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Hybrid model membrane combining micropatterned lipid bilayer and hydrophilic polymer brush Toshiki Nishimura, Fuyuko Tamura, Sawako Kobayashi, Yasushi Tanimoto, Fumio Hayashi, Yuki Sudo, Yasuhiko Iwasaki, and Kenichi Morigaki Langmuir, Just Accepted Manuscript • Publication Date (Web): 17 May 2017 Downloaded from http://pubs.acs.org on May 19, 2017

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Hybrid model membrane combining micropatterned lipid bilayer and hydrophilic polymer brush

Toshiki Nishimura1, Fuyuko Tamura1, Sawako Kobayashi1, Yasushi Tanimoto1, Fumio Hayashi2, Yuki Sudo3, Yasuhiko Iwasaki4, Kenichi Morigaki 1, 5 *,

1: Graduate School of Agricultural Science, Kobe University, Rokkodaicho 1-1, Nada, Kobe 657-8501, Japan 2: Graduate School of Science, Kobe University, Rokkodaicho 1-1, Nada, Kobe 657-8501, Japan 3: Graduate School of Medicine, Dentistry, and Pharmaceutical Sciences, Okayama University, 1-1-1 Tsushima-naka, Kita-ku, Okayama 700-8530, Japan 4: Faculty of Chemistry, Materials and Bioengineering, Kansai University, 3-3-35 Yamatecho, Suita 564-0836, Japan 5: Biosignal Research Center, Kobe University, Rokkodaicho 1-1, Nada, Kobe 657-8501, Japan

*Corresponding author: Kenichi Morigaki: E-mail: [email protected], Fax: +81-78-803-5941

1 ACS Paragon Plus Environment

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Abstract Substrate-supported planar lipid bilayers (SPBs) are being utilized as a versatile model system of the biological membrane. However, the proximity between the solid support and membrane limits utility of SPBs for the functional analyses of membrane proteins. Here, we present a model membrane that can enlarge the distance between the substrate surface and the membrane by combining a stable scaffold of polymerized lipid bilayer with a hydrophilic polymer brush. A micropatterned SPB was generated by the lithographic polymerization of diacetylene lipids and subsequent incorporation of natural

(fluid)

lipid

bilayers.

Hydrophilic

polymer

brush

of

poly-2-methacryloyloxyethyl phosphorylcholine (poly(MPC)) was formed on the surface of polymeric bilayer by the in-situ atom transfer radical polymerization (ATRP) in aqueous solution, in the presence of embedded fluid lipid bilayers. A model membrane protein (Haloquadratum walsbyi bacteriorhodopsin: HwBR) could be reconstituted into the polymer brush-supported bilayers with significantly reduced immobile molecules. Furthermore, the polymer brush terminals could be functionalized by successively polymerizing MPC and 2-aminoethyl methacrylate (AMA). The reactive amine moiety of poly(AMA) enables to conjugate a wide range of biological molecules and surfaces to the membrane. The combination of micropatterned bilayer and polymer brush mimics the two-dimensional and three-dimensional structures of the biological membrane, providing a platform to assay membrane proteins in a truly biomimetic environment.


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1. Introduction Substrate-supported planar lipid bilayers (SPBs) are being developed as a versatile model system of the biological membrane that can be applied to evaluate the membrane organization and functions in a sensitive and quantitative manner. 1-3 Micropatterning of SPBs has been exploited as a promising route to generate complex and functional model membranes.

4-7

However, the proximity between solid support and membrane limits

utility of SPBs, including the functional analysis of membrane proteins. Generally, SPBs are adsorbed on a hydrophilic surface (e.g. silica) by the van der Waals interaction. The distance between the membrane and substrate is typically 1-2 nm, which can ensure the lateral diffusion of lipid molecules but is often too small for the functional incorporation of membrane proteins.

8-10

To amend this technical drawback, lipid

bilayers have been deposited on polymeric materials (polymer cushion), 11-14 or directly anchored to the substrate with a oligomeric spacer group (tethered bilayer). 15-20

Here, we report a hybrid model membrane that combines micropatterned lipid bilayer and hydrophilic polymer brush. We previously developed a micropatterned SPB composed of polymeric and natural (fluid) lipid bilayers.

21

The polymeric bilayer was

formed from diacetylene lipids (DiynePC and DiynePE), and acted as a framework to stabilize the model membrane, whereas the embedded lipid membranes were composed of natural phospholipids and retained the physicochemical properties of the biological membrane. In the present study, we demonstrate the formation of “polymer brush” on the surface of polymeric bilayer. The polymeric bilayer acts as a stable scaffold for both the

fluid

bilayer

and

the

polymer

brush.

