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Amphiphobic Septa Enhance the Mechanical Stability of Free-standing Bilayer Lipid Membranes Daichi Yamaura, Daisuke Tadaki, Shun Araki, Miyu Yoshida, Kohei Arata, Takeshi Ohori, Ken-ichi Ishibashi, Miki Kato, Teng Ma, Ryusuke Miyata, Hideaki Yamamoto, Ryugo Tero, Masao Sakuraba, Toshio Ogino, Michio Niwano, and Ayumi Hirano-Iwata Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b00747 • Publication Date (Web): 17 Apr 2018 Downloaded from http://pubs.acs.org on April 17, 2018

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Amphiphobic Septa Enhance the Mechanical Stability of Free-standing Bilayer Lipid Membranes Daichi Yamaura 1, Daisuke Tadaki 1, Shun Araki 1, Miyu Yoshida 1, Kohei Arata 1, Takeshi Ohori 1, Ken-ichi Ishibashi 2, Miki Kato 1, Teng Ma 3, Ryusuke Miyata 1, Hideaki Yamamoto 4, Ryugo Tero 5, Masao Sakuraba 1, Toshio Ogino 6, Michio Niwano 7 & Ayumi Hirano-Iwata 1,3* 1 Laboratory for Nanoelectronics and Spintronics, Research Institute of Electrical Communication, Tohoku University, 2-1-1 Katahira, Aoba-ku, Sendai, Miyagi, 980-8577, Japan. 2 Hang-Ichi Corporation, 1-7-315 Honcho, Naka-ku, Yokohama, Kanagawa, 231-0005, Japan. 3 Advanced Institute for Materials Research, Tohoku University, 2-1-1 Katahira, Aoba-ku, Sendai, Miyagi, 980-8577, Japan. 4 Frontier Research Institute for Interdisciplinary Sciences, Tohoku University, 6-3 Aramaki-Aza-Aoba, Aoba-ku, Sendai, Miyagi, 980-8578, Japan. 5 Department of Environmental and Life Sciences, Toyohashi University of Technology, Toyohashi, Aichi 441-8580, Japan 6 The Instrumental Analysis Center, Yokohama National University, Yokohama National University, Tokiwadai 79–5, Hodogaya-ku, Yokohama 240-8501, Japan 7 Kansei Fukushi Research Institute, Tohoku Fukushi University, 6-149-1 Kunimi-ga-oka, Aoba-ku, Sendai, Miyagi, 989-3201, Japan. *Authors to whom correspondence should be addressed. Email: [email protected].

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Abstract Artificial bilayer lipid membranes (BLMs) provide well-defined systems for investigating the fundamental properties of membrane proteins, including ion channels, and in screening the effect of drugs that act on them. However, the application of this technique is limited due to the low stability and low reconstitution efficiency of the process. We previously reported on improving the stability of BLM based on the fabrication of microapertures having a tapered edge in SiO2/Si3N4 septa and efficient ion channel incorporation based on vesicle fusion accelerated by a centrifugal force. Although BLM stability and incorporation probability were dramatically improved when these approaches were used, some BLMs ruptured when subjected to a centrifugal force. To further improve BLM stability, we investigated the effect of modifying the surface of the SiO2/Si3N4 septa on the stability of BLM suspended in the septa. The modified surfaces were characterized in terms of hydrophobicity, lipophobicity and surface roughness. Diffusion coefficients of lipid monolayers formed on the modified surfaces were also determined. Highly fluidic lipid monolayers were formed on amphiphobic substrates that had been modified with long-chain perfluorocarbons. Free-standing BLMs formed in amphiphobic septa showed a much higher mechanical stability, including tolerance to water movement and applied forces with and without proteoliposomes, than those formed in septa that had been modified with a short alkyl chain. These results demonstrate that highly stable BLMs are formed when the surface of the septa has amphiphobic properties. Since highly fluidic lipid monolayers that are formed on the septa seamlessly connect with BLMs in a free-standing region, the high fluidity of the lipids contributes to decreasing potential damage to BLMs when mechanical stresses are applied. This approach to improving BLM stability increases the experimental efficiency of BLM systems and will contribute to the development of high-throughput platforms for functional assays of ion channel proteins.

