pubs.acs.org/Langmuir © 2009 American Chemical Society
Supported Bilayers Formed from Different Phospholipids on Spherical Silica Substrates )
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Gopakumar Gopalakrishnan,†,‡,§, Isabelle Rouiller,^ David R. Colman,‡,§ and R. Bruce Lennox*,†,§, Department of Chemistry, 801 Sherbrooke Street West, H3A 2K6 and ‡Montreal Neurological Institute & Hospital, 3801 University Street, H3A 2B4 and §McGill Program in Neuroengineering and Centre for Self-Assembled Chemical Structures (CSACS) and ^Department of Anatomy & Dentistry, 3640 University Street, H3A 2B2, McGill University, Montreal, Canada )
†
Received February 26, 2009. Revised Manuscript Received April 3, 2009 Spherical supported bilayer membranes (SS-BLMs) are very attractive candidates in modern bioanalytics and biorecognition studies. A uniform, facile method of preparing different SS-BLMs on silica beads is reported. Confocal fluorescence microscopy and cryo-TEM imaging have been used to characterize these SS-BLMs. Thermal analysis data and FRAP experiments show that the bilayer properties of the SS-BLM are consistent with those of lipid vesicles from which they are formed. The possibility of modulating the size, lipid type and functionality, and mechanical stability makes these rigid liposomes very attractive candidates in biosensors, drug screening, and gene delivery-related applications. This is especially true in work with native vesicle membranes derived from living cells because the existing methods can only accommodate anionic membranes to a limited extent.
Lipid vesicles (liposomes) derived from synthetic lipids have been widely used as a model system for studying various cell membrane events in vitro and as intracellular delivery vectors.1-4 Despite their extraordinary usefulness, problems with size dispersity as well as physical and chemical stability often arise. Supported bilayer membranes (S-BLMs) made from liposomes, both on planar and spherical substrates, are better candidates for addressing some of these issues.5 Like liposomes, S-BLMs in principle can offer a wide range of opportunities to systematically vary the charge type, charge density, and particular functional groups that a potential artificial cell membrane will encounter. S-BLMs also offer the possibility of introducing specific receptors/membrane proteins into the supported membrane.6 These structures are thus attractive candidates in high-throughput drug screening assays. At the same time, they avoid the polydispersity issue and increase the physicochemical stability. One of the main drawbacks of S-BLMs, however, is their adherence to planar geometry, thus lessening their versatility in applications. Spherically supported S-BLMs (SS-BLMs)7 are particularly interesting because they combine all of the positive features of *Corresponding author. E-mail:
[email protected]. (1) (a) Cevc, G.; Marsh, D. Phospholipid Bilayers: Physical Principles and Models; Wiley: New York, 1987. (b) Gennis, R. B. Biomembranes: Molecular Structure and Function, Springer; New York, 1989. (c) Mouritsen, O. G. Life as a Matter of Fat: The Emerging Science of Lipidomics, Springer; Heidelberg, 2005. (2) (a) Korlach, J.; Schwille, P.; Webb, W. W.; Feigenson, G. W. Proc. Natl. Acad. Sci. U.S.A. 1999, 96, 8641. (b) Hac, A. E.; Seeger, H. M.; Fidorra, M.; Heimburg, T. Biophys. J. 2005, 88, 317. (3) (a) Torchilin, V. P. Nat. Rev. 2005, 4, 145. (b) Kunisawa, J; Masuda, T.; Katayama, K.; Yoshihawa, T.; Tsutsumi, Y.; Akashi, M.; Mayumi, T.; Nakagawa, S. J. Controlled Release 2005, 105, 344. (4) Gopalakrishnan, G.; Danelon, C.; Izewska, P.; Prummer, M.; Bolinger :: P.-Y.; Geissbuhler, I.; Demurtas, D.; Dubochet, J.; Vogel, H. Angew. Chem. Int. Ed. 2006, 45, 5478. (5) (a) Sackmann, E. Science 1996, 271, 43. (b) Groves, J. T.; Boxer, S. G. Acc. Chem. Res. 2002, 35, 149. (6) (a) Terrettaz, S.; Ulrich, W.-P.; Guerrini, R.; Verdini, A.; Vogel, H. Angew. Chem. Int. Ed. 2001, 40, 1740. (b) Geiss, F.; Friedrich, M. G.; Heberle, J.; Naumann, R. L.; Knoll, W. Biophys. J. 2004, 87, 3213. (c) Wise, A. R.; Nye, J. A.; Groves, J. T. ChemPhysChem 2008, 9, 1688. (7) Gilbert, G. E; Drinkwater, D.; Barter, S.; Clouse, S. B. J. Biol. Chem. 1992, 267, 15861.