We

polymerized

2-methacryloyloxyethyl-phosphorylcholine (MPC) from the surface of polymeric


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bilayer by the atom transfer radical polymerization (ATRP) (Figure 1).

22

Poly(MPC) is

hydrophilic and highly biocompatible, suppressing nonspecific adsorption of proteins. 23-25

The initiator was grafted onto the surface of polymeric bilayer that contained an

ethanolamine moiety (DiynePE).

26

The ATRP reaction could be carried out in an

aqueous solution by adding a reducing agent (ascorbic acid) (activator regenerated electron transfer-ATRP: ARGET-ATRP) (Figure 1),

27-30

enabling to generate polymer

brush in the presence of fluid bilayers. The surface-initiated ATRP should form a dense and uniform layer of poly(MPC) on the polymeric bilayer. We could reconstitute a model membrane protein (Haloquadratum walsbyi bacteriorhodopsin: HwBR) into the polymer brush-supported bilayers, showing lateral mobility and significant suppression of nonspecific adsorption. Furthermore, orientation of the brush could be controlled by the distribution of DiynePE in two monolayer leaflets of the polymeric bilayer, and the brush could be functionalized for attaching biological molecules and surfaces. These unique features would help to realize a versatile model system that mimics two-dimensional and three-dimensional structures of the biological membrane, offering possibilities to assay membrane functions in a truly biomimetic environment.


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2. Materials and methods 2.1 Materials 1,2-bis(10,12-tricosadiynoyl)-sn-glycero-3-phosphocholine

(DiynePC),

1,2-bis(10,12-tricosadiynoyl)-sn-glycero-3-phosphoethanolamine

(DiynePE),

and

1,2-dioleoyl-sn-glycero-3-phosphocholine (DOPC) were purchased from Avanti Polar Lipids (Alabaster, AL). Texas Red-1,2-dihexadecanoyl-sn-glycero-phophoethanolamine (TR-PE) and streptavidin-Alexa Fluor 594 conjugate were purchased from Molecular Probes (Eugene, OR). Hilyte Fluor 750 C2 maleimide (HL750) was purchased from Anaspec (Fremont, CA). 2-Methacryloyloxyehtyl phosphorylcholine (MPC) was purchased from NOF Corporation (Tokyo, Japan). 2,2’-Bipyrydyl, copper (I) bromide, 2-aminoethyl

methacrylate

(AMA), tris(2-carboxyethyl)phosphine

hydrochloride

(TCEP) and Cyclooctatetraene (COT) were purchased from Sigma-Aldrich (St. Louis, MO).

N-hydroxysuccinimidyl-2-bromo-2-methyl-propionate

synthesized according to the previously described method.

31

(NHS-BMP)

was

Triethylamine (TEA),

N,N-dimethylformamide (DMF), 2-Mercaptoethanol, and methanol were purchased from Wako Pure Chemical Industry (Osaka, Japan). L-ascorbic acid, sodium dodecyl sulfate (SDS), octyl-β-D-glucopyranoside (OG), dodecyl-β-D-maltopyranoside (DDM), glucose oxidase, catalase, and glucose were purchased from Nakalai Tesque (Kyoto, Japan). Microscopy coverslips were purchased from Matsunami (Osaka, Japan). Silicon substrates with a native oxide film (P-doped, (100)-oriented) were purchased from Chiyoda Trading Corporation (Tokyo, Japan). Deionized water used in this experiments was ultrapure Milli-Q water (Millipore) with a resistance of 18.2 MΩ cm.


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2.2 Substrate cleaning The

substrates were cleaned with a commercial detergent solution, 0.5%

Hellmanex/water (Hellma, Mühlheim, Germany), for 20 min under sonication, rinsed with deionized water, treated in a solution of NH4OH (28%)/H2O2 (30%)/H2O (0.05:1:5) for 10 min at 65 °C, rinsed extensively with deionized water, and then dried in a vacuum oven for 30 min at 80 °C. Before use, the substrates were further cleaned by the UV/ozone treatment for 20 min (PL16−110, Sen Lights Corporation, Toyonaka, Japan).

