Are you boiling your samples prior to SDS-PAGE? Many membrane proteins aggregate and this can result in not being able to observe them on SDS-PAGE. Loading your samples without boiling can often improve results as the SDS in the loading buffer is quite efficient at keeping the protein soluble and providing a net charge.

After the ultracentrifugation step, what are your solubilization conditions?

@ Peters: Thanks for your reply. I perform one wash with TBS-Na2CO3 followed by a second ultracentrifugation, a wash in TBS, a third ultracentrifugation and final resuspension in TBS.

The membrane pellet from the ultracentrifugation step won't dissolve in TBS, you need some detergent. I used the "Popular Detergent Kit" from Anatrace to see which detergent can get my membrane protein back into solution, another round of ultracentrifugation and then ran the supernatants on a regular 6% polyacrylamide gel.

Can you not try TAP-tagging for your protein complex of interest?

In addition to Jim White's comments, we have had to run some membrane proteins in 8M urea to get them to migrate normally on SDS-PAGE (loading buffer and the actual gel). This comes after the purification/capture step so you can keep nondenaturing conditions throughout preparation. Interestingly for us, this was much more an issue with recombinant protein from coli than native protein from eukaryotic cells, so expression level and possibly composition of/in bacterial membranes could impact what you see on a gel. Membrane proteins frequently behave differently in SDS than cytosolic proteins.

You have to use detergent to isolate the membrane protein, buffer alone it doesn't work. If you haven't screen the suitable detergent for your target yet, you should first screen few detergents.

There could be several reasons can you give brief description about the steps, buffer and method. So that it will be easy to see which component is possibly contributing to such result

The attached protocol might help. :)

Purification of putative membrane protein complexes (in order to study possible protein-protein interactions) is even more difficult than purification of integral membrane proteins with low abundance. You must get rid off all soluble proteins and disrupt the membrane structure in a very special way; only as much as would separate the putative membrane protein complexes from each other and/or from the membrane proteins that are not belonging to any complex. I know only one detergent that have already been used successfully in separation of large membrane complexes; it is digitonin. After washing the hopefully 10,000g-100,000g pellet with 0.005 % (w/v) TX-100 in TBS (in order to open the sealed membrane vesicles and to remove the soluble proteins from the lumen of membrane vesicles), you have to establish the proper digitonin concentration fpr the solubilization of "membrane domains" that might contain your putative membrane protein complexes. Never forget include glycerol in your buffer if your work cannot be completed without freezing the samples (cryoprotection). However, how do you want to test what membrane proteins do belong to which membrane domains? First you have to establish a protocol to "separate" the putative membrane domains that contain those proteins which belong to a putative membrane protein complex. You have to think about a technique that is able to move "particles" with size of between about 200 kDa and 2,000 kDa (it is only my guess for the size). PAGE does not seem to me a proper technique. Some sedimentation technique should be applied. Then as soon as you have the separated "membrane domains" you can use PAGE tevhniques for analyzing the components.

In addition to Jim and David's comments I would also ensure you have effective lysis of the original coli cell pellet. Is your cytosolic fraction as expected on a gel - i.e. tons of protein? And how does that fraction compare to the supernatant after the 100,000g spin. Gels with all your fractions will confirm where you are losing protein.

my question may sound silly, but I do find it a little difficult trying to figure SDS-PAGE as the tool of analysis when you dont want to use 'denaturing methods' as you want to study protein-protein interaction. Maybe you are missing something in the description.

secondly, your is that of low yield and improper solubilization. A lot of learned answers have already ben posted here and you may notice that put together it probably answers everything. What you have told us here does not indicate low expression as you are not getting enough representation on your gel. I dont rule out low expression but you have to get a decent gel first. The first thing is to ensure a high percentage of cell breakage. washing with TBS certainly wont help you recover protein complexes. you need to solubilize the membrane with a gentle (nonionic?) detergent like triton or a glucoside buffer. SDS is a highly denaturing protocol and i wonder why you are using the protocol that you are.

with all humility, I would also suggest doing some reading before you launch into your experiment. There are several excellent protocols for isolation of membrane protein complexes from E. coli. Go through them. Bigger things will come later. if you detail your procedure maybe youll get better and more targeted responses.

