Hydrogel Microelectrodes for the Rapid, Reliable, and Repeatable

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Hydrogel Microelectrodes for the Rapid, Reliable, and Repeatable Characterization of Lipid Membranes Elio J. Challita, and Eric C Freeman Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b02867 • Publication Date (Web): 23 Nov 2018 Downloaded from http://pubs.acs.org on November 24, 2018

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Hydrogel Microelectrodes for the Rapid, Reliable, and Repeatable Characterization of Lipid Membranes Elio J. Challita , Eric C. Freeman a 1

1,

School of Environmental, Civil, Agricultural, and Mechanical Engineering, College of

1

Engineering, University of Georgia, 110 Riverbend Road, Athens, GA, 30605 a

Corresponding Author: [email protected]

Abstract

Model lipid bilayer membranes provide approximations of natural cellular membranes that may be formed in the laboratory to study their mechanics and interactions with the surrounding environment. A new approach for their formation is proposed here based on the self-assembly of lipid monolayers at oil-water interfaces, creating a lipid-coated hydrogel-tipped electrode that produces a stable lipid membrane on the surface when introduced to a lipid-coated aqueous droplet. Membrane formation using the hydrogel microelectrode (HME) is tested for a variety of lipids and oils. The channel-forming peptide alamethicin is added to the membrane and its functionality is verified. Finally asymmetric membranes are created using varying lipid compositions and the capacity for repeated quantification of membrane structure is demonstrated. The proposed hydrogel microelectrodes are compatible with multiple oils and lipids, simple to use, and suitable for detecting the presence of both biomolecular transporters and dissolved lipid compositions within aqueous droplets.

Keywords: Biosensing, Model Membranes, Droplet Interface Bilayers, Asymmetric Membranes, Bioelectrodes, Electrophysiology

ACS Paragon Plus Environment

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Introduction

Lipid bilayer membranes are amphiphilic structures assembled from two lipid leaflets that provide a near-impermeable barrier to ion transport and provide distinction between the cellular exterior and the cytosol. These membranes are the basis for cellular interactions with the surrounding environment, and are able to host a variety of proteins, pores, peptides and channels within the membrane to regulate the transmembrane exchange. Approximations of these cellular membranes in the laboratory from individual components provide model systems to study membrane biophysics and interactions in a controlled environment.

1, 2, 3

There are a variety of

techniques that support the creation of model membranes through self-assembly principles, including but not limited to painted bilayers, tethered membranes, 4

5, 6, 7

giant unilamellar vesicles,

8, 9, 10

and droplet interface bilayers (DIBs).

11, 12, 13

DIBs are built from collections of aqueous droplets in an oil reservoir with dissolved phospholipids in either phase serving as a biomolecular surfactant. The dissolved lipids assemble into ordered lipid monolayers at the oil-water interface on the droplet surfaces, and manipulation of these droplets into contact produces lipid bilayer membranes. Advantages of this system include the ease of formation

13, 14

and compatibility with microfluidic platforms.

15, 16, 17

The DIB technique has

been used successfully for screening droplet contents and membrane qualities by forming and reforming the membrane as needed with a collection of droplets in sequence.

18, 19

Inspired by DIBs, hydrogel microelectrodes (HMEs) are proposed in this work as a model membrane platform with an emphasis on rapid membrane formation, repeatable membrane assembly and disassembly, and reduced membrane dimensions.

These exploit the same

microfluidic mechanisms found in DIBs using a probe with an extruded patch of polyethylene glycol dimethacrylate (PEG-DMA) with an embedded silver/silver-chloride (Ag/AgCl) wire


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insulated within a sharpened glass capillary. The HME produces hydrogel-supported lipid membranes

