Biomimetic Membrane Arrays on Cast Hydrogel Supports - American

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Biomimetic Membrane Arrays on Cast Hydrogel Supports Monique Roerdink Lander,†,^ Sania Ibragimova,‡,§,^ Christian Rein,§,||,^ J€org Vogel,†,§ Karin Stibius,‡,§ Oliver Geschke,†,§ Mark Perry,§ and Claus Helix-Nielsen*,‡,§ DTU-Nanotech and ‡DTU-Physics, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark § Aquaporin A/S, Ole Maaløes Vej 3, DK-2200 Copenhagen N, Denmark Nano-Science Center and Department of Chemistry, University of Copenhagen, Universitetsparken 5, DK-2100 Copenhagen Ø, Denmark

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ABSTRACT: Lipid bilayers are intrinsically fragile and require mechanical support in technical applications based on biomimetic membranes. Tethering the lipid bilayer membranes to solid substrates, either directly through covalent or ionic substrate
lipid links or indirectly on substrate-supported cushions, provides mechanical support but at the cost of small molecule transport through the membrane
support sandwich. To stabilize biomimetic membranes while allowing transport through a membrane
support sandwich, we have investigated the feasibility of using an ethylene tetrafluoroethylene (ETFE)/hydrogel sandwich as the support. The sandwich is realized as a perforated surface-treated ETFE film onto which a hydrogel composite support structure is cast. We report a simple method to prepare arrays of lipid bilayer membranes with low intrinsic electrical conductance on the highly permeable, self-supporting ETFE/hydrogel sandwiches. We demonstrate how the ETFE/ hydrogel sandwich support promotes rapid self-thinning of lipid bilayers suitable for hosting membrane-spanning proteins.

’ INTRODUCTION Biomimetic membranes based on lipid bilayers are increasingly being recognized as a platform not only for the study of reconstituted membrane-associated proteins but also as building blocks in devices for sensor and separation applications.1,2 Lipid bilayer membranes are extremely fragile and require mechanical support,3,4 and deposition of planar lipid bilayers onto supports allows for the biofunctionalization of surfaces, providing a natural environment for the immobilization of highly specialized membrane-associated proteins.5 Support surfaces with nanoscale smoothness based on gold, silicon, or cleaved mica have been investigated intensively with several membrane
protein systems.6 Such systems are well-suited for biomimetic sensors where the sensor readout is based on, e.g., detecting electrochemical impedance changes induced by ligand binding to membrane-stabilized proteins of the supported membrane
protein complex via gold electrodes or detection based on optical (fluorescence) signals. Membrane-spanning proteins often have hydrophilic moieties that may hinder optimal support of bilayers with reconstituted proteins on solid surfaces. This has led to the development of “cushions” in the form of polymeric structures that effectively separate the bilayers from the solid surface using, for example, polyacrylamide and agarose as the cushion material.7
9 In these approaches the bilayers were formed across a single aperture (100
500 μm diameter), and the cushion supported the r 2011 American Chemical Society

membrane in the aperture area. Later developments include the formation of membranes where the bilayer is cushioned by poly(ethylene glycol) (PEG)-conjugated lipids that effectively separate the bilayer from a solid surface.10,11 A different approach consists of using supports with submicrometer porosity where the bilayers are formed directly on top of, for example, nanoporous alumina substrates.12,13 Bilayer formation has also been demonstrated using ordered arrays of submicrometer-sized pores formed in silicon substrates with well-defined pore diameters ranging from 250 to 1000 nm.14,15 In this case, the membranes span the pores without additional support. Supports using hydrogels have also been demonstrated.16,17 Applications relying on mass transport across planar biomimetic membranes mediated by transmembrane proteins require that the supported membrane is stable against external forces, such as osmotic and hydrostatic forces, and concomitantly sufficiently porous to allow for vectorial transport of solutes and solvents. Tethering the lipid bilayer membranes to solid substrates, either directly through covalent or ionic substrate
lipid conjugates or indirectly by substrate-supported cushions, provides mechanical support but is not optimal for large-scale Received: December 22, 2010 Revised: March 24, 2011 Published: April 28, 2011 7002

