Transparent nanopore cavity arrays enable highly parallelized optical

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Transparent nanopore cavity arrays enable highly parallelized optical studies of single membrane proteins on chip Tim Diederichs, Quoc Hung Nguyen, Michael Urban, Robert Tampé, and Mark Tornow Nano Lett., Just Accepted Manuscript • DOI: 10.1021/acs.nanolett.8b01252 • Publication Date (Web): 09 May 2018 Downloaded from http://pubs.acs.org on May 10, 2018

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Nano Letters

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Transparent nanopore cavity arrays enable highly parallelized

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optical studies of single membrane proteins on chip

3 4 Tim Diederichs1,6, Quoc Hung Nguyen2,6, Michael Urban1, Robert Tampé1,3,*, Marc Tornow2,4,5*

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1

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Institute of Biochemistry, Biocenter, Goethe-University Frankfurt, Max-von-Laue-Str. 9, 60438

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Frankfurt/M., Germany; 2Molecular Electronics, Technical University of Munich, Theresienstr. 90,

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80333 Munich, Germany; 3Cluster of Excellence Frankfurt (CEF) Macromolecular Complexes; Goethe-

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University Frankfurt, Max-von-Laue-Str. 9, 60438 Frankfurt/M., Germany; 4

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Fraunhofer Research Institution for Microsystems and Solid State Technologies (EMFT), Hansastr.

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27d, 80686 Munich, Germany; 5Center for NanoScience (CeNS), Ludwig-Maximilians-University,

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Geschwister-Scholl Platz 1, 80539 Munich, Germany

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* Correspondence should be addressed to R.T. ([email protected]) and

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M.T. ([email protected])

These authors contributed equally to this work

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Key words: silicon-on-insulator chips, transport kinetics, membrane proteins, optical readout,

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supported lipid bilayers, nanopores, microcavities

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Abstract

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Membrane proteins involved in transport processes are key targets for pharmaceutical

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research and industry. Despite continuous improvements and new developments in the field of

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electrical readouts for the analysis of transport kinetics, a well-suited methodology for

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high-throughput characterization of single transporters with non-ionic substrates and slow

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turnover rates is still lacking. Here, we report on a novel architecture of silicon chips with

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embedded nanopore microcavities, based on a silicon-on-insulator technology for

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high-throughput optical readouts. Arrays containing more than 14,000 inverted-pyramidal

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cavities of 50 femtoliter volumes and 80 nm circular pore openings were constructed

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via high-resolution electron-beam lithography in combination with reactive ion etching and

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anisotropic wet etching. These cavities feature both, an optically transparent bottom and top

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cap. Atomic force microscopy analysis reveals an overall extremely smooth chip surface,

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particularly in the vicinity of the nanopores, which exhibits well-defined edges. Our

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unprecedented transparent chip design provides parallel and independent fluorescent readout

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of both, cavities and buffer reservoir, for unbiased single-transporter recordings. Spreading of

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large unilamellar vesicles with efficiencies up to 96% created nanopore-supported lipid

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bilayers, which are stable for more than one day. A high lipid mobility in the supported

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membrane was determined by fluorescent recovery after photobleaching. Flux kinetics of

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α-hemolysin were characterized at single-pore resolution with a rate constant of

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0.96 ± 0.06×10–3 s–1. Here, we deliver an ideal chip platform for pharmaceutical research,

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which

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single-transporter resolution.

features

high

parallelism

and

throughput,

synergistically

combined

with


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1. Introduction

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Up to date, membrane proteins (MPs) are targeted by more than half of all developed

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drugs,1 underlining their outstanding importance for pharmaceutical industry and basic

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research. They are involved in a multitude of essential cellular processes including

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bioenergetics, transport, and communication. Especially, their role in trafficking substrates

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across lipid bilayers or in artificial nanocontainers became an important research focus for the

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identification of new drug targets as well as for biosensing applications.2

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Multiple techniques for the characterization of MP kinetics are available. For instance,

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patch clamp is able to directly analyze ion channel activity in cell membranes, however, it is

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limited to single or several simultaneous channel recordings.3-6 High-throughput screening

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applications are confined by the complexity of interfaces between electronics and ionic

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systems,7 except for some advanced techniques using whole-cell or automated patch-clamp

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recordings.8-11 For parallel analysis of electrogenic transporter activities, an alternative

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electrophysiological method has been based on solid-supported membranes.12,13 Yet, this

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technique is unable to achieve the single-transporter level. While these electrode-based

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approaches are capable of measuring ionic fluxes or the translocation of ionic substances they

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still lack the ability to characterize transporters with non-electrogenic solutes. Other tools for

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the analysis of MPs have been utilized, including tethered bilayers,14,15 native vesicle

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arrays,16-21 and micro black-lipid membranes,3,22-25 which are compatible with non-ionic

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cargos. So far, these applications are unsuitable for highly parallelized kinetic studies of MPs.

