Effect of polyelectrolyte structure on formation of supported lipid bilayers on polyelectrolyte multilayers prepared using the layer-by-layer method

Journal Pre-proofs Effect of polyelectrolyte structure on formation of supported lipid bilayers on polyelectrolyte multilayers prepared using the layer-by-layer method Ataru Seimei, Daisuke Saeki, Hideto Matsuyama PII: DOI: Reference:

S0021-9797(20)30232-0 https://doi.org/10.1016/j.jcis.2020.02.079 YJCIS 26074

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Journal of Colloid and Interface Science

Received Date: Revised Date: Accepted Date:

16 October 2019 20 January 2020 19 February 2020

Please cite this article as: A. Seimei, D. Saeki, H. Matsuyama, Effect of polyelectrolyte structure on formation of supported lipid bilayers on polyelectrolyte multilayers prepared using the layer-by-layer method, Journal of Colloid and Interface Science (2020), doi: https://doi.org/10.1016/j.jcis.2020.02.079

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Effect of polyelectrolyte structure on formation of supported lipid bilayers on polyelectrolyte multilayers prepared using the layer-by-layer method Ataru Seimeia, Daisuke Saekib,c*, Hideto Matsuyamaa*. a

Research Center for Membrane and Film Technology, Department of Chemical Science and

Engineering, Kobe University, 1-1 Rokkodai, Nada, Kobe 657-8501, Japan b

Research Initiative for Supra-Materials (RISM), Interdisciplinary Cluster for Cutting Edge

Research, Shinshu University, 4-17-1 Wakasato, Nagano 380-8553, Japan c

Department of Materials Chemistry, Faculty of Engineering, Shinshu University, 4-17-1

Wakasato, Nagano 380-8553, Japan *Corresponding authors E-mail address: [email protected] (D. Saeki). Phone: +81-26-269-5395. E-mail address: [email protected] (H. Matsuyama). Phone & Fax: +81-78-803-6180.

Abbreviations1

1 AqpZ Aquaporin Z, BLM black lipid membrane, CLSM confocal laser scanning microscopy, FR fluorescence recovery ratio,

FRAP fluorescence recovery after photo-bleaching, LbL layer-by-layer, PDDA poly(diallyldimethylammonium chloride), PEI polyethyleneimine, PEM polyelectrolyte multilayer, POPC 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine, POPG 1-palmitol2-oleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (sodium salt), PSS poly(sodium 4-styrenesulfonate), PVAm polyvinylamidin, Rhod-PE 1,2-dioleoyl-sn-glycero-3-phosphophenolamine-N-(lissamine rhodamine B sulfonyl) (ammonium salt), SLB supported lipid bilayer

1

GRAPHICAL ABSTRACT

2

ABSTRACT

We investigated the formation behavior of supported lipid bilayers (SLBs) on polyelectrolyte multilayers (PEMs) prepared using the layer-by-layer method. The SLBs were formed using the liposome fusion method, which was driven by electrostatic interactions. We used three types of cationic polyelectrolytes to prepare PEMs: poly(diallyldimethylammonium chloride), which consists of a linear short polymer chain with quaternary ammonium cations, polyvinylamidin, which presents a linear, long polymer chain and polyethyleneimine which features a branched polymer chain. Poly(sodium 4-styrenesulfonate) was used as an anionic polyelectrolyte. First, we evaluated the effect of the molecular structure of the polyelectrolytes on the formation of SLBs. The formation of SLBs was evaluated using water permeability data, the lateral diffusivity of lipid molecules on the PEMs was determined using a fluorescence recovery after photo-bleaching assay and the amount of lipid molecules adsorbed on the PEMs. We revealed that both the molecular structure and charge density of the polyelectrolytes affected the formation of SLBs. Furthermore, we could form SLBs on high permeable PEMs by combining different cationic polyelectrolytes. These SLBs would be applicable for a platform to immobilize lipophilic biomolecules.