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liposomes with those of S-BLMs. SS-BLMs have been successfully formed on silica, polystyrene, and magnetic beads whose sizes range from tens of nanometers to several micrometers.8 SSBLMs resemble unilamellar vesicle membranes and can be formed in the range of sizes observed for vesicles (from ∼50 nm to ∼80 μm).1a,1b Because an SS-BLM diameter is determined by the support diameter, homogeneous membrane structures can be obtained when the supports are monodisperse. They also offer an opportunity for enhanced membrane stability because of their robust inorganic core, around which the bilayer membrane is wrapped. This makes them interesting materials that are useful for long-term studies and for studies that require the modulation of pH, temperature, and so forth. Because SS-BLMs are significantly easier to handle than their planar counterparts, they offer more freedom to work with different microscopy and spectroscopy techniques.8-10 The existing protocols for preparing SS-BLMs usually rely on electrostatic attraction between the bead (often negatively charged) and SUVs.8,9 They are therefore generally restricted to cationic and neutral membranes and only to a limited extent to anionic membranes.8,9b,10 Moreover, the anionic lipid contents of cell plasma membranes often exceed 20% whereas the SS-BLM format has to date been restricted up to a maximum of 20%.10 In this report, we describe a simple, uniform, versatile, and facile method of making SS-BLMs on silica beads. Unlike previous reports,8,9b this method applies equally well to cationic, neutral (zwitterionic), and negative lipid membranes, even at higher ratios of charged and phosphatidylethanolamine (PE) lipids that are comparatively difficult to accommodate in bilayers at higher concentrations. Scheme 1 shows the experimental steps used in forming the SS-BLMs on silica beads. In a typical procedure, an aqueous dispersion of silica beads (Bangs Laboratories Inc.) was (8) Troutier, A.-L.; Ladaviere, C. Adv. Colloid Interface Sci. 2007, 133, 1. (9) (a) Baksh, M. M.; Dean, C.; Pautot, S.; DeMaria, S.; Isacoff, E.; Groves J. T. Langmuir 2005, 21, 10693. (b) Baksh, M. M.; Jaros, M.; Groves, J. T. Nature (London) 2004, 427, 139. (10) Mornet, S; Lambert, O.; Duguet, E.; Brisson, A. Nano Lett. 2005, 5, 281.
Published on Web 4/21/2009
DOI: 10.1021/la9006982
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Letter Scheme 1. Scheme Illustrating the Preparation of SS-BLMsa
a
Not to scale.
washed several times, resuspended in PBS, and incubated overnight with avidin. The uniformity of the avidin coating was tested using avidin-FITC.11 After removing the excess avidin through centrifugation, the pelleted beads were resuspended in PBS and then incubated with biotinylated SUVs. After a short incubation, the mixture was vortex mixed, centrifuged, and then washed to remove any free lipid from solution. The resulting SS-BLMs, suspended in PBS, are stable for at least 2 weeks at 4 C, as confirmed using DSC and confocal experiments. There are parallel reports in which biotin-streptavidin chemistry is used to make planar S-BLMs12 and to immobilize transmembrane proteins such as GPCRs and bacteriorhodopsin in tethered lipid bilayers in an S-BLM environment.13 There are, however, no studies to date addressing the issue of developing a uniform, facile technique for making SS-BLMs that can be applied to a range of lipid groups. The method used here is derived from protocols to optimally immobilize lipid vesicles onto planar solid substrates.14 Extension of that approach allows one to make SS-BLMs from different classes of lipids and also helps in subsequent immobilization on planar substrates. Moreover, further functionalization via the free biotin moieties on the outer surface of SS-BLM is possible. High-precision microarrays of such membrane structures with enhanced mechanical stability have great potential for use in modern bioanalytical measurements.14b This approach has been tested by varying the lipid composition in the membranes with different charges, chain lengths, and functional groups. Figure 1 shows representative confocal fluorescence images of neutral, anionic, and cationic SS-BLMs formed on silica beads. The homogeneous fluorescence of the image indicates that a uniform bilayer membrane is formed around the entire bead. The confocal images of other SS-BLMs (different charge, different functional groups, etc.) closely parallel those shown in Figure 1. When negatively charged SS-BLMs (as shown in the inset of Figure 1a) were prepared using positively charged poly-L-lysine (PLL) instead of avidin to coat the bead surface, the anionic SUVs simply aggregated on the bead surface and eventually produced nonuniform fluorescence in confocal images (Figure SI.1). To confirm the formation of bilayers on the molecular level, we have performed cryo-TEM investigations of SS-BLMs of different lipids and lipid mixtures on beads spanning a range (11) FITC-labeled avidin (0.05 mg/mL) was used to coat the beads and was imaged using the same confocal imaging settings described in the Experimental Section of the Supporting Information. :: (12) Fisher, M. I.; Tjarnhage, T. Biosens. Bioelectron. 2000, 15, 463. (13) (a) Bieri, C.; Ernst, O. P.; Heyse, S.; Hofmann, K. P.; Vogel, H. Nat. Biotechnol. 1999, 17, 1105. (b) Mirzabekov, T.; Kontos, H.; Farzan, M.; Marasco, W.; Sodroski, J. Nat. Biotechnol. 2000, 18, 649. (c) Sharma, M. K.; Jattani, H.; Gilchrist, M. L. Bioconjugate Chem. 2004, 15, 942. (14) (a) Yang, Q.; Liu, X.-Y.; Miyake, J. Supramol. Sci. 1998, 5, 769. (b) Stamou, D.; Duschl, C.; Delamarche, E.; Vogel, H. Angew. Chem. Int. Ed. 2003, 42, 5580.
