INTRODUCTION
Cell membranes define the size and shape of the cell. In addition to this structural role, the membrane has a crucial regulatory role determining what information, nutrients, and waste can permeate this barrier. The cell membrane consists of a lipid bilayer and proteins, which can be either transmembrane or associated with one leaflet of the bilayer. The dynamic organization of proteins and lipids into domains (e.g., rafts) within the bilayer is important for multiple cellular processes, such as recognition and signaling events.
The chemical recognition process occurring on the surface of membranes is the basis of a versatile and specific sensor system for the cell. Lipid bilayer systems that mimic certain aspects of cell membrane function have been employed in biosensor schemes and continue to generate great interest in the nanotechnology field. Chemical recognition events can also cause structures to form, providing a mechanism for creating controllable, dynamic nanoscale architectures. The two main platforms for studying the dynamic properties of membranes for both nanotechnology and nanoscience applications are vesicular structures, called liposomes, and supported lipid bilayers.
This article aims to illustrate the importance of dynamic nanoscale structures in biological and model biological membranes. Applications of such structures for drug screening, biosensors, and microanalysis will be discussed. The emphasis will be on understanding what triggers structural reorganization on the nanoscale and how the temporal and spatial aspects of such reorganization can be controlled. The sophistication of the nanoscale machinery of the cell membrane offers many lessons that can be applied to the emerging field of nanotechnology.
LIPIDS, LIPOSOMES, AND SUPPORTED BILAYERS
Lipids are amphiphilic molecules, having a hydrophobic tail and a polar head group (Fig. 1A). Most commonly the tail consists of two fatty acid chains with an even number of carbon atoms (14-18 atoms long) and with various degrees of unsaturation.[1] In biological membranes, the key components are phospholipids, which have a phosphate at the head group position that is connected to the hydrophobic tails through a glycerol backbone. Often, another moiety, such as choline, serine, or inositol, is attached to the phosphate to form PC, PS, or Pi-type lipids, respectively.1-2-1
In an aqueous solution, these amphiphilic lipids self-assemble into liposomes (Fig. 1B).[2] The spherical bilayer structure minimizes unfavorable interactions of the hydrophobic tail region with the water. The head group and tails determine the properties of the bilayer membrane, such as fluidity, charge density, and permeability. As recognized by Singer and Nicolson,[3] the membrane is a fluid mosaic of lipids and proteins. This fluid-mosaic model does not, however, preclude the existence of structured regions (domains or rafts) within the lipid bilayer. In fact, it is the membrane’s fluidity that enables the creation of dynamic nanoscale structures. The lipids can exist in three distinct phases: a tightly packed, ordered gel phase, an intermediate liquid ordered phase, and a disordered liquid phase. The transition temperature for gel to liquid phase transitions (Tg) decreases with decreasing chain length and degree of unsaturation. The steric hindrances, electrostatic charge, and hydrogen bonding of the head groups can also affect the transition temperature as well as the phase separation within the membrane. Such phase-separated domains (e.g., rafts) can undergo compositional fluctuations.1-4-1 It is key to recognize that under physiological conditions, the membrane is a heterogeneous,[4] nonequilibrium[5] system.
As previously mentioned, liposomes and supported bilayers are the two main systems of interest. Supported lipid bilayers can be prepared by the classic Langmuir-Blodgett method. Alternately, liposomes can be fused with surfaces to form supported bilayers.[6] The fusion schemes for hydrophilic surfaces outlined in Fig. 2A and C result in opposite orientations of membrane faces and any incorporated proteins. In Fig. 2C, the orientation of the leaflets toward the bulk aqueous solution is preserved upon fusion. In the mechanism detailed in Fig. 2A, however, the orientation of the leaflets is reversed; that is,the inside leaflet of the liposome becomes the top leaflet of the supported bilayer. Vesicle fusion to hydrophobic surfaces (Fig. 2B) can also be accomplished but the liposomes must rupture resulting in attachment and spreading of the two leaflets of the lipid bilayer. Liposome composition, surface chemistry, vesicle size, temperature, osmotic pressure, and the presence of calcium ions are all factors that influence vesicle fusion.[7,8] Typically, supported bilayers formed by vesicle fusion maintain a thin layer of water 10 A) between the substrate and the adjacent membrane surface.’9-11-1 This supported bilayer structure is very stable (days to months) in an aqueous environment, but unstable in the presence of detergents or in air. The lateral mobility of lipids within the bilayer is maintained enabling the molecules in the membrane to diffuse over long distances.
