In Situ Monitoring of Membrane Protein Insertion into Block Copolymer

Subscriber access provided by UNIV OF SOUTHERN INDIANA

Applications of Polymer, Composite, and Coating Materials

In situ monitoring of membrane protein insertion into block copolymer vesicle membranes and their spreading via potential-assisted approach Tayebeh Mirzaei Garakani, Zhanzhi Liu, Ulrich Glebe, Julia Gehrmann, Jaroslav Lazar, Stephanie Mertens, Mieke Möller, Niloofar Hamzelui, Leilei Zhu, Uwe Schnakenberg, Alexander Böker, and Ulrich Schwaneberg ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.9b09302 • Publication Date (Web): 22 Jul 2019 Downloaded from pubs.acs.org on July 23, 2019

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 48 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

ACS Applied Materials & Interfaces

Submission to: ACS Applied Materials & Interfaces Manuscript type: Full Paper

In situ monitoring of membrane protein insertion into block copolymer vesicle membranes and their spreading via potential-assisted approach

Tayebeh Mirzaei Garakani1,2‡, Zhanzhi Liu1‡, Ulrich Glebe3,4, Julia Gehrmann1, Jaroslav Lazar5, Stephanie Mertens1, Mieke Möller1, Niloofar Hamzelui1, Leilei Zhu1,6, Uwe Schnakenberg5, Alexander Böker3,4 and Ulrich Schwaneberg1,2* ‡ Authors have contributed equally 1

Institute of Biotechnology, RWTH Aachen University, Worringer Weg 3, D-52074 Aachen, Germany.

2

DWI - Leibniz Institute for Interactive Materials, Forckenbeckstraße 50, D-52074, Aachen, Germany

3

Fraunhofer Institute for Applied Polymer Research IAP, Geiselbergstraße 69, 14476 Potsdam-Golm, Germany. 4

Chair of Polymer Materials and Polymer Technologies, University of Potsdam,

Institute of Chemistry, Karl-Liebknecht-Str. 24-25, 14476 Potsdam-Golm, Germany 5 Institute

of Materials in Electrical Engineering 1, RWTH Aachen University, Sommerfeldstraße 24, 52074 Aachen, Germany

Present Addresses: 6 Tianjin Institute of Industrial Biotechnology, Chinese Academy of Sciences, 32 West 7th Ave., Tianjin 300308, China * Corresponding author: [email protected]

1 ACS Paragon Plus Environment

ACS Applied Materials & Interfaces 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

Page 2 of 48

ABSTRACT Synthosomes are polymer vesicles with transmembrane proteins incorporated into block copolymer membranes. They have been used for selective transport in or out of the vesicles as well as catalysis inside the compartments. However, both the insertion process of the membrane protein, forming nanopores, and the spreading of the vesicles on planar substrates to form solid-supported biomimetic membranes have been rarely studied yet. Herein, we address these two points and, first, shed light on the real-time monitoring of protein insertion via isothermal titration calorimetry. Second, the spreading process on different solid supports, SiO2, glass and gold, via different techniques like spin- and dip-coating as well as a completely new approach of potentialassisted spreading on gold surfaces was studied. While inhomogeneous layers result via traditional methods, our proposed potential-assisted strategy to induce adsorption of positively charged vesicles by applying negative potential on the electrode leads to remarkable vesicle spreading and their further fusion to form more homogeneous planar copolymer films on gold. The polymer vesicles in our study are formed from amphiphilic

copolymers

poly(2-methyloxazoline)-block-poly(dimethylsiloxane)-

block-poly(2-methyloxazoline) (PMOXA-b-PDMS-b-PMOXA). Engineered variants of the transmembrane protein ferric hydroxamate uptake protein component A (FhuA), one of the largest β-barrel channel proteins, are used as model nanopores. The incorporation of FhuA Δ1-160 is shown to facilitate the vesicle spreading process further. Moreover, high accessibility of cysteine inside the channel was proven by linkage of a fluorescent dye inside the engineered variant FhuA ΔCVFtev and hence 2 ACS Paragon Plus Environment

Page 3 of 48 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


preserved functionality of the channels after spreading. The porosity and functionality of the spread synthosomes on the gold plates has been examined by studying the passive ion transport response in the presence of Li+ and ClO4- ions and electrochemical impedance spectroscopy analysis. Our approach to form solid-supported biomimetic membranes via potential-assisted strategy could be important for the development of new (bio-) sensors and membranes.

KEYWORDS: synthosomes, solid-supported biomimetic membranes, polymersome spreading, electrochemical impedance spectroscopy, FhuA



Page 4 of 48

INTRODUCTION Hybrid membranes with incorporated transmembrane proteins acting as nanopores are a promising platform for future applications in biosensors, nanoreactors as well as for drug delivery and water purification. As proteins are exactly monodisperse, they provide an excellent opportunity to function as uniform pores in artificial membranes. As a mimic of natural membranes, lipid vesicles (liposomes) with incorporated membrane proteins were developed.1-4 In the last two decades, polymersomes were evolved into an alternative system exploiting the higher stability of the formed spherical features.5 Amphiphilic block copolymers are used to form polymersomes and have the advantage that the molecular weight of each block can be tailored in addition to easily achievable chemical modifications. Similar to the lipid systems, the permeability of the polymer membrane can be tuned by the insertion of protein nanopores.6-11 Thus, several membrane proteins have been successfully incorporated into polymer membranes with the driving force of stabilizing the hydrophobic region of the protein in the polymer membrane.12,

13

The

triblock

copolymer

poly(dimethylsiloxane)-block-poly(2-methyl

poly(2-methyl oxazoline)

oxazoline)-block-

(PMOXA-b-PDMS-b-

PMOXA) was shown to be particularly suitable for insertion of membrane proteins due to the high flexibility of the hydrophobic PDMS block that can adapt to the structure of the protein nanopore.14 Kumar et al. reported that the incorporation of a membrane protein into block copolymer vesicles affects their morphology.15 Polymersomes were formed from polybutadiene-block-polyethyleneoxide

(PB-b-PEO)

and

PMOXA-b-PDMS-b4

ACS Paragon Plus Environment

Page 5 of 48 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


PMOXA, respectively, and aquaporin-0 (AQP0) water channels inserted. The selfassembled structures were depending on the ratio of protein to polymer. Synthosomes were introduced as polymersomes with an engineered transmembrane protein acting as selective gate.16 Enzymatically catalyzed reactions can take place in these nanocompartments via an enzyme which is entrapped in the vesicle interior with a selective compound recovery or release through the transmembrane protein. A similar approach for enzymatically active self-assembled structures consists of enzymepolymer conjugates that form proteinosomes.17, 18 The conjugates were crosslinked at emulsion interfaces and the proteinosomes used as reaction spaces, for cascade reactions when using multiple enzymes, or protocell models.19 A powerful, label-free online technique to study membrane–surfactant,20, 21 membrane– protein, protein–polymer22, 23 or membrane protein–ligand24 interactions is isothermal titration calorimetry (ITC). ITC is a physical technique used to determine thermodynamic parameters of interactions in solution offering unique accuracy, resolution, and sensitivity over other experimental methods.25 This model-independent method can measure the enthalpy change arising from a binding process.24 Incorporation of a channel protein in vesicles made from lipid molecules was exemplified for KcsA, a homotetrameric K+ channel protein, and monitored by ITC in real time.26 However, to the best of our knowledge, an ITC analysis of the insertion process of protein nanochannels into polymer membranes has not been reported yet.



Page 6 of 48

Ferric hydroxamate uptake protein component A (FhuA) is one of the largest monomeric β-barrel transmembrane proteins with an elliptical cross-section of 39-46 Å and a height of 69 Å.27, 28 In the outer membrane of Escherichia coli (E. coli), FhuA mediates the active transport of ferrichrome bound iron and also acts as the receptor for phage T5, phi 80 and T1 as well as colicin M.29 FhuA consists of an N-terminal inner cork domain which blocks the interior channel and 22 ß-strands forming the barrel structure.28 Due to its rigidity and stability,30 FhuA is an attractive protein scaffold for protein engineering.31-35 Large passive diffusion channels of different size were generated by removing the cork domain (e.g., FhuA Δ1-160).36 Furthermore, we could show that up to eight additional β-strands can be inserted to increase the radius of the protein channel to (2.7 ± 0.6) nm.33 In addition to its function as passive diffusion channel3, 16, 36, 37 for transporting molecules such as ssDNA,36 triggered release systems with FhuA variants based on reduction and light were developed.2, 38 More recently, hybrid FhuA-polymer conjugates were successfully generated by controlled radical polymerization (CRP) of N-isopropylacrylamide (NIPAAm) from lysine residues on FhuA Wild Type and engineered variants.32 Such building blocks could be used to stabilize Pickering emulsions. After crosslinking of the polymer chains, stable micro-compartments were obtained.34 All in all, synthosomes, microcompartments and proteinosomes18 are promising for applications such as nanoreactors and drug delivery. However, stable planar membranes are more promising systems for applications such as separation processes and biosensors.


