Nanoscale Imaging of Domains in Supported Lipid Membranes

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InVited Feature Article Nanoscale Imaging of Domains in Supported Lipid Membranes Linda J. Johnston* Steacie Institute for Molecular Sciences, National Research Council of Canada, 100 Sussex DriVe, Ottawa, Ontario, Canada K1A 0R6 ReceiVed January 14, 2007. In Final Form: February 27, 2007 The formation of domains in supported lipid membranes has been studied extensively as a model for the 2D organization of cell membranes. The compartmentalization of biological membranes to give domains such as cholesterolrich rafts plays an important role in many biological processes. This article summarizes experiments from the author’s laboratory in which a combination of atomic force microscopy and near-field scanning optical microscopy is used to probe phase separation in supported monolayers and bilayers as models for membrane rafts. These techniques are used to study binary and ternary lipid mixtures that have gel-phase or liquid-ordered domains that vary in size from tens of nanometers to tens of micrometers, surrounded by a fluid-disordered membrane. Examples are presented in which these models are used to investigate the distribution of glycolipid membrane raft markers and the preference for peptide and protein localization in ordered versus fluid membrane phases. Finally, the enzyme-mediated restructuring of membranes containing liquid-ordered domains provides an in vitro model for the coalescence of membrane rafts to give signaling platforms. Overall, the results demonstrate the importance of using techniques that can probe the nanoscale organization of membranes and of combining techniques that yield complementary information. Furthermore, the ability of supported lipid bilayers to model some aspects of membrane compartmentalization provides an important approach to understanding natural membranes.

Introduction The compartmentalization of cell membranes plays an important role in regulating a wide range of biological processes, including signal transduction, cell adhesion, lipid trafficking, and viral and bacterial entry. Among the many types of membrane compartments, cholesterol-rich microdomains known as lipid rafts have attracted substantial attention over the past decade.1-6 Following much discussion and controversy over their properties and even their very existence, the following consensus definition is emerging. Membrane rafts are small (10-200 nm), heterogeneous, dynamic sterol- and sphingolipid-enriched domains that compartmentalize cellular processes.7 Rafts are generally believed to exist in a liquid-ordered phase with properties that are distinct from those of the surrounding fluid-disordered membrane. Both lipids and proteins contribute to the formation of these membrane domains, and their functional role in modulating cellular processes via both the spatial and temporal compartmentalization of membrane components is an important part of the raft definition. Although some of the original evidence that led to the raft hypothesis came from the isolation of detergent-resistant membrane fractions, it is now recognized that this may actually induce domain formation.8 During the past decade, many biophysical techniques have been used to attempt to detect, identify, and characterize rafts in cellular membranes.9 However, * E-mail: [email protected]. (1) Simons, K.; Ikonen, E. Nature 1997, 387, 569-572. (2) Simons, K.; Vaz, W. L. Annu. ReV. Biophys. Biomol. Struct. 2004, 33, 269-295. (3) Edidin, M. Annu. ReV. Biophys. Biomol. Struct. 2003, 32, 257-283. (4) Brown, D. A.; London, E. J. Biol. Chem. 2000, 275, 17221-17224. (5) Anderson, R. G. W.; Jacobson, K. Science 2002, 296, 1821-1825. (6) Munro, S. Cell 2003, 115, 377-388. (7) Pike, L. J. J. Lipid Res. 2006, 47, 1597-1598. (8) Lichtenberg, D.; Goni, F. M.; Heerklotz, H. Trends Biochem. Sci. 2005, 30, 430-436.

progress in this area has been hampered by the small size and transient nature of raft domains in cellular membranes. The difficulty of visualizing small, dynamic lipid domains in the complex environment of a cellular membrane has motivated studies in simpler model systems such as phospholipid monolayers, vesicles, and supported bilayers.2,3,10-12 Although an artificial membrane containing only lipids cannot duplicate the functional aspect of membrane rafts, it does provide a tractable system for understanding how molecular associations control lipid domain formation and is compatible with studying the interaction of specific peptides or proteins with lipid domains. Fluorescence microscopy has been widely employed for the direct visualization of domains and membrane morphology for a variety of model membranes and has convincingly demonstrated the presence of coexisting fluid and liquid-ordered or gel phases in both giant vesicles and supported membranes.2,3,10,11 Despite the wealth of useful information provided by these studies, it is increasingly clear that many model membranes have nanodomains that are not detectable with diffraction-limited techniques. Atomic force microscopy (AFM) of supported phospholipid monolayers or bilayers on solid supports provides an alternate approach for investigating phase separation on the nanoscale. The high spatial resolution of AFM (several nanometers in the lateral dimension and approximately 0.1 nm in the vertical direction)13 allows one to distinguish membrane phases on the basis of their difference in thickness and to resolve domains in the 10-200 nm regime, (9) Lagerholm, B. C.; Weinreb, G. E.; Jacobson, K.; Thompson, N. L. Annu. ReV. Phys. Chem. 2005, 56, 309-336. (10) Veatch, S. L.; Keller, S. L. Biochim. Biophys. Acta 2005, 1746, 172-185. (11) Heberle, F. A.; Buboltz, J. T.; Stringer, D.; Feigenson, G. W. Biochim. Biophys. Acta 2005, 1746, 186-192. (12) London, E. Biochim. Biophys. Acta 2005, 1746, 203-220. (13) Atomic Force Microscopy in Cell Biology; Jena, B. P., Ho¨rber, J. K. H., Eds.; Academic Press: San Diego, CA, 2002; Vol. 68.

10.1021/la070108t CCC: $37.00 Published 2007 by the American Chemical Society Published on Web 04/12/2007

