Formation of pore-spanning lipid membrane and cross-membrane water and ion transport

Author’s Accepted Manuscript Formation of Pore-Spanning Lipid Membrane and Cross-Membrane Water and Ion Transport Rona Ronen, Yair Kaufman, Viatcheslav Freger

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S0376-7388(16)30610-X http://dx.doi.org/10.1016/j.memsci.2016.09.059 MEMSCI14780

To appear in: Journal of Membrane Science Received date: 7 June 2016 Revised date: 21 September 2016 Accepted date: 28 September 2016 Cite this article as: Rona Ronen, Yair Kaufman and Viatcheslav Freger, Formation of Pore-Spanning Lipid Membrane and Cross-Membrane Water and Ion Transport, Journal of Membrane Science, http://dx.doi.org/10.1016/j.memsci.2016.09.059 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting galley proof before it is published in its final citable form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Formation of Pore-Spanning Lipid Membrane and CrossMembrane Water and Ion Transport Rona Ronena,b, Yair Kaufmana,b and Viatcheslav Fregera,b* a

Department of Chemical Engineering, Technion - Israel Institute of Technology, 32000 Haifa, Israel.

b

Zuckerberg Institute for Water Research, The Jacob Blaustein Institutes for Desert Research, Ben-Gurion University of the Negev, Sede Boqer Campus 84990, Midreshet Ben-Gurion, Israel.

ABSTRACT: Supported lipid bilayers have the potential to deliver a breakthrough in separation processes, e.g., in desalination. Yet, formation of macroscopically large lipid membranes for use in separations and understanding of their long-term mechanical stability, especially, when they host membrane proteins, is still a challenge. Arrays of microscopic

pore-spanning

lipid

membranes

are

a

promising

upscalable

configuration, in which lipid membranes do not contact support directly, making it attractive for studying their function and stability.

Here we report on (1) the

formation mechanism of an array of pore-spanning phospholipid membranes via ‘vesicle fusion’, and (2) a microfluidic device that is used to assess the stability of the pore-spanning lipid membrane under flow and osmotic gradient. It is shown that the formation of pore-spanning lipid membranes via ‘vesicles fusion’ proceeds in three steps: First, small vesicles merge into giant ones of about the size of the substrate pore size. The giant vesicles then settle at the pore mouths and flatten. Last, the flattened giant vesicles rupture and form a lipid membrane that closes the pore. Exposing the 1

membrane to combined osmotic and shear flows in a microfluidic device, which simulates common osmotic process conditions, shows that, in addition to pores remaining open, a fraction of pore-spanning membranes ruptures. Possible ways to avoid such rupture and minimize fraction of open pores are discussed.

Fluidic setup containing a biomimetic membrane KEYWORDS: self-assembly · pore-coverage · pore-spanning lipid membrane · separation · vesicle fusion

1.

INTRODUCTION

Biological membranes in their native structure, i.e., vesicles/liposomes, exhibit excellent salt rejection (~99.999%), with up to 10-1 cm/s water permeability [1], which motivates development of their biomimetic analogues [2]. Biomimetic membranes, composed of a lipid-like matrix with incorporated aquaporins (waterchannel membrane proteins) or their biomimetic counterparts, have the potential to become an energetically efficient alternative to current commercial reverse osmosis and nanofiltration membranes for separation applications [3].

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Recent studies on biomimetic membranes have focused on constructing membranes incorporating ion- or water-channels, e.g., aquaporin [4], usually supported by a water-permeable substrate made of dense [5] or porous [6-8] materials. Protein channels are usually incorporated to the lipid or lipid-like membrane, using the 'vesicle fusion' technique. In this method, proteins are first incorporated in a lipid vesicle to form proteoliposome, which under certain conditions can adhere to a substrate, rupture and form a supported planar biomimetic membrane [9]. Vesicle fusion is carried out in a surfactant solution, which prevents the exposure of the proteins to denaturing media [10], in contrast to alternative technique such as, Langmuir-Blodgett [11], Langmuir-Schaefer, and lipid painting [12].

Ruptured

vesicles retain the bilayer structure on hydrophilic surfaces such as Si oxide [13] or mica [5, 14, 15]. Unfortunately, the adhesion of the bilayer to hydrophilic surfaces is relatively weak, hence in the absence of lipids in the bulk solution, lipids can desorb from the surface and hinder separation applications. In contrast, hydrophobic (e.g., thiolated) surfaces strongly attract phospholipids through hydrophobic interactions, but promote formation of a monolayer, with hydrophobic lipid tails facing the surface and hydrophilic heads facing water. Nevertheless, when the hydrophobic substrate has open pores, the monolayer extends into the pores as a pore-spanning bilayer that encounters water (hydrophilic environment) on both sides with tails hidden inside. Compared to “regular” planar bilayers supported on permeable hydrophilic substrate, “pore-spanning” configuration [16-18] may have many potential advantages. For instances, bilayer membrane in each pore remains microscopically small and free-standing, contacting the substrate only at their rim. As such, they offer straightforward upscaling and minimal interferences by the substrate.

