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Langmuir 2006, 22, 7078-7083
Biosensing Using Lipid Bilayers Suspended on Porous Silicon Oliver Worsfold,*,† Nicolas H. Voelcker,‡ and Takako Nishiya†,§ Frontier Research DiVision, Fujirebio Inc., 51 Komiya-cho, Hachioji-shi, Tokyo, Japan 192-0031 and School of Chemistry, Physics and Earth Sciences, Flinders UniVersity, Adelaide 5001, Australia ReceiVed January 12, 2006. In Final Form: May 14, 2006 We demonstrate for the first time the formation of a fluid lipid bilayer membrane on mesoporous silicon substrates for bioapplications. Using fluorescence recovery after photobleaching, the diffusion coefficients for the bilayers supported on oxidized, amino-, and biotin-functionalized mesoporous silicon were determined. The biodetection of a single human umbilical vein endothelial cell was accomplished using confocal microscopy and exploiting Foerster resonance energy transfer effects after the incorporation of RGD covalently linked lipid soluble dyes, with fluorescence donor and acceptor components, within the fluid membrane. A signal response of greater than 100% was achieved via the clustering of RGD peptides binding with areas of high integrin density on the surface of a single cell. These results are a testament to the usefulness of such functional molecular assemblies, based on mobile receptors, mimicking the cell membrane in the development of a new generation of biosensors.
Introduction Structure and function studies of individual biologics, within their natural environment, is essential for applications including drug discovery, protein analysis, biodetection, and single-cell studies. The self-assembly of lipid bilayer membranes on solid supports as biomimetics has been well documented1 with numerous novel strategies for supporting the membrane including mixed self-assembled monolayers,2 soft polymer cushions,3 and streptavidin biolinkers.4 Supported lipid bilayers membranes are a prime focus of research at present in a number of areas including biosensors5 and protein-protein characterization6 technologies because of their similarity to naturally occurring cellular and intracellular membranes. A recent application of mixed lipid bilayers highlights the potential for rapid nanopatterning of stripes and channels over macroscale lengths.7 The formation of lipid bilayers on various planar substrates including glass8 and micropatterned substrates9 has been extensively reported. Recent advances in this area include the formation of lipid bilayers directly onto the surface of micro-10 and nanosized11 silica beads. The primary disadvantage of these systems relates to the proximity of the lower lipid leaflet to the substrate surface, which limits the ability of these systems in the study of biologically important transmembrane proteins. * To whom correspondence should be addressed. E-mail:
[email protected]. Tel: +81-426-4774. Fax: +81-426-46-8325. † Fujirebio Inc. ‡ Flinders University. § Present address: Research Division, NanoCarrier Co. Ltd, 5-4-19 Kashiwanoha, Kashiwa, Chiba, Japan 277-0882. (1) Sackmann, E. Science 1996, 271, 43. (2) Jenkins, A. T. A.; Boden, N.; Bushby, R. J.; Evans, S. D.; Knowles, P. F.; Miles, R. E.; Ogier, S. D.; Schonherr, H.; Vancso, G. J. J. Am. Chem. Soc. 1999, 121, 5274. (3) Naumann, C. A.; Prucker, O.; Lehmann, T.; Ruhe, J.; Knoll, W.; Frank, C. W. Biomacromolecules 2002, 3, 27. (4) Proux-Delrouye, V.; Laval, J. M.; Bourdillon, C. J. Am. Chem. Soc. 2001, 123, 9176. (5) Schmidt, J. J. Mater. Chem. 2005, 15, 831. (6) Beddow, J. A.; Peterson, I. R.; Heptinstall, J.; Walton, D. J. Anal. Chem. 2004, 76, 2261. (7) Moraille, P.; Badia, A. Langmuir 2003, 19, 8041. (8) Tamm, L. K.; McConnell, H. M. Biophys. J. 1985, 47, 105. (9) Groves, J. T.; Boxer, S. G. Acc. Chem. Res. 2002, 35, 149. (10) Buranda, T.; Huang, J.; Ramaran, G. V.; Ista, L. K.; Larson, R. S.; Ward, T. L.; Sklar, L. A.; Lopez, G. P. Langmuir 2003, 19, 1654. (11) Baksh, M. M.; Jaros, M.; Groves, J. T. Nature 2004, 427, 139.