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Introduction The cell membrane is composed of a bilayer lipid membrane (BLM), which is self-assembled via hydrophilic and hydrophobic interactions.1 The resulting structure provides membrane proteins with an environment that allows them to retain their structure and functions. An ion channel is a membrane protein that regulates ion traffic across a cell membrane. Because ion channels play a crucial role in maintaining the concentration gradient of ions in living cells, they have attracted substantial interest as a major target for drug design.2–4 Electrical recordings of ion permeation through ion channels are the most common approach to characterizing ion channel functions. Although the patch-clamp method has been the gold standard for evaluating ion channel activities,5,6 this method also has limitations, in that the observed currents may depend on the condition of the target cells and/or other channels that are also present in the cell membrane.5 Reconstituting ion channels into artificial bilayer lipid membranes (BLMs) represents an alternative to the patch-clamp method.7–10 The BLM system has the advantage that the composition of the system can be precisely controlled. In addition, the super high resistance of BLMs make them suitable systems for recording ion channel activities at the single-channel level. However, the BLM reconstitution system suffers from two major drawbacks, i.e., it suffers from instability of the BLM system and low probability of channel incorporation, which impede their wide application as drug screening devices. The simplest way to enhance BLM stability is to decrease the diameter of the apertures in which they are formed,11–15 However, the improved BLM comes at the expense of incorporation efficiency, because a decreased BLM area makes it more difficult to incorporate ion channels, especially those delivered by proteoliposomes.10 Numerous attempts have been made to overcome this trade-off, including the use of a nonvolatile hydrocarbon to strengthen the contact between BLMs and the supporting septa,12,15 and fabricating apertures in which the edge is tapered in a micro-scale.16–19 We addressed this problem by designing and fabricating a microaperture that contains a smooth nano-tapered edge.20,21 The nano-tapered edge functions to reduce the mismatch between the thickness of the aperture edge and that of the lipid bilayer and to reduce distortion of the BLM around the point of contact with the aperture edge. Mechanically stable solvent-free BLMs with ion channel proteins incorporated within them have been successfully formed in nano-tapered apertures fabricated in nanometer-thick silicon nitride (Si3N4) septa.21,22

One of the most critical steps in the BLM reconstitution system is the incorporation of ion channels into BLMs. A number of approaches have been proposed to enhance the likelihood of such an incorporation, including introduction of an osmotic salt gradient across BLMs23,24 and the addition of nystatin-ergosterol complexes in proteoliposomes.25 We recently reported on a chemical-free approach for incorporating ion channels into BLMs that uses centrifugation.22 Centrifugal force results in proteoliposomes being concentrated near the BLMs, which leads to an increase in the probability of proteoliposomes to dock at the BLMs. However, some weaker BLMs were ruptured due to mechanical stress during proteoliposome fusion under conditions of centrifugation. Therefore, further enhancement in BLM stability would be necessary if a high throughput system for recording 3


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ion channel currents were to be realized. Systematic investigations of the relationship between the conditions associated with surface treatment and the stability of BLMs have been initiated. Aspinwall et al. reported that the interaction between lipid membranes and their underlying surface play an important role in the properties of BLMs and that surface modification of the underlying surface can be an effective strategy for enhancing the BLM stability26,27 In this study, we combined a surface modification approach with nanotapered microapertures to further improve BLM stability. Free-standing BLMs were formed in a nanotapered aperture fabricated in an SiO2/Si3N4 septum in which the surface was modified with different silane coupling agents. Since the free-standing BLMs seamlessly connect with lipid monolayers supported on the modified SiO2/Si3N4 septum (Fig. 1), the effects of the modified SiO2/Si3N4 surfaces on the stability of the free-standing BLMs were investigated. The applicability of our approach as the drug-screening platforms for ion channel is also discussed.

