Missle

I expressed an 18kD protein in e.coli with a 6x His on the N-terminus.  It purified fine w/IMAC but I did need to reduce the imidazole in the sample and binding buffer a little from my standard protocol or a lot of it went in the flow thru.  I then tried to run a western blot to confirm if an aggregate band was a contaminant or an aggregated form of my protein and used an anti-His tagged antibody that I've used many times before.  My monomeric protein isn't really being detected even at ridiculous loading amounts (I titered from 5ug to 50ng) and the positive control is fine.  I know there is plenty of protein there and enough histidine residues that it was able to bind to the nickel resin so I'm not understanding why the antibody can't detect it.  Any ideas?

I was also thinking of denaturing a sample with a strong denaturant and seeing if that would allow for antibody binding but I don't have much experience with this and wasn't sure if it was a good idea.

Any suggestions or recommended reading would be much appreciated.

Thanks!

I've been trying to express a couple of different proteins using a T7-based expression vector in E.coli.  They both are predicted to be good candidates for bacterial expression (size, RaCC, hydrophobicity, etc...) I've looked (in parallel) at BL21 (DE3), Rosetta 2 (DE3), and BL21 (DE3) pLysS.  I have 3 samples I'm analyzing: non-induced, induced w/0.05mM IPTG, and induced w/ 0.5mM IPTG.  I grow at 37C until an OD600 of 0.5 to 1.0 and then induce overnight at 18C.  I haven't acheived good expression for either target but what is bothering me is the trend I'm seeing with the three cell lines.  For both proteins, the OD600 at harvest shows very little reduction between induced and non-induced in the BL21 & pLysS strains but good 50% + reduction with the Rosetta which should indicate that the Rosetta strain is expressing my protein (shouldn't it?) but if it is I can't tell.  I don't see it in the insoluble or soluble fraction.  At first I thought it was toxicity but if that were the case, shouldn't BL21 look more like Rosetta interms of OD600 reduction?

I appreciate any ideas!

I am resuspending bacterial pellets (from 25ml cultures) after they've been frozen at -80C.  Sometimes I have up to 40 samples and it takes a long time to get each of them thoroughly resuspended.  I'm resuspending in 1.2ml with a P1000.  Does any one have any tricks/suggestions of how to do it faster?

I've got an odd problem.  I'm evaluating multiple lysis methods (sonication, Lysozyme w/Benzonase, SoluLyse, B-Per, and Bug Buster) on induced and non-induced BL21 (DE3) cells.  The soluble lysates were withdrawn without disturbing the cell pellets after centrifugation at 14,000 x g for 20 minutes and stored at -80C overnight.  With the B-Per and Bug Buster samples (which both lysed very well) when the samples were thawed and warmed to RT, a sizeable loose pellet formed at the bottom of the tube.  I ran SDS-PAGE on it's sup and resuspended and purified the resuspended material and am confident it doesn't contain my protein of interest - and the pellet formed in both induced and non-induced samples.  I centrifuged the 'soluble' lysate again (14,000 x g for 5 min) and the diffuse pellet became very small and smeared to the side of the tube.  I withdrew the lysate and attempted to resuspend the pellets both in TBS and in SDS Sample buffer and it won't resuspend.  The pellet dislodges from the side of the tube but stays flat and breaks into flat flakes when mixing.  I've never seen a pellet that looks like these.  Can anyone tell me what this pellet could be?

Does anyone have any creative ideas/ways of pipetting 100% glycerol for the purposes of creating bacterial stocks?  Using a P1000 pipet with a large 1000ul tip takes forever when adding my 100ul glycerol to each cryovial and I can't think of a faster way.  Any ideas?
Thanks!