Using the acronym qPCR to reference real-time (or quantitative) polymerase chain reaction helps to distinguish it from reverse-transcription PCR (RT-PCR). For my molecular biology experiments, I performed reverse transcription using GE Healthcare’s First-Strand cDNA Synthesis Kit to generate cDNA. I then used these products as cDNA templates for quantitative PCR, so the whole process is called RT-qPCR. Bio-Rad provides an overview of the whole process, from RNA isolation to cDNA generation to analysis of the samples. For this post, I will highlight helpful tips for setting up the real-time PCR/quantitative PCR (qPCR) analysis.
- Prepare your reagents and calculations beforehand. In this case, know the number of targets and experimental replicates. Use a “calculation worksheet” with a table corresponding to each of the 96 plate wells. Depending on the amount of samples, the loading plan may change, so fill in each cell of the table every time. Then color-code them to make it easier to visualize, organize, and keep track as you go through pipetting. This will also help once you scale from just one plate to several plates.
- As with PCR, make more than enough master mix. It is actually much more important to do so for qPCR because the emphasis is on consistency. Going back to make and add just a little bit more master mix both interrupts workflow and introduces experimental error, rendering results unreliable. For my calculations, I factor in an additional five wells for every 20-30 wells. Master mix discrepancies can also be due to old or unserviced multi-channel pipettes, so make sure those are calibrated, up-to-date, and properly operational.
- Remember to use different pipettes for standards versus experimental samples. This is very important for avoiding cross-contamination. Furthermore, designate these pipettes for qPCR only.
- Use filtered tips!
- When making serial dilutions, mix thoroughly and completely. As with PCR, perform quick vortexing and collection using a tabletop centrifuge.
- When you attach the tips to your multi-channel pipettor, double-check that each tip is fastened tightly and consistently every single time. Check manually and visually. Develop a consistent routine for this.
- This is just one of the myriad experiments where consistency and organization are crucial. Keep track of which direction you are pipetting because if anything somehow drips or leaks, it will contaminate that well where your pipette (if malfunctioning) was just passing over. Try pipetting standards first on the top row, switch pipettes, and then pipette samples right below. This way, you won’t accidentally contaminate samples with standard because the standards are already placed.
- To minimize exposure and contamination, I used the unopened seal for the 96-well plate to cover the plate in between every step while I was pipetting and moving around. This guards against microscopic debris that can accidentally fall in—especially in a busy lab with people whizzing by, coughing, sneezing, etc.
- After the master mixes and samples are pipetted, double-check the plate by looking underneath. This will help catch any potential mistakes. Make sure all the wells are consistently loaded with the same amount. If they’re not, make a note.
- Once the seal is applied to the top of the plate, use a plastic press tool to tightly seal the plate and make sure every individual well is sealed. To do this, press down firmly on top of the seal, swipe from one edge to the other, and then press down between each row and column and around the corners and edges of the plate. Fun fact: This is the same tool used by automotive detailers!
For my experiments, I used a Bio-Rad machine to perform the reads. I used SYBR Green Supermix from Bio-Rad as well. Finally, we saved values Cq and C(t) for further calculations.
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Mike has a Ph.D. in Biomedical Sciences from the University of California, Riverside, a M.S. in Cell and Molecular Biology from San Francisco State University, and a B.A. in English from the University of California, Berkeley.