I have some problems expressing a His-tagged protein. I cloned the
sequence in-frame into the pET-20b(+) expression vector, which I used
to transform E.coli BL21 (DE3) cells with. I thoroughly checked that
insert and His-tag are in-frame. I grew the culture at 30 °C, and
although it grew slowly, it did grow continuously as monitored by OD
measurements. So, I assume the protein to be expressed is non-toxic. I
lysed the cells completely. I purified some lysate using Ni-NTA
magnetic agarose beads, which should highly enrich the His-tagged
protein. I ran several SDS-PAGE gels with purified and unpurified
lysates (with non-reducing and reducing sample buffer). The
silver-stained gel did not show any band with significantly increased
intensity (but then again there were many other bands of course that
may obscure this).
Two other gels were immunoblotted and a) incubated with an antibody
against a peptide segment of the protein to be expressed as well as b)
incubated with an anti-His-tag antibody. Western blot a) showed faint
bands of the appropriate size whereas Western blot b) showed
absolutely no bands for the purified lysates and bands of the wrong
size for the unpurified lysates (too low MW).
This is very confusing and I am not sure where the problem could be.
The expression vector seems to be OK, but I cannot detect a His-tagged
protein of the expected size using anti-His-tag antibody.
Does anyone have any idea where the problem could be or can you
recommend a good troubleshooting guide for prokaryotic protein