The potential of nanoporous-substrate-supported lipid bilayers as biosensors was indicated by Doshi et al.,12 who formed a lipid bilayer of simple composition on a fabricated nanoporous silica thin film and demonstrated a continuous bilayer via neutron reflectivity measurements. Lipid bilayers on nanoporous Al2O3 filters13 were also studied using NMR experiments and were shown to form continuous lipid bilayers where a thick water layer was present below the lower lipid leaflet. Our present design involves the formation of a porous substrate support using a simple electrochemical etching technique,14 followed by surface oxidation, silanization, and finally the formation of a lipid bilayer on the mesoporous surface.15 Oxidation and aminosilanization are common surface-modification procedures from silicon, silica, and indeed porous silicon, which is why the present study focuses on these two methods. The further biotin functionalization has been carried out in order to move the bilayer further away from the substrate by introducing a biotin-streptavidin layer. This enables the versatile production of supported lipid bilayers on substrates with arbitrary dimensions featuring arrays of nanoscale biomimetic compartments providing accessibility to the lower lipid leaflet. It is of prime importance for such biomimetic systems is to maintain the fluid nature of the resultant lipid bilayer within the device. Lateral mobility is demonstrated using fluorescence recovery after photobleaching (FRAP) techniques, which are well known as suitable vectors for membrane fluidity.16 To demonstrate the potential applications of the unique macroscaled mesoporous biomimetic device, the biosensing action of a lipid-soluble peptide incorporated within the lipid bilayer toward integrin-loaded human umbilical vein endothelial cells (HUVEC) was investigated using confocal microscopy and Foerster resonance energy transfer (FRET) as the interrogation technique. FRET is based on a nonradiative mechanism for energy transfer from a donor to an acceptor molecule and is regulated by an exponential power relationship (12) Doshi, D. D.; Dattelbaum, A. M.; Watkins, E. B.; Brinker, C. J.; Swanson, B. I.; Shreve, A. P.; Parikh, A. N.; Majewski, J. Langmuir 2005, 21, 2865. (13) Gaede, H. C.; Luckett, K. M.; Polozov, I. V.; Gawrisch, K. Langmuir 2004, 20, 7711. (14) Steinem, C.; Janshoff, A.; Lin, V. S. L.; Voelcker, N. H.; Ghadiri, M. R. Tetrahedron 2004, 60, 11259. (15) Romer, W.; Lam, Y. K.; Fischer, D.; Watts, A. D.; Fischer, W. B.; Goring, P.; Wehrspohn, R. B.; Gosele, U.; Steinem, C. J. Am. Chem. Soc. 2004, 126, 16267. (16) Wagner, M. L.; Tamm, L. K. Biophys. J. 2000, 79, 1400.
10.1021/la060121y CCC: $33.50 © 2006 American Chemical Society Published on Web 06/27/2006
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Figure 1. Schematic representation of the fluid lipid bilayer on porous silicon (a) before biosensing and (b) after cell biosensing has occurred.
