Given PCR’s sensitivity, how do you deal with potential contamination issues?
Dr. Bronfin: We perform all our work in biosafety cabinets and we have an SOP in place for cleaning them. We have separate suits for handling samples, preparing master mix, and making controls and standards. As a rule, during extraction and setting the plate only one tube is open at any given time. We always prepare samples in the following order: negative control and/or blank, samples, positive control. Gloves are changed after each step and if contamination is suspected.
Dr. Mahajan: Since we share the building with our pDNA manufacturing facility, potential contamination issues are a particular challenge. This is especially true with our limits assays such as integration or cleaning verification where the expected result is a low or negative PCR result.
In addition to the standard negative control samples in each run, our primary method of eliminating potential contamination issues is through strict adherence to procedural controls (i.e., dedicated and isolated work spaces, rigorous cleaning of work spaces before and after reaction set up, barrier tip pipette tips, etc.). These practices have served us well so far, and we have not had any significant contamination issues.
Ms. Meda: PCR has changed a lot in 20 years. You no longer need three separate rooms to carry out PCR, and reagents have improved tremendously. Even today, you can’t expect PCR to work well in a contaminated environment. However, reagents have improved tremendously, and the method has become more robust. The advent of Roche AmpliPrep, and many of the TaqMan real-time PCR reagents have improved the technology tremendously as well.
Mr. Murakami: There are two possible sources of contamination in our work: the DNA template or reagents. Carefully collected tissue samples should not contain genetic contamination, while premixed reagents containing buffer, MgCl2, and Taq polymerase minimize contamination. It becomes essential to improve handling and processing through less pipetting, particularly for the master mix preparation. Automated dispensing is just one option we’re using for sample preparation.
Dr. Palchevskiy: Since many things can inhibit qPCR reaction, it is necessary to dilute your cDNA several fold and use it as a standard curve to check efficiency. If any contaminants are present, the curve and the Ct vs. concentration would not be what you expect it to be, and it will point to the potential source and solutions. Diluting out the cDNA sample is one way to minimize any inhibition effects that might be introduced by RT reaction.
Mr. Papenfuss: Contamination should rarely be an issue if you follow good laboratory practices. Often, filtered pipette tips and “good lab hands” are sufficient, but that’s obviously not the case in all labs with all operators. Filtered tips are an absolute must. A good first practice is to use a commercially available surface decontaminant, designed to remove DNA, on surfaces and pipettors before use.
Wear gloves—not to protect yourself from the reaction, but to protect the reaction from the operator. Obviously, if any solutions containing DNA get on the gloves, they should immediately be changed before spreading any contamination. While setting up the reactions, be aware of where you are moving the pipettors. For example, a pipette tip full of DNA should never be passed directly over a negative reaction or other unknowns.
In some circumstances, it makes sense to have separate areas in the lab for addition of unknown samples and positive controls, lowering the risk of contaminating the area where clean master mixes and negative reactions are prepared. Once finished, decontaminate again.