Three-Dimensional, Bifunctional Microstructured Polymer Hydrogels

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Interfaces: Adsorption, Reactions, Films, Forces, Measurement Techniques, Charge Transfer, Electrochemistry, Electrocatalysis, Energy Production and Storage

Three-Dimensional, Bifunctional Microstructured Polymer Hydrogels Made From Polyzwitterions and Antimicrobial Polymers Vania T. Widyaya, Claas Müller, Ali Al-Ahmad, and Karen Lienkamp Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.8b03410 • Publication Date (Web): 18 Dec 2018 Downloaded from http://pubs.acs.org on December 24, 2018

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Three-Dimensional, Bifunctional Microstructured Polymer Hydrogels Made From Polyzwitterions and Antimicrobial Polymers Vania Tanda Widyaya,† Claas Müller, ‡ Ali Al-Ahmad,§ and Karen Lienkamp†,* †

Bioactive Polymer Synthesis and Surface Engineering Group, Department of Microsystems Engineering (IMTEK) and Freiburg Center for Interactive Materials and Bioinspired Technologies (FIT), Albert-Ludwigs-Universität Freiburg, Georges-Köhler-Allee 105, 79110 Freiburg, Germany

‡

Laboratory for Process Technology, Department of Microsystem Engineering (IMTEK), Albert-Ludwigs-Universität Freiburg, Georges-Köhler-Allee 103, 79110 Freiburg, Germany

§

Department of Operative Dentistry and Periodontology, Center for Dental Medicine of the Albert-Ludwigs-Universität, Freiburg, Hugstetter Str. 55, 79106 Germany.

*E-mail: [email protected]

KEYWORDS: antimicrobial polymers, biofilms, microcontact printing, polyzwitterions, protein-repellent polymers. ABSTRACT: Biofilm-associated infections of medical devices are a global problem. To prevent such infections, biomaterial surfaces are chemically or topographically modified to slow down the initial stages of biofilm formation. In the bifunctional material here presented, chemical and

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topographical cues are combined, so that protein and bacterial adhesion as well as bacterial proliferation are effectively inhibited. By changing the surface topography parameters and investigating the effect of these changes on bioactivity, structure-property-relationships are obtained. The target material is obtained by microcontact printing (µCP), a soft lithography method. The antimicrobial component, a poly(oxanorbornene) based Synthetic Mimics of an Antimicrobial Peptide (SMAMP), was printed onto a protein-repellent polysulfobetaine hydrogel, so that bifunctional 3D structured polymer surfaces with 1 µm, 2 µm, and 8.5 µm spacing are obtained. These surfaces are characterized with fluorescence microscopy, surface plasmon resonance spectroscopy, atomic force microscopy, and contact angle measurements. Biological studies show that the bifunctional surfaces with 1 µm and 2 µm spacing are 100% antimicrobially active against Escherichia coli and Staphylococcus aureus, 100% fibrinogenrepellent, and non-toxic to human gingival mucosal keratinocytes. At 8.5 µm spacing, the broad-band antimicrobial activity and the protein-repellency are compromised, which indicates that this spacing is above the upper limit for effective simultaneous antimicrobial activity and protein-repellency of polyzwitterionic-polycationic materials.

TOC image Ph

Ph

O

O n

n O

O

O

NH3

O

N

O O

N+

Antimicrobial SMAMP O

S

O O-

Protein-repellent Polysulfobetaine (PSB)


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MANUSCRIPT:

INTRODUCTION Background. Biofilm formation on medical devices such as catheters or implants is a serious problem in modern healthcare, leading to severe infections and the death of hundred thousands of patients worldwide every year.1-2 Biofilms are so dangerous because the bacteria inside these films are protected by an extracellular matrix; they are therefore much more difficult to eradicate than planktonic bacteria and require 100 to 1000 times higher doses of antibiotics.3 The problem is even worse when antibiotic-resistant bacterial strains are involved, which are unfortunately also on the rise.4-5 Consequently, it is important to inhibit protein and bacterial adhesion as well as bacterial proliferation on surfaces, which are the initial steps of biofilm formation on biomaterials.6 The ability of bacteria to attach to a surface depends on the surface chemistry and topography.7-8 Strategies to fight biofilm formation by modifying the surface chemistry include, for example, surface functionalization with contact-active antimicrobial or protein-repellent polymers, notably polyzwitterions.9-11 Contact-active antimicrobial polymer coatings are polycationic. They kill bacteria by interaction with their negatively charged bacterial cell envelope, although the precise mechanism of that interaction is still under debate.910, 12-16

Polyzwitterions, on the other hand, have an equal number of positive and negative

charges. They are strongly hydrophilic and bind significant amounts of water without disturbing its hydrogen bond network; therefore, proteins and bacteria do not gain sufficient free energy when interacting with the polyzwitterionic surfaces, so that these materials are overall proteinrepellent.17-21 While these chemical surface modification strategies are very useful for short time exposure to bacteria-containing fluids, they are insufficient for long term applications. Cationic antimicrobial polymer surfaces 'capture' negatively charged dead bacteria and other biomolecules by electrostatic interactions. This causes contamination and deactivation of the surface, and can initiate biofilm formation.9 Protein-repellent polymers become vulnerable once 3 Environment ACS Paragon Plus

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a single bacterium manages to settle and proliferate, as this one pathogen can form a biofilm in less than 24 hours.22 The need for long-term prevention of biofilm formation has encouraged researchers to develop bifunctional materials with combined antimicrobial activity and protein-repellency.9, 23-26

The idea of this concept is that “It takes walls and knights to defend a castle”,27 i.e. that

protein and bacteria-repellent 'walls' and lethal 'knights' incorporated into the same material would be able to slow down biofilm formation for a longer time than monofunctional materials. Bifunctional materials can be obtained, for example, by embedding leachable antimicrobial agents inside protein-repellent polymer matrices,28-29 or by releasing covalently bound antimicrobial agents hydrolytically from a protein-repellent material.30-33 Another design involves switching the materials' surface charge from cationic/contact-killing to zwitterionic/protein-repellent.33-34 Surfaces with switchable bioactivities triggered by temperature changes have also been reported.35-36 These concepts are highly interesting from an academic perspective, yet difficult to implement in real-life applications or in clinical settings. Materials that contain covalently bound antimicrobial and protein-repellent moieties, and simultaneously present these two functions to their environment have also been reported, e.g. by biocide tethering to non-fouling hydrophilic polymers,37-39 'grafting-onto' approaches of one component onto the other,27 bifunctional block and graft copolymers,26, 40 or layer-by-layer approaches.41-43 In materials built via these concepts, the physicochemical characteristics of the two functional moieties and the surface architecture dictate the self-organization of the polymer-liquid interface. Even when the bulk composition of the materials can be controlled, the ratio of functional moieties at the interface is difficult to predict and to precisely balance, so that complete simultaneous antimicrobial activity and protein repellency could, to our knowledge, not be obtained by any of these materials. For example, block copolymers from protein-repellent poly(ethylene glycol) (PEG) and cationic antimicrobial polymers were only


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fully protein-repellent at high PEG content, yet at that composition they were only active against Gram-negative Escherichia coli, while their activity against Gram-positive Staphylococcus aureus was compromised.40 In that light, the concept to obtain simultaneously protein-repellent and antimicrobial bifunctional polymer surfaces by deposition of the two functions side-by-side44-45 has gained increasing interest. Pre-structuring of a surface dictates the dimensions and distribution of each surface feature, and thus allows more precise control over the polymer-water interface. Additionally, such approaches allow simultaneous modification of the surface chemistry and topography. Surface topography has been long recognized as an important factor to control adhesion of bacteria and other organisms.46-49 Surface topography influences bacterial adhesion through the size, shape, and orientation of the surface features.7, 50 For instance, features larger than bacterial cells deter proliferation;49-50 equally sized or smaller features often affect bacterial orientation on the surface and reduce bacterial adhesion.51-52 Nano-sized surface features with high aspect ratios were found to be bactericidal since they were able to rupture bacterial membranes.53-56 A recent example of a microstructured side-by-side bifunctional surface made form contact-killing polycations and protein-repellent polyzwitterions is a checkered material consisting of 2 x 2 µm squares made from poly(quaternary ammonium salts) and polysulfobetaines that were grafted onto a layer-by-layer assembly made from poly(allylamine hydrochloride)/poly(styrene sulfonate).44 On this material, adhesion of E. coli was reduced by 70-93% compared to untreated polyamide.44 We recently reported bifunctional micro- and nanostructured polymer surfaces from polycationic Synthetic Mimics of Antimicrobial Peptides (SMAMPs) and zwitterionic poly(sulfobetaines) (PSB).45 The submicrometer-sized surface features of these materials were obtained by colloidal lithography. To obtain structure-propertyrelationships, both the structure spacings (200 nm to 1 µm) and the polymer position were varied.45 The protein repellency of the materials was near-quantitative, particularly for the