of the donor-acceptor intermolecular distance and the fluorescence signal.17 This allows for signal amplification, which is important for increased response sensitivity. Human umbilical vein endothelial cells (HUVECs) contain significant quantities of integrin Rvβ3, a cell-surface-bound adhesion receptor important in cellular-cellular and cellularextracellular matrix interactions.18 The expression of integrin Rvβ3 in tumor cells is thought to be important in cell migration and is one of the targets for the inhibition of tumor growth.19 The biosensing selectivity of RGD (arginine-glycine-aspartate) containing peptides for integrin Rvβ3-loaded HUVECs has been previously demonstrated.20 However, it is worth noting that in our previous work the biosensing mechanism relied on the binding of only one RGD- linked dye molecule (either as a FRET donor or acceptor) whereas in the model presented in this study both the FRET donor and acceptor molecules contain RGD-linked dye molecules that we believe will increase the sensitivity of the biosensing response, thus enabling macroscale interrogation of a potential biosensor. The schematic in Figure 1a shows the structure of the mesoporous silicon substrate covered with a lipid bilayer suspended over the pores. Figure 1b illustrates that in the presence of target cells the RGD-tagged dye molecules aggregate at sites of high integrin intensity (corresponding to a single HUVEC), increasing the local density of the dye-labeled RGD ligands and significantly decreasing the distance between the donor and acceptor molecules and thus increasing the Foerster resonance energy transfer (FRET) from the donor to acceptor moieties. The aggregation of donor and acceptor dye molecules should therefore collectively augment both the donor and acceptor fluorescence emissions in the vicinity of the bound cell. Experimental Section Materials. All reagents purchased from commercial sources were used as received. Egg phosphatidylcholine (EPC), dioleoyl phosphatidylethanolamine (DOPE), N-3-(2-pyridyldithio)propionyl-1,2dipalmitoyl-sn-glycero-3-phosphoethanolamine (PDP-PE), and 1,2dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(cap biotinyl) sodium salt (BPE) were obtained from Avanti (Birmingham, AL). Cholesterol (chol), potassium hydroxide (KOH), caprylic acid, potassium chloride, sodium phosphate (mono- and dibasic forms), 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC), bipyridyl disulfide, n-octyl-D-glucopyranoside (OG), N-(2-hydroxyethyl)piperazine-N′-2-ethanesulfonic acid (HEPES), 2-morpholinoethanesulfonic acid (MES), and all silanes were obtained from Sigma Chemical (St. Louis, MO). Methanol and chloroform were obtained in their highest purity forms from Wako (Osaka, Japan). 4,4-Difluoro-5-(2-thienyl)-4-bora-3a,4a-diaza-s-indacene-3-dodecanoic acid (BODIPY 558/568) and 5-(((4-(4,4-difluoro-5-(2-thienyl)-4-bora3a,4a-diaza-s-indacene-3-yl)phenoxy)acetyl) amino)pentylamine hydrochloride (BODIPY TR cadaverine) were purchased from Molecular Probes (Eugene, OR). Sephadex G-25 and Sephadex G-75 were obtained from Pharmacia Biotech (Uppsala, Sweden). N(17) Selvin, P. R. Nat. Struct. Biol. 2000, 7, 730. (18) Hynes, R. O. Cell 1992, 69, 11. (19) Brooks, P. C.; Clark, R. F.; Cheresh, D. A. Science 1994, 264, 561. (20) Worsfold, O.; Toma, C.; Nishiya, T. Biosens. Bioelectron. 2004, 19, 1505.
Hydroxysulfosuccinimide (NHSS) was obtained from Pierce Chemical (Rockford, IL). The SPSDGRG and cyclic RGD(D-phe)C peptide sequences (HPLC purity > 90%) were synthesized by Qiagen (Tokyo, Japan). Purified water was obtained using a Millipore ELIX 10 water purification system. The phosphate-buffered saline (PBS) solution was composed of 10 mM Na2HPO4, 10 mM NaH2PO4, and 150 mM KCl with the pH adjusted to 7.4. Synthesis of RGD-Tagged Lipids. Prior to incorporation into small unilamellar vesicles (SUVs), the SPSDGRG peptide sequence was covalently attached to the BODIPY 558/568 lipid-soluble dye using the following method. NHSS (3.5 mg) and EDC (7.7 mg) were added to BODIPY 558/568 solubilized with 200 µL