I am using the 96.96 Fluidigm gene expression array with samples from various mouse tissues (not single cell), as well as a set of purchased fetal cDNA. In the attached image, the group of amplification curves with plateaus of ~0.02 consist of my mouse tissue samples, whereas those that reach a plateau of 0.04-0.06 are the fetal cDNA samples. Can anyone explain the discrepancy? It appears PCR is inhibited in my mouse tissue samples.
Mouse tissue RNA was isolated with TRIzol, phase separated with phase-lock gel, and run through RNeasy columns. 260/230 and 280/230 are >2 for most samples; RINs are 8.5-10. Made cDNA with 200ng RNA and 20 pre-amp cycles, all with Fluidigm kits. Pre-amp'd samples were serial diluted from 5-fold to 120-fold.
Fetal cDNA (20ng) was pre-amp'd with 14 cycles and serial diluted from 3-fold to ~5 million fold. This follows Appendix 1 of the single cell analysis document attached.
Are the mock RT CT values, electrophoresis data and dissociation curves in line with expectations? Regardless, I recommend sequencing the PCR products. Mother nature constantly throw's "curveballs" at us.
I ran a plate-based qPCR and all samples looked good. I've since run two more 96.96 chips and all samples showed good amp curves, and expression matches published results. I still don't know why I had dampened curves for my original run, but it appears everything is working. I settled on 10 pre-amp cycles with 10-fold dilution after Exo I treatment.
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