20, 21, 22, 23

through a “harpoon” mechanic, where lipid-coated aqueous droplets containing

various species are punctured by the lipid-coated hydrogel probe. This electrode is immersed in an oil reservoir to produce an oil-water interface at the surface of the extruded hydrogel, and the lipids dissolved in either the hydrogel or oil reservoir self-assemble into a lipid monolayer on the hydrogel surface. This lipid-coated hydrogel is brought into contact with a lipid-coated aqueous droplet containing a ground electrode, merging the two opposing lipid monolayers into a stable lipid bilayer. This lipid bilayer is able to both form and detach within a fraction of a second as shown by membrane capacitance measurements in Figure 1. Membrane Formation/Separation

a

b

Displacement

Bilayer

Hydrogel Hydrogel

Aqueous

Hydro Hydrogel gel

Aqueous

Oil

b

a 25 pA

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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Oil

a

100 ms

Figure 1 – The proposed electrodes create hydrogel-supported lipid membranes as a lipid-infused gel contacts an aqueous reservoir coated with a lipid monolayer. Membrane formation is measured as a function of the capacitive current response to a 10 mV 100 Hz AC signal applied across the membrane. The membrane formation is rapid, reliable, and repeatable. While similar platforms exist that employ tethered or supported lipid membranes on the surface of the electrode or pore,

24, 25, 26, 27, 28, 29

these do not allow for the separation and reformation of the lipid

membrane. The platform described here forms the membrane as needed, reducing the chance of membrane rupture over time. Since the membrane is formed on the surface of an aqueous hydrogel,


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complications with surface defects are avoided. In addition, tailored dissolution of the lipids 30

within the aqueous phase and the hydrogel allows for variable leaflet components and the simple formation of asymmetric membranes. Results and Discussion Membrane Formation and Mechanics The HME is attached to a linear piezoelectric actuator to measure the relative probe location and provide oscillatory motion as needed using a function generator. As the HME advances and retracts, the membrane capacitance is measured using a triangle-wave voltage (100 Hz, 10 mV) with a patch-clamp amplifier in voltage-clamp mode. The expected current indicating membrane formation is a square wave whose amplitude corresponds directly to the membrane dimensions. Results are exported and processed in MATLAB to provide the membrane capacitance as a function of time. This approach provides simultaneous measurements of the relative tip location and the capacitance of the lipid bilayer membrane formed on the HME surface. The HME was placed just within the droplet surface to form a membrane then oscillated with a 125 µm, 0.2 Hz sinusoidal displacement, advancing and retracting within the droplet while measuring changes in the membrane size. Results demonstrate that the membrane remains intact on the hydrogel surface of the probe even as the HME is fully enveloped by the aqueous droplet as shown in Figure 2. As the probe is pushed further into the droplet, the measured capacitance reaches a plateau, providing a fixed maximum membrane size determined by the area of the extruded hydrogel patch.


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a.

Micropipette for Droplet Deposition

Agarose Coated Electrode

+

-

Aqueous Droplet

400 μm

400 μm

Hydrogel Microelectrode Oil Crate Substrate

b.

Bilayer

Oil

c.

Aqueous

Monolayer

Hydrogel

Figure 2 – a.) The bilayer is formed on the extruded hydrogel path at the tip of the HME and remains intact as the electrode is fully engulfed within the droplet as shown in b) the schematic. c.) A 125 µm sinusoidal oscillation is applied to the HME at the droplet surface using a piezoelectric actuator. The membrane dimensions are limited by the area of the extruded hydrogel patch, leading to a plateau in the membrane capacitance as the HME advances into the droplet. Colors are added to the micrographs to improve clarity, with green denoting the PEG-DMA hydrogel and blue denoting the aqueous droplet. Membrane formation was then tested for a variety of oils and lipids as shown in Figure 3, testing membrane assembly and disassembly for each case. The ground electrode is placed on the opposite side of the membrane within a holding pipette to anchor the aqueous droplet, minimizing droplet motion. In each case the electrode tip begins just outside the droplet then is rapidly engulfed within the droplet using a square wave displacement supplied by the actuator, reaching a saturated membrane size dependent on the available area of the extruded hydrogel patch. After 10 seconds the HME is withdrawn from the droplet, separating the monolayer leaflets. Membrane area was compared to the extruded area of the patch by measuring the hydrogel area using the Leica LAS


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X software and multiplying by a standard specific capacitance for 50:50 hexadecane/silicone oil AR20 then comparing to the maximum membrane dimensions.