of 2% (wt/vol) OG in 50 mM MES buffer at pH 5.5, and the mixture was incubated for 10 min at room temperature. BODIPY 558/568 C12 with an NHSS-activated carboxylic derivative was purified using a Sephadex G-25 column with 50 mM HEPES/0.1%(wt/vol) OG at pH 8.0 and was added to 5 mg of RGD-containing peptide SPSDGRG. The resultant solution was incubated for 12 h at 4 °C with gentle stirring and purified by dialysis for 12 h at 4 °C against PBS. MS data: calcd for C50H71BF2N12O13S, 1129.04; found, 1129.69. Cyclic RGD(D-phe)C (0.5 mg) solubilized with 0.2 mL of PBS was added to 0.5 mg of BODIPY TR cadaverine IA and solubilized with 0.2 mL of DMSO. The mixture was incubated for 2 h at room temperature, and the nonconjugated materials were removed by dialysis against PBS for 12 h at 4 °C. MS data: calcd, 1158.17; found, 1159.42. Synthesis of (2-Pyridyldithiopropionamidopropyl)dimethylmethoxysilane. Bipyridyl disulfide (3.75 g, 17 mmol) was dissolved in 20 mL of 99.5% ethanol and 0.5 mL of glacial acetic acid. Upon vigorous stirring, 0.9 g of propionic acid in 5 mL of ethanol was added slowly. The reaction mixture was stirred for 12 h at room temperature. After the removal of the solvent, the product was purified via neutral alumina column chromatography and dried under high vacuum. 2-Pyridyl-2-carboxyethyl disulfide (0.77 g) was dissolved in methylene chloride, and 668 mg (3.5 mmol) of EDC was added. The mixture was stirred under argon for 30 min before 0.6 mL (3.5 mmol) of (3-aminopropyl)dimethylmethoxysilane was injected via syringe. After being stirred overnight, the organic phase was washed several times with a saturated NaHCO3 aqueous solution, dried over Na2CO3, and evaporated. The crude product was purified by silica chromatography. Yield: 0.25 g (70%). 1H NMR (200 MHz, CDCl3): δ 8.41 (m, 1H), 7.62 (m, 2H), 7.08 (m, 1H), 3.52 (s, 9H), 3.23 (m, 2H), 3.03 (t, 2H), 2.55 (t, 2H), 1.60 (m, 2H), 0.63 (m, 2H). Porous Silicon Preparation. Silicon wafers (p++ type, resistivity 0.6-1.5 mΩ cm) from Virginia Semiconductors (Fredericksburg, VA) were cleaned in hot piranha solution for 30 min and rinsed with copious amounts of DI water. Wafers were then treated for 1 min with 40% HF and rinsed with DI water and 2-propanol. Anodization was carried out in the dark in a custom-made etching cell at 180 mA/cm2 to a charge of 4.5 C/cm2. The samples were subsequently rinsed thoroughly with methanol, acetone, and methylene chloride and finally dried in a stream of nitrogen. Porous Silicon Functionalization. A. Oxidation. The predominantly hydride-terminated porous layer was exposed to ozone from a Fischer ozone generator with a flow rate of 7.7 g/h for 10 min. The oxidized samples were vigorously rinsed with methanol, acetone, and dichloromethane and then dried in a stream of nitrogen. B. Amino Functionalization. The oxidized porous silicon samples were functionalized by immersion in a 100 mM solution of (3aminopropyl)trimethoxysilane in dry toluene for 5 min. Afterward,
7080 Langmuir, Vol. 22, No. 16, 2006 the chips were extensively rinsed in toluene, methanol, acetone, and dichloromethane and dried in a stream of nitrogen. C. Biotin Functionalization. The oxidized mesoporous silicon samples were immersed in a 100 mM solution of (2-pyridyldithiopropionamidopropyl)dimethylmonomethoxysilane in toluene followed by refluxing for 3 h. Afterward, the chips were extensively rinsed in toluene, methanol, acetone, and dichloromethane followed by drying in a stream of nitrogen. Reduction of the disulfide in a 10 mM dithiothreitol solution released 2-thiopyridone, which could be quantified by its UV absorption at 343 nm ( ) 8080 M-1 cm-1). Mesoporous silicon functionalized with (2-pyridyldithiopropionamidopropyl)dimethylmethoxysilane was exposed to a 10 mM DTT solution for 1 h and washed extensively with DI water. Subsequently, the reduced surface was submerged in a 400 µM solution of biotinHPDP (Pierce) for 30 min. The biotinylated surfaces were washed extensively with DI water. Surface Analysis. FTIR spectra were acquired with a Thermo Nicolet