I am immobilizing biotinylated phosphoserine lipids on a streptavidin surface, but the wells are plastic (polystyrene). My pipettes, etc. are glass, and if I use plastic pipettes I lose the samples. I see a very long time course for binding to streptavidin (24 hrs) instead of the usual 1 minute or so. It is possible that much of my sample is bound to the plastic and is in slow equilibrium with buffer. I use 0.1% BSA and 0.1% Tween-20 in buffer. Perhaps I just increase the amount of Tween-20? Any suggestions for ways to block binding to plastic or better solubilize lipids in solution?
Polystyrene is a very hydrophobic surface, so lipids will surely bind to it. Proteins will, too. When doing enzyme assays in polystyrene plates, I always include a nonionic detergent to coat the surface so the proteins won't bind. However, I don't necessarily want to have detergent micelles in the assay because they might bind up hydrophobic inhibitors that I'm testing. It is sufficient to block protein binding to use a detergent concentration below the critical micellar concentration (CMC). For example, I would use no more than 0.01% Triton X-100, a bit below the CMC. Perhaps you are getting slow binding of your biotinylated lipids to the streptavidin surface because the Tween-20 concentration is above the CMC (0.0074%) and the lipids are bound up in detergent micelles. Consider experimenting with lower detergent concentrations.
Interesting, I had imagined that more Tween would be better. I will try less. I was worried the lipids themselves would form micelles (they are at 1~20 uM), and the detergent would solubilize and prevent this. I was also thinking that BSA would facilitate binding, acting as a carrier. Once bound, I will be washing so the initial buffer could be anything that doesn't damage streptavidin or lipid.
Right now I am seeing binding that increases over 24 hrs, as monitored by SPR
If you have long chain biotinylated lipid, it will certainly form aggregates. Critical aggregate concentration (CMC for micelles, CBC for bilayers) is roughly in the absence and presence of 100-150 mM salt 5 mM and 100 µM for PS with six carbon chains, and subnanomolar in both cases for PS with 16 carbon chains.
One question I must ask is that is it absolutely necessary for your experiment to do this. Is polystyrene necessity? How about it you just make phosphatidylserine small unilamellar vesicles including a few mol% headgroup-biotinylated PEG-lipid, to form a supported lipid bilayer on the bottom of streptavidin-coated eight-well plate? Of course you would need sufficient salt to block the electrostatic repulsion, but this should not be a problem. (Of course you would need to wash the 8-well plates with Helmanex-II first to make them clean enough.)
It is fairly tricky to block the binding. With quartz surface and proteins only thing that has really worked well has been the PEGylation of the quartz. If the kinetics of the binding to polystyrene and dissociating from it are fast enough for your blocking agent, it wouldn't work very well. I am afraid that you would either need to make the surface hydrophilic or preincubate with something amphipilic that dissociates from the surface extremely slowly. Custom-made imaging chamber might also help.
Albumin is useful to block lipids binding to Polystyrene. You must try several albumin concentratios to be sure about the experimental results.
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