2.3 Preparation of polymeric bilayer Bilayers composed of monomeric DiynePC and DiynePE (4:1) were deposited onto substrates from the air/water interface by the Langmuir−Blodgett (LB) and subsequent Langmuir−Schaefer (LS) methods using a Langmuir trough (HBM-AP, Kyowa Interface Science, Asaka, Japan). The temperature of the subphase (deionized water) was controlled at 16 °C by circulating thermostatted water. The surface pressure was controlled at 30 mN/ m. Polymerization of DiynePC/ DiynePE bilayers was conducted by UV irradiation using a mercury lamp (UVE-502SD, Ushio, Tokyo, Japan) as the light source. The applied UV intensity was typically 10 mW/cm2 at 254 nm, and the UV dose was modulated with the illumination time. After UV irradiation, nonpolymerized DiynePC and DiynePE molecules were removed from the substrate surface by immersing in 0.1 M sodium dodecylsulfate (SDS) solution at 30 °C for 30 min and rinsing extensively with deionized water. The polymerized bilayers were stored in deionized water in the dark at 4 °C.


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2.4 Preparation of fluid bilayers Lipids dissolved in chloroform were mixed in a round-bottom flask, dried under a stream of nitrogen, and subsequently evaporated for at least 4 h in a vacuum desiccator. The dried lipid films were hydrated in PBS overnight (the lipid concentration was 1 mM). Lipid membranes were dispersed by five freeze/thaw cycles, and the suspension was extruded by using Liposofast extruder (Avestin, Ottawa, Canada) with 100 nm polycarbonate membrane filter (10 times) and 50 nm polycarbonate filter (15 times). Extruded vesicles were applied onto a substrate having a patterned polymeric bilayer to form supported planar lipid bilayers (SPB).

2.5 Formation of polymer brush on polymeric bilayer The initiator (bromide) was attached to polymeric bilayers by immersing the samples in a solution of NHS-BMP (18.9 mM in DMF) containing 28.4 mM TEA overnight at room temperature. 32 The substrates were cleaned in 0.1 M SDS solution at 30 °C for 30 min, and rinsed ten times with deionized water. For the polymer brush formation by ARGET-ATRP, 0.8 g MPC and 7 mg NaCl were dissolved in 3.6 mL deionized water. 4 mg CuBr and 34.8 mg 2,2’-bipyrydyl were dissolved in 4.0 mL methanol. 130 mM of L-ascorbic acid (AA) was purged with argon for 30 min for removing oxygen. The MPC solution and 400 µL of the methanol solution were mixed in a 10 mL sample tube (the final concentrations of MPC, NaCl, CuBr and 2,2’-bipyrydyl were 0.67 M, 30 mM, 0.7 mM and 5.6 mM, respectively). The brominated substrates were immersed in the MPC solution. The sample tube was sealed with a rubber septum, and a needle connected to a syringe pump was inserted into the solution through the rubber septum.


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AA solution was injected with the syringe pump at the flow rate of 1 µL/ min and the ARGET-ATRP reaction started. After the reaction, the substrates were removed from the sample tube, and washed extensively with deionized water. For the successive polymerization of MPC and AMA, the solutions were exchanged after finishing the polymerization of MPC and polymerization of AMA was started in a new solution having the same conditions as MPC, except for the AMA concentration of 0.067M. The average film thickness of polymeric bilayer and polymer brush were measured by the ellipsometry (M-220K, JASCO, Hachioji, Japan) using silicon wafers as the substrates. The optical constants (refractive index (n) and extinction coefficient (k)) of Si (n=3.865, k=0.02), SiO2 (n=1.465, k=0), poly(MPC) (n=1.49, k=0), and polymeric bilayer (n=1.49, k=0) were used to calculate the thickness of polymeric bilayers and poly(MPC) brushes.

2.6 Fluorescence microscopy Fluorescence microscopy observation was performed using an upright microscope (BX51WI, Olympus, Tokyo, Japan) equipped with a xenon lamp (UXL-75XB, Olympus), a 60x water-immersed objective (NA 0.90), and a CCD camera (DP 30BW, Olympus). Two types of filter sets were used: (1) excitation 470-490 nm/emission 510-550 nm (green fluorescence), (2) excitation 545-580 nm/emission >610 nm (red fluorescence). Fluorescence images were processed with the MetaMorph program (Molecular Devices, Sunnyvale, CA). The fluidity of lipid bilayers was determined by the fluorescence recovery after photobleaching analysis using the boundary profile evolution method (FRAP-BPE).