Page 7 of 48 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


The spreading of vesicles on solid supports is a highly promising strategy to develop such hybrid planar membranes. Spreading of vesicles was originally developed for lipid systems.39 Therefore, liposomes were spread onto a solid support and the membranes formed by fusion of the lipid vesicles.40-43 The interaction of lipid and solid support is highly affected by the composition of the lipid, the surface charge of the supporting substrate material as well as temperature, pH and ionic strength.44 Spreading of liposomes on a solid support is not straightforward and all mentioned parameters have to be optimized to achieve lipid bilayer formation. Although numerous reports demonstrate formation of solid-supported lipid membranes,45 the disadvantages of lipids, for example being prone to oxidation and dehydration,46 limit their use in applications. Consequently, the use of polymersomes with improved stability and mechanical robustness compared to the lipid systems is the expected following step.47 However, there are only few examples of polymersome spreading48, 49 on solid supports to obtain polymer membranes.50 PB-PEO vesicles with hydroxyl groups were spread on hydrophilic glass surfaces through physisorption and lipoic acid-functionalized PBPEO vesicles were spread on gold substrates through formation of covalent bonds. The latter were further incubated with the peptide polymyxin B and interactions between the solid-supported membrane and the peptide detected.48 Here, we firstly analyzed the insertion of FhuA Δ1-160 into PMOXA-b-PDMS-b-PMOXA polymer vesicles with ITC and subsequently performed a detailed study on the spreading of polymersomes as well as synthosomes on different surfaces. FhuA Δ1-160 was exemplarily used as a passive diffusion channel. An electrochemical-based immobilization method suitable 7 ACS Paragon Plus Environment


Page 8 of 48

for polymer vesicle spreading on gold surfaces was developed. In this study, we demonstrate that charge of the PMOXA-b-PDMS-b-PMOXA polymer could be utilized for immobilizing and spreading of vesicles (with and without FhuA) on an electrode surface taking advantage of electrostatic interactions. Our results provide insight that it is possible to establish a reproducible potential-assisted polymer vesicle spreading protocol that is much more efficient and straightforward than the commonly applied spreading methods. In generated planar membranes, a high surface coverage was reached and presence of the protein nanopores was confirmed. EXPERIMENTAL SECTION 1. Materials All chemicals were of analytical grade and obtained from Sigma-Aldrich, unless stated otherwise. For the current study, dialysis buffer or MPD buffer is defined as a buffer with 10 mM sodium phosphate buffer (pH 7.4, 1.9 mM NaH2PO4, 8.1 mM Na2HPO4), 1 mM EDTA and 50 mM 2-methyl-2, 4-pentanediol (MPD). Water of Milli-Q grade was used for preparation of all aqueous solutions described in this paper. 2. Preparation methods Block copolymer vesicles preparation and FhuA Δ1-160 incorporation Polymersomes were prepared via direct dispersion method16 and using two different ABA block copolymers of PMOXA-b-PDMS-b-PMOXA (hereafter denoted as A19B65-A19 and A14-B65-A14) with number-averaged molecular weight of Mn= 8000 and 7200, respectively (both supplied from Polymer Source Inc. Canada). 1 ml of sodium 8 ACS Paragon Plus Environment

Page 9 of 48 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


phosphate buffer (pH 7.4, 10 mM) was added dropwise into 10 mg of ABA block copolymer at room temperature in a glass vial and the suspension was stirred (Cimarec i Poly 15 and Multipoint Stirrers, 300 rpm, Thermo Scientifics, MA, USA) in darkness overnight. Then, the suspension was sonicated twice (Sonics vibra cell VCX130, 40% power, 1 min, Connecticut, USA) with 15 min interval stirring followed by freeze-thaw cycles described previously.16, 51, 52 The polymersome sample was dipped into liquid nitrogen for rapid cooling for 3 min, then it was transferred to a bath at 60°C for thawing for 3 min. The solution was frozen again and this operation was repeated 10-times. Subsequently, the polymersome suspension was extruded through 0.2 µm PCTE filter (Avanti polar lipids extruder, Alabama, USA) for 6-times and purified with size exclusion column packed with sepharose CL-6B (Sigma-Aldrich Chemie, Steinheim, Germany) prior to investigation. For the FhuA Δ1-160 incorporation experiments, 100 µl protein solution in dialysis buffer with the concentration of 55.8 µM (unless stated otherwise) was added into 1 ml dispersion of copolymer vesicles (1.35 mM block copolymer in 10 mM sodium phosphate buffer, pH 7.4) after which the mixture was vortexed until a homogeneous dispersion was formed. The obtained FhuA-containing polymersomes are denoted as synthosomes in this paper. Substrate preparation Glass (microscope slides, VWR International GmbH, Germany) and silicon wafers (ptype, CrysTec GmbH, Germany) were cleaved, cleaned with ethanol and Milli-Q water and dried in the nitrogen stream. Surface cleaning was accomplished by carbon dioxide 9 ACS Paragon Plus Environment


Page 10 of 48

snow-jet after which oxygen plasma treatment was applied directly before spreading in order to remove any organic traces. For this purpose, substrates were put in an air plasma oven (Plasma Activate Flecto 10 USB with a 100 W HF generator, Plasma Technology GmbH, Germany) and activated at 0.2 mbar for 60 s at room temperature. Silicon wafers (Sigma-Aldrich Chemie, Steinheim, Germany) coated with a 30 nm thick titanium followed by a 100 nm thick gold layer on the polished sides were used for this study. To clean the gold electrodes, they were immersed in a tube filled with water and placed in a water bath of ultrasonic cleaner for 10 minutes. Subsequently, electrodes were rinsed extensively with water over 30 s followed by electrochemical cleaning using cyclic voltammetry in H2SO4 (0.5 M, -0.3 to 1.6 V vs. Ag/AgCl/saturated KCl, 100 mV s-1) until stable voltammograms were obtained. The electrodes were then dried in nitrogen flow before each modification step. Spreading of polymersomes and synthosomes All spreading experiments were performed with polymersome samples prepared via direct dispersion method explained above and by diluting 1 ml of as-prepared sample to a final volume of 8.0 ml via addition of sodium phosphate buffer (pH 7.4, 10 mM). For spreading of synthosomes, first 100 µl protein solution in dialysis buffer with the concentration of 55.8 µM was added into 1 ml dispersion of copolymer vesicles (1.35 mM block copolymer in 10 mM sodium phosphate buffer, pH 7.4) after which the mixture was vortexed until a homogeneous dispersion was formed. Likewise, 1 ml of the as-prepared synthosome sample was diluted to a final volume of 8.0 ml via addition


Page 11 of 48 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


of sodium phosphate buffer (pH 7.4, 10 mM) before performing the spreading experiments. To spread on glass, silicon and gold slides via spin-coating and dip-coating, 1 ml of diluted polymer dispersion was mixed with different amounts of NaCl (0.0, 0.5, 8.0 and 45.0 mg) followed by vortexing until the salt dissolved completely. For spin-coating, slides fixed in a spin-coater (WS-650 mSz-6NPP Lite, Laurell Technologies Corporation, PA, USA) were utilized with 15 μl of sample and a rotational speed of 2000 rpm for 60 s at room temperature. In order achieve spreading via dip-coating, the substrates were immersed in the dispersion for a specific incubation time (up to 1000 s) and pulled out with a speed of 166 µm s-1. All spreading experiments through dip-coating and spin-coating methods were accomplished by incubation in an oven (Thermo Scientific Heraeus, GmbH, Germany) at 45ᵒC for approximately 10 minutes. This should have increased the fluidity of the membrane and facilitated the closure of the gaps in more efficient manner.49 To spread copolymer vesicle self-assemblies on cleaned gold substrates via applying an electrical potential, the depositions were carried out using a three-electrode system in combination with a PGSTAT 128N Autolab spectrometer (Eco Chemie, The Netherlands). The gold substrate acted as working, a platinum rod as counter, and an Ag|AgCl electrode containing KCl saturated aqueous solution (Eᵒ= 0.222 at 25ᵒC) as reference electrode, respectively. The electrodes were subjected in 8 ml sodium phosphate buffer (pH 7.4, 10 mM) serving as electrolyte solution. The electrochemically active area on the gold substrates was defined to 1×1 cm2. Potential11 ACS Paragon Plus Environment


Page 12 of 48

assisted spreading was carried out by chronoamperometry (CA) at a potential of -0.75 V (vs. Ag|AgCl) with a pulse duration of 1000 s in mixture of copolymer vesicles. After each spreading, the coated substrate was rinsed thoroughly with sodium phosphate buffer (pH 7.4, 10 mM) in order to remove the unspecifically bound vesicles from the electrode surface, followed by drying under nitrogen stream for 20 s. 3. Characterization Methods Dynamic Light Scattering (DLS) and zeta potential measurement The average hydrodynamic diameter and zeta potentials of polymersomes/synthosomes was measured with a Zetasizer Nano ZS (Malvern Instruments Ltd, Worcestershire, UK) with noninvasive back-scatter technology at a scattering angle of 173ᵒ using the Zetasizer software with version 7.01. Experiments were performed at room temperature. Samples were stirred for 1 h before transfer to cuvettes. In the zeta potential measurement, the electronic field was 5 V cm-1 in constant-current mode. Both DLS and zeta potential were measured in sodium phosphate buffer (pH 7.4, 10 mM). For the FhuA Δ1-160 incorporation, 100 µl protein solution in dialysis buffer with a concentration of 55.8 µM (unless stated otherwise) was added into 1 ml dispersion of copolymer vesicles (1.35 mM block copolymer in 10 mM sodium phosphate buffer, pH 7.4) after which the mixture was vortexed until a homogeneous dispersion was formed. At least, three analyses were measured as representative value for the hydrodynamic radius for all measurements.


