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.
Suggest ways to obtain acceptable results from samples of suspect quality.
Dr. Bronfin: That’s a tricky one. There are some steps you may take to make it better, but at some point if your sample is poor quality nothing will work. If you start with RNA, extraction is important to keep your samples at 4°C during all the process and use a temperature-controlled centrifuge. Nested PCR protocol and increasing the number of cycles may also help.
Dr. Mahajan: Every now and again we receive samples for testing that are either in an uncharacterized matrix or in a matrix known to affect the efficiency of PCR. Our primary solution has been to dilute the sample to a level where we have diluted out any inhibitors while retaining sufficient target sequences to measure in the quantitative range of the assay.
We verify the acceptability of the result by running an aliquot of the diluted test sample spiked with a positive control pDNA. We compare the result of the spiked sample with the result of a sample of water spiked with the same positive control pDNA.
Ms. Meda: With solid tissue samples, the best way to assure results is by training those who take and prepare samples. This is also true for blood or other types of samples. No matter how wonderful your detection system, a lot of things can go wrong during sample collection and preparation in order to obtain a satisfactory answer.
Mr. Murakami: Sequencing of PCR products from gel electrophoresis helps, as does melting curve analysis with SYBR Green with qPCR.
Dr. Palchevskiy: It is difficult to trust any samples containing low-quality RNA. It is easier for DNA samples since for standard PCR you are looking preferably for the presence or absence of the gene or DNA region. But for RNA, because it is message-level expression, low quality might degrade and compromise measuring the exact expression levels.
If suspect quality means inhibition products from either purification or RT reaction, it is always preferable to re-purify the sample either by column (with DNase digestion for RNA) or precipitation. If suspect quality means degradation (more concern for RNA than for DNA samples) then I would say change the protocol either of sample collection or storage.
I spoke with Core facilities at UCLA and they mentioned that there are specific kits for microarray based on degraded samples. However, you need to analyze the data with extreme caution.
Mr. Papenfuss: The easiest, most straightforward approach is a commercially available PCR cleanup column. It may be worth trying a few to see which works best with your sample types and PCR system. If cleaning up your sample is not possible, perhaps structure the experiment so a yes or no answer is sufficient instead of going after actual Ct values.
Do you employ concentration or other preparative methods for dilute samples?
Dr. Bronfin: During the DNA/RNA extraction step, for some low-concentration samples, we use less extraction volume compared to the original sample volume. It’s important to remember that although lower extraction volume may give you a higher concentration, it usually yields less total DNA. We don’t use any concentration column or other similar methods.
Dr. Palchevskiy: Our laboratory prefers not to concentrate samples as any such procedure results in more manipulation of RNA and even greater risk of degradation. However, if necessary and if there are enough samples, we might use lithium chloride or ethanol precipitation (mostly for DNA).
Another alternative if the sample is dilute and not pure, we might employ re-purification of the sample and re-suspension in the smaller volume. It is our practice to re-suspend samples in the smaller (minimum) volume (usually 20 µL) since it is easier to dilute than to concentrate sample.
Mr. Papenfuss: The most basic concentration method, simply evaporating off some of the water in the sample, works for very clean samples but may lead to problems with some real-world samples, due to the fact that it also concentrates any PCR inhibitors/contaminants present.
A PCR cleanup column is one of the best solutions. The sample gets cleaned up, which is a bonus, and then you elute the sample into water or buffer, concentrating it as much as needed. Another possibility, depending on experimental goals, is to look into digital PCR, which allows for excellent quantification of fairly dilute samples.