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Clickable Antifouling Polymer Brushes for Polymer Pen Lithography Uwe Bog, Andrés de los Santos, Summer Mueller, Shana Havenridge, Viviana Parrillo, Michael Bruns, Andrea E Holmes, Cesar Rodriguez-Emmenegger, Harald Fuchs, and Michael Hirtz ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b01184 • Publication Date (Web): 15 Mar 2017 Downloaded from http://pubs.acs.org on March 19, 2017

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ACS Applied Materials & Interfaces

Clickable Antifouling Polymer Brushes for Polymer Pen Lithography Uwe Bog,†,∇ Andres de los Santos Pereira,§,∇ Summer Mueller,¦ Shana Havenridge,¦ Viviana Parrillo,§ Michael Bruns,╓ Andrea E. Holmes,¦ Cesar Rodriguez-Emmenegger,*,ǁ Harald Fuchs,‡ and Michael Hirtz*,† †

Institute of Nanotechnology (INT) & Karlsruhe Nano Micro Facility (KNMF), Karlsruhe

Institute of Technology (KIT), Germany § Department of Chemistry and Physics of Surfaces and Biointerfaces, Institute of Macromolecular Chemistry ASCR, v.v.i., Czech Republic ¦

Department of Chemistry, Doane University, Crete, Nebraska, and the Center for Nanohybrid

Functional Materials (CNFM), University of Nebraska-Lincoln, USA ╓

Institute for Applied Materials (IAM) & Karlsruhe Nano Micro Facility (KNMF), Karlsruhe

Institute of Technology (KIT), Germany ǁ DWI – Leibniz Institute for Interactive Materials and Institute of Technical and Macromolecular Chemistry, RWTH Aachen University, Aachen, Germany ‡

Physical Institute & Center for Nanotechnology (CeNTech), University of Münster, Germany

KEYWORDS Polymer Brushes, Polymer Pen Lithography, PPL, Antifouling, Biofunctional interfaces

ACS Paragon Plus Environment

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ABSTRACT Protein-repellent reactive surfaces that promote localized specific binding are highly desirable for applications in the biomedical field. Nonspecific adhesion will compromise the function of bioactive surfaces, leading to ambiguous results of binding assays and negating the binding specificity of patterned cell-adhesive motives. Localized specific binding is often achieved by attaching a linker to the surface and the other side of the linker is used to bind specifically to a desired functional agent, as e.g. proteins, antibodies, fluorophores, depending on the function required by the application. We present a protein-repellent polymer brush enabling highly specific covalent surface immobilization of biorecognition elements by strain-promoted alkyneazide cycloaddition click chemistry for selective protein adhesion. The protein-repellent polymer brush is functionalized by highly localized molecular binding sites in the low micron range using polymer pen lithography (PPL). Due to the massive parallelization of writing pens, the tunable PPL printed patterns can span over square centimeter areas. The selective binding of the protein streptavidin to these surface sites is demonstrated while the remaining polymer brush surface is resisting non-specific adsorption without any prior blocking by bovine serum albumin (BSA). In contrast to the widely used BSA blocking, the reactive polymer brushes are able to significantly reduce non-specific protein adsorption, which is the cause of biofouling. This was achieved for solutions of single proteins as well as complex biological fluids. The remarkable fouling resistance of the polymer brushes has the potential to improve the multiplexing capabilities of protein probes and therefore impact biomedical research and applications.


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1. INTRODUCTION Molecular surface patterning requires reliable and stable coupling chemistry for localized chemical functionalization. Simultaneously, for many applications in the biomedical field, the surface should provide high protein repellence in non-patterned areas to prevent unwanted unspecific adhesion (fouling).1,2 This is usually achieved in a two-step approach: First, a desired surface pattern is created, allowing specific binding, and second, the other parts of the surface have to be chemically treated to make them inert and resistant to protein adsorption. Various strategies are usually exploited, the most common in biological labs being the passivation with an inert protein (frequently used are bovine serum albumin (BSA), goat serum, casein or other milk proteins).3–5 These proteins usually prevent further adsorption after they saturate the surface resulting in an interface resembling an antimetamorphic system. Blood compatibility is critical for medical implants and many other biomedical and bio-analytical devices.6 However, blood contacting surfaces require much more stringent passivation strategies, due to the analyte’s high molecular complexity. Avoiding these saturation and blocking steps would be highly desirable, as they can reduce the efficiency of the specific receptors bound to the surface7 and elicit undesired responses on their own.8 Importantly, although culturing cell patterns using BSA blocking for passivation is possible in serum-free media, pattern fidelity completely vanishes in serum-containing media, indeed cells may actively remove the blocking proteins over time.9,10 Therefore, a surface exhibiting chemically active moieties to allow for covalent functionalization, while being simultaneously protein-repellent would be highly desirable operating in complex media.11–13 Moreover, the adsorbed proteins (albumin) change conformation and lead to an activation of the coagulation cascade even if no further protein


