Model lipid bilayer

A model lipid bilayer is any bilayer assembled in vitro, as opposed to the bilayer of natural cell membranes or covering various sub-cellular structures like the nucleus. They are used to study the fundamental properties of biological membranes in a simplified and well-controlled environment, and increasingly in bottom-up synthetic biology for the construction of artificial cells. A model bilayer can be made with either synthetic or natural lipids. The simplest model systems contain only a single pure synthetic lipid. More physiologically relevant model bilayers can be made with mixtures of several synthetic or natural lipids. A model lipid bilayer is any bilayer assembled in vitro, as opposed to the bilayer of natural cell membranes or covering various sub-cellular structures like the nucleus. They are used to study the fundamental properties of biological membranes in a simplified and well-controlled environment, and increasingly in bottom-up synthetic biology for the construction of artificial cells. A model bilayer can be made with either synthetic or natural lipids. The simplest model systems contain only a single pure synthetic lipid. More physiologically relevant model bilayers can be made with mixtures of several synthetic or natural lipids. There are many different types of model bilayers, each having experimental advantages and disadvantages. The first system developed was the black lipid membrane or “painted” bilayer, which allows simple electrical characterization of bilayers but is short-lived and can be difficult to work with. Supported bilayers are anchored to a solid substrate, increasing stability and allowing the use of characterization tools not possible in bulk solution. These advantages come at the cost of unwanted substrate interactions which can denature membrane proteins. The earliest model bilayer system developed was the “painted” bilayer, also known as a “black lipid membrane.” The term “painted” refers to the process by which these bilayers are made. First, a small aperture is created in a thin layer of a hydrophobic material such as Teflon. Typically the diameter of this hole is a few tens of micrometers up to hundreds of micrometers. To form a BLM, the area around the aperture is first 'pre-painted' with a solution of lipids dissolved in a hydrophobic solvent by applying this solution across the aperture with a brush, syringe, or glass applicator. The solvent used must have a very high partition coefficient and must be relatively viscous to prevent immediate rupture. The most common solvent used is a mixture of decane and squalene. After allowing the aperture to dry, salt solution (aqueous phase) is added to both sides of the chamber. The aperture is then 'painted' with a lipid solution (generally the same solution that was used for pre-painting). A lipid monolayer spontaneously forms at the interface between the organic and aqueous phases on either side of the lipid/solvent droplet. Because the walls of the aperture are hydrophobic the lipid/solvent solution wets this interface, thinning the droplet in the center. Once the two sides of the droplet come close enough together, the lipid monolayers fuse, rapidly excluding the small remaining volume of solution. At this point a bilayer is formed in the center of the aperture, but a significant annulus of solvent remains at the perimeter. This annulus is required to maintain stability by acting as a bridge between the ~5 nm bilayer and the 10's of micrometer thick sheet in which the aperture is made. The term “black” bilayer refers to the fact that they are dark in reflected light because the thickness of the membrane is only a few nanometers, so light reflecting off the back face destructively interferes with light reflecting off the front face. Indeed, this was one of the first clues that this technique produced a membrane of molecular-scale thickness. Black lipid membranes are also well suited to electrical characterization because the two chambers separated by the bilayer are both accessible, allowing simple placement of large electrodes. For this reason, electrical characterization is one of the most important methods used in conjunction with painted lipid bilayers. Simple measurements indicate when a bilayer forms and when it breaks, as an intact bilayer has a large resistance (>GΩ) and a large capacitance (~2 µF/cm2). More advanced electrical characterization has been particularly important in the study of voltage gated ion channels. Membrane proteins such as ion channels typically cannot be incorporated directly into the painted bilayer during formation because immersion in an organic solvent would denature the protein. Instead, the protein is solubilized with a detergent and added to the aqueous solution after the bilayer is formed. The detergent coating allows these proteins to spontaneously insert into the bilayer over a period of minutes. Additionally, initial experiments have been performed which combine electrophysiological and structural investigations of black lipid membranes. In another variation of the BLM technique, termed the bilayer punch, a glass pipet (inner diameter ~10-40 µm) is used as the electrode on one side of