Avatar 370 in diffuse reflectance mode (diffuse reflectance smart accessory). Diffuse reflectance spectra are reported in Kubelka-Munk units. Atomic force microscopy (AFM) images were obtained under ambient conditions using a Nanoscope IV Multimode scanning probe microscope (Veeco Corp., Santa Barbara, CA) operating in tapping mode. Mikromasch (Madrid, Spain) hi-res tips (325 kHz) were used as purchased. A Philips (Amsterdam, Netherlands) XL20 scanning electron microscope was used for the electron microscopy imaging. Images were acquired without evaporation of a gold layer. Preparation of Small Unilamellar Vesicles (SUVs). From stock solutions in chloroform, the following mol % ratios of each component were added to a glass vial. EPC, 47 mol %; DOPE, 12 mol %; chol, 29 mol %; BPE, 11 mol %; and dye lipid, 1 mol %. The solvent was removed under N2 and further dried using a vacuum desiccator for 2 h. The dried lipids were rehydrated in 3 mL of PBS solution vortexing as necessary to aid dissolution. The occluded solution was tip sonicated at 4 °C at 15% power until the solution became clear (40 min). The solution was ultracentrifuged at 4 °C for 2 h at 60 000 rpm to remove the titanium particles and large/multilamellar vesicles using a Beckmann-Coulter ultracentrifuge. Various dye lipid donor/ acceptor ratio combinations were produced as SUVs and examined spectroscopically with the optimum biosensing results produced using a donor/acceptor ratio of 1:1. Particle size analysis of the SUVs was performed using a Beckmann-Coulter N4 PLUS submicron particle analyzer, and SUVs with a diameter of 45 nm (SD ) 5.5 nm) were routinely produced using this method. Lipid Bilayer Formation. The mesoporous silicon samples were washed extensively in DI water and assembled in the in-housedesigned sample holder. The biotin-functionalized mesoporous silicon substrates were incubated with 1 mL of 50 µg mL-1 streptavidin in PBS solution for 60 min and washed extensively in PBS prior to vesicle incubation. The amine- or biotin/streptavidin-terminated samples were incubated with 1 mL of PBS buffer solution for 120 min prior to the addition of vesicle solutions. The vesicle solution (0.028 mg mL-1) was added to the sample holder and incubated with the substrates for 15 min. The sample was extensively washed in PBS buffer to remove excess vesicles prior to FRAP or biosensing experiments. For reference, silicon wafers [Si(100), Kyodo Int., Japan] were diced into 1 cm2 squares and cleaned using an ozone cleaner (Bionanoforce Inc.) for 30 min. The same lipid composition as described in the above preparation section was then incubated with the cleaned silicon samples for 15 min. FRAP Experiments. Using an Olympus confocal laser microscope (LSV-1000), excitation was performed using a He/Ne laser at 488 nm at 44% intensity. Emission with ×10 magnification, using a scan rate of 12.5 µs pixel-1, was collected with 550-640 nm filters and a 580 V PMT, 3× gain, and 3% offset. During photobleaching, the 488 nm laser was set at 100% intensity for 15 s. Integrin-Loaded HUVEC Biosensing. Biosensing experiments were performed by removing aliquots of the PBS solution for biosensor equilibration and adding equal volumes of the HUVEC suspension to the sample holder at the specified concentration. Using an Olympus confocal laser microscope (LSV-1000), excitation was
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Figure 2. Diffuse reflectance IR spectra of porous silicon. (A) Freshly etched porous silicon, (B) ozone-oxidized porous silicon, and (C) aminosilane-functionalized porous silicon. The bands under the asterisks correspond to IR reflective interference effects with evenly sized broad peaks.28 performed using a He/Ne laser at 488 nm at 64% intensity. Emission with ×10 magnification, using a scan rate of 12.5 µs pixel-1, was collected in channel 1 filtered between 562 and 588 nm, a 680 V PMT, 1× gain, and 0% offset in channel 2 with a barrier filter >610 nm and a 640 V PMT, 2× gain, and 0% offset. All fluorescence experiments were performed at 298 K.
