Resuspending primers is straightforward: add the right volume of buffer or water to the lyophilized pellet in the tube, mix gently, and store at −20°C. The key step most people want help with is calculating how much liquid to add, which depends on the nanomole (nmol) yield printed on the tube or spec sheet. Once you know that number, everything else follows a simple formula.
What You Need Before You Start
Primers arrive from the manufacturer as a dried-down (lyophilized) pellet at the bottom of a small tube. The tube label or accompanying spec sheet lists the total yield in nanomoles. This number is all you need to calculate your resuspension volume. Have nuclease-free water or TE buffer on hand, along with a calibrated micropipette, a vortex mixer, and a microcentrifuge.
Choosing TE Buffer or Nuclease-Free Water
Either works well under normal lab conditions. At −20°C, primers remain stable for at least 24 months whether stored dry, in TE buffer, or in nuclease-free water, with no significant difference in performance by qPCR. At 4°C, stability extends beyond 60 weeks regardless of medium.
The difference shows up only at higher temperatures. At 37°C, primers in TE buffer and dried primers held up for about 25 weeks, while primers in nuclease-free water began losing function after roughly six weeks. So if your freezer is unreliable or you expect your primers to sit at room temperature during shipping or benchwork, TE buffer (10 mM Tris, pH 8.0, 0.1 mM EDTA) offers a safety margin. The slightly alkaline pH helps prevent the slow acid-catalyzed breakdown of DNA, and the small amount of EDTA chelates metal ions that could promote degradation.
One caveat: if you’re using your primers in an assay that’s sensitive to EDTA (some enzyme reactions, for example), nuclease-free water is the safer choice for the working stock you’ll pipette directly into reactions.
Calculating the Resuspension Volume
Most researchers make a 100 µM stock solution. The math is simple: multiply the nmol yield on the tube by 10, and add that many microliters of liquid.
Here’s why that works. A 100 µM solution equals 0.1 nmol per µL. So if your tube contains 29.4 nmol of primer:
29.4 nmol ÷ 0.1 nmol per µL = 294 µL
Or, equivalently: 29.4 × 10 = 294 µL. Add 294 µL of TE buffer or nuclease-free water, and you have a 100 µM stock.
If your yield is 42.7 nmol, add 427 µL. If it’s 15.0 nmol, add 150 µL. The factor-of-ten shortcut saves you from setting up a full concentration equation every time.
Step-by-Step Resuspension
Give the sealed tube a brief spin in a microcentrifuge before opening it. This pulls any dried material that may have settled on the cap or walls down to the bottom, so you don’t lose primer when you pop the lid.
Add the calculated volume of TE buffer or nuclease-free water directly to the pellet. Then mix gently by pipetting up and down several times or by vortexing at a low speed. Avoid vigorous vortexing, which creates air bubbles and can make it harder to pipette accurately later. After mixing, give the tube another brief centrifuge spin to collect any liquid clinging to the walls. Let the tube sit for a few minutes at room temperature to ensure the pellet is fully dissolved, especially for high-yield tubes where you’re adding a larger volume.
Label the tube clearly with the primer name, concentration (100 µM), date, and your initials. Store it at −20°C.
Making a Working Stock
You don’t want to pipette directly from your 100 µM master stock for everyday PCR reactions. Each time you open that tube, you risk contamination and introduce a freeze-thaw cycle. Instead, make a 10 µM working stock by diluting 1:10.
Add 10 µL of the 100 µM stock to 90 µL of nuclease-free water in a fresh tube. Vortex briefly and spin down. You now have 100 µL at 10 µM, which is a convenient concentration for most PCR setups that call for final primer concentrations in the 0.1 to 0.5 µM range. When this aliquot runs out, make a fresh one from the master stock.
This two-tier system keeps your master stock frozen and untouched most of the time, extending its useful life well beyond what you’d get if you thawed it for every experiment.
Storage and Freeze-Thaw Limits
At −20°C, your resuspended master stock is stable for at least two years. At 4°C, it holds for over a year. Keep working stocks at −20°C as well, or at 4°C if you’re using them within a few weeks.
Freeze-thaw cycles are less damaging than many people assume. One study found that primer-probe mixes subjected to monthly freeze-thaw cycles for five months showed no significant change in DNA quantification by qPCR. That said, minimizing freeze-thaw is still good practice because it reduces the chance of nuclease contamination from repeated tube openings. Making small working aliquots is the easiest way to avoid the issue entirely.
Verifying Primer Concentration
If an experiment is failing and you suspect your primers might be off-concentration, you can check them on a spectrophotometer. Measure absorbance at 260 nm (the wavelength where DNA absorbs light most strongly). To avoid using too much primer, dilute a small aliquot first. For example, adding 10 µL of primer solution to 990 µL of TE buffer gives a 1:100 dilution in a 1 mL volume suitable for a standard cuvette.
The actual concentration calculation requires the extinction coefficient for your specific primer sequence (provided on the spec sheet) and the path length of your cuvette. The formula is:
Concentration (mol/L) = (dilution factor × A260) ÷ (extinction coefficient × path length in cm)
Most researchers only bother with this step when troubleshooting. If your PCR is working fine, the manufacturer’s reported yield is reliable enough for routine use.
Common Mistakes to Avoid
- Skipping the initial spin. Lyophilized primer can stick to the cap. If you open the tube without centrifuging first, you may lose material and end up with a lower concentration than you calculated.
- Using DEPC-treated water. Residual DEPC can modify DNA bases and interfere with enzymatic reactions. Use nuclease-free water that was not treated with DEPC, or use TE buffer.
- Storing at room temperature. Primers degrade faster above 4°C, especially in water. If you accidentally leave a tube on the bench overnight, it’s probably fine, but don’t make a habit of it.
- Pipetting from the master stock for every reaction. This wastes your full-concentration stock and increases contamination risk. Always make a diluted working stock.