Are 6-FAM-labeled oligonucleotides freely taken by mammalian cells in culture in the absence of transfection agent ? (e.g. a liposome). In other words is 6-FAM permeable to the cells?-Carlos

Dear Carlos,
6-FAM isn’t hydrophobic enough to difuse an oligo across the cell membrane. Some modifications that help with cell uptake are multiple incorporation’s of spermine, or hydrophobic labels like stearyl and cholesterol. Please contact us if you have any additional questions.

Kindest regards,


What’s your guaranteed minimum yield (OD) for oligos ~60bp in length after HPLC purification on a 200nmol and a 1umol scale? -Ben

Dear Ben,

At the smaller scales, 15 umole and below, there is no guarantee of final yield. Each oligo has unique synthetic properties based on its sequence and modifications; therefore it is not easy to predict how much material will result from a particular synthesis.

Our general expectations for HPLC purified unmodified DNA oligonucleotides are as follows:

0.2 µmole scale 5 – 15 OD260 units (˜0.15 – 0.5 mg)
1.0 µmole scale 20 – 60 OD260 units (˜0.66 – 2 mg)

If you require a specific yield, please let us know when you place your order or request a quote.



Hi, can Trilink synthesize a RNA di-nucleotide containing at least one intrisically fluorescent ribo-nucleotide analogue? Thank you-Egor

Dear Egor,

We are able to synthesize an RNA dinucleotide containing an intrinsically fluorescent ribo-nucleotide analog.

I have included a link to the product page for a few options we can offer:

Isomorphic-Intrinsically Fluorescent

For a formal quotation, please contact us through

Best Regards,

Tiffany Teng

Product Specialist I

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.

How does a quencher work?

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 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.

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.

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.

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.

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.

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 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 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.

What is Tm?

Tm is shorthand for melting temperature. The melting temperature of a nucleic acid duplex is the temperature at which half of the strands are in duplex form, and the other half are single stranded. This is determined by taking advantage of a spectral phenomenon called hypochromicity, which is the reduction in UV absorbance that is observed when nucleic acid bases form a duplex. The act of hydrogen pairing with the opposing strand and the change in electron patterns results in the observed reduction in the spectral reading at 260 nm. By cooling a sample containing complimentary strands in a cuvette until the strands are fully hybridized (absorbance is stable), then heating the sample slowly and recording the absorbance during the process, one obtains a sigmoidal shaped plot when temperature is plotted against absorbance. The transition temperature at the inflection point is the Tm.

How do I hybridize my oligonucleotide?

Resuspend the two complimentary strands in a neutral pH buffer. Measure the absorption of the solution using a spectrophotometer and calculate the concentration in moles using the provided extinction coefficient. Mix the two oligonucleotides together in equal molar ratios. Hybridization can be confirmed by running 3 samples on a gel under native conditions. Sample 1: single stranded oligonucleotide 1; Sample 2: single stranded oligonucleotide 2; Sample 3: hybridized oligonucleotide 1 and oligonucleotide 2. If hybridization was successful, the hybridized oligonucleotide Sample 3 lane will run slower compared to the single stranded Samples 1 and 2. If a little of one of the strands is observed in the duplex lane, titrate with the other strand until only duplex is observed. In some cases, it will be necessary to chill the gel to ensure duplex stability.

What oligonucleotide QC analyses do you provide?

We confirm quality of each oligonucleotide, regardless of level of purification, by PAGE analysis and our in house mass spectral analysis. Please contact us to review data from a previous order. For complex oligonucleotides that require mass spectral analysis not available at TriLink, please request mass spec at time of quote or order. An additional fee may be required. For mid-scale (50 mg or greater) syntheses, we also offer HPLC analysis at no additional cost.

We offer a variety of analyses including HPLC (AX, RP or Ion Pairing), Gel Densitometry, Conductivity and Enzymatic Digest. Please request any additional analysis at time of quote or order as an additional fee may be required.

We will often obtain data during the process that we do not routinely provide the customer, such as the uv/vis absorption spectrum of dye labeled oligonucleotides. If you would like to see a particular piece of data, please contact us, and we will do our best to accommodate you. If you would like to see this data routinely, we will add it to the specifications for your program and include any additional fees as required.

What is the turnaround time for receiving my oligonucleotide once I place my order?

