Fabrication of multicomponent, spatially segregated DNA and protein

Jul 23, 2018 - Incubation of microbeads coated with supported membrane resulted in an exchange of lipid contents with planar, membrane corrals present...
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Article Cite This: Langmuir 2018, 34, 9781−9788

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Fabrication of Multicomponent, Spatially Segregated DNA and Protein-Functionalized Supported Membrane Microarray Kabir H. Biswas,*,†,‡ Nam-Joon Cho,‡,§ and Jay T. Groves*,‡,∥ †

Mechanobiology Institute, National University of Singapore, Singapore 117411, Singapore School of Materials Science and Engineering, Nanyang Technological University, Singapore 639798, Singapore § School of Chemical and Biomedical Engineering, Nanyang Technological University, 62 Nanyang Drive, Singapore 637459, Singapore ∥ Department of Chemistry, University of California, Berkeley, California 94720, United States Langmuir 2018.34:9781-9788. Downloaded from pubs.acs.org by UNIV OF SUNDERLAND on 09/05/18. For personal use only.

‡

ABSTRACT: Deoxyribonucleic acid (DNA) has been used as a material for a variety of applications, including surface functionalization for cell biological or in vitro reconstitution studies. Use of DNA-based surface functionalization eliminates limitations of multiplexing posed by traditionally used methods in applications requiring spatially segregated surface functionalization. Recently, we have reported a stochastic, membrane fusion-based strategy to fabricate multicomponent membrane array substrates displaying spatially segregated protein ligands using biotin−streptavidin and Ni-NTA− polyhistidine interactions. Here, we report the delivery of DNA oligonucleotide-conjugated lipid molecules to membrane corrals, allowing spatially segregated membrane corral functionalization in a membrane microarray. Incubation of microbeads coated with the supported membrane resulted in an exchange of lipid contents with planar membrane corrals present on a micropatterned substrate. Increases in the system temperature and membrane corral size resulted in alterations in the rate constant of lipid exchange, which are in agreement with our previously developed analytical model and further confirm that lipid exchange is a diffusion-based process that takes place after the formation of a long “fusion-stalk” between the two membranes. We take advantage of the physical dimensions of the fusion-stalk with a large aspect ratio to deliver DNA oligonucleotide-conjugated lipid molecules to membrane corrals. We believe that the ability to functionalize membrane corrals with DNA oligonucleotides significantly increases the utility of the stochastic fusion-mediated lipid delivery strategy in the functionalization of biomolecules such as DNA or DNA-conjugated protein ligands.

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INTRODUCTION Deoxyribonucleic acid (DNA) as a material has found diverse applications owing to its physicochemical properties1,2 and amenability to large-scale production.3 For instance, DNAbased assemblies have been devised to create three-dimensional structures with nanoscale features,4−6 machinelike behavior,7−9 or computational gating.10,11 They have also been used for functionalization of synthetic surfaces for applications such as measuring forces exerted by cells on individual adhesion receptors12−15 or assembly of proteins in vitro.16−19 In addition to these, DNA-based protein anchoring has emerged as a strategy of choice to display ligands on synthetic, supported lipid bilayers (SLBs) in a variety of applications,20 including reconstitution of receptor−ligand interactions,21,22 tethering of vesicles,23,24 and assembly of DNA origami structures.25,26 Supported lipid bilayers are a two-dimensional assembly of phospholipids on a solid support, which are often used as surrogates for cellular membranes.27−30 Retention of many of the cell membrane features such as two-dimensional mobility © 2018 American Chemical Society

of lipid molecules and electrical insulation has made them extremely useful in biological and biotechnological applications. These include reconstitution of receptor−ligand interaction in a hybrid format, wherein one or more ligands of interest are displayed on a supported membrane to engage the cognate receptor present on the cell membrane28,31−37 or biochemical reactions involving various membrane or cytosolic proteins.38−43 To increase the utility of a supported lipid bilayer, a number of efforts have been made toward the production of supported membranes displaying multiple, spatially segregated protein ligands by employing a variety of micropatterning techniques.44−46 These include separation of lipid molecules induced by an electrical field,47−49 photochemical degradation of a previously deposited supported lipid bilayer using a photomask, followed by deposition of a second bilayer of a different composition,50,51 by either microcontact Received: April 26, 2018 Revised: June 27, 2018 Published: July 23, 2018 9781

