The comments about detergents are completely on target. While detergents are necessary to extract and maintain the protein outside of the lipid environment, they are also denaturants. This is due, in part to the fact that they form structures with an extremely high radius of curvature compared relatively flat (I use this loosely) native membranes, and unlike membranes they are really 3D structures not 2D. This means that with time they will tend to "solubilize" individual helices denaturing the proteins. Depending upon your down stream application (this won't work for crystallography) you can extract the protein then reconstitute it into nanodiscs (Steve Sligars work is a good example, although other groups have been coming up with alternate ways to make these). For this you need to fine 1 detergent that is good for solubilizing the protein out of the native membrane (many of the good detergents for extracting from a membrane are poor for long-term stability) and another that you can transfer into to allow reconstitution into nanodiscs. This latter detergent either has a low CMC (removed by dialysis) or, can be extracted with biobeads. I prefer the latter since reconstitution into nanodiscs by dialysis in our hands is less efficient. In principal if you are not going to do much purification upon extraction you could use the extracting detergent. Sligar's group have reconstituted whole membrane extracts into nanodiscs and then pulled out their tagged protein by affinity chromatography. We have not tried that ourselves, but some of my colleagues have had less then stellar success with this approach. On that note, there were several presentations at this years biophysics meeting that used alternate nanodisc strategies to achieve this goal.

Purified membrane protein nano-disc preparations are remarkably stable, allow the protein to be maintained at high concentration (Wagner's group uses the for NMR studies and others have used them for EM one of the Harvard EM groups), in our hands are typically more active then detergent preparations. This may reflect both the presence of native lipids, which can affect activity, but also its widely recognized that detergents tend to affect the helical packing often promoting the an in active conformation (see the crystallographic literature). Now if you are trying to make the protein for crystallography many people have tried nanodics but I do not know of any successful structures determined in this manner.

Adding (or changing) a surfactant to ged rid of the leftover membrane lipids may help. Different membrane/protein combinations require different surfactants to get solubilized. First try some that are usually around in the lab like TritonX100 or Tween before you order some more expensive ones.

As you did not provide any details on your protein/purpose of the purification, I would suggest you read this review on the solubilization of membrane proteins by Michael I. Schimerlik published in Current protocols in Neuroscience.

See attachment, it has a pdf file.

Good luck.

I had an identical problem when I tried isolating the main protein component of lens fiber cell membranes in my 20 year study of those membranes. It was very frustrating to lose the proteins to protein aggregates, clumps, and precipitates. Adjusting the pH did nothing to prevent the aggregation, and indeed both higher and lower pH's caused irreversible aggregations. I finally thought about it and decided that the proteins were too homophobic and when deprived of the protein-phospholipid interaction which normally kept them from aggregating was the real problem. I then experimented with the use of very small concentrations of detergents which I thought would form micelles including the proteins with them, the detergents replacing the phospholipids in providing a safe environment for the proteins and preventing their aggregation. It worked. But the only caveat is that a variety of detergents have to be tried depending on the size, nature, and homophobicity (a word invented by me) of the particular protein in question. I finally settled on SDS at a reasonable concentration, the game is to start at the higher concentrations of the detergent (say 5%) and work one's way down to the smallest concentration at which the aggregations are minimal (say from 0.5% to no more than 2.5%). The problem is that at the higher detergent concentrations (between 3% and 5%), the action of the detergent actually denatures the proteins too much for the proteins to be useful in whatever tests you desire to perform with them. I would peruse the literature on the different detergents available as SDS is a particularly strong one, but then I was dealing with an extremely homophobic intrinsic membrane protein. I hope this explanation helps you, and I would like to serve as a knowledgeable supervisor of your progress if you so wish. The saying is that an expert is the guy that can tell you what doesn't work, but in me you will find an expert that can tell you what does work. Good luck . . . although if you follow my advise you won't need it (luck that is) . . .

The protein is bacterial

Maby you can obtain your protein as a fusion protein with MBP or NusA which increase the solubility. If not then detergents:)

What is the detergent you are using for solubilization and purification?

I think tagging a protein with MBP is a better option to enhance the sollubility of protein. Second, you may have to try some different detergents with varying concentration during the sollubilzation process. In this way you can sort out a optimize the detergent for purification.

