Biomimetic Molecular Assemblies on Glass and Mesoporous Silica

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Biomimetic Molecular Assemblies on Glass and Mesoporous Silica Microbeads for Biotechnology† Tione Buranda,*,§ Jinman Huang,‡ G. V. Ramarao,‡ Linnea K. Ista,‡ Richard S. Larson,§ Timothy L. Ward,‡ Larry A. Sklar,§ and Gabriel P. Lopez*,‡ Center for Micro-Engineered Materials, Department of Chemical and Nuclear Engineering, and Cancer Center and Department of Pathology, University of New Mexico, Albuquerque, New Mexico 87131 Received August 14, 2002. In Final Form: November 25, 2002 This paper describes the use of glass and mesoporous silica microspheres (typically 1-50 µm) as supports for biomimetic lipid bilayer membrane architectures for use in biotechnological applications. We present methods and characterization of lipid bilayer membranes supported on commercially available glass beads and mesoporous silica beads formed by an aerosol process that takes advantage of self-assembly of surfactant template phases in sol-gel synthesis. Methods for controlling the concentration of fluorescent lipids, ligands, receptors, and transmembrane proteins in the bead-supported bilayer assemblies are discussed, along with methods for measuring the concentration of these species using flow cytometry. Diffusion of molecular species both within the lipid bilayer and within the mesoporous bead structure is probed using fluorescence recovery after photobleaching. Flow cytometry and confocal fluorescence microscopy are used to examine dye uptake of the porous beads and the stability of the encapsulating lipid bilayer membranes to proton and fluorophore leakage. The studies presented herein form the basis for the use of several new types of biomimetic bead-supported bilayer architectures in a variety of biotechnological applications including microimmunoassays and fluorescence-based high-throughput screening of biochemical recognition and protein function.

Introduction There is widespread interest in the design and use of microbeads (i.e., synthetic ceramic or polymeric microspheres of diameters of 1-50 µm) as solid-state supports for biomolecular assemblies and molecular recognition systems in a wide variety of biotechnological applications including biochemical synthesis, drug discovery, genomics, proteomics, biosensing, separations, and drug delivery.1-23 * To whom correspondence should be addressed. E-mail: gplopez@ unm.edu, [email protected]. † Part of the Langmuir special issue entitled The Biomolecular Interface. ‡ Center for Micro-Engineered Materials, Department of Chemical and Nuclear Engineering. § Cancer Center and Department of Pathology. (1) Albert, K. J.; Walt, D. R.; Gill, D. S.; Pearce, T. C. Anal. Chem. 2001, 73, 2501-2508. (2) Andersson, H.; Jonsson, C.; Moberg, C.; Stemme, G. Electrophoresis 2001, 22, 3876-3882. (3) Bayerl, T. M.; Bloom, M. Biophys. J. 1990, 58, 357-362. (4) Buranda, T.; Lopez, G. P.; Keij, J.; Harris, R.; Sklar, L. A. Cytometry 1999, 37, 21-31. (5) Buranda, T.; Lopez, G. P.; Simons, P.; Pastuszyn, A.; Sklar, L. A. Anal. Biochem. 2001, 298, 151-162. (6) Buranda, T.; Huang, J.; Perez-Luna, V. H.; Schreyer, B.; Sklar, L. A.; Lopez, G. P. Anal. Chem. 2002, 74, 1149-1156. (7) Dickinson, T. A.; Michael, K. L.; Kauer, J. S.; Walt, D. R. Anal. Chem. 1999, 71, 2192-2198. (8) Eldefrawi, M. E.; Azer, N. L.; Nath, N.; Anis, N. A.; Bangalore, M. S.; KP, O. C.; Schwartz, R. P.; Wright, J. Appl. Biochem. Biotechnol. 2000, 87, 25-35. (9) Fan, Z. H.; Mangru, S.; Granzow, R.; Heaney, P.; Ho, W.; Dong, Q. P.; Kumar, R. Anal. Chem. 1999, 71, 4851-4859. (10) Harma, H.; Aronkyto, P.; Lovgren, T. Anal. Chim. Acta 2000, 410, 85-96. (11) Hetzer, M.; Heinz, S.; Grage, S.; Bayerl, T. M. Langmuir 1998, 14, 982-984. (12) Jiang, G. F.; Harrison, D. J. Analyst 2000, 125, 2176-2179. (13) Lee, M.; Walt, D. R. Anal. Biochem. 2000, 282, 142-146. (14) Levy, D.; Mosser, G.; Lambert, O.; Moeck, G. S.; Bald, D.; Rigaud, J. L. J. Struct. Biol. 1999, 127, 44-52. (15) Linseisen, F. M.; Hetzer, M.; Brumm, T.; Bayerl, T. M. Biophys. J. 1997, 72, 1659-1667.

In several of these applications, it is becoming increasingly desirable to construct biomolecular assemblies of higher complexity to mimic the biochemical functions of cellular systems. In this report, we describe the fundamental aspects of a biomimetic approach that aims to enable such biomolecular assemblies of high complexity. Glass and polymeric beads functionalized with various chemical (or biological) groups are commercially available. However, it is often the case that beads of a given characteristic (e.g., size, composition, appropriate surface (bio)chemical modification) that are required by a particular application are not commercially produced. One goal of this work is to present facile methods of biofunctionalizing beads with user-defined size and characteristics to meet the varied needs of fundamental research. In the current state of the art, beads of a required size are not always commercially available. For example, in recent flow through microimmunoassay applications, beads ranging in size from 2 to 50 µm in diameter have been used,6,21,24,25 (16) Loidl-Stahlhofen, A.; Hartmann, T.; Schottner, M.; Rohring, C.; Brodowsky, H.; Schmitt, J.; Keldenich, J. Pharm. Res. 2001, 18, 17821788. (17) Loidl-Stahlhofen, A.; Schmitt, J.; Noller, J.; Hartmann, T.; Brodowsky, H.; Schmitt, W.; Keldenich, J. Adv. Mater. 2001, 13, 18291834. (18) Ohmura, N.; Lackie, S. J.; Saiki, H. Anal. Chem. 2001, 73, 33923399. (19) Phillips, T. M. J. Biochem. Biophys. Methods 2001, 49, 253262. (20) Potter, R. M.; Key, T. A.; Gurevich, V. V.; Sklar, L. A.; Prossnitz, E. R. J. Biol. Chem. 2002, 277, 8970-8978. (21) Sato, K.; Tokeshi, M.; Odake, T.; Kimura, H.; Ooi, T.; Nakao, M.; Kitamori, T. Anal. Chem. 2000, 72, 1144-1147. (22) Sklar, L. A.; Vilven, J.; Lynam, E.; Neldon, D.; Bennett, T. A.; Prossnitz, E. Biotechniques 2000, 28, 976. (23) Stuart, M. C. A.; Reutelingsperger, C. P. M.; Fredrick, P. M. Cytometry 1998, 33, 414-419. (24) Sato, K.; Tokeshi, M.; Kimura, H.; Kitamori, T. Anal. Chem. 2001, 73, 1213-1218. (25) Hayes, M. A.; Polson, N. A.; Phayre, A. N.; Garcia, A. A. Anal. Chem. 2001, 73, 5896-5902.

