Langmuir 2008, 24, 1629-1634
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Silica Colloidal Crystals as Three-Dimensional Scaffolds for Supported Lipid Films Eric E. Ross*,† and Mary J. Wirth Department of Chemistry, Gonzaga UniVersity, Spokane, Washington 99258, and Department of Chemistry, UniVersity of Arizona, Tucson, Arizona, 85721 ReceiVed October 28, 2007. In Final Form: December 18, 2007 This report describes the assembly of laterally diffusive lipid layers within the pores of colloidal crystals for potential application in membrane-based sensing. The amount of lipid encapsulated within colloidal crystals depends upon the method used to introduce the lipid to the crystalline substrate. Relative to a planar supported lipid bilayer, lipid loading in a 6.6 µm thick crystal was 15-73 times greater, as observed by fluorescence microscopy. Protein adsorption studies indicate that the crystal pores are open and that the silica surface of the crystal is passivated with respect to adsorption of a model protein when coated with POPC. Furthermore, the mesoporous environment of the colloidal crystal is found to protect lipid films from drying and rehydration processes that destroy planar supported lipid bilayers. The potential of colloidal crystal encapsulated lipid films for chemical sensing is demonstrated by a model protein binding assay.
Introduction The assembly of phospholipid bilayers on porous silica supports has been recently reported by several research groups.1 This format of supported bilayer should reduce the interaction of membrane-spanning proteins with the inorganic support and provide experimental access to aqueous compartments on both sides of the bilayer to address intra- and extracellular domains of incorporated membrane proteins.2 Parikh and co-workers recently discovered that phospholipid vesicles rupture at the surface of a colloidal crystal (CC, also known as a synthetic opal) composed of 330 nm silica colloids to form continuous and fluid two-dimensional lipid bilayers that span the crystal pores.1a While CCs have been extensively examined for purposes related to their optical properties,3 this is one example in a host of emerging applications based on their uniform and adjustable mesoscale porosity. Other examples include perm-selective membranes or sieving matrices for proteins.4,5 By virtue of their large and solute-accessible surface area, CCs have demonstrated potential in applications involving adsorption or molecular binding processes, such as highly efficient electrochromotagraphic separation of peptides,6a and sensitive protein arrays.6b Asher and co-workers have pioneered the use of polymerized crystalline colloidal arrays for sensing applications using a variety of receptor chemistries, including biomolecules.7 * To whom correspondence should be addressed. E-mail: rosse@ gonzaga.edu. † Gonzaga Univeristy. (1) (a) Brozell, A. M.; Muha, M. A.; Sanii, B.; Parikh, A. N. J. Am. Chem. Soc. 2006, 128, 62-63. (b) Davis, R. W.; Flores, A.; Barrick, T. A.; Cox, J. M.; Brozik, S. M.; Lopez, G. P.; Brozik, J. A. Langmuir 2007, 23, 3864-3872. (c) Weng, K. C.; Stålgren, J. J. R.; Duval, D. J.; Risbud, S. H.; Frank, C. W. Langmuir 2004, 20, 7232-7239. (2) For a recent review on lipid biosensors, see; Castellana, E. T.; Cremer, P. S. Surf. Sci. Rep. 2006, 61, 429-444. (3) Two reviews, (a) Joannopoulos, J. D.; Villeneuve, P. R.; Fan, S. Nature 1997, 386, 143. (b) Xia, Y.; Gates, B.; Yin, Y.; Lu, Y. AdV. Mater. 2000, 12, 693. (4) (a) Newton, M. R.; Bohaty, A. K.; White, H. S.; Zharov, I. J. Am. Chem. Soc. 2005, 127, 7268. (b) Cichelli, J.; Zharove, I. J. Am. Chem. Soc. 2006, 128, 8130-8131. (c) Cichelli, J.; Zharov, I. J. Mater. Chem. 2007, 17, 1870-1875. (5) Zeng, Y.; Harrison, D. J. Anal. Chem. 2007, 79, 2289-2295. (6) (a) Zheng, S.; Zhang, H.; Ross, E.; Van Le, T.; Wirth, M. J. Anal. Chem. 2007, 79, 3867-3872. (b) Zheng, S. P.; Ross, E.; Legg, M. A.; Wirth, M. J. J. Am. Chem. Soc. 2006, 128, 9016-9017. (7) Walker, J. P.; Asher, S. A. Anal. Chem. 2005, 77, 1596-1600.