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materials with the smaller spacings.45 This is in line with the idea that the adhesive SMAMP patches of these materials would be small and well shielded by the PSB patches. The material with the 1 µm spacing was fully antimicrobially active against E. coli, while the activity of the materials with smaller spacing was reduced.45 Apparently, the small SMAMP patches of these materials were not able to sufficiently damage the bacterial membranes and could therefore not effectively maintain the antimicrobial activity. Thus, these materials demonstrated that the lower limit for sufficient antimicrobial activity was below 1 µm. The weakness of this study was that the fabrication process was rather complicated and not practical on a large scale. Also, since the materials were made from surface-attached polymer monolayers, they were quite susceptible to defects and damage. Thus, a straight-forward process for the fabrication of microstructured bifunctional surfaces, leading to more robust materials, would be desirable. Such a study should also attempt to find the upper limit for dual antimicrobial activity and protein-repellency of bifunctional materials made from cationic antimicrobial polymers and protein-repellent polyzwitterions. Aim and Design of the Study. The aim of this work was to develop a process to obtain robust microstructured bifunctional surfaces made from protein-repellent polyzwitterions and cationic antimicrobial polymers; to explore the structure-property relationships of these surfaces; and to develop a bifunctional material that is fully antimicrobially active, protein-repellent and cell compatible. We here report the fabrication of such bifunctional polymer surfaces using contactkilling cationic antimicrobial poly(oxanorbornene) SMAMPs,57-59 and protein-repellent poly(oxanorbornene)-based poly(sulfobetaines)60-61 by microcontact printing (µCP), a soft lithography technique (Figure 1a and b).62-64 In this process, the antimicrobial SMAMP polymer (NBD-SMAMP or BP-NBD-SMAMP, respectively, Figure 1b and d, green; NBD = green nitrobenzoxadiazole, BP = benzophenone) is printed on top of a protein-repellent polymer base layer (BP-COU-PSB, Figure 1b and c, blue; COU = coumarin). The key idea of this fabrication


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process is not so much the microstructuring process, but the design of the polymers used as base layer and printing ink. Each polymer contains a small fraction of repeat units that carry built-in UV cross-linker groups (e.g. benzophenone, BP; black part of the polymer structure in Figure 1c and d). When UV irradiated, these polymers can cross-link and form networks with covalent bonds to the polymer-substrate interface (grey dots in Figure 1a and b) and throughout the polymer volume (blue and green dots in Figure 1a and b) through C,H-insertion reactions, so that 3D polymer hydrogels65-66 and not just monolayers are formed. For this study, bifunctional polymer hydrogels with varying height and spacing were synthesized. Their physical and biological properties were studied in detail to obtain structureproperty-relationships for microstructured materials made from polyzwitterions and cationic antimicrobial polymers. To isolate the effects of the polyzwitterion base layer, monofunctional materials were also prepared by stamping the SMAMP polymer inks directly onto the desired substrate. The properties of these mono- and bifunctional polymer surfaces were then compared. Notably, some of the bifunctional materials thus obtained were fully active against Gram-negative E. coli and Gram-positive S. aureus, repelled 100% of the protein fibrinogen, and were highly compatible with human gingival mucosal keratinocytes.


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Figure 1. Cartoon illustration of the chemical composition and build-up of the targeted bifunctional surface: a) The protein-repellent polyzwitterion BP-COU-PSB containing UV cross-linkable benzophenone groups (BP, black part of the polymer structure in c) was applied to a BP-functionalized substrate. Upon UV irradiation, covalent bonds to the surface BP groups (grey dots) formed, and the BP groups within the polymer chains formed polymer-polymer cross-links (blue dots). b) An antimicrobial polymer (NBD-SMAMP or BP-NBD-SMAMP) was printed onto the BP-COU-PSB base layer by microcontact printing (µCP). Upon UV irradiation, covalent bonds to the base hydrogel and within the (BP-)NBD-SMAMP patches formed through the BP or NBD groups. All polymers were synthesized via ring-opening metathesis polymerization: c) BP-COU-PSB from PSB, COU and BP monomers and d) BPNBD-SMAMP from SMAMP, NBD and BP monomers, as well as NBD-SMAMP from SMAMP and NBD monomers; e) the SMAMP patches were activated on the surface with HCl, which removed the Boc protective groups and yielded antimicrobially active NH3+ groups. 8 Environment ACS Paragon Plus

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EXPERIMENTAL METHODS General. All chemicals and solvents (reagent grade) were obtained from Sigma-Aldrich (Munich, Germany) or Carl Roth (Karlsruhe, Germany) and used as received unless otherwise indicated. NMR spectra were recorded on a Bruker 250 MHz spectrometer (Bruker, Madison, WI, USA). Gel permeation chromatography (GPC) was used to determine the polymer molecular weight of (BP-)NDB-SMAMP (chloroform, SDV columns (PSS, Mainz, Germany), PMMA standards) and BP-COU-PSB (trifluoroethanol (TFE), 0.1 M sodium trifluoroacetate, PFG columns (PSS, Mainz, Germany), PMMA standards). Single-side polished silicon wafers (orientation of [100] and thickness of 525 ± 25 µm) were obtained from Si-Mat (Kaufering, Germany). The spin-coater used was a SPIN150-NPP (SPS-Europe, Putten, Netherlands). The UV irradiation unit used was a BIO-LINK-box (Vilber Lourmat GmbH, Eberhardzell, Germany) with 254 nm light sources. Polydimethylsiloxane (PDMS) stamps were fabricated by molding the PDMS solution (Elastosil RT601, Wacker Chemie AG, Stuttgart, Germany) on a silicon master (15 × 15 cm2) which was patterned using 2 steps laser interference lithography. The obtained structured PDMS stamps had a parabolic pattern with 1 µm, 2 µm, and 8.5 µm spacing (peak-to-peak distance, Figure S1 in the Supporting Information). It was then cut into 2 × 1.5 cm2 pieces. The stamps were also characterized with contact angle measurements and the results are listed in Table S1. Polymers Synthesis. The polymers NBD-SMAMP and BP-NBD-SMAMP (Figure 1d, respectively) were synthesized as described elsewhere.66 The benzophenone-carrying monomer (BP)67 and the coumarin-carrying monomer (COU)68 (Figure 1c) were synthesized and characterized as previously published. The polysulfobetaine (PSB) monomer precursor exo-4(2-dimethylaminoethyl)-10-oxa-4-aza-tricyclo[5.2.1.02,6]dec-8-ene-3,5-dione (DMAETDD), was

synthesized

as

described

in

the

literature.69

Briefly,

exo-3,6-epoxy-1,2,3,6-

tetrahydrophthalic anhydride (10 g, 60 mmol) was dissolved in methanol (MeOH) and tetrahydrofuran (THF) (250 mL, 1:1) and heated to 65oC. Then N,N-dimethylethylenediamine 9 Environment ACS Paragon Plus

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(5.3 g, 60 mmol) was added slowly to the stirred anhydride solution. The reaction mixture was refluxed at 65°C for 24 h. The solvent was then evaporated under reduced pressure. The product was re-dissolved in a small amount of a dichloromethane (DCM) and MeOH mixture (5:1 v/v) and precipitated into ice-cooled n-hexane. The precipitated solid was separated from the nhexane, re-dissolved again in a small amount of DCM/MeOH mixture (5:1 v/v), and then precipitated into ice-cooled n-hexane. All n-hexane solutions were kept in ice bath for 3 days from which white crystals of DMAETDD were formed. The 1H-NMR data obtained matched those in the literature.69 PSB monomer (Figure1c) was synthesized under nitrogen using standard Schlenk techniques using a previously published procedure.70 In short, DMAETDD monomer (2.5 g, 11 mmol) was dissolved in anhydrous acetonitrile (15 mL). To the solution, 1,4-butane sultone (1.7 g, 12 mmol) and 1,3-dinitrobenzene (40 mg, 0.2 mmol) were added, and the mixture solution was stirred at 50°C for 15 h. The product was washed with ethyl acetate and filtered. It was then dried under high vacuum. The 1H-NMR signals of the monomer matched those in the literature.70 BP-COU-PSB terpolymer was synthesized by ring-opening metathesis polymerization (ROMP) at room temperature under nitrogen using standard Schlenk techniques. PSB monomer (500 mg, 1.3 mmol), BP monomer (30.8 mg, 0.08 mmol), and COU monomer (55.8 mg, 0.16 mmol) were dissolved in dry trifluoroethanol (TFE, 5 mL). The Grubbs’ third generation catalyst (synthesized as previously reported,71 8.5 mg in 2 mL of dry DCM, 12 mmol) was added in one shot into the stirred monomer mixture solution. After 1 h, excess ethyl vinyl ether (1 mL, 10 mmol) was added to terminate the ROMP reaction, and the mixture was stirred for another 1 h. TFE was then evaporated under reduced pressure. The product was re-dissolved in a small amount of TFE and precipitated into ice-cold diethyl ether. The precipitated polymer was filtered and dried under high vacuum. The peak assignment of the NMR signals of BP-COU-PSB can be found in Figure S2. Target molecular mass: 50 kg mol-1, GPC analysis: Mn = 72.5 kg mol-1, PDI = 1.33.


