg or 1.39 mg.
How do I prepare a solution of my oligonucleotide with a predetermined molarity?
First determine the number of μmoles you have. Next, convert the units of your desired stock solution to M. For example, if you wish to prepare a 10 mM stock solution first convert your units to M by dividing by 1000 (resulting in 0.01 M in this example). Now simply divide the number of μmoles you have by the desired concentration of your final solution in M to determine the μL of buffer you will need to achieve that concentration. For example, if you have 0.2 μmoles of product and you wish to prepare a 10 mM stock solution, divide 10 mM first by 1000 to convert to 0.01 M, and then divide 0.2 μmoles by 0.01 M to obtain the number of μL needed, 20 μL in this case. This is a simplified, working version of the string of mathematical equations that underlie the procedure described. If you require more information, please contact us.
What is the extinction coefficient?
The extinction coefficient, ε, is a physical constant that is a key component of Beer’s Law regarding the relationship between optical absorption and concentration; A = ε [conc]. By knowing the ε of an oligonucleotide one can readily convert the optical absorbance reading into concentration, and then into mass. The ε is based on the actual bending and vibration of the bonds and takes into account the absorbance of the individual bases and the effects of neighboring bases. The ε is derived from the exact nucleotide composition of the oligonucleotide so it is unique to every oligonucleotide sequence. The units for the ε are expressed normally as OD260 units /μmole. TriLink calculates the ε of an oligonucleotide using the nearest neighbor model which is considered to be the most accurate method without actually determining the ε empirically, which requires a great deal of experimental effort.
What is the purpose of measuring the oligonucleotide in OD260 units?
It is a very accurate and convenient method of quantifying the oligonucleotide because each of the bases has an absorbance at or near 260 nanometers, so it is the average maximum wavelength of long DNA strands. This method has traditionally been used because oligonucleotides are too large for standard small molecule techniques, and they do not contain enough material to use classic methods to determine mass, such as using a balance.
How long is my oligonucleotide stable once received?
Since just about every oligonucleotide we synthesize here at TriLink is unique, we are unable to provide stability information or run stability studies, as this data could differ with each individual sequence. The most important precaution to take is to properly store your oligonucleotide. As long as your oligonucleotide is stored at -20°C or lower for long term storage and multiple freeze thaw cycles are avoided, we have seen that dye labeled oligonucleotides can be stable up to 6 months and unmodified oligonucleotides stable up to 1+ years.
Why is having too many G’s in my sequence problematic?
Poly guanosine and deoxyguanosine stretches (anything over four in a row) have the propensity to form tetraplexes (complexes consisting of four different strands all bound at the poly G region. These complexes are very tight and make oligonucleotide synthesis difficult. We highly recommend against these sequences. We have been successful in some cases, but we are hesitant to guarantee results.