Supported lipid bilayers are useful as biomimetic models for the study of membrane structure and protein function.8 The study of dynamic membrane processes such as multivalent binding events is enabled by the lateral mobility of lipids in supported bilayers.9 This feature can also be exploited for the detection of binding events; for example, Groves and co-workers have utilized changes in the diffusion rate of a membrane probe to measure ligand-receptor binding.10 Because phospholipid bilayers are generally resistant to nonspecific protein adsorption and supportive of tethered or imbedded protein activity,11 they are appealing supports for biosensors.2 Furthermore, advances in the formation of microscopic domains of fluid lipid bilayers12,13 and lipid modification of microfluidic channels14 provide a promising outlook for the use of lipid films in sensing and diagnostic applications. Two commonly cited drawbacks of supported lipid films are the inherent fragility of the molecular bilayer and the minimal surface area afforded by a smooth, flat interface. The later often affects sensor performance through the generation of relatively weak analytical signals from ligand binding events at planar interfaces; sensitive instrumentation is required to accurately measure binding events to ∼10-12 mol/cm2 immobilized receptor. A single bilayer (∼5 nm thick) also underutilizes the energy in evanescent field based techniques such as surface plasmon resonance or total internal reflectance fluorescence, thus not fully realizing the potential sensitivity of such formats. Furthermore, limited dynamic range in planar assays results from ligand saturation of immobilized receptors.15 With respect to bilayer fragility, progress is being made in rendering supported bilayers (8) (a) McConnell, H. M.; Watts, T. H.; Weis, R. M.; Brian, A. A. Biochim. Biophys. Acta 1986, 864, 95-106. (b) Sackmann, E. Science 1996, 271, 43-48. (9) Shi, J.; Yang, T.; Kataoka, S.; Zhang, Y.; Diaz, A. J.; Cremer, P. S. J. Am. Chem. Soc. 2007, 129, 5954-5961. (10) Yamazaki, V.; Sirenko, O.; Schafer, R. J.; Groves, J. T. J. Am. Chem. Soc. 2005, 127, 2826-2827. (11) Glasma¨star, K.; Larsson, C.; Ho¨o¨k, F.; Kasemo, B. J. J. Colloid Interface Sci. 2002, 246, 40-47. (12) Groves, J. T.; Boxer, S. G. Acc. Chem. Res. 2002, 35, 149-157. (13) Ross, E. E.; Mansfield, E.; Huang, Y.; Aspinwall, C. A. J. Am. Chem. Soc. 2005, 127, 16756-16757. (14) Yang, T. L.; Jung, S. Y.; Mao, H. B.; Cremer, P. S. Anal. Chem. 2001, 73, 165. (15) See for example, (a) Loomans, E.; Gribnau, T.; Bloemers, H.; Schielen, W. J. Immunol. Methods 1998, 221, 119. (b). Bonroy, K.; Friedt, J.-M.; Frederix, F.; Laureyn, W.; Langerock, S.; Campitelli, A.; Sara, M.; Borghs, G.; Goddereris, B.; Declerck, P. Anal. Chem. 2004, 76, 4299.