10.1021/la026405+ CCC: $25.00 © 2003 American Chemical Society Published on Web 01/17/2003

Biomimetic Molecular Assemblies

where commercial sources typically offer biofunctionalized beads of less than 20 µm in size. While it is often useful for a user to define such beads in terms of site coverage of biomolecules that are obtained after a preparation, the surface or compositional characteristics of commercially available beads may not be suited to the desired biochemical modification. Researchers often resort to using nonspecific adsorption methods as a means of biofunctionalization. These methods can result in a number of potentially active sites being eliminated due to protein denaturation, inactive protein orientation, or receptor displacement and thus make control of receptor site density problematic. Lipid bilayers supported on flat or spherical silica substrates continue to be the subject of research activity as models of cell membranes and as biomimetic membrane platforms for biotechnological applications.3,15,26-32 Supported bilayer membranes retain many of the characteristics of natural systems such as lateral fluidity and impermeability to ionic species. On hydrophilic glass surfaces, the typical membrane is generally believed to be separated from the substrate by a thin (10-30 Å) layer of water and, like the natural membrane, to possess macroscopically long-range fluidity with mobile components of both leaflets freely diffusing over the entire surface of the support.33-35 This paper reports on the biofunctionalization of glass and porous silica beads through the application of supported lipid bilayers that incorporate ligands, fluorescent dyes, and transmembrane proteins. The goals of this report are threefold. The first is to present general methods for controlling and measuring the site density of receptors on microbeads coated with fluid bilayer membranes containing mixtures of phospholipids. The second is to demonstrate that transmembrane proteins can easily be incorporated into these supported lipid bilayer architectures. Finally, this paper aims to characterize and establish a new class of mesoporous silica beads as versatile hosts for supported lipid bilayer membranes in which the porous structure of the beads forms an isolated cytosol-like compartment that can be used to store ions, dyes, drugs, and biological molecules and thus increase the functional versatility of the microbeads in biotechnology. We have recently developed a versatile method for forming monodisperse silica microbeads that exhibit a well-defined ordered mesoporous nanostructure via an aerosol synthesis that exploits sol-gel templating of selfassembled surfactant phases.36 It was recently suggested that porous microbeads coated with lipid bilayers can form the basis for rapid screening of biomolecular interactions.17 We view the present study as a first, necessary step in the utilization of the mesoporous microbeads formed by aerosol-assisted surfactant self-assembly and sol-gel (26) Merkel, R.; Sackmann, E.; Evans, E. J. Phys. 1989, 50, 15351555. (27) Sackmann, E. Science 1996, 271, 43-48. (28) Kung, L. A.; Kam, L.; Hovis, J. S.; Boxer, S. G. Langmuir 2000, 16, 6773-6776. (29) Boxer, S. G. Curr. Opin. Chem. Biol. 2000, 4, 704-709. (30) Groves, J. T.; Boxer, S. G. Acc. Chem. Res. 2002, 35, 149-157. (31) Hirn, R.; Schuster, B.; Sleytr, U. B.; Bayerl, T. M. Biophys. J. 1999, 77, 2066-2074. (32) Kasbauer, M.; Junglas, M.; Bayerl, T. M. Biophys. J. 1999, 76, 2600-2605. (33) Groves, J. T.; Boxer, S. G.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1998, 95, 935-938. (34) Smith, L. M.; Parce, J. W.; Smith, B. A.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1979, 76, 4177-4179. (35) Subramaniam, S.; Seul, M.; McConnell, H. M. Proc. Natl. Acad. Sci. U.S.A. 1986, 83, 1169-1173. (36) Rama Rao, G. V.; Lo´pez, G. P.; Bravo, J.; Pham, H.; Datye, A. K.; Xu, H.; Ward, T. L. Adv. Mater., submitted.