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fouling occurs.14,15 Therefore, other strategies based on hydrophilic synthetic polymers have been explored. Remarkable advances have been made in surface modifications that are able to prevent nonspecific protein adsorption.16 In particular, antifouling polymer brushes, grafted from surfaces via living radical polymerizations, have shown superior performance, being the most effective strategy capable of resisting adsorption even from undiluted complex biological fluids.17,18 They have found application in controlling cell adhesion even in serum-containing media19 and labelfree biosensing, allowing real-time detection in real clinical samples.20 Analogously as for superhydrophobic reactive surfaces,21 their effective functionalization remains a substantial challenge, as protein repellence and chemical reactivity are not readily combined. Conventional functionalization methods of antifouling polymer brushes impair their properties, as extensive chemical modifications are required to covalently bind receptors to a protein-repellent surface.7 As a means to tackle this challenge, hierarchical polymer brushes combining an inert antifouling segment and a reactive segment in a diblock architecture show great promise in biosensing.22,23 Owing to the modularity of this approach, the chemical functionality of the reactive polymer block can be selected according to the requirements of the application. To exploit this promising concept for precise surface patterning in the low micron- to nanoscale, we employed polymer pen lithography (PPL).24 PPL is a scanning probe lithography method combining favorable aspects of microcontact printing (µCP) and dip-pen nanolithography (DPN),25 especially retaining the large area printing capability of µCP in conjunction with high precision positioning and high lateral resolution as common in DPN. In contrast to µCP, PPL can generate patterns not predefined on the stamp. This offers a higher pattern flexibility, as the desired result is composed by repeated printing of dot features from the same stamp, enabled by the high precision and


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control over the stamp obtained from the scanning probe setup. By overlapping single dot features, bigger, yet solid features can be composed and moving the stamp along while in contact with the substrate to generate lines is also feasible.26 Additionally, feature sizes can be tuned by the applied contact pressure and varying contact time.27 PPL was previously employed in click– chemistry-based patterning using Cu(I)-catalyzed alkyne-azide cycloaddition (CuAAC),28,29 and Diels-Alder reaction.30 While CuAAC is one of the most common and robust click-chemistry approaches,31,32 it relies on the presence of Cu(I) ions, which might be undesired for potential toxicity issues having biological applications in mind. Moreover, the Cu-based catalyst may disrupt the conformation of certain biomolecules,33,34 making it a potential limitation for the application of this technique for the creation of functional protein patterns. This can be overcome by employing the strain-promoted alkyne-azide cycloaddition (SPAAC)35 reaction instead, giving the added benefit of not needing to add catalyst reagents into the PPL inks. We recently demonstrated the effectiveness of SPAAC as compared to CuAAC for surface immobilization in arrays generated by microchannel cantilever spotting (µCS).36 Herein, we describe the precise surface patterning of reactive yet antifouling polymer brushes via click–chemistry-based covalent immobilization by means of PPL. To this end, we employ hierarchically structured polymer brushes presenting a diblock architecture achieved by means of subsequent surface-initiated atom transfer radical polymerization (SI-ATRP) steps. The bottom block is comprised of an antifouling polymer (oligo(ethylene glycol) methylether methacrylate, MeOEGMA) inert to functionalization, while the top block (3-azido-2-hydroxypropyl methacrylate) displays reactive azide groups, which can participate in the SPAAC reaction. While enabling highly specific chemical functionalization, the brushes remain protein-repellent without any additional background saturation or blocking steps.


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2. MATERIALS AND METHODS 2.1. Preparation of Polymer Brushes. Firstly, a self-assembled monolayer (SAM) of ATRP initiator was attached on the surfaces. Glass and silicon wafer substrates were cleaned by rinsing with ethanol and water twice and activated in a UV/O3 cleaner for 20 min. Without delay, they were

immersed

in

a

1 µg mL–1

solution

of

(11-(2-bromo-2-methyl)propionyloxy)

undecyltrichlorosilane in anhydrous toluene and kept in a dry environment for 3 h. Subsequently, they were rinsed with toluene and acetone, twice with ethanol and water, and then dried under a stream of nitrogen. For the preparation of the bottom polymer block by SI-ATRP, 5 mL of methanol deoxygenated by bubbling Ar for 1 h were added to a previously deoxygenated roundbottom flask containing 2,2-bipyridyl (155 mg, 991 µmol), CuBr2 (16.8 mg, 75 µmol), and CuBr (53.8 mg, 375 µmol). The contents were then stirred until complete dissolution to obtain the catalyst solution. A solution of MeOEGMA (5.9 g, 19 mmol) and 5 mL of deionized water deoxygenated by bubbling Ar for 1 h was added to the flask containing the catalyst solution and the obtained polymerization solution was transferred to individual Ar-filled reactors containing the initiator-SAM-coated substrates. After 20 min of reaction at 30 oC, the polymerization was stopped by removing the substrates, rinsing them with ethanol and water and drying them under a stream of nitrogen, obtaining a dry thickness of poly(MeOEGMA) of 22 nm. The top polymer block was prepared by SI-ATRP glycidyl methacrylate (GMA) using the poly(MeOEGMA) brushes as macroinitiators, followed by nucleophilic epoxide ring opening with azide. N,Ndimethylformamide (DMF, 10 mL), 2,2’-bipyridyl (191 mg, 1222 µmol), CuBr2 (21.8 mg, 98 µmol), and GMA (6.7 mL, 49 mmol) were deoxygenated by bubbling with Ar for 1 h. Subsequently, CuBr (70.1 mg, 489 µmol) was added and the mixture was stirred until full dissolution and transferred to the reactors containing the poly(MeOEGMA)-coated substrates