31, 32

The membrane area is dependent

on the size of the extruded hydrogel patch, which allows for further minimization as needed and provided membranes with areas ranging from 1/200 mm and 1/500 mm over the course of this th

2

th

2

study. Membrane formation lags slightly behind the HME introduction to the droplet interior. The two monolayers are forced into contact as the HME displacement forces the oil out from between the lipid leaflets. Once the membrane begins forming it expands rapidly forming a membrane across the entire hydrogel surface. This process continues to envelop the sharpened capillary, but the detectable membrane size is limited by the hydrogel area. The HME is coated by an insulating lipid membrane while inside the droplet which does not allow for mixing of the contents in the hydrogel and aqueous regions.

a.

b.

Hydrogel-Filled Micropipette

Piezoelectric Actuator

+

-

Oil

Hydrogel Microelectrode

Aqueous

Hydrogel

Aqueous

Hydrogel

DPhPC

DPhPC

DOPC

DOPC

Crate Substrate

c. 500 μm

500 μm

Figure 3 – a.) Membranes are formed using a HME and stationary anchored droplet. Colors are added to the micrographs to improve clarity, with green denoting the PEG-DMA hydrogel and blue denoting the aqueous droplet. The HME punctures the droplet, initiating membrane formation. b.) This approach was compatible with both saturated and unsaturated lipids, as well as c.) all oils traditionally used in DIB studies. The results shown here are for DPhPC.


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Results demonstrated that the HME is compatible with all standard oils used for DIBs as well as both saturated (1,2-diphytanoyl-sn-glycero-3-phosphocholine, DPhPC) and unsaturated (1,2dioleoyl-sn-glycero-3-phosphocholine, DOPC) lipids as shown in Figure 3. The rate of formation appears dependent on the viscosity of the oils and energy of adhesion of the membrane, exhibiting 31

a two-step formation process similar to previous experiments. Withdrawing the HME from the 33

droplet causes the membrane to rapidly disconnect. Transporter Reconstitution and Functionality To further ensure that these membranes were faithful recreations of lipid membranes, pore activities were measured using alamethicin (alm) as shown in Figure 4. Alm provides a voltagedependent pore within the membrane,

15, 34, 35, 36

gating and allowing for transport once the voltage

surpasses a threshold value. Alm was mixed with liposomes in the aqueous droplet and the HME was inserted into the droplet with a clamped potential of 100 mV to generate transient gating events as measured by jumps in the recorded current. Measurements of individual pore activations exhibiting multiple conductance levels

36, 37

were recorded. Measurements taken after puncturing

droplets containing alm with the HME then swapping to droplets without alm did not exhibit any gating or channel-related activity.

As long as the membrane does not rupture during the

measurements, the contents of the droplet and HME interior are separated by the insulating lipid membrane and minimal mixing occurs. This experiment demonstrates that the HME forms bilayer lipid membranes that are capable of incorporating biomolecular transporters. The reconstituted transporters demonstrate their expected functionalities, and the reduced membrane area allows for high-fidelity recordings with minimal capacitive noise.

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b. 5 seconds

100 mV

e

0.5 seconds d

a

b

e

1233 pS

d

474.3 pS

c

c

b

a

160.1 pS 85.96 pS

Figure 4 – Alamethicin provides a voltage-gated pore within the membrane. a.) Current recordings were taken with an applied potential of 100 mV, producing series of transient gating events at various conductivities. b.) A conductance histogram is constructed from the current trace, showing the multiple conductance levels and their frequency of occurrence. The histogram data is truncated on the x axis to limit the ~0 pS measurements where no gating occurred. Repeatability of Membrane Formation Repeated HME membrane formation was tested by rapidly moving the hydrogel tip from just beyond the perimeter of the droplet anchored to the holding pipette to being fully engulfed within the droplet and back out again at the maximum actuator velocity. Figure 5 shows the formation and separation of the membrane occurring repeatedly at a frequency of 1 Hz with a step function supplied to the piezoelectric actuator. This was combined with the 100 Hz, 10 mV triangle wave voltage signal applied between the electrodes to assess the membrane size using capacitive current. In each case, the membrane rapidly grew as the electrode pierced the droplet as measured by the increase in the capacitive current then diminished as the HME was retracted. Figure 5 shows the membrane formation as a function of the membrane capacitance compared to the position of the HME. This was repeatable several hundred times without membrane rupture. It is important to note that precise placement of the electrode tip was not necessary – the membrane was able to