Results and Discussion Porous silicon was prepared by anodization in HF/ethanol solution under conditions that lead to the formation of mesoporous21 silicon. We used mesoporous silicon here because macroporous silicon, which could also be prepared from the same wafer, did not allow the preparation of stable suspended lipid bilayers whereas pore diameters in microporous silicon were deemed insufficient to allow the lateral diffusion of transmembrane components across the suspended regions. Mesoporous silicon substrates were characterized by FTIR in diffuse reflectance mode. Freshly etched porous silicon displayed diffuse reflectance spectra consistent with a hydride-terminated surface. Upon ozone oxidation, the bands corresponding to bending and stretching modes of Si-Hx (∼2100 cm-1) were removed and replaced a peak at 1080 cm-1 and a shoulder at 1160 cm-1 attributed to asymmetrical Si-O-Si and Si-OH stretching modes, respectively (Figure 2). Silanization with (3aminopropyl)trimethoxysilane led to the appearance of two peaks at 2940 and 2890 cm-1 corresponding to aliphatic V(C-H) stretches. The pore size of the mesoporous silicon was estimated from the microscopy images in Figure 3. AFM imaging (Figure 3a) demonstrates that the pores were, on average, 20 nm in diameter and hence within the mesoporous regime, and the crosssectional SEM image (Figure 3b) indicates a pore depth of approximately 3 µm. Suspended lipid bilayers were prepared by the vesicle adsorption and spreading technique using SUVs made from phosphatidylcholines and phosphatidylethanolamines. The SUVs (21) The terminology used in this report to describe the type of porous silicon produced (i.e., mesoporous) is based on the IUPAC nonmetric system and is often referred to as nanoporous silicon in other studies.
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Figure 3. Determination of the pore size for the porous silicon film. (a) Three-dimensional AFM image showing mesopores with a 20 nm mean diameter. (b) Cross-sectional SEM image showing a true depth profile.
Figure 4. FRAP kinetics. (a) Fluorescence micrographs taken directly after photobleaching and 2000 s after photobleaching of the lipid bilayer supported on amino-functionalized nanoporous silicon. (b) Fractional fluorescence recovery curve for dye-tagged lipids in a lipid bilayer on amino-functionalized nanoporous silicon.
also contained RGD-tagged lipid-soluble BODIPY dyes. Therefore, vesicle spreading on the substrate surfaces resulted in the formation of fluorescent suspended lipid bilayers. The membrane’s fluidity is essential for biosensing clustering of bilayer-bound ligands. We therefore determined lipid bilayer fluidity by FRAP techniques. This involves photobleaching a small (known) section of the fluorescent lipid bilayer and measuring the recovery of the fluorescent molecules into the photobleached section in time lapse. This measurement enables the diffusion constant (D) for the system to be calculated via the following equation22
D)
( ) r2 4t50
where D is the diffusion coefficient, r is the radius of the photobleached area, and t50 is the time required to achieve 50% fluorescence intensity recovery. The fluorescence microscope images shown in Figure 4a demonstrate the fluidity of the lipid bilayer suspended on amino-functionalized mesoporous silicon by recovery of the fluorescence measured immediately after photobleaching and 2000 s after photobleaching. The t50 time is calculated from the fractional fluorescence recovery curve shown in Figure 4b, which gives a diffusion coefficient for this system of D ) 1.4 × 10-8 cm2 s-1. The diffusion constants for a lipid bilayer on oxidized porous silicon substrate and for a biotinstreptavidin-supported lipid bilayer on a mesoporous silicon substrate were also obtained (Table 1). The low diffusion constant (22) Gyorvary, E.; Wetzer, B.; Setyr, U.; Sinner, A.; Offenhausser, A.; Knoll, W. Langmuir 1999, 15, 1337. and refs therein.