Keywords: Liposome; Supported lipid bilayer; Layer-by-layer method; Polyelectrolyte multilayer

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1. Introduction Biological membranes consist of lipid bilayers and various lipophilic biomolecules [1], such as membrane proteins [2], peptides [3], glycolipids [4], or glycoproteins [5]. Lipid bilayers separate the interior and exterior of cells and maintain intracellular homeostasis; moreover, lipophilic biomolecules involved in the lipid bilayer effectively recognize specific molecules and present various important properties, such as selective reactivity, adsorption, and permeation [6][7]. The structures and properties of biological membranes have evolved in time. Investigating the properties of biological membranes could be useful not only for revealing their mechanism but for their potential applications for artificial devices, including drug screening devices [2][8], biosensors [7][9][10], and separation membranes [11][12]. However, it is difficult to isolate only a specific function and to use biological membranes for artificial devices directly. To use the superior functions of biological membranes for artificial devices, biomimetic membranes, which are simplified membranes that mimic biological membranes, have been studied [13]. Biomimetic membranes consist of lipid bilayers and lipophilic biomolecules, and their lipid composition can be easily controlled by changing the lipids to maximally express the functions of the target biomolecules. Black lipid membranes (BLMs) [2][14] and liposomes [15][16] are frequently used to form biomimetic membranes. BLMs are flat lipid bilayers formed on 100-1000 μm hydrophobic pores, which allow to control the solutions on both sides of the lipid bilayer and to measure the transport of materials across the membrane [17] or the membrane current [18]. To date, BLMs have been used as a simple platform to evaluate the functions of highly sensitive lipophilic biomolecules [19] (e.g., α-hemolysin [20] and gramicidin [2]). Conversely, liposomes are vesicular lipid bilayers dispersed in aqueous solvents and are also widely used as a simple biomimetic membrane platform [21]. Liposomes can not only hold lipophilic biomolecules on the

4

lipid bilayer, similar to BLMs, but can also encapsulate various substances in the inner phase of their lipid bilayer. Liposomes have been studied for various applications, including drug carriers [22] and cosmetics [23]. However, conventional biomimetic membranes are not sufficiently stable to be used for engineering devices, such as sensors and separation membranes, because they are free-standing membranes. Supported lipid bilayers (SLBs), which are planar lipid bilayers spread on flat, non-porous substrates, such as silicon wafers [24], mica [25], silica [26], or glass [27], are effective approaches for stabilizing biomimetic membranes. Generally, SLBs are obtained using the Langmuir–Blodgett method [28][29], spin coating method [24], or liposome fusion method [30][31][32]. The liposome fusion method is the most frequently used and can generate uniform lipid bilayers without the use of organic solvents. First, liposomes interact with, and are adsorbed on the surface of the substrate via various physical interactions, such as van der Waals forces [27][33][34], electrostatic interactions [32][35][36], or hydrophobic interactions [37]. Then, the liposomes rupture and are transformed into planar lipid bilayers. Generally, SLBs are more stable than conventional unsupported biomimetic membranes, such as BLMs or liposomes [38][39], and are expected to present more practical applications for bio-screening [6], sensors [7][9], and purification membranes [40]. However, the hard substrate negatively affects the ability of lipophilic biomolecules incorporated into SLBs [5][6][41]. To prevent the substrate and lipophilic biomolecules from coming into contact, recently, porous substrates, such as inorganic or polymeric water purification membranes, have been used as supports for SLBs [40][42][43]. The structure and ability of lipophilic biomolecules on SLBs on porous substrates is maintained without deactivation [44]. However, the roughness of the surface of these substrates affects the formation of uniform and planar SLBs, particularly for SLBs obtained using the liposome fusion method,