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The amount of defects in supported lipid membranes of regular planar configuration and their effect on permeability and, especially, selectivity remains an open question and a challenging problem. The relatively high water permeability measured for supported biomimetic membranes, as compared to that of vesicles made of the same lipids, may suggest presence of defects [5, 19-23]. For instance, the measured permeability of 1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC) lipid bilayer in vesicles is 10-3-10-5 cm/s [24], whereas for a biomimetic DMPC membrane supported on a dense polymeric substrate is about 10-1 cm/s[19]. This becomes even more complicated with insertion of channel proteins, which can also induce defects in the lipid membrane, thus permeability may be increased by transport through both channels and defects, not easily distinguished. In addition, a direct contact of proteins with a dense substrate or reinforcing elements often added to supported biomimetic membranes [25] may interfere with its functioning as a selective channel [26]. Many of these uncertainties could be avoided in the pore-spanning configuration, in which the functional part of a supported biomimetic membrane is essentially free-standing, and protein-support and lipid-support contacts are completely avoided. In this report, we explore pore-spanning lipid membranes as a platform for osmotic separations and studies of lipid membrane formation, functioning and stability. This study does not include membrane proteins, however, incorporation of membranes proteins that impart biomimetic membranes with unique functionalities is the main motivation and will be a natural extension for future work. The fact that porespanning membranes behave essentially as free-standing will also help eliminate interferences by the support that have obscured previous results on supported bilayers [27]. However, we deemed it necessary to clarify first the preparation and stability

4

issues using a simple lipid-only system, which is the main purpose of the present study. First, we describe the formation mechanism of a pore-spanning lipid membrane from vesicles that are several times smaller than the substrate pores. Subsequently, we evaluate the percentage of substrate pores that are not covered by lipids (i.e., ‘open’), pores that are covered by pore-spanning lipid membrane, and pores in some transient state. Last, we look into the effect of shear and osmotic flow, commonly encountered in membrane separation processes, on the stability and integrity of the lipid membrane.

2.

EXPERIMENTAL SECTION

2.1.

Materials

1,2-dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1,2-dimyristoyl-sn-glycerophosphoethanolamine-N-(Lissamine-Rhodamine B Sulfonyl) (Ammonium salt), referred to as Rh-PE, were purchased from Avanti Polar Lipids (Alabaster, AL). Texas

Red

conjugated

1,2-Dihexadecanoyl-sn-Glycero-3-Phosphoethanolamine

(DHPE) lipid, referred to as TR, was purchased from Invitrogen (Karlsruhe, Germany). All solutions were prepared from Milli-Q water (>18.2 MΩ cm-1). Vesicle suspending media was phosphate buffer saline (PBS, 10 mM Na2HPO4, 2mM KH2PO4, 137 mM NaCl and 2.7 mM KCl, pH 7.4).

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2.2.

Substrate Preparation

Nuclepore Polycarbonate Track-Etched (PCTE) (Whatman, Kent, England) substrates of 25 mm diameter, 10 µm thickness and 800 nm nominal pore diameter were used to support the membrane (see Supporting Information, Figure S1). Support was rendered conductive by physical vapor deposition (PVD) of a gold layer. Prior to deposition the membranes were rinsed with solvents (methanol, acetone, isopropanol in sequence), washed with double distilled water and dry etched with O2 plasma (Axis Asher) at 200mTorr at 100W for 2 min to strip organic compounds. The substrate was coated with a 10 nm thick adhesive Titanium layer and then a 50 nm gold layer generated at a rate 1 Å/s in a PVD chamber (FC-1800, BOC coating technology, Airco Temescal) using electron beam evaporation. The coating process was conducted in a clean room. Coated substrates were kept in a desiccator until use.

2.3.

Gold Surface Thiolation

Gold-coated substrates were rinsed in ethanol, dried and cleaned in a UV-ozone cleaner (Jelight, model 42-220) for 5 min. A self-assembled monolayer (SAM) was formed on top of gold coat by immersing gold-coated substrate in 1 mM 1-octanethiol (OT, Sigma-Aldrich) solution overnight. Samples were then thoroughly rinsed with ethanol and blown dry with nitrogen before use [28].

2.4.

Lipid Membrane Formation by Vesicle Fusion.

DMPC dry lipid powder was hydrated in PBS for 30 minutes to yield 1.5 mM DMPC solution. DMPC vesicles were extruded 21 times through a 100 nm filter at 40 °C. The average vesicle diameter reduced from >103 nm (wide size distribution) to about 120 nm (polydispersity index 0.045), as examined by dynamic light scattering

6

(Zetasizer, Nano, Malvern). Fluorescent samples were produced by adding fluorescent-labeled lipids Rh-PE or TR (1 mol% relative to the total lipid) during lipid dissolution in chloroform. Pore-spanning lipid membranes were prepared by exposure of OT-gold surface to appropriate vesicle solution for a prescribed time followed by rinsing the excess vesicles with a lipid-free buffer.

2.5.

Atomic Force Microscopy (AFM)

Imaging measurements were performed with Innova AFM (Bruker, Santa Barbara, CA) equipped with a 100-micron scanner. Images were acquired at ambient conditions (temperature 23oC) in the tapping mode with target amplitude 3V and set point ratio around 0.7 in liquid at 512×512 pixels resolution. Cantilevers with nominal spring constants 0.35 N/m were used (DNP type, Bruker, Santa Barbara, CA).

2.6.

Electrochemical Measurements

Lipid membrane formation was monitored using electrochemical impedance spectroscopy (EIS) and cyclic voltammetry (CV) in a custom-designed cell, as shown in Abstract Figure. EIS and CV were performed using a BioLogic SP-300 potenitiostat/impedance analyzer (Bio-Logic Science Instruments) equipped with ECLab software v.10.36. In CV the potential of the gold coated membrane serving as a working electrode versus an Ag/AgCl reference electrode was linearly ramped from 0.2 to 0.6 V and back using a platinum wire as the counter electrode. The electrolyte solution contained 1 mM Fe(CN)63- and 1mM Fe(CN)64- in 100 mM NaCl supporting electrolyte. EIS results were recorded using the same setup in a frequency range 10 -2 to 105 Hz, 6 points per decade, at AC amplitude 25 mV.

7

2.7.

Fluorescence Microscopy

Thiolated substrates were incubated with DMPC vesicles containing a fluorescent lipid marker (Rh-PE or TR) in PBS for 4 hrs. Excess vesicles were rinsed prior to measurement.