The amount of time depends on the type of oligonucleotide you order. Small-scale, unmodified oligonucleotides can take as little as three days. If purification is requested, the oligonucleotides will take slightly longer to be shipped. A midscale order (50 mg – 10 g), will take approximately four weeks. Highly modified or conjugated oligonucleotides can also take longer. We always ship your oligonucleotides out as quickly as possible, but we refuse to sacrifice quality. Upon placing the order you will receive a confirmation email that typically includes an estimated shipping date. We will update you as soon as possible if there are any changes to the estimated shipping date.

What is the difference between ordering an oligonucleotide at a scale amount versus in milligrams?

Ordering by a scale amount means you are requesting a starting scale synthesis. The final yield will not equal the starting amount, since there are synthetic, purification and processing losses that differ from compound to compound and often from batch to batch of the same material. Because of this, the price is calculated based on the pre-determined cost of material and labor for a synthesis at that particular scale. Our small-scale orders include all syntheses up to the 15 µmole scale. When you place an order in milligrams or grams you are requesting an amount of final product. We have set prices starting at 50 mgs for large-scale orders. If the oligonucleotide has any modifications, then a custom quotation will be necessary. You may request a specific final yield for the smaller scale syntheses, such as 1 μmole, however please keep in mind that it may require a 5 µmole starting scale or more to guarantee that yield. When you specify the final yield, we must begin with a large enough scale to ensure the yield is sufficient. Prior experience with your particular compound will help us determine a fair price.

Is it be possible to synthesize an RNA oligo of 60-90 nucleotides long?

Is it be possible to synthesize an RNA oligo of 60-90 nucleotides long? In addition, is it be possible to label some inner selected nucleotides as we desire? –  Francisco Hernandez-Torres, University of Jaen

Yes, we can synthesize a 60-90mer RNA oligonucleotide. We have successfully made up to a 110mer unmodified RNA oligonucleotide and continue to try longer sequences as our customer’s request them. Yes, internal modifications can be incorporated. Read More…

What solution do you have for a DNA oligo with a fluorescent dye attached for a fluorescence polarization assay?

I need a DNA oligo with a fluorescent dye attached for an fluorescence polarization assay. The linker between DNA and dye should be as rigid as possible and the dye most likely a BODIPYdye, dansyl or fluorescein dye. Attachment either internal or at the 5′ of the oligo. What solution do you have for that? – Chris Gloeckner, Illumina

A shorter linker will provide the most rigidity between an oligo and a dye. Our shortest linker is the dT-C2 amino linker. The linker arm is attached to the Thymidine base and the dye would be conjugated onto the amino group post-synthetically. Another option would be to use our C7 internal amino linker. Read More…

Have a question about Oligonucleotides? Post a comment below.

How do I reduce my thiol modified oligo?

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.

Have a question? Post a comment below or email us at

I would like to know what condition (and aqueous solution) you recommend for suspension of the lyophilized oligos for in vitro use?

I would like to know what condition (and aqueous solution) you recommend for suspension of the lyophilized oligos for in vitro use? – Michael Dickinson

Prior to resuspending your oligo be sure to centrifuge it briefly to ensure all product is down in the bottom of the tube. We recommend bringing your oligo up in any aqueous solution with a neutral pH. If you use just water, be sure that it is nuclease free. Read More…

Have a question about oligonucleotides? Post a comment below.

An Antigene-based Oligonucleotide Therapeutic

Though the phenomenon of siRNA is now a key tool in drug development, it does have a fundamental disadvantage. Since siRNAs silence expression at the mRNA stage, it is presumed that siRNA drugs would need to be administered indefinitely. In a recent review article by N. Kolevzon and E. Yavin, the concept of a photoactivated TFO is proposed as a viable therapeutic option… Read More.

Have a question about therapeutic oligonucleotides? Post a comment below.

What is the Site-Directed Mutagenesis?

Site-directed or oligonucleotide-directed mutagenesis is a key method in the study of protein structure/function and gene expression control. If the nucleotide sequence of the gene is known, an oligonucleotide can be manufactured to introduce a single base change in the codon corresponding to the specific amino acid to be altered. The oligonucleotide is then used to generate a set of clones that can be identified and propagated. Learn More.

What is the use of a Trimer Oligo Solution?

While randomers are often used in protein mutagenesis procedures, they have several short comings. First, it is very difficult to achieve a truly random mixture. The second and more problematic issue is the inevitable introduction of stop codons. Fortunately, the ability to direct combinatorial mutagenesis using randomized oligonucleotides has been advanced by the use of trimer oligonucleotides. Read More.