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and could cause defects in SLBs [35]; therefore, a promising method to form SLBs on porous substrates has not been developed yet. To form SLBs on various surfaces, polyelectrolyte multilayers (PEMs) prepared using the layer-by-layer (LbL) method have been used as SLB substrates [45][46][47][48]. When the LbL method is used to prepare PEMs, cationic and anionic polyelectrolytes are alternately stacked via electrostatic interaction on substrate surfaces [49]. When SLBs are formed on PEMs, PEMs act as a cushion to prevent the direct contact between the lipophilic biomolecules incorporated in SLBs and the substrate surface [6]. Flat and non-porous substrates, such as grass [6], mica [50], and silica [47][48] have been used for the formation of SLBs on PEMs. Recently, porous substrates, such as hydrolyzed polyacrylonitrile has been used for the formation of SLBs on PEMs for water purification membrane applications [45][46]. Porous supports present molecular permeability themselves, which allows effective molecular access to the biomolecules on SLBs; therefore, such supports have attracted the attention of researchers and have been studied for further progressive applications, such as biosensors [51][52], a platform for protein studies [53], and element separation [45][46]. M. Wang et al. reported the formation of SLBs with water channels (Aquaporin Z; AqpZ) on PEMs on a porous substrate for reverse osmosis water purification membranes [45]. They revealed that the water permeability of SLBs with AqpZ was 40 times smaller than that of liposomes with AqpZ because some AqpZ escaped from the lipid bilayers during the proteoliposome rupture process. S. Wang et al. reported the fabrication and characterization of AqpZ-incorporating forward osmosis membranes on PEMs as a substrate [46]. The obtained membranes presented high reversed salt solution flux owing to the presence of defects on the SLB. Although the formation of SLBs on PEMs on porous substrates has been

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demonstrated, the studies cited above have not properly described the formation and structure of SLBs. In this study, we investigated the effect of the molecular structure of PEMs prepared on porous substrates on the formation of SLBs. First, PEMs were prepared on nitrocellulose membranes as porous substrate, using anionic poly(sodium 4-styrenesulfonate) (PSS) and three types

of

cationic

polyelectrolytes,

poly(diallyldimethylammonium

chloride)

(PDDA),

polyvinylamidin (PVAm) and polyethyleneimine (PEI) utilizing the LbL method; moreover, their water permeability was used as the index of molecular permeability. Then, SLBs were formed onto these PEMs and the lipid bilayer structure was evaluated using water permeability measurements, lipid fluidity evaluation, and phosphorus quantification assays. To improve the molecular permeability of PEMs as substrates of SLBs, combinations of different cationic polyelectrolytes were also used to prepare PEMs.

2. Materials and methods 2.1.

Materials A nitrocellulose membrane (pore size 0.05 m, cellulose acetate and nitrocellulose mixed

membrane, Merck, Darmstadt, Germany) was used as a porous substrate for PEMs. We used PDDA (Mw=450 kDa, Sigma-Aldrich, St. Louis, MO, USA), which consists of linear short polymer chains with quaternary ammonium cations, PVAm (Mw = 3500 kDa, Mitsubishi Chemical Corporation, Tokyo, Japan), which features a linear, long polymer chain, and PEI (Mw = 750 kDa, Sigma-Aldrich) which presents a branched polymer chain, as cationic polyelectrolytes,

7

and PSS (Mw = 200 kDa, Sigma-Aldrich) as anionic polyelectrolyte. In addition, 1-palmitoyl-2oleoyl-sn-glycero-3-phosphocholine (POPC; NOF, Tokyo, Japan) and 1-palmitoyl-2-oleoyl-snglycero-3-phospho-(1'-rac-glycerol) (sodium salt) (POPG; NOF) were used as electrically neutral and anionic lipids, respectively, and 1,2-dioleoyl-sn-glycero-3-phosphophenolamine-N-(lissamine rhodamine B sulfonyl) (ammonium salt) (Rhod-PE; Avanti polar lipids, Alabama, USA) was used as fluorescent lipid. Other reagents were purchased from FUJIFILM Wako Pure Chemical (Osaka, Japan) and were used without further purification. All aqueous solutions were prepared using Milli-Q water (Merck). The chemical structures of the polyelectrolytes used in this study are presented in Fig. 1. (A) PDDA+

(B) PVAm+

(D) PSS -

(C) PEI+

Fig. 1. Chemical structures of cationic polyelectrolytes (A) poly(diallyldimethylammonium) (PDDA+), (B) polyvinylamidin (PVAm+) and (C) branched polyethyleneimine (PEI+), and anionic polyelectrolyte (D) poly(sodium 4-styrenesulfonate) (PSS-).

2.2.