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templating as versatile platforms for high-throughput assays of protein function based on fluorescence measurements, including flow cytometry and spectrofluorimetry. Materials and Methods L-R-Phosphotidylcholine (egg PC), 1,2-dioleoyl-sn-glycero-3phosphoethanolamine-N-(carboxyfluorescein) (fluorescein PE), and 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(biotinyl) (biotin PE) were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL). Glass beads (5, 10, and 20 µM diameter) were obtained in dry form from Duke Scientific Corp. (Palo Alto, CA). Biotin, fluorescein, and rhodamine 6G were purchased from Molecular Probes (Eugene, OR) and used without further purification. Texas Red labeled and fluorescein-labeled streptavidin was purchased from Pierce Chemicals (Rockford IL). Biotinylated, fluorescein-labeled FLAG peptide (DYKDDDDK) was synthesized in house as described previously.5 ICAM-1 was extracted and purified from human tonsils as described elsewhere.37 Monoclonal antibodies (R6.5) directed against ICAM-1 were purified from hybridoma lines (ATCC, Rockville, MD) as previously described.38 Preparation of Glass Beads. To obtain clean hydrophilic surfaces, glass beads were washed and centrifuged several times with ethanol and deionized water, respectively. Subsequently, the beads were suspended in a 40 mL solution of 4% H2O2 and 4% NH4OH and placed in an 80 °C water bath for 10 min. After being rinsed with distilled water, the beads were resuspended in a 40 mL 4% H2O2, 0.4 M HCl solution and placed in the 80 °C water bath for 10 min. The beads were then rinsed with deionized water several times. After final centrifugation, the beads were resuspended in 10 mL of Tris buffer (50 mM Tris, 50 mM NaCl, pH 7.4). A hemocytometer was used to determine the number of beads per unit volume. Mesoporous Silica Beads. Porous beads were prepared as described elsewhere.36 Briefly, precursor solutions were prepared by addition of Brij-58 (CH3(CH2)15-(OCH2CH2)20-OH, Aldrich) to an acidic silica sol (A2**), as reported by Lu et al.39 In a typical preparation, tetraethyl orthosilicate (TEOS, Aldrich), ethanol, deionized water, and dilute HCl (mole ratios 1:3.8:1:0.0005) were refluxed at 60 °C for 90 min to provide the stock sol. Then, 10 mL of stock sol was diluted with ethanol, followed by addition of water, dilute HCl, and aqueous surfactant solution (1.5 g of surfactant dissolved in 20 mL of water) to provide final overall TEOS/ethanol/H2O/HCl/surfactant molar ratios of 1:22:55:0.0053: 0.06. This sol was stirred for about 10 min before beginning a powder synthesis run. Monodisperse droplets were generated by means of a vibrating orifice aerosol generator (TSI model 3450). The solution was forced through a small orifice (20 µm diam) by a syringe pump, with syringe velocities of approximately 8 × 10-4 cm/s (∼4.7 × 10-3 cm3/s). This delivery rate was adjusted to provide a stable operating pressure of 340-420 kPa. The liquid stream was dispersed into uniform droplets by the vibrating orifice using a frequency range of 40-200 kHz, with the final setting adjusted to eliminate satellite droplets. The droplets were then injected axially along the center of a turbulent air jet to disperse the droplets and to prevent coagulation. Following the mixing of the dispersed droplets with a much larger volume of filtered dry air, the droplet-laden gas stream flowed through a 2.5 cm diameter quartz tube into a three-zone furnace (0.9 m heated length) maintained at 500 °C (A2** runs) or 420 °C (TEOS solution runs). This provided a mean residence time of approximately 0.3 s in the heated zone. The particles were collected on a filter maintained at approximately 80 °C by a heating tape. Collected particles were calcined in air at 400-450 °C for 4 h to remove the surfactant template. Prior to use in further experimentation, the porous beads were washed in deionized water. Preparation of Unilamellar Lipid Vesicles. Small unilamellar vesicles (SUVs) were prepared from a 1 mM solution of

(37) Larson, R. S.; Corbi, A. L.; Berman, L.; Springer, T. J. Cell Biol. 1989, 108, 703-712. (38) Larson, R. S.; Brown, D. C.; Sklar, L. A. Blood 1997, 90, 27472756. (39) Lu, Y.; Fan, H.; Stump, A.; Ward, T. L.; Reiker, T.; Brinker, C. J. Nature 1999, 398, 223.

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egg phosphatidylcholine (egg PC) in chloroform (200 µL total volume) in a clear glass tube.3 Dry nitrogen gas was bubbled through the solution to dryness, leaving a white film at the bottom of the glass tube. The film was subsequently vacuum-dried at room temperature for half an hour. After addition of 1 mL of Tris buffer (pH 7.5), the vesicle suspension was sonicated to optical clarity in a sonication bath (Branson Cleaning Equipment Co., Shelton, CT). Biotin PE (e.g., 0.001 mol % of egg PC) and fluorescein PE (0.01 mol %) were also included in some of the membrane preparations for specific experiments described below. Preparation of Microsphere-Supported Lipid Bilayers (Lipobeads). Beads were added to the SUV dispersions with vortexing for 2 min in microfuge tubes. In this manner, the SUVs are reported3 to spontaneously collapse into a continuous bilayer surrounding the beads. After sitting for 10 min, the beads were then centrifuged and resuspended in buffer, repeating five times to remove unbound SUVs, thus leaving single bilayer covered beads.3 For lipid compositions containing biotin PE, molar excess streptavidin was added to aliquots of lipobeads and incubated with vortexing for 30 min to allow streptavidin to firmly bind to the biotin on the beads. The samples were centrifuged and resuspended in 1 mL of Tris buffer (100 mM Tris-HCl, 150 mM NaCl, 0.1% bovine serum albumin (BSA), pH 7.4) three times to remove unbound streptavidin. Incorporation of ICAM-1 into Bead-Supported Lipid Bilayer Membranes. Proteins were reconstituted with egg PC by detergent dialysis.40 SUVs prepared from 0.1 mM egg PC were serially diluted over 3 decades in 50 mM Tris buffer (1% octaglucoside) to create samples spanning the 10-5-10-7 M concentration range in SUV dispersions. Detergent-solubilized intercellular adhesion molecule-1 (ICAM-1, ≈10.0 nM) was added to each of the vesicle dispersions in final volumes of 150 µL. The various samples of lipid-protein complexes were transferred to dialysis capsules (Pierce) and dialyzed overnight at 4 °C against detergent-free Tris buffer. The detergent-depleted samples of lipid-protein complexes were transferred to microfuge tubes. Then, 5.2 µm glass beads (4.0 × 105 beads in 0.5 µL volume suspensions) were added to the microfuge tubes, with brief vortexing. The samples were left standing for 30 min. This was followed by a brief centrifugation and resuspension of lipobeads in buffer, repeating at least five times to remove unbound lipids. The lipid/protein-bearing beads were finally resuspended in blocking buffer (0.5% BSA containing 50 mM Tris buffer) for an hour. Fluorescein-labeled anti-ICAM-1 monoclonal antibodies (R6.5) were added to the bead samples for 10 min prior to analysis by flow cytometry. Confocal Microscopy. Samples were viewed and imaged using a Zeiss LSM510 system equipped with an argon ion laser. Additional excitation sources included HeNe1 and HeNe2 lasers for the respective 538 and 633 nm excitation wavelengths. Fluorescence recovery after bleaching (FRAP) experiments were performed at room temperature (∼20 °C). Imaging experiments were typically performed using a ×63/1.3 oil immersion objective with the diaphragms set to allow the probing of ∼1 µm wide vertical slices of the imaged beads. Flow Cytometry. Bead suspensions were analyzed by flow cytometry using a Becton-Dickinson FACScan flow cytometer (Sunnyvale, CA) interfaced to a Power PC Macintosh using the CellQuest software package. The FACScan is equipped with a 15 mW air-cooled argon ion laser. The laser output is fixed at 488 nm. Experimental details of these analyses have been described elsewhere.4,41 We have shown elsewhere how molecular assemblies on beads can be analyzed in quantitative fashion by flow cytometry. The average fluorescence on a single bead is converted to the number of fluorophores per bead on the basis of flow cytometric calibration beads.4,41