10.1021/la7033609 CCC: $40.75 © 2008 American Chemical Society Published on Web 01/17/2008
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more robust by utilizing tethered polymer16 and protective protein layers;17 however, some applications may suffer from a constrained lipid composition or possible impediment of solute diffusion to the membrane. Another approach for stabilizing membrane components of sensors is encapsulation within a protective framework such as a hydrogel or sol-gel matrix. Hydrogel encapsulation was found to impart enhanced durability to lipid membranes and allow single molecule studies of membrane proteins.18 For many analytical applications, background fluorescence from organic gels, or swelling in dynamic buffer conditions is a drawback.19 Sol-gel encapsulation of liposomes has been reported,20 and ion sensing using channel peptides21 and environmental fluorophores22 have been demonstrated with these materials. However, the sol-gel process does not generally produce materials with a uniform distribution of mesoscale pores, and equilibration time for binding or sensing studies are frequently several hours due to relatively slow diffusion through sol-gel monoliths.23 CCs have been shown to have very high solute diffusion rates,24 suggesting minimal impediment of ligand diffusion to immobilized receptors. In this work, we describe successful efforts to modify the internal surface area of silica CCs with lipid films. This builds on the work of Parikh and co-workers1 who showed that a single bilayer was not formed upon crystals composed of larger (5.6 µm) colloids and consequently, larger pores. Instead, the vesicles permeated the crystal and colloids were individually modified by lipid layers; no long-range lipid diffusion was observed. These observations support an argument relating vesicle permeation of the CC to the relative size of vesicles and the crystal pores, which are related geometrically to colloid size.25 We show that the internal surfaces of CC films with nanoscale particles can be uniformly coated with lipid and that lateral diffusion is significant. The increase in the bilayer surface area accessible to a diffusing protein for a 6.6 µm thick CC was found to be nearly 40 times greater than a planar supported bilayer. Experimental Section Silica particles synthesized by the Sto¨ber method were deposited on quartz microscope slides by a meniscus evaporation procedure from ethanol.26,27 Prior to deposition, particles were calcined three times at 600 °C, and the deposited crystal was sintered at 900 °C overnight.28 This procedure increases crystal robustness but essentially eliminates surface hydroxyl groups, which were subsequently regenerated by 48 h immersion in a dilute tetrabutyl ammonium hydroxide solution (pH 9) at 50 °C. CCs were rinsed extensively and stored in deionized water. Prior to use, CCs were dried using an air stream and were cleaned in a UV-ozone cleaner (16) Albertorio, F.; Diaz, A. J.; Yang, T.; Chapa, V. A.; Kataoka, S.; Castellana, E. T.; Cremer, P. S. Langmuir 2005, 21, 7476-7482. (17) Holden, M. A.; Jung, S.-Y.; Yang, T.; Castellana, E. T.; Cremer, P. S. J. Am. Chem. Soc. 2004, 126, 6512-6513. (18) Jeon, T.-J.; Malmstadt, M.; Schmidt, J. J. J. Am. Chem. Soc. 2006, 128, 42-43. (19) Honore, P.; Granjeaud, S.; Tagett, R.; Deraco, S.; Beaudoing, E.; Rougemont, J.; Debono, S.; Hingamp, P. BMC Genomics 2006, 7. (20) Brennan, J. D. Acc. Chem. Res. 2007, 40, 827-835. (21) Besanger, T. R.; Brennan, J. D. Anal. Chem. 2003, 75, 1094-1101. (22) Nguyen, T.; McNamara, K. P.; Rosenzweig, Z. Anal. Chim. Acta 1999, 400, 45-54. (23) Besanger, T. R.; Easwaramoorthy, B.; Brennan, J. D. Anal. Chem. 2004, 76, 6470. (24) (a) Newton, R. M.; Morey, K. A.; Zhang, Y.; Snow, R. J.; Diwekar, M.; Shi, J.; White, H. S. Nano Lett. 2004, 4, 875. (b) Bohaty, A. K.; Zharov, I. Langmuir 2006, 22, 5533-5536. (25) The pore diameter is related to colloid diameter geometrically in an fcc packed CC and is ∼0.15d, where d is the diameter of the colloids used in the CC. (26) Stober, W.; Fink, A.; Bohn, E. J. Colloid Interface Sci. 1968, 26, 62-7. (27) Jiang, P.; Bertone, J. F.; Hwang, K. S.; Colvin, V. L. Chem. Mater. 1999, 11, 2132-2140. (28) Van Le, T.; Ross, E. E.; Velarde, T. R. C.; Legg, M. A.; Wirth, M. J. Langmuir 2007, 23, 8554-8559.