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form and remain intact even when the tip was fully engulfed within the droplet (Figure 2), puncturing the surface at the maximum approach velocity offered by the piezoelectric actuator. A video of this process is provided as a supplementary file.

a.

b. 100 μm

100 μm

100 μm

Figure 5 – a.) Positioning the electrode just outside of the reservoir and applying a repeated step displacement to the electrode allows for the rapid assembly and disassembly of the membrane. b.) Membrane formation is depicted by an increase in the capacitance between the electrodes. This was repeatable for several hundred displacement cycles without membrane failure. Colors are added to the micrographs to improve clarity, with green denoting the cured PEG-DMA hydrogel and blue denoting the aqueous droplet.


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Characterizing Droplet Contents This ability to rapidly form and separate membranes with minimal risk of membrane failure or disruption may be applied towards characterizing membrane properties. This has been examined previously in DIB systems through electrowetting

31, 39, 40

and mechanoelectricity, and has been used

to study membrane adsorption and variation in leaflet properties.

33

41, 42

Here this is applied towards

quantifying the properties of the lipids or other amphiphilic molecules in the aqueous droplet. The HME provides a fixed maximum membrane size as illustrated by the maximum capacitance plateau in Figure 2.c, so membrane thinning under a DC voltage or electrocompression must be 32

used as an alternative to electrowetting

. The minimum capacitance in these cases corresponds

32, 39, 40

to the thickest membrane, or the case where there is no electric field across the membrane interior driving electrocompression generated by the membrane asymmetry. The HME allows for an asymmetric membrane composition with the contents of one monolayer provided from within the HME and the contents of other provided from within the aqueous droplet or sample. This allows for quantifying the properties of dissolved lipids or surfactants within the aqueous droplet through measurements of membrane asymmetric. This is tested here by dissolving various lipids within a collection of aqueous droplets and monitoring their compositions using the HME. The HME was hydrated with a DPhPC solution and then used to sequentially puncture droplets containing

ground

electrodes

and

either

1,2-di-O-phytanoyl-sn-glycero-phosphocholine

(DOPhPC) or DPhPC. A DC potential across the membrane was changed in a stepwise fashion from -150 mV to 150 mV after membrane formation and the equilibrium membrane capacitance was recorded for each potential. The minimum capacitance with respect to the potential corresponds to a case where the electrostatic asymmetry present in the membrane is compensated by the values at the boundary. For a symmetric DPhPC/DPhPC membrane this corresponds to 0 42


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mV, and for DPhPC/DOPhPC this corresponds to 135 mV

33, 43

due to differences in the lipid

structure. This demonstrates the ability to quantify membrane asymmetry using the HMEs. This 33

is accomplished by modifying the location of the lipids to produce asymmetric membranes, and may be used to probe lipid or other surface-active molecule properties in a sequential fashion. The hypothesized mechanics for membrane separation is that the leaflets rapidly detach as the HME is retracted from the droplet interior. Consequently, the leaflet composition on the HME surface should change minimally due to the slow rate of flip-flop in these synthetic systems without pores or defects. To test this hypothesis, the HME was moved sequentially from the DPhPC 40

droplet to the DOPhPC droplet, repeating the asymmetry measurements as described previously for each droplet. There was minimal drift in the measured electrostatic offset, confirming that the approach is suitable for screening the contents of several droplets given the rapid rate of membrane formation and minimal time necessary for the measurements. a.

2

1 Oil

Oil

DOPhPC

b.

3 Oil

DOPhPC

DOPhPC

c. Aqueous

Hydrogel

DOPhPC

DPhPC

+

-

DPhPC

Asymmetric Bilayer

DPhPC

DPhPC

+

+

-

-

Aqueous

Hydrogel

DPhPC

DPhPC

1

2

3

Symmetric Bilayer

Figure 6 – Representative capacitance traces for electrocompression measurements with symmetric and asymmetric membranes. a.) The HME is alternated between droplets containing dissolved DPhPC and DOPhPC lipids. b.) This creates symmetric and asymmetric membranes on the HME surface, which c.) is reflected in shifts in the minimum membrane capacitance as a


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function of DC potential. Tetradecane was used in this study as the surrounding oil to amplify the electrocompression of the membrane.