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Figure 1. Binding of biotinylated peptide to lipobeads. (A) Representative binding curve (raw flow cytometric data) of a fluorescently labeled, biotinylated FLAG peptide binding to streptavidin on 10 µm beads (200 000) coated with lipid bilayers (0.001 mol % biotin PE). Open circles: total FLAG peptide bound. Closed circles: nonspecific binding of FLAG peptide to biotin-blocked streptavidin. (B) The analysis of the binding data to obtain a dissociation constant of ≈0.04 nM.

Control of Receptor Site Densities on Lipobeads. An advantage of using beads as platforms for bilayersupported assemblies is the ready control of the number

of biotin sites that is afforded by managing stoichiometric relationships. In these experiments, varying the mole fraction of biotin PE that is mixed with egg PC regulates the number of receptors per bead. Thus bead-supported bilayers derived from a mixture of egg PC and biotin PE of known stoichiometry can result in a bead population with a reasonably defined surface coverage of biotin (and thus streptavidin and streptavidin-linked biotinylated receptors). A desired surface coverage of biotin on supported bilayers can be achieved by determining the total number of lipids in the bilayer surrounding a single bead (Scheme 1). This is established from dividing the surface area of the bead (4πr2) by the area of the average lipid headgroup (≈55 Å2).3 The desired surface coverage of biotin PE is then simply effected by using the appropriate mole fraction of biotin PE in the vesicle preparation. By extension of this approach, one can also ensure that an excess amount of lipids (at least 10-fold) is always mixed with beads. Figure 1A shows a representative binding curve (raw flow cytometric data) of a fluorescently labeled, biotinylated FLAG peptide binding to 10 µm glass beads coated with lipid bilayers (0.001 mol % biotin PE bound to streptavidin). The method for analysis of flow cytometry data from similar beads has been described in detail elsewhere.4,5,41 Figure 1B shows the analysis of the binding data to obtain the dissociation constant of ≈0.04 nM. From the law of mass action, the concentration of bound peptide is determined at saturation to be ≈1.0 nM. This value is in agreement with the receptor concentration expected from the use of 200 000 beads in each of the 200 µL samples.42

(40) Mimms, L. T.; Zamphighi, G.; Nozaki, Y.; Tanford, C.; Reynolds, J. A. Biochemistry 1981, 20, 833-840. (41) Buranda, T.; Jones, G.; Nolan, J.; Keij, J.; Lopez, G. P.; Sklar, L. A. J. Phys. Chem. B 1999, 103, 3399-3410.

(42) The calculation is as follows: surface area of 10 µm beads ) 3.14 × 1010 Å2; lipids per bead ) 3.14 × 1010/55 ) 5.7 × 108; biotins per bead (0.001 mol %) ) 5.7 × 105; molarity of streptavidin sites (200 000 beads in 200 µL) ) 0.9 nM.

Results


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Scheme 1. Lipid Bilayer Assembly of L-r-Phosphatidylcholine (egg PC) and 1,2-Dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(biotinyl) (biotin PE) on Glass Beadsa

a The stoichiometry of egg PC and biotin PE can be used to regulate the number of streptavidins and consequently limit the number of biotinylated ligands that can be bound to each bead. The limit of the desired number of receptor sites can be empirically determined prior to experiment from the known surface area of the beads and the total site occupancy of closely packed lipids that comprise the bilayer.

Figure 2 demonstrates the varying stoichiometry based on fluorescence resonance energy transfer (FRET). FRET is a distance-dependent interaction between the electronic excited states of two dye molecules in which excitation is transferred from an excited donor (D*) molecule (e.g., fluorescein) to an acceptor (A) molecule (e.g., Texas Red) without emission of a photon. The characteristic distance at which the donor fluorescence and FRET are equally probable is defined as R0.43 The R0 value for the fluorescein and Texas Red donor/acceptor pair has been calculated to be ≈46 Å.5 For FRET donor and acceptor molecules randomly distributed on a surface, the average surface density of (43) Lakowicz, J. R. Principles of Fluorescence Spectroscopy, 2nd ed.; Plenum Press: New York, 1999.

acceptors must be less than ≈0.02 acceptors per R02 so that there is little random transfer when the donor and acceptor are not aggregated.44,45 The results shown in Figure 2 demonstrate that surface coverage is dependent on the fraction of biotin PE and that the distribution of the streptavidin molecules on the fluid lipid bilayer surface is random based on the lack of FRET when the number of acceptors per R02 is less than 0.02 (Figure 2D). Fluidity of Supported Bilayers. Figure 3A shows a fluorescence micrograph of a homogeneous lipid bilayer surrounding a bead when viewed as a 1 µm slice at the focal depth at the equator of a bead. The photobleaching (44) Sklar, L. A.; Doody, M. C.; Gotto, A. M.; Pownall, H. J. Biochemistry 1980, 19, 1294-1301. (45) Wolber, P. K.; Hudson, B. S. Biophys. J. 1979, 28, 197-210.