33

Fluorescence images of TR-PE in fluid bilayers in a patterned membrane (line pattern


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with the width of 10 µm) were obtained by using the red fluorescence filter set. Photobleaching through a rectangular slit was performed by keeping the shutter open for 5 or 10 seconds without a ND filter (full power of lamp). The photobleached area was sufficiently large for observing a boundary without being affected by the other boundaries (the observed boundary was perpendicular to the line pattern). After photobleaching, the changes in the fluorescence profiles at the boundary region between the bleached and unbleached areas were observed. The collected boundary profiles were fitted to a Gaussian error function by using the Origin program (OriginLab Corporation, Northampton, MA) for determining the diffusion depth w which is defined as

2

F ( x, t ) − Fbleached  x − xb  = erf   +1 Funbleached − Fbleached  2w 

w = Dt

where D is the diffusion coefficient and t is the time after photobleaching. From the obtained w values, the w2 values were plotted versus the time t. The diffusion coefficient D of fluorescent molecules was determined from the slope of linear dependency.

2.7 Reconstitution of bacteriorhodopsin into patterned bilayer A seven-transmembrane alpha-helical protein from Haloquadratum walsbyi named HwBR was heterologously expressed in Escherichia coli BL21 (DE3) cells as reported previously.

34

The 41st valine residue of HwBR on the cytoplasmic B-C (2nd-3rd) loop

was mutated to cysteine using the Quickchange site-directed mutagenesis method for attaching a fluorophore (Supporting Information, Figure S1). During the DNA


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manipulation, six histidines (His-tag) were attached to the C-terminus of HwBR for purification. The constructed plasmid was analyzed using an automated sequencer (ABI 3100) to confirm the expected nucleotide sequences. Preparation of crude membranes and purification of protein were done as described previously. 35 Briefly, the sample was solubilized with DDM and purified with a Ni affinity column (HisTrap, GE Healthcare), an anion exchange column (HiTrap-Q, GE Healthcare), and a gel-filtration column (Superdex-200, Amersham Biosciences). Because the absorption spectrum of V41C was almost identical to that of the Wild-type HwBR, we concluded that the mutation did not show significant effects on the structure of HwBR (Supporting Information, Figure S1). The purity of the protein was estimated as 90 % from the molar extinction coefficient of 50,000 M-1 cm-1 at 550 nm (ε). Then the purified HwBR was labeled with HL750. HwBR in buffer A (50 mM Tris, 150 mM NaCl, and 0.98 mM DDM, pH=7.0) containing TCEP was mixed with HL750 dissolved in dimethyl sulfoxide (HwBR/ HL750/ TCEP = 1:9:8). The reaction proceeded for 90 min at 25°C, and was stopped by adding 7.2 µmole of 2-mercaptoethanol. After ultrafiltration with Amicon Ultra 30 k (14,000 g, 15 min at 4°C), unreacted HL750 was removed by the gel permeation chromatography using a superose 6 column at room temperature. Collected HL750-HwBR was concentrated and washed with buffer B (50 mM Tris, 1M NaCl, 60 mM OG, pH=7.0) three times by ultrafiltration. The molar ratio of dye/HwBR was estimated to be ca. 0.3 using ε = 50,000 M-1 cm-1 at 550 nm for HwBR and 280,000 M-1 cm-1 at 750 nm for HL750. HL750-HwBR was stored at -30 °C as a 50% glycerol stock solution. HL750-HwBR was reconstituted into a preformed SPB by the rapid dilution of detergent-solubilized molecules. A polydimethylsiloxane (PDMS) chamber was used for


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the reconstitution and microscopic observation. HL750-HwBR was diluted in buffer C (50 mM Tris, 150 mM NaCl, 20 mM OG, pH=7.0) to the final concentration of 760 nM. 2 µL of the diluted solution was quickly mixed with 160 µL of buffer C in the chamber, while vigorously stirring the solution with a magnet. After incubation for 30 sec, excess HL750-HwBR molecules were rinsed 5 times with buffer C. Reconstituted HL750-HwBR was observed by the total internal reflection fluorescence microscopy (TE2000-V; Nikon, Tokyo, Japan) equipped with a near-infrared laser diode (SL750nm 100T; Shanghai Sauk Kaser, Shanghai, China). Images were acquired by an electron-multiplying charge-coupled device camera (C9100-12; Hamamatsu Photonics, Hamamatsu, Japan) at 33 frames s-1 frame rate and 76 nm pixel-1 spatial resolution. To minimize photobleaching and blinking during the observation, 2.5 µg/mL glucose oxidase, 2.5 µg/mL catalase, 70 µg/mL glucose and 10mM COT in DMSO were added. 36


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3. Results and discussion 3.1 Poly(MPC) brush formation on polymeric bilayer Poly(MPC) brush was formed from the surface of polymeric bilayer by ARGET-ATRP. Figure 2 shows the film thickness (dried) after varied ARGET-ATRP reaction time at two MPC concentrations (0.3M and 0.67M). The initial film thickness of ca. 50 Å corresponded to the polymeric bilayer. The total film thickness, including polymeric bilayer and poly(MPC) brush, grew with time and reached a plateau, presumably due to crowding or radical termination at the terminal residues of the brush.