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In the last decade, supported lipid bilayers (SLBs) have gained attention due to their

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long-term stability and their ability to mimic native membranes.26,27 However, the

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incorporation of transmembrane proteins is inherently limited because of their close proximity

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to the solid support.15,28 To overcome this drawback, silicon-based microcavities were

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spanned with membranes creating a platform of supported and free-standing lipid bilayers

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with techniques such as spreading of giant unilamellar vesicles (GUVs),29 large unilamellar

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vesicles (LUVs),30-32 or shear-driven SLB formation.33 In order to transfer the methodology

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from laboratory research to industrial screening applications, it is required to have in vitro

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systems (i) revealing highly automated channel recordings for high-throughput applications,

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(ii) minimizing the sample consumption, and (iii) possessing long-term stability to ensure

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repeatability without suffering from degradation of sensor interfaces.34 Recent advances in

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semiconductor fabrication techniques of micro- and nanostructures with well-defined

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dimensions as well as the development in the field of solid supported membranes provide a

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promising direction for studying MP mediated transport processes.28,35


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Silicon-based chips containing arrays of nanopores with microcavities combined with

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fluorescent readout are in the focus of particular interest, as they enable highly parallel protein

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recordings across lipid membranes. Non-electrogenic membrane transport proteins can be

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incorporated via self-spreading of liposomes and their kinetics can be investigated, which is,

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however, not feasible with electrical set-ups.34 Only a few examples for multiplexed analysis

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using silicon arrays were published, including the characterization of the mechanosensitive

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channel of large conductance (MscL),36 the FOF1-ATP synthase, and α-hemolysin.37,38

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Recently, SLBs were produced with the aid of organic solvent across femtoliter or attoliter

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sized cavities,39,40 suffering from the fact that organic solvents may be harmful to MPs.

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Kleefen and colleagues used nanopores with femtoliter compartments, fabricated by

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electron-beam lithography (EBL), reactive dry, and anisotropic wet etching of a Si3N4 covered

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silicon chip.38 In particular, the bulk bottom handle wafer substrate was fully opaque for

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visible light, hence not accessible for common backside fluorescent microscopy but limited to

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upward oriented microscopes.38 Due to this limitation and owing to the large buffer

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background fluorescence intensity, it was not possible to separate fluorophores in cavities and

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buffer reservoir for unbiased single-transporter recordings. Nevertheless, fluxes of fluorescent

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substances through α-hemolysin and saponin were characterized at single-protein resolution.

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Urban and colleagues utilized silicon-on-insulator (SOI) substrates, which were processed via

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reactive ion etching (RIE) creating cylindrical cavities.36 Chemical vapor deposition of silicon

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dioxide (SiO2) was applied to reduce the cylindrical micrometer openings to nanometer

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scales.36 Thereby, rough surfaces with intrusions of SiO2 were obtained, leading to diminished

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spreading abilities as well as limited diffusion properties for encapsulated substances.

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Nevertheless, an analysis of MP kinetics was presented for MscL and α-hemolysin.36,41

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Here, we present a novel architecture of silicon chips containing macroscopic-scale arrays

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of optically transparent nanopore microcavities that feature a superior homogeneity in shape

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and size, constituting well-defined femtoliter compartments in SOI substrates. Our

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unprecedented chip architecture combines the advantages of both previously reported chip

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structures: (i) extremely smooth, low-strain surface and pore surroundings with well-defined

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and tapered pore edges, and (ii) top and bottom chip transparency, allowing for full optical

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readout access for inverted microscopy. Hence automated and multiplexed readout as well as

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screening become possible. Cavity volume, nanopore diameter, and the patterning of cavity

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arrays (size and shape) can be precisely controlled on wafer scales via high-resolution EBL in

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combination with RIE and anisotropic wet etching. Our chips contain more than 14,000

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inverted-pyramidal cavities arranged in a rectangular pattern with pore distances of 10 µm


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and openings of 80 nm. As validated by atomic force microscopy (AFM) analysis, the

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surfaces are extremely smooth on pore surroundings with well-defined and tapered pore

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edges, hence providing optimal spreading conditions for lipid bilayers.42 The material

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properties of the employed SiO2 and Si3N4 layers allow an optically transparent chip design

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for parallel fluorescence readout of both, cavities and buffer reservoir, for unbiased

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single-transporter recordings in a highly parallel fashion. We show that as anticipated, our

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chip platform indeed features high spreading efficiencies of large unilamellar vesicles (LUV),

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high lipid mobility, and long-term stability of the free-standing, nanopore-supported lipid

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bilayer. As proof of concept, kinetics of the model α-hemolysin pore were analyzed down to

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single-pore resolution in high-throughput.


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2. Experimental section

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2.1 Silicon-on-insulator substrate

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Si3N4 coated, prime-quality 4-inch diameter SOI wafers (Si-Mat, Kaufering, Germany)

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were composed of a top 3 ± 0.5 µm thick undoped silicon (100) device layer, which was

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separated from the 380 ± 15 µm undoped bulk silicon (100) substrate by a 100 ± 10 nm buried

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oxide layer (SiO2, “BOX”). Stoichiometric Si3N4 (thickness ∼50 nm) was then deposited on

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both sides of the substrate by low pressure chemical vapor deposition (LPCVD). The wafers

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were sawed into 13 x 13 mm2 pieces for the following nanopore cavity structure fabrication.

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2.2 Fabrication and characterization of nanopore cavity arrays with transparent bottoms

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The fabrication process of nanopore cavity arrays is schematically depicted in Figure 1.