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under Ar. After 11 h at 60 °C, the substrates were removed from the reactors and rinsed with DMF and dichloromethane twice and dried under a stream of nitrogen. The total dry thickness of the poly(MeOEGMA-b-GMA) brushes was 38 nm, corresponding to a poly(GMA) block thickness of 16 nm. The ring-opening of the epoxide groups was carried out by placing the substrates in a 3.4 mg mL–1 solution of NaN3 in anhydrous DMF at 60 °C for 24 h. The substrates were subsequently removed from the solution and rinsed copiously with DMF, twice with ethanol and water, and dried under a stream of nitrogen. For the quantification of nonspecific protein adsorption via surface plasmon resonance, polymer brushes were grown following the same procedure from gold-coated glass sensor chips from an initiator selfassembled monolayer of ω-mercaptoundecyl bromoisobutyrate (synthesized using a previously published protocol).37 2.2. Patterning by Polymer Pen Lithography. Ink solutions for PPL were produced by dissolving carboxytetramethylrhodamine dibenzocyclooctyne (TAMRA-DBCO) and BiotinDBCO, respectively, (both from Jena Bioscience, Germany) in dimethyl sulfoxide (DMSO), obtaining stock solutions of 1 mg mL–1. These stocks were further diluted with deionized (DI) water to a concentration of 500 µg mL–1 for use in lithography. PPL stamps were prepared as previously reported in the literature.24 First, poly(dimethylsiloxane) (PDMS) prepolymer was prepared by mixing 1 g vinylmethylsiloxane dimethylsiloxane copolymer (ABCR GmbH, Germany), 5 µL 2,4,6,8-tetramethyltetravinylcyclotetrasiloxane (Sigma Aldrich, Germany) and 2 µL platinum-divinyltetramethyldisiloxane complex in xylene, (ABCR GmbH, Germany). The mixture was stirred for 30 min and afterwards degassed in a desiccator at 0.1 bar for 15 min. The crosslinking of the prepolymer was initiated by addition of 0.3 g of the crosslinking agent methylhydrosiloxane dimethylsiloxane (ABCR GmbH, Germany). After a 30 s stirring under


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vacuum the resulting liquid polymer was deposited onto pre-fabricated silicon masters, featuring 5 × 5 mm2 arrays of pyramidal holes with a periodicity of 100 µm. Cut microscopy glass slides were uniformly pressed onto the coated masters to homogeneously fill the holes and remove all air inclusions. These sandwiches were then baked overnight at 70°C on a hotplate. Afterwards, the glass slides were separated from the silicon masters, with the hardened PDMS residing on the glass surfaces. Consequently, the cast PDMS layers now exhibit arrays of pyramidal pens for PPL. Prior ink loading the respective PDMS-glass stamp was treated with oxygen plasma at 10 sccm, 0.2 mbar and 200 W for 2 min to provide sufficient hydrophilicity of the PDMS to improve stamp wetting. Then, the PPL stamps were inked with 2 to 4 µL of the respective ink solutions by using an automatic pipette. PPL was performed with a commercial DPN setup (NLP 2000, Nanoink Inc., USA) modified with a custom-made stamp holder. First, one PPL stamp was glued to a standard microscope glass slide with a two-component epoxy resin adhesive (Uhu, Germany). Then, the whole glass slide is glued onto the custom-made holder and mounted to the instrument. To achieve parallel alignment to the substrate, the stamp was leveled following an optical alignment procedure monitoring the elastic tip deformation upon contacting a clean sacrificial silicon oxide substrate.38 Patterning was then performed at a controlled humidity of 70 to 80% relative humidity (RH) and with a dwell time of 5 to 10 s for the TAMRA-DBCO ink and 30 s for the biotin-DBCO ink. After lithography, samples were allowed to rest for 15 min to promote binding, and then washed by dipping into DMSO and subsequent rinsing with ultrapure water to remove excess ink. 2.3 Spectroscopic Ellipsometry. The dry thickness of the polymer brush layers was measured on Si wafer chips by spectroscopic ellipsometry with a J.A. Woollam M-2000X instrument in air at room temperature at angles of incidence of 60, 65, and 70° in the wavelength range


