Supported Lipid Bilayers: Development and Applications in Chemical Biology
Volker Kiessling, Marta K. Domanska, David Murray, Chen Wan and Lukas K. Tamm, University of Virginia, Charlottesville, Virginia
Lipid bilayers supported on solid substrates were developed almost a quarter century ago as a new model membrane system to study fundamental properties of biological membranes and their constituent lipid and protein molecules, as well as for numerous practical applications. In this review, we summarize the development of supported bilayer-based model membrane systems of increasing complexity while always keeping in mind their primary purpose, which is to reproduce the complex fluid structure of biological membranes to mimic cell membrane-based molecular processes as close to physiological reality as possible. This process must include maintaining the structure and fluidity of the lipid bilayer, as well as the structure and function of reconstituted membrane proteins. We start the review with a general description of a contemporary picture of a generic biological membrane and proceed with an account of how supported membranes recapture different physiological aspects of biological membranes, such as membrane fusion, cell adhesion, or membrane permeability. Supported membranes are particularly good models for reproducing the fluidity of cell membranes even when different liquid lipid phases coexist. They are also superb models for measuring ligand-receptor interactions between membrane-bound receptors and soluble or membrane-bound ligands. The chemical biology of these and many other lipid-lipid, lipid-protein, and protein-protein interactions that have been investigated in supported bilayers are summarized in several sections that form the main body of this review. The article closes with summaries of practical procedures to prepare and characterize supported bilayers.
Since the lipid bilayer was accepted as the basic principle of design for biological membranes more than 40 years ago, researchers have developed model systems to study numerous aspects of biomembrane structure and function. Large multilamellar liposomes have been used frequently in early X-ray diffraction and calorimetric studies that characterize the structure and thermodynamic properties of lipid bilayers. Sonication of these structures leads to small unilamellar vesicles, which are excellent model systems for numerous spectroscopic and binding studies with model membranes. Other experiments require larger or even “giant” unilamellar vesicles, produced by extrusion through filters or swelling from solid lipid deposits with or without applied electric fields. For some applications, such as recording electrical currents across membranes or measuring the mobility and lateral organization of membrane constituents, planar bilayer geometry is advantageous. To this end, single unsupported planar bilayers are suspended with or without solvent in a small orifice in a Teflon septum that separates two chambers that are electrically connected to equipment that can record currents from single molecular channels (1, 2). A wealth of information on the molecular properties of ion channels, toxins, and other membrane-interactive proteins and substances has been obtained using such planar bilayer recordings.
A different planar model membrane design was developed in the early 1980s mainly from research conducted in Harden McConnell’s laboratory at Stanford University—the supported lipid bilayer. The biological motivation driving these early studies was to provide planar model membranes for observation by epi-fluorescence microscopy and to use them as surrogate target membranes to study molecular interactions in immunological synapses between antigen-presenting cells and T-cell receptor-bearing lymphocytes. A precursor of the supported lipid bilayer has been the supported lipid monolayer, in which a lipid monolayer is deposited on a long-chain alkylated substrate (3). This early design using silane chemistry for alkylation was later superseded with sulfur chemistry on slides with thin evaporated gold surfaces. However, because these systems mimic only half a bilayer and cannot accommodate integral membrane proteins, which provide so many key functions to biological membranes, they are not considered in this article.
In a quest to produce a model system that more faithfully resembles biological membranes and that also allows the functional reconstitution of integral membrane proteins, we have developed the supported bilayer as a new model membrane system (4, 5). Supported bilayers have since become widely used in many different areas of chemical, biological, and materials research. Applications range from fundamental studies of bilayer and membrane properties to their phase behavior; membrane protein binding; membrane protein structure; electrophoresis and microanalysis of membrane components; functional studies of membrane receptors, pores and channels; interactions with components of bound cells, viruses, and cellular organelles; and so on. This article is organized in four main sections: We will first briefly review current concepts of cell membranes and how supported bilayers can serve as models for cell membranes. We will next describe biophysical properties and chemical interactions that govern the structure, stability, and use of supported bilayers in chemical and biological analysis. The article will close with two sections on procedures that are commonly used to prepare and characterize supported bilayers.
Supported Bilayers as Models for Cell Membranes
Biological membranes are formed by a lipid bilayer with embedded proteins. The bilayer-forming lipids have polar headgroups that face the aqueous surroundings of the membrane as well as two hydrophobic tails that face the interior of the membrane. A key feature of cell membranes is that they are both fluid but still highly ordered in the membrane plane, as has been captured in the early conception of the fluid mosaic model of biological membranes (6). Lipids move laterally at a fast rate with a lateral diffusion coefficient on the order of 1 pm2/s, but they cross the membrane by flip-flop only once every few hours (7). The slow kinetic rate of lipid flip-flop allows the cell to establish asymmetry across the bilayer, which, with the input of energy, can be maintained for the entire lifetime of a cell. Membrane proteins come in different varieties. Some have sequences that cross the membrane, and therefore they are called integral membrane proteins. Other membrane proteins, such as the peripheral membrane proteins, are attached to the membrane surface either electrostatically or by means of covalent linkage to single or multiple lipid tails. A third class of membrane proteins dip into only one leaflet of the bilayer, where they are stabilized by hydrophobic interactions with the lipids in the bilayer. Regardless of the specific nature of membrane attachment and incorporation, membrane proteins can diffuse relatively rapidly in the plane of fluid membranes. However, they never cross the lipid bilayer except during biogenesis, which requires specialized machines called translocons that translocate newly synthesized polypeptide chains across cell membranes in a complicated and still poorly understood energy-requiring process (8).