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First, a large pyramidal pit was structured on the backside of the chip by a combination of

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optical lithography, RIE, and aqueous potassium hydroxide (KOH (aq)) wet etching. For this

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step, Shipley S1818 photoresist was spun onto the wafer at a speed of 6000 rpm for 60 s

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followed by a soft-bake at 115 °C for 3 min. We then exposed the resist to UV light

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(wavelength 365 nm, power density ∼6.2 mW/cm2) for 15 s through a square mask

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(1.8 x 1.8 mm2) at the center of the wafer using a mask aligner (SUSS MicroTec, Garching,

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Germany). Subsequently, the chip was developed in DEV 351 solution (diluted 1:5 vol% in

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DI water) for 30 s, followed by a RIE (Oxford Instruments, Oxfordshire, UK) step using a

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C4F8/O2 gas mixture for 85 s to remove the exposed Si3N4 layer. After rinsing with acetone,

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isopropanol, and drying with nitrogen flow, the chips were anisotropically chemically etched

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in 20 wt% KOH (aq) solution at 80 °C to remove bulk silicon from the backside, resulting in a

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large pyramidal etch pit. The etching was stopped after 3 h when the etch pit reached the

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BOX layer which served as an etch stop.

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For the fabrication of cavity arrays, the chips were extensively rinsed with water to

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remove KOH residues prior to the following EBL process. For EBL, an e-beam resist (AR-P

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6200, Allresist, Strausberg, Germany) was spun on the chip front side at 3000 rpm for 60 s

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and prebaked at 150 °C on a hotplate for 1 min, resulting in a resist thickness of ∼100 nm.

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Arrays of nanopores (120 x 120 pores) with diameter of 80 nm and pore distance of 10 µm

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were patterned by an e_LiNE system equipped with the Elphy Quantum pattern generator

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(Raith, Dortmund, Germany) at an acceleration voltage of 30 kV and with beam currents of

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∼30 pA. The development of the patterns was done in developer AR 600-546 (Allresist,

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Strausberg, Germany) for 1 min. Subsequently, the chips were rinsed with isopropanol and

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dried under nitrogen flow. After a post-baking at 130 °C for 1 min, the nanopore patterns 6 ACS Paragon Plus Environment

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were transferred to the Si3N4 layer by RIE, as described above. The residual resist was then

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removed by immersion in acetone and isopropanol for 10 min each, followed by drying with

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nitrogen. Finally, arrays of femtoliter cavities with inverted pyramidal shape were formed by

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an anisotropic wet etching step of the Si device layer in 15 wt% KOH (aq) at 50 °C, using the

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Si3N4 nanopore as an etch mask. Under these conditions, the etching of mainly Si(111) planes

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proceeds at a very low rate due to the high etching anisotropy (0.9 ± 0.1 µm h–1). The final

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cavity volume is adjusted by the etching time. For a 50 femtoliter cavity volume, the etching

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takes about 3.5 h, resulting in cavities with free-standing Si3N4 membranes of ∼36 µm2 in

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size, forming the cavity tops, and transparent cavity bottom areas of ∼3 µm2.

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The nanopore cavity arrays were characterized by scanning electron microscopy (SEM)

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using the e_LiNE system (Raith, Dortmund, Germany) at an acceleration voltage of 5 kV. The

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surface topography of the Si3N4 membranes on top of the cavities, including the single

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nanopores, was imaged by high-resolution AFM, utilizing a Veeco Dimension V system

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(Veeco Instruments Inc., New York, United States) equipped with a Veeco Nanoscope V

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controller in Tapping ModeTM. For all topographic measurements, Si cantilever probes

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(model: SSS-NCLR (super sharp), NanosensorTM, Neuchâtel, Switzerland) were used, having

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nominal force constant of 48 N/m and resonance frequency of 190 kHz. These probes feature

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a tip radius of about 2 nm, which helps to minimize tip/sample convolution effects. The

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surface roughness was determined as root-mean-square (rms) value.

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2.3 Liposome preparation LUVs

composed

of

1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine

(POPC),

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1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE), and 1-palmitoyl-2-oleoyl-

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sn-glycero-3-phosphoglycerol (POPG) were purchased from Avanti Polar Lipids Inc.

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(Hamburg, Germany), and mixed in molar ratios of 4:3:3 to a concentration of 5 mg/ml and

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subsequently resolved in HEPES buffer (20 mM HEPES/NaOH, 150 mM NaCl, pH 7.4).

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LUV formation was conducted as described.38,43 Liposomes were extruded 21 times through

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100 nm polycarbonate membranes at the LiposoFast-Basic extruder (AVESTIN, Mannheim,

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Germany). Vesicle sizes were analyzed using the nanoparticle tracking system NanoSight

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LM10 (Malvern, Herrenberg, Germany).

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2.4 Chip preparation

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The SOI chips were glued on 8-well sticky-slides (ibidi, Planegg/Martinsried, Germany),

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followed by oxygen plasma cleaning for 2 min at 0.3 mbar and at 80% power settings with a


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plasma cleaner (Diener Electronics, Ebhausen, Germany). After immersion in 300 µL of

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ethanol for 5 min, successive exchange of ethanol was conducted by dilution with HEPES

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buffer including 5 mM CaCl2 (total volume 500 µL). This step was repeated 15 times to

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completely dilute ethanol and ensure wetting of the microcavities. Different dyes were

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encapsulated via spreading (1 h incubation) of LUVs (1 mg/mL), followed by buffer

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exchange to remove lipid and fluorophore excess. Optical readout was performed with the

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confocal laser scanning microscope (CLSM) LSM 880 (AxioObserver from Zeiss, Jena,

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Germany) equipped with a Plan-Apochromat 20x/0.8 M27 air objective or with the automated

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NyONE epifluorescence microscope (SYNENTEC GmbH, Elmshorn, Germany) equipped

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with a 20x magnification air objective (Olympus UPlan SApo 20x/0.75). Images were

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recorded at different time intervals dependent on the experiment. Each assay was stopped by

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adding 25 µL Triton-X 100 10% (v/v) to induce SLB rupture, that induce rapid efflux of

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fluorescent molecules and represented additional proof for the absence of unspecific binding.