Page 13 of 48 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


Calcein release assay A14-B65-A14 polymersomes were prepared using the direct dispersion method exactly same as mentioned above except for that sodium phosphate buffer (pH 7.4, 10 mM) contained 50 mM calcein. After purification by size exclusion chromatography, 50 µl A14-B65-A14 polymersomes and 150 µl sodium phosphate buffer (pH 7.4, 10 mM) were mixed for each measurement in black microtiter plate (MTP). The fluorescence signal was recorded by TECAN infinite M200 (excitation: 495 nm; emission: 525 nm; program: shaking 3 s, kinetic cycle: 15 x 20 s, manual gain 49%, Tecan Trading AG, Switzerland). In total, 4 batches of 10 µl FhuA Δ1-160 solution with the concentration of 55.5 µM were added stepwise and the fluorescence signal was measured after each addition. As negative controls, the same experiment was performed with the addition of sodium phosphate buffer (pH 7.4, 10 mM) on the one side and dialysis buffer (pH 7.4, 10 mM sodium phosphate buffer, 1 mM EDTA and 50 mM MPD) on the other side. Isothermal Titration Calorimetry (ITC) ITC was carried out on TAM III (TA Instruments, Huellhorst, Germany) after gentle degassing of all samples for 10 min at room temperature. Block copolymer A14-B65-A14 was dispersed in sodium phosphate buffer (pH 7.4, 10 mM) to a final concentration of 1.35 mM. Vesicles were prepared via direct dispersion method according to the procedure described above. Incorporation of the membrane protein was accomplished by titrating as-prepared copolymer vesicle dispersion with FhuA Δ1-160 concentration of 55.8 µM in dialysis buffer (pH 7.4, 10 mM sodium phosphate buffer, 1 mM EDTA 13 ACS Paragon Plus Environment


Page 14 of 48

and 50 mM MPD). In this context, 280 μl of aqueous FhuA Δ1-160 solution was filled into a glass syringe followed by removing gas bubbles inside the syringe. 500 μl of polymer vesicle dispersion were filled in the ampoule of stainless steel. 700 μl (i.e., 500 μl + injection volume) solution of sodium phosphate buffer (pH 7.4, 10 mM) was used as a reference in reference ampoule. The measurement was repeated with FhuA Δ1160-free dialysis buffer in the syringe to determine heats of dilutions and influence of MPD under the same condition. In total, 40 successive injections were used (injection volume: 5 µl; stirring speed: 50 rpm; injection duration: 10 s). The first injection was ignored for the analysis. Measurements were carried out at 25°C and the system equilibrated for at least 6 h. Between measurements, the signal was allowed to return to the baseline in 20 min. Automated baseline adjustment and peak integration were done with Originlab software to yield normalized reaction heats as a function of protein concentration and converting raw calorimetric data (heat evolved or consumed for each titration step, ΔH) to a binding isotherm. Scanning Force Microscopy (SFM), Tapping-mode SFM analysis of the samples was performed in intermittent mode on a Bruker Dimension Icon with Nanoscope 8.10 software and OTESPA tips with a nominal spring constant of 12–103 N/m (Bruker Corporation, Billerica, USA). The cantilever was operated at typical scan rates ranging from 0.7 to 1.2 line s-1. Images were recorded in height, amplitude, and phase modes with a size of 512 × 512 pixels. All analyses were performed on height images; however, some processes were visible with better contrast in phase mode. After spreading, the samples were washed three times with Milli-Q 14 ACS Paragon Plus Environment

Page 15 of 48 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


water to remove the wetting layer of the polymersome dispersion and dried at room temperature. Different areas of the sample were analyzed. Experiments were performed in the dry state at average air humidity of 40-50% and temperature of 22°C. At least, five samples were tested for each system. Electrochemical properties The quality and electroactivity (charge storage ability) of the spread vesicles on gold substrates were monitored in a three-electrode system by cyclic voltammetry (CV) assays using the same PGSTAT 128N Autolab spectrometer (Eco Chemie, The Netherlands) explained above. CVs were conducted with the gold substrate as working electrode, a platinum rod as counter electrode, and an Ag|AgCl electrode containing KCl saturated aqueous solution (Eᵒ= 0.222 at 25ᵒC) as reference electrode in a sodium phosphate buffer (pH 7.4, 10 mM) solution containing 0.1 M of LiClO4. Owing to the nondestructive manner and high diffusional rate of Li+ and ClO4¯ ions in solution, LiClO4 was used as typical supporting electrolyte. Accordingly, the CV assays reflect the facility to exchange ions between the obtained solid-supported films and the electrolyte solution because of bulk diffusion of Li+ and ClO4¯ ions through the nanochannels integrated in the film, which is associated with the β-barrel structure of the incorporated protein. The initial and final potential was 0.2 V, while the reversal potential was 1.4 V. All measurements were performed at 25ᵒC using scan rates of 25 mV s-1.



Page 16 of 48

Functionalization of FhuA ΔCVFtev via ThioGlo1® coupling Accessibility of free cysteine group inside the FhuA ΔCVFtev channel was determined following the established procedure.53 The thiol reactive fluorescent dye ThioGlo1® (MMBC, Berry and Associates, USA) specifically couples to the sulfur group of cysteine. In fact, FhuA Δ1-159 C545 scaffold was optimized to improve accessibility of the Cys545 (N548V) which is covalently bonded to an engineered FhuA ΔCVFtev and the binding site around Cys545 of the FhuA ΔCVFtev was rationally reengineered to ensure accessibility of the thiol group. ThioGlo1® (15 µM in 10 mM sodium phosphate buffer, pH 7.4) was added dropwise to the polymersomes-/ synthosomescoated gold wafers. Samples were incubated light protected (1 h, room temperature). After incubation, samples were washed by dipping into fresh sodium phosphate buffer for three times (solution was exchanged between the washing steps) to avoid unspecific binding of the fluorescent dye and directly used for confocal microscopy. Confocal Laser Scanning Microscopy (CLSM) Visualization of the ThioGlo1® fluorescent dye, which specifically couples to the sulfur group of cysteine, was achieved by using a Leica TCS SP8 microscope (Leica Microsystems CMS GmbH, Mannheim, Germany). Samples were excited at 379 nm and 20% laser intensity (argon laser). A PMT detector was used with an emission of 513 nm.


Page 17 of 48 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


Electrochemical Impedance Spectroscopy (EIS) The gold electrodes were connected to a Solartron 1260A impedance analyzer (Ametek, Farnborough, UK) in a combination with a front end EG&G potentiostat (Princeton Applied Research, Oak Ridge, TN, USA) serving as stable voltage and current source as well as precious measurement device. The measured spectrum was obtained between 0.01 Hz and 100 kHz using 10 mV AC signal added to 0.4 V DC bias voltage against open circuit potential. Impedance measurements were carried out in a three electrode system with the blank/coated gold electrode as the working electrode, a platinum mesh as a counter electrode, and Ag/AgCl reference electrode in a sodium phosphate buffer (pH 7.4, 10 mM) solution containing 0.1 M of LiClO4. LiClO4 was used as electrolyte to render the results comparative to those which were obtained from the electrochemical characterization of the obtained films via running CV assays and in order to estimate the ability of the obtained solid-supported membrane to impede ion transport at the interface between the electrolyte and the membrane. The custom-made PMMA measurement cell has been made for a buffer volume of 5 ml. The data were read out to a PC by Z-Plot (Scribner Software, Farnborough, UK). Raw data were further analyzed and simulated using ZView software package (Version 3.2, Scribner Associates, USA). The experimental data have been analyzed and fitted using a Randles equivalent circuit model. The different components can be assigned to individual parts of the membrane architecture and the surrounding environment.



Page 18 of 48

RESULTS PMOXA-PDMS-PMOXA triblock copolymers consist of hydrophilic poly(2methyloxazoline)

outer

blocks

(block

A)

and

a

flexible

hydrophobic

poly(dimethylsiloxane) core (block B). Polymersomes were prepared by direct dispersion from A14-B65-A14 polymers which have a hydrophilic ratio of 33% that is known to be suitable for the formation of vesicles54 (see Supporting Information for details). The presence of polymersomes was verified by cryo-TEM measurements (Figure S2). First, we investigated two copolymers A19-B65-A19 and A14-B65-A14 to visualize their self-assembled structures via cryo-TEM and, thereby, to select a block copolymer with proper chain length. Next, the insertion of FhuA Δ1-160, having an open protein channel, was optimized and followed by calcein release from the vesicles. Subsequently, efforts have been made to monitor the incorporation of FhuA Δ1-160 in the polymer vesicles of A14-B65-A14 block copolymer in real-time via ITC method. DLS gave us another hint revealing that FhuA Δ1-160 is successfully inserted into the polymersomes. Finally, solid-supported planar hybrid membranes were prepared by means of spreading pre-organized polymer superstructures. The spreading procedure was optimized with respect to the type of substrate material and its pre-treatment conditions as well as varying spreading methodology. SFM was used to examine the surface morphology and topography of membranes prepared by spin-coating on different substrates, namely silicon wafer, glass and gold. Since the resulting polymer vesicles are charged (based on zetasizer measurements), we expected the interactions 18 ACS Paragon Plus Environment

Page 19 of 48 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


between the vesicles and the substrates to be dominated by electrostatic forces. Therefore, a potential-assisted spreading pathway was investigated to produce A14-B65A14 polymer membranes on gold substrates and the results were compared to those yielded on the same material by spin- and dip-coating. To further determine the density of incorporated protein channels within the polymer layer, the variant FhuA ΔCVFtev, which can be dye-labeled inside the channel, was used. ThioGlo1® can be linked to a free cysteine placed at position 545 of FhuA ΔCVFtev and fluorescence was detected using confocal microscopy. Besides, in order to understand the film structure further, a combination of contact angle, confocal microscopy and EIS methods was used to elucidate hydrophobicity change of the substrate, distribution of channels within the polymer film as well as electrochemical behavior of conductive surfaces covered with the resulting film. The main results obtained from these experiments are discussed in the following sections. FhuA Δ1-160 insertion into polymersomes: calcein release assay Reconstitution of FhuA Δ1-160 in a polymersome membrane was monitored in situ via performing calcein release assay.16 For this purpose, polymersomes of A14-B65-A14 were formed with entrapped calcein at a self-quenching concentration. Purification was performed by size exclusion chromatography to remove calcein outside the vesicles. Subsequently, 55.5 µM FhuA Δ1-160 in dialysis buffer were added stepwise (10 µl for each batch). The fluorescence signal was recorded at 525 nm as shown in Figure 1 and the addition of FhuA Δ1-160 was indicated (arrows in Figure 1). Increase in fluorescence intensity was caused by diffusion of calcein through the incorporated 19 ACS Paragon Plus Environment


Page 20 of 48

FhuA Δ1-160 channels into the outside of polymersomes where it was diluted and provided a fluorescence signal.

