Is there anyone with experience in flotation assays with PS lipsomes? In detail I want to test different protein constructs (wt vs. mutants) for their ability to bind to PS lipsomes in vitro. In my pilot experiment I used a sucrose gradient (30% - 25% - PBS) and a 1:80 protein:lipid molar ratio. I incubate the protein with lipids for 10 min at RT, then add sucrose to final conc. of 30% then overlay the batch carefully with 25% sucrose and finally overlay the batch with PBS. I spin the batch for 1h at 4°C using a TLS-55 swinging bucket rotor and collect the fractions from top to bottom (Hamilton syringe) right after centrifugation. Unexpectedly, the protein can be found in all fractions (top, middle, bottom) in nearly identical quantities (see attachment for the wt construct), but I would expect the protein to be found mainly in the top fraction. Furthermore I dont' see a difference w/ or w/o PS liposomes! It's already known for the wt construct to be bound to PS liposomes so something seems to be wrong with my setup. I am wondering if the high sucrose concentration may hinder binding of the protein to the lipids? Any comments (i.e. different floatation agents; Accudenz???) are highly appreciated.
Hi,
We did a floatation assay for POPC:CHOL 2:1 liposomes also in sucrose (LUVs with 100 nm diameter): from bottom to top: 100 uL sample-25 uL buffer-175 uL sucrose 2.4 M, then 400 uL sucrose 0.8 M and finally 300 uL sucrose 0.5 M. The rotor was TLA120.2 and 100000 rpm 3 hours at 4ºC.
Anyway, If your protein induces fusion or aggregation to some extent, maybe you get protein in every sample....
Thanks Beatriz! I am also not very convinced that floatation assay works with proteins tubulating membranes, which is the case for the protein I am investigating. Therefore another question arises: is a floatation assay a reasonable way to proof membrane binding for proteins which tubulate membranes?
Hi Chris,
there are several things you can test. The high sucrose conc. shouldn't be a problem. Is your protein stable in the sucrose solution? For me it helped to make the sucrose solution in my working buffers (i.e. PBS or Hepes with 150mM NaCl) to keep the protein stable. If your protein is fine @ RT then you can also do the spin @ RT. Sometimes liposomes don't like the temp. change. Usually I spin for 2h in the same rotor that you use. Did you test what the lowest sucrose percentage is that your liposomes can penetrate? I didn't use PS liposomes but in my case the sucrose percentage was 10%, therefore I mixed the proteins to a final conc. of 37% sucrose (1 vol), this was overlayed with a 12% sucrose layer (12 vol), overlayed by PBS (2 vol). Maybe the difference between your sucrose conc. is not high enough? How do you detect your liposomes? Sometimes membrane dyes can do strange things to the membrane. Did you check if your liposomes are ok (for example by EM)? Or has the size of the liposomes an influence on the binding of your protein?
I hope I could help you a bit!
Cheers,
Ben
Hi Benjamin, thanks for the tips!
I also prepare the sucrose in the protein's buffer (PBS or 150mM NaCl/HEPES). Good idea to check all fractions via EM for proper liposome formation! I didn't test the lowest sucrose percentage so far since I just began with that assay and our labeled lipids didn't arrive yet. (I am at the very early stage of the establishment of the assay with my protein)
What lipid ratio (labeled:non-labeled) do you suggest? The size of the lipids (at least unlabeled) has no influence of binding of the protein, since I never filter them and the wt protein is always tubulating the lipids (which is a reasonable proof for binding).
I used 0.2 mol% acyl chain labeled NBD-PE for my liposomes. This is enough to see them after the flotation. This also allows for quantification with an fluorescence imager. Usually I was able to recover ~60% of my lipos from the top fraction.
Good luck!
Hi Chris,
If your floatation assay is giving problems, you may consider doing a pull-down experiment. In short, this means mixing your protein with liposomes, followed by a high speed centrifuge step to pellet those liposomes. If you analyse the pellet fraction and supernatant for the presence of your protein (using SDS-PAGE/Western) it should be possible to quantify the interaction. You can find an example of such an experiment here: http://www.nature.com/nsmb/journal/v17/n3/full/nsmb.1770.html, the results can be found in figure 2g.
I hope this was helpful!
Hi Guus,
thanks for your post! I already did spin assays. Anyway, one of "my" reviewers want me to do floatation assays as well "to proof the findings independently".
Well, of course I am happy to them dear reviewer, this will make the paper much more convincing!
To bring this to an end.
Finally I found out, that the assay is not very applicable at all with protein I am working with. I checked for proper floatation of the lipids containing 1-2% fluorescently labeled PS (thanks again Benjamin for the advise) without protein and there is a nice band visible in UV in the top phase. However if I incubate the batch with my protein fluorescence is equally distributed in all fractions. This is probably due to the protein's ability to deform the lipid vesicles making them not floating to the buffer fraction on top. Instead they are distributed in all three fractions so the protein does.
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