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Surface Functionalization. Selective Surface Functionalization with Benzophenone CrossLinkers. Silicon wafers and glass substrates were chemically modified with triethoxy benzophenone silane (3EBP-silane).72 The silane solution (20 mg mL-1 in toluene) was spin coated (3000 rpm, 1000 rpm s-1, 30 s) onto the samples' surface. The samples were then cured at 120°C for 20 min on a preheated hotplate and subsequently washed with toluene. The silicon wafers were then cut into 1.5 cm × 1.5 cm pieces. Gold-covered substrates for surface plasmon resonance spectroscopy (SPR) were functionalized with benzophenone attached to lipoic acid (LS-BP, 10 mg mL-1 in toluene)27 by immersing the substrates in a LS-BP solution for 12 h. The substrates were then rinsed with toluene and dried under nitrogen flow. Formation of BP-COU-PSB Base Layer. Bifunctional surfaces were prepared by first spin coating BP-COU-PSB solution (10 mg mL-1 in TFE; 3000 rpm, 1000 rpm s-1, 10 s) onto the benzophenone-functionalized substrates, followed by UV irradiation (λ = 254 nm, 3 J cm-2). Any loosely attached polymer chains were rinsed off by immersing the substrates in stirred TFE for 18 h. The samples were then dried under nitrogen flow. Microcontact Printing (µCP). All steps of µCP process were conducted under ambient conditions. The substrates for fabrication of monofunctional surface were functionalized with either 3EBP or LS-BP. Bifunctional surfaces were obtained with substrates carrying BP-COUPSB base layers. The SMAMP inks NBD-SMAMP or BP-NBD-SMAMP were used in their tertbutyloxycarbonyl (Boc)-protected form. µCP with PDMS stamps having 1 µm or 2 µm spacings was performed as previously reported.66 In short, the stamps were loaded via an ink pad and brought into conformal contact with the substrate (ink concentration: 5 mg mL-1 for 1 µm spacing stamp and 2 mg mL-1 for 2 µm spacing stamp, solvent was a mixture of 30% v/v DCM and 70% v/v toluene ; 30 N printing pressure, 5 s of printing time). For µCp with the PDMS stamp having 8.5 µm spacing, the stamp was loaded with one drop of the respective SMAMP ink (NBD-SMAMP or BP-NBD-SMAMP, respectively; 3 mg mL-1 in THF) using a 11 Environment ACS Paragon Plus

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pipet. The solution was allowed to evaporate and the ink-loaded stamp was brought onto conformal contact with a substrate (15 N printing pressure, 20 s printing time). The microstructured substrate was then UV-irradiated (λ = 254 nm, 3 J cm-2). Any loosely attached SMAMP chains were removed by immersing the material in stirred toluene for 18 h. The samples were then dried under nitrogen flow. Surface Activation. To remove the Boc protective groups on the SMAMP inks, the samples were immersed in HCl (4 M in dioxane) for 15 h to completely deprotect the SMAMP amine group. They were then washed three times with ethanol and dried under nitrogen flow. Formation of Unstructured SMAMP. SMAMP solution (NBD-SMAMP or BP-NBDSMAMP; 10 mg mL-1 in 30% v/v DCM - 70% v/v toluene) was spin coated onto benzophenone-modified substrates (3000 rpm, 1000 rpm s-1, 30 s). Afterwards, the substrate was UV-irradiated (λ = 254 nm, 3 J cm-2) and washed in stirred toluene for 18 h. The samples were then dried under nitrogen flow. NBD-SMAMP gave a monolayer (SMAMP ML), BPNBD-SMAMP yielded a polymer network (SMAMP NW). Physical Characterization. Fluorescence Microscopy. Fluorescence microscopy images of polymer surface (in the protected state) were taken using a Nikon Eclipse Ti-S inverted microscope (Nikon GmbH, Düsseldorf, Germany) with a green fluorescent protein (GFP, green) and a 4′,6-diamidin-2-phenylindole (DAPI, blue) filter at 40 fold magnification (for features with 8.5 µm spacing) and 60 fold magnification (for features with 2 µm spacing). The images were processed using ImageJ software. Brightness and image contrast were adjusted for better visualization. Confocal Laser Scanning Microscopy. Confocal microscopy images of protected surface features with 1 µm spacing were taken using a Zeiss LSM 880 with AiryScan inverted


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microscope (Carl Zeiss Microscopy, Jena, Germany) at 63 fold magnification. The images were processed with ImageJ software; brightness and contrast were adjusted for better visualization. Atomic Force Microscopy (AFM). A Dimension Icon AFM (Bruker, Karlsruhe, Germany) was used to record the surface topography images in ScanAsyst tapping mode in air. Commercial ScanAsyst Air cantilevers (length: 115 µm; width: 25 µm; spring constant: 0.4 N m-1; resonance frequency: 70 kHz) were used. The obtained images were analyzed and processed using the Nanoscope Analysis 1.5 software. The topography of PDMS stamps with 1 µm and 2 µm spacing were imaged using the same AFM, those with 8.5 µm spacing were measured in tapping mode in air using a Nanowizard 4 AFM (JPK Instruments, Berlin, Germany). Commercial Nanosensors cantilever with tilt compensated high aspect ratio was used (length: 125 µm; width: 30 µm; force constant: 10 – 130 N m-1; resonance frequency: 204 – 497 kHz; aspect ratio at 2 µm: 5:1, tilt compensation: 13°). The topography images of the PDMS stamp were analyzed and processed with Gwyddion 2.45 software. The baseline of all height profile images were vertically translated to zero position using OriginPro 2015G. Contact Angle (CA) Measurements. The CA of all samples were measured using a CA system OCA 20 (Dataphysics GmbH, Filderstadt, Germany). The average value of the CA (static, advancing, and receding) was obtained from 5 samples. The static CA was determined by depositing 5 µL of water droplet onto the test sample. The CA value was then calculated using the Laplace-Young equations. For measuring the advancing CA, the dropping needle was immersed into the center of the water droplet. Water was then continuously dropped with a dosing rate of 0.5 µL s-1 until the droplet expanded, ideally on both sides of the droplet. The advancing CA was then evaluated using the elliptical method. Receding CA was determined by pumping back the water droplet into the water reservoir until the droplet receeded, ideally on both sides of the droplet. The receding CA was then calculated using the tangent method.


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Surface plasmon resonance (SPR) spectroscopy. To study the build-up of the bifunctional materials, a full reflectivity curve was measured after each fabrication step using the SPR instrument RT2005 in Kretschmann configuration (Res-Tec, Framersheim, Germany) with a He-Ne laser (λ = 632.8 nm). SPR substrates were made by the Clean Room Service-Center (RSC) of the Department of Microsystem Engineering, University of Freiburg, using a CS 730 S device (Von Ardenne, Dresden, Germany). In short, LaSFN9 glass (Hellma GmbH, Müllheim, Germany) was coated with 1 nm of Cr and 50 nm of Au. The angular reflectivity curves were simulated based on the Fresnel equations using the Winspall software. Biological Characterization. General. All biological assays were performed after the activation of the SMAMP polymers. Protein Adhesion Test. Protein (fibrinogen) adhesion measurements were conducted using surface plasmon resonance (SPR) spectroscopy Protein adsorption was studied in situ in the kinetics mode, and by taking full angular reflectivity scans of the dry surfaces between before and after the kinetics experiment, as previously reported.73 Details are given in the supporting information. Antimicrobial Activity Assay. The antimicrobial activity of the unstructured/structured polymer films were performed using a modified version of the Japanese Industrial Standard JIS Z 2801:2000 ‘Antibacterial Products Test for Antibacterial Activity and Efficacy’ as previously reported.74 Details are also given in the supporting information. Cell Compatibility Assay (Alamar Blue Assay). The Alamar Blue assay was performed according to previously reported procedure.73 Details are also given in the supporting information. Live-Dead Staining of Keratinocytes Grown on Substrates. The live-dead stain was performed as reported previously.73 Details are also given in the supporting information. 14 Environment ACS Paragon Plus

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RESULTS AND DISCUSSION Material Design and Polymer Synthesis. The target material consisted of a protein-repellent polyzwitterionic base layer, onto which microstructured antimicrobial 3D polymer patches were applied by microcontact printing (µCP). The base layer of this bifunctional material was made from the polyzwitterionic terpolymer BP-COU-PSB (Figure1c). Besides the zwitterionic PSB repeat units (Figure 1c, light blue), this polymer contained repeat units carrying benzophenone (BP, Figure1c, black), and repeat units carrying the blue fluorescent coumarin dye (COU, Figure 1c, blue). The synthesis scheme of BP-COU-PSB is shown in Figure 1c. BP and COU monomers (Figure 1c) were obtained as reported previously.67-68 The PSB monomer was synthesized by a combination of literature procedures69,70 as described in the Experimental. To obtain the BP-COU-PSB terpolymer, these three monomers were dissolved in dry trifluoroethanol in the desired ratio. Grubbs’ third generation catalyst, synthesized as previously reported,71 was added in one shot to the stirred monomer mixture solution. The reaction was quenched with ethyl vinyl ether, and the product was recovered by precipitation. The target molecular weight was 50 kg mol-1; GPC results were in reasonable agreement with that value (Mn = 72.5 kg mol-1, PDI = 1.33; calibrated against PMMA standards). The antimicrobial polymers used to formulate the inks for microcontact printing were the poly(oxanorbornene)-based Synthetic Mimics of Antimicrobial Peptides (SMAMPs) NBDSMAMP and BP-NBD-SMAMP (Figure1d).57-59 Their synthesis has been reported previously.66 In short, the relative amounts of the BP, SMAMP and NBD monomers were dissolved in dichloromethane and polymerized via ring-opening metathesis polymerization, giving polymers with an average molecular mass Mn of 75-92 kg mol-1 and a polydispersity index (PDI) of 1.2-1.3 (determined by gel permeation chromatography in chloroform against PMMA standards).66 In their activated form, these SMAMP polymers carry one primary ammonium group and one hydrophobic group on each SMAMP repeat unit.57 For microcontact 15 Environment ACS Paragon Plus