Table 1. Lipid Bilayer Diffusion Constants (D) for Various Surface-Terminated Porous Silicon (PSi) Substrates and Reference Samples substrate
(D)/ × 10-8 cm2 s-1
oxidized surface PSi amine-terminated PSi biotin-streptavidin on PSi ozone-cleaned silicon flat glass1
2.4 1.4 0.5 2.8 3.0
(D ) 0.5 × 10-8 cm2 s-1) of the biotin-streptavidin-supported lipid bilayer is probably due to steric effects of the lipid attached to a protein support. These steric effects are caused by the “pinning” of the lipid bilayer to the streptavidin via the biotinylated lipid incorporated into the lipid bilayer. The diffusion constant (D ) 2.3 × 10-8 cm2 s-1) of the lipid bilayer on oxidized mesoporous silicon is greater than that obtained for aminoterminated mesoporous silicon, but the oxidized surface was found to be unstable in an aqueous medium and pore corrosion was evident from the generation of hydrogen gas bubbles. The limited stability of the porous layer severely affects the stability of the suspended lipid bilayers on such substrate surfaces. Previous interferometric reflectance spectroscopy studies have shown that the stability toward corrosion in an aqueous environment increases in order of freshly etched, oxidized, and silane-linker-modified porous silicon.12,23 The diffusion coefficient values on mesoporous silicon are only slightly lower than the value recorded for simple lipid bilayers on glass (3 × 10-8 cm2 s-1)1 and are less than the value for the (23) Janshoff, A.; Dancil, K.-P. S.; Steinem, C.; Greiner, D. P.; Lin, S.-Y.; Gurtner, C.; Motesharei, K.; Sailor, M. J.; Ghadiri, M. R. J. Am. Chem. Soc. 1998, 120, 12108.
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same lipid composition on ozone-cleaned flat silicon substrates (2.8 × 10-8 cm2 s-1, Table 1). This decrease in mobility is probably due to the surface roughness of the porous substrate disrupting the continuity of the lipid bilayer. Alternatively, the use of multicomponent lipid bilayers could influence the lateral mobility. In either case, the small loss in membrane fluidity of the mesoporous substrate compared with that of flat surfaces is more than compensated for by the presence of nanocompartments enabling the incorporation of proteins such as the important ligandgated ion channel acetylcholine receptor,24 which is involved in synaptic neurotransmission. The study of the acetylcholine receptors in planar lipid bilayers25 is hampered by steric effects caused by the substrate interacting with the protein’s extramembrane components, disrupting the 3D structure of the protein. However, using the porous matrix described in the present study will enable the study of these types of transmembrane proteins in a biomimetic environment by providing accessible areas below the lipid bilayer for the extramembrane components of transmembrane proteins. This is not achievable with a flat surface as a membrane support. The PC + PE mixed vesicles used here have a low, negative electrokinetic potential at low ionic concentrations and pH 7.4.26 It is well known that electrostatic interactions between liposomes and surfaces are important factors in the formation of supported lipid bilayers. The PC + PE mixed vesicles should, from an electrostatic point of view, favor fusion on positively charged surfaces such as protonated amino groups of the functionalized mesoporous silicon. However, vesicle fusion on the inner pore walls was to be avoided at the same time, and this was achieved through size exclusion effects. Light scattering experiments showed the extruded SUVs to be about 45 nm in diameter (SD, 5.5 nm). Given the mean diameter of pores on porous silicon of 20 nm (Figure 3), it is anticipated that because of size exclusion vesicle fusion is limited to occur on top of the pore holes instead of within the pores, therefore generating a suspended lipid bilayer. In addition, fluorescence recovery experiments demonstrated the continuity of the lipid bilayer spanning the grid-shaped top surface rather than entering into the individual pore holes. If the lipid bilayers were formed only within the pores, then photobleaching of a large area as described in our experiments would not lead to fluorescence recovery whereas the recovery data indicates that the fluorescent bilayer is continuous over large areas and not within the pores. After determining that the membrane fluidity of the suspended lipid bilayers was within acceptable values, we attempted to detect interactions of the RGD-dye-labeled lipid bilayer on mesoporous silicon with integrin expressing cells. We used HUVECs because these cells express large amounts of RGD binding integrin units (1 cell contains approximately 6 × 105 integrins). Figure 5 shows the effect of exposing the suspended lipid bilayers to 1500 cells mL-1 integrin-loaded HUVECs determined by means of confocal laser scanning microscopy. The image in the left panel shows a fluorescence microscopy image of the lipid bilayer on mesoporous silicon prior to the addition of HUVECs. The image in the right panel of Figure 5 demonstrates the increase in fluorescence of specific areas on the lipid bilayer where single cells have attached. In this image (Figure 5, right panel), eight areas have been labeled that vary in size from 30 µm consistent with the expected size range for spreading HUVECs. A time scan of fluorescence intensity shown in Figure (24) Leite, J. F.; Cascio, M. Mol. Cell. Neurosci. 2001, 17, 777. (25) Peterson, I. R.; Beddow, J. A. System Aspects of Supported Membrane Biosensors. In Planar Lipid Bilayers (BLMs) and Their Applications; Tien, H. T., Ottova, A., Eds.; Elsevier: New York, 2003; p 735. (26) Egawa, H.; Furusawa, K. Langmuir 1999, 15, 1660.