Letters for 15 min. Colloid diameter was determined by scanning electron microscopy. CC thickness was measured by optical microscopy (see Supporting Information). Lipid modification of CCs was performed three ways. These include incubation of CCs with a buffered solution of lipid vesicles, immersion of the CC in a buffered surfactant-solubilized lipid solution, and evaporation of a chloroform lipid solution in the presence of a CC. The lipid 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC, Avanti Polar Lipids, Inc.) was used throughout the study. Lipid vesicles were formed by extrusion through polycarbonate filters (0.1 µm pores) of a 1 mg/mL suspension of POPC in 10 mM HEPES (pH 7.4) buffer that had been subjected to three freeze-thaw cycles. Lipid surfactant mixtures were prepared by addition of 1 mM n-dodecyl-β-D-maltoside (cmc 0.17 mM) in HEPES buffer to dried films of POPC in small vials and vortexing the suspension for at least 2 h until the solution was clear. Organic solutions of lipid were prepared by dilution of stock lipid solutions in chloroform. The CC was placed vertically in a lipid chloroform solution (0.5 mg/mL) so that ∼2 cm2 of the crystal was submerged, and the solvent was allowed to evaporate completely. Residual chloroform was removed with a nitrogen stream and storage in vacuum (50 mTorr) for 6 h. Lipid dopants (1% relative to POPC in these studies) included fluorescent Texas Red 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine triethylammonium salt (TR-DHPE, Invitrogen) or the biotinylated lipid 1,2-dioleoyl-sn-glycero-3-phosphoethanolamineN-(Biotinyl)(Sodium Salt) (Biotinyl PE, Avanti Polar Lipids, Inc.). Planar supported lipid bilayers were formed by vesicle fusion to quartz microscope slides. Fluorescein-labeled avidin (Avidin-FITC, Invitrogen) in HEPES buffer (10 mM, pH 7.4) was used at a concentration of 0.1 mg/mL in adsorption and binding studies. CC or lipid modified CCs were incubated with protein solutions for 30 min prior to exchange of the bulk solution to standard buffer for 10 min and subsequent fluorescence measurements. A Nikon Eclipse TE 2000-U microscope, equipped with a 100 W mercury arc lamp (HBO) and a Cascade 512b (Roper Scientific) CCD camera, was used to collect fluorescence images. Either a 10× (N.A. 0.25) or 2× (N.A. 0.10) objective was used. The 514.5 nm line from an air-cooled Krypton-Argon laser (Midwest Laser Products, LLC) was directed through the rear port of the microscope and used for bleach pulses in fluorescence recovery after photobleaching (FRAP) experiments. A sliding mirror allowed rapid switching between the laser (bleach) and arc lamp (probe) light sources. A FRAP data set typically contained 500-1000 images that were acquired using 0.05-0.2 s camera exposures and a 300 ms photobleach pulse. The diameter of the bleached spot in all experiments was 12 pixels, corresponding to 19.2 µm. The FRAP data sets were converted from the Winview file format to an ASCII format and were further analyzed using a Matlab (version 6.5, The MathWorks, Inc.) program that was written to acquire average intensity values from specified pixels for each frame in the data set. The output of the Matlab program was then plotted using a standard software package to generate fluorescence intensity as a function of time. For the FRAP studies, the data were normalized to the initial (pre-photobleach) value, which enables the percentage of photobleaching and the percentage fluorescence recovery within the laser region to be determined. Time constants for recovery curves were obtained by fitting the FRAP data to single and double exponential expressions. Additional experimental details and procedures are located in the Supporting Information.
