Optical and Scanning Probe Analysis of Glycolipid Reorganization

Nov 19, 2002 - Molecular level associations between components within the lipid bilayer as proteins complex to the membrane surface are fundamental to...
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Optical and Scanning Probe Analysis of Glycolipid Reorganization upon Concanavalin A Binding to Mannose-Coated Lipid Bilayers† Bruce Bondurant,‡ Julie A. Last,‡ Tina A. Waggoner,‡ Andrea Slade,§ and Darryl Y. Sasaki*,‡ Sandia National Laboratories, Biomolecular Materials Department, MS 1413, Albuquerque, New Mexico 87185, and Department of Biochemistry, Institute of Biomaterials and Biomedical Engineering, University of Toronto, Toronto, Canada M5S 3G9 Received July 11, 2002. In Final Form: September 25, 2002 Molecular level associations between components within the lipid bilayer as proteins complex to the membrane surface are fundamental to the complete understanding of protein-membrane interactions. To characterize these protein-induced membrane-organizational processes, we have prepared a new fluorescently labeled glycolipid (PSMU) that enables monitoring of glycolipid aggregation within the lipid membrane. The glycolipid’s mannosamine headgroup was specifically recognized by the lectin concanavalin A (Con A). Fluorescence studies with liposomes composed of 5 mol % PSMU/distearylphosphatidylcholine found that the membrane reorganized in response to Con A adsorption. Initially aggregated structures of glycolipid were dispersed as a consequence of specific affinity with the lectin through steric restrictions imposed by other bound Con A, the distance between mannosyl receptor sites, and possible protein insertion events. The protein binding and membrane reorganizational process was slow (ca. days). An association constant for Con A with the glycolipid membrane was estimated to be 3 × 106 M-1, around 2 orders of magnitude higher than that for methyl-D-R-mannopyranoside. Nanoscale imaging with the atomic force microscope found that the glycolipid formed 10 nm wide dendrite structures throughout the membrane and that the bound Con A was associated with those nanoscale features.

Introduction Protein-membrane interactions are the basis of nearly all cell-cell and cell-environment communication. Through specific interactions with membrane receptors or by reorganization of membrane components through nonspecific interactions, proteins can trigger cell signaling cascades that result in cellular activation. These molecular reorganizations within the lipid membrane must be highly orchestrated in order for the signaling process to occur. Understanding how these interactions produce specific coordination in space and time is key to revealing how cell signaling functions. Beyond the biological implications, incorporation of some of the recent insights of proteinmembrane interactions into synthetic materials has already produced a number of unique applications for drug delivery systems,1 separation materials,2 and sensor platforms.3 In the study of protein-membrane interactions, it is important not only to measure the binding strength of the protein to the membrane surface but also to evaluate the reorganization of the membrane components in response * To whom correspondence should be addressed. Fax: (505)8445470. E-mail: [email protected]. † Part of the Langmuir special issue entitled The Biomolecular Interface. ‡ Sandia National Laboratories. § University of Toronto. (1) (a) Lasic, D. D. Liposomes: From Physics to Applications; Elsevier Science: Amsterdam, 1993. (b) Medda, S.; Mukherjee, S.; Das, N.; Naskar, K.; Mahato, S. B.; Basu, M. K. Biotechnol. Appl. Biochem. 1993, 17, 37-47. (2) (a) Pidgeon, C. Enzyme Microb. Technol. 1990, 12, 149. (b) Pidgeon, C.; Venkataram, U. V. Anal. Biochem. 1989, 176, 36. (3) (a) Gu, Y.; LaBell, R.; O’Brien, D. F.; Saavedra, S. S. Angew. Chem., Int. Ed. 2001, 40, 2320-2322. (b) Tauskela, J. S.; Thompson, M. Anal. Chim. Acta 1992, 264.

to the binding event. How strongly are the specific interactions coupled to membrane reorganization and how much of a role do nonspecific interactions play? To what extent do phase separation, electrostatic interactions, and steric effects contribute to the resultant molecular organization? There has been considerable research on understanding how surface-bound proteins reorganize membrane components, that is, membrane proteins and/ or lipids.4 Research has focused on isolating the nature of the host-guest complex,5 phase change of “bound” lipids,6 and aggregation of lipids and proteins.7 However, evaluating the change in relative membrane component positioning at the molecular level has been difficult. The linking of functional ligands to optically responsive lipid membranes can provide a method to evaluate the effect of protein binding on membrane reorganization. Typically, a ligand is attached to a lipid body at the headgroup position with a spacer arm to allow access to (4) (a) Vanderkooi, J.; McLaughlin, A. Use of Fluorescent Probes in the Study of Membrane Structure and Function; Chen, R. F., Edelhoch, H., Eds.; Marcel Dekker: New York, 1976; Vol. 2, p 737. (b) Devaux, P. F.; Seigneuret, M. Biochim. Biophys. Acta 1985, 822, 63-125. (c) Irvine, D. J.; Hue, K.-A.; Mayes, A. M.; Griffith, L. G. Biophys. J. 2002, 82, 120-132. (5) (a) Massey, J. B.; Gotto, A. M., Jr.; Pownall, H. J. Biochemistry 1981, 20, 1575-1584. (b) Kalb, E.; Engel, J.; Tamm, L. K. Biochemistry 1990, 29, 1607. (c) Mustonen, P.; Virtanen, J. A.; Somerharju, P. J.; Kinnunen, P. K. J. Biochemistry 1987, 26, 2991. (d) Mosmuller, E. W. J.; Pap, E. H. W.; Visser, A. J. W. G.; Engbersen, J. F. J. Biochim. Biophys. Acta 1994, 1191, 45. (e) Verbist, J.; Gadella, T. W. J., Jr.; Raeymaekers, L.; Wuytack, F.; Wirtz, K. W. A.; Casteels, R. Biochim. Biophys. Acta 1991, 1063, 1. (6) (a) Wiener, J. R.; Pal, R.; Barenholz, Y.; Wagner, R. R. Biochemistry 1985, 24, 7651. (b) Sui, S.-f.; Urumow, T.; Sackmann, E. Biochemistry 1988, 27, 7463-7469. (c) Chicken, C. A.; Sharom, F. J. Biochim. Biophys. Acta 1984, 774, 110-118. (7) (a) Rehorek, M.; Dencher, N. A.; Heyn, M. P. Biochemistry 1985, 24, 5980-5988. (b) Birrell, G. B.; Griffith, O. H. Biochemistry 1976, 15, 2925-2929. (c) Jones, M. E.; Lentz, B. R. Biochemistry 1986, 25, 567.