Confocal

images

were

acquired

on

an

inverted

confocal

LSM 700 microscope (Zeiss, Oberkocahen, Germany) using an EC Plan-Neofluar 40×/1.30 oil-immersed objective (x400 magnification). A green laser (555 nm) with laser power of 2.6%, PMT gain of 750 and pinhole of 2 Airy units was used for excitation. The fluorescence emission was collected with a long-pass filter of 585 nm. In fluorescence recovery after photobleaching (FRAP) experiments, a spot of 25 µm diameter was bleached at 100% laser power for 20 iterations with pixel dwell of 12.6 µsec. Series of confocal images were recorded immediately after the bleaching process within 5 sec intervals for at least 2.5 min. The FRAP results of Rhodamine-PE were fitted to the model of a single-mode diffusion-dominated recovery [29]. For a circular bleach spot, the following closedform solution involving modified Bessel functions [30] was used:

f  t   exp( D / 2t )  I 0 ( D / 2t )  I1 ( D / 2t  ,

(1)

where t is the elapsed time since the sample was bleached, f  t   F (t ) / F is the normalized fluorescence intensity, F (t ) is the average fluorescence intensity of the bleached area right after the sample was bleached (i.e., at t = 0), and F is the average fluorescence intensity after recovery when it saturates back at sufficiently long time after bleaching. Note that F is not necessarily identical to intensity before bleaching, if other unobserved recovery modes, much slower than diffusion, are involved (e.g., 8

flip-flop between the upper and lower leaflets). I0 and I1 are the modified Bessel functions. The diffusion coefficient, Df, was calculated using the relation Df = 2 / D , where  is the radius of the bleached area.

2.8.

Flow Experiment

The experiments were performed after lipid membrane formation to estimate its permeability and pore coverage. The formation process was monitored using EIS during substrate incubation in vesicle solution for two different times, 4.5 and 8 hrs. Osmotic flow between the feed and draw solutions pumped through respective compartments separated by the membrane, and electrochemical measurements were performed simultaneously in a 2-compartment microfluidic cell. The draw solution of 100 mM NaCl was pumped through to the upper compartment (draw) and doubledistilled water through the lower compartment (feed, Figure 3). The draw and feed compartments were of identical dimensions and both solutions were pumped at the same inlet flow rate of 44 µL/min using two identical 8 mL stainless steel syringes (Harvard-Warner) driven by a dual-syringe pump Fusion 400 Touch (Chemyx Inc., Stafford, Texas) in order to eliminate pressure difference between compartments and minimize effect of flow rate fluctuations. The conductivities of both solutions were measured both at inlet and outlet. The outgoing solutions were collected and weighed every 15 min.

9

Figure 1. The proposed mechanism of a biomimetic membrane formation by vesicle fusion on a porous substrate consists of three steps: (A) Step 1: large vesicles (smaller than the substrate pore) fuse and form giant vesicles with a size commensurate with to the size of substrate pores. (B) Step 2: the giant vesicles adhere to the pore mouth, deform and form a disk-like structure with high curvature at the contact with the pore edge. Step 2 typically lasts 30 minutes after the porous substrate is exposed to a vesicle solution. (C) Step 3: the disk-like vesicles rupture and form a pore-spanning lipid bilayer. A non-completed process of porespanning membrane formation, transient states, and some open pores were also observed. After 4.5 hrs of incubation: 37.5% of the pores are covered by a bilayer, 42.5% of the pores are in transient state, and 20% of the pores are open. Addition of parallel and osmotic flow leads to coverage reduction of 12.5%. (D) After 8 hrs of incubation 60% of the pores are covered with a lipid bilayer, stable under osmotic flow. This mechanism is supported by AFM micrographs (E-G) and cross-section of the pores (E1-G1).

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3.

RESULTS AND DISCUSSION

3.1.

Formation Mechanism of Pore-Spanning Lipid Membranes

The formation of pore-spanning lipid membranes via fusion of vesicles was first examined by AFM. As schematically depicted in Figure 1 A to D, the process proceeds through three distinct steps. The thiolated gold coated polycarbonate tracketched (PCTE) substrate, prior to vesicle exposure, show pores of diameter 0.85 ± 0.26 µm (Figure 1 E). The conical profile, 400-500 nm deep, seen in Figure 1 E1, is a convolution of the pore shape with the pyramidal AFM tip. At step 1, the surface was exposed to extruded DMPC vesicles of fairly uniform initial size ~120 nm and polydispersity index 0.045, as measured by dynamic light scattering. At step 2, these vesicles both aggregate and fuse to form giant vesicles (800 nm or more) that block the pore mouths (Figure 1 B, F and F1). Other vesicles fuse on the surface and form a hybrid lipid membrane, consisting of a lipid monolayer over the hydrophobic OTcapped gold surface (see Supporting Information, section 2). The vesicle fusion in the bulk is possible due to the low zeta potential (-4.06 ± 1.95 mV) of the vesicles [31] [32] (thus weak double layer repulsion between the vesicles) and their high curvature after extrusion, which is unfavorable, given the packing parameter of DMPC lipids that is close to 1 [33]. At step 3, the giant vesicles that block the pores’ mouths (all other vesicles rinsed away) flatten to a disk-like structure and eventually rupture at the contact line between the lipids composing the vesicle and the OT-capped hydrophobic monolayer (Figure 1 B), resulting in a pore-spanning membrane (Figure 1 C). The uniform pore size of the PCTE membrane dictates uniform vesicle size on the surface. The entire multi-step pore-closure process apparently has a characteristic time of a few hours, which is also confirmed by in situ electrochemical impedance measurements (see Section 3.3). However, not all pores reached this final state, as 11

