Rapid and Automated Formation of Suspended Lipid Bilayer Arrays for Parallel Ion Channel and Protein Nanopore Recording G

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RAPID AND AUTOMATED FORMATION OF SUSPENDED LIPID BILAYER ARRAYS FOR PARALLEL ION CHANNEL AND PROTEIN NANOPORE RECORDING G. Baaken1,2*, E. Zaitseva1,2, S. Petersen1,2, J.M. del Rio Martinez1, J.C. Behrends1 1Membrane Physiology and Technology, Dept. of Physiology, University of Freiburg, GERMANY and 2Ionera Technologies GmbH i.G., Freiburg, GERMANY

ABSTRACT We here report on a novel method for the automated and highly reliable formation of arrays of suspended lipid membranes of diameters of 6-50 µm that allow high fidelity electrophysiological recordings from embedded ion channels or protein nanopores using voltage clamp (potentiostatic) electronics. The method is adapted to a recently developed microelectrode cavity array (MECA) device and allows reliable automation of an important, hitherto badly-controlled step in the realization of parallel and high-throughput recordings of ionic currents through ion channels and biological nanopores.

KEYWORDS: suspended lipid membrane, high-throughput, automation, microelectrode cavity array, ion channel, nanopore

INTRODUCTION After decades of being largely superseded by the patch clamp technique which allows direct recordings from ion channels in native cell membranes, electrophysiology on membrane proteins reconstituted in synthetic lipid bilayers is, for more than one reason, regaining popularity. First, there is increasing interest in a number of ion channels and pore- forming proteins from bacteria, viruses and also from eurkaryotic organelles that are not easily amenable to classical or automated cell-based patch-clamp electrophysiology.[1], [2] Second, due to limitations regarding integration density and miniaturization as well as to relatively complex process control inherent in the microfluidic cell positioning method common to all of them, current cell-based automated patch clamp (APC) devices, while enabling significantly enhanced throughput, are not HTS-capable in the sense of >10.000 data points/d so that cell-free bilayer recording systems are increasingly considered as an alternative at least for certain ion channel types[3]-[7]. Third, and perhaps most importantly, the burgeoning field of single-molecule analytical techniques based on measuring ionic currents through biological nanopores such as the bacterial toxins and porins a-hemolysin, aerolysin and mycobacterium smegmatis porin A (MspA)

with applications ranging from polymer sizing to DNA Figure 1: A: : Schematic of one cavity. Lithographical- sequencing[8] constitute a strong motivation to search for ly structured substrate (SU8) is shown in light grey, simple and efficient technologies for the formation of Ag/AgCl electrode in dark grey, gold in yellow and glass and electric readout from arrays of lipid bilayers. Several in light yellow. Membrane is shown containing an α- groups have in the past decades developed various Hemolysin pore interacting with a polymer (PEG) B: approaches aimed at bilayer miniaturization, Background: active area of the MECA Chip. 16 Cavities parallelization and automation, ranging from are contacted by coplanar strip lines; Foreground: Histo- miniaturized and parallelized versions of classical gram showing peaks of residual current during block “vertical bilayer” devices via horizontal microfluidic events by PEG molecules of different oligomeric size. approaches to droplet bilayers.[9] We have recently introduced a a chip-based platform containing an array of Ag/AgCl electrodes in microcavities (MicroElectrode Cavity Array, MECA, Fig. 1) structured in SU8 and demonstrated its application in parallel single molecule detection [10], [11]. For these experiments, bilayers were formed manually using the painting procedure (see below). We now show that bilayer formation on the MECA can be automated using remote painting via magnetically controlled sweeping motion of a teflon-coated effector.

THEORY There are two classical approaches to the de-novo formation of suspended lipid membranes: the Müller-Rudin or “painting” technique[12], where the bilayer is formed by spreading from a bulk solution of lipid in organic solvent and the Montal-Müller technique[13], [14] where it is formed from monolayers at the air-water interface. Both techniques require the presence of nonpolar solvent such as to enable formation of a so-called annulus at the rim of the aperture in a substrate such as Teflon, over which the free-standing bilayer is formed.[15] With the spreading technique, the classical observation is that the bimolecular lipid layer forms by gradual thinning of a lipid/solvent film that is mechanically