Liposome preparation

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Liposome suspensions were prepared using the extrusion method [54]. Solutions of POPC, POPG, and Rhod-PE in chloroform were mixed (molar ratio of 29.85:69.65:0.5) in a flask. A lipid thin film was formed on the bottom of the flask by evaporating the solvent at 40 °C for 15 min using a rotary evaporator (N-1100; Tokyo Rikakikai, Tokyo, Japan). The obtained thin film was dried under vacuum overnight using a desiccator. Multilamellar liposomes were formed via hydration with Milli-Q water. The obtained liposome suspension was frozen in liquid nitrogen for 3 min followed by thawing in a water bath at 55 °C for 5 min. This process was repeated five times. Afterward, the liposome suspension was extruded 10 times through a polycarbonate tracketched membrane (pore diameter: 0.1 μm; Merck) using an extruder (Lipex Biomembranes, Vancouver, BC, Canada) at 55 °C. The final lipid concentration of the liposome suspension was 0.4 mmol/L.

2.3.

PEM preparation on porous substrate PEMs were prepared by repeating the adsorption of the cationic and anionic polyelectrolyte

layers using the LbL method. Porous substrates were hydrophilized using vacuum plasma (YHSR, Sakigake-semiconductor, Kyoto, Japan) for 1 min and were immersed in Milli-Q water for 15 min to facilitate the adsorption of polyelectrolytes. The treated surface changed more hydrophilic because of the generation of carboxyl or hydroxyl groups (Fig. S1). The porous substrates were immersed in 40 mL of each of the cationic polyelectrolyte aqueous solutions, Milli-Q water, the anionic polyelectrolyte aqueous solution, and Milli-Q water in this order every 5 min. The concentrations of the polyelectrolyte solutions were 1 g/L. We added NaCl to increase the thickness of PEMs to cover the pore of the substrate. Na+ or Cl- ions shield the intra-polyelectrolyte

9

electrostatic repulsion, and more cationic or anionic polyelectrolytes adsorbed on the counterionic surface [55]. The PEMs consisted of “N” layers of cationic and “N-1” layers of anionic polyelectrolytes and were denoted as “PEMN” or “[polyelectrolyte]N” (e.g., PDDA4) (Fig. 2).

2.4.

Formation of SLBs on PEMs The formation of SLBs on the surface of PEMs occurred via the electrostatic interactions

between the cationic surface of the PEM and the anionic surface of the liposomes. First, The PEMs were immersed in 15 mL 10 mmol/L MgSO4 aqueous solution at room temperature for 1 h, as pretreatment. Then, the PEMs were immersed in 10 mL liposome suspensions at room temperature for 2 h. The SLBs formed on the PEMs were denoted as “PEMN-SLB” or “[polyelectrolyte]NSLB” (e.g., PDDA4-SLB) (Fig. 2).

Fig. 2. Schematic illustration of preparation of polyelectrolyte multilayers that consisted of “N” layers of cationic and “N-1” layers of anionic polyelectrolytes (PEMN) and formation of SLBs on PEMN (PEMN-SLBs).

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2.5.

Water permeability test The water permeability of the PEMs and PEM-SLBs was measured using a laboratory-

made cross-flow type of permeability test cell [56]. The diameter and effective area of the PEMs and PEM-SLBs were 25 mm and 3.14 cm2, respectively. Milli-Q water was used as the feed solution and permeated through the PEMs or PEM-SLBs by applying the hydraulic pressure of approximately 1 bar using a back pressure valve. The surface of the PEMs or PEM-SLBs were stirred at 200 rpm in the cells. The permeated mass was recorded to calculate the water permeability (L/m2/h/bar) using equation (1): Water permeability =ΔV/(A·Δt·P),

(1)

where, V, A, and P are the permeated liquid volume, the projected area of the porous substrate, and applied pressure, respectively.

2.6.

Evaluation of lipid fluidity using fluorescence recovery after photobleaching assay If a planar lipid bilayer is formed on the surface of a porous substrate, lipid molecules can

diffuse laterally on the surface of the substrate. The lateral diffusivity of lipid molecules on the surface of PEMs was evaluated using a fluorescence recovery after photobleaching (FRAP) assay utilizing a confocal laser scanning microscopy (CLSM; FV1000D, Olympus, Tokyo, Japan) instrument [57]. The fluorescence of the lipid molecules on the PEM surface was partially bleached using 543 nm laser radiation with maximum output power in a 30 μm diameter circular shape.