Fig. 1 (A) Cartoon of a lipid molecule showing the hydrophobic tail region and polar head group and the chemical structure for a typical lipid (1,2-Distearoyl-sn-Glycero-3-Phospho-choline, DSPC). (B) The liposome (or vesicle) that spontaneously forms when lipids are placed in aqueous solutions.
Fluorescence microscopy provides a means to attain real-time data on the dynamical structures occurring in lipid bilayer systems. Single-molecule sensitivity is achievable. Fluorescence resonance energy transfer (FRET) between a fluorescently tagged donor component and a fluorescently tagged acceptor component yields accurate distance information for short length scales. In fluorescence recovery after photobleaching (FRAP), an area on the bilayer surface is photobleached and then monitored for recovery of fluorescence intensity. The fluorescence recovery is due to fluorescently labeled molecules diffusing into the bleached area and can thus be used to measure diffusion rates to evaluate lateral fluidity.
Spatial resolution of fluorescence techniques are, however, diffraction limited to a few hundred nanometers. Unfortunately, this length is often the same size as the dynamic structures. The atomic force microscope (AFM), on the other hand, can be used to characterize features in supported bilayers with subnanometer resolution.1-12-1 The AFM typically requires about a minute to capture an image. Thus it is suitable for imaging relatively static systems, or slow dynamics. Fluorescence and AFM imaging are the two techniques most relevant to the work discussed in this article, but other methods, including neutron scattering and nuclear magnetic resonance (NMR) spectroscopy,[13- have also been applied toward dynamic nanostructures in bilayer systems.
Computer simulations are also an effective tool for understanding dynamic nanostructures in bilayer membranes.'141 Nielsen and coworkers simulated binary mixtures of lipid bilayers revealing dynamic microphase separations with length scales of tens of nanometers.'4-They further showed that this nanoscale structure affects the functional properties of the membrane. Through simulations and theoretical modeling, Gil et al. analyzed protein organization in lipid bilayers.[5] The lateral organization of transmembrane proteins can be explained by the properties of the lipid bilayer. Hydrophobic matching between lipids and the hydrophobic region of proteins can lead to an enrichment of one lipid species near the protein. As such, the lipids can mediate protein attraction or repulsion. Also, the wetting of a protein by one lipid component can lead to larger protein organization patterns.
DYNAMIC NANOSTRUCTURES IN BIOLOGICAL AND MODEL BIOLOGICAL MEMBRANES
In this section, a few examples will be given to illustrate the importance of dynamic nanostructures in biological systems. Rafts, domains, and hierarchical structures are prevalent in the cell membrane and are the predominant sites of biological activity. Subczynski and Kusumi[15] reviewed the three types of rafts (Fig. 3) found, to date, in plasma membranes. In unstimulated cells there are small, unstable (lifetimes of less than 1 msec) lipid rafts (a, a’, and a" in Fig. 3) that may have associated proteins. When receptor molecules in these unstable rafts react with ligands, they can create stabilized rafts (Fig. 3b) with lifetimes of minutes. The coalescence of these two types of rafts creates transient confinement zones (TCZs), which serve as signaling rafts (Fig. 3c) by assembling the necessary constituents to switch on a downstream signaling pathway.
Fig. 2 Schematic representation of the possible mechanisms for planar bilayer formation from liposomes on hydrophilic and hydrophobic surfaces. The drawings are not drawn to scale. Lipid molecules on the support are enlarged approximately 50-fold compared to a liposome. (A) Vesicle fusion on a hydrophilic surface with the leaflet orientation reversed. (B) Vesicle fusion on a hydrophobic surface. (C) Vesicle fusion on a hydrophilic surface with the leaflet orientation preserved.