Figure 1. Incorporation of FhuA Δ1-160 into A14-B65-A14 polymersomes loaded with 50 mM calcein leads to release of calcein through FhuA nanochannels and hence increase of fluorescence. For four-times insertion with each batch 10 µl FhuA Δ1160 (55.5 µM) in dialysis buffer, stepwise increase of fluorescence could be observed. The addition of FhuA Δ1-160 or buffer is indicated by arrows. Control experiments with insertion of only buffer did not lead to increase in fluorescence. In situ monitoring of membrane protein insertion into block copolymer vesicles by Titration Calorimetry ITC experiments were performed to study the insertion of FhuA Δ1-160 into polymersomes formed from A14-B65-A14 block copolymers. The experiments were carried out at 25°C by titrating protein solutions into the sample cell containing polymersomes. In general, a major challenge to study systems containing membrane proteins is the need to keep their hydrophobic patches covered in order to keep proteins correctly folded in presence of detergents.24, 55 In the present study, a water-miscible amphipathic alcohol, MPD, was selected as stabilizing agent for FhuA and to avoid interference with subsequent protein analysis steps. Recently, MPD was shown to successfully stabilize different FhuA variants and be beneficial for performing specific characterizations of protein modification.32


Page 21 of 48 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


Figure 2. (left) Raw calorimetric data accompanying insertion of FhuA Δ1-160 into A14-B65-A14 copolymer vesicles (solid line) and the corresponding experiment with the injection of MPD buffer without FhuA Δ1-160 (bold line). Titration of 500 µl stock solution of polymersomes made from 1.35 mM block copolymer in sodium phosphate buffer (pH 7.4, 10 mM) in the reaction cell with FhuA Δ1-160 (55.8 µM) in dialysis buffer solution (10 mM sodium phosphate buffer, pH 7.4, 1 mM EDTA and 50 mM MPD) at 25ᵒC is illustrated, wherein 200 µl of 55.8 µM FhuA Δ1-160 were injected in equal steps of 5 µl. (right) Isotherm obtained by integration of the peaks of the thermogram shown in the left diagram plotted against the volume of added FhuA Δ1160 solution. The squares represent the integrated heat changes during FhuA Δ1-160 incorporation and the line corresponds to the curve fit to the binding isothermal function. Figure 2 (left panel) shows the ITC thermogram displaying differential heat flow per injection of FhuA Δ1-160 (55.8 µM) into copolymer vesicles versus time as well as control experiment of injection of only dialysis buffer into the same polymersome sample. Each “positive” peak corresponds to the heat absorbed upon each inserted 5 µl FhuA Δ1-160 solution indicating that the insertion of FhuA Δ1-160 into polymersomes is endothermic and enthalpically disfavorable. The heat of dilution of the polymersome dispersion during titration of MPD dialysis buffer was shown to be negligible compared to titration of FhuA Δ1-160-containing dialysis buffer solution as depicted in Figure 2 (left panel). As can be seen from the titration curves shown in Figure 2, the illustrated systems exhibit different heat change profiles. Injection of FhuA Δ1-160 involves endothermic processes, smoothly decaying heat signals, which reflect the incorporation of FhuA Δ121 ACS Paragon Plus Environment


Page 22 of 48

160 into polymersomes and the stepwise saturation of copolymer vesicles via injected membrane protein. Throughout plateau range, the reaction heats remained uniform because each injection gives rise to transfer of a constant amount of FhuA Δ1-160 into copolymer vesicles. The insertion of membrane proteins into polymersomes might be of both positive enthalpy and entropy, which can show that the insertion procedure can be driven by a favorable conformational entropy change which compensates for the unfavorable enthalpy change. This should be a balance between two major effects56 which has been discussed in terms of cooperative interaction.57, 58 The protein incorporation event in the polymer vesicle membranes involved two simultaneous processes: On the one hand, detergents, counter ions or water molecules tightly associated with proteins and vesicles should be released.58 For solvent reorganization, energy is required to disrupt the welldefined solvent shells to give rise to positive enthalpy. On the other hand, increase of translational degrees of freedom, upon hydrophobic interactions, generates positive entropy changes.56 The hydrophobic nature of interaction between the hydrophobic region of the membrane protein and the hydrophobic block of the copolymer might play a predominant role in the incorporation of FhuA Δ1-160 in vesicle membrane. Even though it is not clear what accurate factors cause the adsorption of heat of the titrations, it is commonly realized that the hydrophobic nature of interaction is accompanied by positive entropy which further supports the assumption that the primary hydrophobic driven interaction forwards the whole insertion process.59 22 ACS Paragon Plus Environment

Page 23 of 48 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


Particle size distributions before and after FhuA Δ1-160 insertion DLS analysis gave us another indication that FhuA Δ1-160 is successfully inserted into copolymer vesicles. As shown in Table 1, the zeta potential of the vesicles decreased from 26.7 mV to 19.7 mV when FhuA Δ1-160 is inserted. This is in agreement with the negative charge of FhuA Δ1-160 at pH 7.4 (isoelectric point of 5.16, calculated on https://web.expasy.org/compute_pi/). Another interesting point is that when FhuA Δ1160 is inserted, the vesicles increase in size. The mean size of the vesicles increased from 188.2 nm to 269.1 nm after insertion of FhuA Δ1-160. These observations are similar to those reported by Jahnke et al. who examined the reconstitution of the channel protein KcsA from the nonionic detergent octyl-β-D-glucopyranoside into vesicles made from Escherichia coli polar lipid extract.26 Moreover, Kumar et al. similarly reported that small vesicular polymer structures grew to larger vesicles upon protein incorporation.15 Table 1. Light scattering and ζ-potential analysis of A14-B65-A14 self-assembled structures in aqueous solution.

Vesicle composition

Rg (nm)

ζ-potential (mV)

A14-B65-A14 polymersome

188.2 ± 1.7

26.7 ± 2.1

A14-B65-A14 synthosome

269.1 ± 7.4

19.7 ± 0.8



Page 24 of 48

Spreading of polymersomes and synthosomes on silicon-wafers, glass and gold slides After analyzing the reconstitution of FhuA Δ1-160 in polymer vesicle membranes, we studied the spreading of polymersomes and synthosomes on different planar supports to form solid-supported membranes. Therefore, we chose vesicles formed from A14B65-A14 copolymer, silicon-wafers, glass and gold as substrates and investigated the conditions for spreading in order to achieve more homogeneous films on the substrates. Dip- and spin-coating were performed on all substrates in parallel for all set of spreading processes. Spin-coating turned out to be a useful technique to optimize polymersome spreading. Furthermore, the addition of NaCl to the vesicle dispersion prior to spreading could improve the homogeneity of the obtained film. It has been reported that the addition of salt resulted in a concentration gradient between the intraand extra-vesicular spaces.49 Osmotic pressure led to a decrease in vesicle volume, while the membrane surface remained constant. This exerted mechanical strain on the membrane surface destabilized the polymer vesicles and facilitated their spreading. Therefore, we tested the influence of different concentrations of salt (0.0, 0.5, 8.0 and 45.0 mg/ml) and found that most efficient spreading was achieved when 0.5 mg of sodium chloride was added into 1 ml of the dispersion prior to deposition (8.5 mM). The substrates were heated up to 45ᵒC by placing them in an incubation oven for about 10 minutes to further improve surface coverage and bilayer homogeneity.49 To remove the non-spread polymer aggregates present on the surface, the samples were subsequently rinsed with water and blown dry under a stream of nitrogen for 24 ACS Paragon Plus Environment

Page 25 of 48 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


approximately 20 s. It has already been observed that quick dehydration removed most of the loosely adsorbed polymer material, while the bilayer itself remained unaffected.49 In general, spreading of polymersomes led to inhomogeneous films on substrates with a mixture of membrane monolayers, bilayers and adsorbed intact vesicles. The white dots observed during SFM characterization (see Figures 3 and 4) correspond to adsorbed and non-collapsed vesicular compartments. Remarkably, the thickness of the film was estimated around 10 nm in some regions supported by measured step-heights in the SFM profile for all three substrate materials depicted in Figures 3 b, d, f, h and Figure 4b, which are approximately 10 nm and can be attributed to the thickness of a single wall patch. In fact, fully stretched triblock copolymer molecule as well as two neighboring U-form blocks sitting on each other would have a length of 10 nm. Moreover, the membrane was partially removed by scratching with a cannula in order to determine the thickness of the membrane deposited on silicon and glass substrates by SFM height profiles. The polymer-layer thickness of about 10 nm (Figure S4) is in good agreement with the layer thickness calculated via cryo-TEM imaging of a vesicular wall (Figure S2b). Therefore, a film thickness of 10 nm presumably corresponds to a strongly adsorbed block copolymer on the surface. This implies that the obtained films partly consist of a monolayer. Since the polymer vesicles showed a positive zetapotential (Table 1), the interaction and hence the spreading should be favored for hydrophilic substrates which are more negatively charged. Silicon and glass tend to have a more negative polarization than 25 ACS Paragon Plus Environment


Page 26 of 48

gold substrate.60 This could explain why more intact vesicles remain on the gold surface, when vesicles are spread via spin-coating method (Figure 4). In principle, synthosomes with inserted FhuA Δ1-160 spread differently compared to polymersomes. Interestingly, the obtained films are more homogeneous with a smoother texture, less non-covered areas and fewer adsorbed intact vesicles when synthosomes were spread (Figures 3e, g and 4b). Hence, insertion of FhuA Δ1-160 into copolymer vesicles leads to more efficient spreading. This tendency goes in line with the observation from Kumar et al. that a nearly planar polymer membrane resulted at a high amount of incorporated protein.15 Incorporation of FhuA provokes structural changes in the vesicles which facilitate the spreading of synthosomes compared to polymersomes. The structural changes are apparent from the size expansion of FhuA Δ1-160-loaded copolymer vesicles (Table 1). All in all, protein incorporation facilitates the spreading of vesicles and leads to more homogeneous films on all tested substrates.