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Figure 1. Lipid exchange between planar- and microbead-supported membranes. (a) Schematic showing the process of lipid exchange between membrane corrals on a micropatterned substrate and microbeads. (b) Schematic representation of the model of lipid exchange between the two membranes. Lipid exchange is proposed to take place through the formation of a narrow membrane tube between the planar- and microbeadsupported membranes. (c) Epi-fluorescence image of an exchanging membrane corral undergoing lipid exchange with a microbead-supported membrane and a nonexchanging membrane corral. The membrane corral contains TR-DHPE fluorescent lipid, whereas the microbead membrane does not contain any fluorescent lipid. Color bars represent fluorescence intensities of the epi-fluorescence images. (d) Graph showing fluorescence intensity of a nonexchanging and an exchanging membrane corral with the progression of lipid exchange. Inset shows the fluorescence intensity of nonexchanging corrals at times 0 and 30 s during the exchange process. Values shown are mean ± standard error of mean (SEM) from multiple individual measurements (N = 17) from a representative experiment.

printing of bilayers using poly(dimethylsiloxane) stamps,52−55 or direct robotic deposition of compositionally different bilayers56 on a previously micropatterned substrate. Additionally, such two-component, micropatterned supported lipid bilayers have been created by pH-controlled scanning probe lifting of the previously deposited bilayer, followed by deposition of a second bilayer with a distinct composition,57 by fluid-flow controlled mixing and surface capture of vesicles,58−60 or by spontaneous lipid exchange between oppositely charged bilayers61 on a previously micropatterned substrate in a microfluidic chamber. In all of these cases, supported lipid bilayers are functionalized with proteins using headgroup-modified lipids, such as 1,2-dioleoyl-sn-glycero-3[(N-(5-amino-1-carboxypentyl)iminodiacetic acid)succinyl] nickel salt (Ni-NTA−DOGS) or 1,2-dipalmitoyl-sn-glycero-3phosphoethanolamine-N-(cap biotinyl) (16:0 biotinyl-CapPE).31−33,62,63 Although they have been utilized successfully in a large number of studies, availability of a limited number of these headgroup-modified lipid molecules imposes a limitation on the number of ligand types that can be independently functionalized in the case of multicomponent, membrane microarray substrates. Use of DNA oligonucleotide, however, alleviates this limitation since virtually any number of protein types could be functionalized on the same substrate by changing the DNA oligonucleotide sequence. DNA oligonu-

cleotides could be functionalized on supported lipid bilayers through either covalent coupling to headgroup-modified lipids, such as 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[4(p-maleimidomethyl)cyclohexane-carboxamide] (PE-MCC) or 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine-N-(6-azidohexanoyl) (azidocaproyl PE), or through insertion of cholesterol-modified oligonucleotide into the bilayer. The bilayer-functionalized single-stranded DNA (ssDNA) oligonucleotide could be hybridized with a complementary functional strand. This strategy has been used in a range of applications, including studying cellular signaling using artificial dimers of epidermal growth factor and ephrinA1,21 in the reconstitution of T-cell immunological synapse with synthetic, DNA-based receptor−ligand pairs,22 synthetic surrogate for membrane fusion protein enabling vesicle fusion,24 or an assembly of origami grids on a two-dimensional surface.26 We have recently reported a straightforward, stochastic membrane fusion-based method to deliver lipid molecules on membrane corrals on a membrane microarray substrate64 that overcame many of the limitations faced by the previously developed technologies. Additionally, we had developed an analytical model to describe the process of lipid delivery by one-dimensional diffusion. Here, we provide further proof for a diffusion-mediated mixing of lipid contents between the microbead and planar membranes. More importantly, we 9782