Such dynamic nanostructures have been shown to be important in a variety of biological systems. Sheets et al. explain how the organization of the plasma membrane likely exerts spatio-temporal control on immunoglobulin E receptor-mediated signal transduction.1-16-1 Lipid rafts also play a key role in the immunological synapse.[17-19] The T-cell antigen receptors (TCR) are located in rafts and they become cross-linked by ligand binding. The cross-linking induces raft aggregation, causing colocalization of signaling proteins. This activates the phosphorylation of tyrosine residues on membrane-associated proteins and starts downstream signaling.
Because the biological membrane is a complex entity, most structure and function studies are performed on simpler, well-defined model membrane systems of lipo-somes and supported bilayers. For example, in studies of lipid raft formation, ternary mixtures of saturated lipids, unsaturated lipids, and cholesterol have been found to spontaneously form rafts over a wide range of specific lipid species and concentrations.1-20-22-1 In these studies, micron-sized domains were imaged via fluorescence microscopy techniques. Nanostructural features in lipid bilayers can also be revealed with AFM, although the domain shape and size have been shown to be somewhat dependent on the substrate and bottom leaflet of the supported bilayer.[23,24]
Not only do lipid-lipid interactions cause domain formation but lipid-protein interactions can also induce nanoscale structures to form. Rinia et al. reported that transmembrane WALP proteins perturb the bilayer, creating striated domains of 25 nm to 10 mm with the nanoscale striations spaced at 7.5-nm intervals.[25] Furthermore, the specific physical properties of the bilayer components can modulate enzyme activity. Honger and coworkers demonstrated that phospholipase A2 enzyme activity correlates with the degree of microheterogeneity within the bilayer.[26]
Fig. 3 Three types of rafts found thus far in the plasma membrane. The first type (a) is prevalent in the absence of extracellular stimulation. They are small (perhaps consisting of several molecules) and unstable (the lifetimes may be less than 1 msec) and may be the kind of raft that monomeric GPI-anchored proteins associate with. The second type of raft (b) may appear when receptor molecules form oligomers upon liganding or cross-linking. The receptors may be GPI-anchored receptors or transmembrane receptors with some affinity to cholesterol and saturated alkyl chains. Oligomerized receptors may then induce small but stable rafts around them, perhaps due to the slight reduction in the thermal motion around the cluster and the subsequent assembly of cholesterol. Given the rather stable oligo-merization of the receptor molecules, the second type of raft may be stable for minutes, although the associated raft-constituent molecules may be exchanged frequently between the raft and the bulk domains. Such receptor-associated rafts are called ”core-receptor rafts.” The third type of raft (c) may be formed around these core receptor rafts (although the core receptor rafts may be undergoing diffusion). Here they are called ”signaling rafts,” because they are likely to be directly involved in downstream signaling from the receptor molecules, by assembling signaling molecules through the (transient and/or more stable) coalescence of rafts that may contain one or two signaling molecules. Small/unstable rafts are also likely to exist in the inner leaflet of the membrane (a’) and could coalesce with the core receptor rafts (b and c), where the signaling molecule in the inner leaflet is activated, which might also leave from the signaling rafts (c and a").
Protein binding to receptor lipids in model membranes can also induce a membrane reorganizational process.[27] Concanavalin A (Con A) protein was found to bind to bilayers composed of the mannosamine-functionalized lipid PSMU and distearylphosphatidylcholine (DSPC).
Initially, PSMU forms aggregates in DSPC, but slowly disperses following Con A adsorption to the membrane surface and binding to the mannosamine head groups. Dispersal is attributed to Con A-Con A steric interactions, distance between receptor sites, and possible protein insertion events.
Another example of a molecular recognition-induced lipid reorganization was developed by Song and co-workers to detect biotoxins.[28,29] Fluorescently tagged receptors are dispersed in a fluid lipid matrix of palmitoyl, 9-octadecenenoyl phosphatidylcholine (POPC). The dispersed receptors exhibit strong fluorescence. Specific, multivalent binding of the toxin to its receptors brings the fluorophores in close proximity, causing a strong decrease in fluorescence intensity due to self-quenching. Nonspecific binding of toxins to lipid bilayers can also be used in a biosensing scheme that employs optical evanescence.’30-Pattern analysis and comparison to standards are required to assign the identity of the bound toxin, but the method is rapid and can be used for multiple toxins. The original work analyzed the binding of six different protein toxins.