Nanoscale Imaging of Domains in Lipid Membranes

as well as the larger micrometer-sized domains observed by fluorescence microscopy. As summarized in several recent reviews, AFM has become the method of choice for studying phase separation in supported monolayers and bilayers.14-17 Although the spatial resolution is comparable to that of electron microscopy, AFM does not require sample labeling, staining, or fixation. More importantly, it is compatible with imaging in a physiologically relevant aqueous environment and allows realtime imaging of the formation or reorganization of membranes. AFM has some limitations for studies of multicomponent samples with complex morphology where the topography alone is insufficient to identify the chemical composition. For some applications, this drawback can be overcome by the use of covalently modified tips that interact with a particular component of the sample (e.g., the use of an antibody-modified tip to identify proteins). However, a more general approach is to combine the sensitivity and specificity of fluorescence with the high spatial resolution of AFM. As a result, hybrid microscopes that allow for simultaneous AFM and confocal or total internal reflection fluorescence imaging are becoming more common;18-20 nevertheless, their optical resolution is limited by diffraction to approximately 300 nm for visible excitation. By contrast, nearfield scanning optical microscopy (NSOM) overcomes the diffraction limit by replacing the AFM tip with a small-aperture optical probe that delivers light to the sample.21,22 The high resolution and the ability to obtain simultaneous optical and topographic images simultaneous offer significant advantages for biological imaging. Our transmission near-field microscope is based on a modified atomic force microscope, allowing the use of bent fiber probes, an approach that minimizes sample damage and facilitates NSOM measurements in an aqueous environment.23-26 Bent optical fiber NSOM probes are fabricated using a combination of two-step chemical etching, aluminum deposition, and focused ion beam milling to fabricate well-defined and reproducible apertures. The probes have transmission efficiencies that are roughly 2 orders of magnitude higher than those of pulled probes of similar aperture size and provide excellent results for fluorescence imaging, with typical resolutions of 50-100 nm for imaging dry samples or those in an aqueous environment. However, the mechanical properties and relatively large size of the NSOM probe significantly limit the scan speed, with 20-30 min required for an individual image, a factor of 5-10 times slower than typical AFM imaging. This article provides an overview of our recent studies of lipid domains in supported monolayers and bilayers. A combination of AFM and fluorescence is used to characterize domains in phase-separated lipid mixtures that model some of the properties of membrane rafts. Similar models are used to investigate the localization of glycolipids, peptides, and proteins in ordered (14) Dufrene, Y. F.; Lee, G. U. Biochim. Biophys. Acta 2000, 1509, 14-41. (15) Connell, S. D.; Smith, A. Mol. Membr. Biol. 2006, 23, 17-28. (16) Milhiet, P. E.; Giocondi, M.-C.; LeGrimellec, C. Sci. World J. 2003, 3, 59-74. (17) Rinia, H. A.; De Kruijff, B. FEBS Lett. 2001, 504, 194-199. (18) Burns, A. R. Langmuir 2003, 19, 8358-8363. (19) Shaw, J. E.; Epand, R. F.; Epand, R. M.; Li, Z.; Bittman, R.; Yip, C. M. Biophys. J. 2006, 90, 2170-2178. (20) Shaw, J. E.; Oreopoulos, J.; Wong, D.; Hsu, J. C. Y.; Yip, C. M. Surf. Interface Anal. 2006, 38, 1459-1471. (21) Lewis, A.; Taha, H.; Strinkovski, A.; Manevitch, A.; Khatchatouriants, R.; Dekheter Ammann, E. Nat. Biotech. 2003, 21, 1378-1386. (22) Dunn, R. C. Chem. ReV. 1999, 99, 2891-2928. (23) Burgos, P.; Lu, Z.; Ianoul, A.; Hnatovsky, C.; Viriot, M.-L.; Johnston, L. J.; Taylor, R. S. J. Microsc. 2003, 211, 37-47. (24) Ianoul, A.; Burgos, P.; Lu, Z.; Taylor, R. S.; Johnston, L. J. Langmuir 2003, 19, 9246-9254. (25) Ianoul, A.; Street, M.; Grant, D.; Pezacki, J.; Taylor, R.; Johnston, L. J. Biophys. J. 2004, 87, 3525-3535. (26) Yuan, C.; Johnston, L. J. J. Microscopy 2002, 205, 136-146.

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Figure 1. Cartoon depictions of lipid bilayers with coexisting gel and fluid-disordered phases (A) and liquid-ordered and fluiddisordered phases (B).

membrane domains and to probe the enzyme-mediated restructuring of supported bilayers.

Domains in Supported Monolayers and Bilayers Many recent studies have visualized phase separation in binary and ternary lipid mixtures as a model for the domains that are believed to exist in cellular membranes.2,3,10-12 For binary mixtures, one typically observes gel-phase domains of a lipid with a high melting temperature surrounded by a fluid phase of a low-melting lipid, usually one with either short or unsaturated acyl chains (Figure 1A). Ternary lipid mixtures comprising a saturated phosphatidylcholine (PC) or sphingomyelin (SM), an unsaturated PC, and cholesterol (Chol) have been shown to exhibit two coexisting fluid phases: a liquid-ordered phase that is rich in Chol and saturated lipid and a liquid-disordered (fluid) phase that is predominantly unsaturated PC (Figure 1B). The liquidordered phase is believed to be analogous to membrane rafts and is characterized by a high degree of ordering of the lipid acyl chains but a degree of lipid mobility that is similar to that of a fluid membrane. The evidence for phase separation comes from a wide range of methods applied to both monolayers and bilayers. Representative examples of our use of AFM and NSOM for the direct visualization of coexisting phases in monolayers and bilayers prepared from binary and ternary lipid mixtures are summarized below. Monolayers. Several studies have shown that SM/DOPC (dioleoylphosphatidylcholine)/Chol monolayers transferred to mica at low surface pressure (10 mN/m) exhibit phase separation to give mixtures of large round or elliptically shaped domains and small nanodomains of a liquid-ordered SM-Chol-rich phase that is slightly thicker than the surrounding DOPC-rich fluid phase.23,27-29 As an example, AFM and NSOM images for a 2:2:1 egg SM/DOPC/Chol monolayer transferred to mica at 10 mN/m are shown in Figure 2A,B.29 The monolayer contains 1% Texas red 1,2-dihexadecanoylphosphoethanolamine (TR-DHPE), a dye-labeled lipid that partitions strongly into the fluid phase, thus providing excellent contrast for fluorescence detection of the (dark) liquid-ordered domains. The ordered domains protrude ∼0.9 nm from the surrounding monolayer, and the small, closely spaced nanodomains can be readily visualized by both AFM and NSOM. Monolayers of this lipid mixture undergo a transition (27) Yuan, C.; Furlong, J.; Burgos, P.; Johnston, L. J. Biophys. J. 2002, 82, 2526-2535. (28) Burgos, P.; Yuan, C.; Viriot, M.-L.; Johnston, L. J. Langmuir 2003, 19, 8002-8009. (29) Coban, O.; Popov, J.; Burger, M.; Vobornik, D.; Johnston, L. J. Biophys. J. 2007, 92, 2842-2853.

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Figure 2. AFM (A, C) and NSOM (B) images showing the evolution from a mixture of micrometer- and nanometer-sized domains at low surface pressure (10 mN/m, A, B) to an interconnected network of nanodomains at high surface pressure (30 mN/m, C) for 2:2:1 egg SM/DOPC/Chol monolayers containing 1% TR-DHPE deposited on mica. Adapted from ref 29.

