We are preparing liposomes (900 nm) of a recombinant protein (55 kDa). After the extrusion of the liposomes, we performed 2 centrifugations to separate our larger liposomes from small liposomes and free protein (pellet: liposomes of interest, supernatant: free protein and small liposomes). We evaluated the protein concentration in the last resuspended pellet and the two supernatants. We found an interference of the lipids with this method (BCA quantitation). So, which should be a good method to evaluate the encapsulation efficiency of the protein in the pellet (larger liposomes) after the separation process? It should be a method were lipids do not interfere. Thanks a lot.
Gel filtration chromatography should easily separate liposomes from free protein. The liposomes will elute first and you can then measure the amount of protein left over that elutes later.
Lipid interference in protein assays of the liposomes may be solvable by using organic solvent such as acetone, methanol, or chloroform to dissolve the lipids and precipitate the protein. Then centrifuge the protein and use the pellet for the protein assay.
You can also try running the liposomes on SDS-PAGE to see how much protein is in them. You may encounter a problem with interference with all the lipid, however. Use the smallest possible amount of liposomes and a sensitive protein stain, such as a fluorescent one, to keep the amount of lipid to a minimum.
Good morning Dr Shapiro. Thanks for your answer. About the separation of liposomes from free protein, we do it through centrifugation with good results. Using GFC should be an option but we also want to separate small liposomes from large liposomes. And if we are going to perform a quantitation of protein in the supernatant or the elute there's going to be lipids from the small liposomes so that's our main problem.
About your other two options that's what we were exploring. About the extraction that you mention, do you have any protocol or papers about it? If we get rid of lipids by doing a typical extraction I think we can perform the BCA quantitation of the water-soluble protein.
About the SDS-PAGE we also took it into account but it's kind of time consuming. But we are going to perform some other tests. We already have a good concentration of pure protein and we hydrate the liposomes with this solution.
Do you think there's a way to analyse the encapsulation efficiency through UV spectrophotometry? The problem about this is that we need to find a solvent that dissolves both the liposomes and the protein and since our protein is water-soluble I don't know if a good idea is to dissolve the lipids in mixtures of ethanol:water.
On the other hand, do you know what should be a good method to improve encapsulation of water-soluble proteins in liposomes? (we are using the hydration of thin lipid film method). Thanks a lot for your answer.
I read them but they don't report too much information about the separation process.
Separation of unincorporated protein from liposomes by gel filtration will require a gel filtration resin that is designed for separating very large proteins, in order to assure that the protein is still in the column when the liposomes elute, like Bio-Gel A5m (http://www.bio-rad.com/webroot/web/pdf/lsr/literature/4006139.pdf). These resins are for gravity elution only, not for FPLC.
Perhaps you can deal with the problem of small and large liposomes complicating the quantitation of protein by treating the liposomes with trypsin to digest the external protein (and itself) down to small peptides. These would easily be separated from both small and large liposomes by centrifugation or gel filtration.
I think you could then measure the incorporated protein using detergent to dissolve the liposomes and the BCA assay to measure the protein released.
After doing protocol that dear Adam mentioned, maybe analytical hplc be helpful.
Dr. Shapiro, thanks for your answers. What are the best detergents to dissolve liposomes without affecting the protein? We have SDS, Triton, Tween, etc and we are gonna read some papers, but I'm asking based on your experience.
It depends on what you want to do next. SDS will denature the protein, but Triton X-100 and Tween (for example) will not. For a protein assay, consult the product literature to find out how much of each detergent can be tolerated. I would start with Triton X-100. It does a good job of dissolving phospholipids. Octylglucoside is another favorite.
We are going to quantify the protein by UV spectrophotometry so I assume we don't have problem if the protein is denatured or not. But it has to be dissolved and out of the liposomes. We performed the analysis with TritonX100, CHAPS and SDS but it seems that the liposomes weren't affected by the addition of these detergents a concentrations slightly above their CMC. We re going to try to optimize this but we want no interference with UV analysis.
I think these detergent not have absorption with UV but you can check it. If these had absorption you can use them for blanking spectrophotometer.
Triton X-100 has UV absorbance due to a phenyl ring, so you should probably avoid it.
You will need to use several times the CMC of the detergent if there is a lot of lipid. For example, you might need 1% Tween-20 to dissolve the liposomes because the CMC is so low compared to the lipid concentration.
According to that. I know Triton X-100 has UV absorbance. Why are going to run some experiments to check if the background is too noisy. About Tween-20, does it absorb? Why is it better than Triton X-100? The lipid concentration is 30 mg/mL. But we are going to employ probably supernatants after centrifugation where lipid concentration is lower. How do I know what's the correct amount of detergent to use according to CMC and lipid concentration? Thanks.
It depends on the detergent and the lipid. I suggest you do some simple experiments using empty liposomes and a few detergents, titrating the detergent until the lipids are dissolved. Tween-20 does not have any UV chromophore is the 280 nm range, so it should be OK for your purpose.
How we can analyze if the liposomes are dissolved? performing DLS experiments? Thanks a lot for your ideas, we've been thinking about them.
You want to know if the liposome-encapsulated protein is aggregated? DLS would be the first thing to try because it is such a simple measurement. It might suffer from interference from the detergent-lipid mixed micelles, so you will have to prepare protein-free control samples.
Another method you could try is size-exclusion chromatography SEC) to separate the aggregated and unaggregated protein. Coupling SEC with a multiangle light scattering detector (SEC-MALS) is a particularly powerful analytical approach to protein aggregation measurement. Keep in mind, however, the possibility that detergent-solubilization of the liposomes could alter the aggregation state of the encapsulated protein. Think about experimental controls for that possible effect.
I don't want to know about the aggregation of the protein encapsulated, just the encapsulation efficiency. First I'm going to perform everything without protein, and taking into account a control without detergent to compare if they are dissolved. Thanks a lot.
Avinash Singh Patel Yes and it didn't work so well, so I'm currently analyzing the protein via SDS-PAGE