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Figure 2. Dependence of FRET efficiency on acceptor density/R02 for R0 ≈ 50 Å. Flow cytometric histograms representing detection of fluorescein (green) and Texas Red (red) tagged streptavidins on beads. (A) At 0.3 mol % biotin PE, fluorescence from a 50/50 mixture of fluorescein-tagged and dark streptavidins. The mean of the histogram associated with the fluorescein donor is 366. (B) A 50/50 mixture (≈0.1 A/R02)70 of fluorescein and Texas Red tagged streptavidins indicating FRET. The mean fluorescein histogram is 260 (quenched by ≈30%). The extent of FRET is consistent with theoretical expectations (refs 44 and 45). (C) At 0.03 mol % biotin PE, fluorescence from a 50/50 mixture of fluorescein-tagged and unlabeled streptavidins. (D) A 50/50 mixture (≈0.01 A/R02) of fluorescein and Texas Red tagged streptavidins indicating no FRET.

Figure 3. Lateral fluidity of lipid bilayers on 10 micron beads. The bilayer membranes are comprised of egg PC and 0.2 mol % fluorescein PE. (A) Confocal micrographs showing photobleach and time-resolved recovery of fluorescence intensity of lipid bilayers. Overall sample bleaching is accounted for in the analysis of the FRAP data by normalizing the spot bleached intensity data to intensities on the opposite pole of the bead. (B) Plot of normalized intensity recovery after spot bleaching (D ≈ 8.0 × 10-8 cm2 s-1).

and recovery of a target region can be monitored in parallel with the region opposite; thus the effects of the overall loss in intensity can be accounted for in the analysis of the fluorescence recovery. Figure 3B shows the photobleaching of a 2 µm wide by 1 µm vertical slice of lipid membrane (with fluorescein PE) in confocal mode and subsequent

complete recovery after 3 min. The diffusive rate constant of the lipid components is obtained, in the simple limit of two-dimensional diffusion into a circular spot, from simple expressions such as the one by Axelrod et al.:46 D (cm2/s) ) γr2/4τ1/2 where γ is a correction factor for the amount of bleaching, r is the radius of the bleached area, and τ1/2 is the half-life for fluorescence recovery. In the case described here, γ is estimated to be on the order of 1350, a quantity that corresponds to the number of lipid monolayers in the (1 µm) slice of bleached space. This is based on the surface area (55 Å2)3 of a lipid headgroup. For lipids that bore fluorescent streptavidin molecules, the bleach recovery was notably slower by an order of magnitude (data not shown). This may be related to the possibility that the streptavidin may be bivalently bound, a fact that would require cooperativity in the lateral translation of the attached lipid pair. Incorporation of ICAM-1 into Bead-Supported Lipid Bilayer Membranes. ICAM-1 has several wellmapped antibody recognition sites for which antibodies are readily available.47-49 It is important to note that although no direct test of biological function of ICAM-1 was performed here, the functionality of this protein has been confirmed elsewhere50-52 when prepared under similar conditions as described and used on plastic substrates. In addition, laterally mobile complexes of major histocompatibility-peptide bound ICAM-1 in supported (46) Axelrod, D.; Koppel, D. E.; Schlessinger, J.; Elson, E.; Webb, W. W. Biophys. J. 1976, 16, 1055-1069. (47) Staunton, D. E.; Marlin, S. D.; Stratowa, C.; Dustin, M. L.; Springer, T. A. Cell 1988, 52, 925-933. (48) Staunton, D. E.; Merluzzi, V. J.; Rothlein, R.; Barton, R.; Marlin, S. D.; Springer, T. A. Cell 1989, 56, 849-853. (49) Staunton, D. E.; Dustin, M. L.; Erickson, H. P.; Springer, T. A. Cell 1990, 61, 243-254. (50) Brown, D. C.; Tsuji, H.; Larson, R. S. Br. J. Haematol. 1999, 107, 86-98. (51) DiVietro, J. A.; Smith, M. J.; Smith, B. R. E.; Petruzzelli, L.; Larson, R. S.; Lawrence, M. B. J. Immunol. 2001, 167, 4017-4025. (52) Winter, S. S.; Sweatman, J. J.; Lawrence, M. B.; Rhoades, T. H.; Hart, A. L.; Larson, R. S. Br. J. Haematol. 2001, 115, 862-871.


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Figure 5. Optical micrographs of commercially available glass beads (10 ( 1 µm, top) and mesoporous silica beads (11 ( 1 µm, bottom). Figure 4. (A) Display of ICAM-1 by titration of lipid vesicle/ protein mixtures to 400 000 beads suspended in 150 µL by flow cytometry. Sample a has no ICAM-1 added. Samples b-d are mixed with 10 nM of ICAM-1. The mean channel fluorescences correspond to the binding of labeled antibodies to ICAM-1 on beads (inset). The columns in each set refer to the concentration of antibody used to detect the protein: (1) 67.0 nM, (2) 33.0 nM, and (3) 6.7 nM. (B) Relative intensity of R6.5 antibodies bound to ICAM-1 on beads versus fraction of ICAM-1 in 10-6 and 10-5 M egg PC vesicle dispersions (samples c and d in (A)). Numbers 1-3 are defined in (A). The analysis suggests a nonstoichiometric correlation between the amount of ICAM-1 displayed on beads and the fraction of ICAM-1 in the original vesicle dispersions.

membranes have been shown to undergo active immunological synapse formation with living T lymphocytes.53 The reconstitution of ICAM-1 in supported lipid bilayers on beads was probed by the use of fluorescein-labeled R6.5 monoclonal antibodies. The results are summarized in Figure 4. The antibody binding data show that the surface coverage of the protein varies according to the fraction of protein in the lipid vesicle dispersions. The variation in the amount of fluorescent antibodies bound to the ICAM1-bearing lipids (columns b-d in Figure 4A) is related to (1) stoichiometric differences between the lipid-protein complexes and (2) surface area limited “lipid binding sites” on ≈400 000 beads (≈6.2 × 10-7 M).54 (53) Grakoui, A.; Bromley, S. K.; Sumen, C.; Davis, M. M.; Shaw, A. S.; Allen, P. M.; Dustin, M. L. Science 1999, 285, 221-227. (54) This value is estimated from the number of lipid headgroups (55 Å2) that can be closely packed on the surface area defined by 400 000 beads (5.2 µm), in a 150 µL volume suspension.