from a mixture of micro- and nanodomains at low surface pressure (5-10 mN/m) to a network of small interconnected domains at high surface pressure.29 Note that monolayer pressures between 30 and 35 mN/m are typically considered to be appropriate for modeling a cellular membrane.30,31 The evolution in domain size is clearly illustrated by comparison of the AFM image in Figure 2C for a 2:2:1 egg SM/DOPC/Chol monolayer transferred to mica at 30 mN/m with that shown in Figure 2A for a similar monolayer deposited at 10 mN/m. An examination of changes in membrane morphology as a function of surface pressure demonstrates a gradual evolution in the size and surface coverage for both nano- and microdomains, prior to the formation of a network of interconnected nanodomains. The decrease in domain size is accompanied by a decrease in the height difference between the domain and the fluid phase. Analogous changes in domain size and height with increasing surface pressure are observed for monolayers in which egg SM has been replaced with dipalmitoylphosphatidylcholine (DPPC). The variation in domain size with surface pressure has been attributed to changes in line tension.29 A theory developed by McConnell to explain phase equilibria at the air-water interface rationalizes the sizes and shapes of domains in lipid monolayers on the basis of competition between the opposing forces of line tension and dipole densities.32,33 The line tension at the domain boundary favors large circular domains whereas dipolar or electrostatic interactions between molecules within domains favor small and/or extended or irregular shapes. The evolution of domain size observed for the monolayers of ternary lipid mixtures is consistent with a higher line tension at lower surface pressure. A recent theoretical analysis has examined the dependence of line tension on the elastic moduli of the raft and the surrounding membrane.34,35 The model considers the deformations of a single monolayer that are necessary to avoid exposing the hydrophobic surfaces to water at the boundary between a thick raft and the surrounding thinner membrane and predicts that line tension will increase quadratically with the difference in thickness between the two. Our AFM data indicate a 0.2 nm decrease in the height difference between domains and the fluid phase when the surface pressure is raised from 10 to 30 mN/m, in excellent qualitative agreement with the prediction that a smaller height mismatch will lead to a decrease in line tension and a concomitant decrease in raft size. (30) Nagle, J. F. J. Membr. Biol. 1976, 27, 233-250. (31) Feng, S.-S. Langmuir 1999, 15, 998-1010. (32) McConnell, H. M. Annu. ReV. Phys. Chem. 1991, 42, 171-95. (33) Benvegnu, D. J.; McConnell, H. M. J. Phys. Chem. 1993, 97, 66866691. (34) Kuzmin, P. I.; Akimov, S. A.; Chizmadzhev, Y. A.; Zimmerberg, J.; Cohen, F. S. Biophys. J. 2005, 88, 1120-1133. (35) Akimov, S. A.; Kuzmin, P. I.; Zimmerberg, J.; Cohen, F. S.; Chizmadzhev, Y. A. J. Electroanal. Chem. 2004, 564, 13-18.

The effect of varying Chol concentration in ternary lipid mixtures has also been examined. Egg SM/DOPC monolayers deposited at 10 and 30 mN/m exhibit gel-phase domains that are higher than the surrounding fluid phase.29 At both pressures, the addition of 20% Chol leads to a small decrease (0.2-0.4 nm) in the height difference between domains and the fluid phase, consistent with the transformation of the tightly packed gel phase to a liquid-ordered phase with Chol inserted between SM molecules. Chol addition also leads to slightly smaller domains at 10 mN/m and to an increase in interconnectivity at 30 mN/m. Related studies of brain SM/DOPC and brain SM/palmitoyloleolyphosphatidylcholine (POPC) raft mixtures have offered similar conclusions.36,37 Increases in the Chol content lead to two effects for monolayers transferred at 30 mN/m: the first is a decrease in the size of individual liquid-ordered domains, and the second is a decrease in the height difference between domains and the fluid phase. The combined data from these studies indicate that phase separation is detectable by AFM for SM/Chol/ unsaturated PC mixtures up to ∼50% Chol in some cases. Note that quantitative comparisons are not possible between the various studies because changes from egg to brain SM (with different distributions of chain lengths and numbers of double bonds) and DOPC to POPC have significant effects on the phase behavior of the mixtures. The morphology of monolayers of some ternary lipid mixtures is extremely sensitive to lipid oxidation. For example, the small interconnected domains in 2:2:1 egg SM/DOPC/Chol monolayers at 30 mN/m are transformed into large micrometer-sized domains upon exposure to air; similar effects are observed for ternary mixtures with DPPC and 1:1 SM/DOPC.29 Literature precedent33 and supporting evidence from the effects of substitution of lipids that are less prone to oxidation and the addition of antioxidants indicate that the changes in monolayer morphology are caused by the formation of oxidized lipids that increase the line tension and give larger domain sizes. Although the isolation of oxidized lipids from monolayers of ternary lipid mixtures is problematic, the range of lipid mixtures examined and the substitution of Chol for dihydrocholesterol indicated that the oxidation of both DOPC and SM contributes to the changes in domain size.29 The changes in monolayer morphology upon exposure to air are similar to observations of domain growth promoted by photoinduced oxidation in vesicles containing dye-labeled lipids.38 The monolayer results are also consistent with an earlier report that concluded that the formation of large domains at high surface pressures (>30 mN/m) for monolayers exposed to air is caused (36) Milhiet, P. E.; Domec, C.; Giocondi, M.-C.; Mau, N. V.; Heitz, F.; Le Grimellec, C. Biophys. J. 2001, 81, 547-555. (37) Lawrence, J. C.; Saslowsky, D. E.; Edwardson, J. M.; Henderson, R. M. Biophys. J. 2003, 84, 1827-1832. (38) Ayuyan, A. G.; Cohen, F. S. Biophys. J. 2006, 91, 2172-2183.

Nanoscale Imaging of Domains in Lipid Membranes

by lipid oxidation.39,40 However, the higher spatial resolution available with AFM indicates that exposing monolayers to air leads to an increase in the size of pre-existing nanodomains,29 rather than a change in the miscibility pressure. The observation that lipid oxidation has such pronounced effects on monolayer morphology for ternary lipid mixtures raises important questions concerning the validity of some literature data. Although we now routinely prepare monolayers containing unsaturated lipids in an inert environment, this does not appear to be the case in some published studies (including some of our own early work). Thus, some of the large domains that are observed at high surface pressure may result from the presence of lipid oxidation products. However, the situation is further complicated by the fact that not all ternary lipid mixtures show pronounced changes in morphology upon exposure to air. For example, we have recently observed that mixtures of egg SM/ POPC/Chol do not show increased domain size upon exposure to air for a significantly longer period of time than is required to cause large changes in SM/DOPC/Chol monolayers.41 This may be due to a significantly slower rate of oxidation (note that POPC has a single unsaturated acyl chain whereas DOPC has two), a reduced sensitivity of the POPC mixture to oxidized lipids, or to some combination of these effects. Resolving these issues is particularly problematic because lipid oxidation gives a complex mixture of products; the analysis of these complex mixtures for monolayers that contain very small amounts of lipid presents a formidable analytical problem. Preliminary results indicate that the addition of antioxidants is a reliable method for preventing oxidation at the air-water interface, which is a more straightforward solution than preparing monolayers in an inert environment.41 Bilayers. Supported lipid bilayers are a more realistic model for a cellular membrane than monolayers and have attracted considerable interest both as model systems and for their potential biotechnology applications.42-44 They are amenable to characterization with a wide range of surface-sensitive techniques and have been shown to retain the activity of reconstituted proteins and to have significant lipid mobility. The lateral lipid mobility is primarily due to the thin water layer (1 to 2 nm) that separates the lower leaflet of the bilayer from the solid support and aids in decoupling the membrane from the surface. Nevertheless, lipid diffusion in fluid-phase supported bilayers is approximately 2 times slower than in a vesicle under the same conditions,45 and the main phase transition for PC bilayers on mica occurs at a higher temperature and over a broader temperature range than in vesicles. 46 There are two main methods of preparing supported lipid bilayers. The first involves the adsorption of small unilamellar vesicles on the surface of hydrophilic solid supports, followed by vesicle rupture and spreading to give a uniform membrane.47 This method is compatible with a wide range of lipid compositions and with the incorporation of integral membrane proteins. Detailed studies using a variety of surface-sensitive techniques have (39) Stottrup, B. L.; Veatch, S. L.; Keller, S. L. Biophys. J. 2004, 86, 29422950. (40) Stottrup, B. L.; Stevens, D. S.; Keller, S. L. Biophys. J. 2005, 88, 269276. (41) Scassozi, E.; Johnston, L. J. Unpublished work, 2007. (42) Sackmann, E. Science 1996, 271, 43-48. (43) Boxer, S. G. Curr. Opin. Chem. Biol. 2000, 4, 704-709. (44) Tanaka, M.; Sackmann, E. Nature 2005, 437, 656-663. (45) Przybylo, M.; Sykora, J.; Humpolickova, J.; Benda, A.; Zan, A.; Hof, M. Langmuir 2006, 22, 9096-9099. (46) Leonenko, Z. V.; Finot, E.; Ma, H.; Dahms, T. E. S.; Cramb, D. T. Biophys. J. 2004, 86, 3783-3793. (47) Brian, A. A.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 6159-6163.

