I am new here, and I am searching these kinds of forums for some input! If anybody has some ideas, suggestion or input I would really appreciate it!
I am a master student, and I am looking at a candidate tumor suppressor gene. I have made stable transfectants, and expanded 23 monoclonal clones. The parental cell line (a sarcoma cell line) has a homozyogus deletion of the gene of interest. All the clones were tested for mRNA using TaqMan gene expression assay (real-time PCR), and all of them have expression. Additionally, two probes are used to cover the length of the transcript, one in the first exon-exon boundary and the other in the last exon-exon boundary. From these results it seems like the whole transcript is expressed. The samples were DNase-treated prior to cDNA-synthesis. However, I did not run the samples on a gel to test whether the DNase treatment really had worked. The ORF was sequenced and is OK.
However, when I perform western blot, only 12 of the 23 actually have the protein. I thought there should be a pretty good correlation between mRNA and protein level?
It does not seem to be a (good) correlation between the Ct-values and the amount of protein seen on western blot. Some clones show really high mRNA expression, but still have no protein expression. The clones are maintained in 50 % of the selective antibiotic.
Is it possible that the cells want to silence the introduced gene, but cannot do it pre-transcriptionally because of the selective agent, and therefore does it post-trascriptionally? Or translational control? Or is it normal? My supervisors have never done transfection, so they have no ideas. I have been reading about gene expression control, but a lot of them involves untranslated regions or phosphorylation of initiation factors of translation, and that doesn't seem relevant..
Thanks for your time :)