The notion of a biological membrane represented by a uniform sea of lipids with proteins freely floating in this environment has been significantly revised in the last decade or so (Fig. 1a) (9). It has become increasingly clear that biological membranes are laterally structured even if they are still predominantly fluid with regard to in-plane diffusion of protein and lipid components. First and foremost, membranes are highly crowded with proteins. Although the crowding varies greatly between different biological membranes (e.g., being very high for energy-converting membranes in mitochondria and photosynthetic organelles and rather low for membranes of myelin sheaths that wrap around and electrically insulate neurons), it is clear that membrane proteins do not act independently in membranes but often form clusters of interacting species. In a typical average cell membrane, about a quarter to half of the total cross-sectional area may be occupied by protein. If one considers proteins that have larger extramembraneous domains than transmembrane domain cross-sections, the area fraction covered with protein may reach half to three quarters of the entire membrane surface. Thus, membrane proteins and lipids are both solvents and solutes at the same time. An alloy of hydrophobic proteins and lipids may describe the true nature of biological membranes more accurately than a dilute two-dimensional solution of proteins in a vast sea of lipids. Second, unlike commonly prepared model membranes, biological membranes contain thousands of different lipid species (10). The lipid species distinguish themselves by many different headgroup classes and chain compositions as well as an expansive combinatorial diversity of these structural elements. A very large amount of data has been accumulated over the past four decades on the structures, energetics, and phase properties of pure single and two- or three-component lipid bilayers as summarized in two monographs (11, 12). We know from this data that high- and low-melting lipids tend to separate into different phases and that the lipid phase properties are often dramatically modulated by cholesterol.
Even though cholesterol (substituted by other sterols in plants) is chemically classified as a lipid, the molecule is better grouped it into a special category, which is distinct from membrane proteins and (phospho- and sphingo-) lipids, when discussing its role in biomembrane structure and function. Cholesterol constitutes between 25% and 40% of the total lipid plus cholesterol fraction of most typical cell membranes. These numbers translate into a 33-66% fraction of the noncholesterol lipids, or considering its smaller size, about a 10-20% area fraction of the lipid-occupied area of a biological membrane. Therefore, biological membranes should be considered two-dimensional liquid alloys of three major component classes: lipids, proteins, and cholesterol.
Figure 1. Modeling cell membranes with supported bilayers. (a) Contemporary model of a biological cell membrane. The lipid bilayer is heterogeneous across and in the plane of the membrane. The outer leaflet (bottom pink) is enriched in sphingomyelin and the inner cytoplasmic leaflet (top yellow) is enriched in phosphatidyl-serine and phosphatidyl-ethanolamine. Cholesterol and phosphatidyl-choline are distributed more evenly between the two leaflets. Cholesterol-rich lipid domains ("rafts") are present in register in both leaflets. Transmembrane and monotopic (partially inserted) membrane proteins (larger structures green) occupy much of the available membrane area and are also clustered into functional complexes in many cases. Lines (brown) on the top symbolize interactions with the cytoskeleton, and branched lines (orange) on the bottom of the membrane symbolize carbohydrates of the glycocalix. (b) Supported bilayer with inserted proteins directly supported on a hydrophilic inorganic substrate (gray), (c) Supported bilayer with inserted proteins supported on a (tethered) polymer cushion (purple) that increase the space between the membrane and the support.
Although some membrane proteins are known to interact with cholesterol, cholesterol interactions with other lipids have been studied much more thoroughly. Most notably, cholesterol interacts strongly with sphingomyelin, perhaps by forming complexes, which results in a liquid-liquid phase separation of cholesterol-rich and cholesterol-poor phases (13). The cholesterol-rich phases exhibit more chain order and therefore are referred to as liquid-ordered (l0) phases, whereas the cholesterol-poor phases are called liquid-disordered (ld) phases. In the cell-biological literature, l0 phases have often been referred to as “lipid rafts” mostly because certain membrane proteins that are functionally linked are coextracted by mild detergent treatment from complex cell membranes. Despite the functional linkage of membrane proteins that are involved, for example, in cell signaling and the observation that they appear in the same biochemical fraction as cholesterol and sphingolipids (14), larger platforms of phase-separated cholesterol-rich lipid phases, or “lipid rafts,” that could assemble these proteins have never been observed directly in cell membranes. Their existence is mainly based on inference from studies with pure lipid-cholesterol systems in which cholesterol-rich l0-phase domains can be observed. Therefore, the concept of “lipid rafts” as a means of organizing cell membranes remains highly controversial. Several excellent reviews critically illuminate the status of this sometimes-confusing held of membrane biology (15, 16).