Page 27 of 48 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


Figure 3. SFM surface characterization after spreading of polymersomes on a silicon wafer (a) and a glass slide (c) as well as spreading of synthosomes on a silicon wafer (e) and a glass slide (g). Spreading was performed by spin-coating after addition of 8.5 mM NaCl to A14-B65-A14 vesicles (e, g with incorporated FhuA Δ1-160). The height profiles (b, d, f and h) were obtained from the indicated lines in SFM images a), c), e) and g), respectively.



Page 28 of 48

Figure 4. SFM surface characterization after spreading of a) polymersomes and c) synthosomes (with FhuA Δ1-160) on gold. Spreading was achieved via spin-coating after addition of 8.5 mM NaCl to A14-B65-A14 vesicles. The height profiles b) and d) were obtained from the indicated lines in images a) and c), respectively. Potential-assisted deposition of A14-B65-A14 vesicles on gold substrate The interest in developing potential-assisted deposition was first revealed by the results of zeta potential analysis. Since the vesicles are charged (Table 1), we expected electrostatic interactions between the vesicles and the substrate to become more important when the substrate is additionally charged. Therefore, an approach involving electrostatic interactions between the polymer assemblies and the substrate was investigated in order to promote vesicle spreading. First, chronoamperograms were recorded after spreading of vesicles on gold and capacitive and charge transfer currents compared (Figure S6). Chronoamperometry results shown in Figure S6 exhibited that the highest current can be achieved when a potential of -0.75 V was applied. Additionally, comparing the SFM images and considering the fact that zeta potential analysis revealed that the block copolymer vesicles are positively charged, a potential of -0.75 was chosen as step potential to be applied to assist the polymersomes spreading. 28 ACS Paragon Plus Environment

Page 29 of 48 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


In fact, we varied the charge of the gold substrate by applying voltage to influence the adsorption and subsequent deposition behavior. In order to maximize the spreading process and inhibit possible damaging of the resulting layers by overoxidation, both deposited films from spreading of polymersomes and synthosomes were prepared under a constant potential for a period of 1000 s. In this set of experiments, vesicles were first prepared from A14-B65-A14 copolymer followed by subjecting them into a threeelectrode system. Films obtained from optimized spreading conditions on gold electrodes were analyzed with SFM. Apart from occasional gaps, the surface coverage depicted in Figure 5, which

corresponds

to

electro-assisted

vesicle

spreading

performed

via

chronoamperometry at potential of -750 mV, increased significantly compared to the obtained spin-coated gold substrate (shown in Figure 4). In conclusion, spreading of PMOXA-b-PDMS-b-PMOXA self-assemblies under the potential-assisted conditions mentioned above yielded high surface coverages. Non-electrostatic interactions between the gold surface and the A14-B65-A14 copolymers are not strong enough to induce spreading of the polymer superstructures to form homogeneous bilayers over large substrate areas (see Figure 4). Figure 5 represents an 3.3 µm × 3.3 μm overview of the deposited structures in topography and phase mode achieved via applying a potential on the gold surface where a more homogenous coverage is obtained when compared to the other spreading methods such as spin-coating. Enlarged 10.0 µm × 10.0 μm overviews of Figures 5a and d are shown in Figures S8a and b. 29 ACS Paragon Plus Environment


Page 30 of 48

One interesting observation is that when polymersomes are spread on gold substrate by applying potential, not only the non-covered areas compared to spreading via spincoating (performed after addition of salt) are much less, but also non-spread vesicles (white spots in SFM pictures) are significantly reduced in number. This almost complete restructuring of the copolymer chains from vesicular form toward aligned self-assemblies on the gold surface pointed out that in such system the electrostatic interactions between charged A14-B65-A14 compartments and the charged gold surface were much stronger than the inter-chain interactions accountable for the conservation of the vesicular structures. Furthermore, the phase images depicted in Figures 5c and f confirm the presence of a soft and uniform film with identical properties of the continuous polymer layer when FhuA Δ1-160-loaded vesicular compartments are electro-deposited and spread on the gold surface.


Page 31 of 48 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


Figure 5. Surface characterization of coated gold with spread a) and c) polymersomes, d) and f) synthosomes. Coated substrates were prepared via applying a potential of 0.75 V for 1000 s. In a) and d) topography, c) and f) 3D SFM phase modes are presented, respectively. The height profiles b) and e) were obtained from the indicated lines in images a) and d). FhuA Δ1-160 and block copolymer A14-B65-A14 were used.

To further analyze the films of spread vesicles on the gold surface, wettability analysis on the A14-B65-A14 membrane-coated substrates was carried out. Details of this method are provided in Supporting Information and illustrated in Table S1 and Figure S10. Contact angle results of gold substrates coated with spread polymersomes and synthosomes revealed that the incorporation of hydrophilic cavities of FhuA Δ1-160



Page 32 of 48

channels across the block copolymer layer leads to a decrease in contact angle from 41.2ᵒ (spread polymersome-coated gold) to 19.1ᵒ (spread synthosome-coated gold). The influence of incorporated protein channels in the electrochemical properties of the electrodeposited FhuA Δ1-160-loaded A14-B65-A14 composite was investigated by cyclic voltammetry (CV) measurements (Figures S7a and b). As illustrated in Figure S7a, both anodic and cathodic processes were influenced when the electrodes coated with spread polymersomes and synthosomes were incubated in the sodium phosphate buffer. We assume that the decrease in both anodic and cathodic current can be attributed to an insulate layer which is formed after the deposition and spreading of polymer vesicles on the conductive gold electrode. Although the electrodeposited and spread A14-B65-A14 phase reveals a compact matrix, FhuA Δ1-160 functionality is expected to enhance the passive ion transport across the formed polymer membranes as the incorporated proteins can act as permanent ion channels from the electrolyte towards the electrode surface and vice versa. The same behavior can be observed from obtained cyclic voltammograms shown in Figure S7b in presence of lithium and perchlorate ions and increase in capacitive currents when FhuA is incorporated within the solid-supported block copolymer membrane. Moreover, the cyclic voltammogram recorded for ionic exchange of Li+ and ClO4¯ ions on spread polymersomes-coated gold electrodes (without FhuA Δ1-160) shows an anodic process with a peak potential of around 1.3 V. The current density of this anodic peak increases in case of using synthosomes-coated gold substrate as working electrode (incorporated with FhuA Δ1-160). This observation shows that the electroactivity of 32 ACS Paragon Plus Environment

Page 33 of 48 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


the spread synthosomes-coated gold is higher than that of spread polymersomes-coated gold electrode. This result, which is practically independent of the scan rate (results at higher scan rate are not shown), evidences the role played by the pore-forming protein in maintaining a flow of ions in and out of the deposited polymer matrix. The first cathodic peak at potential of around 0.7 V illustrated in Figure S7b does not shift in all cases. However, it is very interesting that the two observed cathodic peaks are more separated for electrodeposited polymersomes than for synthosomes. This cathodic process at the potential of around 0.7 V can be associated with the redox reaction occurring in buffer as it can be observed almost in the same potential range of all cyclic voltammograms depicted in Figure S7a. The second appearing cathodic peak at around 0.3 V shows, most probably, the ability of the obtained film to exchange Li+ from the electrolyte in and out of the spread membranes. As depicted in cyclic voltammograms in Figure S7b, the cation-exchange ability takes place at more negative potentials for deposited polymersomes compared to synthosomes. The presence of pores provided by FhuA Δ1-160 embedded within the polymer layer forms pathways for such ion-exchange. This leads to flow of cations through the pores at higher cathodic potentials revealing that less energy is needed for ion movement within the FhuA Δ1-160-loaded polymer matrix.



Page 34 of 48

Cysteine accessibility analysis of FhuA ΔCVFtev after spreading of synthosomes on gold Overall, calcein release and ITC results as well as DLS and topographical observations revealed that FhuA Δ1-160 was incorporated within the copolymer-compartments. More specifically, successful FhuA Δ1-160 incorporation could be indicated on gold via electrochemical properties of the electrodeposited FhuA Δ1-160-loaded A14-B65A14 composite determined by cyclic voltammetry assays. To further illustrate the density and distribution of incorporated protein channels across the spread films on the gold substrate, the variant FhuA ΔCVFtev, which can be dyelabeled inside the channel, was designed. FhuA ΔCVFtev was obtained by the deletion of the cork domain from FhuA WT and insertion of Tobacco Etch Virus (TEV)-protease cleavage sites in loops 7 and 8. The lysine at position 545, which is located inside of the β-barrel domain, was substituted by cysteine for covalent binding, and mutations N548V, E501F were performed to improve the binding accessibility.53 ThioGlo1® (denoted as ThioGlo) can be linked to a free cysteine placed at position 545 (C545) in FhuA ΔCVFtev as demonstrated earlier.53 Occupation of C545 via anchoring to the ThioGlo was already confirmed by titration with ThioGlo.61 This fluorescent dye has a maleimide function to selectively bind to cysteine and generates a strong fluorescence signal. Linkage of ThioGlo was performed on the films formed on the substrates followed by extensive washing to remove non-covalently bound dye. As represented in Figure 6a, no fluorescent signal can be detected in case of spread polymersomes, demonstrating that the dye can’t be linked when there is no FhuA ΔCVFtev incorporated 34 ACS Paragon Plus Environment

Page 35 of 48 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


within the polymer membrane. However, a clear fluorescence can be detected in the films from spreading synthosomes via spin-coating as well as potential-assisted deposition (Figure 6b and c). The visualization of fluorescence via confocal microscopy supports the presence of accessible cysteine which is located in the channel interior, thus verifying the porous structure of the obtained films. Comparison of the images shown in Figure 6b and c provides a clear indication that FhuA ΔCVFtev is more homogenously distributed across the polymer membrane when the spreading is performed via applying potential rather than via spin-coating. Spreading of synthosomes on glass and silicon wafers was likewise analyzed by linkage of ThioGlo (Figure S5). Similarly, no homogeneous film was found highlighting the superior spreading on gold via applying potential. The homogeneous FhuA ΔCVFtev distribution in the films obtained via electro-assisted approach supports the SFM surface analysis and is, furthermore, in good agreement with the amount of incorporated FhuA Δ1-160 in the films estimated via BCA assay (Figure S9).