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λ = 245-1000 nm. Data sets were analyzed using multilayer models in CompleteEASE software. The optical properties of the polymer layer were fitted with a Cauchy relation model. 2.4. XPS Characterization. A K-Alpha+ spectrometer (Thermo Fisher Scientific, East Grinstead, UK) was used to perform XPS measurements. The samples were analyzed using a micro-focused, monochromated Al Kα X-ray source (400 µm spot size). The kinetic energy of the electrons was measured using a 180° hemispherical energy analyzer operated in the constant analyzer energy mode (CAE) at 50 eV pass energy for elemental spectra. Thermo Avantage software was used to analyze the spectra. The spectra were fitted with one or more Voigt profiles (binding energy uncertainty: ±0.2 eV). The analyzer transmission function, Scofield sensitivity factors,39 and effective attenuation lengths (EALs) for photoelectrons were applied for quantification. EALs were calculated using the standard TPP-2M formalism.40 All spectra were referenced to the C 1s peak of hydrocarbons at 285.0 eV binding energy controlled by means of the well-known photoelectron peaks of metallic Cu, Ag, and Au. 2.5. Protein Incubation Protocols. To characterize protein binding on the patterns and nonfunctionalized controls, 1 µL of 1 mg·mL-1 fluorescently labelled streptavidin-cy3 (SigmaAldrich, Germany) was diluted in 100 µL of phosphate buffered saline (PBS, pH 7.4) and incubated on the PPL-patterned brush samples for 10 min. Afterwards samples were rinsed with ultrapure water, dried with a nitrogen stream and inspected by fluorescence microscopy. For the control experiments with conventional blocking, 200 µL of 10 wt% BSA in PBS was incubated on the samples for 20 min prior to implementing the above described procedure for streptavidincy3 incubation.


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2.6. Fluorescence Microscopy. The fluorescence microscopy was performed with a Nikon Eclipse 80i upright fluorescence microscope equipped with an Intensilight illumination (all components from Nikon, Japan) and a CoolSNAP HQ2 camera (Photometrics, USA) using a Texas Red filter set (Nikon Y-2E/C). 2.7. Surface Plasmon Resonance. Non-specific protein adsorption was measured using an instrument based on the Kretschmann geometry of attenuated total reflection and spectral interrogation at a temperature of 25 °C. The shift of resonance wavelength was recorded and adsorption was calculated from the difference between the baselines in pure PBS (converted to mass deposited on the surface) before and after contact with the tested solutions for the specified times: streptavidin (non-conjugated, Thermo Fisher Scientific, 10 µg mL–1 in PBS, pH 7.4, 10 min), human blood plasma (Sigma-Aldrich, undiluted, 15 min), and fetal bovine serum (Sigma-Aldrich, 10% in PBS, pH 7.4, 60 min). The solutions were driven by a peristaltic pump at a constant flow rate of 25 µL min–1. The preparation of the BSA-passivated surfaces for surface plasmon resonance was performed in situ by flowing a solution of BSA (10 mg mL–1 in PBS, pH 7.4) for 20 min over the surfaces.

3. RESULTS AND DISCUSSION 3.1. Polymer Brush Synthesis and Characterization. Surfaces combining both resistance to protein fouling, especially from complex biological media, and reactive groups to undergo covalent functionalization are required for many applications in biological research and biosensing. While antifouling polymer brushes are highly resistant to non-specific protein adsorption, conventional methods of functionalization generally impair their properties as the


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functionalization procedures interfere with the brush structure and non-specific reactivity remains even after specific binding has occurred. The incorporation of an inert antifouling polymer block and a reactive top block in a hierarchical architecture was previously introduced as an effective means to limit the deterioration of the properties following functionalization.22,23 In the present work, we extended the approach to incorporate “clickable” end groups in the top polymer block in order to create an antifouling surface suitable for state-of-the-art reactive patterning methods (Figure 1). The antifouling bottom polymer block achieves the shielding of the substrate from the approach of proteins.41 Since the second block (poly(3-azido-2hydroxypropyl methacrylate)) is dense and hydrophilic, for a protein to adsorb it must first remove water molecules from the polymer layer and penetrate the brush, further restricting the conformational freedom of the already stretched polymer chains, with high enthalpic and entropic penalties.


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Figure 1. Scheme of functionalization strategy and the chemical makeup of the polymer brushes. (a) A pattern of protein binding surface features is printed to the reactive protein-repellent polymer brush surface by highly controlled repetitive contacting of a polymer pen lithography stamp onto the sample. (b) Protein can now bind specifically to the patterned spot features, while still being repelled by the background polymer brush. (c) The chemical functionalization route for the generation of protein-repellent reactive polymer brushes. (d) Scheme of the SPAAC immobilization of functional compounds onto the protein-repellent reactive polymer brushes.