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Langmuir take advantage of the physical structure of long “fusion-stalks” formed between the exchanging membranes to deliver DNA oligonucleotide-modified lipid molecules, which are significantly larger than the previously delivered lipid molecules. Therefore, the ability to deliver DNA-modified lipid molecules reported here greatly enhances the utility of the multicomponent membrane array fabrication technology.

ij 1 dC 1 yzz w = jjj + z j dx A1 zz{ k A2

Article

−1

−D

×

−Dw jij 1 1 yzz dΔC + jjj zzzdt = l k A2 A1 { ΔC

■

dΔC dt

(4)

Giving us

RESULTS AND DISCUSSION Incubation of microbeads coated with synthetic membranes resulted in the delivery of their lipid contents to planarsupported membrane corrals present on a glass substrate in a stochastic fashion. For instance, membrane corrals of 1,2dioleoyl-sn-glycero-3-phosphocholine (DOPC) lipid doped with Texas Red-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (TR-DHPE) lipid (used for fluorescence detection) present on a micropatterned membrane array substrate showed an exponential decay in their fluorescence intensity (Figure 1a). Although there are other possibilities, the phenomenon could be modeled using Fick’s law of one-dimensional diffusion of lipid molecules through the formation of a narrow fusion-stalk (Figure 1b), as has been done previously64 giving J = −D

dC w dx

ΔCt = ΔC0e

transfer between the two membranes as k =

where C1 and C2 are the concentration of TR-DHPE in the microbead and bilayer disk, respectively; n1 and n2 are the number of TR-DHPE molecules in the microbead and bilayer disk, respectively; and A1 and A2 are the total surface areas of the microbead and the bilayer disk, respectively. Also, n1 + n2 = N, where N is the total number of TR-DHPE molecules in the system and is conserved. The change in the concentration difference of TR-DHPE over time could be written as n yz dΔC d ij n = jjj 2 − 1 zzz j dt dt k A 2 A1 z{

dn dn dΔC 1 1 = × 2 − × 1 dt A2 dt A1 dt

(

(3)

Now, dn2 is the flux of TR-DHPE from the bilayer disk to the dt microbead. Therefore, dn dC w= 2 dx dt

Substituting −D

dn 2 dt

−Dw l

(

1 A2

+

1 A1

).

We note that in the current experimental setup, the fluorescence intensity of nonexchanging corrals remained constant over the period of 30 s, indicating that the decrease in fluorescence intensity seen with exchanging corrals is not due to photobleaching effects (Figure 1c,d). A key realization of the analytical modeling of the lipid exchange kinetics data was that fusion-stalks formed by the exchanging membranes were long tubes with a high aspect ratio. To further confirm the diffusional basis of lipid exchange observed with the membrane corrals, we first varied the temperature of the system and monitored lipid exchange kinetics by measuring the fluorescence intensity of TR-DHPE lipid by epi-fluorescence microscopy (Figure 2a). Temperature of the system fundamentally controls the diffusion of lipid molecules in a fluid membrane,66,67 and an increase in temperature of the system is expected to increase the rate of lipid exchange. This is based on the Einstein relation D = kBT/ λ, where kB is the Boltzmann constant, T is the temperature, and λ is the viscous drag coefficient. This model describes the Brownian motion in a simple viscous fluid, which should be appropriate for lipid membranes over the moderate range of temperatures used here. Indeed, increasing the temperature of the system from 23 to 37 °C resulted in increases in the rate of lipid exchange (Figure 2b), with a concomitant decrease in the half-life t1/2 of relative fluorescence intensity decay (Figure 2c). Further, increases in the temperature of the system also resulted in increases in the lipid exchange rate constant k (Figure 2d), which is in agreement with the analytical model, wherein the rate constant k is directly proportional to the diffusion constant D of lipid molecules in the membrane ÄÅ ÉÑ ÅÅ Ñ ÅÅk = −Dw 1 + 1 ÑÑÑ. Second, we increased the size of the ÅÅÇ l A2 A1 Ñ ÑÖ membrane corrals on the micropatterned, membrane microarray substrate (Figure 3a). A diffusion-mediated exchange of lipid molecules between the two membranes will predict an increase in the half-life t1/2 of the fluorescence intensity due to an increase in the total number of lipid molecules to be exchanged and increase in the effective distance that individual lipid molecules need to diffusively transverse to reach the fusion-stalk before they could be exchanged. To test this, we fabricated membrane microarrays with different membrane corral sizes by the UV-etching method using photomasks containing predetermined sizes of transparent regions, leading to proportionate sizes of UV-etched regions on the PLL-g-PEG polymer substrate.32 Supported membranes on both the