I would try adding some salt e.g., ammonium salts and in certain cases addition of amino acid, e.g., proline have some beneficial effects. Both are documented in literature with success, but you have to try to see if it works in your case.

Try CHAPS to solublize the membrane protein and prevention of aggregation during purification.

im already using Chaps 20x cmc

Hi Joe,

Long ago when I purified sercoendoplasmic reticular Ca-ATPase from bovine pulmonary artery I studied extensively different proteo-lipid compositions by selectively solubilizing the lipid using an array of detergents like Triton-X100, C12E8 and DHPC. the purified enzyme retained maximum activity without aggregation when it had been reconstituted in proteo-liposomes prepared by DHPC-DOPC.

I agree with all the contributors in this thread that each protein's environment should be customized by not only selecting proper detergent but also by selecting proper ratio of lipid-detergent to prevent aggregation for retaining maximum activity.

I published this data in 2006 Biochim Biophys Acta paper which can be freely downloaded from my RG page.

Good luck!!!

Got to mention if it is integral membrane protein or peripheral membrane protein. I assume you are talking about Integral membrane protein. In this case keep following in mind:

1) Use High salt concentration (may be 500mM NaCl or more)

2) It may be Glycerol sensitive (try with and without 5% glycerol and check ).

3) you may try adding fusion partner but I never have any good luck with it so far.

4) One thing you can do is that you can do a sequence alignment with the homologues and find some unconserved region at either N or C terminal. truncate those residues and see if it changes anything. You may also play with the disorder loops in between.

Have you tried any strong detergents like sarkosyl or SDS?

Joe, overnight induction at 20 C helps to reduce the aggregation of recombinant proteins. If you are not worried about the activity of the protein, addition of urea (2M-8M) keeps the protein in solution during purification and refolding by dialysis in buffers with sequential reduction of urea is a good method to retrieve the activity of your protein.

Try with surfactant such as chaps or triton x-100 for solubility of membrane protein. these detergents also prevents aggregation of membrane protein. You also try with buffer at different pH, acidic, neutral or basic. some time ions, salt or detergent also involved in aggregation of protein

Dear Joe

I think that the triton x-100 will go help you. Regrads

You might want to try adding phospholipids to your buffers during your purification procedure in case aggregation occurs during purification. Reducing the expression level by growing cells at lower temperature, shorter induction time, less inductor might also help.

I use triton x-100 (from 1% to 4%) during purification, and 10% glycerol in the elution buffer.

I would try detergents like dodecyl-maltopyranoside (DDM), LDAO, and Fos-Choline.

Use these at a concentration of ~ 3 times their CMC for the purification.

For solubilization, I would use 1% of DDM for example.

Your description is very short and your question is very unspecific. Thus my answer can be as well very "unspecific". When I do purification of integral membrane proteins I try

- to keep the osmotic and ionic strength condition of the homogenization buffer as close to the in vivo conditions as possible,

- to have appropriate buffering capacity of solutions,

- to have reducing conditions to keep -SH groups reduced,

- to do solubilization of membrane proteins at protein concentration of about 1 mg/ml, the detergent-to-protein ratio between 2:1 and 5:1 (w/w) (of course, you must pay attention to the c.m.c. as well), the temperature between 5 C and 10 C, and for at least 120 minutes,

- to have some cryo-protective agent in buffers after solubilization.

This bring back fond memories...

I would suggest 500 mM NaCl, 20 mM Tris-Cl (we used pH 7.9), 0.02% n-Dodecyl-beta-D-Maltoside, 10% glycerol. We did His-tag purification so we also added imidazole.

Check out:

http://www.jbc.org/content/272/3/1761.full

Yoram

Assuming it is an integral membrane protein the one you are interested in, do you start the purification from native membranes? Playing with pre-extraction conditions may help you having a better starting material. A Key feature is if you need to keep protein activity or not. If not, as mentioned, anionic detergents such as sodium dodecyl sulfate (SDS) or high concentrations of chaotropic agents (Urea) should help you avoiding precipitation.

