There’s piloting a brand new technique for the first time. Then, there’s jumping through hoops trying to get an established lab technique to work. The former, in contrast to the latter, is expected to be fraught with hardships. Yet troubleshooting an old lab technique that isn’t working anymore, is frustrating at a whole new level. Find comfort in that it probably happens more than you’d expect. In fact, it happened with me and my (now) good buddy qPCR.
Is it Me or the Transcript?
Years of our lab’s data had been produced using qPCR. So much so that it was assumed there were two outcomes: it either showed clear results or nothing amplified because you messed up. When I joined the lab, I became interested in a gene that I now know is expressed in very low levels in my tissue of interest. None of our old qPCR techniques gave consistent results, but it was clear something was there. It was like the data just wasn’t clean enough to be meaningful. I was terrified it was just me.
When it comes to highly reliable and sensitive qPCR, many qPCR experts agree that template quality is priority number one.1 Template-lurking inhibitors can greatly decrease your ability to detect small differences between samples or, in my case, lowly expressed genes. These inhibitors could include reagents left over from your extraction (like phenol, chloroform, or detergents) and/or things that tagged along with sample collection (like cell culture media, bile salts, and heparin). This list of things that will inhibit your qPCR reaction is pretty impressive and won’t necessarily show up in your A260/A280.2
Inhibitors will also keep you from achieving the efficiency assumed by the widely used quantification method, delta-delta Ct. Using this method, you assume each template is amplified with 100% efficiency — every DNA template makes two DNA copies each cycle. If amplification efficiency is less than that, you have to get into some nitty gritty mathematical corrections.
Never fear, though, because a clever group of researchers came up with a simple test to see if your qPCR reaction contains inhibitors. It’s called the SPUD assay and here’s how it works.
How to Make SPUD Your Bud
Nolan et. al. 2006 described this delightfully simple technique, thus ending the long reign of chaos and bringing about the uninhibited qPCR golden age.3 To see if your qPCR reaction contains inhibitors, you can toss in a completely separate qPCR reaction with your samples and see how it amplifies. The SPUD assay does this using a potato’s (get it?) photoreceptor gene. Unless you happen to be studying potatoes, then your template won’t contain this gene (which leads to the potato root turning purple). You include the gene’s synthesized amplicon and primers within your sample reaction. Then you compare its crossing point in the presence of your sample to its crossing point when amplified alone. If, for example, you have residual phenol left over from your RNA extraction in amounts too small to be picked up on the A260/A280, but enough to inhibit your qPCR reaction, then the crossing point of the potato gene run with your sample will be larger.
So whether you’re frustrated by poor amplification efficiency or if you’re just trying to increase the sensitivity of your qPCR reactions, the SPUD assay might be just what will get your goals off the ground. In my case, it turns out I wasn’t troubleshooting. I was simply optimizing!
- Bustin SA and T Nolan (2004) Pitfalls of quantitative real-time reverse-transcription polymerase chain reaction. Journal of Biomolecular Techniques. 15(3): 155–166.
- Rossen L, et al. (1992) Inhibition of PCR by components of food samples, microbial diagnostic assays and DNA-extraction solutions. International Journal of Food Microbiology. 17(1): 37–45.
- Nolan T et al. (2006) SPUD: a quantitative PCR assay for the detection of inhibitors in nucleic acid preparations. Analytical Biochemistry, 351 (308–310).