10.1021/la0262295 CCC: $25.00 © 2003 American Chemical Society Published on Web 11/19/2002

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the protein receptor site. At the other end of the molecule is an optical tag that is imbedded in the hydrophobic layer of the bilayer to isolate it from direct interaction with the bound protein and other aqueous phase analytes. Such functionalized lipids can be used to measure the aggregation of lipids upon protein binding.8 Moreover, such lipid bilayer systems can be used as unique biosensors to detect proteins in solution (e.g., his-tagged proteins9 or cholera toxin10). As a sensor material, these functionalized lipid bilayers offer excellent biocompatible properties, reversible affinity and response, and an optical detection scheme intrinsic to the material alleviating the need for protein tagging. We have an ongoing interest in trying to understand the change in the organizational state of molecular components in a lipid membrane in response to chemical recognition events at the membrane surface using fluorescently labeled lipids and nanoscale imaging.11,12 It is our expectation that specific chemistries of the lipids, host-guest interactions, and membrane compositions will give rise to unique insights into how cellular membranes react to specific ligands and environmental stimulation. Of particular interest is how membranes respond to adhesion in which protein-carbohydrate complexation plays a large role.13 Herein, we describe the preparation of a new fluorescently tagged glycolipid (mannosaminyl headgroup), its incorporation in phosphatidylcholine lipid bilayers, the selective binding of the lectin concanavalin A (Con A), and the reorganization of the membrane in response to the binding of the protein. Fluorescence data reveal that the glycolipid disperses at a slow rate within the bilayer upon Con A binding, while scanning probe imaging of the surface finds that the protein binds to regions in the bilayer initially rich with the glycolipid. Experimental Section General. All compounds were of reagent grade purity and used as supplied by the manufacturer unless stated otherwise. Organic solvents were of reagent grade from Fisher Scientific. Concanavalin A (Canavalia ensiformis) and bovine serum albumin (BSA) were obtained from Sigma (St. Louis, MO) in powder form and used as received. Distearylphosphatidylcholine (DSPC) and stearyloleylphosphatidylcholine (SOPC) were obtained from Avanti Polar Lipids (Alabaster, AL). Aqueous solutions were prepared from water purified through a Barnstead type D4700 NANOpure analytical deionization system with an ORGANICfree cartridge registering an 18.0 MΩ cm resistance. Fluorescence measurements were performed on a SPEX Fluoromax II spectrophotometer (Instruments SA, Edison, NJ) equipped with a jacketed cell. NMR analyses were performed on a Bruker DRX 400 (Billerica, MA) with a resonance frequency of 400.1 MHz for 1H and 100.6 MHz for 13C. Infrared absorption spectra were taken on a Perkin-Elmer 1750 FTIR spectrometer (Norwalk, CT). Elemental analyses were performed by Desert Analytics (Tucson, AZ). Carbonic Acid 8-[1-Octadecyl-2-(9-(1-pyrenyl)nonyl)rac-glyceroyl]-3,6-dioxaoctan-1-yl Ester 2,5-Dioxopyrrolo(8) Antes, P.; Schwarzmann, G.; Sandhoff, K. Chem. Phys. Lipids 1992, 62, 269. (9) Ng, K.; Pack, D. W.; Sasaki, D. Y.; Arnold, F. H. Langmuir 1995, 11, 4048-4055. (10) (a) Song, X.; Nolan, J.; Swanson, B. I. J. Am. Chem. Soc. 1998, 120, 11514-11515. (b) Song, X.; Nolan, J.; Swanson, B. I. J. Am. Chem. Soc. 1998, 120, 4873-4874. (11) Sasaki, D. Y.; Waggoner, T. A.; Last, J. A.; Alam, T. M. Langmuir 2002, 18, 3714-3721. (12) Last, J. A.; Waggoner, T. A.; Sasaki, D. Y. Biophys. J. 2001, 81, 2737-2742. (13) (a) Kreis, T.; Vale, R. Guidebook to the Extracellular Matrix, Anchor, and Adhesion Proteins, 2nd ed.; Kreis, T., Vale, R., Eds.; Oxford University Press: New York, 1999. (b) Lectins: Biology, Biochemistry, Clinical Biochemistry; Bog-Hansen, T. C., Freed, D. L. J., Eds.; Sigma: St. Louis, 1988; Vol. 6, p 757.