Another important component of cell membrane structure and function is not integral to the membrane but is adjacent. Cell membranes are often linked to elements of the cytoskeleton and the extracellular matrix via specialized embedded proteins. Actin networks on the inside of cell membranes and extracellular proteins on the outside confine diffusive and directed motions of many integral membrane proteins either by direct and indirect attachment to these structures or simply by putting up fences on the already crowded membrane surface (17). The effect of these extramembraneous protein networks is often manifested in irregular hop-diffusion or anomalous diffusion when analyzed in cell membranes with sufficient time and spatial resolution. Such membrane-attached protein networks may be less significant in the case of many intracellular organelle membranes, although for some of these structures (e.g., for clathrin- or COP-coated endocytic vesicles), similar restrictions on protein motion may apply.
Supported bilayers with and without embedded membrane proteins offer new opportunities to study several of the aforementioned aspects of biological membranes (Figs, lb and lc). Because of their planar geometry and stability provided by the solid support, lateral diffusion of membrane components can be measured in a straightforward manner by fluorescence recovery after photobleaching (FRAP) or single particle tracking (SPT). It has been known since the inception of supported bilayers that phase transitions of lipids are preserved in these systems as evidenced by abrupt changes in lateral diffusion coefficients and morphological changes (4). Lateral phase separations between l0 and ld phases and lipid asymmetry may also be conveniently studied in supported bilayers (18, 19). Supported bilayers continue to contribute to the current understanding of the effect of cholesterol on complex lipid mixtures, and interesting new insights may be expected in the near future. Secondary structures and orientations of secondary structure elements of membrane proteins and membrane-bound peptides are conveniently measured by polarized Fourier-transform infrared (FUR) spectroscopy in supported bilayers (20). More recently, helix-helix interactions of signaling proteins have been measured in supported membranes (21). Helix interactions, for example in receptor dimerization, are thought to be important in transmembrane signaling of activated receptors, but relatively few techniques can measure such interactions in membrane environments. Another area where supported bilayers offer distinct advantages is studies of the kinetics of ligand binding to membrane-bound receptors (22). Supported membranes have also been used successfully to measure protein-mediated fusion of vesicles, for example in virus entry (23) or exocytosis (24).
On the analytical practical side, there is much interest in using supported membranes with incorporated ion channels as biosensors for various analytes (25-27). Another interesting application is to separate membrane components in situ by electrophoresis in supported membranes (28, 29). Such techniques could have a significant future impact on the proteomics of membrane proteins, that is, an area that is currently underdeveloped because of difficulties with appropriately separating membrane proteins in complex mixtures by more standard techniques. Finally, supported membranes continue to provide very interesting engineerable substrates that mimick natural cell surfaces to trigger the differentiation of adjacent cells such as in the previously mentioned immunological synapses (30).
Interactions Between Lipid and Protein Components in Supported Bilayers
A key feature of biological membranes is their graded fluidity. Any useful membrane model system must strive to preserve the characteristic fluidity of lipid and protein components. This concept clearly has been a great challenge in this field. Although it is relatively easy to maintain the fluidity of the lipid components and peripherally attached protein components as demonstrated already in the earliest publications on supported bilayers (5, 31), to achieve the same result with integral membrane proteins has proven to be much more difficult. It is known that the gap between supported membranes and solid glass or quartz supports is on the order of 1 to 2 nm (32, 33), and this gap is filled with a layer of water that is sufficient to lubricate the lower leaflet of the bilayer to permit rapid lipid diffusion (Fig. 1b). However, this gap is not large enough to accommodate integral membrane proteins with significant extramembraneous domains. Such proteins are generally immobilized by interaction with the glass support. Much research activity has been directed in the last decade to uncouple such unwanted nonphysiological interactions, by increasing the gap distance with intercalated water-soluble polymers, either by simple physisorption or covalently attached to one or both surfaces by forming tethers between the substrate and the membrane (Fig. 1c). In the following section, we summarize strategies that have been successful for creating polymer-supported bilayers. The subsequent sections proceed in turn to brief summaries of lipid-lipid interactions, lipid-protein interactions, and protein-protein interactions in supported bilayers, and then to studies of membrane fusion and cell adhesion using supported membranes as a model for one membrane in these membrane-membrane interactions. The final sections summarize recent advances in patterning supported membranes and using supported membranes as novel analytical tools in chemical biology. Depending on the particular application, bilayers directly supported on solid substrates or (tethered) polymer-supported bilayers are preferred as will be mentioned in each case.
Polymer-supported bilayers with and without tethers
The idea of using polymer-supported bilayers has been around for more than a decade (34), but it became practical for chemical and biological applications only more recently. Early versions have used relatively short tethers to link the membranes to the solid substrate and thereby increase their durability for practical applications (25, 35). Because these approaches do not increase the gap distance between substrate and membrane and therefore have not been used to reconstitute integral membrane proteins functionally, they will not be discussed here.