where J is the flux, D is the diffusion coefficient of TR-DHPE molecules, dC/dx is the concentration gradient of the TRDHPE molecules, and w is the width of the channel. Given the dC ΔC homogenous case, dx = l , where l is the length of the channel. n n ΔC = C2 − C1 = 2 − 1 A2 A1 (2)

J = −D

(5)

where Ct is the intensity of TR-DHPE at time t and C0 is the initial TR-DHPE intensity.65 Thus, the fluorescence intensity decay observed in these corrals (Figure 1c,d) could be fitted to the equation, 1 1 ΔCt = ΔC0e−Dw / l( A2 + A1 )t , to determine the rate of lipid

(1)

dΔC 1 zyz dn2 ji 1 = jjj + z× j dt A1 zz{ dt k A2

−Dw / l( 1 + 1 )t A 2 A1

from eq 3

dn dC w= 2 dx dt 9783

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Figure 2. Dependence of the lipid exchange rate on temperature. (a) Schematic showing lipid exchange between a membrane corral and microbead-supported membrane. (b) Graph showing the decay in the fluorescence intensity of the membrane corrals at the indicated temperatures. Note that the rate of fluorescence intensity decay increases with the increase in temperature, indicating a diffusionmediated mixing of the lipid contents. (c, d) Graphs showing t1/2 (c) and k (d) obtained from fitting multiple, individual fluorescence intensity decay curves at different temperatures. Values shown are mean ± SEM from multiple individual measurements (N = 21 for 23 °C, 12 for 27 °C, and 20 for 37 °C) from a single experiment.

Figure 3. Dependence of the lipid exchange rate on membrane corral size. (a) Schematic showing lipid exchange between membrane corrals of different sizes and microbead-supported membranes. (b) Graph showing the decay in the fluorescence intensity of the membrane corrals of the indicated sizes. Note that the rate of fluorescence intensity decay increases with the decrease in corral size, indicating a diffusion-mediated mixing of the lipid contents. (c, d) Graphs showing t1/2 (c) and k (d) obtained from fitting multiple, individual fluorescence intensity decay curves against the reciprocal of different corral sizes. Values shown are mean ± SEM of multiple individual measurements (N = 10 for 4.8 μm2, 4 for 23 μm2, and 5 for 33 μm2) from an individual experiment.

micropatterned substrate and microbeads (5.2 μm in diameter) were deposited as mentioned in the previous section, and final sizes of the membrane corrals were determined from epi-fluorescence images of the substrates. Increases in the membrane corral size lead to slower lipid exchange kinetics between the two membranes (Figure 3b), resulting in increases in the half-life, t1/2 (Figure 3c). Importantly, the rate constant k increased linearly with the membrane corral size (Figure 3d), which is in agreement with the inverse correlation between two parameters, k and A2, in ÄÅ ÉÑ Å −Dw 1 1 Ñ the analytical model ÅÅÅÅk = l A + A ÑÑÑÑ. Thus, these results 2 1 Ñ ÅÇ Ö clearly establish that lipid exchange between the planar membrane corral and microbead-supported membrane takes place through diffusion-mediated mixing. Having established that the delivery of lipid molecules to the membrane corrals is due to diffusion of the lipid molecules through the formation of a long fusion-stalk, we wondered if it is possible to deliver lipid molecules covalently modified with DNA oligonucleotides. While desirable, it is not trivial since the functional lipid molecules that we have successfully delivered so far (Ni-NTA−DOGS or biotinyl-Cap-PE) contain a significantly smaller headgroup than a DNA oligonucleotide. We, therefore, attempted to deliver DNA oligonucleotidemodified lipid molecules onto the membrane corrals to create the DNA oligonucleotide and protein-displaying multicomponent, membrane microarray substrates. For this, microbeads were coated with 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine-N-[4-(p-maleimidomethyl)cyclohexane-carboxamide]

