Doing the sonication in water instead of PBS may help. You can add the PBS afterwards as a concentrate.

The result of sonicating DMPC in aqueous medium is a solution of liposomes. The clarity of the solution improves as the size of the liposomes decreases, which is what sonication achieves. The picture you showed is essentially clear considering that it is fairly concentrated at 5-10 mg/ml DMPC. It may not be possible or necessary to improve on this very much.

Doing the sonication in water instead of PBS may help. You can add the PBS afterwards as a concentrate.

The result of sonicating DMPC in aqueous medium is a solution of liposomes. The clarity of the solution improves as the size of the liposomes decreases, which is what sonication achieves. The picture you showed is essentially clear considering that it is fairly concentrated at 5-10 mg/ml DMPC. It may not be possible or necessary to improve on this very much.

I agree with Adam Shapiro., it looks like you have a nice suspension of small unilamellar vesicles (SUV), which is what you obtain by sonicating o solution of phospholipids. Adding organic solvent you may obtain a clearer solution because you are disrupting the vesicles and solubilizing the lipids. This may be fine or not depending on what you want to achieve: many vesicles forming molecules (and definitely phospholipids) change their structure and their mode of interaction depending if they are solubilized or structured in vesicles.

OK thanks. The goal is to bind DMPC to ApoE, my assumption is that ApoE directs the mode of interaction, i.e. I am not trying to form some particular type of liposome, so as long as the lipid is in some form that can be bound by ApoE I would think that sufficient. The assay is to look at binding to an ApoE receptor, TREM2, which is occurs only when ApoE is properly lipidated.

Unfortunately, this latest solution, when mixed with ApoE, also failed to confer binding ability on ApoE (other species of proteins that are already lipidated did bind).

I wonder if there is a simple biophysical assay that can tell you when your lipid has bound to a protein. I could use biotinylated or fluorescent derivatives etc. but is there a simple assay to tell you a lipid has bound to something like ApoE?

It depends where you come from and what you consider simple. If you are a mass spectrometrist this is a very simple experiment, but since you are asking I guess you are not ;-)

Alternatively, capillary electrophoresis or RP-HPLC are much easier but very effective techniques for something like what you want to do. Once you optimized your run conditions, which might require some work.

Good suggestion. I'm a lowly molecular biologist; but I have colleagues who do mass spectrometry. I guess we could run protein alone, lipid alone and then see what the mixture looks like. We don't have an actual positive control here. But might not be too hard, esp. since I would not be doing the actual work.

If you can separate your protein from any unbound lipid, you can extract it with organic solvent, run the extract on a silica thin-layer chromatography plate, and stain it for lipids. I don't have a protocol, but I'm sure you can find one in the literature. You can run lipid standards side by side to confirm the identity of the lipid.

Another way would be to include some radiolabeled lipids, and measure the radioactivity associated with the protein after removing unbound lipid.

Phosphatidylcholine (as in DMPC) can be assayed in a sample using one of the commercially availble PC assay kits.

It may not be possible to transfer DMPC from liposomes to apoE. You might have to mix the apoE with an water-miscible organic solution of the lipid, or a detergent solution. Another possible method would be to prepare a film of the lipid on the surface of a container (by evaporating the solvent), adding the protein (and some detergent?) and mixing for a while. Elevating the temperature above the phase transition temperature of DMPC may help.

If you want to dissolve the lipid in detergent, starting from the liposome suspension is convenient.

Thanks

Apparently the lipidation of ApoE is a pretty standard method and has been done routinely since the 1980s. So I don't imagine I need to change the protocol for lipidation. But I will look into staining for lipids extracted off the protein, or most simply give the sample to a colleague to run on LC/MS.

Attached is an example of the lipidation protocol from 1978. Seems simple enough....(?)

The standard protocol to make liposomes is to dissolve the lipid at 10 in chloroform or cyclohexane and swirl it in a round flask until the solvent has evaporated, leaving a thin lipid film on the glass wall. Then add buffer and sonicate (bath sonicator should suffice, but tip sonicators are more effective. You are using saturated lipid (C14:0), but still it is a good idea to perform all operations under a N2-protective atmosphere (with non-saturated lipids it would be imperative to prevent lipoperoxide formation).

For protein reconstitution into liposomes see

@ARTICLE{Rig-98,

title = {Membrane proteins: functional and structural studies using reconstituted proteoliposomes and 2-{D} crystals},

author = {J.L. Rigaud},

journal = {Braz. J. Med. Biol. Res.},

year = {2002},

pages = {753-766},

volume = {35},

doi = {10.1590/S0100-879X2002000700001},

language = {engl},

url = {http://www.scielo.br/pdf/bjmbr/v35n7/4512.pdf},

}