Oligonucleotide Protocols and Best Practices for Resuspending Primers

Review selected protocols that are commonly used to spectrophotometrically quantify the concentration of an oligonucleotide or primer solution: how to concentrate or precipitate an oligo, how to PAGE purify oligos, and how to build adaptors with your oligos.

To resuspend primers, follow these steps for optimal results:

1. Prior to resuspension, spin down the tube in a microcentrifuge to collect all material at the bottom.
2. Resuspend the primer in sterile TE buffer, consisting of 10 mM Tris-HCl (pH 8.0) and 1 mM EDTA. It is recommended to use TE instead of deionized water. This is due to the slightly acidic pH of deionized water, which can potentially lead to hydrolysis of the primer.
3. After adding the water or buffer, gently mix the contents by pipetting up and down or by vortexing at a low speed. Avoid vigorous mixing to prevent the formation of air bubbles.
4. Finally, centrifuge the tube briefly to collect any residual liquid that may be present on the walls of the tube, and then store the resuspended primers at –20°C until further use.

Use our handy oligo resuspension calculator to determine the buffer volume needed for your desired stock concentration.

1. First, ensure that the primers are stored at –20°C to maintain their stability. Prior to resuspension, allow the primer stock solution to thaw on ice.
2. Next, gently vortex the primer tube to mix any settled material.
3. Calculate the volume of stock solution (V1) to dilute using the following equation: V1=(M2*V2)/M1, where M2 is the final molar concentration of your diluted solution, V2 is the total volume of your diluted solution, and M1 is your stock molar concentration.
4. Add the calculated volume of stock solution (V1) to an empty tube.
5. Add enough water or appropriate buffer to bring up to the total final volume (i.e., add V2–V1). Pipette up and down to mix.
6. Finally, centrifuge the tube briefly to collect any residual liquid that may be present on the walls of the tube, and then store the diluted and stock primers at –20°C until further use to help maintain stability.

To see examples of calculations related to oligo resuspension and dilution, please see our FAQs.

Oligonucleotide quantitation protocol using a spectrophotometer

1. Add an aliquot of the resuspended oligonucleotide to a final volume of 1,000 µL with water. Calculate the volume of water needed by subtracting the volume of oligonucleotide added from 1,000 µL.
2. Vortex or pipette up and down for 15 seconds to ensure thorough mixing.
3. Read the absorbance of this dilution at 260 nm (A260).
4. Obtain the Weight per OD value from the Certificate of Analysis.
5. Calculate the dilution factor by dividing 1,000 by the volume of resuspended oligo added for the dilution in step 1.
6. Use the formula below to calculate the concentration of the oligonucleotide in the stock solution:

Concentration in µg/mL = A260 × Weight per OD × dilution factor

Protocol for ethanol precipitation of oligonucleotides

For most applications, ethanol precipitation is not required prior to use. However, if ethanol precipitation is desired, the following protocol may be used for oligonucleotides longer than 20 bases and with a quantity exceeding 0.1 OD. In cases where there is a minimal amount of material, a carrier such as tRNA should be used.

1. Dry the oligonucleotide in a microcentrifuge tube.
2. Redissolve the pellet in 0.2 mL of 0.3 M sodium acetate (pH 7.0).
3. Add 0.6 mL cold absolute ethanol (–20°C) and place at 4°C for 30 minutes.
4. Centrifuge at 10,000 × g for 30 minutes.
5. Carefully remove the supernatant without disturbing the pellet.
6. Add 1 mL of cold 70% (v/v) ethanol.
7. Centrifuge at 10,000 × g for 10 minutes.
8. Carefully remove the supernatant (without disturbing the pellet) and evaporate to dryness.
9. Store the oligonucleotide at –20°C.

Protocol for PAGE purification of oligos

The following protocol, excerpted from the Invitrogen GeneTrapper Manual, outlines the steps for PAGE purification of oligos. Please note the resolution is poor on precast mini polyacrylamide gels for smaller (<25 nt) oligos.

1. Use a 1.5 mm gel with well forming combs (not sharkstooth combs).
2. Prepare a 12% acrylamide gel (19:1) with 8 M urea and 1X TBE.
3. Dissolve a 5–10 OD of oligo (approximately 1 OD = 33 µg) in 25 µL TE. Add equal volume of formamide to the oligo and heat to 95°C for 1 min. Chill on ice. To avoid masking the location of the oligo (when detected by UV shadowing) do not add dye to the formamide. We suggest an outside lane not containing oligo have the dye added for tracking purposes.
4. Flush wells before loading oligos. Always leave a space between lanes containing different oligos. Electrophorese 20–30 cm.
5. Place gel on a piece of plastic wrap and then on an X-ray intensifying screen (intensifying side up towards gel). Examine gel with short wave UV light from above (without ethidium bromide staining). The oligo band(s) will appear as a dark shadow.
6. Excise the areas of the gel containing the full-length oligo. Crush the gel slice and elute the oligo in 1 mL TE overnight at 37°C with shaking.
7. Transfer the eluted solution to a fresh tube with a 1 mL pipette. Wash the gel slice with a 200 µL of TE and combine with the eluted solution. Measure the total volume of the eluted oligo.
8. After washing a PD 10 column with 12 mL of autoclaved water, add the eluted oligonucleotide to the column and discard the flow through.
9. Wash the column with autoclaved, distilled water, using a volume equal to 2.5 minus the total volume of the eluted oligonucleotide which was loaded onto the column. Discard the wash flow through.
10. Elute the oligonucleotide by adding 1 mL of autoclaved, distilled water to the column and collect this fraction in a sterile tube. Add another 1 mL of autoclaved distilled water and collect it in a second tube.
11. Dry the oligo under vacuum.
12. Dissolve the oligo in 100µL of TE.
13. Add 100 µL of phenol:chloroform:isoamyl alcohol and vortex thoroughly. Spin at room temperature for 5 minutes at 14,000 x g. Remove 90 µL of upper aqueous layer and transfer it to a fresh tube.
14. Add 45 µL of 7.5 M NH₄OAC and 350 µL absolute ethanol. Vortex to mix, store on ice for 10 minutes, and spin for 10 minutes at 14,000 x g at 4°C.
15. Remove supernatant, wash pellet with 70% ethanol, and dry the oligo.
16. Dissolve the oligo in 60 µL TE. Verify the oligo concentration using OD measurement, making sure the oligo concentration is greater than 0.5 µg/µL.

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Protocol for adapter production/DNA duplex

This protocol outlines the steps to produce adapters and the formation of DNA duplexes:

• PAGE purify oligos prior to use in building adapters. Alternatively, commercially available PAGE purified, cartridge purified, or HPLC purified primers can be used.
• Verify the DNA concentration by preparing duplicate dilutions and determining the A260.
• Calculate the amount of oligo (nmol) using the formula: Amount of oligo (nmol) = A260 x nmol per OD x dilution factor.
• Calculate the volumes of the oligonucleotides needed based on their concentrations and the total volume of the annealing reaction. Scale the reaction as needed.

• Bring the oligo solution to 65°C in a water bath. Maintain the temperature at exactly 65°C for 10 minutes. It is critical to maintain the oligo at exactly 65°C for the entire 10-minute duration.
• Remove the solution from the water bath and allow it to cool slowly at room temperature for 1–2 hours.
• Store the adapter at –20°C until ready to use.

*10X Annealing Buffer Recipe
Mix the following components for a 50 mL stock solution: