Visualizing the Solid–Liquid Interface of Conjugated Copolymer Films

Subscriber access provided by NORTH CAROLINA A&T UNIV

Article

Visualizing the Solid-Liquid Interface of Conjugated Copolymer Films Using Fluorescent Liposomes Yi Zhang, Achilleas Savva, Shofarul Wustoni, Adel Hama, Iuliana Petruta Maria, Alexander Giovannitti, Iain McCulloch, and Sahika Inal ACS Appl. Bio Mater., Just Accepted Manuscript • DOI: 10.1021/acsabm.8b00323 • Publication Date (Web): 15 Oct 2018 Downloaded from http://pubs.acs.org on October 17, 2018

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 21 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 Bio Materials

Visualizing the Solid-Liquid Interface of Conjugated Copolymer Films Using Fluorescent Liposomes Yi Zhang‡, Achilleas Savva‡, Shofarul Wustoni‡, Adel Hama‡, Iuliana P. Maria§, Alexander Giovannitti§, Iain McCulloch§⊥ and Sahika Inal‡ ‡

Biological and Environmental Sciences and Engineering Division, King Abdullah University of

Science and Technology (KAUST), Thuwal 23955‐6900, Kingdom of Saudi Arabia §

Department of Chemistry and Centre for Plastic Electronics, Imperial College London, London

SW7 2AZ, UK ⊥Physical

Science and Engineering Division, KAUST, Thuwal 23955-6900, Kingdom of Saudi

Arabia KEYWORDS: conjugated polymer, lipid, ethylene glycol, fluorescence recovery after photobleaching, n-type organic semiconductor

ABSTRACT: Conjugated polymers are promising engineering tools for establishing bilateral electrical communication with living systems. The free energy of their films, the roughness and charge density play a major role in determining their interactions with lipid bilayers which form the membrane barrier around every living cell allowing for molecular exchange with the extracellular environment. In this work, we investigate lipid bilayer formation from synthetic lipid vesicles (liposomes) on a series of amphiphilic copolymer films based on naphthalene 1,4,5,8 tetracarboxylic diimide bithiophene (NDI-T2) backbone where the alkyl side chain is gradually exchanged for an ethylene glycol-based side chain. As the concentration of ethylene glycol in the composition changes, the surface energy of the films varies drastically which, in

ACS Paragon Plus Environment

1

ACS Applied Bio Materials 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 21

turn, effects the interactions with liposomes. By imaging the interactions of fluorophore-labeled liposomes with these surfaces via a fluorescence microscope, we show that the films can be cast such that ethylene glycol-rich regions, which liposomes favor, are accumulated on the surface and extract information on the wettability behavior that has not been possible using other surface sensitive techniques. This approach uncovers the solid/liquid interface of a promising class of electron transporting conjugated polymer films and suggests synthetic strategies to maximize the number of lipid-polymer contacts for the formation of supported lipid bilayers.


2

Page 3 of 21 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


Introduction Conjugated polymers are excellent candidates for interfacing with biomolecules and cells due to their softness, low-to-absent toxicity, as well as their ability to process ionic (biological) signals.1-6 They have been used as smart, mechanical supports for cells in the form of scaffolds for regulating and reporting cell growth elecrically.7-10 It is well established that the surface properties such as charge, morphology, topography and wettability of the conjugated polymer films play a critical role in how living systems interact with the synthetic material surface, governing cell adhesion and spreading as well as specific cell responses.11-14 The surface properties are even more detrimental for the formation of supported lipid bilayers.15 The lipid bilayer is the essential structure of the cell membrane, which serves as a barrier between the interior and exterior of the cell while selectively permeating small compounds (e.g. ions) through embedded membrane proteins.16 Studying the lipid bilayer thus gives access to information about how cells communicate with the outside world and the function of membrane proteins, for instance their interactions with pathogens. We and others have shown that conjugated polymers can be used to interface with lipid assemblies.15,17,18 A lipid bilayer assembled on a substrate, i.e. a supported lipid bilayer (SLB), is typically insulating while its conductance can be modulated by the activity of the membrane proteins, for instance due to the transport of ions through ion channels or by protein binding events. Since the lipid bilayer formed on top of the conjugated polymer film blocks the vertical ion flow from the electrolyte, changes in membrane conductance directly affect the electrical properties of the film underneath. Thus, conjugated polymer based electronic platforms provide a label-free approach for monitoring the integrity of the bilayer against certain compounds as well as studying the functionality of the membrane proteins in vitro in native or model environments.


3


Page 4 of 21

In general, SLBs are formed via vesicle fusion on hydrophilic and smooth surfaces such as those of glass,19-20 silica,21 and mica.22 When the surface is incubated with vesicle suspension, upon sufficient coverage, vesicle adsorption, deformation, rupture and spreading occur followed by SLB formation.23 For SLB formation, surface wetting is a prerequisite, as for the adherence of cell adhesion proteins to a substrate.23-24 Due to their typical hydrophobicity, conjugated polymers may not be regarded as the best candidates for interfacing lipid bilayers. Through synthetic efforts, however, their chemical structure can be altered based on the application requirements. For instance, conjugated polymers bearing biologically active chemical units (e.g. amino acids, amine or carboxylic acid residues) have enabled specific interactions with cells or the extracellular matrix components.25-27 Over the past decade, another approach to alter the hydrophobicity of the films without compromising the electrical properties has been side chain functionalization of hole (p-type) as well as electron (n-type) transporter conjugated polymers with hydrophilic ethylene glycol (EG) side chains.28-31 The functionalization rendered the polymers useful for applications at the electrolyte interface. Because of EG side chains, the polymers can uptake water (swell like a hydrogel),28,31 which facilitates ion transport along the hydrated chains and thus electrochemical activity in aqueous electrolytes. As such, they became interesting to be applied in state-of-the-art devices of bioelectronics such as the organic electrochemical transistor (OECT), an electrolyte gated transistor used as an amplifying transducer of biological events.32 We have recently shown the use of OECTs comprising an n-type copolymer functionalized with EG side chains for enzymatic sensing of lactate.33 The high performance of these sensors was reasoned with the EG-induced hydrophilicity of the surface which caused close interactions with the enzyme, facilitating for the first time direct electrical communication of an n-type conjugated


4

Page 5 of 21 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


polymer with an enzyme. Such n-type OECTs are particularly interesting to study lipid bilayer functionality, e.g., opening of an ion channel for the transport of cations, as they respond specifically to cation fluxes in the electrolyte.30,31 For the formation of lipid bilayers on these surfaces, however, understanding and controlling the impact of the side chains on surface properties and organization within the films is crucial. Furthermore, to be able to extend the applications of this promising group of polymers at the biotic interface, a thorough characterization of solid/liquid interface is on demand. In this work, we shed light onto the surfaces of a series of recently developed n-type copolymers containing various EG chain densities,31,33 with the aim to explore their potential to study lipid bilayers. As the concentration of EG in the composition changes, the surface energy of the films varies drastically, affecting the vesicle/film interactions. We show that the films can be cast such that EG-rich regions, where liposomes favor, are exposed on the surface by imaging the interactions of fluorophore-labeled liposomes with these surfaces via a fluorescence microscope. As such, we extract information on the wettability behavior of the films that play a major role in vesicle adhesion and subsequently the formation of SLBs. Results and Discussion The polymer series comprises random copolymers of a dibromonaphthalene 1,4,5,8 tetracarboxylic diimide bithiophene (NDI-T2) backbone motif in which a fraction of the alkyl side chain on the imide group is gradually substituted for an EG-based side chain (Figure 1a). The EG side chain percentage in the polymer composition vary as 0, 10, 50, 90, and 100 with respect to the alkyl side chains (therefore those with 0 and 100% EG are homopolymers). The topography of each NDI-T2 based polymer film was investigated with AFM, in both dry and hydrated states as shown in Figure 1b. We observe nanofibrillar structures with a width on the


5


Page 6 of 21

order of tens of nanometers and distinct domains of several hundred nanometers for all samples characterized in air (Figure 1b, upper panel). The nanofibers are intertwined and inter-connected to each other. Films with 0% and 10% EG side chains (P-0 and P-10) present more aligned structures than the ones with higher EG content. The corresponding phase images reveal a similar trend where, accordingly, more ordered structures are observed for P-0 and P-10 (Figure S1). Overall, dry films show very fine textures but no domain-like structures that are observed in topography images. Notably, the differences in topography when switching from air to phosphate buffered saline (PBS) solution are maximized for the films with 90% and 100% EG side chains (P-90 and P-100), indicating increased incorporation of water into the films through hydrophilic regions (compare Figure 1b, upper and lower panels). The polymer films expand in volume when immersed in aqueous salt solution, particularly P-90 and P-100. We have previously shown that, in NaCl solution, below and at 50% of EG concentration in the structure (e.g. P-10 and P-50), the swelling percentage is smaller than 10%, while it increases dramatically for P-90 and P-100 (42% and 102% of swelling, respectively).31 TEM images reveal that the films have a fibrillar morphology (Figure S2). The fibrils are estimated to be within 20-30 nm range in width, consistent with the polymer length. We postulate that they are comprised of π-stacked polymer with a width dictated by the polymer backbone length, and a length dictated by the number of πstacked backbones. Similar structures with long-range ordering of the lamella were observed by other groups for NDI-T2 based polymers.34-35 The overlapping structure is highly evident for P100.


6

Page 7 of 21 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 1. (a) Chemical structure of the five NDI-T2 based polymers with 0, 10, 50, 90 and 100% EG chain percentages. [Reprinted with permission from ref 31. Copyright American Chemical Society.] (b) AFM topography images of the polymer films characterized in air (upper panel) and in PBS (lower panel). The arrow shows the increase in EG percentage in the chemical structure. Scale bar = 500 nm. The color scale is adjusted for each image to enable better comparison. Note the increase in roughness for P-100.

We investigate lipid bilayer formation on the films using fluorophore labeled zwitterionic lipids via fluorescence recovery after photobleaching (FRAP) measurements. This fluorescence microscope approach involves monitoring the recovery dynamics of a fluorescent region of a film once it is photobleached by laser irradiation. The technique is commonly used in assessing


7


Page 8 of 21

the formation of SLBs, as the rate of fluorescence recovery informs about the lateral mobility (diffusion coefficient and mobile fraction) of the lipid bilayer. Here, we use zwitterionic lipids including 1,2-diphytanoyl-sn-glycero-3-phosphocholine (DPhPC) and 1,2-diphytanoyl-snglycero-3-phosphoethanolamine (DPhPE) to prepare the liposomes (Figure 2a). These two phytanoyl lipids are common options for assembly of planar lipid membranes due to their high stability and low ion permeability.36 For FRAP, 4,4-difluoro-4-bora-3a,4a-diaza-s-indacene (BODIPY)

labeled

1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine

(DHPE)

was

incorporated into the mixture of DPhPC and DPhPE to prepare the vesicles. Copolymer films were incubated with this vesicle mixture overnight and rinsed thoroughly to remove the excess vesicles before imaging. Remarkably, each copolymer film exhibits distinct and reproducible fluorescent patterns when the vesicles are deposited on top (Figure 2b). The bright patterns undergo a remarkable change with the EG content, i.e. from island (P-10) to network (P-50) to partially connected island (P-90), while we do not observe any patterns for fully alkylated or glycolated polymers, P-0 or P-100. The proportion of these bright, i.e. fluorescent, regions increases with the percentage of EG chains in the copolymer. The surface coverage for the bright and dark regions on P-10, P-50 and P-90 surfaces is summarized in Table S1. The areas are bright because they are populated by the fluorescent vesicles, i.e., they offer the most favorable surface for the vesicles to adhere onto. We postulate that these patterns are formed due to the amphiphilic nature of the copolymers. As the size of the domains is not comparable to that of the chains, the images are representing only how the relatively more hydrophilic units might be oriented/assembled on the surface. Once the films are exposed to water, glycol/alkyl side chains cluster to form larger polar/non-polar regions at the surface, leading to areas with distinct wettability behavior. In order to understand whether the observed trend is a general phenomenon


8

Page 9 of 21 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


associated with surface hydrophilicity, we investigated a commercially available amphiphilic block copolymer, polystyrene-block-polyethylene glycol (PS-b-PEG). PS-b-PEG contains hydrophobic polystyrene (98 wt%) and hydrophilic polyethylene glycol (2 wt%) moieties. Although this type of polymers is not comparable to our NDI-T2 copolymers due to large hydrophilic/hydrophobic oligo-segments that they contain, we chose PS-b-PEG as an extreme case for an amphiphilic polymer film known to show phase separation at the liquid interface due to the incompatibility of its segments (controlled by the solvent choice and film preparation conditions).37-39 When the film was imaged with BODIPY-labeled lipids in the same manner, we observe bright islands of vesicles (Figure 2c). Moreover, AFM images show distinct features when the film is fully hydrated in PBS (Figure S3, A- D).


9


Page 10 of 21

Figure 2. (a) Chemical structures of the lipid vesicles DPhPC and DPhPE and BODIPY-labeled DHPE (b) Fluorescence images of polymers after incubation with 0.5 mg/mL of the vesicle solution in PBS. (c) Fluorescence image of the polystyrene-block-polyethylene glycol (PS-bPEG) film upon incubation with the same vesicles and its chemical structure. Scale bar in all images is 50 µm. The adsorption behavior of zwitterionic vesicles on the films can be explained by the wettability of the surfaces. As shown in Figure 3a, the surface free energy increases with the EG content indicating the film surfaces becomes gradually more hydrophilic. This is verified by an increase in the polar component of surface energy with the EG content (Figure 3b). To evaluate how the vesicle solution spreads on the film surface, we measured the contact angle formed by a drop of the vesicle solution on the surface of each polymer film. The contact angle decreases with an increase in the EG content (Figure 3c, see Figure S4 for digital images). Consequently, the work of adhesion of the liquid-solid phase boundary, i.e. the measure of the strength of the contact between two phases,40 increases with the EG content, calculated based on the Young and Dupree equations.41 Taken into account that the only polar component in the copolymer structure is the EG moiety, the hydrophilicity of the surface evidences the presence of EG therein. The increase in surface hydrophilicity is in favor of interactions with the vesicles.


10

Page 11 of 21 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. (a) Surface free energy (SFE) and (b) SFE components decomposed into dispersive (left axis) and polar parts (right axis) as a function of the EG content in the polymer composition. (c) Contact angle (left axis) and the work of adhesion (right axis) of the lipid vesicle solution on top of each film.

X-ray photoelectron spectroscopy (XPS) analyzes the elemental composition of surfaces (extending maximum 10 nm from the top),42 and can thus verify whether the EG chains are located at the uppermost surface. From the high-resolution C 1s (Figure 4a) and O 1s spectra (Figure S5a), we clearly observe the variation of relative intensities and peak shapes with EG content in the polymer composition. With more EG in the chemical structure, we detect an increase in the concentration of C-O-C bonds accompanied with a decrease in C-C bonds, which are attributed to the EG and alkyl side chains, respectively (Figure 4b). The elemental surface composition, which was obtained from survey spectra, revealed an increase in oxygen atoms and decrease in carbon atoms with increasing content of EG in the polymer (Figure S5b). The data are in agreement with the FRAP and surface energy observations, that more EG chains are located at the surface of the films when going from P-10 to P-90.


11


Page 12 of 21

Figure 4. (a) High resolution XPS spectra of polymer films in C 1s region. Binding energy positions for C-C and C-O-C bonds are labeled with the vertical dashed lines. The deconvolution of each spectra is shown in Figure S6. (b) The percentage of deconvoluted peaks of C-C (black dots) and C-O-C (blue dots) bonds relative to the total composition of the XPS C 1s spectra. These two bonds are typical for the hydrophobic alkyl and the hydrophilic EG side chains, respectively.

It is intriguing that although P-100 provides the most hydrophilic surface with the highest work of adhesion (Figure 3) a result of its EG-rich surface (Figure 4b), vesicles do not seem to adsorb onto this film (Figure 2b). In fact, when using quartz crystal microbalance with dissipation monitoring (QCM-D) to monitor the mass changes on the film upon interactions with vesicles, we see that P-100 shows the smallest change in oscillation frequency, in other words, it is the most unfavorable polymer for vesicles (Figure S7). Upon introducing the vesicle solution, the mass accumulated on P-10 is the lowest while it is the highest for P-90 (Figure S7). These results suggest that the surface hydrophilicity alone may not be sufficient to ensure optimal vesiclesurface interactions.23 As none of these surfaces carries an ionic charge, we look into their


12

Page 13 of 21 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


roughness. Compared with other films, the topography of P-100 film differs dramatically in aqueous media (Figure 1b). The roughness of the films was evaluated based on the AFM images obtained in PBS and varies between 1.9 nm and 2.1 nm for P-0, P-10, P-50 and P-90 (Figure S8). For P-100 film, on the other hand, the RMS roughness is ca. 2.9 nm. The rougher surface of this film when hydrated is attributed to the higher amount of EG chains on its surface that interact and swell with ions/water molecules. Although the difference is not large, this value is on the order of the size of a single lipid molecule and has been considered as a factor hindering the formation of lipid bilayers.19, 43

Our results show that EG chains are exposed on the surface and contribute to the wetting behavior, and the increase in their concentration in the composition improves vesicle adsorption. It is, however, worthy to note that when the films are drop cast from the same solutions, rather than spin cast, we do not observe any of these fluorescent patterns. For instance, the FRAP image of a drop-cast P-50 film is featureless (Figure S9). XPS data show a significant drop in the concentration of C-O-C bonds at the surface of this film compared to the spin-cast one, suggesting less EG moieties exposed at the surface (Table S2). Additionally, the drop-cast films (from all the copolymer solutions under investigation) exhibit lower surface free energy and work of adhesion than the spin-cast films (Figure S10). Both XPS and surface free energy data indicate the drop-cast films present unfavorable surfaces for vesicle fusion/interactions. In dropcast film formation, the polymer has more time to rearrange into its thermodynamic minimum before the solvent evaporates and the solid phase is set. Thus, the alkyl chains can migrate to the film surface, lowering the surface energy. In spin-coated film formation, there is less time during drying, and the structure is more kinetically driven on solvent evaporation, and therefore


13


Page 14 of 21

is more likely to have some glycol groups at the surface. This shows that the importance of film preparation conditions on the final surface properties of the film.

Figure 5. Recovery of fluorescence observed for the lipid membranes assembled on the NDI-T2 copolymer films, P-10, P-50 and P-90. The dark spot indicates the laser irradiation which bleaches the fluorescence of the particular spot that it hits. Scale bar = 50 µm. The size of the laser spot was adjusted to show the recovery behavior of only bright, dark or mixed regions in the films.

Finally, even when vesicles adsorb on the surface, the bilayer formation is inefficient determined by bilayer diffusion coefficient and mobile fraction. Diffusion coefficient provides information about the lateral mobility of the lipid membrane. For the areas where we could see recovery, the FRAP analysis shows that the average diffusion coefficient of the bilayer formed on P-90 is 1.2


14

Page 15 of 21 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


± 0.2 µm2/s and decreases to 0.6 ± 0.1 µm2/s for P-50 and to 0.2 ± 0.1 µm2/s for P-10 film. The mobile fraction, an indication of the portion of the lipid molecules moving within the bilayer, varies between 50% and 60%. For a comparison, the same liposomes form a bilayer on the standard polymer of organic bioelectronics, poly(3,4-ethylenedioxythiophene) doped with poly(styrene sulfonate) (PEDOT:PSS), with a diffusion coefficient of ca.1.67 µm2/s and a mobile fraction of 95%.15,17 The only trend that could be extracted is that the majority of recovery is observed along the bright areas (notice that the black areas exposed to the laser irradiation are steady over time and that it is only the bright areas that show fluorescence recovery), confirming that these are the regions that are populated by vesicles (Figure 5). When the vesicles are, however, not inter-connected, trapped inside islands defined by EG-rich areas, a continuous bilayer cannot be formed. The fluorescence signal arises from a combined effect of intact vesicles and patches of lipid bilayers. Therefore, either due to insufficient wettability or due to roughness, these surfaces are not ideal for lipid bilayer formation from zwitterionic liposomes. An ideal surface should be hydrophilic (contact angle close to 0) and contain charges, such as the one of PEDOT:PSS, while having minimized roughness. For the NDI-T2 polymers, even the polymer that has the highest concentration of EGs might be still relatively hydrophobic to form a mobile bilayer. Future synthetic efforts aim at increasing the surface energy via decorating EG based side chains with charged units as well as increasing their length.

Conclusions In this work, we investigated the impact of the side chains on surface properties of a recently developed, high performance electron transporting materials, that is NDI-T2 copolymers. We characterized the solid/liquid interface of these materials with the aim to form lipid bilayers. We


15


Page 16 of 21

observed an intriguing phenomenon where the interaction of zwitterionic vesicles with amphiphilic polymer surfaces leads to distinct and reproducible patterns, informing about the wettability of the film. By imaging the interactions of fluorophore-labeled liposomes with these surfaces, we not only show that the films can be cast such that ethylene glycol-rich regions are accumulated on the outmost surface but also evaluate what type of surfaces are required for the formation of lipid bilayers. The wettability information that gained using the amphiphilic vesicles can be used in the future to understand how biological systems interact with synthetic polymers. The ethylene glycol chains are promising to selectively interact with biological systems; controlling their percentage in the polymer composition as well as their surface exposure can enable control over protein adsorption, cell adhesion and spreading. For the formation of continuous lipid bilayers based on these vesicles, future work will focus on designing electron transporting conjugated polymers decorated with ionic charges which exhibit smooth surfaces when hydrated.

Supporting Information. The Supporting Information is available free of charge on the ACS Publications website at DOI: Experimental methods, AFM phase and TEM images of NDI-T2 copolymer films, surface coverage of bright and dark regions observed in the fluorescence images, AFM topography and phase images of PS-PEG film, pictures of a vesicle drop on the copolymer surfaces, XPS O1s and complete C1s spectra of the copolymer films, QCM-D data evidencing vesicle-copolymer interactions, roughness of the copolymer films in PBS, a fluorescence image of drop-cast P-50 film, comparison of surface free energy as well XPS analysis of drop cast versus spin cast films.


16

Page 17 of 21 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


Corresponding Author *[email protected] Notes The authors declare no competing financial interest. ACKNOWLEDGMENT We thank Mr. Sankara Arunachalam for assistance in acquiring surface free energy data and Dr. Nimer Wehbe for performing XPS measurements. We acknowledge Jokubas Surgailis for the analysis of fluorescence images.

REFERENCES (1) Liao, C.; Zhang, M.; Yao, Y. M..; Hua, T.; Li, L.; Yan, F. Flexible Organic Electronics in Biology: Materials and Devices. Adv. Mater. 2014, 27 (46), 7493 -7527. (2) Berggren, M..; Dahlfors, R. A. Organic Bioelectronics. Adv. Mater. 2007, 19 (20), 32013213. (3) Vaquero, S.; Bossio, C.; Bellani, S.; Martino, M.; Zucchetti, E.; Lanzani, G.; Antognazza, M.R. Conjugated Polymers for the Optical Control of the Electrical Activity of Living Cells. J. Mater. Chem. B 2016, 4, 5272-5283. (4) Inal, S.; Rivnay, J.; Suiu A.O.; Malliaras, G. G.; McCulloch, I. Conjugated Polymers in Bioelectronics. Acc. Chem. Res. 2018, 51 (6), 1368-1376. (5) Simon, D. T.; Gabrielsson E. O.: Tybrandt, K.; Berggren, M. Organic Bioelectronics: Bridging the Signaling Gap between Biology and Technology. Chem. Rev. 2016, 116 (21), 13009-13041. (6) Higgins, M. J.; Molino, P. J.; Yue, Z.; Wallace, G. G. Organic Conducting Polymer–Protein Interactions. Chem. Mater. 2012, 24 (5), 828-839. (7) Mawad, D.; Stewart, E.; Officer, D. L.; Romeo, T.; Wagner, P.; Wagner, K.; Wallace, G. G. A Single Component Conducting Polymer Hydrogel as a Scaffold for Tissue Engineering. Adv. Funct. Mater. 2012, 22 (13), 2692-2699. (8) Inal, S.; Hama, A.; Ferro M.; Pitsalidis, C.; Oziat, J.; Iandolo D.; Pappa, A.‐M.; Hadida, M.; Huerta, M.; Marchat, D.; Mailley, P.; Owens, R. M. Conducting Polymer Scaffolds for Hosting and Monitoring 3D Cell Culture. Adv. Biosyst. 2017, 1 (6), 1700052,. (9) Wan, A. M.-D.; Inal, S.; Williams, T.; Wang, K.; Leleux, P.; Estevez, L.; Giannelis, E. P.; Fischbach, C.; Malliaras, G. G.; Gourdon, D. 3D Conducting Polymer Platforms for Electrical Control of Protein Conformation and Cellular Functions. J. Mater. Chem. B 2015, 3 (25), 50405048.