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2.5 Spreading efficiency, long-term stability, and lipid fluidity of the SLB

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The spreading of LUVs and the resulting SLB were visualized via fluorophore

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encapsulation. 10 µM of ATTO655 (ATTO-TEC, Siegen, Germany) and 5 µM of Oregon

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Green (OG) dextran (10 kDa) (ThermoFisher Scientific, Darmstadt, Germany) were added,

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followed by liposome spreading and buffer exchange. Sealing efficiencies were calculated as

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ratio of dye encapsulated cavities regarding total cavities per field of view. Pearson

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correlation coefficients were determined for the location of both fluorophores in the

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individual cavities with ImageJ 1.51o. The long-term stability was monitored at different time

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points (1, 3, and 24 h) after vesicle spreading with the NyONE microscope. For this purpose,

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5 µM of ATTO655 was encapsulated inside the cavities.

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FRAP experiments were conducted at the CLSM with LUVs composed of

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POPC/POPE/POPG

as

described

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oxacarbocyanine perchlorate (DiO 0.5 mol%) (ThermoFisher Scientific, Darmstadt,

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Germany). After spreading, small circular areas of 8 to 15 µm2 diameter were bleached within

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4 s with high laser intensities. Fluorescence recovery was monitored for ~ 1 min in ~ 100 ms

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intervals dependent on the bleached area. The normalized fluorescence intensity (It) was

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calculated according eq. 1,

  t    t  I t =  I o + I inf     1 +   τ 1/ 2    τ 1/2   

and

additionally

doped

with

3,3ꞌ-dioctadecyl-

−1

(1)


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In which Io is the initial fluorescence, Iinf the fluorescence intensity after recovery, and τ½ the

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time at half-maximal fluorescence intensity. The lateral lipid diffusion coefficient (D) was

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determined regarding eq. 2,44-46

D= 4

ro2γ 4τ1/2

(2)

with ro as the radius of the excitation spot and γ as the bleaching parameter.47

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2.6 Single-transport recordings

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10 µM of ATTO655 (0.6 kDa) and 5 µM of OG dextran (10 kDa) were encapsulated via

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liposome spreading. After buffer exchange, α-hemolysin was added to final concentrations of

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500, 100, or 50 nM. Data were collected in 11 s intervals for 3-4 h with the CLSM.

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Unspecific binding of enclosed dyes was foreclosed by addition of 25 µL Triton-X 100

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10% (v/v) after the experiment. The resulting time traces were processed via the Zen 2.1 black

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software from Zeiss and the corresponding mean grey values were calculated via ImageJ,

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followed by plotting and fitting of the translocation kinetics with the Nanocal software

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(Nanospot GmbH). Two different types of exponential decays were identified: (i) sharp

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mono-exponential decays and (ii) more sigmoidal decreasing curves abruptly converting to

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sharp exponential decays. Both types of curves were fitted with mono-exponential equations,

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whereas the point of changing slope of curve type (ii) was used as lower fit boundary (see

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SI Figure 6). First-order rate constants (keff) were further processed with Origin 9.1 Pro

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(OriginLab), including descriptive statistics to count traces in individual intervals, peak

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analysis (Origin 9.1 Pro, peak analyzer), and Gaussian fitting.38 Rate constant errors were

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obtained for both, the fitting error of the single exponentials and the Gaussian distribution fit

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of the histograms. To provide an upper bound estimate for the total error, we list the larger

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derived value of both in each case, respectively.


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3. Results and Discussion

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3.1 Device fabrication and characterization

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Nanopore cavity arrays with transparent cavity bottoms were realized by a combination of

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different processes that included optical lithography, EBL, RIE, and KOH (aq) wet etching.

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The fabrication flowchart is schematically shown in Figure 1 and detailed in the Experimental

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Section. A SOI wafer with 3 µm silicon (100) device layer and 100 nm buried oxide (BOX)

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was used as a starting substrate, which was then coated with 50 nm Si3N4 by LPCVD on both

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sides. A square window in the Si3N4 layer on the backside of the wafer was processed by

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optical lithography and RIE. Subsequently, an anisotropic wet etch in KOH solution was

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performed to remove the thick bulk Si through this window, resulting in a large etch pit of

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pyramidal shape truncated at the BOX layer (etch stop layer; Figure 1A). Here, the shape of a

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large pyramidal cavity is formed because the etch rate of Si perpendicular to the (100) crystal

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planes is much higher than perpendicular to the (111) planes.48 In 20% (wt/v) aqueous KOH

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solution at 80 °C, we observed that the ratio of etch rates (100)/(111) ranged from 45:1 to

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50:1.