Our group has developed a polyethylene glycol (PEG)-based polymer support for bilayers on glass, quartz, or oxidized silicon (36). PEG that consists of 77 subunits bridges a phospholipid on one end and a silane group on the other end, which allows the covalent tethering of the lipopolymer to free silanols on the silicon dioxide surface. In all, 25% of the reconstituted integral membrane protein cytochrome b diffused freely in supported bilayers that contained 3% of the lipopolymer in the proximal leaflet. Integral membrane SNARE proteins were 80% mobile with a diffusion coefficient of 0.8 μm2/s in this system (37). The distance between the substrate and membrane increased from 2 to 4 nm without and with the polymer, respectively (33). A similar approach was introduced by Naumann’s group, who used dioctadecylamine(poly(ethyloxazoline)-8988) and octadecyl(poly(ethyloxazoline)-5822) with 85 and 56 monomer units, respectively, as lipopolymers in their bilayers (38). In this case, the lipopolymers were photo-cross-linked to the glass via a benzophenone silane photocoupling agent. The tethered polymer obstructs lipid diffusion at high concentrations but not at low concentrations (39). The increased stability of the monolayer that faces the substrate allows the preparation of asymmetric supported bilayers that contain separated regions of liquid-ordered and liquid-disordered phases. Lipid raft mixtures in the substrate-proximal layer induce registered raft domains in the distal layer (40, 41). Sackmann’s group used lipopolymers that contain polymerized 2-methyl-2-oxazoline of different lengths to integrate the large transmembrane cell receptor integrin αIIβ3 (42). In all, 20% of the integrins were mobile with a relatively low diffusion coefficient of 0.03 μm2/s. A different approach to form a cushion between the solid support and membrane uses the Langmuir-Blodgett technique to transfer several layers of trimethylsilyl cellulose onto a glass substrate. When supported bilayers were formed on this substrate by direct vesicle fusion, 25% of the reconstituted integrins were mobile with a diffusion coefficient of 0.6 μm2/s (43).
Studies of raft-like lipid domains in supported bilayers
Substantial literature has been published on studies of lipid domains in supported bilayers. Many have investigated lipid mixtures with coexisting gel and liquid-crystalline phases by atomic force (AFM), epifluorescence, and near-field fluorescence microscopy (NSOM) (see, for example, References (44-47)). Other studies have focused on (“raft”) lipid mixtures that form coexisting lo and ld phase domains in the presence of cholesterol (19,48-50). AFM studies are usually carried out on mica substrates, which have the advantage of being atomically flat but are less hydrated than glass or quartz substrates that are commonly used in fluorescence microscopic studies. Although higher resolution is achieved by AFM than by optical microscopy, the bilayers are more tightly coupled to the substrate and are not always fluid on mica. Fluorescence has the advantage that different molecular fluorescent dyes can be used to probe different aspects of complex lipid bilayers, such as diffusion, order, and phase partitioning, but it has the obvious disadvantage that the behavior of the probe is observed, which may or may not reflect the behavior of the host lipids accurately. Judicial choices must be made when selecting lipid probes for different purposes. Typical “raft” lipid mixtures contain about equimolar sphingomyelin, phosphatidylcholine, and cholesterol. Studies have shown that the area of l0 phases in such bilayers depends linearly on the cholesterol concentration and that a percolation threshold exists at about 25 to 30 mol % cholesterol, where l0 domains become connected and ld domains become disconnected (19). Because this mixture is at about physiological concentrations of cholesterol, cells may use shifts in cholesterol concentration as a switch to connect different groups of membrane proteins and thereby regulate their function.
An exciting new development in supported bilayer technology is that bilayers with asymmetric lipid distributions can be prepared. Although it is not so difficult for gel phase lipids with very low diffusion coefficients, it is much more challenging with fluid phase bilayers. However, these challenges have been solved recently (18). It is now possible to study phase coupling across the mid-plane of bilayers with coexisting l0 and ld phases (40, 41). The observation that l0 phases can be induced in lipid mixtures that mimick the inner leaflet of cell membranes that do not exhibit such phases on their own has important potential consequences on signal transduction in cell membranes. Although they do not prove the raft-hypothesis of signal transduction in cells, these experiments provide the first experimental evidence that fluid lipid bilayers with physiological lipid compositions can transmit signals across the mid-plane of membranes by inducing new lipid phases on the other side.
Protein-lipid interactions in supported bilayers
Protein-lipid interactions and particularly peptide-lipid interactions have been studied in supported bilayers by attenuated total reflection (ATR) FTIR spectroscopy. A slightly dated, but still valid comprehensive review on this method applied to supported bilayers has been published (20). Because IR light probes the vibrational properties of different classes of covalent bonds, this method is useful to examine lipids, peptides, and interactions between the two in the same sample. The most common parameter for assessing lipid structure and order is to study the stretching vibrations of the lipid acyl chains, for example as a function of peptide concentration or temperature. Such studies have lead to the conclusion that fusion peptides from viruses increase the lipid chain order of fluid phase bilayers and that this property correlates with the biological activity of the viral peptide sequences (51). Another prominent band in the FTIR spectra of lipids develops from the ester carbonyl vibration. This band is sensitive to the hydrogen-bonded structure and thus hydration properties of the bilayer interface. The carbonyl ester band has been shown to change during interaction with some fusion peptides (52). Lipids also have an effect on peptide structure. For example, influenza and HIV fusion peptides are induced to form a-helices in bilayers at low concentration, but they transition into β-sheets at higher concentration (53, 54). Such structural transitions are readily followed by monitoring the amide I band of the peptide bonds, which is highly sensitive to peptide conformation and secondary structure.