dx.doi.org/10.1021/la1050699 | Langmuir 2011, 27, 7002–7007

Langmuir mass transport across the membrane. Large-scale transport across biomimetic membranes also requires that the entire system is readily upscalable. Approaches using nanoporous alumina require arduous preparation,13 and the use of nanopore silicon arrays is limited by material cost and wafer size. These are issues that need to be addressed to harness the unique opportunities that lipid bilayer membranes offer in technological applications where large-scale transport is desired. Previously, we reported the fast fabrication of dense arrays of 300 μm diameter circular apertures in 50 μm thick partitions made of ethylene tetrafluoroethylene (ETFE) for biomimetic membranes using a CO2 laser ablation method that is costeffective and that can be easily scaled up to square centimeters.18 We also reported successful formation of planar, free-standing black lipid membranes (BLMs) in these hydrophobic multiaperture partitions.19,20 In this study we aimed at stabilizing lipid membranes further against pressure-induced disruptions in a construction that tolerates large flux through the supported BLMs using a relatively simple upscalable porous support structure. Specifically, we formulated a composite hydrogel that demonstrates high water permeability. The hydrogel support is formed by in situ polymerization of an aqueous solution of 2-hydroxyethyl methacrylate (HEMA) and poly(ethylene glycol) dimethacrylate (PEG-DMA) in the presence of silicon dioxide particles. The composite hydrogel was cast using a mold such that the polymerized hydrogel would form a supportive layer across apertures formed in an ETFE array as shown in Figure 1A
C. The rationale behind this design was to optimize the lipid
partition interactions at the aperture rim while providing maximal support for the membranes across the aperture arrays. To ensure good contact between the hydrophilic composite hydrogel and the hydrophobic multiaperture ETFE partition, we modified the ETFE surface using a plasma surface treatment on the partition side facing the hydrogel. We demonstrate how the composite support is capable of promoting rapid self-thinning of BLMs and that the supported BLMs are suitable for hosting membrane-spanning proteins.

’ EXPERIMENTAL SECTION Materials. PEG-DMA (Mw(PEG block) = 1000 g/mol) was pur-

chased from Polysciences, Inc. (Warrington, PA). N,N,N0 ,N0 -Tetramethylethylenediamine (TEMED), R-hemolysin (Staphylococus aureus), 10 phosphate-buffered saline (PBS), pH 7.4, HEMA, 1,4-butanediol diacrylate (BDDA), silicon dioxide particles (0.5
10 μm diameters), ammonium persulfate (APS), n-decane, and ethanol were purchased from Sigma-Aldrich Denmark (Brøndby, Denmark). Tefzel ETFE LZ200 for fabrication of multiaperture arrays and Viton A fluoroelastomer used for the production of rubber chamber-sealing O-rings were supplied by DuPont Fluoropolymers (Detroit, MI). 1,2-Diphytanoyl-snglycero-3-phosphocholine (DPhPC), 1,2-dioleoyl-3-(trimethylammonium)propane (DOTAP), 1,2-distearoyl-sn-glycero-3-phosphoethanolamineN-[biotinyl(poly(ethylene glycol))-2000] (ammonium salt) (DSPEPEG2000-biotin), and 1-oleoyl-2-[6-[(7-nitro-2,1,3-benzoxadiazol-4-yl)amino]hexanoyl]-sn-glycero-3-phosphocholine (NBD-PC) were purchased from Avanti Polar Lipids, Inc. (Alabaster, AL). All materials were used as received. Fabrication of the ETFE/Hydrogel Sandwich. In the first production step the ETFE partition was plasma-treated on one side of the ETFE with HEMA in a custom-built 50 Hz two-phase plasma chamber21 using 1 sccm of Ar as the activation/carrier gas with 10 Pa of total pressure and 10 W of plasma power (20 mA, 500 V). The partition and monomer solution were left overnight to cure. Casting was performed