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printing, the SMAMP polymers were used in their Boc protected form (Figure1d), by which better wetting of the PDMS stamp used for printing was obtained. The two variants of the SMAMP were used because they differed in their cross-linking efficiency, so that 3D polymer patches with different heights could be obtained.66 NBD has a low cross-linking efficiency when UV irradiated,66 thus NBD-SMAMP gives short 3D surface features. Using BP-NBDSMAMP, with the more efficient BP cross-linker, taller features can be obtained.66 For visualization purposes, both the NBD-SMAMP and the BP-NBD-SMAMP ink contained a fluorophore, the green fluorescent nitrobenzoxadiazole (NBD). The first ink polymer, NBDSMAMP (Figure1d, respectively) contained 90 mol% SMAMP moieties and 10 mol% NBD; the second polymer, BP-NBD-SMAMP, had 80 mol% SMAMP, 10 mol% NBD, and 10 mol% BP (Figure 1d, respectively). Microcontact Printing and Fabrication of Reference Materials. The µCP process is shown schematically in Figure 2. Both SMAMP-containing inks were used in their tertbutyloxycarbonyl (Boc)-protected form. First, a PDMS stamp with periodic conical surface features of different spacings (1 µm, 2 µm and 8.5 µm) was fabricated as reported previously.66 Atomic force microscopy images of the stamps thus obtained are shown in Figure S1 of the supporting information. To load the stamp, two different inking methods were applied: contactinking for the stamps with 1 µm and 2 µm spacing, wet-inking for the 8.5 µm spacing stamp (Figure 2a). Contact-inking was carried out by placing the stamp onto an ink pad (e.g. a microscope slide loaded with a thin layer of polymer ink, Figure 2a; stamp = light blue, ink = green layer). This method helped to localize the ink on the protruding areas of the stamp, and thus minimized lateral ink diffusion on the substrate during stamping.75-76 For the largest spacing (8.5 µm), the contact-inking method failed, and the patterns obtained contained many defects. This is consistent with literature reports: contact-inking was initially developed to load stamps with smaller features dimensions,76 as it provided better resolution (i.e. less ink


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diffusion) compared to wet-inking, yet often had a higher defect density.77 This was also observed in the case of the 8.5 µm stamp used here: with contact-inking, the small amount of ink on the ink pad was not adsorbed uniformly by the larger surface features. When wet-inking was used, where sufficient ink solution was spread onto the stamp using a pipette (Figure 2a),6364

regular patterns were accessible (Figure 3). The substrates used for µCP were either gold, silicon, or glass. They were chemically

modified with benzophenone anchor groups following literature procedures.27, 72 Silicon, glass, or other substrates with OH groups were treated with triethoxy-benzophenone silane (3EBP), where the ethoxy group of 3EBP reacted with the OH groups of the substrate upon curing at 120°C (Figure 2b). Gold substrates were functionalized with benzophenone attached to lipoic acid disulfide (LS-BP), where the sulfur atoms formed a covalent bond when in contact with gold (not shown). First, monofunctional polymer surfaces containing only SMAMP patches were produced. To obtain these surfaces, the SMAMP-loaded stamp was directly brought into conformal contact with a benzophenone-modified surface with a defined printing pressure. The stamp was removed from the surface after a defined printing time, and the transferred ink features were UV-irradiated at λ = 254 nm, so that 3D microstructured SMAMP hydrogel patches were obtained (Figure 2b). To prepare bifunctional surfaces, first BP-COU-PSB was spin coated onto a benzophenone-modified substrate and cross-linked by UV irradiation (Figure 2c, blue network), so that a homogeneous polyzwitterionic hydrogel was formed. The two different SMAMP inks were then printed onto this base layer in the same way as described for the monofunctional materials (Figure 2c). Notably, the ink concentration was carefully optimized for each stamp spacing (5, 2 and 3 mg mL-1 for the 1, 2 and 8.5 µm stamps, respectively) to find the concentration that yielded the least amount of structural defects. After UV irradiation, any loosely attached polymer chains were removed by a washing step using a good solvent 17 Environment ACS Paragon Plus

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(trifluoroethanol for BP-COU-PSB, toluene for the SMAMP inks). Altogether, 12 types of microstructured surfaces with different spacing and height were obtained: 1µm-Short, 2µmShort, 8.5µm-Short and 1µm-Tall, 2µm-Tall, 8.5µm-Tall, each as monofunctional and bifunctional surfaces (Figure 2d).

Figure 2. Cartoon illustration of the microcontact printing (µCP) process (not drawn to scale): a) PDMS stamps (light blue) were loaded with NBD-SMAMP or BP-NBD-SMAMP ink (green) via contact-inking (1 μm or 2 μm spacing) or wet-inking (8.5 μm spacing). b) Fabrication of monofunctional surfaces: the substrate was functionalized with benzophenonecontaining anchor groups (BP, orange) followed by μCP, giving a monofunctional microstructured surface. The SMAMP formed 3D hydrogel patches (green) that were covalently attached to the substrate via 3EBP after UV irradiation. c) Fabrication of bifunctional surfaces: BP-COU-PSB (blue network) was spin-coated onto a BP functionalized substrates and UV irradiated to form a surface-attached polymer network. The respective SMAMP inks were then applied by μCP. They were UV irradiated and thereby simultaneously cross-linked and surface-attached to the PSB base layer, forming 3D patches (green). d) Depending on the stamp spacing and the SMAMP ink used for the µCP process, mono- and bifunctional microstructured surfaces with different heights and spacing were obtained. NBD-SMAMP gave short surface features, BP-NBD-SMAMP, which had a higher cross-linking efficiency, gave tall surface features.


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Three unstructured polymer surfaces were prepared as reference surfaces for the biological studies: a polyzwitterionic BP-COU-PSB network, a SMAMP monolayer (SMAMP ML) made from NBD-SMAMP, and a SMAMP network (SMAMP NW) made from BP-NBDSMAMP. To activate the SMAMP patches (i.e. to remove the Boc protective groups on the amines), the surfaces were immersed into HCl solution (4 M in dioxane). This generated antimicrobially active, polycationic SMAMPs with NH3+ groups (Figure 1e).27 Physical Characterization. The structured polymer-functionalized surfaces were first characterized by fluorescence or confocal microscopy to probe the large-scale homogeneity of the printed patterns. Representative results are shown in Figure 3a, indicating that regular and well-defined surface features with the expected pattern and spacing were obtained both for the monofunctional and the bifunctional surfaces. In case of the monofunctional microstructured surfaces, the black space between the green-fluorescent polymer dots are the non-functionalized parts of the silicon wafer. For the bifunctional materials, a blue-fluorescent polyzwitterionic background with bright green-fluorescent SMAMP patches was observed (Figure 3a). The average thickness of the material added after each fabrication step was determined using surface plasmon resonance spectroscopy (SPR). This is shown for one representative sample (1µmShort) in Figure 3b. Gold-coated substrates, which allowed excitation of surface plasmons, were used as substrate for these experiments. As expected, the reflectivity minimum of the curves shifted to higher angles after each fabrication step, indicating an increase of the average layer thickness on the gold substrate. The results obtained for the other bifunctional surfaces are shown in Figure S3. The thickness of each layer was determined by modelling the reflectivity curves with Fresnel equations using the Winspall software; the results thus obtained are listed in Table S2.


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Figure 3. a) Optical micrographs of the BP-COU-PSB hydrogel and the microstructured monoand bifunctional materials obtained with NBD-SMAMP ink, on silicon wafers (1 μm spacing: confocal microscopy; 2 and 8.5 µm: fluorescence microscopy with green GFP and blue DAPI filters); b) Surface plasmon resonance (SPR) reflectivity curves taken after each process step during build-up of the bifunctional materials. Average layer thickness (d) of each layer: gold 46 nm; BP-COU-PSB network - 78 nm; NBD-SMAMP - 31 nm.


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More details of the surface morphology of each sample type were revealed by atomic force microscopy (AFM, Figure 4). For the unstructured BP-COU-PSB hydrogel, a relatively high surface roughness of 14  5 nm was observed due to partial dewetting on the 3EBP functionalized substrate, which however did not have an adverse effect on the proteinrepellency of the material (see below). AFM also revealed that the height of the printed polymer patches was quite uniform within each sample (see height profiles in Figure 4), while the height of the patches on different samples varied with the stamp spacing and the ink used. Since the mono- and bifunctional materials obtained from the same stamp/ink combination had similar heights, it can be excluded that the surface properties of the substrate had a major effect on the feature heights. The highest 3D patches were obtained using PDMS stamp with 2 µm spacing (80 ± 20 nm when using the NBD-SMAMP ink (= 2µm-Short), 750 ± 250 nm with the BPNBD-SMAMP ink (=2µm-Tall)). The 1µm-Short features had a height of 45 ± 15 nm, and the 8.5µm-Short features were only 25 ± 10 nm. The 1µm-Tall samples had an average feature height of 200 ± 50 nm; the 8.5µm-Tall sample features were only 100 ± 20 nm high. It is difficult to make out a trend between polymer ink concentration, stamp morphology and the height of the printed features since the optimized printing conditions required different ink concentrations for each sample. Also, not only did the spacing of the PDMS stamp vary; at the same time, the height of the PDMS surface features was not uniform so that their relative aspect ratio could be maintained (see Figure S1 for AFM images of the PDMS stamps). Thus, the surface area of the stamps varied widely, resulting in a different amount of ink per unit area of substrate transferred for each stamp. The effect of the roughness of the BP-COU-PSB layer on the morphology of the SMAMP patches of the bifunctional surfaces can be also seen in the AFM height images. While the SMAMP patches of the monofunctional surfaces had quite sharp features, those found in the ‘short’ bifunctional microstructures were more blurred, indicating that these 3D patches followed the morphology of the underlying polyzwitterionic network (Figure 4). This is plausible since the base layer thickness was about 80 nm, and that of the 21 Environment ACS Paragon Plus

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SMAMP patches 25-80 nm. Such defects were not visible in the ‘tall’ patterns, which were of equal height or higher than the underlying network (100-750 nm, respectively).