978-0-9798064-6-9/µTAS 2013/$20©13CBMS-0001 1520 17th International Conference on Miniaturized Systems for Chemistry and Life Sciences 27-31 October 2013, Freiburg, Germany stretched out over the aperture. This spontaneous but protracted thinning process, which can be followed by monitoring the increasing electrical capacitance of the layer over minutes, is thought to involve a flow of lipid solution towards the periphery due to a centripetal hydrostatic pressure difference imposed by the curvature of the annulus (Plateau-Gibbs border). Additionally, in later stages, the film is thought to be compressed by long-range van-der-Waals attraction of the aqueous media on either side.[15] Our initial experiments using manual painting on 4x4 MECA with apertures of 16 µm diameter had shown that success rates in bilayer formation were high (>90%) and mechanical bilayer stability was superior to classical systems, requiring no vibration insulation[10]. Usually, a sweeping motion on the with a Teflon whisk that had been immersed in lipid solution resulted in Figure 2: Schematic of the set-up with mag- simultaneous formation of bilayers on the majority of cavities. netic stirrer and countermagnet on the turnable Interestingly, as evidenced by the immediate appearance of rod. channel activity when bilayers were formed in the presence of channel formers, film thinning was a very fast process. We were thus interested in ascertaining whether this painting process could be automated using remotely controlled spreading of lipid, reasoning that a magnetically moved teflon-coated metal bar such as used for stirring purposes might be suitable.

EXPERIMENTAL The general layout of the apparatus used is shown in Fig. 2. A turnable rod below the MECA chip carries an excentrically mounted countermagnet. The arrangements holds the chip approximately 1 mm above the countermagnet. Turning the rod actuates the magnetic "stirring bar" lying on the MECA chip. The rod could be turned using an electromotor; its turning speed was controlled with a tunable resistor. Experiments were performed in a variety of KCl-based electrolyte solutions ranging from 0.05 to 4 M concentration (buffered to between pH 7 and 8.5 using TRIS-base or HEPES (10 mM) as appropriate. The MECA chip was connnected to a 16-channel voltage clamp amplifier (Jet-16 , Tecella, San Diego USA) which was run from a Personal Computer using Tecellalab software for voltage control and data acquisition. In one embodiment of this technique, a small teflon coated metal rod, i.e. a "stirring bar" of dimensions 6 x 3 Figure 3: Left: Cross sectional schematic of the mm is positioned on top of the MECA chip in the presence MECA illustrating bilayer formation by remotely ac- of electrolyte solution. Following pipetting of a small tuated painting (see text). Right: Sketch of the remote- amount (<=1-3 µl) of lipid in solvent to the chip surface in ly actuated movement of the teflon-coated bar induced the immediate vicinity, the bar is moved across the aper- by one turn of the counter magnet (see Fig. 2) tures in a circuitious fashion (Fig. 2) by performing one slow (45-180°/s) rotation of a counter magnet positioned below the chip.

RESULTS AND DISCUSSION Using Di-Phytanoyl- or Di-Oleoyl-Phosphocholine dissolved in octane, decane, hexadecane or hexadecene (other solvents were not yet employed) at 2-10 mg/ml we obtained bilayer forming success rates always in excess of 90%, generally achieving instant bilayer formation over all 16 cavities present in the rectangular 800 x 800 µm active area of the MECA chip. Seal resistances obtained are in excess of 10 GOhm. Interestingly, unlike with the classical painting method, there is no prolonged thinning-out of the lipid film before bilayer formation but it is fully established in after the initial increase in resistance. This is illustrated in Fig. 4, where channels mediated by the pore-forming peptide gramicidin appeared directly after bilayer formation. The first rectangular channel current due to the dimerization of two gramicidin peptides residing in the opposite leaflets of the double membrane[16] is observed 1050 ms after the initial decrease of conductance due to formation of the lipid film. Because of the "hydrophobic mismatch" between the dimensions of the gramicidin dimer and the phosphatidylcholine bilayer [17], the appearance of gramicidin channels signals maximal thinning of the bilayer over roughly 1 s. Simultaneously, the resistance increases further from an initial 20 GOhms immediately after rotation to 150 GOhms upon channel opening, signalling a concomitant increase in lipid packing density. Despite this rapid formation process, the resulting bilayers present a 'classical' morphology including a lipid-solvent annulus that appears continuous with the lipid-solvent mixture wetting the SU8-surface. We observed that solvent wetting of both the MECA surface and the rotor element is important for the success of this bilayer forming procedure. In experiments with glass-coated rotors, we observed a sharp decline of success rate. The same result is seen when the SU8- surface is made more hydrophilic using UV/ozone treatment (not shown).

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CONCLUSION We conclude that it is possible to automate the Müller-Rudin technique of bilayer formation using a simple rotation mechanism pro- vided that both the rotor element and the substrate surface are suffi- ciently apolar to permit wetting by organic solvents as a prerequisite for ideal spreading of lipid over the aperture of the cavity.