37,38

The results

suggest that the average molecular weight of polymer chains can be controlled by the monomer concentration and the reaction time. In the following studies, MPC concentration was fixed at 0.67 M.

We could control poly(MPC) brush formation by the lipid compositions of polymeric bilayer (DiynePC/DiynePE). For example, symmetric and asymmetric brushes could be prepared from the combination of monolayers composed of DiynePC/DiynePE mixture (4:1) and pure DiynePC. We prepared four types of bilayers by the successive deposition of monolayers (top/ bottom monolayers: PC/PC, PC/PC-PE, PC-PE/PC, PC-PE/PC-PE). The average film thickness before and after ARGET-ATRP was compared (Figure 3). We obtained the largest thickness increase for PC-PE/PC-PE, which could form poly(MPC) brushes on both sides of the bilayer. The increase was minimum for PC/ PC, as expected. The slight increase in the film thickness was either due to the adsorption of poly(MPC) from the aqueous phase or partial reactioin of NHS-BMP to DiynePC. Although the initiator was present only on the surface of polymeric bilayers, we observed that polymerization proceeded in the bulk solution as


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well by the radical transfer from the substrate surface (data not shown). The thickness increase was intermediate for the asymmetric bilayers (PC-PE/PC and PC/PC-PE), in which DiynePE was present only in one monolayer. Bilayers with DiynePE in the top monolayer facing the aqueous phase (PC-PE/PC) had a slightly larger increase than those with DiynePE in the bottom monolayer facing the substrate (PC/PC-PE). This difference may stem from the fact that MPC has to diffuse through the bilayer in the case of PC/PC-PE. We also studied the brush formation with varied DiynePC/DiynePE compositions in the polymeric bilayer (PC-PE/PC-PE). The average film thickness after the ARGET-ATRP reaction increased linearly with the DiynePE composition up to 20% (it was not possible to prepare a stable polymeric bilayer with a higher DiynePE content due to the unfavorable molecular packing of PE for forming a bilayer structure), clearly demonstrating that poly(MPC) formed from DiynePE molecules in the membrane (Supporting Information, Figure S2). Furthermore, the amount of poly(MPC) formed was proportional to the average thickness (surface coverage) of the polymeric bilayer before the ARGET-ATRP reaction (Figure S3). These results corroborated the premise that poly(MPC) brushes uniformly formed from the surface of polymeric bilayer domains.

3.2 Poly(MPC) brush formation in the presence of fluid bilayer ARGET-ATRP in an aqueous solution enables to generate poly(MPC) brush from a patterned membrane containing polymeric and fluid bilayers (Figure 4). The fluid bilayers (DOPC/ TR-PE (1mol%)) were introduced into the corrals between polymeric bilayers after brominating the polymeric bilayers. We then initiated the ARGET-ATRP reaction by introducing monomer (MPC), catalyst (CuBr), and reducing agent (AA).


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Figure S4 (Supporting Information) compiles the film thickness increase on four types of polymeric bilayers (PC/PC, PC/PC-PE, PC-PE/PC and PC-PE/PC-PE (top/ bottom monolayer)). The largest thickness increase was obtained for PC-PE/PC-PE, and the increase was minimum for PC/ PC. (It should be noted that the average thickness was smaller than the bilayers in Figure 2, because the patterned membranes contained ca. 75% polymeric and ca. 25% fluid bilayers). The thickness increase was intermediate for the asymmetric bilayers (PC-PE/PC and PC/PC-PE). Again, the thickness increase was larger for bilayers having DiynePE in the top monolayer facing the aqueous phase (PC-PE/PC) than those with DiynePE in the bottom monolayer, presumably due to the fact that diffusion of MPC through the bilayer is limited in the case of PC/PC-PE. The influences of fluid bilayers on the brush formation were assessed by comparing the relative film thickness before and after ARGET-ATRP for fully polymeric bilayers and patterned bilayers (Supporting Information, Figure S4 (B)). The relative film thickness was obtained by dividing the thickness after ARGET-ATRP (polymeric bilayer plus brush) with that of polymeric bilayer before ARGET-ATRP. Since the relative film thickness of fully polymeric bilayers and patterned bilayers was mostly the same, we conclude that the fluid bilayers did not affect the brush formation on polymeric bilayers.

