Graphene-Templated Supported Lipid Bilayer Nanochannels - Nano

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Graphene-templated supported lipid bilayer nanochannels Wan Li, Jean K Chung, Young Kwang Lee, and Jay T Groves Nano Lett., Just Accepted Manuscript • DOI: 10.1021/acs.nanolett.6b01798 • Publication Date (Web): 30 Jun 2016 Downloaded from http://pubs.acs.org on July 3, 2016

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Graphene-templated supported lipid bilayer nanochannels Wan Li, Jean K. Chung, Young Kwang Lee, and Jay T. Groves* Department of Chemistry, University of California, Berkeley, California 94720, United States *Corresponding author: [email protected]

ABSTRACT. The use of patterned substrates to impose geometrical restriction on the lateral mobility of molecules in supported lipid membranes has found widespread utility in studies of cell membranes. Here, we template-pattern supported lipid membranes with nano-patterned graphene. We utilize focused ion beam milling to pattern graphene on its growth substrate, then transfer the patterned graphene to fresh glass substrates for subsequent supported membrane formation. We observe that graphene functions as an excellent lateral diffusion barrier for supported lipid bilayers. Additionally, the observed diffusion dynamics of lipids in nanoscale graphene channels reveal extremely low boundary effects, a common problem with other materials. We suggest this is attributable to the ultimate thinness of graphene.

KEYWORDS. Supported lipid bilayer, graphene, diffusion barrier, nanochannels

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MAINTEXT As one of the most important naturally-occurring self-assembled structures, the phospholipid bilayer defines the biological cell membrane, and hosts much of the machinery for cellular communication and transport.1 Supported lipid bilayers2-7 provide a highly controllable in vitro system for modeling processes at the cell surface, including cell signaling,8-10 ligand– receptor interactions,11 and enzymatic reactions.12, 13 A powerful feature of supported membranes is that patterns on the underlying substrate can be used to impose geometrical restrictions on the lateral mobility of molecules in the membrane. Partitioning and controlling the lateral diffusion of supported lipid bilayers14-16 enables the spatial manipulation of reconstructed biological systems without changing the key fluidic nature of lipid membranes, and has proven to be a valuable tool in elucidating the spatial and mechanical aspects of biological processes.17-21 While local scratching/blotting can be used to pattern a supported lipid bilayer,22-24 the resultant structures are often unstable and sensitive to the environment.25,

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Physical barriers, e.g., patterned metal, metal oxides, polymers, and

proteins, deposited onto the underlying substrate prior to assembly of the bilayer, have been highly successful in confining the bilayer into stable, micrometer-scale patterns through local depletion and/or selective immobilization, depending strongly on the vesicle-substrate interactions.27-34 Despite of the varieties of barrier materials, the patterned barriers are often taller in height than the lipid bilayer itself (~4 nm), and patterning at the nanoscale remains challenging. We report the use of graphene as an atomically thin barrier to partition and confine the lateral diffusion of supported lipid bilayers on glass surfaces into nanoscale patterns. Graphene


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possesses outstanding electrical, optical, and mechanical properties.35,

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indicated that graphene has high surface energy and completely alters the wettability of the substrate.37, 38 Since supported bilayer assembly is driven by surface properties,25 we anticipated that such perturbations would prevent fluidic supported membrane formation and thus allow graphene to function as a barrier material.

Figure 1. Fabrication of nano-patterned graphene devices that are compatible with both the formation of supported lipid bilayers and fluorescence microscopy by a pattern transfer method. (a) Schematic of the fabrication process flow. (b) SEM image of the generated graphene pattern on copper substrate [as in Step 2 in (a)]. (c) Optical microscopy image of the completed device on coverglass. Scale bars: 5 μm.


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To achieve high definition of the patterned graphene nanostructures while maintaining a clean surface, which is critical for the preparation of supported lipid bilayers, we developed a pattern transfer fabrication method in which graphene was first patterned on its growth substrate and then transferred to a freshly prepared coverglass surface. Briefly, the process begins with chemical vapor deposited (CVD) monolayer of graphene grown on copper substrates (Fig. 1a). 39 Next, a focused ion beam was used to pattern graphene on copper substrates (Fig. 1a, Step 2) by milling down ~10 nm into the material. Complete removal of graphene was evident from scanning electron microscope (SEM) images (Fig. 1b), where visible structures in graphene (e.g., wrinkles) are observed to be completely removed from ablated regions. The patterned graphene was then transferred to a freshly cleaned, highly hydrophilic coverglass substrate following standard PMMA-based transfer methods (Fig. 1a Steps 3-6, and Fig. 1c).40 Raman spectroscopy indicated that devices were of high quality, monolayer graphene (Fig. S1).41 Besides ensuring that the patterned graphene is directly deposited onto a fresh coverglass surface without the need of further fabrication steps, our design also circumvents e-beam lithography, the common method to pattern graphene at sub-micrometer resolution.42 Due to surface charging, e-beam lithography is difficult to apply to the fully insulating coverglass substrates, which are necessary for fluorescence microscopy studies of the lipid. Supported lipid bilayers were immediately prepared on graphene devices via vesicle fusion from a solution of small unilamellar vesicles (SUVs).43, 44 The supported lipid bilayer was comprised of 1,2-Dioleoyl-sn-glycero-3-phosphocholine (DOPC) supplemented with 5.5 % 1,2dioleoyl-sn-glycero-3-phospho-L-serine (DOPS). 0.5% Texas Red 1,2-dihexadecanoyl-snglycero-3-phosphoethanolamine (TR-DHPE) was added for fluorescence detection. The device was maintained in Tris-buffered saline (TBS) during the lipid diffusion experiments below.