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Figure 3. AFM images of 1:1 DOPC/DPPC bilayers prepared by vesicle fusion at room temperature on mica and imaged in water. Images A and B were obtained immediately and 24 h after preparation of the bilayer, respectively. The cross section (A′) for the line indicated in image A shows domains with two different heights (0.8 and 1.2 nm) above the fluid phase. Adapted from ref 49.

examined this process in some detail and have provided at least a qualitative picture of the manner in which the lipid composition and concentration, temperature, electrostatic effects, solid support, and presence of divalent cations (particularly calcium) affect the speed of bilayer formation and the overall morphology of the resulting membrane.48 Figure 3A shows images of a phaseseparated bilayer comprising saturated (DPPC, C16, TM ) 41 °C) and unsaturated (DOPC, C18 with 1 double bond, TM ) -18 °C) phosphatidylcholines and prepared by vesicle deposition.49 The membrane shows raised domains with two distinct heights extending 1.2 and 0.8 nm above the surrounding fluid phase. The presence of domains with different heights is attributed to a compositional asymmetry in the upper and lower leaflets of the bilayer; the higher domains have DPPC in both leaflets whereas the lower domains result from DPPC in one leaflet and DOPC in the other. The presence of symmetric domains is very sensitive to the thermal history of the sample, as shown by the observation of predominantly symmetric domains with DPPC superimposed in the two leaflets when higher temperatures (45-60 °C vs room temperature) or longer equilibration times (Figure 3B for a bilayer equilibrated at room temperature for 24 h) are used for vesicle and/or bilayer formation.49,50 The coupling of domains between upper and lower leaflets for a DSPC/DLPC mixture was similarly shown to depend on the thermal history of the sample.51 Although it is usually assumed that the lipid composition is the same in both leaflets for bilayers prepared from vesicles, there is evidence of significant asymmetry in the interleaflet distribution of PS for DOPC/dioleolylphosphatidylserine (DOPS) bilayers on mica.48 It has been postulated that calcium-mediated interactions between DOPS and the support are responsible for this asymmetry. A second technique used to prepare supported lipid bilayers is based on the sequential transfer of two monolayers from the air-water interface.52 Depending on the lipid composition, either the Langmuir-Blodgett (LB, transfer by vertical dipping) or Langmuir-Blodgett/Schafer (LB-LS, successive vertical and horizontal dips for the transfer of the two monolayers) technique (48) Richter, R. P.; Berat, R.; Brisson, A. R. Langmuir 2006, 22, 3497-3505. (49) Choucair, A.; Chakrapani, M.; Chakravarthy, B.; Katsaras, J.; Johnston, L. J. Biochim. Biophys. Acta 2007, 1768, 146-154. (50) Giocondi, M.-C.; Vie, V.; Lesniewska, E.; Milhiet, P.-E.; Zinke-Allmang, M.; Le Grimellec, C. Langmuir 2001, 17, 1653-1659. (51) Lin, W.-C.; Blanchette, C. D.; Ratto, T. V.; Longo, M. L. Biophys. J. 2006, 90, 228-237. (52) Schwartz, D. K. Surf. Sci. Rep. 1997, 27, 245-334.

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Figure 5. AFM (A) and NSOM (B) images for 2:2:1 egg SM/ DOPC/Chol bilayers prepared by vesicle fusion on mica and imaged in aqueous solution. The bilayer imaged by NSOM contains 1% TR-DHPE.

Figure 4. NSOM (A, B) and AFM (C) images in aqueous solution for a hybrid bilayer prepared by transferring a 7:3 DLPC/DPPC monolayer containing 2% TR-DHPE at 30 mN/m to a DPPE monolayer on mica. A comparison of AFM and NSOM images indicates that the large domains contain gel-phase DPPC whereas some of the small, dark islands observed in the NSOM images are bilayer defects. Adapted from ref 24.

is used. Transfer of the upper monolayer generally works best when a tightly packed gel-phase lipid is used for the lower leaflet because this minimizes defects caused by the loss of material from the lower leaflet during the deposition of the second layer.53,54 Figure 4 shows an example of a phase-separated hybrid bilayer with a mixture of short-chain (dilauroylphosphatidylcholine, DLPC) and long-chain (DPPC) PCs transferred by vertical dipping onto a dipalmitoylphosphoethanolamine (DPPE) monolayer.24 The NSOM images (Figure 4A,B) show large, dark gel-phase DPPC domains and smaller dark islands surrounded by a fluorescent fluid bilayer. AFM (Figure 4C) indicates that many of these dark islands correspond to bilayer defects. We have used similar methods to prepare phase-separated hybrid bilayers with DLPC/dipalmitoylphosphatidylserine (DPPS) and with several other ternary lipid mixtures with Chol or SM in the top leaflet.55,56 The Langmuir methods are, in principle, useful for the fabrication of asymmetric bilayers that have different lipids distributions in the two leaflets, an important factor for mimicking the asymmetry of a native cell membrane. However, an early study of bilayers containing anionic phospholipids showed that there was a significant exchange of lipids between the upper and lower leaflets around defects.54 Two more recent studies have (53) Bassereau, P.; Pincet, F. Langmuir 1997, 13, 7003-7007. (54) Rinia, H. A.; Demel, R. A.; van der Eerden, J. P. J. M.; De Krujiff, B. Biophys. J. 1999, 77, 1683-1693. (55) Murray, J.; Cuccia, L.; Ianoul, A.; Cheetham, J. C.; Johnston, L. J. ChemBioChem 2004, 5, 1-6. (56) Yuan, C.; O’Connell, R. J.; Feinberg-Zadek, P. L.; Johnston, L. J.; Treistman, S. N. Biophys. J. 2004, 86, 3620-3633.