transient states of bilayer formation and open pores were also observed (Figure 1 C, G). A cross-section of AFM micrograph (Figure 1 G1) showed that the bilayer was sunken about 40 nm below the surface plane. Mey et al. [34] reported a similar ‘sinking’ of a lipid bilayer spanning similar pores. Note that this is unlikely to be an AFM artifact, as this would result in a different curved topography, deeper at the center of the pore. A possible reason is that the lower leaflet of the lipid bilayer, curving near the rim of the pore, might pull the pore-spanning lipid bilayer into the cavity, as illustrated in Figure 1 C (the leftmost pore) [17]. Indeed, the gold of the PCTE membrane coating produced via vapor deposition process and subsequently converted to a thiolated surface coats the inner pore surface to some depth, about 1020 nm, in addition to the 60 nm of the total coating thickness above the initial PCTE surface. The thiolated surface then extends about ~70 nm into the pores, as indicted by SEM images (supporting information Figure S3), and gets coated by a monolayer. Balancing the tensions of two parts of the monolayer extending into the pore as upper and lower leaves of the pore-spanning bilayer requires that it sinks somewhat into the pore, as indeed observed.

3.2.

Ex situ Estimation of Pore Coverage Without Shear Forces

Fluorescence microscopy was used to verify the presence of a thin contiguous lipid membrane and estimate the percentage of pores that are covered by a pore-spanning lipid membrane without shear flow and osmotic pressure. Micrographs of lipid membranes, probed with a fluorescent-labeled lipid Rhodamine-PE, show that on average 80% of the pores (based on 7 samples) were covered with a lipid membrane (covered by pore-spanning lipid bilayer or by giant vesicle(s)) in comparison to the bare substrate (see Figure 2 A and Supporting Information, Figure S1). 12

Fluorescence recovery after photobleaching (FRAP) technique was used to verify the fluorescence came from a contiguous planar membrane, in which case the lipid should have a lateral diffusivity of about D ~ 10-8 cm2/s [27, 35, 36], as compared to the much lower ~10-11 cm2/s diffusivity of vesicles [37]. Such a large difference comes from the fact that the rapid lipid diffusion within a vesicle is limited to the vesicle size (<0.8 m in the present case), while diffusion over larger has to involve much slower diffusion of whole vesicles or lipid exchange with solution. Therefore, for the 20 m photobleached spot, the primary recovery is associated with a contiguous lipid layer, in which there is no no such limitation on diffusion distance, and recovery of lipids that belong to “bleached” vesicles is not observable. The measured lateral diffusion coefficient, D = 3.9±0.5×10-8 cm2/s, deduced from the recovery kinetics (Figure 2 B2), is much closer to the former. Figure 2 B2 also shows that the recovery was 90%, which indicates that most fluorescent lipid was found in a contiguous lipid membrane and not vesicles. This diffusion coefficient is valid for the entire area of the hybrid lipid membrane including both the pore-spanning membranes and the monolayer over the thiolated solid substrate (85% of the substrate). Given that the ratio between coefficients of suspended bilayer and supported monolayer is approximatly 1:5 [27, 35, 36], the pores are only 15% of the area, and the bleached area contains many (~100) pores, the measured average effective diffusivity 3.9±0.5×10-8 cm2/s should be within the range of the values reported for supported bilayers (1.32-4.6 μm2/s [27, 36]), and significantly different from that for unruptured vesicles. The result then confirms the formation of hybrid membranes over most of the surface.

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The morphology of the hybrid membrane was also studied using a different fluorescent lipid, possessing a Texas Red (TR) head-group, which allowed distinguishing between different states of pore closure [38]. At distances approximately below 15 nm from the gold surface, TR fluorescent head-groups undergo long range dipole-dipole coupling with gold via Förster resonance energy transfer mechanism, which quenches the fluorescent head-group [39]. The black, nonfluorescing background, over the solid surface in Figure 2 C, which does fluoresces in the case of Rhodamine-PE (Figure 2 B), indicates the lipid was only present as a thin membrane (i.e., a layer of thickness less than 15 nm) [40]. Yet, TR does fluoresce while it is within the pore-spanning membranes and in unruptured vesicles hanging over the pores (Figure 2 C1). Only the pore-spanning lipid membranes can show recovery in FRAP. The FRAP kinetics for TR showed the degree of recovery was low, 15±4% based on 7 experiments (Figure 2 C2). Due to slow flip-flop between the leaflets in the pore-spanning membrane (days) [41], only the upper leaflet, an extension of the monolayer part of the hybrid membrane, could participate in the fluorescence recovery [23], while the lipids in the lower leaflet and unruptued vesicles could not. Time dependent analysis of the TR micrograph (Figure 2 D) revealed that at least 20% of the fluorescent spots, as shown in Figure 2 C, are moving objects, i.e., vesicles weakly attached to the surface. If the remaining 80% of the objects in Figure 2 C are pore-spanning membranes, only the upper leaflet i.e, about half or 40% of intensity prior to bleaching, should recover, however, only about one third of this recovery, ~15%, was observed. This discrepancy presumably results from pores that were in a transient state covered with unruptured vesicles. The results then suggest that 4.5 hrs of vesicle incubation were insufficient to form a defect-free membrane 14

over all pores, as is also evident from AFM results (Figure 1 C and G1) and only about one third of the pores were closed with fully formed pore-spanning membranes.