Bondurant et al. dine-1-yl Ester. 8-[1-Octadecyl-2-(9-(1-pyrenyl)nonyl)-racglyceroyl]-3,6-dioxaoctan-1-ol, or PSOH9 (1.6 g, 2.0 mmol), was dissolved in dry toluene (50 mL) in a 200 mL round-bottom flask, and the solvent was evaporated under reduced pressure. The atmosphere was restored with nitrogen gas. Dry DMSO (20 mL) was added followed by disuccinimidyl carbonate (0.77 g, 3.0 mmol) and dry pyridine (0.24 g, 3.0 mmol). The reaction was stirred under nitrogen at room temperature for 24 h, at which time it was found to be complete. The solvent was evaporated at 0.1 mmHg and 37 °C. Ethyl acetate (50 mL) was added, and the solution was filtered. The filtrate was washed with aqueous saturated NaHCO3 (20 mL) followed by deionized water (3 × 20 mL). The organic phase was dried with brine (20 mL) followed by anhydrous MgSO4. The solvent was removed under reduced pressure to produce 2.1 g of crude amber oil. The product was purified rapidly by vacuum-flash chromatography on silica gel in a 35 mL sintered glass funnel. The product was loaded in 5:1 hexanes/ethyl acetate (20 mL) and then eluted with a gradient ending with 1:3 hexanes/ethyl acetate. Carbonic acid 8-[1octadecyl-2-(9-(1-pyrenyl)nonyl)-rac-glyceroyl]-3,6-dioxaoctan-1yl ester 2,5-dioxopyrrolodine-1-yl ester (Rf ) 0.54 in 60% ethyl acetate/hexanes) was recovered in 90% yield (1.7 g, 1.8 mmol). 1H NMR (CDCl ): δ 8.287-8.261 (d, J ) 10.3 Hz, 1H, Py-H), 3 8.140-8.086 (m, 4H, Py-H), 8.012-7.970 (m, 3H, Py-H), 7.8667.846 (d, J ) 7.9 Hz, 1H, Py-H), 4.441-4.418 (t, J ) 4.8 Hz, 2H, C(O)OCH2), 3.756-3.732 (t J ) 4.8 Hz, 2H, C(O)OCH2CH2), 3.635-3.628 (d, J ) 3 Hz, 9H, glycerol OCH2), 3.568-3.399 (m, 10 H, triethyleneglycol OCH2), 3.341-3.302 (t, J ) 7.8 Hz, 2H, Py-CH2), 2.748 (s, 4H, succinimide CH2), 1.88-1.81 (p, J ) 7.4 Hz, 2H, Py-CH2CH2), 1.550-1.476 (m, 7H, OCH2CH2), 1.3661.238 (m, 43 H, aliphatic CH2), 0.890-0.856 (t, J ) 6.8 Hz, 3H, stearyl-CH3) ppm. 13C NMR (CDCl3): δ 168.7, 152.2, 137.6, 131.6, 131.1, 129.9, 129.2, 128.8, 128.4, 127.7, 127.4, 127.3, 126.7, 125.9, 125.5, 125.3, 125.0, 124.8, 123.7, 78.1, 77.6, 77.2, 76.9, 71.9, 71.6, 71.1, 71.0, 70.8, 70.4, 68.5, 33.8, 32.2, 30.3, 30.0, 29.9, 29.7, 29.6, 26.3, 25.6, 22.9, 14.3 ppm. FTIR (KBr): 2924.4, 2854.0, 1813.0, 1790.6, 1745.0, 1225.0, 1109.6 cm-1. O-[8-[1-Octadecyl-2-(9-(1-pyrenyl)nonyl)-rac-glyceroyl]3,6-dioxaoctan-1-yl] N-Mannosaminocarbamate (PSMU). Carbonic acid 8-[1-octadecyl-2-(9-(1-pyrenyl)nonyl)-rac-glyceroyl]-3,6-dioxaoctan-1-yl ester 2,5-dioxopyrrolodine-1-yl ester (0.47 g, 0.50 mmol) and mannosamine hydrochloride (0.16 g, 0.75 mmol) were combined in toluene (20 mL) and dry DMF (10 mL) in a round-bottom flask. The toluene was evaporated under reduced pressure, and the atmosphere was replenished with nitrogen. Dry diisopropylethylamine (0.13 g, 1.0 mmol) was added, and the reaction was stirred under N2 at room temperature for 3 h. The solvent was evaporated under reduced pressure. The crude brown oil was redissolved in chloroform, filtered through neutral Celite, and purified by column chromatography on silica gel with an elution gradient of pure CHCl3 to 80% CHCl3/ methanol. Further purification was achieved by reverse phase chromatography on C18 silica with an elution gradient starting at 30% ethanol/methanol and ending with 100% ethanol. Pure PSMU was recovered as an amber-colored oil in 73% yield (0.37 g, 0.37 mmol). 1H NMR (CDCl3): δ 8.283-8.260 (d, J ) 9.2 Hz, 1H, Py-H), 8.159-8.077 (m, 4H, Py-H), 8.035-7.952 (m, 3H, PyH), 7.865-7.846 (d, J ) 7.6 Hz, 1H, Py-H), 6.026-5.928 (m 1H, OC(O)NH), 5.751-5.730 (d, J ) 8.2 Hz, 0.3H, mannosyl C(1)OH), 5.247-5.204 (m, 1H, mannosyl C(1)-H), 4.853-4.835 (d, 7.2 Hz, O.5H, mannosyl -OH), 4.41-4.0 (m, 5H, mannosyl -OH), 4.0-3.40 (m, 27H, -OCH2, mannosyl -CH), 3.34-3.30 (t, J ) 8 Hz, 2H, Py-CH2), 1.88-1.81 (m, J ) 7.4 Hz, 2H, PyCH2CH2),1.54-1.46 (m, 6H, -OCH2CH2), 1.37-1.23 (m, 41 H, aliphatic CH2), 0.890-0.85 (t, J ) 6.8 Hz, 3H, -CH3) ppm. 13C NMR (DMSO): δ 156.4, 137.0, 130.9, 130.4, 129.2, 128.0, 127.4, 127.3, 127.1, 126.4, 126.0, 124.8, 124.7, 124.2, 123.3, 93.2, 92.9, 77.3, 72.8, 72.3, 70.5, 70.1, 69.7, 69.3, 68.8, 66.8, 63.2, 61.2, 55.7, 40.1, 39.9, 39.7, 39.5, 39.3, 39.1, 38.9,32.7, 32.3, 31.5, 31.3, 29.6, 29.0, 28.8, 28.7, 25.6, 22.1, 13.9 ppm. FTIR (KBr) 3355.8, 2923.9, 2852.9, 1700.7, 1541.3, 1463.7, 1250.6, 1109.1, 843.2 cm-1. Elemental Anal. Calcd for C59H93O12N: C, 70.27%; H, 9.30%; N, 1.39%. Found: C, 70.70%; H, 9.05%; N, 1.53%. Liposome Preparation. Thin films of lipid were prepared by volumetrically combining stock solutions of lipid in chloroform in a 10 mL round-bottom flask and evaporating the solvent under