Phospholipases bound to supported bilayers hydrolyze distinct paths through the bilayer substrate as observed by AFM (55). The membrane-embedded portions of integral membrane proteins are protected from amide hydrogen exchange, which can be measured by a shift in the amide I and II IR bands when the system is changed from an H2O to a D2O buffer (56). Binding of proteins to supported bilayers may be studied quantitatively by total internal reflection fluorescence microscopy (TIRFM), as shown, for example, in binding studies of antibodies to lipid haptens (57, 58). Fast kinetics of antibody fragment binding and unbinding to supported bilayers have been studied in detail by combining fluorescence correlation spectroscopy with TIRFM (59).
Another aspect of lipid-protein interaction that is conveniently studied in supported bilayers is the lateral diffusion of proteins and lipids and their influence on each other. The regulatory lipid phosphatidyl-inositol-biphosphate (PIP2) slows the diffusion of syntaxin in supported bilayers (37). Conversely, increasing syntaxin concentrations decrease the diffusion of PIP2 and to a lesser extent that of phosphatidylserine. In another system, in which the transmembrane domain of the fibroblast growth factor receptor was incorporated into supported bilayers, lipid and protein diffusion were measured (60). Although protein diffusion was slow (0.006 μm2/s), lipid diffusion was fast (2.6 μm2/s).
Protein-protein interactions in supported bilayers
Much current interest exists in lateral interactions between two or more integral membrane proteins. Several membrane-bound receptors are activated by such interactions during ligand binding. In addition, the interaction between individual transmembrane helices is key to membrane protein folding and has thus received a lot of attention. Although these kinds of interactions are more often studied in nonsupported model membranes, the Hristova group has recently used supported bilayer formats to examine helix-helix interactions between the transmembrane domains of the human fibroblast growth factor receptor (21). Fluorescence resonance energy transfer measurements of fluorescein- and rhodamine-labeled peptides in supported bilayers show that these transmembrane domains dimerize with a sequence-specific dimerization energy of ~3.6 kcal/mol. Meaningful measurements of such interaction energies require that the proteins are laterally mobile in the supported membranes, which was verified in this system (60).
Membrane fusion with supported bilayers
Membrane fusion is a key process in cell biology that takes center stage in membrane biogenesis, fertilization, virus entry, and other events. Our group began studying membrane fusion in supported bilayers 14 years ago. Influenza virus hemagglutinin (HA) (i.e., the protein that mediates virus entry into cells by membrane fusion) was reconstituted into supported bilayers. The physiological pH-dependent fusion of liposomes to the planar HA-containing membranes was demonstrated (23). Several fusion-related structural transformations of HA were recorded by FTIR spectroscopy (61, 62). The fusion of single virions to supported bilayers has also been reported recently (63).
Recently, SNARE-mediated fusion (i.e., the process that leads to neurotransmitter release in synaptic transmission) has been reconstituted in a supported bilayer format in four different laboratories including our own. The general design of these experiments is illustrated in Fig. 2. Fix et al. (24) observed that 15% of the docked vesicles fused within 15 seconds, which yielded a fusion rate of ~7 x 10-5 s-1 in POPC bilayers at a coexpressed syntaxin1/SNAP25 protein/lipid concentration of 1:300. The fusion probability increased 40-fold after Ca2+ addition. Bowen et al. (64) observed thermally induced fusion with syntaxin (protein/lipid ratio 1:14,000) in lipid bilayers composed of eggPC and brainPS. In all, 5-15% of docked vesicles fused with a rate of 0.07 s-1 within 120 sec after triggering. Liu et al. (65) showed that 77% of the reconstituted synaptobrevin vesicles fused within 100 ms after docking (rate, 40 s-1) to syntaxin or syntaxin1/SNAP25 complexes in POPC/DOPS bilayers at a protein/lipid ratio of 1:30,000. The results of these three groups are different from each other, and each of these experiments had several problems, some of which may be caused by the immobile reconstitution of the SNARE proteins in the supported bilayers (66). The independence of fusion on the presence of SNAP25 in the reconstitutions of Bowen and Liu is puzzling because it contradicts in vivo as well as in vitro fusion results. Preliminary work from our laboratory is in much better agreement with the biological literature on this process.
Figure 2. Studying membrane fusion with supported bilayers. A supported bilayer is suspended from a quartz substrate (top, gray background) and illuminated by the evanescent wave of a totally internally reflected laser beam (angled cylinders red). A membrane vesicle is observed to approach, hemifuse, and then fully fuse with the supported membrane. Vesicle contents, lipids, or proteins may be labeled fluorescently to monitor this process.
