I have transducted a cell line (TGBC2) with lentivirus with different shRNA sequences and controls. My controls were scramble and empty. The backbone is pLKO.1-puro. If it were just the shRNAs and controls it would be okay, 3/5 shRNA sequences show 80% reduction relative to control or more of the protein (GPC1), so that's good. The problem is that when comparing the empty and scramble to the WT they show almost 3 times less protein expression than the WT. I have no idea why this could be. I don't know if this could be because of the backbone or maybe the transduction process itself. For the scramble MAYBE it could be related to some unknown off-target but for the empty vector I have no explanation. The cells didn't look to be contaminated and grew as they always did but I can't completelly discard that theory. What could be causing this?
Firs of all, perform the transduction of the empty lentivirus and measure the changes of expression of your gene of interest. If there are no significant changes between wild type and transduced cells, your vector per se cannot affect the transcription, and the problem can be in your control shRNA. Try to clone another control, sometimes there are effects, based on interference of control shRNA and noncoding RNAs. Some of them could affect the expression of your gene, or even group(s) of genes.
The change between the WT and empty control is wild, 3x and 6x on each cell line. The vectors have been validated by the provider (Sigma) but from what I've been gathering quite apparently the design of the shRNAs isn't the best and I think they might produce a lot of off-target. Since one of the cell lines I worked isn't even published yet there isn't much information on it. For one of the cell lines the empty caused 90% silencing of my protein, but the scramble was alright; and when I redid the WB they both were pretty similar. Thank you very much for all your suggestions!
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