(

(PE-MCC) lipid containing supported membrane. Marina blue-1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (MB-DHPE) was included at a concentration of 1 mol % for the purpose of monitoring lipid exchange. A thiolated ssDNA oligonucleotide was conjugated to PE-MCC lipid present in the SLB in the presence of tris(2-carboxyethyl)phosphine (TCEP). Additionally, microbeads coated with a biotincontaining bilayer were simultaneously incubated for the functionalization of membrane corrals with protein (streptavidin) molecules. We used slightly larger microbeads (7 μm in diameter compared with the 5.2 μm for the ssDNA-modified lipid bilayer-coated microbeads) for the delivery of biotin lipid to distinguish the two types of the microbeads during imaging. The large size of the microbeads compared with the size of the membrane corral will preclude exchange between a single corral with multiple microbeads and therefore the two types of microbeads were simultaneously incubated with a TR-DHPEcontaining membrane microarray substrate (Figure 4a,b). This resulted in exchange of lipids between the microbeads and membrane corrals on the micropatterned substrate, as determined from the decrease of TR-DHPE intensity and an increase in the MB-DHPE intensity in some membrane corrals (Figure 4c). The emergence of low TR-DHPE fluorescence intensity of individual membrane corrals after microbead incubation confirmed the exchange of lipid between some of the membrane corrals and microbeads (Figure 4d,e). Following lipid exchange, we incubated the substrate with the fluorescently labeled complementary DNA oligonucleotide

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Figure 4. Microfabrication of DNA- and protein-functionalized multicomponent, membrane microarray substrate. (a) Schematic showing steps in the microfabrication of DNA- and protein-functionalized, spatially segregated, multicomponent membrane microarray substrates. (b) Epifluorescence image of TR-DHPE membrane microarray substrate before lipid exchange. (c) Epi-fluorescence (red, TR-DHPE; blue, MB-DHPE), reflection interference contrast microscopy (RICM) and bright field images of membrane microarray during lipid exchange. DNA-functionalized microbeads were marked by the inclusion of MB-DHPE, whereas biotin lipid-functionalized microbeads could be detected from their larger (7 μm) diameter. Lipid exchange between the DNA-functionalized microbead and bilayer corral results in the transfer of MB-DHPE into the bilayer corral (blue fluorescence). (d, e) Graph showing frequency distribution of TR-DHPE fluorescence intensity in individual corrals before (d) and after lipid exchange (e). (f) Epi-fluorescence images of a DNA- and protein-functionalized, four-component, micropatterned SLB substrate (red, TR-DHPE; cyan, SA (streptavidin-Dylight 680); green, DNA-Alexa Fluor 488; blue, MB-DHPE; dark, PLL-g-PEG-RGD) after lipid exchange and incubation with streptavidin and DNA. The bilayer corrals are 2 μm in diameter with an intercorral spacing of 2 μm. (g) Intensity linescan of DNA-containing multicomponent, membrane microarray substrate. (h) Graph of MB-DHPE intensities vs cDNA intensities, showing a correlated exchange of MBDHPE and DNA-conjugated lipid molecules. (i) Graph showing frequency distribution of fluorescence intensity of complementary DNA oligonucleotide bound to ssDNA on individual corrals.

