Western blot trouble shooting - (Aug/28/2012 )

Hello

I am expressing a plant protein in E.Coli. I found out that my protein is expressing as inclusion bodies rather than being a soluble protein. My protein is His-tagged. I want to do a nice western blot for my protein. I purified my protein using the denaturing buffers in the Qiagen Ni-NTA Fast start handbook. As control I have inclusion bodies from the same bacteria but without IPTG. THe inclusion bodies from the control was resuspended in buffer containing Tris and sucrose only. I used 5 micro litre of my samples and it has a distinct band of moderate size not a fat band. Now I use the TMB substrate kit for peroxidase from the vector laboratories for western blot. I use PVDF membrane for western blotting.

Now in my membrane just when it starts staining I can see distinct bands in the front (the side that was actually in contact of my gel). Just after seeing those bands I take out my membrane from the staining but when it dries the lanes form a blue color that spreads uniformly from top till the place where my protein is supposed to be, so more than half of the membrane.

But in the back of the membrane I have nice distinct bands at the places where I saw them in the front at the start of my staining.

My control sample lane remains absolutely white and the the portion of my denatured sample lanes below my protein remains white too.

I use skimmed milk for blocking my membrane overnight at 4 degrees. I use 1:6000 dilution (.17 micro grams) for my primary antibody and 1:10000 dilution for my secondary antibody. I wash 6 times 5 minute each with TBST in the washing steps after incubation with the antibodies.

I think its not a problem of high background as otherwise I should get blue color across my controls as well as in my ladder lane too. I also think its not due to overloading as I have a moderate size band not like a thick band.

Any suggestions that can help improve my western blot is highly appreciated.

Thanks.

Bioscientist

It looks to me that you are transferring too long. Smaller proteins get transferred faster than the large ones, so you might need to decrease your blotting times in case you have such a protein. To check for this you can use 2 membranes on top of each other and check both with the antibody. If you get your fat band on the 2nd membrane, then I was right.

In addition, it seems that you are describing that you have a smear of your protein in the lanes you are loading your samples. You might have overloading. I have that behavior when I have good purification yields. You can see this in the sample before loading if you have either a more viscous than usual sample or even a precipitate of proteins in the sample after boiling it in the loading buffer. Try to dilute your protein before you add the SDS-loading dye. Alternatively, I have this smearing also in the fractions containing aggregates (after gel filtration the first samples after the void volume contain aggregates, so you can be sure that after Ni-NTA you have a mixture of aggregates and properly folded proteins). Actually, I always purify using a gel-filtration as polishing even though I have only my protein in the elution of the Ni beads. In the end: you are refolding your protein, you are bound to have more than average aggregates. If you do not have a gel filtration column and you want to avoid this, you can remove aggregates (not perfectly) by spinning your sample at the max speed of a table-top centrifuge or by syringe filtering it with a 0.2 um filter.

Andreea

Thanks Andreea.

My protein is about 29 KDa and I transfer for 1 hour at 25 volts. I can reduce it to 30 minutes and see what happens.

And my samples doesn't look viscous compared to the control. I will decrease my loading amount of sample.

And I was also wondering if I cut the gel and extract my protein from the gel and use that to run western blot, do you think it can reduce the smearing effect?

Thanks.

With the cutting of the band: it is like cheating by omission with the purpose of beatifying your WB; do not do it.

29 kDa is a bit small, even though 1 h should be still ok... either try a bit less or the double membrane strategy I was telling you about: at least like this you know where the protein goes. Also stain the gel after transfer with coomassie. Like this you can follow where your protein went. If it is not on the membrane, then it is on the gel, if it is not on either, it went through.

With the loading of less sample: this is also ok, but what I mean about diluting your sample is that when you boil your sample with sample buffer and you have too many aggregates, the SDS/mercaptoethanol or DTT are not enough to completely denature your aggregates. If you dilute your sample, let's say 1 to 9 or 1 to 4 (before boiling it with loading buffer), then you also dilute the aggregates and you make it easier for the loading buffer to denaturate your aggregates.

Andreea