Hello all,
I am trying to knock down a protein that has been documented to be heavily upregulated in oral squamous cell carcinoma (confirmed via western blot). I have attempted to knock down this protein using two different Sigma shRNA (77% knockdown or 99% knockdown documented in A549) but have not had any success. I wanted to validate these shRNA in my cells via transient transfection before moving to stable knockdown and have tried 6 hour polyethyleneimine transfection (1DNA:4 PEI), Effectene transfection at 1:10, 1:25, and 1:50 ratios (24 hour incubation), and electroporation collection after 3, 24, and 48 hour incubation, all with parallel scrambled controls. None of these efforts have resulted in any knockdown of the protein evident via western blot. I have not been able to find any small molecule inhibitors for this protein and am a bit at a loss as to how to get this knockdown working, as the cancer cells are vehemently opposing knockdown. I am currently working on stable transfection (psPAX2 and pMD2G) but this will take multiple days and is not a guarantee. Any ideas would help me greatly!
What's your transfection efficiency? Do you have a marker of some kind to assess successful transfection? It's unclear to me if this is a failure of your transfection or of your shRNAs.
I was going to suggest doing CRISPR knockdown if the problem is in your shRNAs, but that's kind of moot if your transfections aren't working.
Unfortunately there is no marker to assess transfection efficiency. I have been assessing knockdown by comparison to scrambled control. I am currently testing the knockdown in A549 cells to confirm functionality of both shRNA. It seems this issue has been a problem in the lab before, as attempts to knockdown a protein upregulated by a similar cancer cell line was equally difficult, and I have been told that stable knockdown did not last long those cells.
I think the problem here is that you don't know what your problem is. If your transfection efficiency is low (which is totally possible, transfection can be very finicky), then you're not going to detect knockdown in the bulk population since only a subset of the cells have the plasmid. Also, it's going to be really difficult to select stables out if the shRNA-positive population is bottlenecked at the transfection phase.
You need to have a way to assess transfection. In the absence of a marker that you can visualize, you might want to consider using qPCR to see if your shRNA is actually being produced in your transfected cells. It might be hard to be quantitative about it, depending on what kind of controls you have available, but it might at least help you get an idea where your problems might be.
Why not just clone a marker like GFP in tandem with your shRNA construct? It's pretty commonplace practice. Even if you don't have access to fluorescence microscopes or flow cytometry, you can still blot for GFP protein.
If you can conclude that your transfection is working great, then the problems probably are with your shRNA and you may want to redesign them or use a CRISPR knockdown strategy. CRISPR knockdown would be stable, but again you'd still need some sort of marker to select successfully modified cells out of the bulk population - you won't knock down 100% of the cells.
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