32, 44

Conclusions The proposed HME allows for the repeated formation of lipid bilayer membranes inspired by the DIB technique. This allows for the formation of the membrane-on-demand, assembling and disassembling the membrane as necessary. Electrodes were fabricated by extruding a patch of PEG-DMA hydrogel containing an embedded Ag/AgCl electrode within sharpened glass capillary. By dissolving lipids either within a surrounding oil reservoir or within the hydrogel, lipid monolayers self-assemble at the surface of the hydrogel patch. Puncturing a lipid-coated aqueous droplet with the lipid-coated hydrogel promotes the rapid formation of a lipid bilayer for analysis. Importantly, bilayer formation is largely independent of the HME position within the droplet, allowing for rapid formation of interfacial membranes with a predetermined size governed by the area of the exposed hydrogel. These membranes were used to quantify the presence of transport biomolecules and assess electrostatic asymmetry within the membrane as demonstrations of their quality. The resulting electrode allows for the rapid formation of lipid membranes on demand and allows for the detection of changes in lipid properties and interfacial characteristics through the creation of asymmetric membranes. Materials and Methods: Hydrogel Microelectrode (HME) Creation The HME consists of a pulled glass micropipette filled with a UV cured hydrogel surrounding the Ag/AgCl electrode embedded inside the micropipette (Figure 7). Ag/AgCl electrodes are commonly used for electrophysiological studies of lipid bilayers. They are created here by immersing silver wires (250 μm diameter, Goodfellow) in bleach for approximately 15 minutes


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before thoroughly rising them with deionized water. Several heat-pull protocols are developed for the micropipette puller (P-1000, Sutter instruments) to consistently fabricate short-taper shaped glass micropipettes (Figure 7) with inner diameters ranging from 10 μm to 30 μm from an initially flat tip borosilicate glass micropipette (OD 1 mm, ID 0.5 mm). Using a MicroFil (World Precision Instruments, 34 gauge, 67mm long), the fabricated glass micropipette is filled with the hydrogel solution. The hydrogel solution is prepared by dissolving PEG-DMA 1000 (Polyscience Inc.) at 40% (w/v) in a buffer solution (500 mM KCl,10 mM MOPS, pH = 7) with 0.5% (w/v) 2-Hydroxy4′-(2-hydroxyethoxy)-2-methylpropiophenone (Irgacure 2959, Sigma Aldrich). Once exposed to UV light (3 minutes, 1 W intensity, 365 nm source, Thorlabs), the hydrogel undergoes free-radical photopolymerization. During this process, crosslinked polymeric structures are induced

45, 46

and the

hydrogel gradually stiffens around the wire electrode. The hydrogel provides a conductive path from the electrode surface to the membrane at the extruded hydrogel surface and fastens the electrode within the capillary. After the curing step, the solidified hydrogel remains trapped inside the micropipette a few microns away from the tip (Figure 7). This is undesirable as this creates a pocket of insulating oil between the electrode and the droplet interior. The wire is manually pushed inside the micropipette to extrude the cured hydrogel into a miniature cylindrical tip out of the capillary aperture. The diameter of the exposed hydrogel corresponds to the inner-diameter of the pulled glass capillary with a height typically ranging from 10 to 40 μm. On the other end of the glass capillary, the extended silver wire is carefully twisted around the micropipette to further secure the hydrogel-electrode assembly. Because water-swollen hydrogels are prone to dehydration in air, the tip of the electrode is submerged in the oil phase. A crocodile clip is used to attach the coiled wire to the ground.


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b.

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1

c.

