The thing is that I am transfecting both protein isoforms separately in 10 cm dishes with COS-1 cells and FuGENE® HD Transfection Reagent. I am transfecting 3 µg of the each vector containing each isoform with 12 µl of the FuGENE® HD Transfection Reagent (1:4 ratio). I am lysing the cells with the NP-40 lysis buffer. I'm using a first mouse anti-Myc tag antibody and a second peroxidase labelled anti-mouse antibody. Well, the point is that at the end I get a strong band signal for the bigger 36.7 kDa protein isoform but I don't get any signal at all for the smaller 15.6 kDa protein isoform. I cannot detect this last one. I sequenced both vectors and both myc tags are perfectly in frame. I also performed a Beta-Tubulin Loading Control and that's not definitely the problem. The Cell line used should be the optimal one. Perhaps, since the smaller protein isoform (15.6 kDa ) is much smaller than the other one, maybe I should use a stronger lysis buffer in order to allow the antibody to be more accessible to the epitop. As could be easily renaturated somehow.
P.D. I am talking about a cytoplasmatic (soluble) protein. Any suggestions?
Do you know if the smaller protein is being expressed? Do you see a stronger band at 15kDa in a coomassie-stained gel from transfected cells vs non-transfected cells, for example? Or maybe do some immunofluorescence microscopy to detect it that way? There are probably a thousand things you can tinker with to optimise the biochemistry, so I would start with checking expression.
Hi Michael!
Yes, I do certainly know that both protein isoforms are being expressed by a qrt-pcr analysis. In fact, the smaller protein isoform is much more overexpressed than the bigger one under the same exact conditions. The thing is that we got the band of the smaller protein isoform once on the western blot but we can't reproduce the experiment anymore. And as I mentioned before should not be a problem of the vectors. We dectect it also with immunofluorescence microscopy but the thing is that we need to get the band in the western blot.
Adding to the last comment by Michael that i think it is important too, have you tried to run a gel with a higher percentage of acrylamide? If you are trying to see a 15.6 Kda protein maybe a 15 % acrylamide gel should do well. I'm a master student and here in my lab we always perform a 15 % acrylamide gel when we want to see small proteins like p11 for example that it's not very far from yours.
NP 40 lysis buffer we usually use it for the whole cell so it should be working well...but you can try another ones like Tris-HCl lysis buffer if you want because is specific for cytoplasmic soluble proteins extracts.
good luck hope it works
A high % gel is a good idea Lina, I didn't think of that.
Iván, can I also ask about your detection method? You're using a myc-tag antibody in the WB, what did you use in the IF? Was that also against myc? Also, is there maybe a protease site between the tag and your protein of interest? (I'm assuming you use protease inhibitors during lysis anyway) I'm just wondering if you're losing the tag along the way.
I was about to say that maybe the epitope was not present in that Isoform, but I see you are detecting the expression through a myc tag and that you saw the expression of the protein by immunofluorescence already. Yes, I agree with the rest, try a higher concentration gel. 12% would do, but better use 15%. I don't think the transfer time is a problem since you still see a very strong band with the 37 KDa isoform, however it would not hurt to reduce the time. Try to load more lysate also. Good luck.
Could the smaller isoform be highly unstable? You do not even see tiny amounts? Tried to enrich by IP?
Hope you solve this problem soon.
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