At first, we performed transfections. To analyze that transfection, we performed RNA isolation, cDNA synthesis and qPCR.
Using the qPCR, we analyze multiple samples, whereof each sample we measured this in triplo. The qPCR samples, we combine the reaction of 2 genes in one well, so we measure target and reference gene in the exact sample (target gene is FAM-labeled and reference gene is VIC-labeled).
The result shows us:
1. The reference gene CT values are very stable in the triplo of one sample.
2. The target gene CT values are very unstable in the triplo of one sample.
Does anyone have a cause of the difference in this stability?
It may be as simple as how high the CT value is for the two probes. The higher the CT value for the probe, the more variable the result will be...
If you figure that there is some level of inherent variability (pipetting error, etc), then that variability is going to be amplified exponentially by every PCR cycle. When you're using a reference probe that has a CT of 20 cycles, it will look very stable. By 30 cycles, it will have significant variability, and by 40 cycles it will be highly variable.
Two questions to ask yourself:
1. Is the probe as efficient as it could be? Two options for this; first, you can (and probably should) run efficiency checks (run the same control sample, preferably with known high level expression, at several dilutions. Ideally the CT should increase by 1 for every 1:1 dilution you make. If the CT is dropping by significantly more than 1, that's a sign that your probe is not very good. In contrast, if the CT drops by less than 1, it's a sign that you've got PCR inhibitors in your samples). A second option is to just make another probe or two and run them head-to-head. Even better, make another probe or two, and test them both by dilution!
2. If you think your probe is good, and the absolute CT is really low, it suggests that the samples you're testing have very low expression. Before you work on carefully optimizing everything, ask yourself whether this is actually the answer you need, or whether you want to spend months trying to carefully measure almost nothing vs almost nothing in a whole bunch of samples. (if so, you may want to explore digital droplet PCR, which can provide more accurate measures of low-abundance transcripts).
Another suggestion: If it's a cdna transcript, you may want to try getting a commercial tissue panel to find a tissue that expresses your transcript at high levels. This is a useful control for troubleshooting, and can be a reference to show that the transcript is weakly expressed in your samples.
It happens when you have very high CTs for the target gene. In this case the problem is usually the low volume of cDNA sample you pipette in the well.
It could be as simple as pipetting error. If one sample has a higher or lower volume than the other two, then it's Ct value will be different.
Or evaporation. Basically, check the volumes of your reactions.
Check your DNA integrity and your concentration, check if you are putting a low volume of reactive for PCR. If the different is high between the triplo (more than one unit) you most likely have a pipetting error. If not, maybe is the final volume of the disolution (reactive for PCR, probe(s), and cDNA), is insufficient for you problem cDNA.
I suggest you a test in different wells of you samples.
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