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Figure 2. Graphene functions as a barrier material for the lateral diffusion of supported lipid bilayer. (a) Fluorescence image of TR-DHPE doped supported lipid bilayer deposited on a device with patterned graphene grids. (b) The field of illumination defined by microscope field-stop aperture for photobleaching. (c) Intensity along the line indicated by blue arrows in (b). (d) and (e) FRAP images at 40 seconds and 6 min 10s of recovery. (f) Intensity along the line indicated by blue arrows in (d). (g) Time dependence of normalized fluorescence intensity in the three boxed regions in (b). The colors of the curves correspond to those of the boxes. Inset: schematic diagram of a supported lipid bilayer partitioned by the patterned graphene grids. The underlining mechanism could be either local depletion or selective immobilization of supported lipid membrane on the graphene surface.27 The former scenario was shown for the schematic diagram. Scale bar in (b): 5 μm. Figure 2a shows the fluorescence image of a device in which 7 µm-sized squares, arranged in a 3x3 array, were removed from the graphene sheet. Fluorescence signal indicates that lipid membranes are formed on the coverglass surface where the graphene is locally


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removed. Uniform fluorescence intensity is observed for these “corral” regions, indicating good quality of the lipid membrane. Meanwhile, the remaining graphene grids correspond to minimal fluorescence, with the detected counts being comparable to the background signal as seen outside the illumination aperture (e.g., Fig. 2b, c). A small amount of immobile bright spots were occasionally observed on the graphene surface, possibly due to the attachment of residual vesicles to local defect sites. Defects, including atomic scale structural defects45 generated in the growth processes and nanoscale cracks and wrinkles generated in the transfer process,46, 47 are common in CVD graphene and may alter local surface properties.48 In comparison, we did not observe such bright spots on pristine, exfoliated graphene sheets (Fig. S2), which are virtually defect-free. These results indicate that monolayer graphene can efficiently prevent the local formation of supported lipid bilayers. A recent quartz crystal microbalance with dissipation (QCM-D) study also indicates that no homogeneous lipid bilayers are formed on graphene surfaces with SUV rupture.49 Fluorescence recovery after photobleaching (FRAP) experiments50 were next conducted to examine whether the patterned graphene grids are also capable of confining the lateral diffusion of the partitioned lipid bilayers. Figure 2b shows the field of illumination for photobleaching defined by the field-stop aperture of a microscope, which included the entire center corral and parts of the other corrals. After continuous illumination for ~1 min, the central corral was entirely bleached, whereas fluorescence intensity gradients were observed in the partly illuminated corrals, indicative of lipid exchange between the bleached and non-bleached regions of the corrals (Fig. 2d, f). The fluidic nature of lipid bilayer was also reflected by the gradually recovering fluorescence intensity (Fig. 2g) of the bleached regions in these corrals (magenta and red boxes in Fig. 2b). Meanwhile, no appreciable recovery of fluorescence was


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observed for the center coral, which was isolated by the graphene grid from non-bleached lipid bilayers (Fig. 2e, g). These results indicate that graphene, despite being only one atom thick, is an effective barrier to the lateral diffusion of fluidic lipid bilayer (Fig. 2g Inset).

Figure 3. Patterned graphene devices for quantitative study of the diffusion of supported lipid bilayer under nanoscale lateral confinement. (a) Fluorescence image of two closely placed graphene devices, each patterned with two reservoirs connected with a thin channel, after the deposition of supported lipid bilayer. (b-e) FRAP measurements of the two devices at 0.5 min, 1.5 min, 8 min and 20 min of recovery. The octagon in (b) indicates the field of illumination for bleaching. (f) Time dependence of the normalized fluorescence intensity in different reservoirs color-labeled in (a).