concluded that it is in fact quite difficult to maintain the initial bilayer asymmetry for bilayers prepared by LB-LS methods for a variety of lipid mixtures with coexisting liquid-ordered and liquid-disordered phases.57,58 Changes in the initial lipid composition occur by the removal of some of the lower monolayer during the transfer of the second and by lipid exchange between layers. However, more stable asymmetric bilayers could be formed in some cases by depositing an initial monolayer by LB transfer to a tethered polymer support, followed by addition of the upper leaflet by vesicle fusion.58 In some cases, these problems have been overcome by preparing hybrid bilayers by the transfer of a lipid monolayer to a glass slide that has been modified by covalent attachment of a hydrocarbon layer.59,60 Figure 5 shows representative AFM and NSOM images for a 2:2:1 egg SM/DOPC/Chol bilayer prepared by the vesicle spreading method. Both topographic and fluorescence contrast show approximately round domains in a range of sizes, with some irregular shapes that most likely arise by the coalescence of two smaller domains. Analogous results are obtained for several other ternary lipid mixtures.17,19,61-64 In each case, the liquidordered domains are coupled through both leaflets with no evidence of the asymmetry that is sometimes observed in phaseseparated bilayers of binary mixtures (e.g., Figure 3A). Bilayers with ternary lipid raft mixtures in one or both bilayer leaflets have also been prepared using LB or LS methods.39,40,59,65 However, the interpretation of these results is complicated by the use of fluorescence microscopy techniques that do not resolve closely spaced nanodomains and by possible complications due to the lipid oxidation of monolayers and the equilibration of lipids between leaflets, as discussed above. Domain Size. The above examples and literature data clearly indicate that domain sizes vary from tens of nanometers to ∼10 µm in supported monolayers and bilayers prepared from ternary lipid mixtures that are used to model the properties of membrane rafts. The transition from microdomains to nanodomains that we observe with increasing surface pressure is qualitatively similar (57) Crane, J. M.; Kiessling, V.; Tamm, L. K. Langmuir 2005, 21, 13771388. (58) Kiessling, V.; Crane, J. M.; Tamm, L. K. Biophys. J. 2006, 91, 33133326. (59) Dietrich, C.; Bagatolli, L. A.; Volovyk, Z. N.; Thompson, N. L.; Levi, M.; Jacobson, K.; Gratton, E. Biophys. J. 2001, 80, 1417-1428. (60) Dietrich, C.; Volovyk, Z. N.; Levi, M.; Thompson, N. L.; Jacobson, K. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 10642-10647. (61) Milhiet, P.-E.; Giocondi, M.-C.; Baghdadi, O.; Ronzon, F.; Roux, B.; Le Grimellec, C. EMBO Rep. 2002, 3, 485-490. (62) Milhiet, P.-E.; Giocondi, M.-C.; Le Grimellec, C. J. Biol. Chem. 2002, 277, 875-878. (63) Saslowsky, D. E.; Lawrence, J.; Ren, X.; Brown, D. A.; Henderson, R. M.; Edwardson, J. M. J. Biol. Chem. 2002, 277, 26966-26970. (64) Weerachatyanukul, W.; Ira; Kongmanas, K.; Tanphaichitr, N.; Johnston, L. J. Biochim. Biophys. Acta 2007, 1768, 299-310. (65) Crane, J. M.; Tamm, L. K. Biophys. J. 2004, 86, 2965-2979.

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to reported changes in domain size for monolayer or vesicle membranes as a function of changes in Chol concentration.37,66,67 For example, studies of DLPC/DPPC/Chol mixtures show large micrometer-sized domains at low Chol concentrations whereas at higher amounts of Chol there are much smaller nanodomains that cannot be visualized by optical microscopy but are detected by either fluorescence resonance energy transfer (FRET) or AFM.67 A range of raft sizes has also been reported for SM/ POPC/Chol mixtures on the basis of a time-resolved FRET study.68 These observations are in contrast to other studies in giant vesicles where large circular domains are frequently observed. Small nanodomains may be more relevant than large micrometer-sized domains for understanding the lipid organization in cellular membranes where rafts are believed to be small. However, the variations between results for various model membranes have led to questions of whether these nanodomains correspond to thermodynamic phases and why they do not rapidly increase in size to full phase separation. A partial answer is provided by a recent theoretical study dealing with phaseseparation kinetics in flat lipid bilayers that has concluded that entropic traps stabilize nanodomains in multicomponent membranes.69 The early nucleation and independent growth stages of phase separation occur rapidly to give nanodomains that are typically less than 50 nm in radius. At longer times, domain mobility and merging become important, and line tension determines the domain size distribution. At low line tension, the decrease in entropy resulting from domain merging is larger than the entropy of merging, and only nanodomains are present; the entropy and boundary energy compete to trap nanodomains for as long as hours. Conversely, for large line tensions, the decrease in boundary energy dominates the unfavorable entropy of merging, and nanodomains rapidly grow to micrometer size. At intermediate line tensions, large and small domains can coexist. In supported membranes, the additional interaction between the support and the lower bilayer leaflet will also contribute to reducing the domain merger, relative to a vesicle bilayer with the same composition. However, the surface-bilayer interaction may to some extent reproduce the interaction of a cellular membrane with the underlying cytoskeleton. The comparison of results for vesicles and supported membranes clearly highlights the importance of applying techniques that are capable of probing domains on a range of length scales for studies of phase separation in raft models. As noted in a recent review, this is particularly important for the measurement of phase diagrams and indicates the limitations of using fluorescence microscopy alone for such determinations.11 In particular, the recent comparison of monolayer and bilayer phase diagrams for DOPC/SM/Chol mixtures will require changes to include phase separation over a wider pressure range for monolayers.40

Distribution of Glycolipid Raft Markers in Phase-Separated Membranes The glycolipid ganglioside GM1 has been widely used as an in vivo marker for membrane rafts and is usually detected by fluorescence after the selective binding of a dye-labeled cholera (66) Milhiet, P. E.; Vie, V.; Giocondi, M.-C.; Le, Grimellec, C. Single Mol. 2001, 2, 109-112. (67) Tokumasu, F.; Jin, A. J.; Feigenson, G. W.; Dvorak, J. A. Biophys. J. 2003, 84, 2609-2618. (68) de Almeida, R. F. M.; Loura, L. M. S.; Federov, A.; Prieto, M. J. Mol. Biol. 2005, 346, 1109-1120. (69) Frolov, V. A. J.; Chizmadzhev, Y. A.; Cohen, F. S.; Zimmerberg, J. Biophys. J. 2006, 91, 189-205.