Glycolipid Reorganization upon Protein Binding reduced pressure. The samples were then dried for 12 h at 0.1 mmHg and then hydrated with 3.0 mL of pH 7.0 PBS buffer (5 mM sodium phosphate, 144 mM sodium chloride, and 400 µM each of calcium chloride and manganese chloride) to make a suspension of 3.0 mM total lipid content. The lipids were subjected to 10 freeze-thaw cycles between -78 and 60 °C for DSPCcontaining liposomes or between -78 and 35 °C for SOPCcontaining liposomes. The hydrated lipids were extruded twice through two stacked 200 nm pore size polycarbonate membranes followed by 10 times through two stacked 100 nm pore size polycarbonate membranes. Extrusion of DSPC- and SOPCcontaining liposomes was done at 60 and 25 °C, respectively. Liposome Leakage Studies. Lipids were combined in chloroform (0.09 mmol total lipid) in a 10 mL round-bottom flask, the solvent was evaporated under reduced pressure, and the resulting film was dried at 0.1 mmHg for at least 8 h. The lipids were hydrated in 3.0 mL of an aqueous solution containing 50 mM carboxyfluorescein (5,6-isomers from Sigma) that had been titrated to pH 8 with NaOH. The lipid suspension was then processed in an identical manner as described above to yield 100 nm unilamellar liposomes of 5% PSMU/DSPC. Liposomes were separated from unencapsulated dye by gel permeation chromatography on Sephadex G-50 gel (50 mL) using isoosmolar PBS buffer (pH 7.5, 400 mOsm). The carboxyfluorescein concentration of the liposome solution was determined using the absorbance at 470 nm ( ) 63 000) of a 10-fold dilution of the liposome solution in pH 9 PBS. Liposomes were added to protein-containing solutions (PBS, pH 7.5) at an amount such that the total carboxyfluorescein concentration was 1.5 µM, while the protein content was variable between 0 and 50 µM. Fluorescence measurements were made with an excitation at 470 nm while monitoring at 515 nm. Measurements were made continuously over the first 30 min and then periodically over the latter 15 h. Fluorescence Response to Con A. Fluorescence measurements of the PSMU-containing liposomes were obtained using an excitation of 346 nm and emission taken between 365 and 550 nm (2 nm slit widths). Measurements were made in a stirred quartz cuvette at 25.0 ( 0.1 °C. The protein was diluted to appropriate concentrations in 3.0 mL of pH 7.0 PBS solution in the cuvette. A 10 µL aliquot of the liposome solution (3.0 mM total lipid concentration) was then added to the protein solution to produce a total lipid concentration of 10 µM. Emission scans were made immediately after addition of the liposomes followed by scans at various time intervals afterward. The monomer emission (M) was measured as the area under the curve between 365 and 437 nm, while the excimer emission (E) was measured between 437 and 550 nm. The reported M/(M + E) data are averages from at least three trials with a standard deviation of 7%. Transmission Electron Microscopy (TEM) Images. The liposomes were stained with ammonium molybdate using a standard TEM preparation protocol.14 TEM images were taken on a Philips CM-30 operated at 300 kV. Atomic Force Microscopy (AFM). AFM experiments were performed with a Nanoscope IIIa Multimode scanning probe microscope (Digital Instruments, Santa Barbara, CA). The images were acquired in tapping mode in solution using a commercially available liquid cell (Digital Instruments) with 120 µm oxidesharpened silicon nitride V-shaped cantilevers. The nominal spring constant of the cantilever was 0.35 N/m. Images were collected with the E scanner, which has a maximum range of 15 um × 15 um, operating at a scan rate of 2 Hz. Data were collected with 512 data points per line. The supported lipid bilayer was prepared via vesicle fusion on a freshly cleaved mica substrate. The clean mica was first imaged in PBS buffer to establish the baseline. The liposome solution was then injected into the AFM liquid cell, and the mica surface was imaged subsequently. Vesicle adsorption and fusion to the mica surface occurred rapidly. The liposome solution was incubated with the mica at room temperature for 1 h to allow optimal time for vesicle fusion, typically resulting in full bilayer coverage of the substrate. The bilayer was heated above 60 °C for 30 min to complete fusion of the bilayer. (14) New, R. R. C. Liposomes, 1st ed.; Oxford University Press: New York, 1990.

Langmuir, Vol. 19, No. 5, 2003 1831 Scheme 1

The addition of concanavalin A was accomplished by flushing a solution containing 9 µM concanavalin A in PBS buffer through the AFM fluid cell. Images were collected immediately after addition of the Con A and at subsequent intervals over a 16 h period.