Pulled glass capillary Ag/AgCl electrode

2

Solidified hydrogel 250 μm

Figure 7 – a.) The electrodes are formed by sharpening a borosilicate capillary before shearing the tip to the desired dimension using a capillary cutting stone b.) The pulled capillary is then filled with the uncured hydrogel then a silver/silver-chloride wire is threaded into the gel. UV light is subsequently used to cure the gel in place before applying a gentle pressure to extrude a hydrogel patch at the tip (1 → 2). c.) The resulting HME may be manually transferred into a pipette holder to manually form on-demand synthetic lipid membranes in oil. Holding Pipettes Pipettes containing the cured hydrogel are also used to anchor aqueous droplets at their tip. A Bunsen burner is used to bend the capillary and the capillary is then filled with the same used in the HME while leaving a small offset at the tip of the pipette as shown in Figure 3. Before the curing step, an Ag/AgCl electrode is embedded in the holding pipette for measurements. During experiments, the holding pipette is submerged in oil and an aqueous droplet is deposited onto the hydrogel located at the tip of the pipette. This provides a fixed anchor for the droplet so it is immobile during the experiment. Lipid-Out Solutions


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The lipid-oil solutions used in most of the lipids-out experiments consists of a mixture of 1:1 hexadecane-silicone oil AR 20 (Sigma Aldrich) with DPhPC (Avanti Polar Lipids, Inc.). Modifications include the mixing of DPhPC lipids with other types of oils including squalene, mineral oil, hexadecane and tetradecane (Sigma Aldrich) or using DOPC (Avanti Polar Lipids, Inc) instead of DPhPC with the same 1:1 hexadecane-silicone oil AR 20. The lipid-oil concentration in all cases is 0.5 mg/mL. Lipid-In Solutions Buffer solutions were prepared from deionized water. 500 mM KCl and 10 mM MOPS was added to the buffer solutions to provide sufficient conductivity for electrical measurements. Lipidaqueous solutions were prepared by dissolving powdered lipids DPhPC (Avanti Polar Lipids Inc.) or DOPhPC (Avanti Polar Lipids Inc.) in the buffer solution at a concentration of 2 mg/mL, then completing five freeze-thaw cycles. The resulting solutions were then extruded 10 times through a filtration block with a 0.01 µm polycarbonate filter (Whatman). This is followed by a bath sonication step for 1 hour or until the solutions were clear. Lipid-hydrogel solutions were obtained by mixing the sonicated lipid-aqueous solutions with the hydrogel solution at a concentration of 2 mg/mL. Both lipid-aqueous and lipid-hydrogel solutions were stored at 4 C until needed. o

Alamethicin Solutions A solution containing alm peptides from the fungus Trichoderma viridae (A.G. Scientific) with a concentration of 100 ng/mL is obtained by diluting a small fraction of a reserved stock (5 mg/mL, stored at −20 °C in ethanol) in a lipid-aqueous solution (2 mg/mL DPhPC, 500 mM KCL,10 mM MOPS, pH = 7). Setup and Electrophysiology


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An Axopatch 200B patch clamp amplifier (Molecular Devices) connected to a Digidata 1440 DAQ (Molecular Devices) is used to collect electrical measurement the membrane level with a sampling frequency of 20 kHz with a 1 kHz low-pass Bessel filter. The recorded data were further analyzed using MATLAB scripts that convert the signals into measurements of membrane properties such as capacitance. The nominal capacitance of the bilayers is evaluated by sending a triangular voltage at a frequency of 100 Hz and peak-to-peak amplitude of 20 mV using a function generator (33120A, Hewlett-Packard) and recording the current response. Alm gating is recorded with a DC potential of 100 mV. Membrane asymmetry is assessed using an accumulating step change in a DC offset (increasing ~13 mV every 30 seconds) provided by a voltage subroutine provided by a compact DAQ (cDAQ-9133, National Instruments) and Labview and combined with the same AC signal used for capacitance measurements. HME position is controlled using a piezoelectric flexure-guided linear actuator (P-601.3SL, Physik Instrumente). A digital controller (E-709.SRG, Physik Instrumente) coupled with a function generator (33220A, Agilent) are used to control the movement of the actuator. Displacement of the tip is recorded using the Digidata 1440 for simultaneously tracking the HME position and interfacial properties. The HME is connected to the actuator using a machined grooved plate. Acknowledgements The authors graciously acknowledge support from the NSF grant # 1537410 and Air Force Office of Scientific Research Basic Research Initiative Grant FA9550-12-1-0464.