Figure 4. AFM height images of the BP-COU-PSB hydrogel (R = roughness) and the monofunctional and bifunctional surfaces with varied spacing and height (1 μm, 2 μm, and 8.5 μm; short and tall). The height profiles of the SMAMP features were generated from diagonal cross-sections; Avg = average. For completeness, contact angles of the functionalized surfaces were also measured and are given in Table S1 in the supporting information. Except for the observation that the Boc22 Environment ACS Paragon Plus

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protected materials were more hydrophobic than the activated ones (which was to be expected), no clear trend with respect of the effect of surface feature heights or spacing is observed in these data. Biological Characterization. General. To understand how the surface structuring affected the bioactivity of the above presented monofunctional and bifunctional polymer surfaces, their protein-repellency (using fibrinogen), compatibility with human gingival mucosal keratinocytes, and their antimicrobial activity against two bacteria was determined. Antimicrobial Activity. The antimicrobial activity of the materials against rod-shaped E. coli, a representative for Gram-negative bacteria, and spherical S. aureus, a member of the Gram-positive bacteria family, was tested using a previously reported standardized assay.74 With this assay, the number of surviving colony forming units (CFUs) on each material compared to an uncoated silicon wafer as growth control was determined (Figure 5). As expected, the unstructured polyzwitterionic hydrogel BP-COU-PSB that was used as base layer was not antimicrobial (E. coli: 82% CFUs, S. aureus: 92% CFUs). The antimicrobial activity of the unstructured SMAMP monolayer (SMAMP ML) and network (SMAMP NW) against E. coli was quite high (< 1% CFUs), i.e. close to previously reported values.74 The monofunctional materials with 1 µm spacing had a similarly high activity against E. coli (0.7% and 0.1% CFUs for 1µm-Short and 1µm-Tall, respectively), while the activity of the monofunctional materials with 8.5 µm spacing was significantly lower (38% CFUs for 8.5µmShort; 5% CFUs for 8.5µm-Tall). In the case of S. aureus, the surviving CFUs on the unstructured SMAMPs were higher than expected (25% CFUs for the monolayer; 16 % for the network) compared to a previous publication.74 Surprisingly, the activity of monofunctional 1µm-Short and 1µm-Tall against S. aureus was significantly higher (7% and 5% CFUs, respectively). At the 8.5 µm spacing, the monofunctional materials were only weakly


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antimicrobial (more than 60% CFUs for 8.5µm-Short and 8.5µm-Tall, respectively) against S. aureus.

Figure 5. Antimicrobial activity of a) monofunctional surfaces and b) bifunctional surfaces against E. coli and S. aureus bacteria, respectively. The results are expressed as percentage of surviving colony forming units (CFUs) normalized to growth control (uncoated silicon wafer). SMAMP ML = SMAMP monolayer made from NBD-SMAMP and SMAMP NW = SMAMP network made from BP-NBD-SMAMP.

When interpreting this data, one has to carefully distinguish between intrinsic polymer properties and structural effects. In our previous work on homogeneously coated SMAMP surfaces, we found that these materials generally had a stronger antimicrobial activity against E. coli than against S. aureus.40, 57, 74 This is also consistent with results from other groups on different cationic polymers,78 and may be related to the higher negative surface potential of E. coli and the larger surface contact area per bacterium compared to S. aureus.79-80 The observed 24 Environment ACS Paragon Plus

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relative difference in activity against the two bacteria seen here is thus not a surface structuring effect. As to the effect of surface feature dimensions, two effects can be seen: first, for all monofunctional materials, whether structured (1 µm, 8.5 µm) or not (SMAMP ML and SMAMP MW), the thicker samples (Tall vs Short, NW vs. ML) had a better antimicrobial activity against E. coli and S. aureus than the thinner ones with the same lateral dimensions. This is in line with previous publications that investigated the effect of layer thickness of unstructured coatings on antimicrobial activity.74 Our original interpretation of this was that the thicker coatings had less defects or covered the surface more completely, so that bacteria would not sense the underlying substrate.74 However, since the microstructured monofunctional surfaces here reported have a lot of intentional 'defects', one should consider an alternative interpretation. This surface feature height-dependency could also be related to the ability of bacteria to interact with these surface features. Soft, thicker layers and higher surface features potentially have a higher deformability and thus might offer a larger contact area and/or number of available charges to the bacteria than thinner layers. A second effect of the surface patterning can be seen when comparing the two monofunctional materials with different spacing. The loss of activity of the materials with the 8.5 µm spacing is probably related to the relative dimensions of bacteria and the pattern. The spacing refers to the peak-to-peak distance of the surface features. For the 8.5 µm samples, the base width of the stamped features was around 3 – 5 µm (Figure 4), so that the uncoated areas have a width of around 3 – 5 µm. Thus, when S. aureus (diameter around 0.6 μm)81 or E. coli bacteria (length 2 – 4.5 µm; diameter 0.8 – 2.3 µm in our samples) came into contact with this material, some of them could adhere to the silicon patches between the protruding SMAMP features and thus maintained their viability. For the monofunctional materials with the 1 µm spacing, this was not possible since the silicon patches between the SMAMP features were too small, so that bacteria were forced into contact with the SMAMP features, and were killed. The somewhat higher activity of the 1 µm SMAMP patches compared to the homogenously coated SMAMPs could be an additional microstructuring effect, 25 Environment ACS Paragon Plus

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as bacterial proliferation could be hindered by the presence of the protruding features, in line with literature reports.49-52 On the bifunctional materials with the BP-COU-PSB base layer and SMAMP patches with the small spacing, quantitative killing of both types of bacteria (0% CFUs, within the accuracy limit of the method) was observed (for 1µm-Short, 1µm-Tall, 2µm-Short, and 2µm-Tall, respectively). This activity became much lower for the materials with 8.5 µm spacing against S. aureus. Here, the percentage of surviving CFUs was 8% on 8.5µm-Short and 2% on 8.5µmTall. Notably, the activity of these materials against E. coli was still significant (0.02% CFUs for 8.5µm-Short, 0% CFUs for 8.5 µm-Tall). The improved activity of the bifunctional materials compared to the monofunctional materials can be clearly assigned to the presence of the polyzwitterionic protein-repellent BP-COU-PSB base layer, which prevents bacterial adhesion (and thus survival) on the areas between the SMAMP features. The trends observed for the monofunctional materials were also found in the bifunctional materials: first, the materials were overall more active against E. coli due to the chemical nature of the polymer; second, taller features had higher antimicrobial activity than the shorter ones; and third, smaller spacings resulted in a better antimicrobial activity. Importantly, the data indicates that the upper limit of full antimicrobial activity against both Gram-positive and Gram-negative bacteria is between 2 µm and 8.5 µm spacing. For S. aureus alone, the upper limit of full antimicrobial activity seems to be at a somewhat lower value (more near 2 µm), possibly due to its smaller size compared to E. coli. E. coli, on the other hand, with a length of up to 4.5 µm, can hardly avoid the SMAMP patches of the bifunctional surfaces with 8.5 µm spacing regardless its position or orientation, and thus gets near-quantitatively killed even at that larger spacing. In summary, the results of the monofunctional materials show that microstructuring indeed helped to reduce the number of surviving CFUs as long as their dimension was close to the bacteria cell dimensions. Additionally, the data indicates that the combination of 26 Environment ACS Paragon Plus

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microstructuring and surface modification with an antimicrobial polymer alone (i.e., the monofunctional surfaces) was not sufficient to fully inhibit bacterial growth. This could be only achieved when the polyzwitterionic BP-COU-PSB layer was present and masked the otherwise inactive substrate background. On these samples, quantitative reduction of bacterial growth was observed when the spacing of the SMAMP patches matched the dimensions of the bacteria. Protein Repellency. The protein-repellency of the materials was studied by exposing the samples to fibrinogen solution and monitoring this interaction with SPR. Two types of experiments were performed. The build-up of protein layers was recorded in situ by SPR kinetics experiments, i.e. by time-dependent measurement of reflective intensity at constant angle. Representative results are shown in Figure 6a both for the monofunctional and bifunctional materials (using 1µm-Short). In a typical experiment, first the baseline reflectivity of the sample against buffer was recorded for 20 – 25 min; then, fibrinogen solution was injected (Figure 6a, first arrow). The monofunctional surface had an increase of reflectivity which indicated that the surface was protein-adhesive (Figure 6a, grey curve). The reflectivity value measured when exposing the bifunctional surfaces to protein remained constant, indicating that no fibrinogen adsorbed (Figure 6a, black curve). After about 15 – 20 min (40-45 min in total), the flow cell was again flushed with buffer to remove any reversibly attached fibrinogen (Figure 6a, second arrow). No decrease of reflectivity was then detected for the monofunctional surface, which confirmed that all fibrinogen molecules were irreversibly adsorbed on that surface. The curve of the bifunctional surface (Figure 6a, black curve) again did not alter, showing that neither reversible nor irreversible protein adhesion had occurred. In addition to these qualitative experiments, full angular reflectivity curves of the dry samples were taken before the protein adhesion experiments, and afterwards when the samples had been washed with buffer followed by water, and dried. By modelling the thus obtained curves using the Fresnel equations (Winspall software), the mass of adsorbed fibrinogen per unit area (ng mm-2) could be


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determined (Figure 6b). An increase of layer thickness due to protein adsorption would cause the plasmon minimum to shift toward higher angles, as seen in the data obtained from the monofunctional surface (Figure 6b, grey solid line = before fibrinogen exposure, black dashed line = after fibrinogen exposure). In contrast, no plasmon minimum shift was observed from reflectivity curves of the bifunctional surface, suggesting no fibrinogen adsorbed on this material (Figure 6b, light blue solid line = before fibrinogen exposure, blue dashed line = after fibrinogen exposure). The kinetics experiments and angular reflectivity curves of other sample surfaces are shown in Figure S4 and S5, together with the data of the unstructured reference samples. The average layer thickness of adhered protein on each material thus obtained is summarized in Figure 6c. The data shows that, expectedly, the protein adsorption on BP-COUPSB was below the detection limit of surface plasmon resonance spectroscopy, i.e. < 0.1 ng mm-2. For the monofunctional SMAMP surfaces, the amount of protein adhesion depended on the thickness of the surface features: more protein adhered to the thin SMAMP ML than onto SMAMP NW, and more protein adhered to the Short features than to the Tall ones. The tall features and the network should have a lower polymer segment density at the polymer-liquid interface than the short features and the monolayer; and being further away from the substrate, probably also a lower local elastic modulus. This may lead to the observed lower amount of adhered protein, despite the higher relative surface area of some of the tall features compared to the short ones (see Calculation S1 in the supporting information). The effect of spacing of the monofunctional materials on the protein adhesion is less clear, as there is no observable trend in the data. Because each of these features has a different height, it is possible that height and spacing effects may interfere with each other, so that the effect of each is veiled.