ACKNOWLEDGEMENTS This research was supported by the BioChance (KMU- Figure 4: Current trace showing the rapid for- Innovativ) program of the Ministry of Education and Research (PTJ, mation of a bilayer with the appearance of gramici- FKZ 0315316B) and by the Exist-Research-Transfer-Program of the din-mediated current steps. 150 mV, 1 M HCl. the Ministry of Economics and Technology BMWi (PTJ, FKZ arrowed bracket signals the artifact resulting from 03EFT9BW44) switching off the power supply of the electromnotor used to rotate the actor, which removes the 50 Hz REFERENCES parasitic signal initially contaminating the signal. [1] M. Schieder, K. Rotzer, A. Brüggemann, M. Biel, and C. Inset: fluorescence micrographs (0.5% rhodamin- Wahl-Schott, “Planar Patch Clamp Approach to Characterize Ionic labelled DPhPC) of a MEC without and with a lipid Currents from Intact Lysosomes,” Science Signaling, vol. 3, no. 151, bilayer formed. The annulus is clearly visible as a p. pl3, Dec. 2010. ring of stronger fluorescence at the rim of the cavi- [2] E. M. Nestorovich, C. Danelon, M. Winterhalter, and S. ty. M. Bezrukov, “Designed to penetrate: time-resolved interaction of single antibiotic molecules with bacterial pores,” Proc Natl Acad Sci U S A, vol. 99, no. 15, pp. 9789–9794, Jul. 2002. [3] S. A. Portonovo, C. S. Salazar, and J. J. Schmidt, “hERG drug response measured in droplet bilayers,” Biomed. Microdevices, vol. 15, no. 2, pp. 255–259, Nov. 2012. [4] A. M. El-Arabi, C. S. Salazar, and J. J. Schmidt, “Ion channel drug potency assay with an artificial bilayer chip,” Lab Chip, vol. 12, no. 13, p. 2409, 2012. [5] S. Leptihn, J. R. Thompson, J. C. Ellory, S. J. Tucker, and M. I. Wallace, “In Vitro Reconstitution of Eukaryo- tic Ion Channels Using Droplet Interface Bilayers,” J. Am. Chem. Soc., vol. 133, no. 24, pp. 9370–9375, Jun. 2011. [6] J. L. Poulos, T.-J. Jeon, R. Damoiseaux, E. J. Gillespie, K. A. Bradley, and J. J. Schmidt, “Ion channel and to- xin measurement using a high throughput lipid membrane platform,” Biosensors and Bioelectronics, vol. 24, no. 6, pp. 1806–1810, Feb. 2009. [7] T. Thapliyal, J. L. Poulos, and J. J. Schmidt, “Automated lipid bilayer and ion channel measurement platform,” Biosensors and Bioelectronics, pp. 1–4, Feb. 2010. [8] J. J. Kasianowicz, J. W. F. Robertson, E. R. Chan, J. E. Reiner, and V. M. Stanford, “Nanoscopic Porous Sen- sors,” Annu. Rev. Anal. Chem., vol. 1, no. 1, pp. 737–766, Jul. 2008. [9] M. Zagnoni, “Miniaturised technologies for the development of artificial lipid bilayer systems,” Lab Chip, vol. 12, no. 6, p. 1026, 2012. [10] G. Baaken, M. Sondermann, C. Schlemmer, J. Rühe, and J. C. Behrends, “Planar microelectrode-cavity array for high-resolution and parallel electrical recording of membrane ionic currents,” Lab Chip, vol. 8, no. 6, pp. 938–944, 2008. [11] G. Baaken, N. Ankri, A.-K. Schuler, J. Rühe, and J. C. Behrends, “Nanopore-Based Single-Molecule Mass Spectrometry on a Lipid Membrane Microarray,” ACS Nano, vol. 5, no. 10, pp. 8080–8088, Oct. 2011. [12] P. MUELLER, D. O. RUDIN, H. TI TIEN, and W. C. WESCOTT, “Reconstitution of excitable cell membrane structure in vitro,” Circulation, vol. 26, no. 5, pp. 1167–1171, 1962. [13] M. Takagi, K. Azuma, and U. Kishimoto, “A new method for the formation of bilayer membranes in aqueous solution,” Ann. Rep. Biol. Works Fac. Sci., vol. 13, pp. 107–110, Jul. 1965. [14] M. Montal and P. Mueller, “Formation of bimolecular membranes from lipid monolayers and a study of their electrical properties,” Proc Natl Acad Sci U S A, vol. 69, no. 12, pp. 3561–3566, 1972. [15] S. H. White, “The physical nature of planar bilayer membranes,” in Ion Channel Reconstitution, 1st ed., no. 1, C. Miller, Ed. New York: Plenum, 1986, pp. 3–35. [16] V. Borisenko, T. Lougheed, J. Hesse, E. Fureder-Kitzmuller, N. Fertig, J. C. Behrends, G. A. Woolley, and G. J. Schutz, “Simultaneous optical and electrical recording of single gramicidin channels,” Biophysj, vol. 84, no. 1, pp. 612–622, Jan. 2003. [17] T. K. Rostovtseva, H. I. Petrache, N. Kazemi, E. Hassanzadeh, and S. M. Bezrukov, “Interfacial polar interac- tions affect gramicidin channel kinetics,” Biophysical Journal, vol. 94, no. 4, pp. L23–5, Feb. 2008.

CONTACT *G. Baaken, tel: +49-761-203-5145; [email protected]

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