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Figure 1. Confocal fluorescence image of a variety of SS-BLMs formed on 5 μm silica beads. (a) The image shows a representative confocal cross-sectional image of a DOPC SS-BLM. The inset shows SS-BLMs formed from an anionic bilayer membrane (60:40% DOPC/DMPS). (b-d) SS-BLMs formed from a cationic bilayer membrane (75:25% DOPC/DOTAP). The fluorescence (b) and DIC (c) channels and their overlay (d) clearly indicate that nearly all beads had been coated with SS-BLMs. In all preparations, 0.1 mol % DSPE-PEG-biotin as well as 0.1 mol % TRITC-DHPE, a fluorescently labeled lipid, was used. The scale bars are 5 μm.
of sizes. Figure 2 shows representative cryo-TEM images of SS-BLMs. These clearly show that a uniform, single bilayer membrane is present on almost every bead examined. This was the case for all neutral, cationic, and anionic lipids tested. Beads of 160 and 500 nm exhibit the same structural properties except that the 500 nm (Figures 2 and SI.2) beads appeared to exhibit a more uniform surface (that is inherited from the bead’s surface roughness), indicating that stable SS-BLMs were formed irrespective of the surface irregularities on the substrate.15 These cryo-TEM images resemble results published elsewhere,10 confirming that our approach yields similar-quality SS-BLMs on beads. Nevertheless, SS-BLMs of anionic membranes have been reported only up to a maximum of 20% in the case of DOPS in a lipid mixture.10 Our preparation route, however, has provided for up to 40% phosphatidylserine (PS) and phosphatidic acid (PA) in the lipid bilayer (illustrated in Figures 1a (inset) and 2d, respectively). We also observe that some intact vesicle populations are present (indicated by arrows in Figure 2d) when PS lipids were used to form SS-BLMs but they remained separate from the SS-BLM. The approach presented here yields a wider range of SS-BLMs utilizing various lipid mixtures than do the existing methods.8,15 Moreover, the yield of bilayer formation on the beads is greater than the conventional electrostatic route (Figure SI.3) for similar classes of lipids. This was noticeable only in the case of 160 nm beads. Confocal imaging of 5 micron beads did not, however, show much difference in the yield. Changes in the thermotropic and lyotropic properties of lipid bilayers are characteristic of structural perturbations.1a,1b To determine whether the SS-BLMs have similar structural (15) Richter, R. P.; Berat, R.; Brisson, A. R. Langmuir 2006, 22, 3497.
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Figure 2. Cryo-TEM images of different SS-BLMs formed on differently sized silica beads. The images show representative free-standing, vitrified samples of different SS-BLMs formed on 160 nm (a-d) and 500 nm (e, f) silica beads. The uncoated beads (a) show a very clear contrast difference compared to the SS-BLM-coated beads (b-d). Mixed SS-BLMs (b-f) formed via the biotin-avidin method resemble the SS-BLMs formed via simple electrostatic spreading (c). Mixed SS-BLMs shown here include a ternary lipid mixture containing 25:25:50% DOPC/DOTAP/DPPE (b, c, e, f) as well as a binary lipid mixture containing 60:40% DOPC/DPPA (d). A close-up view (f) of the selected area from image e gives a better understanding of the bilayer spreading on the support. The entire image in (e) is shown in the Supporting Information (Figure SI.2) because both the bilayer and the support are more clearly visible as a result of the higher resolution of the image. Scale bars are 50 nm.