17


Page 18 of 21

(10) Guex, A. G.; Puetzer, J. L.; Armgarth, A.; Littmann, E.; Stavrinidou, E.; Giannelis, E. P.; Malliaras, G. G.; Stevens, M. M. Highly Porous Scaffolds of PEDOT:PSS for Bone Tissue Engineering. Acta Biomater. 2017, 62, 91-101. (11) Bendrea, A.-D.; Cianga, L.; Cianga, I. Review Paper: Progress in the Field of Conducting Polymers for Tissue Engineering Applications. J. Biomater. Appl. 2011, 26 (1), 3-84. (12) Golabi, M.; Turner, A. P. F.; Jager, E. W. H. Tunable Conjugated Polymers for Bacterial Differentiation. Sens. Actuators, B 2016, 222, 839-848. (13) Maione, S.; Fabregat, G.; Valle, J. d. V.; Bendrea, A.‐D.; Cianga, L.; Cianga, I.; Estrany, F.; Aleman, C. Effect of the Graft Ratio on the Properties of Polythiophene‐g‐poly(ethylene glycol). J. Polym. Sci., Part B: Polym. Phys. 2015, 53 (4), 239-252. (14) Balint, R.; Cassidy, N. J.; Cartmell, S. H. Conductive Polymers: Towards a Smart Biomaterial for Tissue Engineering. Acta Biomater. 2014, 10 (6), 2341-2353 (15) Zhang, Y.; Wustoni, S.; Savva, A.; Giovannitti, A.; McCulloch, I.; Inal, S. Lipid Bilayer Formation on Organic Electronic Materials. J. Mater. Chem. C 2018, 6 (19), 5218-5227. (16) Chan, Y-H. M.; Boxer, S. G. Model Membrane Systems and Their Applications. Curr. Opin. Chem. Biol. 2007, 11 (6), 581-587. (17) Zhang, Y.; Inal, S.; Hsia, C.-Y.; Ferro, M.; Ferro, M.; Daniel, S.; Owens, R. M. Supported Lipid Bilayer Assembly on PEDOT:PSS Films and Transistors. Adv. Funct. Mater. 2016, 26 (40), 7304-7313. (18) Pitsalidis, C.; Pappa A-M,: Porel M.; Artim C. M.; Faria G. C.; Duong D. D.; Alabi C. A.; Daniel S.; Salleo A.; Owens R. M. Biomimetic Electronic Devices for Measuring Bacterial Membrane Disruption. Adv. Mater. 2018, 30, 1803130. (19) Cremer, P. S.; Boxer, S. G. Formation and Spreading of Lipid Bilayers on Planar Glass Supports. J. Phys. Chem. B 1999, 103 (13), 2554-2559. (20) Schönherr, H.; Johnson, J. M.; Lenz, P.; Frank, C. W.; Boxer, S. G. Vesicle Adsorption and Lipid Bilayer Formation on Glass Studied by Atomic Force Microscopy. Langmuir 2004, 20 (26), 11600-11606. (21) Rapuano, R.; Carmona-Ribeiro, A. M. Physical Adsorption of Bilayer Membranes on Silica. J. Colloid Interface Sci. 1997, 193 (1), 104-111. (22) Richter, R. P.; Brisson, A. R. Following the Formation of Supported Lipid Bilayers on Mica: A Study Combining AFM, QCM-D, and Ellipsometry. Biophys. J. 2005, 88 (5), 34223433. (23) Tero, R.; Watanabe, H.; Urisu, T. Supported Phospholipid Bilayer Formation on Hydrophilicity-controlled Silicon Dioxide Surfaces. Phys. Chem. Chem. Phys. 2006, 8 (33), 3885-3894. (24) Webb, K.; Hlady, V.; Tresco, P. A. Relative Importance of Surface Wettability and Charged Functional Groups on NIH 3T3 Fibroblast Attachment, Spreading, and Cytoskeletal Organization. J. Biomed. Mater. Res. 1998, 41 (3), 422-430 (25) Hackett, A. J.; Malmström, J.; Travas-Sejdic, J. Functionalization of Conducting Polymers for Biointerface Applications. Prog. Polym. Sci. 2017, 70, 18-33 (26) Lee, J. Y.; Schmidt, C. E. Amine‐Functionalized Polypyrrole: Inherently Cell Adhesive Conducting Polymer. J. Biomed. Mater. Res. Part A 2015, 103 (6), 2126-2132. (27) Lee, J.-W.; Serna, F.; Nickels, J.; Schmidt, C. E. Carboxylic Acid-Functionalized Conductive Polypyrrole as a Bioactive Platform for Cell Adhesion. Biomacromolecules 2006, 7 (6), 1692-1695.