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X-ray photoelectron spectroscopy (XPS) was employed to confirm the success of the different steps in the preparation of the surfaces. Initially, an ad-layer of silane ATRP-initiator was grafted on the surfaces. The presence of the initiator moiety on the surface is evidenced directly by an overlapping spin-orbit doublet in the Br 3d region of the spectrum at 70.3 eV, characteristic of C-Br (Figure S1 in the Supporting Information). In the C 1s spectrum, the initiator gave rise to a signal at 285.0 eV from C–C, C–H in the alkane backbone and C–O and O=C–O from the ester group at 286.8 eV and 289.0 eV, respectively (Figure 2a). In accordance with the chemical structure, which includes a long alkyl spacer and a single ester group, the C–C, C–H component strongly predominates over the C–O and O=C–O components. The bottom polymer block was composed of poly(oligo(ethylene glycol) methyl ether methacrylate) (poly(MeOEGMA)) and a thickness of 22 nm was employed as it was previously found to provide excellent resistance to fouling. The C 1s core-level XPS spectrum reveals a predominance of the signal at 286.5 eV arising from the C–O in the polymer side chains. The signal at 289.0 eV can be assigned to the O=C–O groups in the methacrylate backbone. Thus, the area ratio between the (C–O):(C–C, C– H) is approximately 2.4, as the oxygen-bound carbon appears all along the side chain of polymer while the aliphatic component arises only from the polymer backbone. Azide groups were selected for the top polymer block as they can participate in the SPAAC reaction. This click chemistry reaction has several advantages that make it an ideal alternative for the functionalization of surfaces for biological applications, as it proceeds rapidly in aqueous solution at room temperature without requiring any catalyst. Moreover, residual azide groups remaining after the patterning are not able to react with any biological molecules nonspecifically in subsequent steps. Adequate selection of top-block functionality plays a critical role, as previous studies have shown that fouling resistance was impaired even on hierarchical


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brushes if functionalizable groups remaining in the brush could react non-specifically with chemical groups present in the proteins.7,22 The bioorthogonality of the SPAAC reaction overcomes this obstacle, ensuring that the functionalizable brush retains its resistance to fouling. For the preparation of the azide-functional block, a 16-nm-thick poly(GMA) block was firstly grown on top of the poly(MeOEGMA) block. It should be noted that while thicknesses are assigned individually to both polymer blocks, some extent of intermixing is likely. Thus, the final properties of the surface are expected to depend on the functionality and thickness of both blocks. The C 1s XPS spectrum of the diblock copolymer confirms the increase in the intensity of the C–C, C–H and O=C–O signals (arising from the methacrylate backbone) with respect to the C–O signal arising from the side chains, which are longer for poly(MeOEGMA) than for poly(GMA). Correspondingly, the area ratio of (C–O):(C–C, C–H) markedly decreases to 1.3. The epoxide ring-opening proceeds in the presence of sodium azide to form the reactive monomer unit of the top polymer block, 3-azido-2-hydroxypropyl methacrylate. This reaction is clearly confirmed by the high-resolution XPS spectrum of the N 1s region, where the peaks at 404.5 eV and 400.9 eV are characteristic of the N atoms in the azide moieties (Figure 2b).42,43 The peak at 399.4 eV is a result of the gradual degradation of the organic azide under X-ray irradiation.44 In contrast, the poly(MeOEGMA-b-GMA) brush lacks any signals in the N 1s region before reaction with sodium azide. Importantly, for all surfaces no remnants of copper were detected in the high-resolution scans of the Cu 2p region (Figure S2 in the Supporting Information). Further confirmation of the success in the preparation of the targeted layer was provided by Fourier-Transform Infrared (FTIR) spectroscopy in the Infrared Reflection– Absorption Spectroscopy (IRRAS) mode, identifying the presence of the expected chemical groups along the whole thickness of the polymer layer (Figure S3 in the Supporting


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Information). While inorganic azides are cytotoxic, their toxicity requires binding to the heme cofactor, essential for the function of various enzymes. Azide-bearing brushes are not expected to present any cytotoxicity as the azide moieties are covalently bound to the surface-grafted polymer chains and cannot be released into the medium, thus not being able to reach the interior of the cells and exert any toxicity. Moreover, similar organic azides were used to carry out the SPAAC reaction inside living cells in vitro and also in vivo.45,46

Figure 2. (a) High-resolution XPS spectra of the C 1s region of ATRP initiator (1), poly(MeOEGMA) (2), poly(MeOEGMA-b-GMA) (3), and poly[MeOEGMA-b-(3-azido-2-


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hydroxypropyl methacrylate)] (4). (b) High-resolution XPS spectra of the N 1s region of poly(MeOEGMA-b-GMA) before (3) and after (4) functionalization with azide. Note: in both panels, the chemical species giving rise to each component in the spectra are highlighted in red.

3.2. Patterning with Polymer Pen Lithography. To demonstrate the capability to introduce localized coupling sites for proteins on the otherwise protein-repellent polymer brushes, we employed PPL (Figure 1a). To establish suitable printing parameters and confirm that the SPAAC reaction takes place as intended, fluorescent TAMRA-DBCO was printed in a grid pattern.

Figure 3. Fluorescent patterns on polymer brushes printed by PPL. (a) Overview image of a part of a fluorescent TAMRA-DBCO grid pattern. Scale bar is 300 µm. (b) and (c) show a unit cell of the pattern in (a) before and after rinsing, respectively. The insets show the fluorescence intensity plot along the line indicated by the white arrow in each image. Scale bars are 40 µm in both images.