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Figure 6. AFM (A-C) and NSOM (D) images of 1:1 DOPC/ DPPC monolayers transferred to mica at 30 mN/m. Monolayers contain either 4% GM1 (A) or 4% GM1-Bodipy. The NSOM image is an overlay of monomer (green, 515 nm) and dimer (red, 630 nm) emission and shows that the aligned branched structures in the AFM images (B, C) are aggregated ganglioside, whereas the monomeric ganglioside is detected only by NSOM. Adapted from ref 71.

toxin, the protein for which GM1 is the natural receptor. Several fluorescence studies of GM1 distribution in supported membranes provided evidence for its uniform distribution throughout large micrometer-sized liquid-ordered domains, results that were inconsistent with estimates for rafts in cellular membranes and led to questions on the validity of model systems.18,59,60,70,71 However, AFM studies of the distribution of GM1 in a variety of phase-separated monolayers provided strong evidence for a heterogeneous distribution of glycolipid within ordered domains. For example, we and others have shown that GM1 is heterogeneously distributed as small islands clustered around the edge and in the interior of the ordered or gel-phase domains in DPPC monolayers with coexisting liquid-ordered and liquid condensed phases and in phase-separated DOPC/DPPC monolayers (Figure 6A).72,73 A similar heterogeneous distribution of GM1 islands or a network of GM1-rich filaments was observed upon addition of GM1 to DPPC/Chol monolayers, as a model for the liquidordered phase.72 These and related studies provide strong evidence for the localization of GM1 in small nanodomains that appear to mimic some of the properties of cellular rafts. Small GM1rich islands within liquid-ordered domains of SM/DOPC/Chol monolayers were also observed by both AFM and two-color fluorescence.28 GM1 was doped with ∼1% of the fluorescent analog GM1-Bodipy (ceramide-chain-labeled) to visualize the glycolipid by fluorescence. The use of only labeled ganglioside dramatically changed the glycolipid partitioning with labeled GM1, giving small aggregates within the fluid phase. Similar changes in the partitioning of GM1 have been observed in bilayer membranes.19,70,74 The effect of dye labeling on glycolipid partitioning has been examined in more detail in a recent study that compared the (70) Samsonov, A. V.; Mihalyov, I.; Cohen, F. S. Biophys. J. 2001, 81, 14861500. (71) Coban, O.; Burger, M.; Laliberte, M.; Ianoul, A.; Johnston, L. J. Langmuir 2007, in press. (72) Yuan, C.; Johnston, L. J. Biophys. J. 2000, 79, 2768-2781. (73) Vie, V.; Mau, N. V.; Lesniewska, E.; Goudonnet, J. P.; Heitz, F.; Le, Grimellec, C. Langmuir 1998, 14, 4574-4583. (74) Burns, A. R.; Frankel, D. J.; Buranda, T. Biophys. J. 2005, 89, 10811093.

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Figure 7. AFM images of GM1-containing bilayers prepared by Langmuir-Blodgett transfer of a DPPC monolayer to a DPPE-on-mica monolayer (A and B: GM1 only in the top DPPC monolayer) and by vesicle fusion for 2:2:1 egg SM/DOPC/Chol (C). Bilayer A contains 10% GM1, which is visible as small raised islands. After incubation with cholera toxin B for 30 min and rinsing with water, the bilayer shows islands that are substantially higher (B, note the change in the z scale), consistent with protein binding. Bilayer C contains 1% GM1 and has been incubated with cholera toxin B. Prior to the addition of protein, the bilayer is similar to that shown in Figure 5A, with no evidence of GM1-rich areas. However, the heterogeneous distribution of protein on the raised domains in image C indicates that GM1 is clustered in the liquid-ordered domains. Note the three heights in the cross section corresponding to the fluid phase, ordered domains, and protein bound to the ordered domains. Adapted in part from ref 75.

distribution of GM1 and GM1-Bodipy in phase-separated monolayers using AFM and NSOM.71 The independent measurement of the Bodipy monomer and dimer emission was used to distinguish between monomeric and aggregated glycolipid. For DOPC/DPPC mixtures, the labeled ganglioside partitions into both fluid-phase (monomer) and gel-phase (dimer) domains at low surface pressure, forming either small islands or long filaments. By contrast, at high surface pressure the addition of GM1-Bodipy leads to smaller gel-phase domains and the formation of a new GM1-rich phase that is distinct from either the fluid or gel phase and consists of small individual domains aligned in striped patterns (AFM, Figure 6B,C). The domain alignment was shown to occur during transfer to the substrate. Near-field fluorescence (Figure 6D) shows that the aggregated GM1 in the aligned domains is surrounded by a more dilute area of monomeric ganglioside that is not detected by AFM. The results for GM1-Bodipy differ significantly from the behavior of unlabeled GM1, which is heterogeneously distributed in small islands throughout gel-phase DPPC domains. In DPPC/DOPC/ Chol monolayers, both GM1 and GM1-Bodipy result in the disappearance of large micrometer-sized domains and the appearance of aligned domains of the GM1-rich phase with a central area of dimer surrounded by monomer. The dramatic effect of GM1 on this particular raft mixture is in contrast to the earlier data for ternary lipid mixtures containing SM. This may be due to inherent differences in the behavior of the DPPC and SM mixtures. This is substantiated by the observation that neither labeled nor unlabeled GM1 modifies the morphology of the small interconnected domains obtained for 2:2:1 SM/DOPC/Chol monolayers at 30 mN/m. It is also possible that oxidation may complicate the earlier results for the SM raft mixtures. Clearly, the effects of ganglioside addition are complex, particularly for the acyl-chain-labeled material, and experiments with a sugarlabeled ganglioside18,70 are necessary to resolve the effects of GM1 on raft monolayer morphology. Nevertheless, the complex partitioning and aggregation of GM1-Bodipy illustrate the importance of using multiple techniques to assess the behavior of multicomponent mixtures.

The addition of GM1 to DPPC and DPPC/Chol bilayers produces small islands of a new GM1-rich phase (Figure 7A).75 These islands are 30-200 nm in diameter, and their apparent height above the surrounding membrane is very sensitive to the imaging force, consistent with a significant electrostatic interaction between the tip and the negatively charged sugar head group of the ganglioside. Cholera toxin binding to the islands leads to a large increase in height (Figure 7B, 6 nm), providing unequivocal assignment to a GM1-rich area of the membrane.75 Similarly, the incubation of bilayers prepared from 2:2:1 SM/DOPC/Chol containing 1-5% GM1 shows selective binding of cholera toxin to liquid-ordered domains (Figure 7C). In this case, the protein is distributed unevenly across the domains (note that the domains show two distinct heights with the higher of the two corresponding to protein-rich areas), providing evidence for a heterogeneous distribution of glycolipid, although on a smaller length scale than is detectable by fluorescence. The observation of selective localization of GM1 in ordered domains in bilayers is consistent with several other studies using a range of methods in supported bilayers.18,19 The distribution of a different glycolipid, sulfogalactosylglycerolipid (SGG), has been examined in supported bilayers prepared from both artificial lipid mixtures and lipids extracted from cellular membranes.64 These experiments were undertaken to provide direct evidence for the association of SGG with raft domains in sperm plasma membranes. A mixture of DPPC/ palmitoyldocosahexaenoylphosphatidylcholine (PDPC)/Chol (2: 2:1 molar ratios) was used to model the plasma membrane, and SGG was demonstrated to localize in the liquid-ordered domains by assessing changes in domain morphology and surface coverage and by the selective binding of anti-SGG. Interestingly, bilayers prepared from lipids extracted from sperm plasma membranes showed SGG-rich domains that were comparable in size to those observed in DPPC/PDPC/Chol bilayers. Furthermore, confocal microscopy demonstrated that similar SGG-rich domains were observed in membranes prepared from plasma membrane vesicles. A disruption of the bilayers formed from both extracted lipids and artificial mixtures by the Chol-sequestering agent methyl(75) Yuan, C.; Johnston, L. J. Biophys. J. 2001, 81, 1059-1069.