Experimental Section Materials L-α-phosphatidylcholine (Egg-PC), L-α-phosphatidylethanolamine (Egg-PE) and 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine Bsulfonyl) (Rhod-DOPE) in chloroform were purchased from Avanti Polar Lipid. 2-[4-(2-Hydroxyethyl)piperazin-1-yl]ethanesulfonic acid (HEPES), acetonitrile, toluene were purchased from Wako Pure Chemicals (Osaka, Japan). Cholesterol (Chol) was obtained from Wako Pure Chemicals and recrystallized three times from methanol. (Tridecafluoro-1,1,2,2-tetrahydrooctyl)dimethylchlorosilane (PFDS), (heptadecafluoro-1,1,2,2-tetrahydrodecyl)dimethyl-chlorosilane (PFDDS), (tridecafluoro-1,1,2,2-tetrahydrooctyl)trichlorosilane (PFTS), 3-cyanopropyldimethylchlorosilane (CPDS) and 3,3,3-trifluoropropyldimethylchlorosilane (FPDS) were purchased from GELEST, Inc (Morrisville, PA). Octadecyltrichlorosilane (OTS) was obtained from Sigma-Aldrich (St. Louis, MO). The structures of each silane coupling agent that were examined are shown in the Supporting Information (Fig. S1). Human embryonic kidney (HEK) 293 cell lines expressing the hERG channel was obtained from Anaxon AG (Berne, Switzerland) and cultured in a 37°C incubator with 5% CO2 according to the manufacture’s protocol. The cell lines were maintained in DMEM/GlutaMAX medium supplemented with 10% FBS, 1% penicillin/streptomycin under an antibiotic pressure of 100 µg/ml geneticin (GIBCO, Waltham, MA). The hERG channels were extracted from the HEK 293 cell lines as membrane fractions, according to the procedures in ref. 22. Surface Functionalization and Characterization FZ Si (100) wafers (>9000 Ω cm, 200 µm in thickness), one side of which was coated with 200 nm-thick Si3N4 and 100 nm-thick SiO2 layers, were purchased from Semitec (Chiba, Japan). These 4


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SiO2/Si3N4/Si substrates were cleaned by ultra-sonication in chloroform, ethanol, acetone and toluene or acetonitrile, followed by ashing using an air plasma (PM100, Yamato Scientific, Tokyo, Japan). After rinsed with toluene or acetonitrile, the substrates were then immersed in 2% (v/v) solutions of silane coupling agents in a solvent at room temperature for 1-6 h in a nitrogen-filled glove box. Acetonitrile was used as the solvent for CPDS and FPDS surface modifications, while toluene was used for OTS, PFDS, PFDDS, and PFTS modifications. After the reaction, substrates were rinsed successively in acetonitrile or toluene, acetone, ethanol, and chloroform and dried with a stream of N2 gas. The static contact angles of pure water and n-hexadecane (0.5 µL) on each modified substrate were measured by the sessile droplet method. The modified substrate was placed on a level stage of a contact angle analyzer (LSE-B100, Nick Co. Ltd., Saitama, Japan) and the static contact angle of a droplet was then determined. The root-mean-square (RMS) surface roughness for each modified surface was measured by tapping mode atomic force microscopy (AFM) (Dimension Icon® AFM System, Veeco Instruments, Inc.). The image scan area was 1 µm2. The chemical state of each modified surface was investigated by X-ray photoelectron spectroscopy (XPS) (AXIS-NOVA, Kratos Manchester, UK) with Al Kα radiation. All data are presented in the form of the mean ± standard deviation. Measurement of Diffusion Coefficient in Lipid Monolayers Formed on Modified Surfaces 1.05 mg of Egg-PC, 0.15 mg of Egg-PE, 5.2 × 10-3 mg of Rhod-DOPE and 0.3 mg of cholesterol were dissolved in chloroform in a glass vial. Chloroform was evaporated with stream of N2 for 5 min followed by evacuation in a vacuum desiccator for at least 6 h. The dried lipid films were dissolved in a 5-mL aliquot of buffer solution (120 mM KCl and 10 mM HEPES/KOH, pH 7.2) to give a lipid concentration of 0.40 mM and vortexed at 50 ℃ for 1 h to prepare multilamellar vesicle suspensions of phospholipids. The vesicle suspension was then frozen and thawed five times in liquid nitrogen and this suspension was extruded five times through an 800 nm polycarbonate filter, followed by extrusion through a 100 nm polycarbonate filter five times to obtain unilamellar vesicles.28 Lipid monolayers were formed on the substrates by the vesicle fusion method.29–33 In brief, SiO2/Si3N4/Si substrates modified with PFDS, PFDDS, PFTS, OTS, CPDS or FPDS were attached to the center of a glass bottom dish (D11130H, Matsunami Glass, Osaka, Japan) with glue. A 400 µL aliquot of the filtered vesicle suspension was then dropped on each substrate followed by incubation for 1.5 h in the dark at room temperature. Finally, the substrate was rinsed with the buffer solution 10 times. The diffusion coefficients of Rhod-DOPE in the lipid monolayer formed on each substrate was measured by fluorescence recovery after photobleaching (FRAP) 34–36 using a laser scanning microscope (A1, Nikon, Tokyo, Japan) with a 561 nm laser as the excitation light sourse. FRAP was performed according to the procedure described in a previous report.34 Preparation of BLMs in Functionalized Chips and Evaluation of BLM Stability. Micro-apertures (Φ: 20-40 µm) were fabricated in Si3N4 septa suspended over Si supports, according 5


