General suggestions for avoiding false positives in your negative template control (NTC) sample:
- Use freshly purified/sterile water, tubes, and reagents.
- We highly recommend that you aliquot your probe and primers into the volumes required for a single experiment. This will decrease the chances of contamination, allow you to start with a fresh tube if you do have contamination, and minimize freeze–thaw cycles that can reduce oligonucleotide quality. Always include appropriate positive and negative controls in your experiment.
- Use separate dedicated PCR work areas for reaction setup, adding the template, and handling the amplification product. Decontaminate PCR work areas regularly with 10% bleach and UV irradiation.
- Use 5% bleach to clean micropipettes regularly. Use sterile, filter tips for pipetting to minimize contamination from aerosols. We recommend having separate pipettes and pipette tips for PCR setup and post-PCR analysis.
- Place the no template control (NTC) wells as far as possible from positive samples.
- Late amplification (beyond cycle 34 for SYBR® Green dye–based assays) may not be indicative of a positive NTC as it could also be a result of dimer amplification. Perform melt curve analysis after PCR to check for primer-dimers.
If you do find contamination in your NTC sample:
- Be sure to replace all reagents and stock buffers and thoroughly clean PCR preparation areas.
- Check whether the probe is degraded. In such situations, there may still be signal from free dye and/or high background. Signal to noise assessment, mass spectrometry, or a fluorometric scan can be used to address possible probe degradation.
- Use a more conserved or novel gene as a control if you are repeatedly seeing false positives.
Author: Darcey Klaahsen, MS, is a Scientific Applications Specialist at IDT.
Avoiding False Positives When Using Universal Primers for Bacterial Identification
In PCR experiments, amplification in the “no template control” (NTC) before the ~38th cycle with probe-based assays (or ~34th cycle when using intercalating dyes) is a sign of false positives and/or contamination. This sidebar specifically addresses false positives that occur during bacterial research when primers and probes are designed to detect common sequences, such as ribosomal RNA (rRNA).
Am I amplifying DNA from my reagents or consumables?
The exponential PCR process can amplify a single copy of DNA to detectable levels. Thus, it is important to consider the pervasiveness of the chosen primer or probe sequences. There are some DNA sequences, such as bacterial genes for 16S or 23S rRNA, that can be found almost anywhere. While not commonly used for RT-PCR or gene expression studies, rRNA sequences are often used for characterizing environmental species diversity, such as bacterial strains within the intestine or in salt water marshes. For such applications, genomic DNA, rather than RNA or cDNA, is used as the sample.
Choose a well-conserved, species-specific gene or novel sequence
In 16S rRNA experiments, a better approach may be to choose a unique sequence from the hypervariable region of 16S rRNA. Alternatively, a conserved, species-specific gene may be used. Blocking oligos and/or clamps can also be used to block amplification of common sequences and enhance the amplification of a rare sequence.
As new sequences are deposited into the NCBI database, it is essential to perform a BLAST search of every primer and probe sequence used in PCR to check for specificity and cross reactivity.
Bacterial ribosomal sequences can be amplified readily from virtually any bacterial source, including bacteria-derived Taq polymerases used to amplify them, as well as nonsterile tubes and pipette tips. Hence, if a positive NTC is observed, testing with different master mixes will help rule out the master mix as the cause of contamination.