I do not work much with proteins and I do not know how the experiment with establishing protein half-life should be done.
I work with HEK293T cells. I want to measure the half-life of my truncated protein which is expressed from plasmid (overexpression). In cells there is also a Wild-Type variant of this protein (slightly longer) with low level of expression.
I did a transfections with wild-type plasmid for overexpression of WT protein and separate transfection with plasmid expressing truncated protein.
After 48h, I added a cycloheximide in conc. 0.1 mg/ml to see the difference in WT and truncated protein stability.
I made 2 time point, 4h, 8h, because I don't know what is the half-life of my WT protein. Maybe a longer time is needed?
When I did a western blot, i did not see any changes in stability of each proteins, though, I did not have a good control of protein with short half-life.
My main question is, whether it is OK to load the same amount of protein (~30ug) for each time-point, no matter how many cells has survive this experiment, (after 8 h the cells are less viable). Should I somehow normalise the loading amount of protein to the total cell number? How?
What kind of controls should I include: protein with long half-life like: GAPDH?, any others?
and protein with short half-life: like tp53, maybe some others, with half-life less than 4h?
Waiting for reply,