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β-cyclodextrin provided further evidence for the involvement of raftlike domains. Overall, the good agreement between the results for bilayers prepared from artificial lipid mixtures and extracted membrane lipids demonstrates the utility of a supported membrane for mimicking at least some of the behavior of a natural membrane.

Enzyme Restructuring of Raft Membranes Ceramide is a hydrophobic sphingolipid that has significant effects on the physical properties of membranes;76,77 in model membranes, these effects include an increase in lipid order, the formation of ceramide-enriched domains, and the formation of nonlamellar structures. In cells, ceramide mediates signal transduction, induces transbilayer flip-flop, and has recently been hypothesized to promote the coalescence of small raft domains to give larger signaling platforms.78 This reorganization of raft domains is initiated by the activation of the enzyme sphinogmyelinase (SMase), which hydrolyzes sphingomyelin, forming ceramide and releasing phosphorylcholine.79 Although a number of studies in model systems have provided clear evidence for ceramide-promoted phase separation and the formation of large ceramide-enriched macrodomains, the coalescence of small raft domains to give large signaling platforms has not been observed directly. Recently, we have used AFM to examine the effects of ceramide incorporation on phase-separated ternary lipid mixtures.80 The enzymatic generation of ceramide on 2:2:1 SM/DOPC/Chol bilayers results in the formation of many small raised subdomains that are predominantly located on the edges of the original liquidordered domains (Figure 8A; compare to an untreated bilayer in Figure 5A). These are similar to the small subdomains that appear throughout the domains upon replacement of 5-10% of SM with ceramide in the vesicles used to form the supported bilayer (Figure 8B). The raised subdomains are assigned to either a ceramide- or SM-enriched phase. The formation of the shorter ceramide from SM would not be expected to lead to an increase in height. However, cholesterol is believed to be displaced from liquid-ordered domains by ceramide,81,82 so the subdomains may correspond to a more tightly packed gel phase. Their localization at the domain edges after SMase treatment is consistent with enzyme activity at the interface between the domain and the fluid phase where disordered lipid packing facilitates enzyme binding to its substrate. Similarly, phospholipase attack has been shown to occur at membrane defects.83 Additional experiments with asymmetric bilayers and with secondary ion mass spectrometry are currently being employed to provide more information on the composition of the domains for ceramide-containing mixtures. In addition to inducing domain heterogeneity, enzymatic ceramide generation leads to a significant restructuring of the membrane, an effect that is not observed upon direct incorporation of ceramide in the initial bilayer. This restructuring gives areas of the fluid phase that are devoid of domains, areas that have a distribution of domains similar to those observed prior to treatment with SMase, and areas containing clusters of domains in a new (76) Cremesti, A. E.; Goni, F. M.; Kolesnick, R. FEBS Lett. 2002, 531, 4753. (77) Bollinger, C. R.; Teichgraber, V.; Gulbins, E. Biochim. Biophys. Acta 2005, 1746, 284-294. (78) Grassme, H.; Jendrossek, V.; Riehle, A.; von Kurthy, G.; Berger, J.; Schwarz, H.; Weller, M.; Kolesnick, R.; Gulbins, E. Nat. Med. 2003, 9, 322-30. (79) Goni, F. M.; Alonso, A. FEBS Lett. 2002, 531, 38-46. (80) Ira; Johnston, L. Langmuir 2006, 22, 11284-11289. (81) Megha; London, E. J. Biol. Chem. 2004, 279, 9997-10004. (82) Chiantia, S.; Kahya, N.; Ries, H.; Schwille, P. Biophys. J. 2006, 90, 4500-4508. (83) Grandbois, M.; Clausen-Schaumann, H.; Gaub, H. Biophys. J. 1998, 74, 2398-2404.

Figure 8. AFM images showing the bilayer restructuring induced by the enzymatic generation of ceramide and a comparison with a bilayer to which ceramide has been incorporated at the stage of vesicle preparation. Images A and C show a 2:2:1 egg SM/DOPC/ Chol bilayer after incubation with SMase and washing with water to remove residual enzyme in the aqueous phase. The bilayer restructuring includes areas of heterogeneous domains with raised edges (image A, cross section A′, and area 1 in image C), areas of domains clustered in a new phase (C, area 2), and areas of the fluid phase with defects (C, area 3). Image B shows a 3:1:4:2 egg SM/ ceramide/DOPC/Chol bilayer (10% of SM replaced by ceramide), indicating that direct incorporation gives small raised subdomains (cross section B′) that are randomly scattered throughout the liquidordered domains. Adapted from ref 80.

phase that has a height intermediate between those of the domain and the fluid phase (Figure 8C). Quantification of the ceramide yield and the lack of effect of SMase on DPPC-Chol liquidordered domains in control bilayers demonstrated that the membrane restructuring is a direct result of the high local concentration of ceramide produced by enzymatic activity. On the basis of literature results indicating that ceramide can expel Chol from liquid-ordered domains,81,82 we hypothesized that the new phase surrounding the domains may be a Chol-DOPC-rich phase. Overall, the observation of ceramide-promoted heterogeneity and the clustering of raft domains in a physiologically relevant model provides strong support for the ceramide-induced formation of signaling platforms in cell membranes. The above observations raised questions about the dependence of the enzyme-mediated restructuring on concentration and time. Experiments in which bilayers were imaged immediately following enzyme addition (rather than after incubation and washing to remove excess enzyme) indicated that membrane restructuring starts almost immediately after exposure but the bilayer continues to evolve over a period of 1 to 2 h.80 The bilayer restructuring can also be followed by TIRF microscopy, which provides a significant advantage because data can be collected rapidly (within ∼30 s after enzyme addition) to follow the early stages of enzyme action and over a period of several hours to follow bilayer restructuring. This is difficult using AFM because tip contamination frequently prevents repeated imaging of bilayers exposed to enzyme. The two techniques are complementary, with AFM providing more information on features that are less than 300 nm in size whereas fluorescence gives the possibility of multiple labeling to aid in the identification of various membrane phases.