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to the procedure described in ref. 9. During the final step in the fabrication process, a thin layer of SiO2 was sputtered on the surface of the Si chip, yielding SiO2/Si3N4/Si structures (Fig. 1). The Si chip thus fabricated was silanized by the same procedure as was used for the SiO2/Si3N4/Si substrates and could be re-used for BLM formation up to 6 times. Solvent-free BLMs were formed by the folding method37 without a coating of n-hexadecane around the apertures. Detailed procedures for the formation of BLMs are given in the Supporting Information. BLM formation was confirmed by measuring membrane resistance, which was calculated by a current difference between +100 and −100 mV with an Axopatch 200B patch-clamp amplifier (Molecular Devices, Sunnyvale, CA). A lipid solution (Egg-PC: Egg-PE: cholesterol =7: 1: 2 (w/w)) in chloroform/n-hexane (1:1, v/v) was used to prepare the BLM. The probability of BLM formation was defined as the percentage of BLMs with a resistance higher than 100 GΩ. Breakdown voltage, lifetime, tolerance to aspiration cycles (ACs) of buffer solution and tolerance to applying centrifugal force (CF) were evaluated as a measure of BLM stability, according to the procedures described in Ref. 9 (see Supporting Information). BLMs were still surrounded by water phases during the ACs, because the BLMs were formed in small recessed parts of the recording chamber. Ion Channel Reconstitution and Current Recording The reconstitution of ion channels into BLMs was performed via vesicle fusion between proteoliposomes containing hERG channels with pre-formed BLMs under centrifugation (44 × g), as described in Ref. 22 (see Supporting Information). Current recordings were performed with an Axopatch 200B patch-clamp amplifier (Molecular Devices, Sunnyvale, CA). Current signals were filtered online at 1 kHz with a low-pass Bessel filter, and stored on-line using a data acquisition system (Digidata 1440A and pCLAMP 10.3). Single-channel currents were filtered offline at a cutoff frequency of 0.7 kHz.38

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Results and Discussion Surface Characterization of Modified SiO2/Si3N4/Si Substrates Fig. 1 shows a schematic illustration of a BLM formed in a Si chip the surface of which had been modified with silane coupling agents. The BLM is seamlessly surrounded by lipid monolayers formed on a modified SiO2/Si3N4/Si septum. The surface properties of the modified SiO2/Si3N4/Si substrate, such as the hydrophobicity, lipophobicity and roughness, and the fluidity of the lipid monolayers on the modified substrate, may significantly affect the stability of the BLMs in a free-standing region. We first investigated the hydrophobicity and lipophobicity of SiO2/Si3N4/Si substrates that had been modified with various silanes by measuring the water and oil (n-hexadecane) contact angles of each modified substrate, respectively (Fig. 2). Table 1 summarizes the water contact angles, oil contact angles, and surface roughness of the substrates modified with PFDS, PFDDS, PFTS, OTS, CPDS and FPDS (Fig. S1). Perfluorinated silanes and CPDS have frequently been utilized for modifying the surface of Si or glass chips that are used for suspending BLMs.12,20,26,39 OTS was also examined, because it is well known that lipid monolayers can be formed on OTS-modified surfaces.30,31 The water contact angles for substrates that were modified with silanes containing a long perfluoro chain (PFDS, PFDDS, PFTS) or a long hydrocarbon chain (OTS) were larger than 90º; while the substrate modified with a silane having a short hydrocarbon chain and a polarized head group (CPDS) showed a smaller water contact angle (74º), indicating that the CPDS-modified substrates were less hydrophobic. On the other hand, oil contact angles larger than 60º were observed in the case of substrates that were modified with PFDS, PFDDS and PFTS, while smaller oil contact angles of 36º and 12º were observed in the case of OTS- or CPDS-modified substrates, respectively. These water and oil contact angles on all modified surfaces are in agreement with previously reported data.26,27,40– 43