Figure 6. Confocal microscopy images of the coated gold surfaces prepared via spreading of A14-B65-A14 a) polymersomes, b) synthosomes via spin-coating and c) synthosomes via applying potential of -0.75 V for 1000 s. Imaging was performed after the fluorescent dye ThioGlo was added to the spread films followed by three washing steps. Successful FhuA ΔCVFtev incorporation is visualized by ThioGlo fluorescence (exaction: 379 nm, emission: 513 nm). 35 ACS Paragon Plus Environment


Page 36 of 48

Impedance spectroscopic characterization of the spread vesicles on gold substrate EIS is a powerful technique for investigation of, e.g., electrode properties,62 biochemical reactions at functionalized polymer layers,63,

64

or block copolymers.65

Since the incorporation of membrane protein in block copolymer vesicles could endow ion pathways on block copolymer platform formed via spreading of polymer vesicles on conductive surfaces, we employed EIS analyses to investigate the passive ion transport response by subjecting the coated electrode in electrolytic medium with Li+ and ClO4¯ ions. Figure 7 shows the Nyquist plots for both spreading approaches of spin-coating and potential-assisted pathways. The spectral curves of each sample were fitted by a simple equivalent circuit inserted in Figure 7 consisting of a resistor Rs representing the electrolyte solution resistance and a resistor Rf simulating the film resistance and therefore also the resistance of the pores inside the film. The electrical double layer at the coated substrate-electrolyte interface was represented by a constant phase element CPEd and the capacitive properties of the copolymer film were simulated by CPEf. The constant phase element CPE is defined by two parameters CPE-T standing for double layer charge capacity and CPE-P defining the non-ideal behavior of the capacity, which is given by the electrode geometry, porosity and surface reactivity.66 The ideal capacitor is given by CPE-P of 1. In opposite, the resistive character shows the CPE-P of 0. In fact, observing the obtained spectra, we can notice depressed semicircle shapes after fitting. This behavior is normally expected for electrodes made of composite


Page 37 of 48 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


materials.67 For this reason, a constant phase element was used in the circuit to reflect the properties of a non-ideal electrode surface.68 A detailed overview of the values and relative errors of the equivalent circuit elements is given in Table S2 in the Supporting Information. The fitting quality was kept with maximal error of 6%. However, in this work we focused on the variation of charge transfer resistance (Rf) between the electrolyte and the electrode surface, since this parameter reflects the ion mobility at the electrode surface and the copolymer vesicles.

Figure 7. Nyquist plots of spread polymersomes-coated electrode achieved via spincoating (black), applying potential (blue) and spread synthosomes-coated electrode obtained from spin-coating (green) and applying potential (red) in a sodium phosphate buffer (pH 7.4, 10 mM) solution containing 0.1 M of LiClO4 (Frequency range: 10510-2 Hz). An electrical equivalent circuit model used for fitting of the data is shown in insert. FhuA Δ1-160 and block copolymer A14-B65-A14 were used. This value in the spectra is reflected by the diameter of the semicircle after fitting. Based on the estimate of the semicircle diameter, the parameter Rf of the copolymer-coated 37 ACS Paragon Plus Environment


Page 38 of 48

gold electrode achieved via spin-coating pathway is smaller (ca. 475 kOhm cm-2) than that obtained via electro-assisted approach (ca. 601 kOhm cm-2). Evidently, a smaller Rf indicates a faster electron transfer rate.69,

70

The electrode surface is physically

blocked by the initially immobilized block copolymers after spreading and access of electrolyte ions to the conductive surface of gold is hence impeded. The spreading of synthosomes with incorporated FhuA Δ1-160 leads to the formation of a polymer layer on the gold containing open channels provided by accommodated membrane protein across the block copolymer network. Rf values for membranes made of synthosomes are lower than those formed by polymersomes via both spreading approaches. This illustrates the superior potential of FhuA Δ1-160 channels distributed within the block copolymer coatings to provide pathways for electrons/ions and therefore increase ion mobility at the electrode surface. Moreover, comparing the two fabrication methods, the Rf of the spread synthosomes via applying potential is lower than the Rf of the spin-coated film (ca. 203 kOhm cm-2 vs. ca. 236 kOhm cm-2). This is additionally supported by the fluorescence results illustrated in Figure 6 which show better efficiency of the electrically driven spreading. Thus, the application of an electrical potential during the deposition of synthosomes and polymersomes means great advance in the field of solid-supported membrane preparation.


Page 39 of 48 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


CONCLUSION The reconstitution of the membrane protein FhuA Δ1-160 into vesicle membranes made from the triblock copolymer PMOXA14-b-PDMS65-b-PMOXA14 was monitored and validated via calcein release, high-sensitivity titration calorimetry in real time, DLS, and zeta potential measurements. Thereby, the ITC method proved to be a reliable technique to follow the insertion of membrane proteins into the polymersome membrane. To our knowledge, this is the first study of membrane protein insertion into block copolymer vesicles via isothermal titration calorimetry. Our study revealed that the insertion of FhuA Δ1-160 into the polymer membrane is an endothermic process which is dominated by the hydrophobic nature of interaction between the hydrophobic region of the membrane protein and the hydrophobic block of the copolymer. The spreading of the polymer vesicles was optimized on silica wafers, glass and gold through the addition of NaCl, spin-coating and heating subsequent to film formation. Interestingly, our new approach of potential-driven spreading of synthosomes enabled to prepare uniform solid-supported hybrid membranes. The polymersomes have a positive zeta potential in aqueous solution. Spreading on gold substrates with applied potential of -0.75 V for 1000 s showed to be suitable to form more homogeneous films. The advantage of the potential-assisted spreading became particularly evident with the linkage of the fluorescent dye ThioGlo to a free cysteine inside the channel of the FhuA ΔCVFtev variant which demonstrated a much more homogeneous distribution of the protein in the membrane on the gold solid support. Furthermore, the labeling suggests that the protein pore structure is still intact after spreading. 39 ACS Paragon Plus Environment


Page 40 of 48

Advantages of potential-driven compared to spreading pathways by spin-coating and dip-coating are the ability to produce more homogeneous and highly reproducible films, easy processability with a standard electrochemical setup and short deposition times. Therefore, this strategy can be adapted to different block copolymers and proteins as well as various conductive materials as solid support. We believe that the potentialassisted spreading strategy using a defined potential might open up novel opportunities in the development of air-stable biosensors and biomimetic membranes for functional separation processes as well as downstream-processing. ASSOCIATED CONTENT Supporting information (SI) The Supporting Information is available free of charge on the ACS Publications website at DOI: Methods: Expression, extraction and refolding of FhuA Δ1-160 and FhuA ΔCVFtev variants, Cryo-Transmission Electron Microscopy (Cryo-TEM), Static contact angle determination, Protein Concentration Determination via Bicinchoninic Acid (BCA) assay. Results: Expression and extraction of FhuA Δ1-160 and FhuA ΔCVFtev variants, Preparation of polymersomes from different block length and characterization via cryoTEM, Spreading of A19-B65-A19 copolymer vesicles on silicon-wafer and glass slides,

Control voltammograms of electro-assisted spreading of A14-B65-A14 vesicles on gold substrate, Surface characterization of copolymer-coated gold via electro-assisted spreading of A14-B65-A14 vesicles, Concentration evaluation of incorporated FhuA


Page 41 of 48 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


Δ1-160 in polymer platform by BCA assay, Wettability studies on the A14-B65-A14 copolymer-coated substrates, Detailed explanation of Impedance spectroscopic characterization of the spread vesicles on gold substrate. References. AUTHOR INFORMATION Corresponding Author [email protected] Present Addresses 6 Tianjin

Institute of Industrial Biotechnology, Chinese Academy of Sciences, 32 West

7th Ave., Tianjin 300308, China Author Contributions ‡ Tayebeh Mirzaei Garakani and Zhanzhi Liu are joint first authors Notes: The authors declare no competing financial interest. ACKNOWLEDGMENT Tayebeh Mirzaei Garakani thanks Mehdi Rashvand Avei for the fruitful scientific discussion. The authors would like to thank Dr. Khosrow Rahimi for help with cryoTEM measurements. Tayebeh Mirzaei Garakani and Zhanzhi Liu acknowledge the Alexander von Humboldt-Stiftung as well as China Scholarship Council (CSC No. 201306350039), respectively, for financial support. The Bundesministerium für Bildung und Forschung (BMBF) is kindly acknowledged for financial support in the framework of the BMBF-Forschertandem “Chiral Membranes” (Förderkennzeichen: 031A164 and 031B0559). Part of this work was performed at the Center for Chemical 41 ACS Paragon Plus Environment


Page 42 of 48

Polymer Technology (CPT) unit of DWI, which was supported by the EU and the federal state of North Rhine–Westphalia (grant EFRE 30 00 883 02). Part of this work was further supported as Fraunhofer High Performance Center for Functional lntegration in Materials.

ABBREVIATIONS PMOXA-b-PDMS-b-PMOXA:

poly(2-methyloxazoline)-block-

poly(dimethylsiloxane)-block-poly(2-methyloxazoline), FhuA: ferric hydroxamate uptake protein component A, PB-b-PEO: polybutadiene-block-polyethyleneoxide, AQP0: aquaporin-0, ITC: isothermal titration calorimetry, E. coli: Escherichia coli, CRP: controlled radical polymerization, NIPAAm: N-isopropylacrylamide, MPD: 2methyl-2, 4-pentanediol, DLS: Dynamic Light Scattering, :CD: Circular dichroism, SFM: Scanning Force Microscopy, CLSM: Confocal Laser Scanning Microscopy, EIS: Electrochemical Impedance Spectroscopy, Cryo-TEM: Cryo-Transmission Electron Microscopy, BCA: Protein Concentration Determination via Bicinchoninic Acid.