Figure 2. After 4.5 hrs of incubation excess vesicles were rinsed off and fluorescent microscopy was applied. (A) SEM image of a bare track-etched polycarbonate membrane (substrate). (B) Fluorescent micrograph of Rhodamine-PE labeled lipid membrane showing 20% uncovered pores (black spots). (B1) Schematics of a covered pore, where the red stars represent the fluorescent labeled lipid. (B2) Fluorescent recovery after photobleaching (FRAP) of a small circular area (25 µm diameter). Micrographs below show the area in three time frames. (C) Fluorescent micrograph of Texas Red (TR) labeled lipid membrane showing circular objects, i.e., covered pores or vesicles (red spots), where the black background is the covered substrate. (C1) TR quenches in proximity to the gold layer (distance between the TR and the gold layer is smaller than 15 nm). Quenched TR is illustrated as black star. (C2) FRAP experiment of TR sample in different time frames. (D) Spot-analysis of micrograph (C) that allows to differentiate between objects that moved more than 1 µm in 2 min (blue), stationary objects up to 800 nm diameter (yellow) and stationary objects above 800 nm (red).

3.3.

In situ Estimation of Pore Coverage Without Shear Forces

A microfluidic cell (Figure 3) allowed monitoring the membrane formation over the surface using non-intrusive electrochemical techniques with or without concurrent tangential or trans-membrane osmotic flows. The cell was an open system used to measure the macroscopic water and salt mass transport across the membrane, by 15

diffusion and convection, e.g., in osmosis between two flowing solutions. The porous substrate, separating two flow compartments, had a surface area 0.2 cm2, 4 orders of magnitude larger than the one tested in FRAP and AFM experiments. Electrochemical impedance spectroscopy (EIS) and cyclic voltammetry (CV), were used to monitor the lipid surface coverage without flow. The electrode configuration in Figure 3 consisted of a bare or thiolated Au layer on top of the substrate, acting as a working electrode, and a Pt counter-electrode and Ag/AgCl reference electrode in the upper (draw) compartment. This configuration was preferred over the one, in which working and counter-electrode are placed in different compartments. Although the current in the latter arrangement has to cross the pore-spanning membranes and measured impedance may seem to be a direct indication of the pore closure, it will be also strongly affected by the very large resistances – both diffusive and electrical - of the supporting membrane and solution in the lower compartment, which are orders of magnitude thicker than the lipid layer, and thus fairly insensitive to the presence of lipid membranes. In contrast, the Au electrode layer coated with a lipid membrane, without such intermediate resistance, and thus may be more sensitive to the lipid coating, even though the contribution of the pore-spanning membranes to the impedance is only fractional, through lateral current collected at the edges of Au coating at pore rims or its part extended into the pore. In addition, electrochemical measurements are obviously very sensitive since to the coverage of the solid part of the substrate, which is prerequisite to pore coverage. Therefore, changes of impedance or CV current associated with the substrate coverage provides an important indication of how the lipid layer formation advances towards formation of a contiguous lipid layer and, eventually, its pore-spanning part.

16

Figure 3. Schematic illustration of the microfluidic setup used for electrochemical measurements of membrane formation and estimation of pore coverage in parallel and osmotic flow conditions. The height of the draw and feed compartments was identical and they were fed form two identical syringes driven by the same dual-syringe pump to minimize the effect of flow rate fluctuations and pressure difference between compartments.

Figure 4A shows the results of electrochemical impedance spectroscopy (EIS) measurements recorded in situ during the formation of a lipid membrane on the OTgold surface (working electrode) from vesicles dispersed in phosphate buffer in the draw compartment, whereas the other compartment contained phosphate buffer free of vesicles. The EIS spectrum was analyzed using an equivalent circuit, shown in the inset of Figure 4 A, to extract the membrane resistance Rm, which is more sensitive to membrane defects than other elements [42], namely, Rs, the constant solution resistance, and Qm, a constant phase element accounting for non-ideal capacitance of the OT-monolayer, the lipid membrane and the double-layer [43]. Qm monotonically increases with the surface coverage, yet Rm increases more significantly (see Supporting Information, Section 4). The membrane resistance, Rm, plotted versus incubation time in Figure 4 A demonstrates that a steady-state resistance was reached within about 2 hrs, after which Rm remained constant. Subsequently, the overall surface coverage (substrate + pores) stabilized, as indicated by fluorescent 17

microscopy and EIS, subsequent subtle and slow changes in the structure of porespanning bilayers, barely detectable by these methods, could still profoundly change defect rate and transport properties of bilayers towards osmotic flow, for which stabilization could then take much longer.

This electrical pattern was well

reproducible in repeated experiments, however, the stabilized membrane resistance varied quite significantly, between 103 to 106 Ω cm2, which is nevertheless consistent with the range of values measured in other studies [18, 44].

Figure 4. Testing surface coverage by a lipid membrane without flow. (A) A plot of membrane ohmic resistance, Rm, versus incubation time. The resistance was extracted using the equivalent circuit. (B) Cyclic voltammograms of bare gold, OT-gold surface and lipid monolayer. Measurements were taken in 0.01 M K3/4[Fe(CN)6] and 0.1 M NaCl, recorded at 100 mV/s. Inset shows a larger potential scale that includes the bare gold voltammogram.