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Conclusions and Perspectives

Figure 9. TIRF and AFM images in aqueous solution for a 1:1 DOPC/DPPC + 0.5% TR-DHPE bilayer after incubation with 3 µM Aβ for 30 min (A, TIRF, 85:15 unlabeled/fluorescein-labeled Aβ) and 19 h (B, AFM, unlabeled Aβ). The TIRF image is an overlay of Texas red (red, 543 nm excitation) and fluorescein (green, 488 nm excitation) showing that Aβ accumulates selectively on the gelphase domains that exclude TR-DHPE. Rapid accumulation of peptide on the gel-phase domains is followed by a much slower aggregation that gives the large peptide aggregates visualized by AFM. Adapted from ref 49.

Peptide and Protein Binding to Domains Combinations of AFM and fluorescence are ideal for studying the interaction of peptides or proteins with supported bilayers and, in particular, for examining the preference for interaction with ordered versus fluid phases. As an example, we have recently shown that β-amyloid peptide Aβ42 interacts selectively with gel-phase DPPC domains in 1:1 DOPC/DPPC bilayers.49 TIRF microscopy using a mixture of dye-labeled and unlabeled Aβ42 and TR-DHPE to visualize the fluid phase shows that the peptide accumulates on the gel-phase domains within several minutes of addition (Figure 9A). AFM indicates that interaction with the membrane promotes a slower aggregation of the peptide that occurs over a period of hours (Figure 9B). The selective interaction of Aβ with DPPC domains is in contrast to the observation that Aβ is distributed randomly throughout both gel domains and the fluid phase when the peptide is reconstituted into DOPC/DPPC vesicles prior to bilayer formation. The preferential accumulation of Aβ on DPPC domains suggests that ordered membrane domains may act as platforms to concentrate peptide and enhance its aggregation and may be relevant to recent studies that implicate lipid rafts in Aβ accumulation and toxicity.84 Similarly, AFM studies have shown that placental alkaline phosphatase is targeted to raft domains in supported lipid bilayers.61,63 In a related study, AFM and NSOM were used to examine the interaction of synapsin, a membrane-associated protein responsible for maintaining a pool of neurotransmitter-loaded synaptic vesicles for use during neuronal activity, with supported bilayers prepared from DLPC and DPPS mixtures.55 This study provided direct visualization of the selective accumulation of synapsin on charged PS domains in the membranes, consistent with a strong electrostatic component of protein binding.85 The height change and increased fluorescence upon binding of dye-labeled protein to the membrane and the use of synapsin-coated tips were employed to visualize charged domains in cases where the height difference between phases was too small to be detected via topographic contrast. (84) Cordy, J. M.; Hooper, N. M.; Turner, A. J. Mol. Membr. Biol. 2006, 23, 111-122. (85) Cheetham, J. C.; Murray, J.; Ruhkalova, M.; Cuccia, L.; McAloney, R.; Ingold, K. U.; Johnston, L. J. Biochem. Biophys. Res. Commun. 2003, 309, 823829.

This article has summarized our recent work using a combination of AFM, NSOM, and TIRF to study the nanoscale organization of supported lipid monolayers and bilayers. There are several general conclusions that emerge from this work and the literature references cited herein. First, our results illustrate the importance of using techniques that can probe the organization of membranes over a range of length scales. It is particularly important to utilize methods with nanometer resolution because small nanodomains that cannot be detected by conventional fluorescence microscopy are observed for many ternary lipid mixtures commonly used to mimic membrane rafts. High spatial resolution is also advantageous for studies of the interaction of peptides or proteins with a lipid bilayer. AFM has unique capabilities for these experiments, being compatible with imaging in an aqueous environment and with the detection of individual biological macromolecules. A second important consideration is the frequent necessity to combine topographic contrast with other measurements such as fluorescence, particularly for multicomponent mixtures. The combination of topographic and fluorescence contrast leads to a more complete description of the system than either technique alone. It is frequently important to carry out simultaneous topographic and optical scans of the same area of the sample so that a hybrid microscope such as NSOM or an AFM directly integrated with a TIRF or confocal platform provides a distinct advantage. The sub-diffraction-limited resolution of NSOM is clearly advantageous for some studies, although there are many examples where a diffraction-limited resolution of ∼300 nm is adequate for model membranes. Because NSOM is currently limited by the slow scan speed and the fragility and mechanical properties of the probes, a more routine and rapid method of fluorescence imaging is preferable when 50 nm spatial resolution is not essential. Although not covered in this article, the ability of secondary ion mass spectrometry to map the chemical composition of a lipid membrane with 100 nm resolution is an exciting recent development.86 The addition of a routine imaging tool with chemical specificity will fill an important gap in existing microscopy techniques. Many of the experiments described above were directed toward understanding the behavior of lipid mixtures that are relevant to membrane rafts. Although one should be mindful of the limitation of a model for mimicking the functional aspect of membrane rafts, there are important aspects of domain formation that can be understood in a model system and that are difficult to study in a native cell membrane. For example, model membranes with lipid compositions that favor nanodomains are important for understanding the factors that control lipid domain formation in cellular membranes. Our work also demonstrates the significant effects that small amounts of additives (e.g., glycolipids, dyes, and oxidized lipids) have on the phase-separation behavior of ternary lipid mixtures. Equally interesting is the ability to test in a well-controlled system the preference of peptides or proteins for localization in ordered (raftlike) versus fluid-disordered phases. The modulation of membrane organization by the enzymatic generation of ceramide is also a promising approach for modeling raft coalescence in cells. As a final comment, it is worth speculating on the utility of nanoscale methods such as AFM and NSOM for probing raft domains in cells. In this context, our recent observation of clusters (86) Kraft, M. L.; Weber, P. K.; Longo, M. L.; Hutcheon, I. D.; Boxer, S. G. Science 2006, 313, 1948-1951.

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of integral membrane proteins and their co-localization with caveolae membrane domains in fixed cardiac cells25,87 is an important step toward the direct visualization of lipid and protein components of rafts in native membranes. The potential for imaging lipid and protein domains at 50 nm resolution has broad implications. NSOM and other optical techniques with subdiffraction-limited resolution88,89 may in the next few years enable the direct visualization of raft domains with various protein and (87) Ianoul, A.; Grant, D. D.; Rouleau, Y.; Bani-Yaghoub, M.; Johnston, L. J.; Pezacki, J. P. Nat. Chem. Biol. 2005, 1, 196-202. (88) Willig, K. I.; Rizzoli, S. O.; Westphal, V.; Jahn, R.; Hell, S. W. Nature 2006, 440, 935-939. (89) Betzig, E.; Patterson, G. H.; Sougrat, R.; Lindwasser, O. W.; Olenych, S.; Bonifacino, J. S.; Davidson, M. W.; Lippincott-Schwartz, J.; Hess, H. H. Science 2006, 313, 1642-1645.

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lipid compositions in cells. Such studies may finally start to resolve many of the outstanding questions on membrane rafts.

Acknowledgment. I gratefully acknowledge the many coworkers and collaborators who have contributed to the studies of supported membranes summarized in this article. This work was supported in part by NRC’s Genomics and Health Initiative and by student stipends from the Natural Sciences Engineering Research Council.

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