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Figure 1. (A) Top view of a section of the ETFE/hydrogel sandwich. The composite hydrogel is attached to the back of the ETFE without penetrating the apertures. (B) Schematic of the ETFE scaffold with PMMA rings (red) and the Teflon mold used for casting of the hydrogel support. The hydrogel-forming solution (green) is contained in a reservoir with a depth of 500 μm. (C) Optical scanning image of a single aperture with the composite hydrogel. The green toroidal structure inside the aperture represents the sloping wall of the aperture. The spikes are measurement artifacts due to missing data points. (D) Profile scan across five apertures. The red dashed lines indicate the surfaces of the ETFE sheet. with a solution consisting of 23 mg of PEG-DMA, 560 μL of HEMA, 8 μL of TEMED, 5 μL of BDDA, 140 mg of SiO2, and 1000 μL of Milli-Q water vortexed to create a milky suspension. To 400 μL of this solution was added 15.3 μL of an initiator solution (170 mg of APS in 1000 μL of Milli-Q water). The mixture was vortexed for 10 s and immediately transferred to a reservoir (2 cm diameter, 0.5 mm height) in a Teflon mold (see Figure 1B). A surface-functionalized ETFE partition with 8  8 apertures (300 μm diameter, 400 μm center-to-center spacing),18 glued between two custom-made poly(methyl methacrylate) (PMMA) rings, was fitted on top of the mold, resulting in direct contact between the monomer solution and the functionalized side of the ETFE partition. This setup allowed for the monomer solution to cover but not fill up the apertures. After 10
20 min the hydrogel-coated partitions could be removed from the mold and were stored in Milli-Q water. The hydrogel surface topography was characterized by focus-variation optical scanning of the surface (InfiniteFocus, Alicona Instruments, GmbH, Graz, Austria), shown in Figure 1C,D. Before membrane formation, excess water was removed from the hydrogel-coated partitions by blotting with tissue. BLM Formation. The membrane formation chamber used was a modified version of the horizontal chamber design described by Hansen et al.19 The chamber was assembled by placing the hydrogel-coated partition with the ETFE side up in the cis chamber between two O-rings. An open lid sealed the assembly. Both the cis chamber and the channel between the cis and trans chambers were filled with 1 mL of 1 PBS 7003

dx.doi.org/10.1021/la1050699 |Langmuir 2011, 27, 7002–7007

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Figure 2. (A) Comparison of mean ( SD specific capacitance values Cm of unsupported free-standing (black circles, n = 9) and hydrogelsupported (triangles and asterisks, n = 8) membrane arrays consisting of 8  8 individual membranes versus time. Two trends were observed for the hydrogel-supported membranes: (1) development of the specific capacitance to a mean value of 0.3 μF/cm2 (red asterisks, n = 4); (2) development of the specific capacitance to a mean value of 0.6 μF/cm2 (black triangles, n = 4). (B) Specific resistance for the membranes shown in (A). (filtered through a 0.2 μm filter). The apertures were pretreated by depositing 4 μL of lipid solution and incubating for 1 h. The lipid solution used in membrane preparation for electrical measurements contained 294 μL of DPhPC (72 mol %), 61 μL of DOTAP (18 mol %), and 145 μL of DSPE-PEG2000-biotin (10 mol %). Lipid solutions in chloroform (10 mg/mL) were dried under nitrogen gas. The dried lipid film was subsequently redissolved in 200 μL of decane (25 mg/mL). The lipid solution used in membrane preparation for fluorescence measurements contained 292 μL of DPhPC (71 mol %), 61 μL of DOTAP (18 mol %), 146 μL of DSPE-PEG2000-biotin (10 mol %), and 39 μL of NBD-PC (1 mol %). Lipid solutions in chloroform (10 mg/mL) were dried and redissolved in decane as described above. Lipid solutions were deposited on top of the apertures in 2 μL aliquots up to 10 μL. One mL of 1 PBS (filtered through a 0.2 μm filter) was added to the cis chamber, which was subsequently sealed. Membrane formation was monitored by electrical capacitance and conductance measurements using standard electrophysiological methods as previously described.22 Briefly, the membranes were voltage clamped using AgCl electrodes in each chamber compartment, and transmembrane currents were recorded using a patch-clamp amplifier. The membrane resistance R and capacitance C were measured by applying rectangular and triangular waveforms, respectively (Vpp= 10 mV). The response signal was low pass filtered at