The surface roughness of the modified substrates was also measured, because the surface roughness of the modified septa could also affect the seal between the lipid membranes and the septa, and consequently the formation of BLMs. Before the surface modification, the SiO2/ Si3N4/Si

substrate exhibited a surface roughness of 0.26 ± 0.01 nm. After modification, all of the modified surfaces with the exception of PFTS showed a surface roughness smaller than 0.7 nm, consistent with previously reported findings.26 The PFTS-modified surface showed a large roughness of 1.30 ± 0.02 nm, probably due to the formation of aggregates. As shown in XPS spectra of the PFTS-modified substrate (Fig. S2), the F1s peak of the PFTS-modified substrate was much more intense than that of PFDS-modified substrate, confirming the formation of PFTS aggregates on the substrate. Diffusion Coefficient of Lipids on Modified SiO2/Si3N4/Si Substrates We next examined the formation of lipid monolayers on the modified surfaces by means of the vesicle fusion method.30–33 When the SiO2/Si3N4/Si substrates modified with PFDS, PFDDS, PFTS, OTS, CPDS or FPDS were treated with vesicles containing Rhod-DOPE, the substrates showed a 7


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uniform fluorescent intensity with no evidence of vesicle precipitates (Fig. 3), indicating that uniform and continuous lipid layers had been formed on the substrates. Since it has been clearly established that lipid monolayers are formed on OTS-modified substrates via vesicle fusion,30,31 the observed uniform fluorescence on the OTS-modified SiO2/Si3N4/Si substrate indicates that lipid monolayers were formed on it. In addition, all of the modified substrates except for the PFTS-modified sample showed a fluorescent intensity in the range of 2.6-3.1 × 103 after the substrates were treated with Rhod-labelled vesicles. Considering that the fluorescent intensity of lipid membranes containing labelled lipids is dependent on the number of lipid layers (Table S1),44 these results indicate that lipid monolayers are uniformly formed on the substrates that were modified with PFDS, OTS, PFDDS, FPDS and CPDS. Although a lower fluorescence intensity of 2.2 × 103 was observed in the case of the PFTS-modified substrate probably due to the formation of large aggregates, the fluorescence after photobleaching was recovered to over 90% (Fig. S3), indicating that lipid monolayers were formed on the substrates. We then evaluated the fluidity of the lipid monolayer formed on the substrates modified with PFDS, PFDDS, PFTS, OTS, CPDS and FPDS. The diffusion coefficient of lipid monolayers was calculated based on FRAP measurements (Fig. 3, Fig. S3 and Table 1). The diffusion coefficient of the lipid monolayer formed on the OTS-modified substrate was calculated to be 1.50 ± 0.45 µm2/s, which was in agreement with previously reported values for OTS-modified surfaces.45,46 Lipid monolayers that were formed on amphiphobic PFDS- and PFDDS-modified surfaces, which showed large water and oil contact angles, had high diffusion coefficients of 1.74 ± 0.43 and 1.48 ± 0.28 µm2/s, respectively. In the case of the FPDS- and CPDS-modified surfaces, which showed smaller water and oil contact angles, the lipid monolayers exhibited much lower diffusion coefficients of 0.94 ± 0.16 and 0.67 ± 0.28 µm2/s, respectively. These results suggest that a lower interaction between lipid tails and modified surfaces allows lipid molecules to glide more rapidly over the surface. The PFTS treatment also resulted in the formation of an amphiphobic surface, but the lipid monolayers formed on PFTS-modified surfaces had less fluid characteristics than those on the PFDS- and PFDDS-modified surfaces, possibly because the high surface roughness interferes with the diffusion of lipid molecules. A similar dependence of lipid fluidity on surface roughness has also been reported by using supported lipid bilayers formed on diamond with a different surface roughness.47 These results indicate that lipid monolayers with a high level of fluidity can be prepared on smooth amphiphobic surfaces. In the experiments outlined below, PFDS- and CPDS-modified surfaces were used for preparing and charactering BLMs, since the lipid monolayers formed on these surfaces showed the largest difference in lipid fluidity. Although the BLMs were formed by the folding method, and consequently, lipid monolayers surrounding the BLM region were formed on SiO2/Si3N4/Si chips by the folding method, such difference in the method used for monolayer formation would have only a negligible effect on the fluidic properties of the lipid monolayer. This is supported by an observation that similar diffusion coefficients were observed for outer lipid monolayers of supported lipid bilayers, irrespective of the methods used to prepare them: either by vesicle fusion with supported 8


