Hello, I have a problem with ddPCR. Usually I dilute cDNA for it, to get rid of PCR inhibitors from revertase mix. But I was taught that inhibitors are less crucial for ddPCR than for RT-PCR because ddPCR detects the final point anyway (I attach the article about this). So, recently I had a few samples with very little RNA concentration, and I was afraid that if I dilute them 1:4 as usual, I wouldn't register my target at all. And I decided to use them for ddPCR undiluted. To confirm that there was indeed no inhibitors effect, later I diluted two of the most concentrated samples from this batch 1:4 and measured them again (on another plate). I expected the ratio to be either 1:4 (if inhibitors weren't crucial) or less than 1:4 (because more concentrated sample would amplify less in the presence of inhibitors). However, the ratio is about 1:8-1:10. The reference sample that was present in both plates demonstrated similar (+/- 10%) copy numbers for 4 targets, so I guess there was no problem with reaction or droplet assessment. Can anybody explain this to me?
I am not sure about your exact experimental procedure, but it appears you are using RNA samples. Could it be that your target has degraded during the dilution step, or there have been different efficiencies of the reverse transcription reaction?
I know this response comes a bit late, but one thing with ddPCR is that since the dynamic range is from only few copies to about 100,000 copies per reaction, the template samples are often very diluted. Diluted DNA samples may give surprising results due to DNA molecules binding to the tube walls. To prevent this, dilutions should be done fresh just before ddPCR using low-binding tubes and a non-template carrier. Another crucial thing is to ensure high pipetting accuracy.
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