On the basis of the behavior of the lipid vesicles alone, one might expect detergent-solubilized proteins, homogeneously dispersed in unilamellar lipid vesicles, to reconstitute onto beads in a way that is directly correlated to the lipid-to-protein ratio in the vesicle dispersions. Thus, fewer proteins are displayed on beads when the concentration of the lipid/protein complexes in the vesicle dispersions is less or much greater (>10×) than the beadlimited binding sites (columns b and d, respectively, in Figure 4A). The optimal amount of ICAM-1 is displayed (column c) when the lipid/protein dispersions are within an order of magnitude greater than the putative binding sites on the beads. To determine whether the ICAM-1 is reconstituted on beads in the same mole fraction as that in the original vesicle dispersions, we have plotted (Figure 4B) the relative amount of R6.5 antibody bound to ICAM-1 on beads versus the original mole fraction of ICAM-1 in the vesicle dispersions. We have used background-corrected data from samples c and d in Figure 4A. On the basis of this limited analysis, the reconstitution of ICAM-1 on beads appears not to be stoichiometrically equal to the amount of protein in the vesicle dispersions. Mesoporous Beads: Membrane Coverage and Analysis of Void Space. The synthesis and physical characteristics of these beads have been described in detail elsewhere.36 Figure 5 presents optical micrographs of both the mesoporous silica beads and the commercially obtained glass beads used for comparative analysis. Figure 6 presents fluorescence histograms obtained from flow cytometry of the fluorescent lipid bilayer supported on porous and solid beads. The similarity of the histograms obtained from the two bead populations indicates similarity in the character of the lipid bilayer architectures formed

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Figure 8. Quantitation of the average void volume of porous beads. Plot of the average absorbed fluoresceins per bead as determined with flow cytometry versus the supernatant concentrations in molecules per liter. The slope of the linear plot is equivalent to the average dye-accessible void volume of the bead samples (≈10 fL).

Figure 6. Flow cytometry histograms obtained from fluorescent bilayer membranes (egg PC and 0.01 mol % fluorescein PE) on commercially available glass beads (top) and mesoporous beads (bottom). The X-axis indicates the fluorescence intensity of the beads, and the Y-axis (event counts) indicates the number of beads with a given intensity.

Figure 7. Flow cytometry dot plots for laser scattering by porous beads (top) and solid glass beads (bottom). The forward scatter (FSC-H, X-axis) is related to the size of the particles. The side scatter (SSC-H, Y-axis) is related to the internal structure (or granularity) of the particles.

on the two types of beads. This similarity suggests that there is not appreciable absorption of the lipid bilayer structure into the porous network of the mesoporous beads. Figure 7 shows flow cytometry dot plots of porous beads and solid glass beads (both ∼10 µm in diameter). The flow cytometer can distinguish particles on the basis of size (FSC in Figure 7) and the internal structure (or granularity) (SSC in Figure 7).55 The dot plots indicate similarity in size and distribution between the commercial beads

(mean diameter of 10 ( 1 µm) and the porous beads (mean diameter of 11 ( 1 µm). The void space of the porous beads is indicated by the lower reading on the granularity scale compared to the result obtained for the similar-sized solid beads. The mesoporous beads were further characterized in terms of their capacity to absorb dye molecules and as robust supports for lipid bilayers. To obtain a reasonable estimate of the average dye-accessible volume of the void space of the porous beads, the beads were incubated in solutions of fluorescein dye (nM-µM concentration range) for 72 h. The beads were centrifuged and washed twice in the 50 mM Tris buffer, after which they were immediately coated with lipid bilayer (egg PC), thus trapping the dye inside. The beads were then analyzed with flow cytometry. Bilayer-coated beads with variable concentrations of fluorescein dye trapped inside their voids were analyzed at the flow cytometer to determine the average amount of dye molecules trapped in the beads. The means of the fluorescence histograms from the flow cytometric analysis were converted to the number of dye molecules per bead using standard fluorescent beads.41 The data are plotted against the original concentration of the fluorescein in the supernatants in which the beads had been equilibrated (Figure 8). The plot of the average absorbed fluoresceins per bead versus the supernatant concentrations was linear for those beads that were allowed to reach equilibrium with the supernatant (72 h incubation), whereas the shorter incubations often exhibited plateaus at higher solution concentrations. This characteristic suggests a relatively slow diffusional exchange of fluorophores between the bulk solution and the confined volume of the porous beads. In this analysis, the slope of the graph in Figure 8 may be considered to be directly proportional in magnitude to the void space that is accessible to the trapped dye. In this manner, the average dye-accessible void volume for each bead was determined to be on the order of 10 fL, comprising a rather small fraction of the total volume (≈3.0 pL) of the average 10 µm bead. The relationship of the slowly accessible volume and the architecture of the beads remains to be clarified. Figure 9 shows confocal micrographs of porous beads containing rhodamine 6G dye. In addition, the beads are coated with a lipid bilayer (egg PC and 0.01 mol % fluorescein PE). The lipid bilayer membranes tended to be uniform over the entire bead surface. Furthermore, (55) Murphy, R. F. Ligand binding, endocytosis, and processing; Wiley-Liss: New York, 1988.


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Figure 9. Confocal micrographs of (egg PC and 0.01 mol % fluorescein PE) lipid bilayer membrane supporting porous beads (11 µm diam) filled with rhodamine 6G dye. The accessibility of the internal volumes of the beads is variable: (a) has limited access to the rhodamine dye as indicated by the dark core, (b) and (c) represent the intermediate case, and (d) represents those beads whose core is most accessible to dye molecules.