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After bleaching, CLSM images were continuously recorded using the 543 nm laser radiation as excitation source and a 560 nm dichroic mirror as emission source. The fluorescence intensity values of the bleached and unbleached areas (Ib and Ir, respectively) were obtained via image analysis using Image J (NIH, Bethesda, MD, USA) software. The relative fluorescence intensity, RI(t) = Ib/Ir, and fluorescence recovery ratio, FR(t) = (RI(t) – RI(0))/(1 – RI(0)), were calculated. The lateral diffusion coefficient (D, cm2/s) of the lipid molecules of the PEMs was determined by fitting the theoretical equation (2): τD = ω2/ 4D, FR(t) = exp(-2τD/t)[I0(2τD/t)+ I0(2τD/t)],

(2)

where τD, I0, and I1, and ω are the characteristic diffusion time, modified Bessel functions, and radius of the bleached spot (15 μm), respectively [57].

2.7.

Evaluation of number of lipid bilayers on PEM surface To calculate the number of lipid bilayers on the PEM surface, the amount of phospholipid

molecules adsorbed on the PEMs was measured using the modified Bartlett method, as previously reported [44]. The number of lipid bilayers was calculated using the following equation: Number of lipid bilayers = Q (Σxiai)/A,

(3)

where Q, xi, ai, and A are the quantified phospholipid amount, molecular ratio of each phospholipid in the lipid bilayers, area occupied by each phospholipid in the lipid bilayer [58], and projected area of the porous substrate, respectively. This value represents the amount of lipid molecules on the surface of PEMs as the ratio of the area to the projected area of the substrate.

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3. Results and discussion 3.1.

Effect of number of layers on structure of PEMs and PEM-SLBs The effects of the number of layers and polyelectrolyte species of PEMs on their water

permeability are illustrated in Fig. 3. The water permeability of PEMs decreased as the number of layers increased, regardless of the polyelectrolyte species. This indicated that the polyelectrolytes were stacked on the substrate surface using the LbL method and filled the substrate pores [59]. The water permeability of the PEMs prepared using PDDA and PVAm decreased by approximately 25%, 65% and 65% when the number of layers increased from one to two, from two to three, and from three to four, respectively. These differences in water permeability were probably caused by the different stacking structures of each layer. On the assumption that the polyelectrolytes are spherical, the diameters of the PDDA (450 kDa), PVAm (3500 kDa), PEI (750 kDa) and PSS (200 kDa) are estimated about 28.9, 77.2, 36.9, and 19.5 nm, respectively (Table S1) [60]. Actually, the polyelectrolytes have intra-molecular repulsion and should not be spherical but be larger. Therefore, the polyelectrolyte sizes are close to the size of the substrate pores (0.05 m), and the substrate pores were covered after the 4-cycle LbL stacking operations. The polyelectrolytes were first adsorbed on the substrate surface and reduced its pore size, which caused the small decrease in the water permeability that corresponded to PEM1 or PEM2. Then, the pores of the substrate became entirely covered with polyelectrolytes, and water permeability significantly decreased, which corresponded to PEM3 or PEM4. The water permeability of the PEMs prepared using PEI was higher than those of the PEMs prepared using PDDA and PVAm. Particularly, the water permeability of PEI4 was much higher than those of PDDA4 or PVAm4.

13

The branched structure of PEI generated sparse PEMs, while the linear chains of PDDA and PVAm generated dense PEMs [61]. Next, we attempted to form SLBs on the obtained PEMs. Fig. 3 presents the effect of the number of layers of PEMs on the water permeability of the subsequently formed SLBs. The water permeability of PDDAN-SLBs decreased greatly from PDDA2-SLB to PDDA3-SLB, probably owing to the different stacking structures of PDDA2, which featured a small size of pores and PDDA3, which presented covered pores. The water permeability of PDDA1-SLB and PDDA2-SLB increased after the formation of SLB. The PEMs with small layer numbers like as PDDA1 or PDDA2 probably peeled off from the substrate surface during the immersion process into the liposome suspension because the polyelectrolyte network was not grown enough to stably adsorb the liposomes. The water permeability of PEM4-SLB was lower than that of PEM4 using all three types of cationic polyelectrolytes. The water permeability of PDDA4-SLB, PVAm4-SLB, and PEI4-SLB were 89.7%, 45.5% and 99.7%, respectively, and were lower than those of the corresponding PEMs. The significant decrease in the water permeability values of PDDA4-SLB or PEI4-SLB compared with those of their corresponding PEMs indicated that the PEM surface was highly covered by a lipid bilayer structure. About the PEMs prepared with PVAm, a few lipid molecules were probably adsorbed on the surface of PVAm4.