Results and Discussion Our observations indicate that vesicle interaction with 6.6 µm thick CCs composed of 330 nm silica nanospheres is dependent upon the initial hydration state of the crystal prior to incubation with lipid vesicle suspensions. Specifically, direct placement of an aqueous vesicle suspension (100 nm diameter vesicles) on a dry CC (void space occupied by air) results in crystal internalization of lipid vesicles, whereas a pre-wet CC (void space
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Langmuir, Vol. 24, No. 5, 2008 1631
Figure 1. (a) Cartoon depicting the incubation of a wet (top) and dry (bottom) CC with an aqueous vesicle solution. (b) Fluorescence image of vesicle deposited bilayer on CC surface before and after (c) contact with cotton swab. (d) Fluorescence image of internalized lipid films before and after (e) damage was inflicted with a metal syringe. The perimeter of the drop contact area is evident by rings of higher fluorescence intensity. Fluorescence intensity scales are not normalized. Table 1. Summary of Collected Fluorescence Dataa single exponential lipid films internalized, formed by vesicle fusion internalized, formed by surfactant depletion internalized, formed by chloroform evaporation planar bilayer on flat surface planar bilayer on CC
normalized lipid loadingb
double exponential
τ (s)
% recovery
R2
τ1 (%)
τ2
% recovery
R2
15.8 ( 2.3
281 ( 24
87 ( 4
0.993
214 ( 8 (42)
3910 ( 665
96 ( 4
0.986
17.7 ( 2
312 ( 13
84 ( 3
0.984
208 ( 15(35)
6100 ( 2100
92 ( 2
0.997
72.5 ( 5.5
168 ( 10
88 ( 4
0.991
68.6 ( 5 (70)
368 ( 33
96 ( 2
0.996
1 ( 0.05
36.3 ( 5
100 ( 1
0.996
42.0 ( 6 (49)
26.5 ( 15
100 ( 1
0.997
1.13 ( 0.09
110 ( 10
100 ( 1
0.961
41 ( 3 (30)
154 ( 8
100 ( 1
0.950
a
Lipid coverage is normalized to a planar supported lipid bilayer on a quartz microscope slide (column 1). FRAP data were treated to nonlinear curve fitting using single and double exponential expressions for recovery. Internalized lipid films refer to those embedded within the colloidal crystal. b Data reported for intensities and recovery fits are the average of three separate films (n ) 3), except for surfactant depletion where n ) 2.
occupied by buffer) supports the fusion of a single bilayer on the crystal surface. Figure 1 shows a cartoon depicting the two incubation conditions (Figure 1a), fluorescent microscopy images of a supported lipid bilayer (Figure 1b), and internalized lipid films (Figure 1d). Evidence for the internalization of lipids is acquired by contacting the surfaces with a cotton swab.29 A supported lipid bilayer is disturbed; contrast in the image results from loss of lipid and lipid dopant (Figure 1c). No change in the image of the internalized lipid film is observed because it is protected by the CC scaffold. Forceful contact with a metal syringe tip such that the CC scaffold is damaged is required to alter the uniform image (Figure 1e). The fluorescence intensities of the films also differ. Table 1 summarizes quantitative data from fluorescence images collected in this work. Lipid coverage is normalized to the intensity observed for a standard planar supported lipid bilayer (PLSB) on a quartz microscope slide; a 15.8-fold increase in fluorescence intensity is observed from the CC with internalized lipid. When the concentration of lipid in the vesicle suspension was increased from 1 to 10 mg/mL, lipid loading increases negligibly, indicating that the CC is saturated with respect to adsorbed lipid at relatively low solution vesicle concentrations. (29) CCs used in this work are stabilized via a sinter process, otherwise the CC itself would be disturbed by this process; see ref 28.
A likely explanation for the internalization of lipid vesicles is that rapid wetting of the dry crystal from strong capillary forces causes deformation and possible extrusion of the 100 nm vesicles through 50 nm crystal pores.30 From geometric considerations, a 6.6 µm thick fcc packed CC composed of 330 nm colloids will theoretically have 88 times greater surface area than a planar surface for a given footprint.6b The available surface area for CC applications that involve modified colloids (for example, immobilized receptor) will be less than that predicted by theory due to imperfect colloid packing and steric constraints near colloid contact points that are inaccessible to finite sized molecules. The 15.8-fold increase in fluorescence observed for internalized lipid films is significantly less than predicted, however, suggesting that the surface coverage is either incomplete or discontinuous or that the lipid films do not rigorously follow the contours of the colloid surfaces. The two possibilities, continuous versus discontinuous bilayer, can be distinguished by the diffusion rates of probe lipids in the films. We observed that the CC internalized bilayers did not exhibit lipid probe mobility when they were initially formed (Figure 2), (30) Repeated passage through polycarbonate membranes can be used to generate a relatively narrow distribution of small unilamellar vesicles (SUVs). An mini-extrusion device from Avanti Polar Lipids was used to prepare vesicles, see MacDonald, R. C.; MacDonald, R. I.; Menco, B. P.; Takeshita, K.; Subbarao, N. K.; Hu, L. R. Biochim. Biophys. Acta 1991, 1061, 297-303.