The fluidity of embedded bilayers after the ARGET-ATRP reaction was assessed by FRAP. As we photo-bleached the fluid bilayer, the boundary between the bleached and non-bleached regions became unclear with time, suggesting that fluidity of the bilayer was retained after the ARGET-ATRP reaction (Figure 4(B)). The diffusion coefficients of the fluid bilayers as determined by FRAP-BPE did not change significantly before and after ARGET-ATRP in the two types of polymeric bilayers PC-PE/PC-PE and PC/PC


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(Supporting Information, Figure S5). These results clearly demonstrate that the bilayer structure and its physicochemical property (fluidity) were retained during the brush formation. If fluid bilayers were introduced into the corrals between polymeric bilayers after ARGET-ATRP, non-raptured vesicles were absorbed on the substrate surface and formation of planar bilayers was hindered, presumably due to the presence of adsorbed poly(MPC) (data not shown). Therefore, in-situ formation of polymer brush in the presence of embedded fluid membranes is important for generating polymer brush-supported model membranes.

3.3 Separation of the membrane from the substrate An important objective of the polymer brush was to separate the lipid membrane from the substrate surface. In order to assess the separation between the polymeric bilayer and the substrate surface, we measured the quenching of fluorescence by the Förster resonance energy transfer (FRET) between the bilayer and silicon substrate. The fluorescence emission from polymeric and fluid bilayers should be quenched by the energy transfer to metallic silicon in a distance dependent manner. 39,40 Figure 5 compares the fluorescence from polymeric bilayer and fluid bilayer (DOPC/ TR-PE) before and after the brush formation. The polymer brush formed on both sides of polymeric bilayer (PC-PE/PC-PE). The fluorescence from polymeric bilayer, which arises from the electronically conjugated ene-yne backbone,

41

was mostly quenched in the case of a bilayer directly adsorbed on

the silicon substrate, since the distance between the membrane and silicon was expected to be less than 10 nm (Figure 5(A) top-left). On the other hand, the fluorescence was significantly enhanced after the brush formation (Figure 5(A) bottom-left and 5(B) left), clearly indicating that the distance between the polymeric bilayer and substrate increased


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due to the poly(MPC) layer. Fluid bilayers also showed enhanced fluorescence in the polymerized region (Figure 5(A), (B) right). The fluorescence should come either from planar bilayers embedded in the small areas between polymeric bilayer domains or from vesicles adsorbed on the polymeric bilayer surfaces. The latter possibility would be less likely, since we generally observed detachment of vesicles from the surface by the brush formation (data not shown). It is plausible that fluid bilayers penetrated into the space between polymeric bilayer domains upon brush formation. The regions near the rim of corrals often had brighter fluorescence (indicated with an arrow in Figure 5(A) bottom-right). This observation also supports the premise that fluorescence came from embedded planar bilayers. On the other hand, fluorescence from the fluid bilayers in corrals did not increase after the brush formation (Figure 5), suggesting that the distance between the bilayer and the substrate did not significantly increase. This may be explained by the fact that the size of fluid bilayer (20 µm) was much larger than the thickness of polymer brush (tens of nanometers). As a control experiment, we observed fluorescence of polymeric and fluid bilayers on a glass substrate (Supporting Information, Figure S6). The fluorescence intensities decreased after the brush formation both for polymeric and fluid bilayers, presumably due to detachment of vesicles. These results also corroborated the conclusion that the fluorescence enhancement observed on silicon substrates was due to the increased distance between membrane and substrate.

3.4 Reconstitution of Haloquadratum walsbyi bacteriorhodopsin (HwBR) For assessing the utility of polymer brush-supported model membrane, we reconstituted the V41C mutant of HwBR (Supporting Information, Figure S1) into the patterned bilayers and observed the lateral diffusion of single molecules (Figure 6). The


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trajectories of HwBR molecules were confined in the corrals in the case of patterned membranes with fully polymerized bilayer (Figure 6 left). In the absence of polymer brushes, most of the HwBR molecules observed were immobile (shown with red color in Figure 6), suggesting that they were not integrated into the bilayer membrane. In the presence of polymer brushes, the number of immobile HwBR molecules drastically decreased and most HwBR molecules were observed to diffuse laterally in the fluid bilayer regions. (We estimated the mobile fractions to be less than 5% without brush and more than 50% with brush, although it was difficult to quantify the fractions of immobile molecules due to their aggregation.) The results suggest that poly(MPC) brush suppressed non-specific adsorption of HwBR onto the polymeric bilayer. The density and diffusivity of mobile HwBR were comparable in the membranes with and without polymer brushes. The number of reconstituted HwBR molecules was rather restricted by the reconstitution conditions, in which HwBR molecules were rapidly diluted in the chamber and mostly incorporated into the coexisting vesicles.

