Hi, I'm having a bit of trouble with my experiments so if anyone has any ideas please let me know!
I've been transiently transfecting cells with my gene of interest, waiting 24 hours and then extracting the total RNA and using qRT-PCR to measure it's expression.
I'm trying to optimise a time course experiment where I ultimately inhibit transcription using a drug (e.g. DRB) and extract total RNA over a time course to measure the transcript degradation rate of the gene I'm interested in.
So far I've been just been trying extracting RNA from cells without transcription inhibition over an 8 hour period, starting 24 hours after transfection (cells have reached100% confluency). I expected that the fold increase (2^ddCt) in expression above endogenous DNA would remain pretty much steady over the time course, but I've been getting wild results.
For example, at 0 hours the fold increase was 157, then 60 at 2 hours, 68 at 4 hours, then up to 229 at 8 hours. On other runs I've also gotten weird results that don't remain constant across time.
I don't know how to explain this, or what to change in my methods to get steady fold differences across the time points. I don't think it's DNA contamination, as I DNase the samples and check for residual contamination with a standard PCR/gel electrophoresis. I'm also not convinced it's RNA degradation as the bands I get when I run the samples on a gel are pretty good (a bit smudgy though).
My only other thoughts are that it might be due to the cell cycle, or because the cells are stressed at 100% confluency and behaving strangely?
If anyone has any thoughts please let me know