My experience is with membrane protein complexes in which activity was needed to be preserved. Here you have some tips:

-Dissolve the membrane protein under optimal conditions for stability at a detergent/protein ration that is not much above minimal detergent/protein ratio required for protein solubilization (this specially applies for membrane protein complexes).

-Perform separations in cold and try to keep isolation time as short as possible.

-Keeping the appropriate pH is necessary for protein stability and solubility. Phosphate and Tris buffers are commonly used.

-Triton X-100 and B-Docecyl Maltoside are very much used mild detergents when activity needs to be preserved. First one is much cheaper and that may be important if you need to scale up.

-Ionic strength is another factor affecting the solubility of membrane proteins in presence of detergents. Since it is a rather freely adjustable parameter, you could try 500 mM NaCl, and then go higher if needed.

One of the surfactants above should work to keep it solubilized during purification, however, without much details of your purification process it is hard to determine what else could be done. For example, you may have to worry about how to get rid of some of the surfactant you are adding - which will be different for example between salting out vs. chromatography. Finally, you may have to pick a different surfactant after purification to maintain structure and activity of the purified protein. One of the suggestion above - to use lauryl maltoside or other similar surfactants - can do wonder, in my experience. All the best.

Add some H2O2 (10%) in the presence of the MnO2 (catalysator) in the cleaning phase. ;-)

Renato Babic; WHY do you think your suggestion would help membrane protein aggregation? WHY would O2 generation prevent something that might have origin in (be governed by) hydrophobic interaction? I would highly appreciate, if you explained (provided details) to your recommendation.

Dear Joe,

If your interest is to study kinetics and affinity of membrane proteins, you do not need to purify. In the label-free cell-based biosensor Attana Cell 200 you can grow cells directly on the sensor surface and characterize the interactions with the membrane proteins still intact in the cell.

Best regards

Teodor

I suggest to changing detergents (3x CMC) or combining detergent used for solubilization and additional detergents during wash and elution step of IMAC. It looks better to use less than 5x CMC is good choice of detergent concentration. There are many detergents. But it would be good idea to select some popular ones (Tx100, Brij35, DDM, CHAPS, OG, sodium cholate, DHPC, Facade). It would be also good to add lipids (0.1mg/ml POPE-it would not soluble but you can use after filter turbid solution for preparation of washing and elution buffer). I hope it helps your experiment.

Sung

The many people who have suggested trying different detergents are correct, but if you are getting inclusion bodies you might also try:

coexpression with chaperonins (e.g., using GROELS plasmid)

lowering the temperature (we often turn down E.coli from 37C to 17-25C at induction

slowing down the rate of synthesis (e.g., changing vectors to use a slower promoter)

if there are cofactors, making sure you supply cells with plenty of precursors.

You didn't specify what system you were expressing in, so we don't even now if your using mammalian cell culture, yeast or bacteria.. You might need to change systems.

Hello,

Does this protein aggregate at certain purification step or is it impossible to purify it at all from cells?

There are lot of suggestions above how to extract it. I would try multiple detergents on small pellets (all the detergents you have in the lab - CHAPS, Tween, Tergitol, Triton, DDM, Cymal, sarcosyl, cholate, deoxycholate) and then run pellet and super after extraction on gel. By the nature of protein you can predict if non-ionic or anionic detergent would work best (total charge of protein an be determined by pH titration freewear).

If you do extract protein and it aggregates in the following steps of purification addition of detergents such as Tween around 0.1% should help. Also some small sugars such as lactose or trehalose stabilize purified proteins during concentrating, buffer exchange or ion-exchange polishing steps. The concentration is in the range of 1 mM but you can titrate using this concentration as starting point. Combination of detergent and monosaccharide is most beneficial.