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Figure 1. Scheme of the fabrication process of nanopore cavity arrays in SOI wafer. (A) A large pyramidal etch pit connecting the chip bottom to the BOX layer is created by optical lithography, RIE, and KOH (aq) wet etching. (B) and (C) nanopores (array, 120 x 120 pores) with 80 nm diameter in the Si3N4 top layer are patterned by EBL and another RIE step. (D) A second wet chemical etch in KOH (aq) of the chip results in the formation of inverted-pyramidal cavity arrays. Since SiO2 serves as etch stop layer, the pyramid tip is truncated at the cavity bottom after etching. Dimensions are not drawn to scale. 10 ACS Paragon Plus Environment

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In the second step, a square array of nanopores (120 x 120 pores) with a pore diameter of

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80 nm and pore distance of 10 µm was patterned in the Si3N4 top layer by high-resolution

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EBL and RIE (Figure 1B-C). Finally, the wafer was etched again in KOH solution to fabricate

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a homogenous nanopore cavity array (Figure 1D). The size of the cavities was well controlled

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by adjusting the etching time (see Experimental Section).

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The nanopore cavity array was characterized by SEM. The cavities form a well-ordered

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array with highly homogenous cavity sizes (Figure 2A). The Si3N4 layers on top of the

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cavities are intact and appear as dark grey squares as they are almost transparent for the

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electron beam. An image of a single cavity reveals a single pore in the very center of the

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Si3N4 membrane, with diameter of ∼80 nm (Figure 2B-C). The magnification image of one

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cavity with the Si3N4 top layer almost removed confirms the inverted pyramidal shape of the

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cavity (Figure 2D). The pyramid tip, visible as a small black square, is truncated at the cavity

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bottom composed of a SiO2 layer.

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Figure 2. Silicon nanopore chip design for multiplexed parallel recordings. 80 nm pores in a Si3N4 layer were fabricated via EBL and RIE, followed by wet etching with KOH (aq) to create more than 14,000 inverted-pyramidal cavities per chip. (A) SEM micrograph of a portion of the whole homogenous array, with nanopore cavities appearing as dark grey squares. (B) and (C) SEM images presenting a single cavity, disclosing a single nanopore in the center of the Si3N4 membrane with diameter of ∼80 nm. (D) Front side view of a single cavity after removal of most of the Si3N4 membrane, disclosing the truncated, inverted pyramidal shape of the cavity with transparent bottom (black square). (E-G) Images of the chip backside after the final KOH etch step showing an array of homogenous but smaller squares compared to the front side. These squares correspond to the truncated, inverted pyramid tops, hence to the transparent cavity bottoms (cf. dark square in Figure 2D). (H) 11 ACS Paragon Plus Environment

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AFM topography showing a 3D rendered image of a nanopore with highly rounded and smooth edges of the pore opening.

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size (Figure 2E-G). The cavity array was well aligned with the BOX layer window exposed

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from the backside, both in the center of the chip. The dimensions of the cavity top and bottom

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squares can be accurately measured by SEM. By further knowing that the angle between the

8

cavity top (Si (100) plane) and the cavity sidewalls (Si (111) plane) is 54.74° and that the

9

height of the cavities is 3 ± 0.5 µm (Si device layer thickness), the volume of the cavities can

10

An SEM investigation from the chip backside verifies that the cavities are homogenous in

be calculated. In this study, chips with cavity volumes of 50 ± 3 femtoliters were used.

11

The topography of the nanopores was investigated by high-resolution AFM in tapping

12

mode using a Si cantilever probe featuring a tip with radius ~2 nm, thereby minimizing

13

tip/sample convolution effects. Figure 2H shows a 3D rendered AFM image of a nanopore,

14

revealing that the nanopore opening has well curved and smooth edges. The topography of the

15

Si3N4 membrane in the area of a certain nanopore cavity was also detailed. SI Figure S1A

16

shows a single cavity with the areas in the middle (freely suspended Si3N4 membrane)

17

appearing to be lower in height than the rest of the Si3N4 surface. From the average height

18

profiles of the marked regions on this image in both horizontal (blue) and vertical (red)

19

direction (SI Figure S1B), we observed that the freely suspended Si3N4 membrane on top of

20

the cavity is weakly bent downwards, from the four edges towards the center of the cavity top

21

where the nanopore is located (downward spikes). This is likely due to small residual stress

22

within the deposited Si3N4 layer resulting in visible strain upon release. Notably, the overall

23

height variance is still extremely small with respect to the lateral dimensions of the cavities,

24

by about a factor of 1/5000 – much less than what we have reported for freely suspended

25

silicon membranes released from SOI.29,49 SI Figure S1C and D present AFM height images

26

of Si3N4 surfaces in a small area close to the pore opening, and in an area where no cavities

27

were structured, respectively, showing ultra-flat surfaces. The rms roughness in both cases is

28

approximately 0.2 nm. Hence together, practically stress-free Si-nitride layers deposited on

29

polished, prime-quality Si wafers constitute an ideal starting system for the preparation of

30

both ultra-flat and smooth suspended solid-state interface as support for lipid membranes. We

31

would like to emphasize that both the highly smooth surface around the pore area and the

32

rounded pore edges are essential for the successful spreading of artificial membranes from

33

LUVs, as confirmed below.

34 35

3.2 SLB: Spreading efficiency, long-term-stability, and mobility


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1

The formation of pore-spanning lipid bilayers occurs via self-spreading of liposomes.