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Figure 6. a) Surface plasmon resonance (SPR) kinetics curves of fibrinogen adhesion on monofunctional and bifunctional 1µm-Short samples. The time point of fibrinogen injection is indicated by the first arrow, and buffer injection is marked by the second arrow. Fibrinogen strongly adhered to the monofunctional material (grey), while no fibrinogen adsorption was observed on the bifunctional material (black). b) SPR angular reflectivity curves of 1µm-Short samples in the dry state before (solid line) and after (dashed line) exposure to fibrinogen injection. c) Average amount of adhered fibrinogen (in ng mm-2) on the unstructured reference surfaces (gold), the monofunctional microstructured surfaces (black), and bifunctional microstructured surfaces (grey) as determined by SPR spectroscopy.

For the bifunctional surfaces with the polyzwitterionic BP-COU-PSB base layer and the 3D SMAMP patches, the amount of adsorbed protein was 0 ng mm-2 in all cases (i.e. below the detection limit of the method of < 0.1 ng mm-2). The fact that no protein adhesion was observed for these samples even at larger spacings was insofar surprising as the size of fibrinogen (with 29 Environment ACS Paragon Plus

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a length of about 50 nm)45 is well below the size of the all spacings used. Thus, this negatively charged protein should be able to come into contact with the cationic SMAMP patches, and adhere to them via electrostatic interactions. Therefore, the inability of the protein to adhere is not a steric effect, but must be energy-related. As mentioned above, the 3D SMAMP patches are not rigid objects with a defined surface, but rather soft materials with a decreasing segment density at the interface. For uncharged hydrophilic polymer networks having such interfaces, it was found that proteins cannot adhere.82 In such cases, the H term in the free energy equation was close to zero, and it was argued that the TS term in the free energy equation for this process is negative because protein adsorption required deformation of the polymer chains, which would lead to an entropy loss due to further chain stretching. This would lead to an overall positive G for the protein adsorption process.82 In the case of the SMAMP patches, the H term should be negative due to the opposite charges of the protein and the SMAMPs; however, the entropy loss of the many freely dangling polymer chain ends at the polymer-liquid interface might outweigh even this relatively large energetic contribution, so that the overall G for the protein adsorption process would still be positive and thus unfavorable. Cell Compatibility. The toxicity of the unstructured and microstructured polymer surfaces was tested with immortalized gingival mucosal keratinocytes (as a representative for human cells) using the Alamar Blue Assay. In this assay, the reduction of the dye resazurin is monitored, which is way to quantify the metabolic activity of the cells.83 For the Alamar blue assay, keratinocytes were plated out on the test surfaces and cultivated for 24, 48, and 72 h, respectively. The relative dye reduction at these time points for cells grown on the here presented sample set (normalized to the dye reduction by cells grown on an uncoated substrate as growth control) is shown in Figure 7. Absolute values of dye reduction can be found in Figure S6. The data shows that the cell metabolism on all monofunctional and bifunctional polymer surfaces was at least on the level of the growth control, i.e. all materials were non-toxic. 30 Environment ACS Paragon Plus

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Notably, the highest dye reduction was observed for monofunctional 8.5µm-Tall, with about 175% dye reduction compared to the control. The cells grew even better on the bifunctional surfaces than on monofunctional surfaces. For the 1 µm and the 8.5 µm spacing, a strong difference is observed between the short and the tall features, with significantly more growth on the tall features. Bifunctional 8.5µm-Tall had the highest relative dye reduction of all samples tested (about 190%). The data were confirmed by optical microscopy images (Figure S7), which illustrated that there was no significant difference in cell morphology between the keratinocytes grown on the functionalized surfaces and those in the growth control. The cell viability was further assessed qualitatively with the live-dead stain (Figure 8 for bifunctional surfaces and Figure S8 for monofunctional surfaces), which indicated that most of these cells were membrane-intact (green), and only a low amount was membrane-compromised (red). Thus, the combined cell compatibility data confirms that cell growth was not adversely affected by the presence of small microstructures (up to 2 µm). The increased proliferation on both types of 8.5µm-Tall surfaces (Figure 7) could be related to the better ability of the mammalian cells, which are much larger than bacteria, to adhere to these larger surface features. Thus, the height (100 ± 20 nm) and morphology is sensed by these cells. Interestingly, the 8.5 µm-Short surface, with a height of only 25 ± 10 nm, did not have this effect. So far, no general structure-activity rules exist as to how mammalian cells behave on micro- and nanostructured surfaces.84-85 Interestingly, the cells grew better on the bifunctional materials than on the monofunctional ones (Figure 7b). This could be attributed to the higher hydrophilicity of the bifunctional surfaces. It has been reported in many studies that more hydrophilic surfaces promoted cell response,86-88 although contradictive results have also been reported.89-90 Alternatively, the better cell growth on the bifunctional materials could be due to their lower elastic modulus compared to the monofunctional surfaces with bare substrate areas. Yet this is merely speculative since cell-surface interactions are rather complex and results depend not only on the cell and surface types used, but also on the exact experimental conditions. 31 Environment ACS Paragon Plus

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Figure 7. Cell viability of immortalized human gingival mucosal keratinocytes on a) monofunctional and b) bifunctional polymer surfaces, determined by the Alamar Blue Assay. Relative resazurin dye reduction was determined after 24, 48, and 72 h incubation time, respectively. The results are reported as percentage of dye reduction normalized to the growth control, an uncoated substrate, which was defined as 100% growth for each time point.


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Figure 8. Live-dead staining images of human cells keratinocytes grown on bifunctional microstructured surfaces after 72 h of incubation time. Growth control was uncoated substrate and dead control was uncoated substrate treated with isopropanol. Syto 16 (green) stained live cells and propidium iodide (red) stained dead cells. Merged images are the overlay of the live and dead cells images. 33 Environment ACS Paragon Plus

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CONCLUSION This study presents the fabrication of bifunctional polymer surfaces consisting of a polyzwitterionic base layer and cationic microstructured three-dimensional polymer patches by microcontact printing (µCP). To obtain these materials, first a protein-repellent polysulfobetaine (BP-COU-PSB, with blue fluorescent coumarin groups and the UV crosslinker benzophenone) was applied to a substrate. It was UV-cross-linked and surface-attached in one step by UV-triggered C,H insertion reactions. Polymer inks containing antimicrobially active polymers (poly(oxanorbornene)-based Synthetic Mimics of Antimicrobial Peptides, SMAMPs) were printed onto this polyzwitterionic base layer using microstructured PDMS stamps with different spacings (1 µm, 2 µm, and 8.5 µm). The thus obtained microstructured SMAMP ink patches were also UV cross-linked to form 3D surface-attached polymer features. As the cross-linking in the UV irradiation step took place throughout the entire polymer volume and not just at the surface, it locked in the 3D surface architecture pre-determined by the microcontact printing step and made the resulting polymer patches stable in both organic solvents and aqueous conditions.66 Since C,H insertion reactions are applicable to many polymers and have been reported for the synthesis of other homogenous,65, 82 multilayered91 and microstructured92-93 materials (yet not in the context of microcontact printing), the here reported process should be applicable to numerous polymer combinations besides the presented example, and thus be of interest also for other applications. The two different SMAMP inks used here for µCP had different cross-linking efficiency: NBD-SMAMP contained repeat units carrying fluorescent nitrobenzoxadiazole (NBD) groups, a moderate UV cross-linker, and BP-NBD-SMAMP contained NBD and the highly efficient UV-active cross-linker benzophenone. With these inks, the height of the SMAMP features could be tuned. Thus, two series of bifunctional microstructured polymer surfaces were obtained: 1µm-Short, 2µm-Short, 8.5µm-Short, and 1 µm-Tall, 2µm-Tall, 8.5µm-Tall. For 34 Environment ACS Paragon Plus