the bilayer in order to isolate a small patch of membrane. This modification of the patch clamp technique enables low noise recording, even at high potentials (up to 600 mV), at the expense of additional preparation time. The main problems associated with painted bilayers are residual solvent and limited lifetime. Some researchers believe that pockets of solvent trapped between the two bilayer leaflets can disrupt normal protein function. To overcome this limitation, Montal and Mueller developed a modified deposition technique that eliminates the use of a heavy non-volatile solvent. In this method, the aperture starts out above the water surface, completely separating the two fluid chambers. On the surface of each chamber, a monolayer is formed by applying lipids in a volatile solvent such as chloroform and waiting for the solvent to evaporate. The aperture is then lowered through the air-water interface and the two monolayers from the separate chambers are folded down against each other, forming a bilayer across the aperture. The stability issue has proven more difficult to solve. Typically, a black lipid membrane will survive for less than an hour, precluding long-term experiments. This lifetime can be extended by precisely structuring the supporting aperture, chemically crosslinking the lipids or gelling the surrounding solution to mechanically support the bilayer. Work is ongoing in this area and lifetimes of several hours will become feasible. Unlike a vesicle or a cell membrane in which the lipid bilayer is rolled into an enclosed shell, a supported bilayer is a planar structure sitting on a solid support. Because of this, only the upper face of the bilayer is exposed to free solution. This layout has advantages and drawbacks related to the study of lipid bilayers. One of the greatest advantages of the supported bilayer is its stability. SLBs will remain largely intact even when subject to high flow rates or vibration and, unlike black lipid membranes, the presence of holes will not destroy the entire bilayer. Because of this stability, experiments lasting weeks and even months are possible with supported bilayers while BLM experiments are usually limited to hours. Another advantage of the supported bilayer is that, because it is on a flat hard surface, it is amenable to a number of characterization tools which would be impossible or would offer lower resolution if performed on a freely floating sample. One of the clearest examples of this advantage is the use of mechanical probing techniques which require a direct physical interaction with the sample. Atomic force microscopy (AFM) has been used to image lipid phase separation, formation of transmembrane nanopores followed by single protein molecule adsorption, and protein assembly with sub-nm accuracy without the need for a labeling dye. More recently, AFM has also been used to directly probe the mechanical properties of single bilayers and to perform force spectroscopy on individual membrane proteins. These studies would be difficult or impossible without the use of supported bilayers since the surface of a cell or vesicle is relatively soft and would drift and fluctuate over time. Another example of a physical probe is the use of the quartz crystal microbalance (QCM) to study binding kinetics at the bilayer surface. Dual polarisation interferometry is a high resolution optical tool for characterising the order and disruption in lipid bilayers during interactions or phase transitions providing complementary data to QCM measurements. Many modern fluorescence microscopy techniques also require a rigidly-supported planar surface. Evanescent field methods such as total internal reflection fluorescence microscopy (TIRF) and surface plasmon resonance (SPR) can offer extremely sensitive measurement of analyte binding and bilayer optical properties but can only function when the sample is supported on specialized optically functional materials. Another class of methods applicable only to supported bilayers is those based on optical interference such as fluorescence interference contrast microscopy (FLIC) and reflection interference contrast microscopy (RICM) or interferometric scattering microscopy (iSCAT). When the bilayer is supported on top of a reflective surface, variations in intensity due to destructive interference from this interface can be used to calculate with angstrom accuracy the position of fluorophores within the bilayer. Both evanescent and interference techniques offer sub-wavelength resolution in only one dimension (z, or vertical). In many cases, this resolution is all that is needed. After all, bilayers are very small only in one dimension. Laterally, a bilayer can extend for many micrometres or even millimeters. But certain phenomena like dynamic phase rearrangement do occur in bilayers on a lateral sub-micrometre lengthscale. A promising approach to studying these structures is near field scanning optical microscopy (NSOM). Like AFM, NSOM relies on the scanning of a micromachined tip to give a highly localized signal. But unlike AFM, NSOM uses an optical rather than physical interaction with the sample, potentially perturbing delicate structures to a lesser extent.