2

Vesicles with an average diameter of ~150 ± 27 nm (SI Figure S2) were used for SLB

3

formation induced by adding CaCl2 (5 mM final). Shielding effects of the divalent cations

4

allow to overcome the repulsion forces and the vesicles adhere via van-der-Waals and electric

5

interactions to the solid support, similar to spreading at SiO2 surfaces.50 The ratio between

6

nanopore opening and vesicle diameter is critical for successful spreading.38,51 Vesicle

7

intrusion into the nanopores was avoided by vesicle sizes larger than the nanopore openings.36

8

A systematic analysis of nanopore and lipid vesicle sizes demonstrated that pores of 80 nm

9

result in the highest spreading efficiency for LUVs of ~150 nm diameter.38

10

Our solvent-free method ensures a spatial compartmentalization of cavities and buffer

11

reservoir, which can be visualized via the strict separation of fluorescent dyes. 10 µM of

12

ATTO655 (0.6 kDa) as well as 5 µM of OG dextran (10 kDa) were selectively trapped by

13

SLB formation inside the cavities. After buffer exchange, cavities with bilayer defects did not

14

show fluorescent signals based on rapid leakage out of the cavities and infinite dilution into

15

the buffer reservoir. In contrast, areas with nanopore-supported membranes show

16

well-defined squared patterns of fluorescence, separated by a center-to-center distance of

17

10 µm. A Pearson correlation coefficient of 0.93 (Figure 3A, merge channel) was obtained for

18

the local correlation of ATTO655 (0.6 kDa) and OG dextran (10 kDa) in the underlying

19

cavities. These findings are consistent with perfect colocalization of both dye molecules in the

20

microcavities. Colocalization deviations, lacking either green or red fluorescence, can be

21

caused by variations of the encapsulation efficiencies. The latter are mainly influenced by the

22

physical properties, of the fluorophores, e.g. size and hydrophobicity, especially of the

23

dextran conjugate. We like to note that regions of interest without a defined colocalization

24

were neglected in further analysis. The homogeneity of fluorophore encapsulation, an optimal

25

cavity filling, and the high signal-to-background ratio of our novel device are illustrated as

26

magnifications of sections of Figure 3A, cf. Figure 3B below.


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1 2 3 4 5 6 7 8 9 10 11 12 13

Figure 3. Sealing efficiency. 10 µM of ATTO655 (0.6 kDa) and 5 µM of OG dextran (10 kDa) were incubated in HEPES buffer with 5 mM CaCl2. Liposomes were added to a total concentration of 1 mg/ml followed by self-spreading, creating a SLB. After buffer exchange, the cavities were imaged by CLSM. Colored cavities (merge channel: both dyes were encapsulated) indicate an intact SLB whereas unsealed cavities or areas with membrane defect and leakage remain dark. (A) Field of view in a CLSM. Overall, a representing sealing efficiency of 94% and a Pearson correlation coefficient of 0.93 for the location of both fluorophores in the individual cavities were observed. (B) The homogeneity of fluorophore encapsulation as well as the high signal-to-background ratios are illustrated as magnifications of sections of the images in A (white squares).

14

perfect wetting abilities and low unspecific binding of fluorescent dyes (SI Figure S3).

15

Furthermore, compartmentalization of fluorophores in cavities and buffer reservoir can be

16

achieved and imaged due to the transparency of the chip design. For this purpose, ATTO655

17

was trapped via vesicle spreading, followed by buffer exchange and OG dextran addition

18

(SI Figure 4, merge channel). The focus area of the CLSM was adjusted horizontally to the

19

cavities to visualize solely the chip cavities. The rounded edge shape and the surface of the

20

pore surroundings with sub-nanometer roughness revealed perfect conditions for liposome

21

spreading with 80-96% sealing efficiency.42

Optimal conditions for optical single-transporter translocation assays were provided by

22

As illustrated before, advanced applications will be the investigations of slow, non-ionic

23

transporters. Therefore, long observation periods with suspended lipid bilayers stable for

24

multiple hours are required. The long-term stability was analyzed for 24 h and visualized via

25

selectively entrapped ATTO655 molecules. Previous studies revealed high SLB stabilities for

26

nanopores in the range of 100 nm.42 Our SLBs are stable for 3 h without any leakage of 14 ACS Paragon Plus Environment

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1

fluorophores (see Figure 4A). Even after 24 h, more than 67% of the cavities are still sealed.

2

Notably, fluorophore bleaching as well as membrane permeability can be neglected due to

3

millisecond illumination times and the low membrane interaction factor of ATTO655.52

4

Unspecific binding was ruled out by adding Triton-X 100 10% (v/v) after 24 h to solubilize

5

the SLB.

6

Continuous homogeneous coverage as well as the dynamic properties of the SLB, which

7

are important to preserve the physiological environment of MPs, were examined via FRAP on

8

the SLBs doped with DiO (0.5 mol%). Fluorescence recovery of 97.1 ± 0.3% and a lateral

9

diffusion coefficient of 0.85 ± 0.20 µm2 s–1 are consistent with an intact homogeneous, fluid

10

suspended lipid bilayer determined in nine independent measurements. These values are in

11

agreement with the lateral diffusion coefficient of lipids measured on glass or silicon based

12

surfaces, which range from 1 to 3 µm2 s–1 (Figure 4B).53-55

13 14 15 16 17 18 19 20 21

Figure 4. Long-term stability and high lipid mobility of the SLB. (A) 5 µM of ATTO655 were encapsulated inside the cavities via self-spreading of liposomes, followed by buffer exchange and imaging with epifluorescence microscopy. Images were recorded at different time points after LUV spreading (1, 3, and 24 h). Adding Triton-X 100 10% (v/v) induces bilayer rupture and release of the compartmentalized fluorophore molecules. (B) LUVs doped with DiO (0.5 mol%) were spread on the chip surface and a representing FRAP was recorded with the CLSM.