This is how I do it:

E. coli cells are separated into periplasm, cytoplasm, and membrane fractions using the previously described lysozyme/cold osmotic shock method (Randall and Hardy, 1986). Briefly, the cells from a 10 ml culture of E. coli are pelleted down by centrifugation (4000 x g) before being resuspended in 0.5 ml of pre-chilled buffer 1 (100 mM tris-acetate pH 8.2, 500 mM sucrose and 5 mM EDTA). 0.5 ml of H2O and 20 µl of a 4 mg/ml stock of lysozyme is added simultaneously to the cell suspension. Samples are incubated in an ice bucket for 5 to 10 minutes. The resulting spheroplasts are treated with 20 µl of 1.0 M MgSO4 and pelleted by centrifugation. The supernatant (the periplasmic fraction) is collected. Spheroplasts are resuspended in 1 ml of buffer 3 (50 mM tris-acetate pH 8.2, and 2.5 mM EDTA) that had been pre-cooled, before briefly sonicating (30 seconds at ~12 microns) to break the plasma membranes. The membrane and cytoplasmic compartments are then separated by ultracentrifugation at 250,000 g for 30 minutes (using a Beckman TL100.3 rotor that had been pre-cooled to 4 °C). In not sure if the membranes will pellet at 100,000 x g but they certainly do at 250,000 x g. The supernatant (cytoplasmic fraction) is collected and the membrane pellets are solubilised in 1.0 ml buffer 3 containing a detergent (for example 2 % Dodecyl maltoside). For the solubilistaion the pellet is resuspended in buffer using a needle and syringe and the sample aggitated for a few hours. You can clean up the membrane sample with a further spin at 50, 000 x g. This time the solubilised membranes will be in the supernatant. You can then run samples from each cell fraction to check whether your protein of interest is inserted into the membrane. You can scale up accordingly for purification.

If you want to use native PAGE to study large membrane protein complexes you could consider Blue-Native PAGE that was developed for exactly this purpose.

Thank you all for your valuable replies. Sorry if my approach was not clear, I will summarize it briefly so that some of your questions could be answered.

I am analyzing several protein complexes and to do so I have cloned some genes which are thought to interact with these complexes. I overexpress these proteins and produce His-tagged recombinant proteins for later purification. I pellet cells, resuspend them in TBS, add lysozyme, PMSF and protease inhibitor and sonicate them. As I am also interested in protein localization (not all proteins are expected to be 100% membrane located) I perform cell fractionation, following the steps above mentioned, and analyze total cell extract, sonicated cell extract, cytosolic fraction and membrane fraction in SDS-PAGE to make sure I have good separation of fractions before continuing with purification. All observed patterns make sense but the one corresponding to the membrane, where I hardly see any band. As I said, I think I lose my membranes during the fractionation steps or I just cannot get good solubilization in the final membrane preparation.

I now reply your comments:

@ Peters: I will consider using this kit you mentioned to find out which detergent I could use, thanks for your advice.

@ Qureshi: the proteins I am trying to purify are His-tagged and purification has worked fine with cytosolic proteins.

@ White: thanks, I will also try preparing my samples following your instructions.

@ Horita: I am performing SDS-PAGE to test if fractionation works fine and see if I am actually obtaining membrane proteins in my membrane fraction preparation, not to analyze purificated samples. I will anyway consider your advice if I have problems when migrating purified membrane proteins, thank you.

@ Debela: Indeed, it seems I must somehow improve my final preparation for the membrane fraction.

@ Mastrogiacomo: thank you very much, your protocol differs pretty much from the one I plan to use but I may pick up some idea from yours.

@ Craig: I check expression in total cell extracts and I obtain a positive signal corresponding to my protein, so expression is not a problem in this case. Also, I break my cells by sonication and pellet (15000 x g, 20 min) cell debris before I proceed with fractionation steps. Thanks for the attached protocols.

@ Bérczi: I will read about this digitonin, I have never heard of it before, thank you. In principle I do not need to separate different membrane domains as I will focus on cytosolic vs membrane localization. As replied to Horita, I am using PAGE to test that I am actually obtaining membrane proteins in my final membrane fraction preparation. I will consider sedimentation approaches though, thanks.

@ Kerr: I am actually quite sure lysis is completed as I see no difference between the band patterns obtained from total cell extract and sonicated cell extract (where cell debris have already been removed). I do see a difference (some bands missing) between the later and cytosolic fraction, difference corresponding to membrane proteins I should (and I do not) see in the membrane fraction. All in all, I think I lose the membranes while fractionation or I do not have a suitable protein preparation.