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lipid bilayers or by Langmuir-Blodgett deposition on supported lipid bilayers.48

Stability of BLMs Formed on PFDS- and CPDS-Modified SiO2/Si3N4/Si Chips The probability of forming solvent-free BLMs on SiO2/Si3N4/Si chips treated with PFDS or CPDS was investigated (Table 2). Solvent-free BLMs could be reproducibly formed on the PFDS-modified chips, with an 86% probability of formation. The probability was much lower (47%) in the case of CPDS-modified chips. We conclude that the difference in formation probability is due to differences in hydrophobicity, because a hydrophobic surface is necessary for producing a uniform tail-down lipid orientation, which is required for forming BLMs. 26,27 Both of the BLMs formed on PFDS- and CPDS-modified chips exhibited similar background noise currents, suggesting that the background noise level was not affected by modifying the surface of the chips. To examine the effect of surface modification of the chips on the stability of free-standing BLMs, we compared the stability of BLMs formed on PFDS- and CPDS-modified Si chips by measuring the average lifetime, breakdown voltage, tolerance to movement of the solutions surrounding BLMs, and tolerance to centrifugal force (CF). We first evaluated static stability in terms of average lifetime and breakdown voltage. The membrane lifetime was determined to be 15.5 ± 13.1 h (n = 7) for BLMs on PFDS-modified chips and 16.8 ± 10.8 h (n = 9) for the CPDS-modified samples. Neither of the BLMs were broken, even when a high voltage (± 1000 mV) was applied for periods in excess of 30 s. These findings indicate that there was no significant difference between the BLMs formed on PFDSand CPDS-modified chips in terms of static membrane stability. We next evaluated the mechanical stability of the solvent-free BLMs formed on PFDS- and CPDS-modified Si chips in terms of tolerance to mechanical shock during repeated movement of the solution-air interface over the BLMs. The buffer solutions surrounding BLMs were removed from the aqueous compartments by aspiration and the same volumes of the solutions were reintroduced. This aspiration-reinjection cycle was then repeated.9,17 The solution movement during the aspiration-reinjection cycle mimics conditions associated with exchanging the solutions surrounding the BLMs. A higher tolerance to repeated solution exchange is preferable for enhancing experimental throughput. It was found that 75% of the BLMs formed on PFDS-modified chips showed a membrane resistance in exceess of 1 GΩ, even after 20 aspiration cycles (ACs). On the other hand, only 50% of the BLMs formed on CPDS-modified chips survived the same treatment. Thus, the BLMs formed on the PFDS-modified surface exhibited a higher tolerance to ACs. As another measure of the mechanical stability of BLMs, we also compared the tolerance of BLMs to a CF with and without proteoliposomes containing human ether-a-go-go-related gene (hERG) channels. Applying a CF to BLMs has been used to improve the incorporation efficiency of ion-channel into BLMs probably due to centrifugal condensation of proteoliposomes in the vicinity of the BLMs, but unstable BLMs rupture during this process.22 As shown in Table 3, when the BLMs were centrifuged at 55 × g in the absence of proteoliposomes, all of the BLMs on the PFDS-modified 9