the bilayer appeared to be impermeable to protons or dye. As an example, beads that were filled with fluorescein dye and covered with an egg PC lipid bilayer did not lose their intensity over time (48 h) nor did exposure to a pH lower than 7.5 (e.g., pH 6.0) affect the intensity of the lipid-bilayer-trapped pH sensitive dye (fluorescein).56 The dye accessibility to the mesoporous volume was variable based on flow cytometry and microscopy measurements. This diversity in accessibility is illustrated among the beads marked a-d in Figure 9. The internal space of bead a has limited access to the rhodamine dye as indicated by the dark core. Beads b and c represent the intermediate case, while bead d represents those beads whose cores are most accessible to dye molecules. Such variability was also apparent in the width of the flow cytometry fluorescence histograms (not shown). The connectivity of the internal pore space was probed by confocal microscopy by imaging the cross section of a bead in 1 µm optical slices along the z-axis. The typical cross section of the typical bead as represented by Figure 10A suggests a homogeneous distribution of the trapped rhodamine dye. This result indicates that while accessibility of the dye to different porous beads may vary, the distribution of dye within the porous structure of each bead is fairly uniform. In addition, the recovery of the fluorescence intensity of a bleached spot inside the bead reflects the pore connectivity within the internal volume of the bead (Figure 10B). Discussion This study is part of an ongoing effort to obtain broad access to well-characterized biofunctional particulate surfaces for use in basic and applied research.4-6,22,41,57-63 As the mechanism by which vesicles spontaneously form (56) Huang, J.; Buranda, T.; Sklar, L. A.; Lopez, G. P. Work in progress. (57) Sklar, L. A.; Edwards, B. S.; Graves, S. W.; Nolan, J. P.; Prossnitz, E. R. Annu. Rev. Biophys. Biomol. Struct. 2002, 31, 97-119. (58) Jackson, W. C.; Kuckuck, F.; Edwards, B. S.; Mammoli, A.; Gallegos, C. M.; Lopez, G. P.; Buranda, T.; Sklar, L. A. Cytometry 2002, 47, 183-191. (59) Nolan, J. P.; Buranda, T.; Cai, H.; Kommander, K.; Lehnert, B.; Nolan, R.; Park, M. S.; Ruscetti, T.; Shen, B.; Sklar, L. A. Proc. SPIEInt. Soc. Opt. Eng. 1998, 3256, 114-121. (60) Nolan, J. P.; Lauer, S.; Prossnitz, E. R.; Sklar, L. A. Drug Discovery Today 1999, 4, 173-180. (61) Nolan, J. P.; Sklar, L. A. Trends Biotechnol. 2002, 20, 9-12.

Figure 10. Characterization of the “cytosolic” space of porous beads. (A) Optical sectioning of a 10 µm bead (1 µm thick slices) by confocal microscopy. The distribution of rhodamine is homogeneous throughout the bead, within the resolution of the microscope (0.2 µm). (B) Photobleaching and recovery of trapped rhodamine dye inside porous beads.

on supported lipid bilayer membranes on glass surfaces is elucidated, this understanding has evolved into the basis of the technology of patterning membrane arrays29,30 and membrane protein microarrays.64 Controlled interactions between supported membranes and transmembrane proteins and the phase behavior of lipid bilayers have provided direct insight into the nature of biological interactions involved in cell signaling and adhesion.65-68 Much of the foundation for this work on beads is due to an early publication by Bayerl and Bloom,3 notwithstanding the pioneering work on planar surfaces by McConnell et al.69 and others.26,27,40 Thus, in this work we have taken advantage of the known behavior of lipid bilayers and (62) Piyasena, M.; Buranda, T.; Huang, J.; Wu, Y.; Sklar, L. A.; Lopez, G. P. Anal. Chem., in preparation. (63) Sklar, L. A.; Seamer, L. C.; Kuckuck, F.; Posner, R. G.; Prossnitz, E.; Edwards, B.; Nolan, J. P. Adv. Opt. Biophys. 1998, 3256, 144-153. (64) Fang, Y.; Frutos, A. G.; Lahiri, J. J. Am. Chem. Soc. 2002, 124, 2394-2395. (65) Brown, D. A.; London, E. Annu. Rev. Cell Dev. Biol. 1998, 14, 111-136. (66) Brown, D. A. Proc. Natl. Acad. Sci. U.S.A. 2001, 98, 1051710518. (67) Dietrich, C.; Bagatolli, L. A.; Volovyk, Z. N.; Thompson, N. L.; Levi, M.; Jacobson, K.; Gratton, E. Biophys. J. 2001, 80, 1417-1428. (68) Simons, K.; Ikonen, E. Nature 1997, 387, 569-572. (69) McConnell, H. M.; Watts, T. H.; Weis, R. M.; Brian, A. A. Biochim. Biophys. Acta 1986, 864, 95-106.

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present a facile method of biofunctionalizing beads in a way that is amenable to ready characterization of beadborne molecular assemblies. Scheme 1 illustrates the key features that favor the applicability of beads as solid supports for lipid bilayers. Uniform-sized beads are used to display egg PC mixed with functionalized lipids (e.g., biotin) with stoichiometry that can be regulated during the preparation of the SUVs. This is supported by the data shown in Figure 2 where the manifestation of FRET is used as an indicator of the close agreement between experimental and empirical regulation of the homogeneous surface density of biotinylated lipid components of a fluid bilayer membrane. FRET increases with acceptor density in a relationship that has been described elsewhere.44,45 Thus, for the data shown, when the acceptor site density is 0.01 acceptors/ R02,70 no FRET is observed (Figure 2D) and approximately 30% transfer efficiency is measured when the acceptor density is 0.1 acceptors/R02 in agreement with theoretical expectations.44,45 The data shown in Figure 2 also suggest that the labeled streptavidins themselves might be developed into a universal docking and FRET transduction platform for the various types of proximity-based assays in a manner similar to the FRET-based proximity sensors for multivalent protein toxins39,40 developed by Song et al. The use of mobile streptavidin linkers labeled with pairs of donors and acceptors might extend the applicability of this approach by easing the requirement that protein analytes must have multiple, specific binding sites for small molecule ligands. The ease with which lipid composition and stoichiometry can be used to control the receptor site coverage on beads is an important step toward rapid assay development. Such assays can be used either in a multiwell format suitable for screening or as components of microimmunoassays. In previous work, we have used flow cytometry and spectrofluorometry to characterize standard commercial 6.2 µm streptavidin-coated beads, prior to use in quantitative microimmunoassay related projects.6 During the course of that study, it became clear that because of the substantial pressure losses suffered during fluid transport, larger biofunctionalized beads were better suited for the application. Therefore, in a related study to be published elsewhere, we have used 20 µm streptavidin-bearing lipobeads prepared as described herein as platforms for molecular assembly in quantitative multianalyte microimmunoassay applications.62 ICAM-1 is an adhesion molecule expressed on vascular endothelial cells in response to physiological insults such as infection and inflammation. ICAM-1 is a multisubunit protein whose extracellular domain is shed into the circulation during the acute phase response, and its level typically changes several-fold over a period of hours to several days. The circulating levels can reach 1 µg/mL (≈70 nM).71,72 In Figure 4, we demonstrate the reconstitution of the protein in lipid bilayers where the surface coverage of the protein is regulated by its stoichiometry relative to the egg PC, albeit not in a simple linear fashion.73 However, because various proteins behave quite