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Figure 2. Fluorescence recovery after photobleaching data for CC internalized lipid films before (open circle) and after (black square) drying and rehydrating the CC.
in agreement with the result of Parikh et al.1 and indicating either the pores are obstructed by unruptured (unfused) vesicles or that the lipid material is discontinuous and separated by gaps that prevent long-range lipid diffusion. After removing the CC from buffer, drying it with gentle air flow, and re-immersion with buffer, two unexpected phenomena were observed. First, no loss of lipid was observed, as evident by consistent and uniform fluorescence intensity equivalent to initial (predry) values. By comparison, near-quantitative removal of supported lipid bilayers on nonporous planar surfaces is observed by passage across the air-liquid interface.31 Second, the internalized lipid is observed to become laterally mobile within the CC as indicated by FRAP measurements (Figure 2). This will be discussed below. The stability of the lipid films to repeated rinse/dry cycles, which included vacuum desiccation to ensure complete dehydration, is likely a result of different drying mechanisms that occur with planar surfaces and highly porous surfaces. It is well known that supported lipid bilayers are not stable to passage through the air-water interface.31 The porous structure of the CC prevents rapid loss of the crystal void liquid during drying, such as occurs with the sheeting removal of liquid from a planar surface. Instead, void liquid leaves the CC by evaporation through 50 nm pores which does not, apparently, result in drainage of lipids from the crystal. The affect desiccation has on the lamellar structure of the lipid film is unknown. A possible explanation for the post-dry recovery of lateral mobility is that disordered or discontinuous lipid material that clogs the pores and prevents long-range lateral diffusion is redeposited as more continuous dried lipid film on the colloids that is laterally mobile upon rehydration. The original models for FRAP data with a Gaussian bleach beam developed by Axelrod and co-workers assume the membrane acts as an infinite plane.32 This model works well for planar supported bilayers, and lateral diffusion coefficients can be calculated from the time constants derived from recovery curves. Alternate algorithms for curved, tubular, or spherical membranes have also been reported.33 At this point, we do not have a suitable model for the geometry of lipid films supported by CCs and have simply reported time constants (τ) returned from single and double exponential recovery (31) Holden, M. A.; Jung, S. Y.; Yang, T. L.; Castellana, E. T.; Cremer, P. S. J. Am. Chem. Soc. 2004, 126, 6512. (32) (a) Axelrod, D.; Koppel, D. E.; Schlessinger, J.; Elson, E.; Webb, W. Biophys. J. 1976, 16, 1055. (b) Koppel, D. E.; Axelrod, D.; Schlessinger, J.; Elson, E.; Webb, W. Biophys. J. 1976, 16, 1315. (33) (a) Berk, D. A.; Clark, A., Jr.; Hochmuth, R. M. Biophys. J. 1992, 61, 1-8. (b) Koppel, D. E.; Sheetz, M. P.; Schindler, M. Biophys. J. 1980, 30, 187192.