386

471

PEM-SLB

400 300

298

200 100 0

89 16

29

3

PDDA 1 PDDA 2 PDDA 3 PDDA 4

Water permeability [L/m2/h/bar]

Water permeability [L/m2/h/bar]

500

PEM

523

600

(C) PEM

410

PEM-SLB

500 400

307

300 200

102

100 0

n.d.

n.d.

n.d.

33

18

PVAm 1 PVAm 2 PVAm3 PVAm4

Water permeability [L/m2/h/bar]

(B)

(A) 600

600

551

500

PEM PEM-SLB

360

400 297

300

225

200 100 0

n.d.

PEI 1

n.d.

PEI 2

n.d.

PEI 3

0.6 PEI 4

14

Fig. 3. Water permeability of the PEMs with different numbers of layers, obtained using (A) PDDA, (B) PVAm and (C) PEI as cationic polyelectrolytes. Here, “n.d.” indicates no data, it means no experiments were conducted. The error bars indicate the standard deviation of the measured values.

3.2.

Effect of cationic polyelectrolyte species on lipid fluidity of SLBs The water permeability values of PDDA4-SLB and the PEI4-SLBs indicated that lipid

molecules were adsorbed on the surface. To investigate the adsorption of lipid molecules, the lipid fluidity on the surface of PEM4-SLB was evaluated using a FRAP assay. Fig. 4 presents the time evolution of FR of the lipid molecules adsorbed on the prepared PEM4 obtained using the FRAP assay. The fluorescence of PDDA4-SLB recovered after photobleaching, which indicated that the lipid molecules on the surface of PDDA4 were able to diffuse laterally and presented fluidity. Because lipid fluidity is characteristic only for planar lipid bilayers, it was concluded that a planar lipid bilayer was formed on the surface of PDDA4 [57]. Conversely, the fluorescence of PVAm4SLB did not recover. Considering the relatively small decrease in water permeability of PVAm4SLB compared with that of PVAm4 (Fig. 3B), we concluded that the liposomes did not interact enough with the surface of PVAm and no planar lipid bilayer formed. Moreover, the fluorescence of PEI4-SLB did not recover, while the water permeability of PEI4-SLB was greatly decreased (Fig. 3C). The liposomes probably adsorbed on the PEM surface but not ruptured owing to its sparse structure, and therefore, they did not possess fluidity. The calculated D value of PDDA4-SLB was 1.9 × 10−9 cm2/s, which was lower than that of the flat glass surface [62]. The fitting curve could not be completely fitted because the fluorescent lipids diffused slower on PDDA4 than on glass

15

because the cationic surface of PDDA4 interacted with, and conferred negative charge of fluorescent lipids. The FR values of PEI4-SLB and PVAm4-SLB were too low to calculate the corresponding D values. A simulation result using the liposome fusion method indicated that the surface charge density of PEMs affected the liposome spreading behavior [31]. The charge densities of PDDA and PEI were large, and were attributed to their quaternary ammonium cation and numerous primary, secondary, and tertiary amines. Conversely, the charge density of PVAm was relatively small, and liposomes did not adsorb on the surface of the PEMs prepared using PVAm. The PEMs prepared using polyelectrolytes with high charge density, such as PDDA, could form planar lipid bilayers. 1.0

1.0

0.8

0.8

PEI2-PDDA 2 PDDA 4 PVAm 4 PEI 4

FR [-]

FR [-]

FRFR[-][-]

FR [-]

0.6 1.0 1.0 2-PDDA 2 2 PEI2-PDDAPEI 1.0 0.4 0.4 4 2-PDDA 2 PEI PDDA 4 PDDA 0.8 4 PDDA 0.8 PVAm 4 PVAm 4 0.8 PVAm 0.2 0.2 PEI 4 4 PEI 4 PEI 4 0.6 0.6 0.6 0.0 0.0 0 30 0.410 0 0.4 2010 20 30 Time [min] 0.4 Time [min] 0.2 0.2 0.2 ratio (FR) of PEM4-SLBs and using two layers of Fig. 4. Time evolution of fluorescence recovery 0.0 0.0 PEI and two layers of PDDA (PEI2-PDDA 0.02-SLB). The solid line is the theoretical line using 0 20 30 010 10 20 30 0 10 [min] 20 [min] 30 equation (2). Time Time Time [min] 0.6

16

3.3.