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chips survived this treatment. In contrast, half of the BLMs on the CPDS-modified chips were broken during centrifugation. In the presence of proteoliposomes containing hERG channels, centrifugation enhanced the probability of proteoliposome fusion, which can have a more severe impact on BLMs, even if lower CF of 40 × g was applied. Decreased survival rates of 71 % and 43 % were observed for the BLMs formed on PFDS- and CPDS-modified chips, respectively. In both experiments, the mechanical stability of the BLMs formed on the PFDS-modified chips was clearly superior to those formed on the CPDS-modified chips. As demonstrated in the FRAP measurements, lipid membranes formed on the PFDS-modified surface showed a considerably higher diffusion coefficient than that formed on the CPDS-modified surface. These results confirm that the fluidity of lipids on the supported region is strongly correlated with the mechanical stability of free-standing BLMs. We hypothesize that the higher fluidity of lipids on septa modified with PFDS is preferable for compensating posible defects by a meachanical force on BLMs or during proteoliposome fusion. Overall, the PFDS-modified surfaces provided a more favorable platform for lipid bilayer formation owing to their smooth and high amphiphobic surfaces. Recording Ion Channel Activities in BLMs Finally, we examined the functionality of BLMs formed on PFDS-modified SiO2/Si3N4/Si chips through the incorporation of hERG channels into the BLMs. The hERG channel is a cardiac voltage-dependent potassium channel that plays a crucial role in electrical activity in the human heart. This channel has recently attracted considerable attention, because its relation to arrhythmic side effects following drug treatment is well recognized.49 Fig. 4 shows example traces of hERG single-channel currents. Stepwise currents with a single conductance of 12 pS were recorded. This conductance level was similar to those (11-13 pS) reported for hERG channels in a BLM reconstitution system21 and hERG channels expressed in Xenopus oocytes.38 The channel activities were completely blocked, when astemizole was added to trans side of the BLM. Astemizole is an antihistamine, whose adverse effect on the hERG channel is well recognized.49 This confirms the functional reconstitution of hERG channels into the BLMs formed in the PFDS-modified Si chips.

Conclusion We report herein on the effects of the surface modification not only on the fluidic nature of lipid monolayers formed on modified SiO2/Si3N4/Si septa but also on the stability of free-standing BLMs surrounded by a modified septum. The diffusion coefficient of lipids on the modified surfaces was affected by several different factors, including hydrophobicity, lipophobicity and surface roughness. Highly amphiphobic surfaces with water and oil contact angles higher than 90º and 60 º, respectively, lead to the formation of highly fluidic lipid monolayers on them, while a large surface roughness of 1.3 nm reduces the fluidity of the lipid monolayers. The findings also show that the stability of BLMs was also affected by the type of surface modification of the septa. BLMs on PFDS-modified 10


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chips had a higher formation probability (>80%) and a higher mechanical stability, such as tolerance to movement of water surrounding BLMs and tolerance to applied centrifugal forces, than BLMs that were formed on CPDS-modified chips. Since the lipid monolayers on the septa seamlessly connect with BLM in a free-standing region, the high lipid fluidity contributes to reducing potential damage to BLMs when mechanical stresses are applied. This simple approach leads to an improved experimental throughput for BLM-based reconstitution systems and contributes to realization of high-throughput drug screening platforms for use in studies of ion channel proteins after a further enhancement of the bilayer stability.

ASSOCIATED CONTENT Supporting Information. Additional description on structure of the silane coupling agents, XPS spectra of the PFDS- and PFTS-modified surfaces, and fractional fluorescence recovery vs. time of lipid monolayers on the OTS-, PFDDS-, FPDS- and PFTS-modified surfaces. This information is available free of charge via the Internet at http://pubs.acs.org. Acknowledgement This work was supported by the CREST program of the Japan Science and Technology Agency (JPMJCR14F3) and Grants-in-Aid for scientific research from the Japan Society for the Promotion of Science, Japan (15H03822). Nation-wide Cooperative Research Projects, Research Institute of Electrical Communication, Tohoku University are also acknowledged. The authors also wish to thank Yoshiaki Okamoto at the Toyohashi University of Technology for the FRAP measurements and useful discussions. The authors also wish to thank Takenori Tanno in the Fundamental Technology Center, Tohoku University for the XPS measurements. Some of the equipment used in this research was manufactured by Kento Abe, a technical staff member in the machine shop division of Fundamental Technology Center, Research Institute of Electrical Communication, Tohoku University.