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Due to

the low number of reconstituted HwBR molecules, it was not possible to statistically analyze their diffusion coefficients. Successfully reconstituted HwBR molecules had rather rapid lateral diffusion, suggesting they are quite mobile owing to their compact structures. Quantitative evaluation of the diffusivity would be a subject for future studies, since the diffusivity significantly changes depending on the reconstitution procedure and the state of lipid bilayer membrane. We reconstituted HwBR also into a patterned composite membrane, in which polymeric and fluid lipid bilayers coexisted as nanometer-sized domains.

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The presence of polymer brush drastically reduced

non-specific adsorption of proteins, while retaining incorporated HwBR molecules mobile (Figure 6 right). HwBR molecules diffused both in the circular corrals of fluid


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bilayers and composite membrane regions. This result demonstrates that the composite membranes remained continuous and fluid after the brush formation.

3.5 Functionalization of polymer brush Functionalization of polymer brush would enable to attach various objects to the membrane, including biological molecules, cells, and microstructures, extending the two-dimensional membrane system into a three-dimensional architecture. In order to attach functional molecules onto polymer brush, we constructed block copolymers of poly(MPC) and poly-(2-aminoethyl methacrylate) (poly(AMA)) by successively polymerizing MPC and AMA by ARGET-ATRP (Figure 7).

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Poly(AMA) has a

primary amine moiety in the side chain, to which other molecules can be covalently bonded. Figure 7 shows the average film thickness of polymeric bilayer (designated as Bilayer), bilayer with poly(MPC) brush (MPC-Bilayer), and bilayer with block copolymer brush of poly(MPC) and poly(AMA) (ARGET-ATRP for 3 hours for MPC and AMA each: AMA-MPC-Bilayer). Although the thickness growth of poly(AMA) was rather small, the amine moiety in poly(AMA) brush could be functionalized with biotin and fluorescently labeled streptavidin (SAF-594) was bound (Figure 8). Fluorescence was negligible both on polymeric bilayer (Bilayer) and bilayer with poly(MPC) brush (MPC-Bilayer), whereas we observed fluorescence of SAF594 on AMA-MPC-Bilayer, clearly demonstrating the presence and functionality of grafted poly(AMA). For Bilayer and MPC-Bilayer, the fluorescence intensity of SAF594 was lower on silicon compared with a glass substrate due to the energy transfer to the substrate (Figure 8(B)). On the other hand, the fluorescence intensities were comparable for glass and silicon substrates in the case of AMA-MPC-Bilayer. Elimination of FRET


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suggests that the poly(AMA) segment of polymer brushes was separated from the substrate.


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4. Conclusions We developed a hybrid model membrane comprising patterned bilayer and polymer brush. The polymer brush was formed on the polymeric bilayer by the surface initiated ARGET-ATRP in the presence of embedded natural lipid membranes. Generation of polymer brush effectively enlarged the distance between the polymeric bilayer and the substrate, as evidenced by the fluorescence enhancement on silicon substrates. The presence of polymer brush significantly reduced nonspecific adsorption of model membrane protein (HwBR) onto the patterned bilayer. The heightened fraction of mobile protein molecules should be advantageous for their functional assays. The density and diffusivity of reconstituted HwBR were comparable for SPBs with and without polymer brush, presumably due to the reconstitution conditions and the compact structure of HwBR molecules. Detachment of fluid bilayer from the substrate should be more critical for more bulky membrane proteins. Incorporation of a wider variety of membrane proteins would be an important future objective. To this end, it is important to analyze the structures of lipid bilayer-polymer brush hybrids in aqueous solution by interfacial techniques such as X-ray reflectivity measurements for elucidating the detailed structural features such as chain density/ length and the mushroom-brush transition. For realizing a model membrane that mimics the structural motif of cytoskeleton-supported cell membrane, functionalization of the polymer brush with a reactive side chain (AMA) should also play an important role. Such model membranes with controlled two-dimensional and three-dimensional structures would offer an ideal platform for incorporating membrane proteins and measuring their functions in a truly biomimetic environment.


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Supporting Information Available. Additional information about the site-directed mutagenesis of HwBR for attaching a fluorophore, formation of poly(MPC) brush, diffusion coefficients of fluid bilayers before and after ARGET-ATRP determined by FRAP-BPE, fluorescence micrographs of patterned bilayers on glass substrate. This material is available free of charge via the Internet at http://pubs.acs.org.