(cDNA) and fluorescently labeled streptavidin and imaged the substrate by epi-fluorescence microscopy. As shown in Figure 4f, some of the membrane corrals were found to bind the fluorescently labeled cDNA molecules whereas others bound fluorescently labeled streptavidin molecules, indicating the

spatially segregated transfer of ssDNA and biotin-lipid from their respective microbeads to the membrane corrals (Figure 4f). Thus, the membrane microarray substrate containing three types of bilayer corrals: (i) double-stranded DNA−lipidfunctionalized, (ii) streptavidin−biotin-functionalized, and (iii) 9785

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slips were then cleaned by extensive rinsing with deionized H2O and were dried under a N2 jet. Vesicle Preparation. Lipid vesicles were prepared by mixing constituent lipid molecules DOPC with 16:0 biotinyl-Cap-PE (5 mol %) or TR-DHPE (1 mol %) in chloroform.31 For DNA exchange experiments, vesicles were prepared with DOPC, PE-MCC (1 mol %), and MB-DHPE (1 mol %). All lipids, except TR-DHPE and MBDHPE (Life Technologies), were purchased from Avanti Polar Lipids (Alabaster, Alabama). A thin lipid film was deposited on a round bottom flask by rotary vacuum. Lipids were then resuspended in deionized H2O to a final concentration of 0.5 mg/mL and sonicated to generate small unilamellar vesicles (SUVs). Vesicle suspensions were centrifuged at 20 000 × g and 4 °C for 4 h; supernatants were collected and stored at 4 °C until further use. Bilayer Deposition, Lipid Exchange and Protein Functionalization. All surfaces (planar glass as well as microbeads) were coated with a bilayer by spontaneous fusion of SUVs. PLL-g-PEGRGD micropatterned substrates were incubated with a 1:1 mixture of 99% DOPC + 1% TR-DHPE vesicles and 2× Tris-buffered saline (TBS) for depositing bilayers on the polymer-cured positions on the substrate. Microbeads (Bangs Laboratories, Inc.) of 5.2 μm diameter, unless indicated otherwise, were coated with biotin or PE-MCC lipids, separately. In the DNA-conjugated lipid exchange experiment, biotin-containing SLBs were coated on 7 μm diameter microbeads for the purpose of detection. Post bilayer deposition, both micropatterned substrates and microbeads were washed with 1× TBS and blocked with 0.001% bovine serum albumin for 30 min. All membrane fusion-mediated lipid delivery61,71,72 experiments were performed on a vibration isolating optical table (Thorlabs, Newton, New Jersey) by simultaneously seeding bilayer-coated microbeads (DOPC + biotin lipid SLB-coated microbeads in an 1:1 ratio) on the micropatterned substrate for 15−30 min at 37 °C. For the DNA functionalization experiment, microbeads coated with DOPC + PE-MCC + MB-DHPE SLB were functionalized with ssDNA, as described previously.21 Briefly, the 5′ thiolated, ssDNA (5′GTA ACG ATC CAG CTG TCA CT-3′) was dissolved in TE buffer (5 mM Tris, 0.5 mM ethylenediaminetetraacetic acid, pH 8) at a concentration of 1 mg/mL, while tris(2-carboxyethyl)phosphine (TCEP) was diluted to a concentration of 10 mM with 50 mM (4(2-hydroxyethyl)-1-piperazineethanesulfonic acid) buffer, pH 8.0. Single-stranded DNA was incubated with 2 mM TCEP for 90 min at 37 °C, following which ssDNA was filtered with Bio-spin six columns (Biorad, Hercules, CA). ssDNA was conjugated with PEMCC lipid molecules by incubation at room temperature for 30 min, and unreacted DNA was removed by buffer wash (thrice with 5 mL of 1× TBS). Following lipid exchange, microbeads were removed by inverting and shaking of the substrate inside a chamber containing 1× TBS and substrates were further washed thrice with 5 mL of 1× TBS. For the DNA functionalization experiment, the substrates were incubated with fluorescently labeled streptavidin and 3′ labeled cDNA (5′-AGT GAC AGC TGG ATC GTT AC-3′) simultaneously. Unbound cDNA were removed by washing the substrate thrice with 5 mL of 1× TBS. We note that the fluorescently labeled cDNA showed some nonspecific binding to the PLL-g-PEG-RGD. Imaging. All images were acquired using an Eclipse Ti inverted microscope (Nikon) attached with an Evolve EMCCD camera (Photometrics, Tucson, AZ). All fluorescence images were acquired in the epi-fluorescence mode using a mercury lamp as the light source and appropriate excitation and emission filters. RICM imaging was performed using a filter cube containing a 530:11 nm excitation filter and a 50:50 beam-splitter dichroic mirror. Images were collected using Metamorph (Molecular Devices, Sunnyvale, CA) and analyzed with either ImageJ (NIH, Bethesda, MD) or Fiji.73 All representative images were contrasted in ImageJ, and epi-fluorescence images were modified using the Smooth function. All data were analyzed using the GraphPad Prism software (version 6.00; La Jolla, CA).