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20. Sarles, S. A.; Stiltner, L. J.; Williams, C. B.; Leo, D. J. Bilayer Formation between LipidEncased Hydrogels Contained in Solid Substrates. Acs Applied Materials & Interfaces 2010, 2 (12), 3654-3663. 21. Sapra, K. T.; Bayley, H. Lipid-coated hydrogel shapes as components of electrical circuits and mechanical devices. Scientific Reports 2012, 2. 22. Bayoumi, M.; Bayley, H.; Maglia, G.; Sapra, K. T. Multi-compartment encapsulation of communicating droplets and droplet networks in hydrogel as a model for artificial cells. Scientific Reports 2017, 7, 45167. 23. Baxani, D. K.; Morgan, A. J. L.; Jamieson, W. D.; Allender, C. J.; Barrow, D. A.; Castell, O. K. Bilayer Networks within a Hydrogel Shell: A Robust Chassis for Artificial Cells and a Platform for Membrane Studies. Angew Chem Int Edit 2016, 55 (46), 14238-14243. 24. Wu, Z.; Wang, B.; Cheng, Z.; Yang, X.; Dong, S.; Wang, E. A facile approach to immobilize protein for biosensor: self-assembled supported bilayer lipid membranes on glassy carbon electrode. Biosensors and Bioelectronics 2001, 16 (1-2), 47-52. 25. Wu, Z.; Tang, J.; Cheng, Z.; Yang, X.; Wang, E. Ion channel behavior of supported bilayer lipid membranes on a glassy carbon electrode. Analytical chemistry 2000, 72 (24), 60306033. 26. Cornell, B. A.; Braach-Maksvytis, V.; King, L.; Osman, P.; Raguse, B.; Wieczorek, L.; Pace, R. A biosensor that uses ion-channel switches. Nature 1997, 387 (6633), 580. 27. Tang, J.; Wang, B.; Wu, Z.; Han, X.; Dong, S.; Wang, E. Lipid membrane immobilized horseradish peroxidase biosensor for amperometric determination of hydrogen peroxide. Biosensors and Bioelectronics 2003, 18 (7), 867-872. 28. Majd, S.; Yusko, E. C.; Billeh, Y. N.; Macrae, M. X.; Yang, J.; Mayer, M. Applications of biological pores in nanomedicine, sensing, and nanoelectronics. Current opinion in biotechnology 2010, 21 (4), 439-476. 29. Haque, F.; Li, J.; Wu, H.-C.; Liang, X.-J.; Guo, P. Solid-state and biological nanopore for real-time sensing of single chemical and sequencing of DNA. Nano today 2013, 8 (1), 56-74. 30. Richter, R. P.; Bérat, R.; Brisson, A. R. Formation of solid-supported lipid bilayers: an integrated view. Langmuir 2006, 22 (8), 3497-3505. 31. Taylor, G. J.; Venkatesan, G. A.; Collier, C. P.; Sarles, S. A. Direct in situ measurement of specific capacitance, monolayer tension, and bilayer tension in a droplet interface bilayer. Soft Matter 2015, 11 (38), 7592-605. 32. Gross, L. C.; Heron, A. J.; Baca, S. C.; Wallace, M. I. Determining membrane capacitance by dynamic control of droplet interface bilayer area. Langmuir 2011, 27 (23), 1433542. 33. Freeman, E. C.; Najem, J. S.; Sukharev, S.; Philen, M. K.; Leo, D. J. The mechanoelectrical response of droplet interface bilayer membranes. Soft Matter 2016, 12 (12), 3021-31. 34. Venkatesan, G. A.; Sarles, S. A. Droplet immobilization within a polymeric organogel improves lipid bilayer durability and portability. Lab on a Chip 2016, 16 (11), 2116-2125. 35. Challita, E. J.; Najem, J. S.; Monroe, R.; Leo, D. J.; Freeman, E. C. Encapsulating Networks of Droplet Interface Bilayers in a Thermoreversible Organogel. Scientific Reports 2018, 8 (1), 6494. 36. Taylor, G. J.; Sarles, S. A. Heating-Enabled Formation of Droplet Interface Bilayers Using Escherichia coli Total Lipid Extract. Langmuir 2015, 31 (1), 325-337.