differently in vesicle reconstitutions and may be sensitive to lipid compositions, generality cannot be inferred from the current findings. However, the sensitivity to the amount of R6.5 antibody used to display it can potentially be used as a basis of competitive bead-based assays to detect circulating levels of the protein in patient serum.74-77 Porous particles ranging in size from submicron to several hundred microns are widely used in chromatographic applications. In the past decade, there have been concerted efforts by several groups, notably Bayerl and co-workers,15,78 to make these particles more biofunctional by coating them with lipid bilayers. Some potential uses of these porous particles in biotechnological applications have been recently described.16,17 The accessibility of internal pore volume from the surface and the ability to incorporate fluorophores and functional biomolecules are features which are central to the use of these particles in chemical sensing and biophysical studies. To optimize their utility as platforms of such applications, the characteristics of their cytosolic space and its accessibility to the bulk surroundings must be well-defined. The combination of flow cytometry and microscopy provides a unique approach to such an analysis. Homogeneity in bead size and cytosolic accessibility are desirable characteristics in quantitative assays. Flow cytometry can be used to sort particles on the basis of scatter (Figure 7) or on the basis of the intensity analysis. Because flow cytometry can measure fluorescence intensity associated with beads on a single particleby-particle basis, it is possible to determine the average quantity of trapped dye molecules and thus to determine the volume of porous space in which the dye is trapped (Figure 8). Fluorescence microscopy of fluorophore-bearing beads can provide a detailed snapshot of the characteristics of a bead sample as illustrated in Figures 9 and 10. The details of Figure 10 suggest that the average bead in the sample is continuously accessible to dye. Furthermore, the diffusive mobility of the sequestered dye is consistent with the equimolar partitioning of dye molecules in the beads and the bulk solution (at equilibrium) as indicated by the linearity of the plot shown in Figure 8.

(70) The surface coverage of FRET acceptors is determined in terms of R02 from acceptors/R02 ) χAcR02/A where χAc is the mole fraction of the acceptor-labeled lipids, R0 ) 46 Å, and A is the area of the lipid headgroup (55 Å2). (71) Marlin, S. D.; Springer, T. A. Cell 1987, 51, 813-819. (72) Sudhoff, T.; Wehmeir, A.; Arning, M.; Bauser, U.; Schlomer, P.; Aul, A.; Schneider, W. Leukemia 1997, 11, 346-351. (73) In other experiments, we have varied the amount of ICAM-1 in 10-6 M vesicle dispersions; the changes in surface coverage of the protein were not linearly correlated to the amount of protein used.

(74) Tacyildiz, N.; Yavuz, G.; Gozdasoglu, S.; Unal, E.; Ertem, U.; Duru, F.; Ikinciogullari, A.; Babacan, E.; Ensari, A.; OkcuogluCavdar, A. Pediatr. Hematol. Oncol. 1999, 16, 149-158. (75) Bruserud, O.; Ulvestad, E. Leuk. Res. 1999, 23, 149-157. (76) Yaris, N.; Buyukpamukcu, M.; Kansu, E.; Canpinar, H. Med. Pediatr. Oncol. 2001, 36, 359-364. (77) Sudhoff, T.; Germing, U.; Aul, C. Int. J. Oncol. 2002, 20, 167172. (78) Grage, S.; Heinz, S.; Bayerl, T. M. Eur. Biophys. J. Biophys. Lett. 1998, 27, 425-428.

Conclusions In this study, we have demonstrated the utility of lipid bilayers supported on glass and mesoporous silica beads as versatile platforms for well-defined molecular assemblies. The supported lipid bilayers can incorporate ligands, fluorescent dyes, and transmembrane proteins. The concentration of these components in the supported lipid bilayer can be easily controlled through varying of their concentration in the vesicle preparation used to form the supported bilayers. The bilayers can be easily formed on microspherical substrates of a variety of sizes, depending on the end use application. The site density of fluorophore-labeled lipids, ligands, receptors, or transmembrane antigens can be quantitatively measured using flow cytometry. The conservation of fluidity by the lipid membrane allows for the incorporation of functional proteins53 into lipid membranes on solid or porous beads.


The tendency of some transmembrane proteins to lose functional activity upon reconstitution in supported lipid bilayers on conventional substrates such as glass has been reported, and considerable research effort is currently devoted to mitigating this setback.17,29,30,64,79,80 Mesoporous beads might provide a solution to this problem.16,17 Mesoporous beads also offer potential as versatile hosts for supported lipid bilayer membranes in which the porous structure of the beads forms an isolated cytosol-like compartment that can be used to store ions, dyes, drugs, and biological molecules and thus increase the functional versatility of microbeads in biotechnological applications. (79) Arora, A.; Tamm, L. K. Curr. Opin. Struct. Biol. 2001, 11, 540547. (80) Wagner, M. L.; Tamm, L. K. Biophys. J. 2000, 79, 1400-1414.

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Acknowledgment. This work was supported by the Air Force Office of Scientific Research (F49620-01-1-0168), the Department of Energy through the U.S./Mexico Materials Corridor Initiative, the National Science Foundation (EEC-0210835, MCB-9907611), NIH-BECON (GM60799/EB00264), and NM State Cigarette tax to the UNM Cancer Center. Some images in this paper were generated in the Cancer Center Fluorescence Microscopy Facility, which received support from NCRR 1 S10 RR14668, NSF MCB9982161, NCRR P20 RR11830, NCI R24 CA88339, the University of New Mexico Health Sciences Center, and the University of New Mexico Cancer Center. LA026405+

Biomimetic Molecular Assemblies on Glass and Mesoporous Silica

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