properties to the analogous bilayers in water, we have examined the thermotropic properties of the SS-BLMs. Unlike previous reports,8 where a small shift in Tm was observed upon SS-BLM formation, our preparation method exhibits no noticeable change in Tm (Figure 3). SS-BLMs formed via simple electrostatic attraction also do not show any shift in the Tm (Figure SI.4). This comparison is important because it demonstrates that the tethering used in this work does not influence the nature and quality of the resulting bilayer membrane. It is also noteworthy that the characteristic pretransition calorimetric feature is as distinct for all of the SS-BLMs as it is for the SUVs. The DSC thermograms of SS-BLMs after 2 weeks at 4 C are unchanged (data not shown) in Tm and net enthalpy change. Equally important in using model membranes is the mobility of the component lipid molecules in the bilayer. This capability is crucial for many signaling pathways mediated by lipid bilayers in living cells. To observe the lipid molecule diffusion within the bilayer on SS-BLMs, we have conducted comparative fluorescence recovery after photobleaching (FRAP) studies9 in a confocal microscope environment. As shown in Figure 4, the fluorescence in the bleached area has recovered after 8 s. The fluorescence intensity drops to zero in the selected area when bleached using seven iterations of 100% laser intensity (λex 488 nm, 7.5 A˚). The images show that the net fluorescence intensity of the bead that undergoes photobleaching is lower than that for the neighboring unbleached ones at the end of the time series. This is probably due to free diffusion of the remaining fluorescent lipids in the membrane until equilibrium is achieved. The overall intensity is thus reduced by about 30%, which corresponds to the bleached area. Whether the diffusion coefficient of lipid molecules is slower because of the “avidin anchor” in our preparation is not presently known. Induced membrane asymmetry has been reported when an intervening polymer layer keeps the supported bilayer distant Langmuir 2009, 25(10), 5455–5458
Figure 3. DSC thermograms of SS-BLMs on silica beads. DSC curves show that the Tm of the SS-BLMs on silica beads is the same as for the bilayer in solution (black). Differently sized silica beads were assessed in these experiments: 5 μm (red), 0.5 μm (blue), and 0.16 μm (green). The ΔT1/2 values of the peaks were measured and show a steady decrease in the peak width when going from DOPC SUVs to 160 nm SS-BLMs in the order of size. The measured values were 0.83, 0.65, 0.60, and 0.57 C for SUVs and 5 μm, 500 μm, and 160 nm for SS-BLMs.
from the underlying substrate.16 It is shown to be very useful in lipid phase domain-related studies.17 The present SS-BLM approach may thus provide a 3D variant of such asymmetric bilayers on solid supports. However, asymmetric bilayers were formed on TiO2 when Ca2+ bridging was used,18 in which negatively charged lipids preferentially occupied the leaflet that (16) Arnaud, C. H. Chem. Eng. News 2009, 87, 31. (17) Collins, M. D.; Keller, S. L. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 124. (18) (a) Rossetti, F. F.; Bally, M.; Michel, R.; Textor, M.; Reviakine, I. Langmuir 2005, 21, 6443. (b) Rossetti, F. F.; Textor, M.; Reviakine, I. Langmuir 2006, 22, 3467.
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Finally, potential SS-BLM formation using native vesicles,19 derived from living cells having negatively charged (net charge) membranes, can readily be prepared using our protocol if prior biotinylation of cellular membranes is performed. Figure 4. n w FRAP profiles of SS-BLMs on silica beads. The confocal cross-sectional images of SS-BLMs made from DOPC/ 0.1% NBD-phosphoethanolamine show fluorescence recovery after a small selected area was bleached under high laser power. The complete movie file of the experiment is included as a web enhanced object. Time series is available as an avi file. NBD is highly photobleachable, and hence the total intensity decreases as the time series progresses.
Acknowledgment. This work is supported by a grant from the Regenerative Medicine and Nanomedicine Initiative of the Canadian Institutes of Health Research (CIHR) to D.R.C. and R.B.L. and a CIHR grant (no. MOP-86693) to I.R. I.R. is recipient of a CIHR New Investigator award. We thank Petr Fiurasek (CSACS) for helping with DSC experiments and Dr. Patricia Yam (MNI&H) for helpful discussions with FRAP measurements.
is in close proximity to the substrate. Our alternative method could provide good tunability in such asymmetries in lipid bilayers on different substrates. In summary, we have revisited the formation of supported bilayers on spherical substrates and have successfully demonstrated a facile route to make SS-BLMs on silica beads using a biotin-avidin tethering protocol. Many positive features including ease of preparation, size selectivity, enhanced stability, and access to a range of lipids make this the most versatile method reported to date. Very importantly, the resulting bilayer membranes maintain the signature thermotropic properties of their electrostatically tethered counterparts and their liposome cousins.
Note Added after ASAP Publication. This Letter was published ASAP on April 21, 2009. Several text changes have been made in the manuscript. The correct version was published on April 27, 2009.
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Supporting Information Available: Detailed experimental details as well as additional data including extended figures. This material is available free of charge via the Internet at http://pubs.acs.org. (19) Pick, H.; Schmid, E. L.; Tairi, A.-P.; Ilegems, E.; Hovius, R.; Vogel, H. J. Am. Chem. Soc. 2005, 127, 2908.
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