Cell adhesion and signaling in supported bilayers
Supported bilayers have been used since their inception as surrogate cell surfaces to stimulate immune cells in artificially created immunological synapses. For example, Brian and McConnell (67) have reconstituted major histocompatibility complex (MHC) proteins into supported bilayers and used these planar membranes to stimulate cytotoxic T-cells via specific T-cell receptor interaction. Antigen presentation and signaling through the T-cell receptor have been studied in this system in several follow-up papers as summarized by Watts and McConnell (68). The approach of these early studies (and several others that followed) has experienced a recent renaissance because of improved imaging technologies and because we have learned in the interim how to manipulate supported membranes to form specific controllable spatial patterns (see the next section). These newer approaches are exciting because they have allowed investigators to manipulate spatial patterns in the immunological synapse and thereby ask biologically important new questions with regard to the mechanism of cell-induced cell signaling. Mossman et al. (30). have used laterally constrained bilayers to induce novel patterns of T-cell receptor and intracellular adhesion molecule (ICAM) distribution in MHC-stimulated cells. The authors used a GPI-linked version of the MHC in the supported bilayers to circumvent problems of lateral mobility of the native integral membrane protein in un-cushioned bilayers. Wu et al. (69) demonstrated the compart- mentalization of IgE-receptors in rat basophilic leukemia cells when they stimulated these cells with lipid-hapten bound IgE in nano-fabricated bilayer patches.
Adhesion of different immune cells to one another or to epithelial cells has also been studied using planar bilayer models. For example, lymphocyte function-associated protein-1 (LFA-1) promotes cell adhesion in inflammation [i.e., a reaction that can be mimicked by binding to purified ICAM-1 in supported membranes (70)]. Similarly, purified LFA-3 reconstituted into supported bilayers mediates efficient CD2-dependent adhesion and differentiation of lymphoblasts (71). This work was followed by a study in which transmembrane domain-anchored and GPI-anchored isoforms of LFA-3 were compared (72). Because this research occurred before the introduction of polymer cushions and because the bilayers were formed by the simple vesicle fusion technique, the transmembrane domain isoform was immobile, whereas the GPI isoform was partially mobile. By comparing results with these two isoforms at different protein densities in the supported bilayer, the authors showed that diffusible proteins at a sufficient minimal density in the supported membrane were required to form strong cell adhesion contacts in this system.
In another study, endothelial cells were bound via their integrin receptors to supported bilayers that presented the RGD-peptide, which is the classic integrin ligand (73). The cells spread on RGD presenting membranes but not on control membranes that lacked this peptide. In a similar approach, a laminin-derived peptide was presented on supported bilayers and shown to mediate the spreading and partial differentiation of neuronal subventricular zone progenitor cells (74). The authors observed a strong nonlinear relationship between surface concentrations of the peptides and conclude that this approach may provide novel conditions for growing stem cells with only a limited and controlled amount of differentiation induction.
Patterning supported bilayers
To use supported bilayers as platforms for screening assays, different approaches have been taken to pattern the membrane on the surface. One goal is to observe the membrane interaction of substances simultaneously with supported bilayers of different composition. Bilayers can be subdivided into different areas by diffusion barriers erected on the substrate or by blotting patches of membrane with polydimethylsiloxane (PDMS) stamps (75). A combination of these techniques may be used to form patterned membranes of different composition. Barriers may be formed from metal or metal oxides by standard lithographic techniques or by stamping proteins onto the substrate (76). Originally, homogeneous supported membranes have been patterned photolithographically using deep-UV light (77). Using photolithography in combination with a polymer lift-off technique, patterned lipid bilayers with patches measuring 1 to 76 pm may be formed (78). Another approach uses the flow of vesicle suspensions within microfluidic devices to deposit membranes of different compositions at the same time (75). Supported membranes of different compositions on the same substrate are conveniently addressed with different solutions of analytes flowed through micro-channels in PDMS (79).
Using supported bilayers as platforms for chemical sensing and analysis
Because of their planar geometry, supported bilayers are predestined for biological and chemical sensor applications. The basic concept is to couple the high specificity and sensitivity of molecular membrane receptors to substrates that integrate opto-electronic circuits (25-27). In combination with the described patterning techniques, supported membranes should provide a nearly ideal physiological environment for high-throughput sensing with biological membrane receptors or channels in chip-based arrays. However, despite their great promise, economically viable commercial applications have so far not yet been used because of a series of obstacles that still must be overcome. Nanoscopic and microscopic defects in fluid supported lipid bilayers most often lead to low electrical resistance and therefore make them unsuitable for high-sensitivity electronic detection. In some approaches, this problem has been overcome by linking the lipid bilayer with very short and dense tethers to the substrate. However, doing so prevents the incorporation of larger membrane receptors and channels.
Another approach is to prepare black lipid membranes with solvent over microscopic or nanoscopic holes in the substrate (80-82), or by using giant vesicles that adhere to the substrate (83). Although high-resistance seals have been obtained, long-term stability has not yet been achieved with these systems. In cases where highly insulating membranes are established, the conductivity of ligand-gated ion channels or larger pores may be recorded (25, 84). Alternatively, the conductivity may be probed by measuring the impedance (26) or with metal-free field-effect transistors that have the advantage of avoiding electrochemical perturbations (83). Optical detection is suitable for applications such as immunoassays when the molecules of interest can be addressed by fluorescent- or gold-labeled antibodies (79). As is true for electronic biosensors, optical biosensors designed for routine practical applications must be robust and stable for a long time. Lipopolymers that stabilize supported bilayers against air exposure are also helpful in this case (85).
Procedures to Prepare Supported Bilayers
Supported lipid bilayers with or without reconstituted membrane proteins are prepared by one of three methods described below and illustrated in Fig. 3. Methods details can be found in Reference 86.