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comparison, the corresponding monofunctional surfaces with the same heights and spacing were fabricated by directly printing the two SMAMP inks onto benzophenone-functionalized substrates. All materials were treated with HCl to activate the antimicrobial function of the SMAMPs. The physical characterization of the surface features (by AFM, fluorescence microscopy, contact angle measurements, and SPR) confirmed that well-defined microstructures with the expected topographies were obtained in all cases. The biological characterization of the materials revealed that the bifunctional surfaces with 1 µm and 2 µm spacing had 100% growth inhibition of E. coli and S. aureus bacteria. At a larger spacing, this broad-band antimicrobial activity was lost. All bifunctional surfaces also were quantitatively protein-repellent, regardless of their spacing and height. Monofunctional reference materials were not only less antimicrobially active, but also showed significant amounts of protein-adhesion at all spacings. These results show that the presence of the polyzwitterionic BP-COU-PSB base layer underneath the SMAMP patches was crucial for the simultaneous antimicrobial activity and protein-repellency of the material. In the protein adhesion experiments on monofunctional surfaces, reduced protein amounts were observed on the taller surface features compared to the shorter ones. This could be related to the relative polymer segment density of these materials at the polymer-liquid interface. It is known from literature that protein adsorption on highly crosslinked hydrogel is energetically favorable, unlike protein adsorption on loosely crosslinked hydrogel.82, 94 The former should also have a higher segment density at the interface, and a higher elastic modulus than the latter. The here presented tall surface features, particularly the bifunctional hydrogels, should have a low polymer segment density at the polymer-liquid interface. The potential entropy loss upon permanent protein adhesion seems to compensate the potential energy gain by the electrostatically driven interaction between SMAMP chain segments and proteins, so that overall the free enthalpy of this process is positive and no protein


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adheres to the bifunctional surfaces. In our previous work on bifunctional polymer surfaces made from polyzwitterions and SMAMPs,45 we found that structured surfaces obtained by colloidal lithography were fully protein-repellent at a spacing of 200 nm and 500 nm, and started to adhere small amounts of protein at 1 µm spacing. These materials were bifunctional polymer monolayers and thus should both have a higher polymer segment density at the polymer-liquid interface (due to steric constraints), and also a higher local elastic modulus (due to the underlying substrate) than the here presented structured polymer hydrogels. This could be an explanation, in line with literatures,82, 94 why the bifunctional materials here presented were fully protein-repellent even at the largest spacings of 8.5 µm. Thus, protein-repellency is strongly affected by the polymer architecture and its effects on physical properties, besides the material spacing and chemical nature of its components. The previously reported bifunctional materials obtained by colloidal lithography had a reduced antimicrobial activity against E. coli at a spacing of 200 and 500 nm and were only fully active at a spacing of 1 µm.45 Presumably, the small antimicrobial polymer patches did not sufficiently interact with the bacterial membranes to cause perturbation or damage. In the here presented data, the upper limit for full activity against E. coli and S. aureus, irrespective of the feature height, was > 2 µm. Thus, the optimum for simultaneous antimicrobial activity and protein repellency of such bifunctional materials, whether obtained by colloidal lithography or by microcontact printing, is at spacings between 1 and 2 µm. In that range, the patch sizes of antimicrobial moieties is sufficiently large to be lethal, yet the materials are still protein repellent due to their surface architecture. In the cell experiments, all monofunctional and bifunctional surfaces here presented were compatible with human gingival mucosal keratinocytes. Thus, this study shows that it is possible to find a 'sweet spot' where the surface characteristics of a material are so well balances that they are conducive to cell adhesion and at the same time, the proliferation of bacterial pathogens is prevented. The remaining challenge


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for this class of materials is to generalize these findings, e.g. by quantify the physical properties that lead to the observed optimal bioactivity. Additionally, further studies such as human blood plasma and serum adhesion, bacterial adhesion, biofilm formation, and long term stability tests will reveal if this kind of material is truly useful for biomedical applications.

ASSOCIATED CONTENT Supporting Information. The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir....... - Methodological details for the biological characterizations: protein adhesion test, antimicrobial activity assay, cell compatibility assay (Alamar Blue Assay), and live-dead staining of keratinocytes grown on substrates. - Figures of PDMS stamp height profiles (AFM); assignment of 1H-NMR shifts of BP-COUPSB; SPR angular reflectivity curves of build-up of bifunctional surfaces; SPR kinetic experiments; angular reflectivity curves of protein adhesion tests; absolute values of dye reduction of cell compatibility assay; optical micrographs of human gingival mucosal keratinocytes grown on mono- and bifunctional surfaces; fluorescence micrographs of human gingival mucosal keratinocytes grown on monofunctional surfaces. - Tables: Contact angle measurements and SPR fitting parameter for the reflectivity curves of the bifunctional surfaces. - Calculation of the surface area of the monofunctional SMAMP materials.

AUTHOR INFORMATION Corresponding Author. *E-mail [email protected] (K.L.). ORCID 37 Environment ACS Paragon Plus

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Karen Lienkamp: 0000-0001-6868-3707

Funding. Funding of this work by the Ministry for Science, Research and Art of the State of BadenWürttemberg, Germany (GenMik I + II graduate school), and the European Research Council (ERC-StG REGENERATE) is gratefully acknowledged.

Notes. The authors declare no competing financial interest.

Acknowledgements Vera Bleicher and Diana Lorena Guevara Solarte are gratefully acknowledged for performing the antimicrobial activity assay. Dr. Sibylle Rau and Dr. Alice Eickenscheidt are gratefully acknowledged for performing the cell compatibility assay and live-dead staining of the cells. Dr. Marie Follo, Core-Facility, Universitätklinikum Freiburg, Germany, is gratefully acknowledged for imaging the 1 µm structures with confocal microscopy.

Received: ((will be filled in by the editorial staff)) Revised: ((will be filled in by the editorial staff)) Published online: ((will be filled in by the editorial staff))

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58. Lienkamp, K.; Madkour, A. E.; Musante, A.; Nelson, C. F.; Nüsslein, K.; Tew, G. N., Antimicrobial Polymers Prepared by ROMP with Unprecedented Selectivity: A Molecular Construction Kit Approach. J. Am. Chem. Soc. 2008, 130 (30), 9836-9843. 59. Lienkamp, K.; Tew Gregory , N., Synthetic Mimics of Antimicrobial Peptides—A Versatile Ring-Opening Metathesis Polymerization Based Platform for the Synthesis of Selective Antibacterial and Cell-Penetrating Polymers. Chem. Eur. J. 2009, 15 (44), 1178411800. 60. Zhang, Z.; Chen, S.; Chang, Y.; Jiang, S., Surface Grafted Sulfobetaine Polymers via Atom Transfer Radical Polymerization as Superlow Fouling Coatings. J. Phys. Chem. B 2006, 110 (22), 10799-10804. 61. Zhang, Z.; Chao, T.; Chen, S.; Jiang, S., Superlow Fouling Sulfobetaine and Carboxybetaine Polymers on Glass Slides. Langmuir 2006, 22 (24), 10072-10077. 62. Alom Ruiz, S.; Chen, C. S., Microcontact printing: A tool to pattern. Soft Matter 2007, 3 (2), 168-177. 63. Kaufmann, T.; Ravoo, B. J., Stamps, inks and substrates: polymers in microcontact printing. Polymer Chemistry 2010, 1 (4), 371-387. 64. Perl, A.; Reinhoudt David, N.; Huskens, J., Microcontact Printing: Limitations and Achievements. Adv. Mater. 2009, 21 (22), 2257-2268. 65. Prucker, O.; Brandstetter, T.; Rühe, J., Surface-attached hydrogel coatings via C,Hinsertion crosslinking for biomedical and bioanalytical applications (Review). Biointerphases 2017, 13 (1), 010801. 66. Widyaya, V. T.; Riga, E. K.; Müller, C.; Lienkamp, K., Submicrometer-Sized, 3D Surface-Attached Polymer Networks by Microcontact Printing: Using UV-Cross-Linking Efficiency To Tune Structure Height. Macromolecules 2018, 51 (4), 1409-1417. 67. Riga, K. E.; Saar, S. J.; Erath, R.; Hechenbichler, M.; Lienkamp, K., On the Limits of Benzophenone as Cross-Linker for Surface-Attached Polymer Hydrogels. Polymers 2017, 9 (12). 68. Riga Esther, K.; Boschert, D.; Vöhringer, M.; Widyaya Vania, T.; Kurowska, M.; Hartleb, W.; Lienkamp, K., Fluorescent ROMP Monomers and Copolymers for Biomedical Applications. Macromol. Chem. Phys. 2017, 218 (21), 1700273. 69. Rankin David, A.; P'Pool Steven, J.; Schanz, H.-J.; Lowe Andrew, B., The controlled homogeneous organic solution polymerization of new hydrophilic cationic exo-7oxanorbornenes via ROMP with RuCl2(PCy3)2CHPh in a novel 2,2,2trifluoroethanol/methylenechloride solvent mixture. J. Polym. Sci. A 2007, 45 (11), 2113-2128. 70. Colak, S.; Tew, G. N., Synthesis and Solution Properties of Norbornene Based Polybetaines. Macromolecules 2008, 41 (22), 8436-8440. 71. Trnka, T. M.; Grubbs, R. H., The Development of L2X2RuCHR Olefin Metathesis Catalysts: An Organometallic Success Story. Acc. Chem. Res. 2001, 34 (1), 18-29. 72. Gianneli, M.; Roskamp, R. F.; Jonas, U.; Loppinet, B.; Fytas, G.; Knoll, W., Dynamics of swollen gel layers anchored to solid surfaces. Soft Matter 2008, 4 (7), 1443-1447. 73. Kurowska, M.; Eickenscheidt, A.; Guevara-Solarte, D.-L.; Widyaya, V. T.; Marx, F.; Al-Ahmad, A.; Lienkamp, K., A Simultaneously Antimicrobial, Protein-Repellent, and CellCompatible Polyzwitterion Network. Biomacromolecules 2017, 18 (4), 1373-1386. 74. Al-Ahmad, A.; Zou, P.; Solarte, D. L. G.; Hellwig, E.; Steinberg, T.; Lienkamp, K., Development of a Standardized and Safe Airborne Antibacterial Assay, and Its Evaluation on Antibacterial Biomimetic Model Surfaces. PLoS One 2014, 9 (10), e111357. 75. Pompe, T.; Fery, A.; Herminghaus, S.; Kriele, A.; Lorenz, H.; Kotthaus, J. P., Submicron Contact Printing on Silicon Using Stamp Pads. Langmuir 1999, 15 (7), 2398-2401. 76. Libioulle, L.; Bietsch, A.; Schmid, H.; Michel, B.; Delamarche, E., Contact-Inking Stamps for Microcontact Printing of Alkanethiols on Gold. Langmuir 1999, 15 (2), 300-304.