Results and Discussion Pyrene-labeled, carbamate-linked D-mannosaminefunctionalized lipid, PSMU, was prepared in two steps from PSOH (Scheme 1). The headgroup was formed by linking mannosamine with the alcohol headgroup of PSOH through disuccinimidyl carbonate. The carbamate linkage between the carbohydrate and the lipid body provided the glycolipid in good yield while maintaining the 3, 4, and 6 hydroxyl groups of the pyranose free for accessible binding to the lectin recognition site.15 The mannosyl group can extend approximately 7 Å further into solution from the surface of the phosphatidylcholine membrane by the triethyleneglycol spacer. PSMU was isolated as an ambercolored oil with all spectral assignments matching the proposed structure. Liposomes of PSMU with DSPC or SOPC were prepared by suspending dried films of the lipid mixtures in PBS buffer (pH 7.0), followed by extrusion through 100 nm pore polycarbonate membranes. The vesicle size range, determined by TEM imaging of the ammonium molybdate stained vesicles, was 100-300 nm (Supporting Information, Figure S1). Lipid bilayers prepared with matrix lipids in the gel phase at room temperature (i.e., DSPC, Tg ) 55 °C) or in the fluid phase at room temperature (i.e., SOPC, Tg ) 6 °C) allowed us to more fully assess the behavior of glycolipid organization in the presence of lectin protein. Fluorescence spectra of 5 mol % PSMU/DSPC and 5 mol % PSMU/SOPC, shown in Figure 1, were obtained with an excitation of 346 nm. The difference in the excimer to monomer intensity ratio in the gel- and fluid-phase membranes reveals the fluid-phase behavior of the PSMU lipid. That is, in the gel-phase DSPC membrane the fluidphase PSMU lipid separates from the DSPC crystalline domains producing a high local concentration of pyrene and a large excimer to monomer fluorescence intensity ratio. In the fluid-phase matrix of SOPC, however, the PSMU lipid mixes readily in the matrix resulting in a low excimer to monomer ratio.16 (15) Hardman, K. D.; Goldstein, I. J. The Structure and Activity of Concanavalin A; Atassi, M. Z., Ed.; Plenum: New York, 1977; Vol. 2, pp 373-416. (16) (a) Galla, H.-J.; Hartmann, W. Chem. Phys. Lipids 1980, 27, 199. (b) Hresko, R. C.; Suga´r, I. P.; Barenholz, Y.; Thompson, T. E. Biochemistry 1986, 25, 3813.

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Figure 1. Fluorescence spectra of liposomes composed of 5% PSMU/DSPC (solid line) and 5% PMSU/SOPC (dotted line) using an excitation wavelength of 346 nm.

Figure 2. Fluorescence spectra of 5% PSMU/DSPC liposomes before and after exposure to 10 µM concanavalin A in PBS buffer (pH 7.0) recorded at different time intervals.

Addition of Con A into solutions of either PSMU/DPSC or PSMU/SOPC liposomes, at a total lipid concentration of 10 µM, produces a loss in pyrene excimer fluorescence intensity with a concomitant increase in the monomer intensity. The change in fluorescence occurred relatively slowly reaching a plateau over a period of several hours for the SOPC bilayers, yet taking longer than 2 days for the DSPC bilayers at 25 °C. The response was distinct with the DSPC bilayers, while that for the SOPC bilayers registered only a change of ∼10%. A series of fluorescence spectra of a stirred solution of the 5% PSMU/DSPC liposomes in the presence of 10 µM of Con A over the period of 56 h is shown in Figure 2. An important note from these series of spectra is the presence of an isosbestic point at 437 nm. This indicates that the fluorescence change was due to changes in the aggregational state of the pyrene moieties without contributions from fluorescence quenching mechanisms,17 such as changes in the polarity of the environment or metal ion interaction, both of which could occur if the bound protein deteriorates the lipid membrane. (17) Kinnunen, P. K. J.; Koiv, A.; Mustonen, P. Pyrene-Labeled Lipids as Fluorescent Probes in Studies on Biomembranes and Membrane Models; Wolfbeis, O. S., Ed.; Springer-Verlag: New York, 1993; p 159.

Bondurant et al.

Figure 3. A time-dependent plot of fluorescence response (M, monomer emission; E, excimer emission) of 5% PSMU/DSPC liposomes in the presence of Con A at 0.1 µM (O), 1.0 µM (]), 3.0 µM (0), 5.0 µM (1), 7.0 µM (9), and 10 µM (b).

With regard to aggregation of the liposomes in the presence of Con A, only a slight turbidity was observed at the highest concentration reported (e.g., 10 µM). Higher concentrations of Con A (50 µM) gave similar results to those produced at 10 µM. The spacer length of the PSMU was apparently long enough for mannosyl recognition by the Con A receptor site18 but sufficiently short such that Con A induced liposome aggregation was not a significant factor.19 The fluorescence response to Con A was a slow process that contrasts the relatively rapid binding of Con A to receptor-laden membranes (40 min) observed by others.20 The fluorescence data reported here do not, however, represent just the binding of protein to a membrane but rather are a monitor of membrane reorganization in response to protein-membrane interaction. The results we observe seem to indicate that the membrane continues to reorganize well after the time at which Con A should have reached a stable binding stoichiometry with the membrane. A plot of the pyrene monomer emission over the total emission of both monomer and excimer (M/(M + E)) versus time for various concentrations of Con A is shown in Figure 3. The very broad fluorescence response curves do not reach plateaus within reasonable time frames of our experiment, and so we arbitrarily confine our results to a sampling time of 30 h, where the response curves begin to flatten. A plot of the fluorescence response with respect to the concentration of Con A in solution (Figure 4) was produced using this sampling time. The data fit a Langmuir adsorption isotherm21 rather well, allowing a coarse estimate of the binding constant of Con A to these glycolipid-functionalized membranes at a value of 3 × 106 M-1. Control experiments using nonlectin proteins and dummy ligands on the membrane surface provided further evidence of the specificity of the lectin-membrane interaction. Addition of 10 µM BSA produced no significant (18) Sundler, R. Biochim. Biophys. Acta 1984, 771, 59-67. (19) Orr, G. A.; Rando, R. R.; Bangerter, F. W. J. Biol. Chem. 1979, 254, 4721-4725. (20) Ketis, N. V.; Grant, C. W. M. Biochim. Biophys. Acta 1983, 730, 359-368. (21) Connors, K. A. Binding Constants; John Wiley & Sons: New York, 1987.