TR-DHPE lipid-containing nonfunctionalized corrals, and a solid phase RGD peptide was fabricated (Figure 4f,g). We note that the quantity of ssDNA transferred from the microbeads to individual membrane corrals correlates well with the amount of MB-DHPE lipid molecules transferred (Figure 4h), with a narrow distribution of densities across individual corrals (Figure 4i).

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CONCLUSIONS To conclude, we report a significant extension of our previously reported method to fabricate multicomponent membrane microarrays for a spatially segregated display of multiple, membrane-anchored ligands. We validated our analytical model of lipid exchange between a planar and microbead-supported membrane on the basis of one-dimensional lipid diffusion by varying the temperature and size of the membrane corrals. We took advantage of the physical structure of long fusion-stalks formed between exchanging membranes to successfully deliver substantially larger, DNA oligonucleotide-modified lipid molecules. This allowed us to fabricate membrane microarray substrates containing both protein and DNA ligands on spatially segregated membrane corrals. The ability to deliver DNA-modified lipid molecules to membrane corrals significantly improves the utility of the method since DNA-based anchoring strategies allow a high degree of multiplexing in a way that is not possible with protein-only ligand anchoring. One of the limitations of the lipid exchangemediated spatially segregated, multicomponent substrate fabrication is the randomness in the functionalization of bilayer corrals, unlike robotics-based56 or microcontact printing-based52−55 methods, wherein individual corrals could be independently addressed. However, this does not pose a significant problem for single cell assays since the size of membrane corrals is much smaller than cells and therefore, it is highly like that individual cells encounter multiple membrane corrals.64 Further, the ease of fabrication without the requirement for highly sophisticated instrumentation makes it extremely useful, especially in settings where advanced technologies such as robotic deposition system or microcontact printing are unavailable. Thus, multiple types of cellular receptor-activated biochemical/signaling reactions could be monitored in single cells in a spatially resolved fashion. We envisage that these substrates could also be applied for simultaneous monitoring of biochemical reactions on membrane surfaces in vitro38,41,46,68 or detecting multiple chemical species in a multiplexed biosensing assay.45,69,70

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METHODS

Micropatterned Membrane Microarry Substrate Preparation. Glass cover slips were first cleaned by sonication in 50% isopropanol, rinsed with a copious amount of deionized H2O, and then incubated overnight in 50% H2SO4, following which they were rinsed with a copious amount of deionized H2O. Finally, cover slips were illuminated with ultraviolet (UV) light in a UV oven for 15−30 min. Cleaned glass cover slips were dried under a N2 jet and coated with PLL-g-PEG-RGD (Susos AG, Dübendorf, Switzerland) by incubation with the polymer (1 mg/mL) for a minimum of 2 h at room temperature. Cover slips were extensively rinsed with deionized H2O to remove unbound polymer molecules and dried under a N2 jet. Quartz photomasks designed to contain centimeter-scale features of transparent circles (2, 4, or 5 μm diameter with equivalent spacing) were then used to create a similar pattern on the polymer-coated glass cover slips by photolithography involving illumination with a highintensity UV light for 8 min in a UV oven. UV-exposed glass cover 9786

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected] (K.H.B.). *E-mail: [email protected] (J.T.G.). ORCID

Kabir H. Biswas: 0000-0001-9194-4127 Nam-Joon Cho: 0000-0002-8692-8955 Jay T. Groves: 0000-0002-3037-5220 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by the National Research Foundation, Singapore through the Mechanobiology Institute, National University of Singapore and Competitive Research Program grant CRP001-084 awarded to J.T.G. and University of California, Berkeley.

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