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Another method used to verify surface coverage, apparently more sensitive to presence of defect, was cyclic voltammetry (CV), in which redox-active hexacyanoferrate species were added to solution. The inset of Figure 4 B shows Fe(CN)63- reduction and Fe(CN)64- oxidation waves on bare gold electrode, which were dramatically suppressed when the bare gold electrode was functionalized with OT-monolayer. Once a DMPC lipid monolayer was added on top, a further strong suppression of the current was observed. A decrease in the redox current was caused by the increase in the charge transfer resistance. The current transfer across the lipid membrane mainly occurs via diffusion of Fe(CN)63-/4- species through defects and electron tunneling through the lipid layer, both being strongly reduced when another blocking lipid membrane is added. These CV voltammogram clearly indicate a lipid membrane was formed on top of OT-capped gold surface

3.4. In situ Estimation of Pore Coverage under Shear Forces and Osmotic Flow Having verified formation of a lipid membrane ex situ and in situ, osmotic flow experiments were conducted in the microfluidic cell by simultaneously pumping a concentrated salt solution into the upper compartment (draw) and double-distilled water (DDW) into the lower compartment (feed) at the same flow rate (44 µl/min, Figure 3), yielding an initial osmotic pressure difference of 248 kPa. Obviously, only a fraction of this difference was translated to the actual osmotic gradient across the lipid membrane due to the fast osmotic flow through the fraction of open pores (see below), as well as diffusional resistances of the supporting membranes and adjacent solution layers (concentration polarization). The salt transport through the membrane was assessed by measuring salt concentration at feed outlet, while repetitive EIS measurements ensured the membrane coverage remained stable for at least 2 hrs. 19

An ideal defect-free membrane, with all pores covered by pore-spanning membranes (100% pore coverage) should be almost impermeable to salt, as evident from the extremely low permeability of DMPC membrane to chloride (Pf ~ 10-10 m/s) [23], which would give a chloride permeation rate of ~ 10-10 g/s. This is far smaller than the salt permeation observed here, as deduced from the salt concentration at the feed outlet. Therefore, it was presumed that the salt transport through the pores that are covered with lipids was negligible compared to open pores. By relating salt passage through a substrate with pore-spanning lipid membranes to that before exposure to lipid vesicles, i.e., with all pores open, the apparent fraction of closed pores, ɸ, could be estimated, as follows:

  1  Cm

Co ,

(2)

where Cm and Co are the salt concentrations in the outlet of feed compartment for a substrate with and without pore-spanning membranes, respectively. The osmotic experiments yielded an average of 25% pore coverage after 4.5 hrs of incubation with DMPC vesicles for 6 experiments in osmotic flow conditions (Figure 1 C). Note that the difference between Cm and Co was significant, with a P-value 0.02 for two-tail ttest.

This  can be compared to 37.5% deduced from fluorescence results after

similar incubation time. The lower value for osmotic experiments could be caused by an extra stress exerted by the osmotic pressure and pressure fluctuations between the compartments that could rupture not fully developed pore-spanning membranes. For instance, a transient state where an unruptured vesicle already blocking the pore could eventually rupture under flow to leave the pore open rather than form a pore-spanning membrane.

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However, extending the incubation time to 8 hrs has increased the percentage of pores that remained covered under flow. This pore coverage was assessed by means of eq. 2 to 60% (Figure 1 D), based on the significant reduction in outgoing feed concentration from 2.8±0.8 mM to 1.1±0.3 mM. This highlights the fact that membrane design and preparation, e.g., vesicle incubation and sufficient time to complete formation of pore-spanning membrane, may be pivotal and increase the chance of membrane integrity under flow. The increase of pore coverage under flow with incubation time, i.e., with higher pore coverage before starting the flow, suggests that pore coverage with and without flow is correlated. In turn, this also suggests that other factor promoting formation of porespanning lipid membrane will increase the chance of having stable pore coverage under flow as well. Thus, it may be presumed that a pore-spanning membrane formed on a substrate with smaller pore radii, rp, is expected to be more robust. Importantly, for transport experiments, it should also be more mechanically stable and withstand higher transmembrane pressures. Indeed, the rupture pressure ΔPmax is given by ΔPmax ~ 2γ/rp, where γ ~4-12 mN/m is the rupture tension of lipid bilayers [45, 46]. In the present work relatively large 800 nm pores were used to facilitate optical microscopy observation, for which ΔPmax was of the order of 0.1 bar and, likely, too small to withstand the osmotic gradient used here. However, in future experiments, a higher pore-coverage could be possibly achieved by using smaller pores, such as 50-100 nm (rp = 25-50 nm), where the membrane can withstand ΔPmax of a few bars.

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CONCLUSIONS AND OUTLOOK In this paper, we shed light on the formation mechanism of an array of pore-spanning lipid membrane and on its mechanical stability under shear and osmotic flow conditions, as encountered relevant in foreseen applications and studies of lipid membrane permeability. The formation mechanism of pore-spanning lipid bilayer via ‘vesicle fusion’ of small vesicles (smaller than the substrate pore size) consists of three steps: First, the small vesicles fuse to giant vesicles (larger than the substrate pore size) in the bulk. Second, these giant vesicles then adhere to the pore mouth and flatten. These two steps occur simultaneously, and, for the studied system and conditions, within ~0.5 hrs most (but not all) of the substrate pores were covered by giant vesicles. In the last step, which can take more than 8 hrs, part of giant vesicles transform to a hybrid membrane including pore-spanning lipid bilayers and a monolayer over the solid substrate. Nevertheless, even after 8 hrs incubation, some of the substrate pores remained ‘open’, i.e., not covered by pore-spanning lipid bilayer nor by giant vesicles. The stability of the pore-spanning lipid membranes was tested by exposing them to tangential flow and transmembrane osmotic gradient. The results show that combined shear force and osmotic gradient rupture significant number of the lipid membranes that cover the pores. However, extending the vesicle incubation time reduces the number of open pores, as deduced from salt transfer; for example, after 8 hrs incubation it dropped by half, compared to 4 hrs incubation. This correlation between completion of the lipid membrane formation and its stability under flow suggests that benefits of increasing incubation time even further are not exhausted.