22

In summary, all results reveal a successful spreading process, resulting in a stable

23

impermeable, and extremely fluidic suspended lipid bilayer, which is able to spatially and

24

temporally compartmentalize molecules in cavities and buffer reservoir. The barrier was used

25

to investigate the transport processes via optical readout and ensures a nature-like

26

environment for integral proteins.


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3.3 Single-transporter recording of the membrane protein complex α-hemolysin

2

α-hemolysin was chosen as model membrane protein complex to investigate MPs at

3

single-transporter resolution and in high-throughput. This bacterial pore-forming toxin binds

4

to the choline groups of phospholipids, followed by oligomerization and spontaneous

5

insertion into lipid bilayers.41 The crystal structure of this heptameric protein revealed a pore

6

with its narrowest opening of 1.4 nm.56-59 Nevertheless, apertures up to 2.9 nm were reported,

7

dependent on lipid composition and buffer/salt conditions, allowing the diffusion of

8

molecules up to 2 kDa in size.56 Prior to SLB formation, 10 µM of ATTO655 (0.6 kDa) and

9

5 µM of OG dextran (10 kDa) were trapped inside the cavities followed by buffer exchange

10

and protein addition. α-hemolysin discriminates fluorophores based on their molecular sizes.

11

Here, ATTO655 acts as a flux reporter whereas the dextran-coupled fluorophore serves as

12

membrane impermeable control. Hence, SLB rupture is indicated by a rapid efflux of both

13

fluorophores, while α-hemolysin-mediated translocation events are classified by both, an

14

exponential decline of the ATTO655 signal and a constant OG signal (Figure 5A). In total,

15

four different situations were observed: (i) α-hemolysin-mediated transport, (ii) membrane

16

rupture, (iii) complex kinetics, and (iv) undefined events. The latter mainly comprised events

17

without control signals of the OG channel, whereas complex kinetics include processes with

18

two declining steps or spontaneous signal increases in one of the two channels. Only traces

19

displaying single-exponential decays in the ATTO655 channel (red) together with a constant

20

signal in the OG channel (green) were accepted for final analysis. Exemplary efflux traces of

21

ATTO655 are shown in Figure 5B. Based on the spontaneous insertion and oligomerization of

22

the toxin, the single-exponential decline occurs at different time points, which is similar to

23

previous studies.38

24

Single-transporter recordings were successively investigated by decreasing the protein

25

concentration. α-hemolysin coalesces to heptameric pores before those insert into the

26

membrane.60 The likelihood for multiple protein insertion per silicon nanopore drastically

27

diminishes with declining protein concentration. We started by applying α-hemolysin

28

(500 nM final), which have been reported to address the range of single-transport

29

recordings.38,41 The efflux rates (keff) of 275 α-hemolysin traces are summarized in a

30

histogram composed of 35 classes. A broad Gaussian distribution was observed with an

31

average efflux value of 3.29 ± 0.23×10–3 s–1 (Figure 5C). Contrary to our observation,

32

single-translocation processes are expected to consist of narrow distributions, as shown for the

33

protein MscL.36 To identify single-transporter events the concentration of α-hemolysin was

34

further decreased successively down to 100 nM and 50 nM (Figure 5D and E, respectively).


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1 2 3 4 5 6 7 8 9 10

Figure 5. Flux kinetics of α-hemolysin in the single transporter regime. 10 µM of ATTO655 (0.6 kDa) and 5 µM of OG dextran (10 kDa) were compartmentalized inside the cavities, followed by buffer exchange and imaging via CLSM. The keff values were determined for single and double incorporations of α-hemolysin after separation in 35 classes for each histogram. (A) Schematic illustration of the transport experiments with possible scenarios including α-hemolysin-mediated transport, rupture, and cavities without fluorescence changes. (B) Exemplary efflux traces of ATTO655 at different pores at the same chip, mediated by α-hemolysin. (C), (D), and (E) show histograms of the ATTO655 efflux rates for 500, 100, and 50 nM of α-hemolysin, respectively. 17 ACS Paragon Plus Environment

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1

204 traces of α-hemolysin-mediated translocation events were analyzed after adding

2

α-hemolysin (100 nM final). The classification as well as the intervals were chosen as

3

described (Figure 5D). Two populations were identified by peak analysis and fitted with

4

Gaussian distributions. The average efflux values of the first Gaussian peak (red) at

5

1.23 ± 0.25×10–3 s–1 clearly suggest that at 500 nM of α-hemolysin (see Figure 5C) single-

6

transporter recordings were superimposed by the elevated statistics of multiple protein

7

insertions per silicon nanopore. This statement is corroborated by similar average efflux

8

values of 3.25 ± 0.44×10–3 s–1 at 100 nM of α-hemolysin (Figure 5D, blue curve) in respect to

9

the average efflux value of 3.29 ± 0.23×10–3 s–1 at 500 nM of α-hemolysin (Figure 5C).

