Dear Dymitrij,

Protein degradation is the simplest explanation, especially if the protein can disturb the cell in the survival.

Best - Jan

Thanks, Jan. It is possible, because the protein is a transcription factor and transcription factors are often unstable in vivo proteins. However, earlier works suggest that this is a relatively abundant protein: about 3000 molecules per cell.

Dear Dima,

Could be your tag is blocking the interaction with another transcription factors, thus leading to the degradation. I would look at the structure, if available to check where is your tag placed in the transcription complex.

Best - Jan

Nice idea. At first glance, the C-terminally tagged protein was successfully used in experiments. Who knows. In principle it is possible. There is, at least, one report that says that this protein interacts with one other transcription factor.

The fact that 3HA is detected in Westerns and there is a consistent mol wt of 40kDa in all experiments, the likelihood of cleavage by a protease is high. Try expressing at low temperatures overnight or so (I m no expert in yeast protein production though).

Alternately, us system could be using an internal start codon in your gene (if it exist) and make a smaller protein.

The reasons may be target protein has been cleaved or digested, Splice variants exist or another protein bearing the same/similar epitope has been detected by antibody. Possible solutions are use a fresh sample which has been kept on ice, add fresh protease inhibitors to the lysis buffer or try alternate antibody

As Bastian pointed out, a lot of possible explanations must be considered. To distinguish between them, I suggest you try the following:

1) Try different extraction buffers and cocktails of protease inhibitors. If the ratio between higher and lower molecular weight bands stays the same, you can rule out post-extraction effects.

2) Try a pulse-chase analysis (pulse-label for a short period with 35S methionine and take a sample at the end, then chase for longer periods with several samples taken at various time-points). Freeze samples in liquid nitrogen or dry ice with ethanol. When you have all samples, extract proteins and instead of doing westerns, do an immunoprecipitation (with your antibody and then protein A sepharose to pellet them down). Wash the pellets several times, and then load on gel, run it, dry it, and expose to film. If you get the same two bands with high and low molecular weight, but the small band becomes intenser at the expense of the higher, then it is degradation in vivo. If the ratio doesn't change, then it is alternative translation....

Enjoy the search, working with proteins is never boring.

Cheers, Jurgen

I have had a similar issue in the past with a different tag that destabilized my protein of interest. I am not sure what you final objective is, but if you only want to purify the protein to study it in vitro (i.e. enzymatic analysis,..) a quick and dirty way of doing it is cloning that protein in front of an inducible promoter, grow the yeast and induce the promoter for a short period of time. Then extract the protein using a strong cocktail of inhibitors and different extraction buffers. Having a lot of cells when doing the induction allows you to maximize the yields and minimize the time of exposure of your protein to the cell proteases.

Hope it helps! Happy cloning!

Mariette

One possible problem is that the sequence Asp-Pro can be cleaved just by processing protein for Western blots. If the protein is in an acidic buffer and then treated with standard loading buffer and heat, that sequence may be cleaved chemically. See if that sequence is near the likely breakpoint, and if it is, make sure acidic buffers are not present just before denaturation. Hard to do if you have to use TCS precipitation, but that should not be the case in your system.

The protein has two Asp-Pro dipeptides in its sequence. However, cleavage at dipeptide that is the closest to the C-terminal should give about 70 kDa tagged fragment. Indeed, we can see and additional band of 70 kDa.

1. Use alkaline/SDS to extract protein from cells at mig-log phase to see if the small band still exist. By this way, the degradation during extraction can be avoided.

2. Overexpression is possibly the reason. Plasmid numbers in yeasts are not uniform. Even cen plasmids exist different copies in different cells. Native promoter will not prevent the overexpression. Integrating the gene into genome will solve the problem, and will also help to interpret other results.

3. If construct is correct, a fraction of protein degradation won't affect the interpretation for most of results.

You may also try to put another tag at the N-terminus, for example, FLAG or myc, and then do a WEstern blot with both, anti HA and anti FLAG abs

Too simple, but possible: Did you verify the sequence of your gene. PCR amplification during the cloning procedure might have introduced a frameshift or stop codon.

Entire gene with flanking sequences were sequenced and no errors were found. Even if stop codon somehow appeared in the part of gene encoding C-terminus of the protein, then we would not see any band, because protein is C-terminally tagged.

Sorry, missed that point in the question text. I favour Bastians idea of an alternative start site but you are having the natural promoter..So everything is natural from promoter, 5'UTR to start site? At least a bit of correct protein should be detectable then. Is a proeolytic processing of mammalian homologues known?

That is OK. I think it is possible that this band may be a result of alternative translation initiation or degradation. It could be easily detected because it transfers onto membrane much easier than whole protein. I should add that apparent mass of the whole protein in PAAG is 125 kDa. Moreover, after cloning into 5'UTR a tandem array of 3 tetracycline aptamers that should regulate translation of this protein, we found that the major band ran at the level of 15 kDa (!). It is reproducible with different colonies. Such strange behavior could be explained by some internal translation initiation.

You are right about smaller proteins easier to be western-transferred. They also absorb antibodies, and make it harder to see the desired band. Cutting off a part of the membrane (the part below 50kb in your case) will help to see the desired band. The principle is similar to using the low percentage gel, so that small band will run out.