Figure 3. Methods for supported bilayer formation and membrane protein reconstitution. (a) and (b) LB/LS method. A lipid monolayer is spread at the air-water interface of a Langmuir trough and transferred to a solid substrate while keeping the surface pressure constant. A second monolayer is transferred by horizontal apposition of the first transferred monolayer and collection of a counter-piece with spacers. (c) Direct VF method. Membrane vesicles are flown into a chamber with a clean surface substrate on top. After about an hour of incubation, they form a supported bilayer on the substrate and excess vesicles are flushed out. (d) LB/VF method. The procedures depicted in panels (a) and (c) are combined leading to an asymmetric bilayer with an asymmetric protein distribution. Although this method can also be performed without a polymer, it is shown here with the polymer transferred during the LB step.
Langmuir-Blodgett/Langmuir-Schafer (LB/LS) technique (4, 5)
This technique is historically the first method to prepare supported bilayers. A lipid monolayer is spread from a desired lipid solution in organic solvent onto a pure water surface in a Langmuir trough. After evaporation of the solvent, the monolayer is compressed slowly to reach a surface pressure of 32 mN/m (thought to be the equivalence pressure of a bilayer) and equilibrated. A carefully cleaned hydrophilic substrate (glass, quartz, oxidized silicon, etc.) is then rapidly submerged into the trough and slowly withdrawn with a dipper mechanism while a constant surface pressure is maintained. This step transfers a single monolayer of lipids known as the LB layer onto the substrate. A second monolayer known as the LS layer is then spread and compressed on the trough in the same fashion. The LB-coated substrate is attached to a suctioning tip, and its face is gently lowered to contact the LS monolayer at the air/water interface for a few seconds. To complete the bilayer, the slide is then pushed through the interface and placed on a cover slip fitted with two spacers of water-resistant double-sided tape that had been previously placed at the bottom of the trough. After removal of the supported bilayer sandwich from the trough, the water between the surfaces may be exchanged by flow-through with any desired buffer while always maintaining full hydration of the bilayer. Some investigators have prepared peptide-containing supported bilayers by placing peptides in the first or second monolayer.
Vesicle fusion (VF) technique (67)
This technique is the simplest method for forming supported bilayers. Much literature is available on the mechanism and kinetics of vesicle spreading on hydrophilic substrates, which is not reviewed here. Although it is simple, the method tends to result in bilayers with more defects, and the orientation of membrane proteins cannot be controlled in this method. Small or large unilamellar pure lipid vesicles or proteoliposomes are prepared by standard liposome preparation or membrane protein reconstitution methods. A clean hydrophilic substrate is placed in a flow-through chamber, and the vesicles are injected and incubated with the surface for 30-60 minutes. Excess vesicles are washed out by extensive rinsing with a buffer. This method has also been used to make polymer-supported bilayers, in which case the surface of the support is pre-treated with the polymer using conditions that depend on the particular polymer being used.
Langmuir-Blodgett/vesicle fusion (LB/VF) technique (87)
This method is a combination of the other two methods. In our opinion, it is the most gentle method to reconstitute membrane proteins into supported bilayers and to prepare supported bilayers with fragile coexisting liquid phases of lipids. A LB monolayer is prepared on a hydrophilic substrate as described above. To prepare tethered-polymer supported bilayers, a suitable lipopolymer may be included at a concentration of a few mol % at this stage. With some lipopolymers, it is necessary to cure the slide (by light, temperature, zero humidity, etc.) at this stage. The monolayer-coated slide is then placed in a custom-built flow-through chamber, and vesicles or proteoliposomes are injected and incubated with the surface for 30-60 minutes (pure lipid vesicles) or 60-120 minutes (proteoliposomes). Excess vesicles are washed out by extensive rinsing with buffer. Because the second monolayer is completely vesicle derived and because membrane proteins are introduced as proteoliposomes only in the second step, they tend to be unidirectionally oriented in the supported bilayer as can be verified with quenching antifluorophore antibodies (23, 37) or by FLIC microscopy (33).
Procedures to Characterize Supported Bilayers
The simplest way to examine the quality and integrity of supported membranes is to include a fluorescent lipid probe such as nitrobenzoxa-diazol (NBD)-PE or rhodamine-PE and to look for their uniform appearance on a standard epifluorescence microscope. Many artifacts can be detected readily and eliminated with this very simple test. Higher-resolution images can be obtained by AFM or NSOM. These techniques are useful to detect small defects in the 10 to 500 nm range that might escape detection by standard wide-field optical microscopy. Because supported bilayers are only stable under water, these imaging modalities must be carried out under water. AFM probes height profiles, but NSOM and epifluorescence microscopy permit the labeling of specific chemical structures or physical properties of the structures by using different fluorescent probes. It is often necessary to discriminate surface from bulk fluorescence, for example when measuring the binding of fluorescent ligands to supported membranes. This result is conveniently achieved with TIRFM, which has a typical 1/e illumination depth of 50 to 100 nm from the surface, which depends on refractive indices, angle of incidence, and wavelength of light. Brewster angle and surface plasmon microscopies are also surface-selective optical imaging techniques, which do not require the fluorescent labeling of the membrane, but rather depend on lateral changes of refractive index in the sample.