Effect of cationic polyelectrolyte species on lipid adsorption on substrate To elucidate the sorption state of lipid molecules, the number of lipid bilayers of the formed

PEM4-SLB was estimated. The calculated number of lipid bilayers of the formed PEM4-SLBs are shown in Fig. 5. The number of lipid bilayers of PDDA4-SLB, PVAm4-SLB, and PEI4-SLB were 14.9, 3.8, and 27.8, respectively. The number of lipid bilayers of all PEM4-SLBs were greater than 1. However, the lipid fluidity data of PDDA4-SLB in Fig. 4 indicates flat lipid bilayers formed. Multiple lipid bilayers probably did not form owing to electrostatic repulsions, because 70% of the lipids in liposomes are anionic lipids [63]. The number of lipid bilayers was calculated using the projected area of the porous substrate instead of the effective area, which includes the surface of the internal pores. We hypothesized that SLB formed along with the shape of the pore; therefore, the number of lipid bilayers calculated using equation (3) became greater than 1. We presented the possible states of the stacking of each polyelectrolyte and liposome sorption from the above results (Fig. 6.). The different adsorption behavior of PVAm can be explained by the molecular structure and charge density. The molecular structure of PDDA, PVAm and PEI are long linear, very long linear, and branched structure, respectively (Fig. 1). The surface charge of PDDA4, PVAm4 and PEI4 around the pH value of ultra-pure water (pH=5-7) were about +10 mV, -15 mV and +20 mV, respectively (Fig. S2). The water permeability of PDDA4-SLB was low and its lipid fluidity was high. Moreover, PDDA4-SLB presented a planar lipid bilayer on the surface of its PEM. PDDA has long linear polymer chain with quaternary ammonium cations through both the PEM preparation and formation of SLB. Therefore, PDDA forms dense and not permeable PEM (Fig. 3), and attract the anionic lipids (Fig. 5), resulting in the formation of a flat lipid bilayer (Fig. 6A). PVAm4-SLB presented high water permeability and no lipid fluidity. The PVAm4 had anionic surface charges (Fig. S2) because PVAm has a very long polymer chain with

17

low cationic density, and peeled off through the PEM preparation processes. Therefore, the prepared PEMs hardly attracted the anionic lipids (Fig. 6B). PEI4-SLB presented low water permeability while no lipid fluidity. PEI has the branched molecular structure, while it has the highest amine ratio, 32.6~36.8 %, comparing with PDDA (8.7 %) and PVAm (10.6 %). Therefore, sparse and high permeable PEMs were obtained from PEI, and lipids adsorbed as liposomes (Fig. 6C). These results indicated that polyelectrolytes with linear molecular structure and high charge density are required to prepare PEMs that could form planar lipid bilayers. In conclusion, we revealed that the structure and charge density of the polyelectrolytes affected the formation of SLBs. We could form flat lipid bilayer on the soft and permeable support even if there is still space to reduce the amount of lipids. We expect it to be applicable for a platform to immobilize various

Number of lipid bilayers [-]

lipophilic biomolecules such as peptides and membrane proteins.

27.8

30 20

17.0

14.9

10 3.8

0

PDDA 4

PVAm 4

PEI4

PEI2PDDA2

Fig. 5. Number of lipid bilayers on polyelectrolyte multilayers with four layers estimated using phosphorus quantification. The error bars indicate the standard deviation of the measured values.

18

Fig. 6. Schematics of the effect of cationic polyelectrolytes on the adsorption behavior of lipids on the PEM surfaces.

3.4.