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Figure 1. Schematic illustration of a BLM in a microaperture fabricated on a Si chip in which the surface was treated with a silane coupling agent. Figure 2. Sessile droplet contact angles on planar SiO2/ Si3N4/Si substrates; (a-d) H2O and (e-h) n-hexadecane. (a,e) PFDS, (b,f) CPDS, (c,g) OTS and (d,h) PFTS. Figure 3. Typical Fluorescence images and fractional fluorescence recovery vs time for Rhod-DOPE-supported lipid monolayers on (a) PFDS and (b) CPDS modified surfaces. Figure 4. Typical hERG single-channel currents recorded at –100 mV after a 300-ms prepulse of +50 mV. (a) Before the addition of astemizole. (b) After adding astemizole to the trans side of the recording chamber. The final concentration of astemizole in the chamber was 8.5 nM.

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Table 1. Surface Characterization of Modified Substrates and Diffusion Coefficients of Lipid Monolayers Formed on the Substratesa surface

water contact

n-hexadecane contact surface roughness

diffusion

modification

angle [degree]

angle [degree]

RMS [nm]

coefficient [µm2/s]

PFDS

107± 2

63 ± 2

0.47 ± 0.22

1.74 ± 0.43

OTS

113 ± 1

36 ± 1

0.39 ± 0.10

1.50 ± 0.45b

PFDDS

109 ± 1

65 ± 3

0.65 ± 0.15

1.48 ± 0.28

FPDS

90 ± 1

40 ± 1

0.53 ± 0.11

0.94 ± 0.16

PFTS

116 ± 1

78 ± 1

1.30 ± 0.02

0.92 ± 0.13

CPDS

74 ± 1

12 ± 1

0.36 ± 0.11

0.67 ± 0.28

a

Number of trials is 5 except for the measurement of lipid diffusion coefficient on the OTS modified-surface. bNumber of trials is 9.

Table 2. Formation Probability, Lifetime, and Background Current Noise Level of the BLMs Suspended on Modified Si Chips surface probability of modification PFDS CPDS a

BLM formation (%) 86 (42/49) 47 (30/64)

lifetime (h) 15.5 ± 13.1 (n=7) 16.8 ± 10.8 (n=9)

noise width a

RMS (pA) 0.88 ± 0.61 (n=44) 0.78 ± 0.42 (n=17)

p-p noise width (pA) 3.5 ± 1.8 (n=44) 3.6 ± 2.1 (n=17)

The RMS noise was calculated from the current trace for 1 s.

Table 3. Mechanical Stability of the BLMs Formed on Modified Si Chipsa Surface tolerance to CF tolerance to CF tolerance to ACsb c modification without proteoliposomes with proteoliposomesd PFDS CPDS

75% (9/12) 50% (3/6)

100% (11/11) 50% (5/10)

71% (12/17) 43% (3/7)

a

Stability of BLMs formed in the PFDS- or CPDS-modified Si chips. Survival probability was calculated by dividing the number of BLMs whose resistance remained over 1 GΩ (lowest sealing resistance required for single-channel recordings) after the tolerance test by the number of BLMs whose resistance was in excess of 100 GΩ before the tolerance test. bNumber of aspiration cycles

(ACs): twenty. cCondition of centrifugation: 55 × g for 10 minutes. dCondition of centrifugation: 40 × g for 10 minutes with proteoliposomes.

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Table of Contents Graphic

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Figure 1. Schematic illustration of a BLM in a microaperture fabricated on a Si chip in which the surface was treated with a silane coupling agent. 326x135mm (150 x 150 DPI)


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Figure 2. Sessile droplet contact angles on planar SiO2/SiN/Si substrates; (a-d) H2O and (e-h) nhexadecane. (a,e) PFDS, (b,f) CPDS, (c,g) OTS and (d,h) PFTS. 389x119mm (150 x 150 DPI)


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Figure 3. Typical Fluorescence images and fractional fluorescence recovery vs time for Rhod-DOPEsupported lipid monolayers on (a) PFDS and (b) CPDS modified surfaces. 459x260mm (150 x 150 DPI)


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