A next and very important level of characterization of supported bilayers is the measurement of the lateral mobility of their constituents. Because biological membranes and their proper function are defined by their fluidity, to recreate this characteristic is imperative for biology-motivated work. In addition, lateral diffusion measurements on supported bilayers can easily detect many artifacts that may go unnoted by simple microscopic inspection. For example, deposited membranes may look completely uniform but may not show any long-range lateral diffusion when vesicles or membrane fragments that are smaller than the resolution of the light microscope are densely packed on the substrate surface. Two techniques are common to determine the diffusion of fluorescently labeled lipids or proteins in supported bilayers: FRAP and SPT. The lateral diffusion coefficient and fraction of mobile molecules are obtained from either measurement.
In FRAP, a brief pulse of intense laser light is used to photobleach fluorophores partially in a small area of the sample. The recovery of fluorescence caused by diffusion of labeled molecules into the bleached area is then observed over time, while care is taken to minimize additional photobleaching. In spot photobleaching the light is focused to a circular spot, which reflects the Gaussian beam profile of the laser. During recovery, the half-width of the bleached area decreases whereas the intensity increases. The diffusion coefficient and mobile fraction are extracted from the time course and the amplitude, respectively, of the recorded recovery curve (88). In a variant called periodic pattern photobleaching, the bleach pulse projects a stripe pattern of a Ronchi ruling onto the sample, which permits integration over a larger area and therefore an increased signal/noise (89).
SPT is the preferred technique when more detail on different populations of moving particles in a heterogeneous system is required. Although the information content of SPT is much higher than that of FRAP, it is much more demanding on instrumentation and statistical evaluation procedures. SPT has been introduced to the characterization of supported bilayers in 1995 (90) and has been used frequently since then. In practice, the technique is best used in combination with TIRF microscopy and high-sensitivity charge-coupled devices. Typical labeling ratios of the lipid bilayer are 1:108fluorescent probes:lipids. The reconstructed particle trajectories and appropriate statistics can be analyzed to distinguish between diffusion, anomalous diffusion, confined diffusion, and directed motions.
Neutron reflectivity has been used to characterize the transverse organization of supported bilayers structurally (32, 91). The lateral structure of lipid bilayers on solid supports may also be characterized by grazing incidence X-ray diffraction, although this technique has so far been mainly used on monolayers at the air-water interface. Vibrational spectroscopies open interesting windows to look at details of lipid structure in supported bilayers. FTIR spectroscopy has several bands that are characteristic of the state of lipid order and hydration in supported bilayers (20). An interesting relatively recent method to study the organization of supported bilayers, particularly with respect to their asymmetry, is sum frequency vibrational spectroscopy (SFVS) (92). Signals develop in this nonlinear form of vibrational spectroscopy only when symmetry is broken (i.e., when the type, structure, and number of lipids is unequal across the mid-plane of the supported bilayer). A complementary technique to study lipid asymmetry, or more specifically fluorescent probe asymmetry, is FLIC microscopy, in which the average distance of fluorophores from an oxidized silicon mirror surface is measured interferometrically (18).
Protein secondary structure and orientation
Two methods have been used to determine the secondary structure and orientation of membrane proteins in supported bilayers: polarized ATR-FTIR spectroscopy and oriented CD spectroscopy. SFVS may also be applied to study peptide and protein structures in supported bilayers. Polarized ATR-FTIR spectroscopy is sensitive enough that high-quality spectra can be obtained from a single bilayer. Beta-sheet structures are readily distinguished from α-helical and random structures, and the orientations of α-helices are determined from the linear dichroism of the peptide amide I bands (20). Multiple stacks of supported bilayers have to be used to gain enough sensitivity to determine the structure and orientation of α-helices in lipid bilayers by oriented CD spectroscopy (60, 93).
Conductance and impedance
Pure lipid membranes are electrical insulators with a specific capacitance of ~1 μF/cm2, which separate two electrolytic compartments. The conductance of biological membranes is mainly determined by highly specialized proteins that act as ion channels. For supported membranes to mimic the electrical properties of a biological membrane, it is necessary to measure its electrical characteristics. Even very small defects that are not visible by microscopy increase the conductance significantly. If the membrane is supported by a conductive substrate like gold or indium tin oxide, then it is possible to measure the impedance by applying A/C voltages with frequencies up to 100 kHz and by measuring the magnitude and phase of the current. The conductance and capacitance of the supported membrane is determined from the evaluation of the entire electrical circuit, which includes resistance and capacitance of the electrode and if necessary alternative electrical paths around the membrane (94). A similar approach uses metal-free field effect transistors that probe the electrical potential at the transistor gates that face the cleft beneath the supported membrane. The electrical membrane parameters are extracted from the voltage transfer from the electrolyte bath to the transistor gates (83).
In conclusion, supported bilayers have evolved into a reliable model membrane system since their first inception almost a quarter century ago. Numerous basic research questions regarding the structure and function of biological membranes and applications that range from biosensing to proteomic analyses of membrane components have been addressed with this system. We anticipate more growth and an even more prominent role of this tool in basic and applied membrane research in the decades to come.
The work in our laboratory was supported by grants from the National Institutes of Health.
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Membranes, Fluidity of
Membrane Proteins, Properties of
Membrane Fusion, Mechanisms of
Lipid Bilayers, Properties of