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77. Fujihira, M.; Furugori, M.; Akiba, U.; Tani, Y., Study of microcontact printed patterns by chemical force microscopy. Ultramicroscopy 2001, 86 (1), 75-83. 78. Kugler, R.; Bouloussa, O.; Rondelez, F., Evidence of a charge-density threshold for optimum efficiency of biocidal cationic surfaces. Microbiology 2005, 151, 1341-8. 79. Dickson, J. S.; Koohmaraie, M., Cell surface charge characteristics and their relationship to bacterial attachment to meat surfaces. Applied and Environmental Microbiology 1989, 55 (4), 832-836. 80. Guo, S.; Jańczewski, D.; Zhu, X.; Quintana, R.; He, T.; Neoh, K. G., Surface charge control for zwitterionic polymer brushes: Tailoring surface properties to antifouling applications. J. Colloid Interface Sci. 2015, 452, 43-53. 81. Tripathy, A.; Sen, P.; Su, B.; Briscoe, W. H., Natural and bioinspired nanostructured bactericidal surfaces. Adv. Colloid Interface Sci. 2017, 248, 85-104. 82. Wörz, A.; Berchtold, B.; Moosmann, K.; Prucker, O.; Rühe, J., Protein-resistant polymer surfaces. J. Mater. Chem. 2012, 22 (37), 19547-19561. 83. Rampersad, S. N., Multiple Applications of Alamar Blue as an Indicator of Metabolic Function and Cellular Health in Cell Viability Bioassays. Sensors 2012, 12 (9), 12347-12360. 84. Martínez, E.; Engel, E.; Planell, J. A.; Samitier, J., Effects of artificial micro- and nanostructured surfaces on cell behaviour. Ann. Anat. 2009, 191 (1), 126-135. 85. Nikkhah, M.; Edalat, F.; Manoucheri, S.; Khademhosseini, A., Engineering microscale topographies to control the cell–substrate interface. Biomaterials 2012, 33 (21), 5230-5246. 86. Paterlini, T. T.; Nogueira, L. F. B.; Tovani, C. B.; Cruz, M. A. E.; Derradi, R.; Ramos, A. P., The role played by modified bioinspired surfaces in interfacial properties of biomaterials. Biophys. Rev. 2017, 9 (5), 683-698. 87. Dhirendra, S. K.; Rajesh, V.; Kirubanandan, S., Improved Biomaterials for Tissue Engineering Applications: Surface Modification of Polymers. Curr. Top. Med. Chem. 2008, 8 (4), 341-353. 88. Gittens, R. A.; Scheideler, L.; Rupp, F.; Hyzy, S. L.; Geis-Gerstorfer, J.; Schwartz, Z.; Boyan, B. D., A review on the wettability of dental implant surfaces II: Biological and clinical aspects. Acta Biomater. 2014, 10 (7), 2907-2918. 89. Chen, S.; Lu, X.; Zhu, D.; Lu, Q., Targeted grafting of thermoresponsive polymers from a penetrative honeycomb structure for cell sheet engineering. Soft Matter 2015, 11 (37), 74207427. 90. Kim, M. S.; Park, S.; Chun, H. J.; Kim, C. H., Thermosensitive Hydrogels for Tissue Engineering. J. Tissue Eng. Regen. Med. 2011, 8, 117-123. 91. Schuh, K.; Prucker, O.; Ruehe, J., Tailor-Made Polymer Multilayers. Adv. Funct. Mater. 2013, 23 (48), 6019-6023. 92. Moschallski, M.; Evers, A.; Brandstetter, T.; Rühe, J., Sensitivity of microarray based immunoassays using surface-attached hydrogels. Anal. Chim. Acta 2013, 781, 72-79. 93. Kleber, C.; Bruns, M.; Lienkamp, K.; Rühe, J.; Asplund, M., An interpenetrating, microstructurable and covalently attached conducting polymer hydrogel for neural interfaces. Acta Biomater. 2017, 58 (Supplement C), 365-375. 94. Pandiyarajan, C. K.; Prucker, O.; Zieger, B.; Rühe, J., Influence of the Molecular Structure of Surface-Attached Poly(N-alkyl Acrylamide) Coatings on the Interaction of Surfaces with Proteins, Cells and Blood Platelets. Macromol. Biosci. 2013, 13 (7), 873-884.


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Figure 1. Cartoon illustration of the chemical composition and build-up of the targeted bifunctional surface: a) The protein-repellent polyzwitterion BP-COU-PSB containing UV cross-linkable benzophenone groups (BP, black part of the polymer structure in c) was applied to a BP-functionalized substrate. Upon UV irradiation, covalent bonds to the surface BP groups (grey dots) formed, and the BP groups within the polymer chains formed polymer-polymer cross-links (blue dots). b) An antimicrobial polymer (NBD-SMAMP or BP-NBDSMAMP) was printed onto the BP-COU-PSB base layer by microcontact printing (µCP). Upon UV irradiation, covalent bonds to the base hydrogel and within the (BP-)NBD-SMAMP patches formed through the BP or NBD groups. All polymers were synthesized via ring-opening metathesis polymerization: c) BP-COU-PSB from PSB, COU and BP monomers and d) BP-NBD-SMAMP from SMAMP, NBD and BP monomers, as well as NBD-SMAMP from SMAMP and NBD monomers; e) the SMAMP patches were activated on the surface with HCl, which removed the Boc protective groups and yielded antimicrobially active NH3+ groups. 173x259mm (96 x 96 DPI)

ACS Paragon Plus Environment

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Figure 2. Cartoon illustration of the microcontact printing (µCP) process (not drawn to scale): a) PDMS stamps (light blue) were loaded with NBD-SMAMP or BP-NBD-SMAMP ink (green) via contact-inking (1 μm or 2 μm spacing) or wet-inking (8.5 μm spacing). b) Fabrication of monofunctional surfaces: the substrate was functionalized with benzophenone-containing anchor groups (BP, orange) followed by μCP, giving a monofunctional microstructured surface. The SMAMP formed 3D hydrogel patches (green) that were covalently attached to the substrate via 3EBP after UV irradiation. c) Fabrication of bifunctional surfaces: BPCOU-PSB (blue network) was spin-coated onto a BP functionalized substrates and UV irradiated to form a surface-attached polymer network. The respective SMAMP inks were then applied by μCP. They were UV irradiated and thereby simultaneously cross-linked and surface-attached to the PSB base layer, forming 3D patches (green). d) Depending on the stamp spacing and the SMAMP ink used for the µCP process, monoand bifunctional microstructured surfaces with different heights and spacing were obtained. NBD-SMAMP gave short surface features, BP-NBD-SMAMP, which had a higher cross-linking efficiency, gave tall surface features. 423x351mm (96 x 96 DPI)


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Figure 3. a) Optical micrographs of the BP-COU-PSB hydrogel and the microstructured mono- and bifunctional materials obtained with NBD-SMAMP ink, on silicon wafers (1 μm spacing: confocal microscopy; 2 and 8.5 µm: fluorescence microscopy with green GFP and blue DAPI filters); b) Surface plasmon resonance (SPR) reflectivity curves taken after each process step during build-up of the bifunctional materials. Average layer thickness (d) of each layer: gold - 46 nm; BP-COU-PSB network - 78 nm; NBDSMAMP - 31 nm. 110x320mm (96 x 96 DPI)


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Figure 4. AFM height images of the BP-COU-PSB hydrogel (R = roughness) and the monofunctional and bifunctional surfaces with varied spacing and height (1 μm, 2 μm, and 8.5 μm; short and tall). The height profiles of the SMAMP features were generated from diagonal cross-sections; Avg = average. 190x275mm (96 x 96 DPI)


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Figure 5. Antimicrobial activity of a) monofunctional surfaces and b) bifunctional surfaces against E. coli and S. aureus bacteria, respectively. The results are expressed as percentage of surviving colony forming units (CFUs) normalized to growth control (uncoated silicon wafer). SMAMP ML = SMAMP monolayer made from NBD-SMAMP and SMAMP NW = SMAMP network made from BP-NBD-SMAMP. 347x429mm (96 x 96 DPI)


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Figure 7. Cell viability of immortalized human gingival mucosal keratinocytes on a) monofunctional and b) bifunctional polymer surfaces, determined by the Alamar Blue Assay. Relative resazurin dye reduction was determined after 24, 48, and 72 h incubation time, respectively. The results are reported as percentage of dye reduction normalized to the growth control, an uncoated substrate, which was defined as 100% growth for each time point. 347x460mm (96 x 96 DPI)


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Figure 8. Live-dead staining images of human cells keratinocytes grown on bifunctional microstructured surfaces after 72 h of incubation time. Growth control was uncoated substrate and dead control was uncoated substrate treated with isopropanol. Syto 16 (green) stained live cells and propidium iodide (red) stained dead cells. Merged images are the overlay of the live and dead cells images. 184x308mm (96 x 96 DPI)


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Three-Dimensional, Bifunctional Microstructured Polymer Hydrogels

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