18

Page 19 of 21 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


(28) Giovannitti, A.; Sbircea, D.-T.; Inal, S.; Nielsen, C. B.; Bandiello, E.; Hanifi, D. A.; Sessolo, M.; Malliaras, G. G.; McCulloch, I.; Rivnay, J. Controlling the Mode of Operation of Organic Transistors through Side-Chain Engineering. Proc. Natl. Acad. Sci. 2016, 113 (43), 12017-12022. (29) Nielsen, C. B.; Giovannitti, A.; Sbircea, D.-T.; Bandiello, E.; Niazi, M. R.; Hanifi, D. A.; Sessolo, M.; Amassian, A.; Malliaras, G. G.; Rivnay, J.; McCulloch, I. Molecular Design of Semiconducting Polymers for High-Performance Organic Electrochemical Transistors. J. Am. Chem. Soc. 2016, 138 (32), 10252-10259. (30) Giovannitti, A.; Nielsen, C. B.; Sbircea, D.-T.; Inal, S.; Donahue, M.; Niazi, M. R.; Hanifi, D. A.; Amassian, A.; Malliaras, G. G.; Rivnay, J.; McCulloch, I. N-type Organic Electrochemical Transistors with Stability in Water. Nat. Commun. 2016, 7, 13066. (31) Giovannitti, A.; Maria, I. P.; Hanifi, D.; Donahue, M. J.; Bryant, D.; Barth, K. J.; Makdah, B. E.; Savva, A.; Moia, D.; Zetek, M.; Barnes, P. R. F.; Reid, O. G.; Inal, S.; Rumbles, G.; Malliaras, G. G.; Nelson, J.; Rivnay, J.; McCulloch, I. The Role of the Side Chain on the Performance of N-type Conjugated Polymers in Aqueous Electrolytes. Chem. Mater. 2018, 30 (9), 2945-2953. (32) Rivnay, J.; Inal, S.; Salleo, A.; Owens, R. M.; Berggren, M.; Malliaras, G. G. Organic Electrochemical Transistors. Nat. Rev. Mater. 2018, 3, 17086. (33) Pappa, A.M.; Ohayon, D.; Giovannitti, A.; Maria, I. P.; Savva, A.; Uguz, I.; Rivnay, J.; McCulloch, I.; Owens, M.R.; Inal, S. Direct Metabolite Detection with an n-type Accumulation Mode Organic Electrochemical Transistor. Sci. Adv. 2018, 4 (6), eaat0911. (34) Takacs, C. J.; Treat, N. D.; Krämer, S.; Chen, Z.; Facchetti, A.; Chabinyc, M. L.; Heeger, A. J. Remarkable Order of a High-Performance Polymer. Nano Lett. 2013, 13 (6), 2522-2527. (35) Deshmukh, K. D.; Qin, T.; Gallaher, J. K.; Liu, A. C. Y.; Gann, E.; O'Donnell, K.; Thomsen, L.; Hodgkiss, J. M.; Watkins, S. E.; McNeill, C. R. Performance, Morphology and Photophysics of High Open-Circuit Voltage, Low Band Gap All-Polymer Solar Cells. Energy & Environ. Sci. 2015, 8 (1), 332-342. (36) Baba, T.; Toshima, Y.; Minamikawa, H.; Hato, M.; Suzuki, K.; Kamo, N. Formation and Characterization of Planar Lipid Bilayer Membranes from Synthetic Phytanyl-Chained Glycolipids. Biochim. Biophys. Acta, Biomembr. 1999, 1421 (1), 91-102. (37) Feng, L. B.; Zhou, S. X.; You, B.; Wu, L. M. Synthesis and Surface Properties of Polystyrene‐graft‐poly(ethylene glycol) Copolymers. J. Appl. Poly. Sci. 2007, 103 (3), 14581465. (38) Richards, R. W.; Rochford, B. R.; Webster, J. R. P. Surface Phase Separation in an Amphiphilic Block Copolymer Monolayer at the Air-Water Interface. Polymer 1997, 38 (5), 1169-1177. (39) Nitta, K.; Kimoto, A.; Watanabe, J. Surface Properties of Amphiphilic Graft Copolymer Containing Different Oligo Segments. Polymer 2016, 96, 45-53. (40) Schrader, M. E. Young-Dupre Revisited. Langmuir 1995, 11 (9), 3585-3589. (41) Young, T. III. An Essay on the Cohesion of Fluids. Philos. Trans. R. Soc. London 1805, 95, 65-87. (42) Gilbert, J. B.; Rubner, M. F.; Cohen, R. E. Depth-Profiling X-ray Photoelectron Spectroscopy (XPS) Analysis of Interlayer Diffusion in Polyelectrolyte Multilayers. Proc. Natl. Acad. Sci. 2013, 110 (17), 6651-6656. (43) Raedler, J.; Strey, H.; Sackmann, E. Phenomenology and Kinetics of Lipid Bilayer Spreading on Hydrophilic Surfaces. Langmuir 1995, 11 (11), 4539-4548.


19


Page 20 of 21


20

Page 21 of 21 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


Table of Contents


21

Visualizing the Solid–Liquid Interface of Conjugated Copolymer Films

Recommend Documents