Acknowledgements. This work was supported by Grant-in-Aid for Scientific research from Japan Society for the Promotion of Science (#23106714, #15K14489), Nippon Sheet Glass Foundation, and Tokyo Ohka Foundation for the Promotion of Science and Technology. We thank Dr. Minoru Mizuhata and Dr. Itsuko Ayabe (Kobe University) for allowing us to use the ellipsometer. Discussion with Dr. Shinichi Yusa (University of Hyogo) is also appreciated.


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Figures:

Figure 1: (A) Schematic of a polymer-brush-supported model membrane constructed by the surface-initiated polymerization. (B) Chemical structures of polymerizable lipids. (C) Monomer structures and the reaction scheme of ARGET-ATRP.


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Figure 2: Formation of poly(MPC) brush on the surface of polymeric bilayer. The average film thickness, including the bilayer and polymer brush, was plotted as a function of the reaction time of ARGET-ATRP. Two MPC concentrations were used (red open circles: 0.3M, blue closed circles: 0.67M).


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Figure 3: Formation of poly(MPC) brushes on four types of polymeric bilayers that contained monolayers of DiynePC (designated as PC) and/ or DiynePC/ DiynePE (80:20) (designated as PC-PE). The symmetric and asymmetric bilayers were generated by combining the monolayers via successive LB and LS depositions. The average film thickness was compared before and after the ARGET-ATRP.


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Figure 4: Polymer brush formation from a patterned membrane composed of polymeric and fluid bilayers: (A) Schematic illustration. (B) Fluorescence micrographs of embedded fluid bilayers (DOPC/TR-PE (1mol%)). The fluid bilayers retained the structural integrity and fluidity after the brush formation as assessed by the FRAP measurements. The fluorescence intensity profiles along the dashed white lines were compared before and after the brush formation.


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Figure 5: Fluorescence micrographs of a patterned bilayer on silicone substrate before and after the brush formation (A). The left and right columns are fluorescence images of polymeric bilayer (green) and fluid bilayer (DOPC/TR-PE (red)), respectively. Before the brush formation, fluorescence of adsorbed bilayers was quenched due to the Förster energy transfer (FRET). After the brush formation, fluorescence in the polymeric bilayer region was enhanced, suggesting the separation of the bilayer from the substrate surface. The arrow indicates the boundary of polymeric regions where brighter fluorescence of fluid bilayer was observed. The size of corrals was 20 µm. (B) Fluorescence intensity profiles of polymeric and fluid bilayers along the dashed lines in (A), before (black) and after (red) the brush formation.


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Figure 6: Single molecule fluorescence microscopy observation of reconstituted HL750-HwBR in patterned bilayers with and without polymer brushes. The polymeric bilayer comprised DiynePC and DiynePE (4:1) in both monolayers (PC-PE/PC-PE). The green, red, and magenta colors in the micrographs represent fluid bilayer (DOPC/ TR-PE), immobile HL750-HwBR, and trajectories of mobile HL750-HwBR, respectively. The micrographs on the right side are observations in patterned membranes composed of fluid bilayers and composite bilayers (nanometer-sized mixture of polymeric and fluid bilayer domains). The size of each image is 13.1 µm.


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Bilayer

MPC-Bilayer

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AMA-MPCBilayer

Figure 7: Block copolymer of poly(MPC) and poly(AMA) was formed by the successive polymerization of MPC and AMA (3h each). DiynePE was present only in the upper monolayer. The average film thickness was compared for brominated polymeric bilayer (Bilayer), bilayer with MPC brush (MPC-Bilayer), and bilayer with AMA-MPC block copolymer brush (AMA-MPC-Bilayer).


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Figure 8: Surface functionalization of AMA-MPC brush: Biotin was covalently bound to the amine moiety of AMA-MPC brush, and fluorescently labelled streptavidin (SAF594) was adsorbed to biotin. (A) Fluorescence micrographs of bilayers/ brushes on glass and silicon. Streptavidin bound only onto the patterned bilayers with AMA-MPC brush. The size of corrals was 20 µm. (B) Fluorescence intensities of SAF594 for the samples in (A). The red and blue bars are the intensities of glass and silicon substrates, respectively.


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TOC Graphic :

ATRP from polymeric bilayer

Polymeric bilayer

Polymer brushsupported bilayer


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Hybrid Model Membrane Combining ... - ACS Publications

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