Enter the amount printed on your primer tube to get how much TE or water to add for a 100 µM stock. Then work out the dilution to a 10 µM working solution and how much to add to each PCR. Amounts in µg or OD260 units work too.
Runs in your browser.
1Resuspend the dry primer
Amount in the tube
Add 250 µL
of TE or nuclease-free water to 25 nmol for a 100 µM stock.
2Make a working solution
10 µL stock
plus 90 µL of water makes 100 µL of 10 µM, a 1:10 dilution.
3Add to each PCR
Most PCR protocols use 0.1 to 0.5 µM of each primer.
0.5 µL per reaction
of each primer at 10 µM gives 0.2 µM in 25 µL.
Volumes are rounded to what a pipette can measure. Everything is calculated in your browser.
Divide the amount of oligo by the concentration you want. A 1 µM solution holds 1 nmol per mL, which is 0.001 nmol per µL, so the factor of 1,000 converts mL to µL.
Volume (µL) = amount (nmol) ÷ concentration (µM) × 1,000
For a 100 µM stock that comes down to a shortcut: add 10 µL per nmol. For 10 µM, add 100 µL per nmol. A tube with 25.3 nmol needs 253 µL for 100 µM.
| Oligo in tube | For 100 µM | For 10 µM |
|---|---|---|
| 5 nmol | 50 µL | 500 µL |
| 10 nmol | 100 µL | 1,000 µL |
| 25 nmol | 250 µL | 2,500 µL |
| 50 nmol | 500 µL | 5,000 µL |
| 100 nmol | 1,000 µL | 10,000 µL |
TE is the safer choice for a stock you will keep. Tris holds the pH near 8, while water is often slightly acidic, and EDTA binds the metal ions that nucleases need. Nuclease-free water is fine for working dilutions and for oligos you will use within a few weeks.
The small amount of EDTA carried into a PCR from a primer in TE is negligible next to the Mg2+ in the buffer.
Some spec sheets and older tubes give the amount as a mass or as OD260 units. Convert to nmol first.
nmol = µg × 1,000 ÷ MW
nmol = OD260 × 1,000,000 ÷ ε260
One OD260 unit is the amount that gives an absorbance of 1.0 at 260 nm in 1 mL with a 1 cm path. For a typical 20-mer that is about 5 nmol, or 30 to 35 µg.
For an oligo without a 5′ phosphate, MW (g/mol) = 313.21 × A + 289.18 × C + 329.21 × G + 304.2 × T − 61.96.
ε260 comes from the nearest-neighbor method: add the values of every pair of neighboring bases and subtract the values of the internal bases (Cantor, Warshaw and Shapiro, 1970). Vendors use slightly different methods, so when your spec sheet lists ε260 or MW, type that value into the calculator.
Modifications such as a 5′ phosphate, fluorophores or biotin change both numbers. Use the spec sheet values for modified oligos.
A 10 µM working solution is easier to pipette into PCRs and protects the stock from repeated freezing and thawing. Use C1V1 = C2V2.
Stock volume = working conc. × final volume ÷ stock conc.
10 µM × 100 µL ÷ 100 µM = 10 µL of stock, topped up with 90 µL of water.
Most protocols use 0.1 to 0.5 µM of each primer in the final reaction. The volume to add is the final concentration times the reaction volume, divided by the working concentration.
Primer volume = final conc. × reaction volume ÷ working conc.
0.2 µM × 25 µL ÷ 10 µM = 0.5 µL of each primer. For 24 reactions that is 12 µL of each, plus about 10% for pipetting loss.
Enter the same numbers in the calculator to check each step.
The spec sheet says 25.3 nmol. You want a 100 µM stock and 100 µL of 10 µM working solution.
253 µL for the stock, then 10 µL of stock plus 90 µL of water.
You have 150 µg of the GAPDH forward primer GAAGGTGAAGGTCGGAGTC (5 A, 2 C, 9 G, 3 T).
Add 252 µL. The stock is 100 µM × 5,957.94 ÷ 1,000 = 595.8 ng/µL.
The same primer, measured as 5 OD260 units. Its nearest-neighbor ε260 is 197,100 L/(mol·cm).
Add 254 µL. If the spec sheet gives a different ε260, type it in; vendors calculate it in slightly different ways.
Multiply the nmol on the tube by 10 and add that many µL. A tube with 25 nmol needs 250 µL of water or TE for a 100 µM stock.
Dilute it 1:10. Mix 10 µL of the 100 µM stock with 90 µL of water or TE to get 100 µL of 10 µM. For a different volume, use one part stock to nine parts water.
TE is better for the stock. Its Tris keeps the pH near 8 and its EDTA binds the metal ions nucleases need, so the oligo stays intact longer. Nuclease-free water is fine for working dilutions and for primers you will use soon.
Divide the mass by the molecular weight and multiply by 1,000: nmol = µg × 1,000 ÷ MW. A 20-mer weighs about 6,000 g/mol, so 30 µg is about 5 nmol. Paste the sequence into the calculator to get its exact MW.
One OD260 unit is the amount of oligo that gives an absorbance of 1.0 at 260 nm in 1 mL with a 1 cm light path. For a typical 20-mer that is about 5 nmol, or 30 to 35 µg. The exact amount depends on the sequence through its extinction coefficient: nmol = OD × 1,000,000 ÷ ε260.
Synthesis yield varies, so vendors measure each oligo after purification and print the amount delivered. Use that number, not the synthesis scale you ordered.
Yes. After shipping, the dried oligo can be on the cap or the walls of the tube. A short spin collects it at the bottom so none is lost when you open it.
Usually 0.1 to 0.5 µM of each primer in the final reaction. From a 10 µM working solution, 0.5 µL in a 25 µL reaction gives 0.2 µM. Check your polymerase protocol: many Taq protocols use 0.2 µM and many high-fidelity enzymes 0.5 µM.
Keep the stock at −20 °C and use a separate working aliquot day to day, so the stock is not thawed and refrozen each time. Frozen in TE, primers stay usable for a long time.
No. The calculator runs in your browser.