@ Sharma: as stated above I am using SDS-PAGE to control I am performing good cell fractionation, not to analyze purified proteins yet. I hope I made this point clear in the description I wrote above. Also, I obtain good WB signals to control expression of my proteins and as replied to Kerr, I am pretty sure of complete cell lysis. Indeed, I must improve my membrane fraction preparation including some gentle detergent.

@ Barnett: I have evidence of membranes pelleting at 150000 x g in 1 h, for reasons related to infrastructure I cannot go faster than 100000 x g, so I have increased the time to 2 h. As you said, it maybe be a problem membrane pelleting ... I appreciate your tips for solubilization, I could include some of your advices in my next experiments.

@ all of you: thanks again for your suggestions, I will let you know whether I obtain better results.

Sorry to say but now, after having read your "working program and activity" I am even more confused about your goals than I had been before writing my first comments. I am just not able to see what protein-protein interactions (or interaction between membrane and some proteins, or membrane protein interaction) you want to study. The precise formulation of a problem might help to receive useful suggestions.

Only one comment in general. The commonly used nonionic detergents (like TX-100, OG, DM, etc.) solubilize biological membranes in a concentration-dependent way. Solubilization depends on both the detergent concentraion and the detergent-to-protein ratio on weight base (w/w). One must follow different solubilization protocols depending on the aim of solubilization. And solubilization is only one step in the series of steps in studying protein-protein interactions where ony partner is a membrane protein. The strategy is completely differen,t if both partners are membrane proteins.

@ Bérczi: I want to purifiy my cloned membrane proteins together with their putative interacting partners. However, I also want to determine the localization of my proteins, as some are suspected to be present in the cytosol too. So I want to purify my proteins and their putative interacting partners both in cytosolic and membrane fractions. The problem I am facing is that I do not obtain a good membrane fraction, I lose membranes during the fractionation steps or I do not solubilize them properly before analyzing the fraction by SDS-PAGE. So I am still in the pre-purification stage of my experiment. I hope I made myself clear now.

Thanks for your tips about detergents, I appreaciate it.

Are you boiling your samples prior to SDS-PAGE? Many membrane proteins aggregate and this can result in not being able to observe them on SDS-PAGE. Loading your samples without boiling can often improve results as the SDS in the loading buffer is quite efficient at keeping the protein soluble and providing a net charge.

@ Craig: I am not analyzing if my protein is in the membranes or not yet, but checking if I actually have any membrane protein in my preparation. Once I would confirm cell fractionation is OK I would proceed with the detection of my protein by WB in the membrane fraction and subsequent purification.

@ McDevitt: I do boil samples prior to SDS-PAGE, I will take your advice into consideration when I charge next samples, thank you.

I work with membrane proteins and this article is really helpful

https://www.mcdb.ucla.edu/Research/Jacobsen/LabWebSite/PDFOthers/GESeminar.pdf

1] try a bunch of screens for solubilization with different detergents...Mitegen/hampton etc

2] perform a quick blot on those screens to check if there is any of your target protein in membrane fraction

3] scale-up with detergent / buffer where your see the most yield / signal

one thing to remember is the yields are always marginal in these type of preps and not as good as purifying a soluble his tagged protein.

good luck

@ all: I have improved membrane vs cytoplasm separation by increasing ultracentrifugation speed up to 150000 x g. I obtain reasonably good protein yields in both fractions, with no noticeable cross-contamination between them.

Thank you all for your advice, I really appreciate it!

I suggest sucrose gradients with ultracentrifugation after breakage of cells with a ball bearing homogenizer, or just dounce homogenization. For bacteria, you may need a French press, but this is pretty harsh. Freeze thaws might be best. The cytoplasm is removed by the first spin, 100000 x g to pellet all the membranes. The cells are broken in 0.25M sucrose, with 25mM HEPES, and a little magnesium. The pellet from the first centrifugation is resuspended in the 0.25m sucrose buffer, and then layered on a sucrose gradient for further purification. a caveat is that I have never done this with bacteria, just cells. But sucrose gradients and breaking cells in sucrose buffer, works for any type of membrane preps, such as for cytochrome P450 from tissues, etc. So I think this will help for uniformity. If you break the cells too harshly, I would worry about the membranes being sheared too much, and not staying intact.

Just saw your note above. I am glad you got it to work.