Glycolipid Reorganization upon Protein Binding

Figure 4. The fluorescence response of the 5% PSMU/DSPC liposomes with varying concentrations of Con A in solution sampled at 30 h.

fluorescence response from the liposomes. Likewise, by substituting PSOH for PSMU, thus removing the carbohydrate headgroup functionality, the fluorescence response of the liposomes for Con A was completely extinguished. Binding of Con A to the PSMU-containing liposomes was inhibited, albeit incompletely, using Dmannose. A 2000-fold excess of D-mannose (10 mM) to Con A (5 µM) in solution inhibited the fluorescence response to 60% of that found in the uninhibited experiment. By removing the bound Con A from the membrane, we would expect the original bilayer fluorescence to return, as we have seen with previous studies with metal-chelating membranes.11,12 However, the fluorescence spectra of the Con A bound 5% PSMU/DSPC bilayers could not be reverted, even slightly, with the addition of 10 mM D-mannose to solution indicating strong affinity of the protein for the membrane. The fluorescence results observed here appear to contradict other reports where lipid or receptor aggregation was induced by Con A binding at the membrane surface.20,22 If one considers specific binding occurring between a protein and a glycolipid, the size mismatch is considerable. Con A is a tetramer of 26 kD proteins formed in a tetrahedral structure at pH 7.0, with the mannosyl binding site at the apex of each protein in the tetrahedron.23 The width of the structure is ca. 100 Å. The width of a PC lipid is ca. 6-7 Å, and the glycolipid is about 50% larger. A schematic drawing of such a protein bound to the 5% glycolipid membrane depicts the contrast in size of the protein and bound glycolipid (Figure 5). It is evident from the drawing that if specific protein-glycolipid binding occurs the glycolipids will be separated either by the steric interactions of the bound proteins for one tetramer binding to one glycolipid or by the physical separation of binding sites of the tetramer binding to two or three glycolipids. The slow response time observed with the fluorescence data could be understood from the steric bulk of bound proteins crowding the membrane surface. The PSMU lipids, aggregated in domains within the DSPC matrix, would cause the proteins to initially bind to these highly functionalized areas producing a canopy-like effect over (22) Kitano, H.; Ishino, Y.; Yabe, K. Langmuir 2001, 17, 2312-2316. (23) (a) Moothoo, D. N.; Canaan, B.; Field, R. A.; Naismith, J. H. Concanavalin A Structure 1BXH, Protein Data Bank, http://www. rcsb.org/pdb/. (b) Kalb, A. J.; Habash, J.; Hunter, N. S.; Price, H. J.; Raferty, J.; Helliwell, J. R. Met. Ions Biol. Syst. 2000, 37, 279-304.

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Figure 5. Schematic illustration of Con A binding to the outer leaflet of the liposomal membrane with initially aggregated PSMU glycolipid. The tetrahedral structure of Con A and the carbohydrate binding site located at the periphery of each subunit induce glycolipid dispersion.

the available glycolipids and subsequently shielding further lectin interactions. The low molecular mobility of lipids in the gel-phase DSPC matrix would, on a slow time scale, permit the glycolipids to reposition themselves in the membrane to interact with free Con A. The more rapid fluorescence response time observed for Con A binding with the fluid-phase 5% PSMU/SOPC bilayers is consistent with the concept of lipid mobility, host-guest accessibility, and protein crowding at the surface. However, other scenarios for the observed membrane reorganization may also play roles, significant or otherwise, in this complex interaction between structurally dynamic systems. A very strong possibility exists for protein insertion or denaturation within the membrane, which may also occur over long time periods. In this case, the inserting protein may disperse the glycolipids through volume displacement or through a deterioration of the membrane (e.g., micelle formation with lipids, thinning of the membrane, hole formation) that results in a permeable structure. A leakage study was thus performed on these glycolipid membranes to assess the membrane quality in response to the binding of Con A. Carboxyfluorescein was entrapped at a high concentration within 5% PSMU/DSPC liposomes followed by liposome separation from nonentrapped dye. The liposomes were then stirred in a solution of Con A at varying concentrations. The liposome by itself tends to be somewhat leaky, releasing about 17% of its content over a period of 11 h. In the presence of Con A, measurable increases in the amounts of leaked dye from the liposomes were observed. Several concentrations of Con A were examined, showing an increase in dye leakage with increasing protein concentration (see Supporting Information, Figure S2). At the highest Con A concentration, 50 µM, a total dye leakage of 36% was found over the 11 h period. The liposomes in the presence of BSA, on the other hand, yielded no increase in dye leakage. The increase in leakage observed with the addition of Con A may be coincident to an effect seen by others studying ion conductivity of bilayer lipid membranes induced by Con A24 but does not mimic any results with phosphatidyl(24) (a) Krull, U. J.; Brown, R. S.; Koilpillai, R. N.; Nespolo, R.; Safarzadeh-Amiri, A.; Vandenberg, E. T. Analyst 1989, 114, 33-40. (b) Deleers, M.; Poss, A.; Ruysschaert, J.-M. Biochem. Biophys. Res. Commun. 1976, 72, 709-713.

