Membrane proteins are a weird bunch. We expressed one as a GST fusion and saw nothing under standard SDS-PAGE. We added a protease to cleave GST from the fusion protein (desperation) and suddenly saw a lot of GST, suggesting that the intact fusion protein was actually expressed but not visualized, likely smeared throughout the lane. We obtained much better visualization using gels with 6-8M urea in the matrix and loading buffer. For some membrane proteins, SDS is like the native state. If you have an antibody, you could try a Western (anti-His on lysed cells will be a nonspecific mess but you still might be able to see a difference between induced/non-induced).

Sequence and the reading frame is ok, right?

Maybe rare codons are the problem? BL21 might have some problems if the sequence is not optimal.

And other people also used BL21 strain?

In order to suggest you something, I would like to know other issues regarding your construction, have you cloned it with its own signal peptide? have you removed it? or have you replaced it for another signal peptide? Anyway, expressing membrane proteins is always a very tricky process, so my general suggestion is to follow protocols that has been proved previously. That will save you a lot of time and efforts.

Also, sometimes I have found very useful to use C41 or C43 strains (Lucigen) instead of BL21 for expression of membrane proteins, but again it depends on the protein.

Most likely the N terminal His tag is interfering with some crucial processing required to have it expressed at the membrane.

I have seen much more E. coli expressed membrane proteins with C-terminal His-tag than attached to the N-terminus. Does the original plasmid have a His-tag? If yes, what is the location?

Swati, a few questions before one can advise:

1. Does the protein also have an N-terminal signal sequence for export/membrane insertion?

2. If not, have you looked for inclusion bodies?

3. Is there a low level of expression but protein is in membrane fraction?

Membrane proteins are a weird bunch. We expressed one as a GST fusion and saw nothing under standard SDS-PAGE. We added a protease to cleave GST from the fusion protein (desperation) and suddenly saw a lot of GST, suggesting that the intact fusion protein was actually expressed but not visualized, likely smeared throughout the lane. We obtained much better visualization using gels with 6-8M urea in the matrix and loading buffer. For some membrane proteins, SDS is like the native state. If you have an antibody, you could try a Western (anti-His on lysed cells will be a nonspecific mess but you still might be able to see a difference between induced/non-induced).

As others asked, give some more information about your experiment. like what are the various factors you already checked? how big is your protein? how are you visualizing the expression of the protein etc.

Ditto above-Not enough information provided to make a good evaluation. Did you verify the DNA fragment by sequencing both directions and checked that the sequence is in correct reading frame? Are you expressing with or without the signal peptide? Did you store the transform E. coli prior to expression, how and for how long?

I would be worried if you're producing any mRNA. Your gene sequence may be correct but is your promoter intact? Following that logic, do you have a high enough induction strength?

Thanks everyone for the suggestions. The native protein does not contain a signal sequence, the 1st TM domain serves as stop transfer sequence. I have sequenced from both sides and its in frame (confirmed using EXPASY translate). The his-tag at N-terminus can be a problem. I have checked for the expression in inclusion body and can't detect it. My collaborators just informed me not to boil the sample before SDS. It seems it has negative effect on Membrane proteins. I will try this and see if it works.

I am using AP-conjugated his-ab for protein detection.

Hi Swati, have you tried to run the SDS-PAGE with the E. coli lysis? Did you try to stain the SDS-PAGE with silver? If your protein were there, it will appear!

Dear Swati, the high hydrophobicity of membrane proteins impair with the expression in bacterial system. In my personal experience, cloning of one of my protein of interest with similar properties troubled me a lot in expression. Later on deletion of N-terminal trans membrane domain and further subcloning into pET 28 vector having ribosome binding site made me smile to see excellent expression in soluble form with N-terminal 6X His tag. Now you may have to look into the hydrophobicity plot of your protein sequence first... all the best.

The fusion partner greatly influences expression levels as weak solubility of TM proteins is often problematic. I used different expression strains and a deep well screening assay to improve expression levels about thirty fold compared to BL21 standard conditions, changing expression detectable by Western Blot only to a prominent band in Coomassie stained gels. I attach the publication if you want to have a look. We have established excellent expression protocols also for other TM proteins in the lab as well as other non-TM protein, so its really widely applicable and should work in your case as well. Good luck !

Article Optimisation of expression and purification of the feline an...

When you say the gene sequence is right, do you mean you've also checked the promoter and other regulatory elements?

Also, there might be expression that you just can't see: are you trying to detect it thanks to its His-tag (i.e. anti-His antibody)? In that case it is possible that the His tag has been cleaved off so you might want to try an antibody against the protein itself if you have one available. Also are you trying to detect it in the soluble fraction after cell lysis? If so you might not be detecting it because it might be insoluble - in that case try whole cell extracts: boil cells directly in lots of loading buffer, then spin, then run the supernatant on a gel and do a western.

Hi

Anti hist antibodies can be problematic in WB due to lack of affinity or hist tag being blocked or lost by degradation etc. A positive control for the hist antibody would be good to use. I take it you cant see an induced band by coomassie staining.

I'm also assuming that you transformed the BL21 cells with plasmid for each expression experiment and did not take from a glycerol stock?

The use of protease inhibitor cocktails are also essential if you are lysing cells.

To increase sensitivity it may be possible to partially purify your tagged protein via IMAC?

A positive control for your induction would also be good: I take it you induce with IPTG? if so make sure this is prepared and frozen in aliquots and not freeze-thawed.

Working on you induction protocol may yield better results: looking at your time of induction/temperature/IPTG conn etc

good luck