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In conclusion, the paper provides several useful insights into the formation mechanism of a pore-spanning lipid membrane on an array of pores via fusion of small (smaller than the pore size) vesicles and limitations and obstacles that still remain. It seems that the focus of future efforts may have to be on achieving improved coverage and stability and complete and robust formation of durable pore-spanning membranes. One such way may be through increasing the vesicle incubation time to over 8 hrs, as suggested by the present results. The kinetics, however, may be improved by actively promoting vesicle rupture, e.g., using ‘osmotic shock’ [47]. Another useful approach could be strengthening the interactions between lipids and substrate by tuning the lipid composition to produce a "stitching" effect or by optimizing the surface potential, as demonstrated in our previous work [10]. Finally, decreasing the size of the substrate pores may greatly accelerate membrane formation and increase durability. Note that, as demonstrated by Steinem et al. [34], monolayer tension exerted on the bilayer at pore rims may revert the trend at some size and make very small pore-spanning membrane more prone to spontaneous rupture. However, it seems that the 800 nm pore size used in this work and mainly selected to allow optical microscopy examination may be reduced by as large as an order of magnitude with significant benefits for kinetics and stability.

ACKNOWLEDGMENTS We thank Nitzan Dahan (LS&E infrastructure unit, Technion) for assistance with fluorescence microscopy and Dr. Arkady Bitler (Technion) for help with AFM imaging.

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REFERENCES AND NOTES [1] C. Maurel, Aquaporins and water permeability of plant membranes, Annual review of plant biology, 48 (1997) 399-429. [2] P. Kuchel, G. Ralston, Theory and Problems of Biochemistry. New York: Schaum's Outline Series, in, McGraw-Hill, 1988. [3] M. Kumar, Biomimetic membranes as new materials for applications in envrionmental engineering and biology, in, University of Illinois at UrbanaChampaign, 2010. [4] G. Sun, T.-S. Chung, N. Chen, X. Lu, Q. Zhao, Highly permeable aquaporinembedded biomimetic membranes featuring a magnetic-aided approach, RSC Advances, 3 (2013) 9178-9184. [5] Y. Kaufman, A. Berman, V. Freger, Supported lipid bilayer membranes for water purification by reverse osmosis, Langmuir, 26 (2010) 7388-7395. [6] H. Wang, T.S. Chung, Y.W. Tong, K. Jeyaseelan, A. Armugam, Z. Chen, M. Hong, W. Meier, Highly Permeable and Selective Pore‐ Spanning Biomimetic Membrane Embedded with Aquaporin Z, Small, 8 (2012) 1185-1190. [7] P.H. Duong, T.-S. Chung, K. Jeyaseelan, A. Armugam, Z. Chen, J. Yang, M. Hong, Planar biomimetic aquaporin-incorporated triblock copolymer membranes on porous alumina supports for nanofiltration, J. Membr. Sci., 409 (2012) 34-43. [8] G. Favero, A. D’Annibale, L. Campanella, R. Santucci, T. Ferri, Membrane supported bilayer lipid membranes array: preparation, stability and ion-channel insertion, Analytica Chimica Acta, 460 (2002) 23-34. [9] Y. Kaufman, V. Freger, Supported biomimetic membranes for pressure-driven water purification, INTECH Open Access Publisher, 2011. [10] R.P. Richter, R. Bérat, A.R. Brisson, Formation of solid-supported lipid bilayers: an integrated view, Langmuir, 22 (2006) 3497-3505. [11] M.S. Khan, N.S. Dosoky, J.D. Williams, Engineering Lipid Bilayer Membranes for Protein Studies, International journal of molecular sciences, 14 (2013) 2156121597. [12] S. Leptihn, O.K. Castell, B. Cronin, E.-H. Lee, L.C. Gross, D.P. Marshall, J.R. Thompson, M. Holden, M.I. Wallace, Constructing droplet interface bilayers from the contact of aqueous droplets in oil, Nature protocols, 8 (2013) 1048-1057. [13] Y. Kaufman, S. Grinberg, C. Linder, E. Heldman, J. Gilron, V. Freger, Fusion of bolaamphiphile micelles: a method to prepare stable supported biomimetic membranes, Langmuir, 29 (2013) 1152-1161. [14] R.P. Richter, A.R. Brisson, Following the formation of supported lipid bilayers on mica: a study combining AFM, QCM-D, and ellipsometry, Biophysical journal, 88 (2005) 3422-3433. [15] D.W. Lee, X. Banquy, K. Kristiansen, Y. Kaufman, J.M. Boggs, J.N. Israelachvili, Lipid domains control myelin basic protein adsorption and membrane interactions between model myelin lipid bilayers, Proceedings of the National Academy of Sciences, 111 (2014) E768-E775. [16] J. Drexler, C. Steinem, Pore-suspending lipid bilayers on porous alumina investigated by electrical impedance spectroscopy, J. Phys. Chem. B, 107 (2003) 11245-11254. [17] M. Kocun, T.D. Lazzara, C. Steinem, A. Janshoff, Preparation of solvent-free, pore-spanning lipid bilayers: modeling the low tension of plasma membranes, Langmuir, 27 (2011) 7672-7680.