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To further understand lipid-graphene interactions and explore whether graphene patterning can be employed to quantitatively study the diffusion of lipid membranes, we prepared graphene devices that contained two lipid reservoirs connected by thin channels of different widths. Figure 3a shows the fluorescence image of two closely placed channel devices (I-II; III-IV) and one non-connected corral (V) after deposition of lipid bilayer. The channel widths (w) of the two devices were 114 nm and 770 nm, respectively. As the thinner channels in our study had widths smaller than the resolution of conventional optical microscopy (~300 nm), the channel widths were determined indirectly based on the measured fluorescence intensity by w = wm (Ichannel / Ireservoir), where wm is the apparent width of the channel, Ichannel is the average fluorescence intensity over the apparent width, and Ireservoir is the bulk fluorescence intensity measured at the reservoirs. FRAP was employed to study the diffusion kinetics of lipid bilayers in the channel devices. Reservoirs II and III, as well as the non-connected Corral V, were first photobleached for 1 min 10 s. Figure 3b-f shows the fluorescence images of the devices at 0.5 min, 1.5 min, 8 min, and 20 min after photobleaching, as well as the time dependence of the fluorescence intensity in each reservoir. No fluorescence recovery was observed for the fully isolated Corral V. In contrast, for the two channel devices, gradual recovery of fluorescence signal in the bleached reservoirs (II, III) was accompanied by decreases in fluorescence signal in the connected unbleached reservoirs (I, IV). The sum amount of the fluorescence signal in each device remained almost constant, except for slow decreases due to photobleaching. These results indicate that the fluorescence recovery was entirely due to diffusion of lipid through the channel. Noticeably different recovery dynamics were observed for channels of different widths, with slower recovery being observed in thinner channels.


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Figure 4. Agreement between experimental and theoretical results indicates that the graphene barrier interferes minimally with the dynamics of the confined bilayer. (a) The fluorescence recovery time constant as a function of the channel width obtained from experiments, Monte Carlo simulation, and simplified 1D diffusion theory. Inset: The diffusion coefficient of supported lipid bilayers measured from 20 devices with channel width ranging from 28 nm to 1070 nm. (b) Representative molecule diffusion trajectories generated in Monte Carlo simulation. The presented device has a channel width of 200 nm. (c) Representative FRAP recovery curves obtained from Monte Carlo simulations.


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To quantify our results, we performed a series of FRAP experiments on 20 graphene devices of varying channel widths. For each dataset, the time constant of fluorescence recovery was determined by fitting the measured recovery time traces with a single exponential (Fig. 3f and Fig. S3). The channel width dependence of the resultant time constants was then compared with theory (Fig. 4a). A simplified model based on macroscopic one-dimensional diffusion, as well as exact, Monte Carlo simulations based on the actual geometry of the devices were both implemented (Supporting information). For the latter, 3,000 particles were initially evenly distributed in one of the two reservoirs, and then each particle was allowed to evolve independently following two-dimensional random walks and elastic collisions at the device boundaries (Fig. 4b). The simulated fluorescence intensity over time, measured as the number of particles in each reservoir, successfully recapitulated our experimental data (Fig. 4c). Fluorescence-recovery time constants were hence extracted from the simulated data to compare with experiment (Fig. 4a). For this comparison, the averaged time constant of the 1 µm-wide devices was used to determine the diffusion coefficient of the lipid, which was used as the only free parameter in plotting the theory results. Good agreement was found between the experimental and theoretical results. In particular, the experimental time constants of the 20 devices fell closely onto the Monte Carlo simulation curve. We thus further calculated the diffusion coefficients of all devices by comparing the actual experimental time constant values to the simulation. Similar diffusion coefficient values (0.5-1.8 µm2/s; Fig 4a inset) were found in all devices with channel widths wider than 50 nm, in agreement with previous results.51,

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These results indicate that the

patterned graphene, while confining the lateral diffusion of supported lipid bilayers, interferes minimally with the dynamics of the confined bilayer. The device with the thinnest channel width


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of 28 nm, however, showed a reduced diffusion coefficient of 0.3 µm2/s. A recent study on the use of dip-pen lithography to produce lipid stripes on mica also noted similar reduction in diffusion coefficient for widths smaller than 50 nm, which may be intrinsic to the lipid due to a tilted phase at the edge of lipid stripes.51 By employing monolayer graphene as a one-atom thick barrier, we have achieved excellent confinement for the lateral diffusion of lipid bilayers. Nanoscale diffusion channels were thus fabricated out of graphene, and the observed diffusion dynamics of lipids was in good agreement with theory with minimal edge effects. These results may be attributed to the fact that graphene is only one atom in thickness: there is zero geometrical variation (roughness) across the height, and the confined lipid bilayers only interact minimally with the graphene edges at the surface. ACKNOWLEDGMENT We thank Prof. Ke Xu for helpful discussions. This work was partly supported by the College of Chemistry at UC-Berkeley. ASSOCIATED CONTENT Supporting Information Detailed experimental processes, theoretical models, computational simulations, and additional experimental data are included in the Supporting Information.


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