REFERENCES 1. Rigaud, J. L.; Levy, D. Reconstitution of Membrane Proteins into Liposomes. Methods Enzymol. 2003, 372, 65-86. 2. Güven, A.; Dworeck, T.; Fioroni, M.; Schwaneberg, U. Residue K556-A Light Triggerable Gatekeeper to Sterically Control Translocation in FhuA. Adv. Eng. Mater. 2011, 13, B324-B329. 3. Krewinkel, M.; Dworeck, T.; Fioroni, M. Engineering of an E. coli Outer Membrane Protein FhuA with Increased Channel Diameter. J. Nanobiotechnology 2011, 9, 33.


Page 43 of 48 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


4. Sperandio, V. Purification and Liposome Reconstitution of a Bacterial MultiTransmembrane Domain Protein for Detection of Autokinase Activity in Response to Signals. Protoc. Exch. 2012, DOI:10.1038/protex.2012.056. 5. Discher, B. M.; Won, Y. Y.; Ege, D. S.; Lee, J. C.; Bates, F. S.; Discher, D. E.; Hammer, D. A. Polymersomes: Tough Vesicles made from Diblock Copolymers. Science 1999, 284, 1143-1146. 6. Kumar, M.; Grzelakowski, M.; Zilles, J.; Clark, M.; Meier, W. Highly Permeable Polymeric Membranes based on the Incorporation of the Functional Water Channel Protein Aquaporin Z. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 20719-20724. 7. Zhang, X.; Fu, W.; Palivan, C. G.; Meier, W. Natural Channel Protein Inserts and Functions in a Completely Artificial, Solid-Supported Bilayer Membrane. Sci. Rep. 2013, 3, 2196. 8. Meier, W.; Nardin, C.; Winterhalter, M. Reconstitution of Channel Proteins in (Polymerized) ABA Triblock Copolymer Membranes. Angew. Chem. Int. Ed. Engl. 2000, 39, 4599-4602. 9. Nallani, M.; Andreasson-Ochsner, M.; Tan, C. W.; Sinner, E. K.; Wisantoso, Y.; Geifman-Shochat, S.; Hunziker, W. Proteopolymersomes: In Vitro Production of a Membrane Protein in Polymersome Membranes. Biointerphases 2011, 6, 153-157. 10. Klermund, L.; Poschenrieder, S. T.; Castiglione, K. Biocatalysis in Polymersomes: Improving Multienzyme Cascades with Incompatible Reaction Steps by Compartmentalization. ACS Catal. 2017, 7, 3900-3904. 11. González-Pérez, A.; Persson, K. M.; Taboada, P. Structural Stability of SoPIP2;1 aquaporin under Reconstitution in Polymersomes. J. Mol. Liq. 2018, 257, 26-31. 12. Baumann, P.; Tanner, P.; Onaca, O.; Palivan, C. G. Bio-Decorated Polymer Membranes: A New Approach in Diagnostics and Therapeutics. Polymers 2011, 3, 173192. 13. Garni, M.; Thamboo, S.; Schoenenberger, C. A.; Palivan, C. G. Biopores/Membrane Proteins in Synthetic Polymer Membranes. Biochim. Biophys. Acta 2017, 1859, 619-638. 14. Itel, F.; Najer, A.; Palivan, C. G.; Meier, W. Dynamics of Membrane Proteins within Synthetic Polymer Membranes with Large Hydrophobic Mismatch. Nano lett. 2015, 15, 3871-3878. 15. Kumar, M.; Habel, J. E. O.; Shen, Y.; Meier, W. P.; Walz, T. High-Density Reconstitution of Functional Water Channels into Vesicular and Planar Block Copolymer Membranes. J. Am. Chem. Soc. 2012, 134, 18631-18637. 16. Nallani, M.; Benito, S.; Onaca, O.; Graff, A.; Lindemann, M.; Winterhalter, M.; Meier, W.; Schwaneberg, U. A Nanocompartment System (Synthosome) designed for Biotechnological Applications. J. Biotechnol. 2006, 123, 50-59. 17. Huang, X.; Li, M.; Green, D. C.; Williams, D. S.; Patil, A. J.; Mann, S. Interfacial Assembly of Protein-Polymer Nano-Conjugates into Stimulus-Responsive Biomimetic Protocells. Nat. Commun. 2013, 4, 2239. 18. Huang, X.; Patil, A. J.; Li, M.; Mann, S. Design and Construction of Higher-Order Structure and Function in Proteinosome-Based Protocells. J. Am. Chem. Soc. 2014, 136, 9225-9234. 43 ACS Paragon Plus Environment


Page 44 of 48

19. Tang, T. D.; Cecchi, D.; Fracasso, G.; Accardi, D.; Coutable-Pennarun, A.; Mansy, S. S.; Perriman, A. W.; Anderson, J. L. R.; Mann, S. Gene-Mediated Chemical Communication in Synthetic Protocell Communities. ACS Synth. Biol. 2018, 7, 339346. 20. Keller, S.; Heerklotz, H.; Jahnke, N.; Blume, A. Thermodynamics of Lipid Membrane Solubilization by Sodium Dodecyl Sulfate. Biophys. J. 2006, 90, 4509-4521. 21. Heerklotz, H.; Tsamaloukas, A. D.; Keller, S. Monitoring Detergent-Mediated Solubilization and Reconstitution of Lipid Membranes by Isothermal Titration Calorimetry. Nat. Protoc. 2009, 4, 686-697. 22. Liu, J.; Postupalenko, V.; Duskey, J. T.; Palivan, C. G.; Meier, W. pH-Triggered Reversible Multiple Protein–Polymer Conjugation Based on Molecular Recognition. J. Phys. Chem. B 2015, 119, 12066-12073. 23. Thiele, M.; Davari, M. D.; Ko, M.; Hofmann, I. N.; Junker, O.; Garakani, T. M.; Vojcic, L.; Fitter, J.; Schwaneberg, U. Enzyme−Polyelectrolyte Complexes Boost the Catalytic Performance of Enzymes. ACS Catal. 2018, 8, 10876−10887. 24. Rajarathnam, K.; Rosgen, J. Isothermal Titration Calorimetry of Membrane Proteins- Progress and Challenges. Biochim. Biophys. Acta 2014, 1838, 69-77. 25. Bončina, M.; Lah, J.; Reščič, J.; Vlachy, V. Thermodynamics of the Lysozyme−Salt Interaction from Calorimetric Titrations. J. Phys. Chem. B 2010, 114, 4313-4319. 26. Jahnke, N.; Krylova, O. O.; Hoomann, T.; Vargas, C.; Fiedler, S.; Pohl, P.; Keller, S. Real-Time Monitoring of Membrane-Protein Reconstitution by Isothermal Titration Calorimetry. Anal. Chem. 2014, 86, 920-927. 27. Ferguson, A. D.; Hofmann, E.; Coulton, J. W.; Diederichs, K.; Welte, W. Siderophore-Mediated Iron Transport: Crystal Structure of FhuA with Bound Lipopolysaccharide. Science 1998, 282, 2215-2220. 28. Locher, K. P.; Rees, B.; Koebnik, R.; Mitschler, A.; Moulinier, L.; Rosenbusch, J. P.; Moras, D. Transmembrane Signaling across the Ligand-Gated FhuA Receptor: Crystal Structures of Free and Ferrichrome-Bound States Reveal Allosteric Changes. Cell 1998, 95, 771-778. 29. Bonhivers, M.; Plancon, L.; Ghazi, A.; Boulanger, P.; le Maire, M.; Lambert, O.; Rigaud, J. L.; Letellier, L. FhuA, an Escherichia coli Outer Membrane Protein with a Dual Function of Transporter and Channel which Mediates the Transport of Phage DNA. Biochimie 1998, 80, 363-369. 30. Tenne, S. J.; Schwaneberg, U. First Insights on Organic Cosolvent Effects on FhuA Wildtype and FhuA Δ1-159. Int. J. Mol. Sci. 2012, 13, 2459-2471. 31. Muhammad, N.; Dworeck, T.; Fioroni, M.; Schwaneberg, U. Engineering of the E. coli Outer Membrane Protein FhuA to Overcome the Hydrophobic Mismatch in Thick Polymeric Membranes. J. Nanobiotechnology 2011, 9, 8. 32. Charan, H.; Kinzel, J.; Glebe, U.; Anand, D.; Garakani, T. M.; Zhu, L.; Bocola, M.; Schwaneberg, U.; Böker, A. Grafting PNIPAAm from β-Barrel Shaped Transmembrane Nanopores. Biomaterials 2016, 107, 115-123.