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Table of Contents/Abstract Graphic

b

a

b

Monolayer

Hydrogel Hydrogel

Aqueous

Oil

Bilayer

a

a

Displacement

Hydro Hydrogel

25 pA

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gel

Aqueous

100 ms

b

Oil

a b

a 500 μm

500 μm

a

b


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Figure 1 – The proposed electrodes create hydrogel-supported lipid membranes as a lipid-infused gel contacts an aqueous reservoir coated with a lipid monolayer. Membrane formation is measured as a function of the capacitive current response to a 10 mV 100 Hz AC signal applied across the membrane. The membrane formation is rapid, reliable, and repeatable. 237x205mm (300 x 300 DPI)


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Figure 2 – a.) The bilayer is formed on the extruded hydrogel path at the tip of the HME and remains intact as the electrode is fully engulfed within the droplet as shown in b) the schematic. c.) A 125 m sinusoidal oscillation is applied to the HME at the droplet surface using a piezoelectric actuator. The membrane dimensions are limited by the area of the extruded hydrogel patch, leading to a plateau in the membrane capacitance as the HME advances into the droplet. Colors are added to the micrographs to improve clarity, with green denoting the PEG-DMA hydrogel and blue denoting the aqueous droplet. 623x275mm (300 x 300 DPI)


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Figure 3 – a.) Membranes are formed using a HME and stationary anchored droplet. Colors are added to the micrographs to improve clarity, with green denoting the PEG-DMA hydrogel and blue denoting the aqueous droplet. The HME punctures the droplet, initiating membrane formation. b.) This approach was compatible with both saturated and unsaturated lipids, as well as c.) all oils traditionally used in DIB studies. The results shown here are for DPhPC. 629x224mm (300 x 300 DPI)


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Figure 4 – Alamethicin provides a voltage-gated pore within the membrane. a.) Current recordings were taken with an applied potential of 100 mV, producing series of transient gating events at various conductivities. b.) A conductance histogram is constructed from the current trace, showing the multiple conductance levels and their frequency of occurrence. The histogram data is truncated on the x axis to limit the ~0 pS measurements where no gating occurred. 395x134mm (300 x 300 DPI)


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Figure 5 – a.) Positioning the electrode just outside of the reservoir and applying a repeated step displacement to the electrode allows for the rapid assembly and disassembly of the membrane. b.) Membrane formation is depicted by an increase in the capacitance between the electrodes. This was repeatable for several hundred displacement cycles without membrane failure. Colors are added to the micrographs to improve clarity, with green denoting the cured PEG-DMA hydrogel and blue denoting the aqueous droplet. 338x277mm (300 x 300 DPI)


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Figure 6 – Representative capacitance traces for electrocompression measurements with symmetric and asymmetric membranes. a.) The HME is alternated between droplets containing dissolved DPhPC and DOPhPC lipids. b.) This creates symmetric and asymmetric membranes on the HME surface, which c.) is reflected in shifts in the minimum membrane capacitance as a function of DC potential. Tetradecane was used in this study as the surrounding oil to amplify the electrocompression of the membrane.32, 44 506x179mm (300 x 300 DPI)


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Figure 7 – a.) The electrodes are formed by sharpening a borosilicate capillary before shearing the tip to the desired dimension using a capillary cutting stone b.) The pulled capillary is then filled with the uncured hydrogel then a silver/silver-chloride wire is threaded into the gel. UV light is subsequently used to cure the gel in place before applying a gentle pressure to extrude a hydrogel patch at the tip (1 → 2). c.) The resulting HME may be manually transferred into a pipette holder to manually form on-demand synthetic lipid membranes in oil. 97x51mm (300 x 300 DPI)


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ToC image 458x197mm (300 x 300 DPI)


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Hydrogel Microelectrodes for the Rapid, Reliable, and Repeatable

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