Trouble detecting Histaq - (Jan/16/2012 )

Hi friends,


I want to do western blot for detecting 4 E. coli expressed proteins . previously i was not able to detect the proteins from induced cells. Then i purified the his taq proteins and was able to detect only the smallest band.

I was thinking the other proteins are high MW so did not transfer well - My transfer conditions are:

20% methanol - tris glycine buffer, 100V for 1.30 hrs - After transfer I could still see some high MW prestain marker bands on the gel. Primary antibody 1:100 for 3 hrs and secondary antibody for 1hr

I tried changing transfer buffer with reduced methanol to 12.5% and transfer time to 2hrs but the assembly got too hot - performed on ice - i could see vapours coming out - IS IT POSSIBLE THAT MY PROTEINS GOT DISTROYED BY HIGH TEMPERATURE

Also, I changed blocking time from 60 min to 30 min - fearing that my antigens could have been suppressed.


Right now I am thinking,
1. Could it be sample preparation - first time when i could see 23 mw protein the sample was 8 min heated but now i am using sample that is 15 min heated
2. The transfer assembly got to hot - and my proteins got distroyed
3. Changing methanol is not necessary for my proteins
4. Instead of using 2 hrs at 100V i can use overnight transfer
5. prolong the antibody incubation times

I would say that you probably only need to boil your samples for 5 minutes in denaturing loading dye.

52 kDa is not a big protein - it should transfer fine in less than an hour at 100 V. You may be "blowing through" the proteins when you are transferring, you could shorten the transfer to 1 hour or use a lower voltage and see how that works for you.

If you are using PVDF as a membrane, try adding 0.01% SDS to the transfer buffer.

Regards your antibodies - have other people in the lab used them with the same conditions you are using? If so did they work for the other people? You may need to titrate the antibody concentration and/or change the blocking solution (mik, BSA, serum...) and incubation times, but most antibodies should work under the conditions you mentioned. Denatured proteins are unlikely to be damaged by the transfer, after all you have just boiled them before loading.

Thank you bob1. My labmates use serum for blocking. As it was not giving results i changed the blocking buffer to milk

I will do this expt again with lower voltage.

Helo friends,

I change the transfer protocol a bit. I used gel equilibration (with transfer buffer) for 15min. After the transfer most of the protein marker upto 130kDa got transferred. However, my proteins still remain in the gel . I am using the samples directly form Nickel sepharose purification. My elution buffer has lysozyme. Even lysozyme is not transferred.

My elution buffer has Tris, NaCl, imidazole, glycerol, tween 20 - Is there any chance that imidazole or tween 20 might be responsible for preventing the protein transfer from gel ?

I checked the lysozyme pI it is also more than 9 - similar to my proteins - would it be better if I change the pH of my transfer buffer.

Still, I havent checked the inclusion of SDS in transfer buffer - will do that today

you may want to try a higher pH transfer buffer (eg caps pH 11). after the methanol in the transfer buffer strips the sds from the proteins the pI becomes more important. you won't be able to transfer a protein with a pI in excess of 9 in a buffer at pH 8.3. the addition of sds to the transfer buffer (it helps when transferring large proteins) may help a little but it will still be stripped by the methanol.