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The grid pattern was implemented by bringing the PPL stamp into contact with the sample surface, generating the first set of dot features corresponding to the 100 µm spacing of the PPL pen array on the stamp. The other dot features within a square subpattern were then generated by repeatedly lifting, moving laterally, and lowering the stamp array until all desired features were printed. Figure 3 shows a typical outcome of such a printing procedure for optimized parameters. PPL allows for large area patterning over square centimeters with homogeneous features and parallel generation of all subpatterns, resulting in rapid printing. The presented pattern was generated with a stamp of size 5 × 5 mm² within 1 min, the overview in Figure 3a shows only a fraction (~2.4 mm²) of the overall pattern. To probe the covalent binding, the patterns were rinsed after lithography. Figure 3b and c show a unit cell before and after rinsing, respectively, and an intensity line plot along eight dot features as insets. Upon rinsing, the intensity is reduced by about 10%, indicating the removal of excess ink that was not covalently bound to the surface. Negative control experiments with TAMRA-DBCO on glass samples without polymer brush and TAMRA-Azide on the polymer brush resulted in near complete removal of the pattern upon rinsing (Figures S4 and S5, respectively, in Supporting Information), indicating that the binding chemistry works as expected. When looking at the intensity profiles, one can observe that the feature rims are of higher intensity than the center part. This is most likely caused by the (reversible) tip deformation of the PDMS stamp on contact with the surface during printing.27 The tip apex flattens and inhibits ink flow to the area covered by the deformed tip itself. In this area, only the ink present on the tip before can transfer and bind to the surface, while at the rims of the contact area additional ink can accumulate by flowing from the upper stamp to the substrate. This accumulation of fluorophore-DBCO conjugate ink on the rims leads to an increased reaction rate and thus a higher fluorescence signal.


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Figure 4. Characterization of pattern conformity and tunable feature size. (a) Example unit cell as used to determine the average feature size. Scale bar is 20 µm. (b) Feature diameter in different areas of the sample, extracted from 20 unit cell as shown in (a) from the different regions stamped with a thick ink layer (red) and a thin ink layer (blue) on the PPL stamp. (c) A pattern demonstrating the tunable feature size by applying different amounts of pressure to the PPL stamp for each feature. Scale bar is 100 µm in the main image and 10 µm in the inset. (d) Graph of the feature diameter versus the dot number as obtained from 20 subpatterns of the sample depicted in (c).


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To characterize the pattern conformity, dot feature sizes in 20 arbitrary unit cells distributed over the pattern were measured and combined into a histogram (Figure 4a and b). To understand the distributions, the inking process for the PPL stamp needs to be considered: when the ink is pipetted onto the stamp, there is a region of a thicker, less homogeneous, ink layer where the ink was amassed by the nitrogen stream when drying the stamp and a (larger) region of thin and homogeneous ink coating on the stamp where only the directly adhering ink remained. In the distributions of dot diameter, a clear dependence on the ink layer homogeneity on the stamp became visible: the average feature diameter was measured to be (7.5 ± 0.8) µm for the center and right side of the pattern that was stamped by the homogeneously coated parts of the stamp. For the outer left side, corresponding to the less homogeneous inked stamp region, a feature diameter of (15.0 ± 1.9) µm was observed. Besides the almost doubled diameter as such, also the distribution width is increased, underlining the importance for homogeneous inking of the stamp for homogeneity of the printing results. When spin coating is not feasible to achieve best ink distribution on the stamp, a sacrificial area on the substrate should be considered or alternatively one edge of the stamp is positioned to remain off substrate to print solely with the homogeneously inked part. The dots in these test grid patterns were intentionally rather large to facilitate the observation during parameter optimization. To elucidate the feasibility of smaller features, PPL printing with different applied pressures was employed.24,47 By raising the stamp by 2 µm after each consecutively generated dot feature, the resulting contact area between a given pen and the substrate diminishes due to less elastic deformation of the pen apex. A typical outcome of such a procedure is given in Figure 4c. In the eight printed features per subpattern, a dot feature diameter range from 9.1 µm down to 3.3 µm was achieved (Figure 4d). After the first printed dot (1) was created for alignment purposes, the following (2-8) were created with less


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and less contact pressure to the substrate as described above. The diameter diminishes in a linear way as expected from previous results for this kind of procedure.24,27,48 While we stopped in downscaling the features after the submicron scale was reached as our means of observation was fluorescence microscopy and thus diffraction limited, it should be noted that PPL patterning can reach well below 100 nm in resolution.24,48 3.3 Binding Experiments. In many applications in biotechnology, medicine, and biological research, it is necessary to achieve a patterned protein immobilization. To enable protein-binding feature sites, biotin-DBCO was patterned after the printing parameters had been established. Biotin shows strong and specific binding to avidin,49 a mechanism widely used in biotechnology for coupling proteins, e.g. antibodies, by tagging them with biotin. Nevertheless, the ability of the surface to prevent the non-specific adsorption of protein is critical, as it is responsible for the specificity of the binding. In this regards, antifouling polymer brushes have shown the best performance in comparison with other state-of-the-art surface modifications.18,50


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Figure 5. Protein binding on biotin patterns. (a) Fluorescence microscopy image of a biotinDBCO pattern on a polymer brush sample after incubation with fluorescently labeled streptavidin-cy3. Scale bar is 40 µm. (b) Comparison of the non-specific protein adsorption (fouling) measured via surface plasmon resonance on bare gold, BSA-passivated surfaces and diblock brush-coated surfaces from streptavidin (10 µg mL–1 in PBS, contact time 10 min), human blood plasma (undiluted, contact time 15 min) and fetal bovine serum (10% in PBS, contact time 60 min). Note: The symbol “*” means that the fouling was below the limit of detection.

