I am investigating whether lipids are bound to my plasma protein of interest and interfering with its quantitation by ELISA. I would like to pretreat samples with something capable of removing bound lipids. I have reason to believe the dissociation rate for my protein+lipids occurs on a reasonable timescale, so competitive binding could be a path forward.
What molecules could bind lipids tightly enough that they will not re-associate with my protein?
Sequestration of the lipids is preferred; I have a paper in which a fat sandwich ((my protein -- lipid -- apolipoprotein)) forms, and I would like to avoid that if possible.
We have tried extraction by activated charcoal and lost our protein.
We have tried extraction by immiscible organic solvent (e.g. chloroform) and irretrievably precipitated our protein.
Hi Jeremy, the major lipid binding protein in plasma is serum albumin.
Serum albumin does not have high affinity for lipids, but if you have it in you assay at high concentrations, it will sequester lipids due to mass action.
If you need proteins in your assay, try casein or IgG.
If you treat your protein with a detergent above its critical micellar concentration, the lipids will dissolve in the detergent micelles. You will then be faced with the necessity of separating the protein from the detergent before you can do the ELISA.
You could try running your protein over a hydrophobic interaction column or a reverse-phase HPLC column. Those should bind the lipids pretty tightly and the protein less tightly.
Steingrimur,
Thank you for your suggestions. I am working with some milk proteins that I have in the lab until I find a beta-lactoglobulin supplier that I like. It may have the kind of sequestration I'm looking for. Our assays are using 1% BSA (fatty acid free), while our protein is diluted to nM concentrations. If this is a lipid issue I'm dealing with, BSA doesn't seem to be sufficient. I'll check with my labmate when he's back from vacation, but I'm pretty sure IgG has been tried with no success.
Adam,
Thank you as well. I think I'll search out some hydrophobic interaction columns to give this approach another go. I've tried what you suggested with SDS and reverse phase spin columns, but my protein stuck too strongly. Taking the organic content of the mobile phase up high enough to remove it denatured it. I still like the approach though.
Hi Jeremy, don't dismiss serum albumin as just a molecular weight marker.
1% FAF BSA can bind a lot of lipids, especially if the ratio is 100,000 to 1
Small molecular weight drugs binding to serum albumin is a critical factor in pharmacokinetics. If a drug does not bind to serum albumin, it is pissed out in no time at all.
I wouldn't use SDS because it denatures proteins. Use a nonionic or zwitterionic detergent. CHAPS is a good choice because the micelles are very small and can be removed by dialysis or gel filtration. You could also try using Triton X-100, which can be removed using Biobeads (but be careful to use highest quality TX-100).
Hi Steingrimur,
I don't mean to dismiss it, and thanks for the insight into small molecule drugs and SA. I'm just noting there is no improvement in quantitation of our protein with the addition of 1% BSA. My thought is that we either need a stronger lipid binder, we have dissociation rate that is too slow for this strategy to work efficiently, or lipids are not the interfering agent.
Adam,
I have some TX-100 (Acros), but there's no indication of it's quality. I had an interesting conversation with Thermo tech support that when something like:
Me: I have a bottle of Triton X with no indication of purity, can you...
Tech: Yeah, that's a detergent, we don't indicate one. What were you looking for?
Me: The amount that is Triton X and the amount that isn't Triton X?
Tech: It's 100%
Me: So 100.0000%?
Tech: I can't give you that many 0's
Me: How many are you willing to commit to?
Tech: Maybe a couple less
So there you have it, about 100.00%!
More seriously, is there something you've come across in lesser quality TX100's? It's good to know your ingredients in the kitchen.
Triton X-100 is one of those detergents that can accumulate peroxides when exposed to air for a long time. These can cause damage to your protein. I would never use Triton X-100 that had been around the lab for a long time, stored in air at room temperature. You should either get a fresh bottle or, if you can afford it, by the product SurfactAmps-100 from Thermo-Fisher. This is a 10% solution of Triton X-100 stored under inert gas in a sealed ampule. Once the ampule is opened, any unused solution can be stored in the freezer to keep it from oxidizing.
if triton X is pored into the protein , as it is detergent .so it will not denature the protein ?
Triton X is particularly used for the concentration of hydrophobic or basic protein
As a nonionic detergent, Triton X-100 is not likely to denature proteins.
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