Good luck,

Galina

Hi Joe,

A week has gone and you have got many interesting ideas and suggestions. However, you have to make your choice. To do so please try to answer (clarify) the following questions before you try any further experiments:

a) Is the membrane protein of your interest an intrinsic (integral) membrane protein or an extrinsic (membrane associated) protein?

b) After a short "in silico" study which would give you some idea about the primary structure and physico-chemical properties (pI, secondary structure elements, number and location of Cys residues, possible phosphorylation and glycosylation sites, etc.) of your protein in interest, make your decision of the pH and composition of buffer.

c) Depending on the size and possible physico-chemical nature of your protein in question, choose a detergent with high or low c.m.c. value, ionic or non-ionic nature. Pay also attention to the size of detergent micelles (the so-called aggregation number of detergent in micelles) which in itself might help you to decide if the detergent would be able to fulfill your request.

d) After solubilization (and after checking the solubilization yield and recovery), each solution (buffer) you apply must contain the detergent used in the solubilization at least twice the concentration of the c.m.c. of the detergent.

e) After solubilization, use also cryo-protective agent (glycerol, sucrose, mannitol, etc.) and appropriate ionic strength in all buffers with buffering capacity of 20-50 mM.

f) Keep your protein in deep-freezer.

If you are over expressing your membrane protein it may be making inclusion bodies that have not folded correctly. Try dialing down your expression (induction) then isolating the membrane protein with the detergent suggestions above. This will help to ensure you are isolating properly folded membrane protein.

try to use DTT or some mild detergent like tween. This will help

Use DTT or some mild detergent like Tween-20 in low concentrations

use 1 mM DTT or try mild detergent like TWeen-20 in low concentration.

I used SDS+ beta mercaptoethanol to extract membrane proteins then ran them on a gel. Showed very little aggrigation

The choice of detergents is quite critical for maintaining the structural integrity of membrane proteins. If you are purifying your protein of interest, you can analyse the protein by SEC to monitor the mono- or polydispersity of your protein. You can also couple your protein to GFP and test out different detergents, lipids, buffering agents, reducing agents, ionic strength, etc. to screen conditions for monodispersity (Kawate and Gouaux. Structure. 2006 Apr;14(4):673-81.).

The comments about detergents are completely on target. While detergents are necessary to extract and maintain the protein outside of the lipid environment, they are also denaturants. This is due, in part to the fact that they form structures with an extremely high radius of curvature compared relatively flat (I use this loosely) native membranes, and unlike membranes they are really 3D structures not 2D. This means that with time they will tend to "solubilize" individual helices denaturing the proteins. Depending upon your down stream application (this won't work for crystallography) you can extract the protein then reconstitute it into nanodiscs (Steve Sligars work is a good example, although other groups have been coming up with alternate ways to make these). For this you need to fine 1 detergent that is good for solubilizing the protein out of the native membrane (many of the good detergents for extracting from a membrane are poor for long-term stability) and another that you can transfer into to allow reconstitution into nanodiscs. This latter detergent either has a low CMC (removed by dialysis) or, can be extracted with biobeads. I prefer the latter since reconstitution into nanodiscs by dialysis in our hands is less efficient. In principal if you are not going to do much purification upon extraction you could use the extracting detergent. Sligar's group have reconstituted whole membrane extracts into nanodiscs and then pulled out their tagged protein by affinity chromatography. We have not tried that ourselves, but some of my colleagues have had less then stellar success with this approach. On that note, there were several presentations at this years biophysics meeting that used alternate nanodisc strategies to achieve this goal.

Purified membrane protein nano-disc preparations are remarkably stable, allow the protein to be maintained at high concentration (Wagner's group uses the for NMR studies and others have used them for EM one of the Harvard EM groups), in our hands are typically more active then detergent preparations. This may reflect both the presence of native lipids, which can affect activity, but also its widely recognized that detergents tend to affect the helical packing often promoting the an in active conformation (see the crystallographic literature). Now if you are trying to make the protein for crystallography many people have tried nanodics but I do not know of any successful structures determined in this manner.

Normally membrane protein aggregation can be prevented using detergents like LDAO and OG. These detergents are specially used for membrane proteins only and a concentration of even less than 0.1% would help the protein to remain in non-aggregated state... Do try it out once.... Wishing you all the best....

Joe S Matarlo ... Please check references below.. hope it will be helpful for you…

Regards…

http://wolfson.huji.ac.il/purification/PDF/Literature/Bondos2003.pdf

http://www.amr-inc.co.jp/dcms_media/other/%E3%80%90%E3%83%97%E3%83%AC%E3%82%BC%E3%83%B3%E8%B3%87%E6%96%99%E3%80%91120726%20JHUPO%20Symposium.pdf