Formation of SLBs on PEMs prepared by combining two polyelectrolytes A planar lipid bilayer could form on the PDDA4 surface; however, the water permeability,

of PDDA4, which is used as the index of molecular permeability, was quite low (Fig. 3A). The molecular permeability of substrates is an important factor for SLB applications for biosensors or as a platform for protein studies and element separation. This was attributed to substrates with low molecular permeability hindering the access to the biomolecules incorporated in SLBs. The PEMs prepared using PEI presented high water permeability (Fig. 3C), although planar lipid bilayers did not form on these PEMs. Therefore, we used PEI as bottom layers of PEMs to obtain PEMs with high water permeability and we used PDDA as top layers of PEMs to obtain PEMs, which could form planar lipid bilayers on their surface. Fig. 7 illustrates the water permeability of the PEMs prepared using PDDA and PEI with the different number of layer ratios: PEIM-PDDAN-SLBs. The water permeability of PEIM-PDDAN decreased as the number of PDDA layers increase, although the water permeability of PEIM-PDDAN was higher than that of PDDA4. Moreover, PEIM-PDDAN presented both sparse PEI layers as bottom layers and dense PDDA layers as top layers. The

19

subsequent SLB formation decreased the water permeability of PEI2-PDDA2 by 99%, which

Water permeability [L/m2/h/bar]

indicated that lipid molecules were present on PEI2-PDDA2. 300

225 193

250

PEM 191

200

PEM-SLB 143

150 100 50 0

0.6

4-0

1.7

3-1 2-2 1-3 PEI M-PDDAN

29 3.0

0-4

Fig. 7. Water permeability of PEMs formed using “M” layers of PEI and “N” layers of PDDA (PEIM-PDDAN) and that of corresponding SLBs (PEIM-PDDAN-SLB). For comparison, water permeability of PEI4 (4-0) and PDDA4 (0-4) are also included. Here, “n.d.” indicates no data, it means no experiments were conducted. The error bars indicate the standard deviation of the measured values.

To characterize the sorption state of lipid molecules on PEI2-PDDA2-SLB, we evaluated its lipid fluidity using the FRAP method. The time evolution of FR of PEI2-PDDA2-SLB is also presented in Fig. 4. The fluorescence of PEI2-PDDA2-SLB recovered by approximately 0.4 after 30 min and hardly further increase; this FR value was half of that of PDDA4-SLB, and the number of lipid bilayers of PEI2-PDDA2-SLB was 17.0 (also shown in Fig. 5). Lipid molecules with fluidity and without fluidity were present on the surface of PEI2-PDDA2. The presence of lipid

20

molecules that exhibited fluidity indicated that the high charge density of PDDA led to the formation of planar lipid bilayers on the surface of PEI2-PDDA2. Conversely, lipid molecules without fluidity were also present on the surface of PEI2-PDDA2. Some liposomes probably did not transform into lipid bilayers but were adsorbed on PEI2-PDDA2. A partial PEI was present on the surface of the PEI2-PDDA2 and the liposome that interacted with PEI did not rupture. However, the surface of PEI2-PDDA2 was entirely covered by lipid bilayers or liposomes because the water permeability of the PEI2-PDDA2 was greatly decreased by the formation of the SLB. Combining two different types of cationic polyelectrolytes, we could form SLBs featuring high water permeability on PEI2-PDDA2.

4. Conclusions

We investigated the formation behavior of SLBs on PEMs prepared with three types of cationic polyelectrolytes using the LbL method. Effect of the molecular structure of the cationic polyelectrolytes on the formation of SLBs was evaluated through water permeability measurement, lipid fluidity evaluation, and phosphorous quantification assay. The molecular structure and charge density of the cationic polyelectrolytes affected the formation of SLBs. The polyelectrolytes with high charge density such as PDDA and PEI attracted the liposome, while polyelectrolyte with low charge density such as PVAm could not attract the liposome. However, sparse PEI prevented the SLB formation on the PEM. Therefore, PDDA was the best cationic polyelectrolyte for the SLB formation. Furthermore, we were able to form SLBs on PEMs with high permeability by combining different types of cationic polyelectrolytes. The obtained SLBs

21

would be applicable for a platform to immobilize lipophilic biomolecules such as peptides and membrane proteins.

Acknowledgments This work was supported in part by a Grant-in-Aid for Young Scientists (A) (No. 17H04963) from the Japan Society for the Promotion of Science to D.S.

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