24

[18] E.K. Schmitt, M. Nurnabi, R.J. Bushby, C. Steinem, Electrically insulating poresuspending membranes on highly ordered porous alumina obtained from vesicle spreading, Soft Matter, 4 (2008) 250-253. [19] Y. Kaufman, S. Grinberg, C. Linder, E. Heldman, J. Gilron, Y.-x. Shen, M. Kumar, R. Lammertink, V. Freger, Towards supported bolaamphiphile membranes for water filtration: Roles of lipid and substrate, J. Membr. Sci., 457 (2014) 50-61. [20] J.C. Mathai, S. Tristram-Nagle, J.F. Nagle, M.L. Zeidel, Structural determinants of water permeability through the lipid membrane, The Journal of general physiology, 131 (2008) 69-76. [21] Y. Graziani, A. Livne, Water permeability of bilayer lipid membranes: sterollipid interaction, The Journal of membrane biology, 7 (1972) 275-284. [22] A. Studer, X. Han, F.K. Winkler, L.X. Tiefenauer, Formation of individual protein channels in lipid bilayers suspended in nanopores, Colloids and Surfaces B: Biointerfaces, 73 (2009) 325-331. [23] S.S. Mansy, Membrane transport in primitive cells, Cold Spring Harbor perspectives in biology, 2 (2010) a002188. [24] M. Jansen, A. Blume, A comparative study of diffusive and osmotic water permeation across bilayers composed of phospholipids with different head groups and fatty acyl chains, Biophys. J., 68 (1995) 997-1008. [25] C.Y. Tang, Y. Zhao, R. Wang, C. Hélix-Nielsen, A.G. Fane, Desalination by biomimetic aquaporin membranes: Review of status and prospects, Desalination, 308 (2013) 34-40. [26] J.J. Gray, The interaction of proteins with solid surfaces, Current opinion in structural biology, 14 (2004) 110-115. [27] M.L. Wagner, L.K. Tamm, Tethered polymer-supported planar lipid bilayers for reconstitution of integral membrane proteins: silane-polyethyleneglycol-lipid as a cushion and covalent linker, Biophys. J., 79 (2000) 1400-1414. [28] Sigma-Aldrich, Molecular self-assembly, in, Material Matters, 2006. [29] D. Axelrod, D. Koppel, J. Schlessinger, E. Elson, W. Webb, Mobility measurement by analysis of fluorescence photobleaching recovery kinetics, Biophysical journal, 16 (1976) 1055. [30] D. Soumpasis, Theoretical analysis of fluorescence photobleaching recovery experiments, Biophysical journal, 41 (1983) 95-97. [31] B. Heurtault, P. Saulnier, B. Pech, J.-E. Proust, J.-P. Benoit, Physico-chemical stability of colloidal lipid particles, Biomaterials, 24 (2003) 4283-4300. [32] T. Deming, Peptide-Based Materials, Springer Science & Business Media, 2012. [33] J.N. Israelachvili, Intermolecular and surface forces: revised third edition, Academic press, 2011. [34] . Mey, M. Stephan, . . Schmitt, M.M. M ller, M. en Amar, C. Steinem, A. Janshoff, Local membrane mechanics of pore-spanning bilayers, J. Am. Chem. Soc., 131 (2009) 7031-7039. [35] T.D. Lazzara, T.-T. Kliesch, A. Janshoff, C. Steinem, Orthogonal functionalization of nanoporous substrates: Control of 3D surface functionality, ACS applied materials & interfaces, 3 (2011) 1068-1076. [36] A. Sonnleitner, G.J. Schütz, T. Schmidt, Free Brownian Motion of Individual Lipid Molecules in Biomembranes, Biophys. J., 77 (1999) 2638-2642. [37] Y. Sakuma, N. Urakami, Y. Ogata, M. Nagao, S. Komura, T. Kawakatsu, M. Imai, Diffusion of domains on nanometer sized vesicle, in: Journal of Physics: Conference Series, IOP Publishing, 2010, pp. 012036.

25

[38] I. Höfer, C. Steinem, A membrane fusion assay based on pore-spanning lipid bilayers, Soft Matter, 7 (2011) 1644-1647. [39] A.E. Oliver, E.L. Kendall, M.C. Howland, B. Sanii, A.P. Shreve, A.N. Parikh, Protecting, patterning, and scaffolding supported lipid membranes using carbohydrate glasses, Lab Chip, 8 (2008) 892-897. [40] K. Vasilev, W. Knoll, M. Kreiter, Fluorescence intensities of chromophores in front of a thin metal film, The Journal of chemical physics, 120 (2004) 3439-3445. [41] A.M. Brozell, S. Inaba, A.N. Parikh, Lipid bilayers on topochemically structured planar colloidal crystals: a versatile platform for optical recording of membranemediated ion transport, Soft Matter, 6 (2010) 5334-5341. [42] G. Wiegand, N. Arribas-Layton, H. Hillebrandt, E. Sackmann, P. Wagner, Electrical properties of supported lipid bilayer membranes, The Journal of Physical Chemistry B, 106 (2002) 4245-4254. [43] B. Hirschorn, M.E. Orazem, B. Tribollet, V. Vivier, I. Frateur, M. Musiani, Determination of effective capacitance and film thickness from constant-phaseelement parameters, Electrochim. Acta, 55 (2010) 6218-6227. [44] A. Carruthers, D. Melchior, Study of the relationship between bilayer water permeability and bilayer physical state, Biochemistry, 22 (1983) 5797-5807. [45] M. Bialecka-Fornal, H.J. Lee, H.A. DeBerg, C.S. Gandhi, R. Phillips, Single-cell census of mechanosensitive channels in living bacteria, PloS one, 7 (2012) e33077e33077. [46] K. Olbrich, W. Rawicz, D. Needham, E. Evans, Water permeability and mechanical strength of polyunsaturated lipid bilayers, Biophys. J., 79 (2000) 321-327. [47] E. Reimhult, F. Höök, B. Kasemo, Intact vesicle adsorption and supported biomembrane formation from vesicles in solution: influence of surface chemistry, vesicle size, temperature, and osmotic pressure, Langmuir, 19 (2003) 1681-1691.

Highlights    

Formation of pore-spanning lipid membrane on thiolated porous substrates via ‘vesicle fusion’ studied. Pore-coverage by a DMPC bilayer examined with and without flow and osmotic gradient. Microfluidic cell used simultaneously monitors membrane formation and osmotic flow. Membrane ability to block osmotic flow correlates with completion of membrane formation.

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