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models in Table 1. These serve as relative time parameters because the bleach spot diameter was held constant. The value of the diffusion coefficient our method returns for a planar supported bilayer is 2.5 × 10-8 cm2/s, which is typical for a fluid lipid bilayer on a silicon dioxide surface. The laterally mobile fraction is greater than 80% for all lipid modified CCs examined in this work. For most of the CC lipid films, the FRAP data fit moderately better to a double exponential recovery which appears to account for a slow approach to the recovery plateau better than a singleexponential fit. Consequently, the double exponential model predicts higher fractions of mobile lipid. The qualitative interpretations are the same for both cases: diffusion is much slower for the internalized lipids in the CC than for the planar lipid bilayer on a flat surface. To test for the possibility of increasing lipid probe mobility, other methods of introducing lipid within the crystal were examined. Detergent depletion is an alternative method to vesicle fusion for the formation of supported lipid films on some supports, such as hydrophobic monolayers or polymer cushions.34 Applied to CCs, this idea involves filling the pores with a surfactant solubilized lipid solution, followed by a dialyzing procedure where the CC is placed in buffer to form lamellar lipid films on the colloids as the more soluble surfactant diffuses out of the crystal. Given that the total pore volume of the CC can be calculated from the crystal dimensions,35 an estimate of the lipid concentration required to form a continuous bilayer on the internal surface of the CC (assuming 100% retention of the lipid and 100% removal of surfactant) is ∼100 mg/mL, which is impractical due to solubility limitations, viscosity of the resulting solution, and the amount of surfactant required to solubilize the lipid. Nevertheless, incubation of a CC with a 20 mg/mL lipid solution using the surfactant dodecyl maltoside, followed by overnight storage in 500 mL of buffer produced internalized lipid films within the CC that appeared uniform by fluorescence microscopy and were observed to be laterally mobile. Faster time constants relative to vesicle modified CCs were not observed (Table 1). The lipid loading level was not appreciably greater than the results obtained by the use of vesicles, although the 17-fold increase in lipid loading relative to a planar bilayer (17:88 ) 19.3% of the estimated increase possible) agrees remarkably well with the theoretical concentration of lipid required to completely coat the CC. Repeating the process of lipid/surfactant exposure did not appreciably increase the loading level but did cause a decrease in fluorescence image homogeneity (see images in the Supporting Information). This approach may be useful for the modification of inverse CCs or for CCs fabricated from larger diameter colloids where the relative void volume to surface area is larger. The final method of lipid loading investigated was solvent evaporation from a solution of lipids in chloroform in the presence of a CC. CCs modified in this manner contained a large amount of fluorescently doped lipid, as evident by the colorful hue of the crystals after deposition. Fluorescence images of the rehydrated CC show the lipid loading is ∼4-fold greater than either of the other loading methods examined, 72-fold greater than a single lipid bilayer, and remarkably uniform over the entire area of the CC that had been immersed in the lipid solution (34) (a) Burgess, J. D.; Jones, V. W.; Porter, M. D.; Rhoten, M. C.; Hawkridge, F. M. Langmuir 1998, 14, 6628-6631. (b) Ataka, K.; Giess, F.; Knoll, W.; Naumann, R.; Haber-Pohlmeiser, S.; Richter, B.; Heberle, J. J. Am. Chem. Soc. 2004, 126, 16199. (35) Mittleman, D. M. B.; Jiang, P.; Hwang, K. S.; Covin, V. L. J. Chem. Phys. 1999, 111, 345-354.
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Figure 3. Fluorescence images of evaporation deposited POPC films doped with 1% Texas Red-DHPE. The image in (a) collected using a 10× objective, the image in (b) with a 2× object near the immersion line of the CC. Both images illustrate the uniformity of the deposited lipid films on CCs. Very slight streaking in image (b) is observed but is less than 4% variant across the image.
Figure 4. (a) Selected frames from fluorescence recovery movie for an evaporation deposited lipid film at 0, 30, 60, 120, and 600 s (left to right) and (b) FRAP plot of the same. Overlaid curve fits in red and green depict single and double exponential expressions for recovery respectively.
(Figure 3). At low magnification some streaking is evident; the intensity across the entire CC surface nevertheless varies by less than 4%. FRAP data (Table 1 and Figure 4) indicate that chloroformdeposited bilayers have the largest percentage of laterally mobile lipid and the smallest diffusion time constants compared to the other methods, suggesting more uniform and continuous lipid films. The time constant derived from a double exponent fit rivals that of the single bilayer supported on top of the CC. As with vesicle and surfactant loading methods, repeatedly drying the crystal did not affect the fluorescence image, indicating no loss of lipid. It was observed that a direct, high-pressure stream of N2 (air flow perpendicular to crystal surface) did have an affect on the homogeneity of the internalized lipid film.36 (see Supporting Information for drying data) This effect was not critically examined, and care was taken to dry films in a consistent (36) The pressure of the nitrogen line used was ca. 50 psi, and the stream was directed using an air gun attachment with a 1 mL disposable pipet tip attached to the end to provide fine stream control through an orifice of ∼1 mm.
