Scenario:
--> 5 tubes, all with the same amount of dried-down lipid (e.g. 1 mg).
--> In the end, each 1 mg of lipid will be solubilized in the same final volume (e.g. 1 mL)
--> in the end, each 1 mg of lipid will be solubilized with the same concentration of the same detergent
--> The only difference between the tubes is the amount of protein being added (solublized in the same buffer with the same concentration of the same detergent as the dried-down lipids).
--> For each tube of dried-down lipid, a certain volume "A" of detergent solution is added, which varies depending on the amount of solubilized protein sample "B".
--> After the addition of buffer volume "A", followed by vortexing to solublize the dried-down lipids, then brief sonication of each vial, ALL 5 tubes still have the same turbidity.
--> The required volume of solublized protein sample "B" is added, yielding different protein:lipid ratios (i.e. same amount of lipid, but with varying total amounts of protein)
--> For ALL 5 tubes, "A" + "B" = 1 mL
--> As the amount of protein relative to the amount of lipid increases, the preparations become increasingly LESS turbid, i.e. they "clear up" (but the same amount of lipid and detergent should be present in each tube).
What could be the reason for this observation? I have done quite a bit of asking around of grad students, post docs, and professors, and none have been able to provide a conclusive answer.
Just speculating:
If your lipid+detergent "solution" is turbid, this means that you have two phases present, i.e. your lipid is not fully in solution (as it would be in e.g. chloroform), but more like an emulsion.
Some proteins (e.g. albumin and apolipoproteins) bind lipids very efficiently.
My guess would be the following: Your protein binds lipids in a molecularly defined fashion (i.e. have one or more lipid binding sites). The forming protein-lipid complexes are really soluble - and not an emulsion. And a "real" solution is not turbid.
Adding more and more protein removes more lipid from the lipid phase of your emulsion, thereby diminishing the amount of lipid phase present and thereby also the turbidity.
This hypothesis should be verifiable using Isothermal Titration Calorimetry (ITC) (http://www.microcal.com/technology/itc.asp).
Just speculating:
If your lipid+detergent "solution" is turbid, this means that you have two phases present, i.e. your lipid is not fully in solution (as it would be in e.g. chloroform), but more like an emulsion.
Some proteins (e.g. albumin and apolipoproteins) bind lipids very efficiently.
My guess would be the following: Your protein binds lipids in a molecularly defined fashion (i.e. have one or more lipid binding sites). The forming protein-lipid complexes are really soluble - and not an emulsion. And a "real" solution is not turbid.
Adding more and more protein removes more lipid from the lipid phase of your emulsion, thereby diminishing the amount of lipid phase present and thereby also the turbidity.
This hypothesis should be verifiable using Isothermal Titration Calorimetry (ITC) (http://www.microcal.com/technology/itc.asp).
Interesting observation; I can only speculate. The increase in turbidity could be an indication that the size of you liposomes increase if you add more solubilised protein (the bigger the particles are, the more light they will scatter, which makes your suspension turbid) After sonication, you should have only very small liposomes and at low concentration, the suspension may look clear. Suppose your protein encourages liposome fusion (less likely), or something else in you solubilised proteins solution does (more likely), you could end up with a ratio-dependent increase in turbidity. The usual suspects that induce liposome fusion are bivalent cations (e.g. Mg2+). If you used such compounds at any point during your protein isolation it can be very difficult to get rid of it. Adding a few mM EDTA would help in that case.
Hi Salim,
If you would really like to get answer to your question, please be more specific. What detergent have you used? Speculation would be different depending on both the c.m.c. value and the "kind" (type) of your detergent.
I'm agree with Guus Erkens, but not only because the protein could induce lipid fusion , but also because your protein could fold in dimers or more depending on the concentration. Are you sure the protein is a monomer in all the concentrations of your experiment (tertiary structure)? Is it always the same secondary structure?
I am not sure if you are using purified synthetic lipids or isolated plasma membrane lipids. In any case, some lipids and some proteins are not soluble in detergents, hence the term "detergent resistant membrane". This type of lipids would include cholesterol and fully saturated phospholipids, sphingomyelin and ceramide, etc. Many membrane associating proteins naturally associate with these lipids in the plasma membrane, hence the term "lipid rafts". Thus, if you mix these lipids and membrane proteins with detergents, such as triton X-100, you WILL NOT get a homogeneous solution, but a turbid suspension.
One way to test this is to increase the temperature. The reason that these lipids and proteins don't dissolve in detergent is that these highly saturated and hydrophobic lipids form gel or liquid-ordered phase at ambient temp. Because of the tight lateral packing, it becomes difficult for detergent molecules to partition between these lipids, hence detergent resistance. Increasing temperature will shift the lipid bilayer phase from gel/liquid-ordered phase to liquid-disordered phase, making it easy to be dissolved.
So try to increase the temp. If you begin to see your lipid/protein mix dissolving into detergent at high temp, that means you are dealing with detergent resistant membrane and lipid rafts. And this detergent resistance is only natural for these molecules. This also potentially means that you may be bale to deduce the localization of these membrane proteins in the plasma membrane.
If your protein in B is a solubilized protein, it means that the buffer contains detergent and if your detergent is in excess in B (X times the cmc) when you add B at increasing volume you contribute to solubilize your lipids (the solution is less and less turbid) and when you have a sufficient quantity of detergent (by the addition of buffer B) you can reach the complete or quasi-complete solubilization of your lipids. The solubilization of your lipids implies that your solution become limpid.
Thank you all for the suggestions and ideas thus far.
Stefan: I originally had similar (but quite vague!) concepts of my detergent-solubilized protein serving to "mop up" the free lipid, such that the more protein that was present, the more lipid would be bound. thus reducing the overall turbidity. Thank you for helping to focus that idea; also, thank you for recommending ITC! (I had completely forgotten about that technique)
Guus and Mercedes: I haven't yet actively generated liposomes at this step. At the moment, the conundrum is that the MORE protein that is present (relative to lipid), the LESS turbid the mixture. Also, as an aside, I have examined my protein-containing liposomes and lipid-alone liposomes under negative staining and transmission electron microscopy, and the lipid-alone liposomes are smaller.
Yong: I've been using egg phosphatidylcholine (EggPC). Unfortunately, this is a bacterial integral inner membrane protein that is not highly expressed (naturally), so localization (in lipid rafts) may be a tad difficult to determine.
Célia: The concentration of detergent is the same in the existing protein purification solution, and the buffer used to first resuspend the dried-down lipids. As my next step though, I proceed to remove the detergent, thus resulting in formation of proteoliposomes.
It's a bad impression or you have less foam when you increase the protein concentration?
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