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How to Make Lipid Bilayers

lipid bilayers

Supported lipid bilayers are a very useful tool in many fields of cell and membrane biology. But how easy is it to make them?

Bilayers can be made quite reproducibly, once you have found a reliable protocol! However, it can take some time to optimize your technique, so to increase your chances, make sure you read up on the literature before you start.

This protocol that I find very useful describes the method to make bilayers for studying T cell immune synapses1, but the principles apply to other applications too.

Choosing Your Bilayer Composition

The first step when setting out to make bilayers is to decide on the lipid composition. Depending on your application, your lipid composition will vary, but most bilayers will primarily contain phosphatidylcholine (PC), which provides the structural framework.

You can use a polyethylene glycol (PEG) cushion to improve the uniformity of your bilayer and reduce protein aggregation. To do so, include a small fraction of lipids with PEG5,000 polymers covalently attached to the lipid headgroups. However, be aware that too much PEG will compromise the fluidity of your bilayer.

You can use different strategies to attach proteins to the bilayers; the two most common are via Ni-NTA lipids (which bind His-tagged proteins) and biotinylated lipids (which bind streptavidin, which in turn captures proteins tagged with biotin).

Creating the Bilayers – it’s a Kind of Magic

Below is a table that shows an example of a typical lipid mixture containing DOPC (1,2-dioleoyl-sn-glycero-3-phosphocholine) as the main structural component, DGS-NTA (1,2-dioleoyl-sn-glycero-3-[(N-(5-amino-1-carboxypentyl)iminodiacetic acid)succinyl] (ammonium salt)) for binding His-tagged proteins, DOPE (1,2-dioleoyl-sn-glycero-3-phosphoethanolamine)-biotin for attaching streptavidin, and PEG5,000-DOPE to improve the uniformity of the bilayer. The lipids are added at specific molar percentages. From the lipid concentrations and molecular weights, you can calculate the required volumes to get a rehydrated lipid mix of the desired final concentration (for example 1 mg/ml).

Lipid ComponentMol %
DOPC96.5
DGS-NTA2
DOPE-biotin1
PEG 5,000-DOPE0.5
Total100

Once you have the individual lipids in hand (which are stored in chloroform), mix them in a glass vial and then allow the mixture to evaporate overnight. This results in lipid sheets being deposited on the bottom of the glass tube – this is called lipid cake. You’ll see, it looks like a white deposit on the side of the vial.

Rehydrate the cake in 1ml of PBS, vortex for 30 seconds, and the cake will swell and form large vesicles of varied sizes. Next, squeeze the lipid mixture through a polycarbonate filter several times, in a process called ‘extrusion’, to generate a homogenous solution of liposomes (small lipid vesicles) of the desired size (typically 50-100 nm, as determined by the size of the filter). Store the liposomes in the fridge for up to two weeks.

In the next step, when you deposit these liposomes on a clean glass coverslip at the right concentration and in the right salt concentration (e.g. 5 mM Ca2+), they will spontaneously explode and fuse to form a supported lipid bilayer.

It’s like magic!

For this step, the preparation of the glass coverslips is extremely important – the glass needs to be clean, dry and hydrophobic. Otherwise, the bilayers may not be uniform or the vesicles could accumulate at the surface without bursting and merging to form the bilayer.

I had some issues with this step, but after contacting a researcher in the US, I was able to improve bilayer mobility by adjusting the cleaning procedure. I clean my slides by adding 10M NaOH for 15 minutes, then 70% ethanol for 5 minutes, followed by a thorough rinse with MilliQ water. I then dry the slides using a nitrogen stream.

After cleaning, you can pipette the lipids directly onto the slides making sure you cover the whole surface, and leave for 30 minutes. This allows enough time for the bilayers to form. After that, rinse thoroughly with plenty of PBS to remove any excess unfused vesicles from the surface.

Are Your Bilayers Mobile?

It’s important to have mobile bilayers so that your lipids and proteins can diffuse in the same way as they would in an actual cell membrane. To check the mobility of your bilayers, use a microscopy technique called Fluorescence Recovery After Photobleaching (FRAP)2, which relies on the diffusion of fluorescently-labeled molecules into a photobleached sample area.. You should check both the mobility of your lipids (by using fluorescently-tagged lipids, e.g. rhodamine-tagged lipids) and the mobility of your attached proteins (by using fluorescent streptavidin or fluorescently-labelled proteins).

You’re in Business…

In summary, bilayers are a cool tool for studying cell-cell interactions or membrane protein dynamics in a reductionist system that you can control. My work with bilayers has helped me show how T cells are able to transmit an activating signal from the outside of the cell to the intracellular signaling pathway.

Maybe bilayers could help you too!

References

  1. Beemiller P et. al. 2012. Imaging and analysis of OT1 cell activation on lipid bilayers. Protoc. Exch. doi:10.1038/protex.2012.028
  2. Axelrod D et.al. 1976. Mobility measurement by analysis of fluorescence photobleaching recovery kinetics. Biophys. J. 16: 1055–69.

1 Comment

  1. Isaac Schaider on September 29, 2017 at 4:31 am

    Hey Sophie,
    A few questions — is the lipid pipetted directly onto the glass cover slip then a glass slide is put on top of the cover slip or is the lipid put onto the glass cover slip and left open to the air with no glass slide needed? Also, for the preparation of the glass cover slips do both the glass slides and the glass cover slips used need to be hydrophobic or just the glass cover slips? Finally, once the lipid bilayer is made can I use HEPES buffer solution to wash away any unbound proteins from my lipid bilayer for the experiment I am performing?

    Thank you!

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