manner, with relatively gentle air flow at an oblique angle to the crystal surface. Solvent evaporation resulted in an increase in lipid loading relative to a planar bilayer that is within reasonable agreement of the expected ratio of CC to planar surface area (72 vs 88). The uncertainties in the scattering cross-section and the degree of crystallinity prevent an exact prediction of the fluorescence ratio. On the basis of the approximate thickness of a POPC bilayer (5 nm), it is expected that a significant area of the CC near colloidal contact points will be too sterically constrained to support a lipid bilayer on both colloids. We estimate the actual CC surface area that could support a lipid bilayer to be 63%, resulting in a membrane surface area enhancement of 55 relative to a planar lipid bilayer (estimate derived in the Supporting Information). By this calculation, if the lipid was distributed uniformly throughout the CC, multilayers and not just single bilayers, would be present to some extent. This raises the question of the amount of CC surface area that is covered with at least one bilayer of lipid, an important issue if the materials are to be used in sensing applications. A large fraction of multilayer lipid present in the crystal would indicate that a significant amount of bare silica CC surface remains uncovered. To gain insight into the extent of lipid film coverage over the CC surface area, protein adsorption studies were conducted. Many proteins, including avidin, are known to adsorb strongly to silica surfaces, whereas phosphorylcholine (PC) based lipid films are known to be resistant to nonspecific adsorption of proteins.11,37 Adsorption of avidin-FITC to bare CCs and to CCs modified with POPC was examined by fluorescence microscopy; results are presented in Figure 5. The POPC-modified CC suppressed 93% of the avidin adsorption relative to an unmodified silica CC. To address the possibility that suppression resulted from the surface area of the modified CC being inaccessible to proteins due to pore obstruction, specific adsorption to POPC films doped with 2% biotinylated phosphoethanolamine was examined. Thirty-seven times greater intensity was observed relative to a planar lipid bilayer of the same lipid composition. At this point, it has not been determined whether the difference between the increases in specific protein binding (37×) and the lipid loading (72×) result from regions of the crystal being inaccessible to protein diffusion due to obstructed pores, steric (37) Scott, P. K.; Quan, C. Anal. Chem. 2005, 77, 327-334.
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and cover a sizable percentage of the crystal surface area, and that diffusion of avidin through the crystal pores is significant.
Conclusion
Figure 5. Graph of normalized fluorescence intensity of adsorbed avidin-FITC to unmodified and lipid coated colloidal crystals (CCs). CCs coated with POPC (second column) suppressed avidin-FITC adsorption relative to an unmodified silica CC (third column), whereas biotin-doped POPC films (fourth column) had a 37-fold increase in binding relative to planar supported bilayers with the same lipid composition (first column).
exclusion of the protein from regions near the colloid contact points, or the presence of multilayers. However, the data presented here demonstrate that the organic deposition procedure produces lipid coatings that are resistant to nonspecific protein adsorption
Lipid coated CCs may be attractive candidates for use in lipid membrane based sensor designs by virtue of the increased membrane surface area which promises stronger analytical signals in binding assays and by virtue of the stability of the films to dry storage, which is an impediment to lipid bilayer sensor applications. Furthermore, these materials provide a large lipid membrane to aqueous volume phase ratio relative to planar bilayers which would provide favorable mass transfer properties in flow-through assays. This would be particularly useful in microfluidics based sensors where minimal time is available for equilibrium binding processes and minimal surface area limits the amount of immobilized receptors. Acknowledgment. We acknowledge Gary Chandler of the University Spectroscopy & Imaging Facilities for SEM assistance. This work was funded by NIH Grant No. R01GM65980. Supporting Information Available: Fluorescence images of lipid modified CCs formed by surfactant deposition, films after drying, and films used to determine CC thickness; FRAP equations and single and double exponential fits; data pertaining to film stability to drying; calculation of theoretical area available for bilayer support. This material is available free of charge via the Internet at http://pubs.acs.org. LA7033609