10

The single-transporter regime was reached by finally decreasing the concentration of

11

α-hemolysin down to 50 nM. Here, 117 efflux traces were analyzed and distributed as

12

described above, and two distinct Gaussian peaks were identified. The first population (red)

13

raises at the expense of the second one (blue) (Figure 5E). The 10-fold lower concentration

14

than previously reported for single α-hemolysin recordings,38,41 in combination with only

15

4.8%

16

single-transporter regime has been reached because the likelihood of protein insertion per

17

silicon nanopore has decreased drastically. We interpreted our findings as incorporation of

18

one or two α-hemolysin molecules inside a single silicon nitride nanopore. Thereby, we

19

identified an average efflux value of 0.96 ± 0.06×10–3 s–1 for single- and an average efflux

20

constant of 2.89 ± 0.12×10–3 s–1 for double-protein recordings (Figure 5E). By decreasing the

21

α-hemolysin concentration, the main peak shifted from two protein insertions per nanopore in

22

panel C to one protein per nanopore in panel E.

of

sealed

cavities

showing

mono-exponential

decays,

indicates

that

the

23

Slight shifts in the exact average efflux values at all three concentrations can be explained

24

by statistic variations, especially for the final concentration of 50 nM α-hemolysin. However,

25

all corresponding values are comparable in the range of error. Nevertheless, the concentration

26

series of α-hemolysin clearly demonstrates that our silicon chips are suitable for

27

single-transporter recordings. Additionally, the efflux values for single α-hemolysin

28

recordings are in the range of previous data for MscL.36 Efflux rates of α-hemolysin in the

29

order of 10–5 s–1 were not reproduced.41 However, a later study showed single-protein

30

recordings of α-hemolysin with rates of 5.5×10–4 s–1.37 Their results are in good agreement

31

with our findings and small deviations occur based on different fluorophores, lipids,

32

buffer/salt conditions, and fluorophore concentrations. Especially, the concentration of

33

fluorescent dyes, which is the driving force of the diffusion through α-hemolysin makes it

34

difficult to compare exact efflux values. Nevertheless, our values are in the range of reported


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1

α-hemolysin efflux rates, which clearly demonstrates that our novel chip design is suitable for

2

single-protein recordings.37

3

We finally tested if the efflux kinetics are altered by encapsulation of two dyes in one

4

cavity. Therefore, 10 µM of ATTO655 was encapsulated inside the cavity and 5 µM of OG

5

dextran (10 kDa) was added after SLB formation (SI Figure S4). α-hemolysin was added

6

(500 nM final) and 186 traces were analyzed resulting in an average efflux value of

7

3.40 ± 0.13×10–3 s–1 (SI Figure S5A and S6), consistent with the results above (Figure 5C).

8

Furthermore, rupture events of the SLB were analyzed revealing more than 10-fold faster

9

efflux rates for the diffusion of 10 µM ATTO655, indicating that our single-protein

10

recordings are not biased by lipid rupture processes (SI Figure S5B). In short, single-pore

11

efflux kinetics of α-hemolysin at different concentrations were presented that demonstrate the

12

viability of the transparent nanopore-chips for high-throughput single-transporter studies.

13 14

4. Conclusion

15

In summary, we have presented a novel SOI-based nanopore cavity platform for highly

16

parallelized single-transporter studies using optical readout. The fabrication of the chips,

17

which comprise large, homogenous arrays of cavities with both, top and bottom optical

18

transparency, were controlled precisely on wafer-scale, delivering ideal conditions for

19

screening approaches. A homogenous coverage of the chip surface with an SLB was verified

20

by fluorescent dye encapsulation experiments. Based on the transparent nature of SiO2 and

21

Si3N4 it is possible to track influx as well as efflux processes of fluorescent solutes with high

22

contrast. Single-protein resolution was clearly demonstrated with the bacterial toxin

23

α-hemolysin. The self-spreading process allows the use of proteoliposomes. Thereby,

24

reconstitution of MPs in liposomes can be used to make the presented chip design suitable for

25

medically relevant transporters, like the transporter associated with antigen processing

26

(TAP),61 the transporter associated with antigen processing like (TAPL),62 or Thermus

27

thermophilus multidrug resistance proteins A and B (TmrAB), the functional homolog of

28

human TAP.63 The high lipid mobility as well as the long-term stability of the SLB create a

29

perfect platform for the characterization of non-ionic transporters in single-protein resolution,

30

which we intend to elaborate on in the future. By reducing the cavity volume and increasing

31

the number of microcavities per field of view, higher parallelism can be accomplished, based

32

on thinner top silicon layer architectures and smaller pore-to-pore distances. Further,


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1

vector-based cluster analysis may become a future alternative approach to data analysis for

2

extracting pore populations with transporters.

3

Acknowledgements

4

The German Research Foundation (SFB 807 to R.T.), LOEWE (DynaMem to R.T.), and

5

a German-Israeli Project Cooperation (DIP, TO 266/8-1 to R.T. and M.T.) supported this

6

work. We thank C. Kutter for useful discussions. We are grateful to all members of the

7

Institute of Biochemistry (Goethe University Frankfurt) for helpful comments on the

8

manuscript.

9 10

Supporting Information

11

The Supporting Information is available free of charge on the ACS Publications website at

12

DOI: 10.1021/acs.nanolett.XXX. Figures S1-S6 (PDF).


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