After patterning and rinsing of the samples, the resulting binding site features are not visible in optical microscopy as the biotin-DBCO is non-fluorescent. Following the patterning, the surfaces were incubated with a solution of fluorescently labeled streptavidin-cy3 in PBS without any prior blocking steps. It must be noted that on a non-functionalized and unblocked bare glass surface, streptavidin-cy3 adsorbed homogeneously to cover the whole unprotected substrate, leading to a significant fluorescent signal, as was revealed in control experiments (Figure S6 in Supporting


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Information). In contrast, a clear and precisely defined pattern emerges on the polymer brushes in fluorescence after incubation, indicating selective binding of the streptavidin to the patterned biotin-bearing features, while the non-patterned background remains dark due to the retained protein repellence of the brushes (Figure 5a). The fluorescence background intensity after incubation with the streptavidin-cy3 solution of non-functionalized antifouling polymer brushes or BSA-passivated substrates without brushes are very similar, while dramatically reduced in comparison to the fluorescence signal obtained from streptavidin-cy3 adhered to bare glass surfaces (Figure S6 in Supporting Information). The labeling of streptavidin with fluorophores (3-9 molecules per protein according to supplier data sheet) could induce conformational changes leading to different tendency to adsorb on surfaces. Thus, for a precise quantitative characterization of protein adsorption, additional direct measurements using unfunctionalized streptavidin were performed by surface plasmon resonance (SPR). These studies confirmed that both BSA passivation and the grafting of azide-functional antifouling polymer brushes fully prevent adsorption of streptavidin (below the limit of detection of SPR, 0.3 ng cm–2) under the conditions employed (Figure 5b and Figure S7 in Supporting Information). In contrast, unfunctionalized streptavidin rapidly adsorbed on bare gold reaching a plateau value of 174.3 ng cm–2 under identical conditions. Thus, the use of azide-bearing antifouling diblock brushes eliminates the need to employ additional passivation procedures. This potentially simplifies the readout of array-based sensing techniques, cutting analysis times and saving extra process steps. However, non-specific protein adsorption from complex biological fluids such as culture media or bodily fluids presents a much more significant challenge to the protein repellence of a surface.50,51 For this reason, additional SPR measurements were performed to compare the


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fouling resistance of the azide-functional diblock polymer brushes with the widely used BSA passivation under conditions that approach real applications (Figure 5b and Figure S7 in the Supporting information). Fouling upon contact with undiluted human blood plasma for a short time of 15 min was examined, as this fluid is of high interest for clinical biochemical and biosensing applications. In these areas, proteins fouling can give rise to a non-specific interfering signal preventing accurate quantification of the analyte of interest.52 The polymer brushes showed a fouling of 25 ng cm–2, which represents a reduction of 92% in comparison to the fouling observed on an unprotected bare gold surface. On the other hand, the fouling from blood plasma on a surface pre-adsorbed with BSA was 2.5 times higher at 64 ng cm–2. Furthermore, mammalian cell culture is generally performed employing media supplemented with animal sera. Such experiments are of critical importance in biological research and would benefit greatly from the power of the PPL-assisted patterning in combination with effective antifouling coatings. Non-specific protein adsorption from the cell culture media would provide additional cellbinding motifs homogeneously on the surface, thus negating the binding specificity of the patterned features.19 Therefore, to assess the potential of the polymer brushes in comparison with BSA passivation the fouling from fetal bovine serum (FBS, diluted to 10% in PBS) was evaluated for a contact time of 60 min. On a non-coated bare gold surface without any passivation, FBS immediately led to a high amount of adsorbed protein (291.9 ng cm–2), reaching a plateau after only a few minutes (sensograms can be found in Figure S7 in the Supporting Information). While BSA passivation considerably slowed down the fouling process, the fouling after 1 h still reached almost 20% of the value as for bare gold. Importantly, the azide-functional diblock polymer brushes managed to further reduce the fouling from FBS by a factor of 30 times with respect to BSA passivation, preventing 99.3% of the non-specific


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adsorption observed for bare gold. The remarkable improvement in the resistance to non-specific protein fouling on micropatterned surfaces highlights the promise of these antifouling reactive polymer brushes for cell culture in serum-containing media.

4. CONCLUSION In summary, we presented a protein-repellent, yet reactive polymer brush and demonstrated its ability to dramatically reduce fouling not only from diluted solutions of single proteins, but also from complex biological media, in contrast to the widely employed BSA passivation. Importantly,

the

protein-repellent

polymer

brush

allows

highly

localized

surface

functionalization (here demonstrated by PPL) via SPAAC click-chemistry, to enable covalently bound arbitrary structured micropatterns. This facilitates the introduction of selective protein binding sites on an otherwise highly protein-repellent polymer brush surface. PPL can achieve a tunable feature size for arrays on such polymer brushes ranging from the micron readily into the low submicron scale, while the overall patterns span over square centimeter areas. The presented results have implications for a wide range of applications where non-specific protein adhesion interferes with the intended function of micropatterned surfaces. This includes screening arrays for antibodies and other proteins without requiring blocking or passivation steps, biosensors designed to operate in complex media, as well as cell culture on micropatterned surfaces, especially when using serum-enriched media. We envision the implementation of such clickable antifouling surfaces to vastly augment the power and capabilities of advanced patterning techniques such as DPN and PPL in biological applications.