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Figure 5. Confocal microscopy image of a dye-tagged RGD-lipid bilayer on a mesoporous silicon matrix. Left-hand and right-hand images were recorded prior to and after the addition of 1500 cells mL-1 of HUVEC, respectively. Excitation is at 488 nm.
Figure 6. Time scan of HUVECs for the fluorescence of the dyetagged RGD-lipid bilayer on an amine-terminated porous silicon matrix. Channel 1 is filtered from 562 to 588 nm, and channel 2 uses a barrier filter >610 nm. Excitation is at 488 nm.
6 that is measured at the point labeled “1” in the right panel of Figure 5 shows a rapid increase in both donor and acceptor fluorescence during HUVEC addition. HUVEC cells were only added after 4000 s of lead-in time to demonstrate the stability of the donor and acceptor fluorescence within the lipid bilayer. There is a large increase in both the donor and acceptor fluorescence intensities because the proposed mechanism utilizes RGD-containing ligands on both fluorescent moieties. Biosensing of integrin-loaded HUVEC cells is signal mediated via both ligands causing simultaneous aggregation of the donor and acceptor dye moieties with subsequent increases in both the concentration and fluorescence output of the excited donor dye and also the acceptor dye via a FRET mechanism. The data shows a >100% increase in the donor fluorescence intensity (channel 1) and a >60% increase in the acceptor fluorescent intensity (channel 2) when biosensing a single HUVEC. This study demonstrates the possibility of using a simple macroscale interrogation technique (confocal microscopy) to investigate nanoscale biofunctions such as the co-localization of ligands or receptors. Previous work has shown that the peptide system described in this study is selective for integrin-loaded HUVECs.27 In addition, this previous study using fluorescence spectroscopy was only able to detect a HUVEC concentration of 10 cells mL-1 whereas the present fluorescence microscopy demonstrates single-cell detection. This novel device matrix with bilayer-capped nanoscale compartments will enable the use of high-throughput techniques in studying nanoscale events such as the clustering of integrin receptors and the characterization of other biologically important transmembrane proteins and biochemical reactions. There is (27) Worsfold, O.; Nishiya, T. J. Biomed. Nanotechnol. 2005, 1, 102. (28) Gao, T.; Gao, J.; Sailor, M. J. Langmuir 2002, 18, 9953.
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further scope to use these nanocompartments to investigate biochemical reactions in femtoliter environments or to pursue electrochemical processes. By incorporating RGD-tagged lipidsoluble dye molecules within the lipid bilayer matrix, we have demonstrated the applicability of using confocal microscopy for single-cell biosensing of integrin-expressing HUVECs. These systems are envisaged for applications in the study of transmembrane proteins, such as the acetylcholine receptor, hormone receptor, and other biologically important G-protein coupled receptors and their interactions with other proteins or analytes, which are not possible with supported lipid bilayers on flat substrates.
Conclusions Here we show the formation of a fluid lipid bilayer membrane on a mesoporous (20 nm pore diameter) silicon substrate with arbitrary macroscale lateral dimensions and the subsequent use of this system for the biodetection of integrin-expressing cells. The fluidity of the lipid bilayer suspended on numerous surfaces including amino-terminated mesoporous silicon was demonstrated. Single-cell biodetection, with over a 100% change in the fluorescence signal due to receptor clustering, was observed via FRET confocal microscopy. LA060121Y