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choline small unilamellar vesicles3a or corked liposomes25 where rapid and complete dye leakage was observed. Con A binding to the bilayers indeed leads to some significant but limited membrane disruption. This might be expected for a protein insertion mechanism but does not suggest any deterioration of the membrane from adverse protein denaturation resulting in lysis or hole formation. To further understand the binding interaction of Con A with this glycolipid membrane, AFM imaging of the protein-bound membranes was conducted. We have previously shown that it is possible to monitor the nanoscale changes in lipid aggregation induced by chemical recognition at the membrane surface.11,12 Similarly, in the present study supported lipid bilayers composed of 5% PSMU/ DSPC were prepared at room temperature on a mica surface via vesicle fusion in a solution cell. The height of the fused membrane measured from the mica surface was approximately 55 Å, consistent with the thickness of a DSPC bilayer floating atop an 8-10 Å water layer.12,26 However, unlike previous membranes that used metal ion chelates (e.g., iminodiacetic acid, 18-crown-6) as headgroups on lipids to produce fused bilayer membranes with only a sparse density of nanometer-size holes, the present system yielded supported bilayers with distinct 10 nm wide gaps, having depths of 1-2 Å, throughout the surface (Figure 6A). Section plots of Figure 6 are shown in Figure S3 in the Supporting Information. The narrow width of the gaps does not permit full depth measurement due to the curvature of the AFM tip. Additional observation finds that the taller glycolipids, estimated to be ∼10 Å taller than DSPC from modeling studies, appear to be aggregated at the gap edges and imaged as the lighter shaded structures. These regions were, however, only 2-3 Å taller than the darker surrounding areas, assumed to be the DSPC matrix. Heating of the bilayer to 60 °C for 30 min resulted in the eventual coalescence of the membrane producing the image of Figure 6B. The lightest shaded regions, appearing in areas that were originally gaps, have become 5-6 Å taller than the DSPC matrix with the same 10 nm width of the original gap structure (Supporting Information, Figure S3B). This would suggest that the glycolipids have become more vertically oriented but still remain shorter than their estimated height difference (10 Å) against DSPC. This is in contrast with earlier studies of similar systems using iminodiacetic acid12 or 18-crown-611 functionalized lipids in a DSPC matrix where height differences corresponded with modeling study estimates of an extended triethyleneglycol spacer and headgroup. In the current system, however, the aggregated lipid structures are considerably different, exhibiting extremely narrow widths that frustrate reliable estimates of the true glycolipid height difference. The PSMU/DSPC bilayer exhibits interesting actuated behavior at the gap region upon AFM scanning. An image such as that seen in Figure 7A reversibly converted to that in Figure 7B, where the light areas corresponding to the glycolipids tend to nearly disappear during the scanning and then reappear after some time period. The gaps in the bilayer and the reversible structure formation of Figure 7 raise a few points for consideration. Why do these lipid membranes resist fusion at room temperature and why do the glycolipids appear to aggregate at the (25) Okahata, Y.; Nakamura, G.-i.; Noguchi, H. J. Chem. Soc., Perkin Trans. 2 1987, 1317-1322. (26) (a) Lis, L. J.; McAlister, M.; Fuller, N.; Rand, R. P.; Parsegian, V. A. Biophys. J. 1982, 37, 657-665. (b) Johnson, S. J.; Bayerl, T. M.; McDermott, D. C.; Adam, G. W.; Rennie, A. R.; Thomas, R. K.; Sackmann, E. Biophys. J. 1991, 59, 289-294.

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Figure 6. AFM images of 5% PSMU/DSPC lipid bilayers supported on a mica surface (A) after they have formed via vesicle fusion at room temperature, with line gaps evident in the structure, and (B) after heat treatment at 60 °C for 30 min, resulting in coalescence of the membrane (height scale ) 5 nm).

edges of these cracks? Moreover, why are the heights of the light areas, corresponding to the glycolipids at the gap edges, considerably shorter than the expected height difference with respect to the DSPC matrix and why do they remain shorter even after the membrane has coalesced with heat treatment? These questions and the unique behavior of this glycolipid membrane, although not apparently having any direct consequence toward the binding of Con A, are of interest since this unusual behavior has not been previously reported. A brief discussion of the possible membrane structure is as follows. The reversible behavior of the membrane at the gap regions might be explained with the illustration of Figure 8. The glycolipid, being both fluid phase at room temperature and rather cone shaped in structure owing to its large, hydrated carbohydrate headgroup, should tend to aggregate at the high curvature edge of the flat DSPC bilayer, which exists in the gel phase at room temperature. This highly hydrated and structurally stable edge would resist membrane fusion, thus leaving an open gap between sections of surface-fused bilayers. Because the glycolipids occupy the bilayer edge, the headgroups will be tilted with respect to the lipid bilayer and thus appear shorter than

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Figure 8. Schematic illustration of the PSMU/DSPC bilayer on the mica surface before and after the AFM tip scans over. The glycolipid tends to aggregate at the bilayer edges due to its fluid-phase nature and cone-shaped structure. The aggregates can be pushed down by the AFM tip, resulting in a loss of height of the structures.

Figure 7. AFM images of 5% PSMU/DSPC bilayers in the same scan area showing the change in height of the lighter shaded regions adjacent to the cracks in the bilayer. (A) Initially, these lighter shaded areas, considered as aggregates of glycolipid, are 2-3 Å taller than the DSPC bilayer membrane. (B) After scanning for a period of minutes, those areas have decreased to the same height as the DSPC matrix (height scale ) 5 nm).

if they were oriented perpendicular to the bilayer surface. If these glycolipids are aggregated at these edges, then the scanning AFM tip could push them down slightly into the bottom leaflet of the bilayer. The bulk of the glycolipid’s headgroup can be accommodated by the ca. 10 Å water layer that exists between the supported membrane and the mica surface. After some time, the bilayer relaxes and the glycolipids return to their original position. Such gaps were not found in fluid-phase bilayers of SOPC with PSMU or any other pyrene-labeled lipids we have previously studied. Instead, these membranes were featureless, indicative of a homogeneous lipid distribution. Hence, it is reasonable to assume that phase separation of the fluidphase PSMU lipid from the gel-phase DSPC matrix is responsible for the formation of these unique membrane features. The studies of Con A binding to PSMU/DSPC bilayers were performed on membranes that were fused to mica and coalesced by 60 °C heat treatment. To provide good visibility of the aggregated glycolipid structures by AFM, supported bilayers composed of 20% PSMU/DSPC were prepared. Con A was then added to the solution cell at a

