I am experiencing trouble in screening for an effective shRNA sequence. There are varios approaches (western/northern blot, flow cytometry...). As RNA interference does not produce an absolute phenotype (ie total absence of the target protein), I think the best approach for an immediate sequence selection is flow cytometry. I have designed 4 shRNA candidate sequences which are currently cloned into an expression vector, also expressing the eGFP. My target protein is cloned into an interim expression vector which does not produces any reporter tag or protein. I tested two out of four sequences by cotransfection in 293 cells and subsequent immunohistochemistry (72h after cotransfection) but none of them produced any clear effect. Nonetheless, I performed cell counts to be sure that my shRNAs were not working at all. I couldnt find any drop in the number of cells expressing my target protein, but I found what seemed to be a slight drop in the intensity of fluorescence.
I think the best course of action now, before I test the last two shRNAs sequences, is to subclone my target protein into an expression vector that produces a fluorescent reporter protein different from eGFP. The target protein must be cloned in-frame with the reporter protein or the latter be expressed from the same messenger via an IRES signal. Then, I should perform a cotransfection assay and analyze the fluorescence levels by flow cytometry. I think that is the faster and most precisse way to compare the fluorescence between different conditions. ¿Do you think I am right? Any advice is welcomed...
Finally, I have a more prosaic and methodology-related question. I have been reading some protocols to test shRNAs by cotransfection. Some of them propose to mix the target cDNA plasmid with the plasmid containing the silencing cassette in 1:5 molar ratio (in my experiment, I used an 1:1 ratio), but they dont explain this point further. ¿Can anyway shed light on this issue?
Thanks in advance.
Edited by litos, 01 July 2009 - 10:55 AM.