Successful PCR runs require more than excellent primer design. Here are seven best practices for both obtaining consistently high-quality data and ensuring the integrity and economic use of your PCR reagents.
- Centrifuge lyophilized primers upon delivery. While the DNA is usually present as a nearly invisible film on the bottom of the tube, it can come loose and fall out when the cap is removed for the first time.
- Resuspend primers in 10 mM Tris pH 7.5, 1 mM EDTA solution (TE buffer) instead of water. The Tris and EDTA prevent acidic water and contaminating DNases from hydrolyzing and enzymatically degrading DNA, respectively.
- Aliquot the resuspended primers into working stocks. This eliminates the need for damaging freeze / thaw cycles of the master stock as the working stocks can be removed from the freezer three to five times without degrading the DNA. If contamination of a working stock occurs, it can be thrown out and replaced with another without compromising the master stock.
- As with your primers, if you purchase large volumes (e.g., 50 mL) of 2X PCR premixes, you can aliquot these reagents into smaller working stocks that are suitable for a single experiment (e.g., a 96-well plate) to avoid contamination.
- For each target gene you are working on, make a “master mix” of your 2X PCR premix, water, and primers. Some people factor in extra replicates or a percentage (~10%) of volume for each master mix to account for pipetting error. You can then add this “master mix” to the tubes/plates first and then add your samples.
- If you want to be “extra-passionate” about the precision between your technical replicates, you can even make a “master mix” for each target and sample—add the 2X PCR premix, primers, water, and template—and then add everything all at once into your tubes/plates. This will require extra tubes, but the intra-replicate variability can be greatly reduced.
- Always keep your pipettes calibrated and take your time. Reagents are expensive, and samples can be limited.