concentration of 9 µM. Representative AFM images of the bilayer before and several hours after addition of Con A are shown in images A and B of Figure 9, respectively. The initial structure of the bilayer contains the same general dendridic pattern of thin lines composed of aggregated glycolipids as the 5% PSMU/DSPC bilayers of Figure 6B, but with a higher density of these structures. Furthermore, at the junction points of the dendritic lines were areas that appear to be high in concentration of glycolipids showing up as 60-90 nm wide dots or ellipsoids. The heights of these areas are ∼16 Å, significantly taller than the 5-6 Å heights measured on the dendrite lines and taller than the modeled height difference of 10 Å between PSMU and DSPC. The large height of the glycolipid regions is likely due to an effect of the dense packing of the carbohydrate headgroups. This observation with high-density receptor regions is not unique and has also been observed in some instances with our metalchelating lipid membranes.27 Our claim that the lightest shaded areas belong to aggregates of PSMU is further corroborated by the measurement of the overall area occupied by the lines and dots amounting to approximately 20% of the total membrane area. Several hours of incubation with Con A produces the image of Figure 9B. An increase in noise and loss of resolution were always observed after the addition of Con A, suggesting that the protein interferes with, and possibly adsorbs onto, the AFM tip. The lightest shaded features were consistently observed over multiple experiments following the addition of Con A, and thus we distinguish them as the protein bound to the membrane surface. Individual structures of circular shape and similar size as the Con A tetrameric structure can be found sporadically (27) Unpublished results.

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overlayer leaving only the bound structures as shown in Figure 9B. The initial behavior of the proteins may have some analogy with reports that Con A binds rapidly and nonspecifically to phosphatidylcholine lipid membranes with a Ka of ca. 105 M-1.29 Thus, the scanning action of the AFM tip may rid the surface of nonspecifically bound protein, leaving behind only the specifically or strongly bound species, such as those that are membrane inserted. This may also aid in explaining the observed decrease in resolution upon imaging of Con A on the bilayer surface as loosely bound or unbound protein may be collecting upon the AFM tip. Con A appears to bind to the PSMU/DSPC membrane as aggregates in structures coincident with the dendritic patterns that the initially observed glycolipid aggregates defined, but considerably wider and more diffuse. This is consistent with the notion of a dispersion of the glycolipid once bound to the lectin. The structures we observed are in contrast to AFM images reported previously of Con A bound to mannose-functionalized LB films.30 However, due to the 20-fold higher magnification reported herein a comparison is not straightforward. In Figure 9B, there also exist large patches of aggregated Con A approximately 200 nm in diameter that appear to correspond to areas that were originally high in density of glycolipid, as seen in Figure 9A. Thus, the AFM data support the claim of specific interaction of Con A to the glycolipid-rich areas in the membrane. However, protein insertion into the membrane, which is supported by the leakage studies and height measurements with the AFM, also seems to play an important role in the binding of Con A. How this protein insertion affects lipid redistribution in the membrane is unclear at this time, nor do we understand what form the Con A tetramer takes when bound to the membrane. Further clarification of these issues will be pursued in our future efforts.

Figure 9. AFM images of a 20% PSMU/DSPC membrane (A) before and (B) after 6 h of exposure time with 9 µM Con A in PBS buffer (pH 7.0) at room temperature. Arrows indicate structures of possible individual Con A tetramers (height scale ) 5 nm).

covering the surface. The measured diameter of the structure, which is 22 nm full width at half-maximum, is comparable to the 10 nm width of Con A considering image convolution from the AFM tip.28 Two of these Con A structures are indicated by arrows in Figure 9B. The height of these individual structures only measures about 1020 Å, considerably shorter than their known diameter, whereas large aggregates in the same image yield heights of ca. 50 Å. While the heights of the individual proteins are difficult to assess due to their small size and possible movement on a soft lipid membrane surface, the general observation of bound protein heights measuring 50 Å or less suggests that Con A does not exist as a tetramer atop the membrane surface. Rather, it either inserts into the membrane or, alternatively, denatures or binds as dimers against the membrane. AFM imaging alone, however, cannot make this distinction. Almost immediately after the addition of Con A and during the first few hours of imaging, the membrane surface was covered with what appears to be an overlayer of protein. At some time later, the surface loses this (28) Warner, M. J.; Gilchrist, M.; Schindler, M.; Dantus, M. J. Phys. Chem. B 1998, 102, 1649-1657.

Conclusion The ability to monitor molecular associations both optically and with nanoscale resolution provides a level of detail that is necessary to fully characterize proteinmembrane interactions. Our pyrene-labeled glycolipid PSMU allows one to evaluate the effect of specific protein interactions upon lipid associations within the membrane. Concanavalin A binding with the mannosyl headgroup of PSMU induced a dispersion of the glycolipid evidenced by both fluorescence and AFM monitoring of the system. Steric interactions between the protein-lipid complexes formed at the membrane and the physical separation of binding sites within the tetrameric structure are the likely reason for the observed dispersion of initially aggregated glycolipid, while possible protein insertion into the membrane may also contribute to lipid reorganization. The binding of protein and the subsequent lipid reorganization of the membrane were found to be slow, proceeding on the order of days. These studies suggest that membrane-protein associations are complex incorporating various phenomena, such as steric crowding, molecular mobility, and protein insertion. These issues are indeed difficult to measure and monitor but are important for understanding the basic principles of cell membrane biochemistry. Acknowledgment. The authors thank Dr. Paul Kotula for the TEM images of the liposomes. This research was supported in part by the Division of Materials Science (29) Lu¨scher-Mattli, M. Biopolymers 1987, 26, 1509-1526. (30) Bakowsky, U.; Rettig, W.; Bendas, G.; Vogel, J.; Bakowsky, H.; Harnagea, C.; Rothe, U. Phys. Chem. Chem. Phys. 2000, 2, 4609-4614.

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and Engineering in the Department of Energy Office of Basic Energy Sciences. A.S. thanks the Natural Sciences and Engineering Research Council of Canada and the Walter C. Sumner Foundation for scholarship support. Sandia is a multiprogram laboratory operated by Sandia Corporation, a Lockheed Martin Company, for the United States Department of Energy under Contract DE-AC0494AL85000.

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Supporting Information Available: TEM image of 5% PSMU/DSPC liposomes, carboxyfluorescein leakage study of 5% PSMU/DSPC liposomes in the presence of Con A at various concentrations, and AFM line scans. This material is available free of charge via the Internet at http://pubs.acs.org.

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