Question 3 of 9 Intermediate Scenario
How would you evaluate primer efficiency and assay specificity?
Short answer
I would validate a qPCR primer pair with a dilution series to estimate amplification efficiency and usable range, then check that it produces the intended product. Specificity needs sequence-level evaluation and experimental evidence such as melt curves, product size or probe-based controls.
Why
Primer design begins with a defined target sequence and an understanding of possible off-target templates, isoforms and genomic DNA contamination. Use a tool such as Primer-BLAST to screen primer binding and evaluate predicted amplicons. Then test a series of accurately prepared dilutions in the relevant matrix.
Plot Cq against log starting quantity and calculate efficiency from the standard-curve slope. Examine consistency, linearity and the limit at which quantification becomes unreliable. A single sharp melt peak can support specificity in dye-based assays, but it is not conclusive proof of identity.
Gel checks or sequencing may be useful when uncertainty remains. Include no-template controls and, for reverse-transcription assays, controls for residual genomic DNA. MIQE 2.0 emphasizes proper controls, assay-specific validation and transparent reporting rather than a single magic efficiency number.
An example
An assay gives a tidy standard curve but a late second melt peak and an NTC signal. The primer pair is not accepted until the unwanted product is understood or eliminated.
Common mistakes
- Claiming a good R-squared value guarantees specificity or assuming a high Cq alone proves contamination
The interviewer might ask next
How would you detect an assay that amplifies a close paralog rather than your target?