Page 45 of 48 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


33. Liu, Z.; Ghai, I.; Winterhalter, M.; Schwaneberg, U. Engineering Enhanced Pore Sizes using FhuA Δ1-160 from E. coli Outer Membrane as Template. ACS Sens. 2017, 2, 1619-1626. 34. Charan, H.; Glebe, U.; Anand, D.; Kinzel, J.; Zhu, L.; Bocola, M.; Garakani, T. M.; Schwaneberg, U.; Böker, A. Nano-thin Walled Micro-Compartments from Transmembrane Protein-Polymer Conjugates. Soft Matter 2017, 13, 2866-2875. 35. Anand, D.; Dhoke, G. V.; Gehrmann, J.; Garakani, T. M.; Davari, M. D.; Bocola, M.; Zhu, L.; Schwaneberg, U. Chiral Separation of D/L-Arginine with Whole Cells through an Engineered FhuA Nanochannle. Chem. Commun. 2019, 55, 5431-5434. 36. Nallani, M.; Onaca, O.; Gera, N.; Hildenbrand, K.; Hoheisel, W.; Schwaneberg, U. A Nanophosphor-Based Method for Selective DNA Recovery in Synthosomes. Biotechnol. J. 2006, 1, 828-834. 37. Joëlle Ruff, A.; Arlt, M.; van Ohlen, M.; Kardashliev, T.; Konarzycka-Bessler, M.; Bocola, M.; Dennig, A.; B. Urlacher, V.; Schwaneberg, U. An Engineered Outer Membrane Pore Enables an Efficient Oxygenation of Aromatics and Terpenes. J. Mol. Catal. B: Enzym. 2016, 134, 285-294. 38. Onaca, O.; Sarkar, P.; Roccatano, D.; Friedrich, T.; Hauer, B.; Grzelakowski, M.; Guven, A.; Fioroni, M.; Schwaneberg, U. Functionalized Nanocompartments (Synthosomes) with a Reduction-Triggered Release System. Angew. Chem. Int. Ed. Engl. 2008, 47, 7029-7031. 39. Brian, A. A.; McConnell, H. M. Allogeneic Stimulation of Cytotoxic T Cells by Supported Planar Membranes. Proc. Natl. Acad. Sci. U.S.A. 1984, 81, 6159-6163. 40. Cremer, P. S.; Boxer, S. G. Formation and Spreading of Lipid Bilayers on Planar Glass Supports. J. Phys. Chem. B 1999, 103, 2554-2559. 41. Reimhult, E.; Höök, F.; Kasemo, B. Intact Vesicle Adsorption and Supported Biomembrane Formation from Vesicles in Solution: Influence of Surface Chemistry, Vesicle Size, Temperature, and Osmotic Pressure. Langmuir 2003, 19, 1681-1691. 42. Burgess, I.; Li, M.; Horswell, S. L.; Szymanski, G.; Lipkowski, J.; Majewski, J.; Satija, S. Electric Field-Driven Transformations of a Supported Model Biological Membrane—An Electrochemical and Neutron Reflectivity Study. Biophys. J. 2004, 86, 1763-1776. 43. Woodward, J. T.; Meuse, C. W. Mechanism of Formation of Vesicle Fused Phospholipid Monolayers on Alkanethiol self-Assembled Monolayer Supports. J. Colloid Interface Sci. 2009, 334, 139-145. 44. Lind, T. K.; Wacklin, H.; Schiller, J.; Moulin, M.; Haertlein, M.; Pomorski, T. G.; Cardenas, M. Formation and Characterization of Supported Lipid Bilayers Composed of Hydrogenated and Deuterated Escherichia coli Lipids. PloS one 2015, 10, DOI: 10.1371/journal.pone.0144671. 45. Belegrinou, S.; Menon, S.; Dobrunz, D.; Meier, W. Solid-Supported Polymeric Membranes. Soft Matter 2011, 7, 2202-2210. 46. White, S. H. A Study of Lipid Bilayer Membrane Stability using Precise Measurements of Specific Capacitance. Biophys. J. 1970, 10, 1127-1148. 47. Zhang, X.; Tanner, P.; Graff, A.; Palivan, C. G.; Meier, W. Mimicking the Cell Membrane with Block Copolymer Membranes. J. Polym. Sci. A 2012, 50, 2293-2318. 45 ACS Paragon Plus Environment


Page 46 of 48

48. Dorn, J.; Belegrinou, S.; Kreiter, M.; Sinner, E. K.; Meier, W. Planar Block Copolymer Membranes by Vesicle Spreading. Macromol. Biosci. 2011, 11, 514-525. 49. Wang, H.; Chung, T. S.; Tong, Y. W.; Jeyaseelan, K.; Armugam, A.; Chen, Z.; Hong, M.; Meier, W. Highly Permeable and Selective Pore-spanning Biomimetic Membrane Embedded with Aquaporin Z. Small 2012, 8, 1185-1190. 50. Palivan, C. G.; Goers, R.; Najer, A.; Zhang, X.; Car, A.; Meier, W. Bioinspired Polymer Vesicles and Membranes for Biological and Medical Applications. Chem. Soc. Rev. 2016, 45, 377-411. 51. Tsamaloukas, A. D.; Keller, S.; Heerklotz, H. Uptake and Release Protocol for Assessing Membrane Binding and Permeation by Way of Isothermal Titration Calorimetry. Nat. Protoc. 2007, 2, 695-704. 52. Traikia, M.; Warschawski, D. E.; Recouvreur, M.; Cartaud, J.; Devaux, P. F. Formation of Unilamellar Vesicles by Repetitive Freeze-Thaw Cycles: Characterization by Electron Microscopy and 31P-Nuclear Magnetic Resonance. Eur. Biophys. J. 2000, 29, 184-195. 53. Philippart, F.; Arlt, M.; Gotzen, S.; Tenne, S. J.; Bocola, M.; Chen, H.; Zhu, L.; Schwaneberg, U.; Okuda, J. A Hybrid Ring-Opening Metathesis Polymerization Catalyst Based on an Engineered Variant of the β-Barrel Protein FhuA. Chem. Eur. J. 2013, 19, 13865-13871. 54. Discher, D. E.; Ahmed, F. Polymersomes. Annu. Rev. Biomed. Eng. 2006, 8, 323341. 55. Kinzel, J.; Sauer, D. F.; Bocola, M.; Arlt, M.; Mirzaei Garakani, T.; Thiel, A.; Beckerle, K.; Polen, T.; Okuda, J.; Schwaneberg, U. 2-Methyl-2,4-pentanediol (MPD) Boosts as Detergent-Substitute the Performance of β-Barrel Hybrid Catalyst for Phenylacetylene Polymerization. Beilstein J. Org. Chem. 2017, 13, 1498-1506. 56. Pourhosseini, P. S.; Saboury, A. A.; Najafi, F.; Sarbolouki, M. N. Interaction of Insulin with a Triblock Copolymer of PEG-(Fumaric-Sebacic acids)-PEG: Thermodynamic and Spectroscopic Studies. Biochim. Biophys. Acta 2007, 1774, 12741280. 57. De, M.; You, C. C.; Srivastava, S.; Rotello, V. M. Biomimetic Interactions of Proteins with Functionalized Nanoparticles: A Thermodynamic Study. J. Am. Chem. Soc. 2007, 129, 10747-10753. 58. Wang, S.; Chen, K.; Xu, Y.; Yu, X.; Wang, W.; Li, L.; Guo, X. Protein Immobilization and Separation Using Anionic/Cationic Spherical Polyelectrolyte Brushes Based on Charge Anisotropy. Soft Matter 2013, 9, 11276-11287. 59. Sperber, B. L. H. M.; Cohen Stuart, M. A.; Schols, H. A.; Voragen, A. G. J.; Norde, W.Overall Charge and Local Charge Density of Pectin Determines the Enthalpic and Entropic Contributions to Complexation with β-Lactoglobulin. Biomacromolecules 2010, 11, 3578-3583. 60. Jensen, W. B. Electronegativity from Avogadro to Pauling: Part 1: Origins of the Electronegativity Concept. J. Chem. Educ. 1996, 73, 11-20. 61. Wright, S. K.; Viola, R. E. Evaluation of Methods for the Quantitation of Cysteines in Proteins. Anal. Biochem. 1998, 265, 8-14. 46 ACS Paragon Plus Environment

Page 47 of 48 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


62. Lazar, J.; Schnelting, C.; Slavcheva, E.; Schnakenberg, U. Hampering of the Stability of Gold Electrodes by Ferri-/Ferrocyanide Redox Couple Electrolytes during Electrochemical Impedance Spectroscopy. Anal. Chem. 2016, 88, 682-687. 63. Lazar, J.; Park, H.; Rosencrantz, R. R.; Böker, A.; Elling, L.; Schnakenberg, U. Evaluating the Thickness of Multivalent Glycopolymer Brushes for Lectin Binding. Macromol. Rapid Commun. 2015, 36, 1472-1478. 64. Lazar, J.; Rosencrantz, R. R.; Elling, L.; Schnakenberg, U. Simultaneous Electrochemical Impedance Spectroscopy and Localized Surface Plasmon Resonance in a Microfluidic Chip: New Insights into the Spatial Origin of the Signal. Anal. Chem. 2016, 88, 9590-9596. 65. Wagner, T.; Lazar, J.; Schnakenberg, U.; Böker, A. In situ Electrochemical Impedance Spectroscopy of Electrostatically Driven Selective Gold Nanoparticle Adsorption on Block Copolymer Lamellae. ACS Appl. Mater. Interfaces 2016, 8, 27282-27290. 66. Nahir, T. M. Impedance Spectroscopy: Theory, Experiment, and Applications, 2nd ed; John Wiley & Sons, Inc.: Hoboken , 2005. 67. Bonanni, A.; Pividori, M. I.; Campoy, S.; Barbe, J.; del Valle, M. Impedimetric Detection of Double-Tagged PCR Products Using Novel Amplification Procedures Based on Gold Nanoparticles and Protein G. Analyst 2009, 134, 602-608. 68. Patolsky, F.; Filanovsky, B.; Katz, E.; Willner, I. Photoswitchable Antigen−Antibody Interactions Studied by Impedance Spectroscopy. J. Phys. Chem. B 1998, 102, 10359-10367. 69. Ding, C.; Liu, H.; Zhu, Y.; Wan, M.; Jiang, L. Control of Bacterial Extracellular Electron Transfer by a Solid-State Mediator of Polyaniline Nanowire Arrays. Energy Environ. Sci. 2012, 5, 8517-8522. 70. Schröder, U.; Nießen, J.; Scholz, F. A Generation of Microbial Fuel Cells with Current Outputs Boosted by More Than One Order of Magnitude. Angew. Chem. Int. Ed. Engl. 2003, 42, 2880-2883.



Page 48 of 48

Table Of Contents


In Situ Monitoring of Membrane Protein Insertion into Block Copolymer

Recommend Documents