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ASSOCIATED CONTENT Supporting Information. Additional XPS spectrum, data on control experiments for covalent binding, comparison of background fluorescence after protein binding on different surfaces, and SPR sensograms of protein binding. This material is available free of charge via the Internet at http://pubs.acs.org.

AUTHOR INFORMATION Corresponding Author * E-mail: [email protected] * E-mail: [email protected] Author Contributions ∇

These authors contributed equally to this work.

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT This work was partly carried out with the support of the Karlsruhe Nano Micro Facility (KNMF, www.knmf.kit.edu), a Helmholtz Research Infrastructure at Karlsruhe Institute of Technology


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(KIT, www.kit.edu). The K-Alpha+ instrument was financially supported by the Federal Ministry of Economics and Technology on the basis of a decision by the German Bundestag. SM, AH and MH acknowledge support by NSF grant #1459838. C.R.-E. acknowledges support of the Center for Chemical Polymer Technology (CPT) under the support of the EU and the federal state of North Rhine-Westphalia (Grant EFRE 30 00 883 02). A.d.l.S.P acknowledges support of the Grant Agency of the Czech Republic (GACR) under contract no. 15-09368Y.

REFERENCES (1)

Hickman, G. J.; Boocock, D. J.; Pockley, A. G.; Perry, C. C. The Importance and Clinical Relevance of Surfaces in Tissue Culture. ACS Biomater. Sci. Eng. 2016, 2, 152–164.

(2)

Wei, Q.; Haag, R. Universal Polymer Coatings and Their Representative Biomedical Applications. Mater. Horiz. 2015, 2, 567–577.

(3)

Cai, Y.; Schwartz, D. K. Influence of Protein Surface Coverage on Anomalously Strong Adsorption Sites. ACS Appl. Mater. Interfaces 2016, 8, 511–520.

(4)

Vasdekis, A. E.; Laporte, G. P. J. Enhancing Single Molecule Imaging in Optofluidics and Microfluidics. Int. J. Mol. Sci. 2011, 12, 5135–5156.

(5)

Daneshtalab, N.; Doré, J. J. E.; Smeda, J. S. Troubleshooting Tissue Specificity and Antibody Selection: Procedures in Immunohistochemical Studies. J. Pharmacol. Toxicol. Methods 2010, 61, 127–135.

(6)

de Mel, A.; Cousins, B. G.; Seifalian, A. M. Surface Modification of Biomaterials: A Quest for Blood Compatibility. Int. J. Biomater. 2012, 2012, 8.


26

Page 27 of 33

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


(7)

Vaisocherová, H.; Ševců, V.; Adam, P.; Špačková, B.; Hegnerová, K.; de los Santos Pereira, A.; Rodriguez-Emmenegger, C.; Riedel, T.; Houska, M.; Brynda, E.; et al. Functionalized Ultra-Low Fouling Carboxy- and Hydroxy-Functional Surface Platforms: Functionalization Capacity, Biorecognition Capability and Resistance to Fouling from Undiluted Biological Media. Biosens. Bioelectron. 2014, 51, 150–157.

(8)

Sivaraman, B.; Latour, R. A. Time-Dependent Conformational Changes in Adsorbed Albumin and Its Effect on Platelet Adhesion. Langmuir 2012, 28, 2745–2752.

(9)

Lussi, J. W.; Falconnet, D.; Hubbell, J. A.; Textor, M.; Csucs, G. Pattern Stability under Cell Culture Conditions--a Comparative Study of Patterning Methods Based on PLL-GPEG Background Passivation. Biomaterials 2006, 27, 2534–2541.

(10)

Nelson, C. M.; Raghavan, S.; Tan, J. L.; Chen, C. S. Degradation of Micropatterned Surfaces by Cell-Dependent and -Independent Processes. Langmuir 2003, 19, 1493–1499.

(11)

Bog, U.; Laue, T.; Grossmann, T.; Beck, T.; Wienhold, T.; Richter, B.; Hirtz, M.; Fuchs, H.; Kalt, H.; Mappes, T. On-Chip Microlasers for Biomolecular Detection via Highly Localized Deposition of a Multifunctional Phospholipid Ink. Lab Chip 2013, 13, 2701– 2707.

(12)

Bog, U.; Brinkmann, F.; Kalt, H.; Koos, C.; Mappes, T.; Hirtz, M.; Fuchs, H.; Köber, S. Large-Scale Parallel Surface Functionalization of Goblet-Type Whispering Gallery Mode Microcavity Arrays for Biosensing Applications. Small 2014, 10, 3863–3868.

(13)

Bog, U.; Brinkmann, F.; Wondimu, S. F.; Wienhold, T.; Kraemmer, S.; Koos, C.; Kalt, H.; Hirtz, M.; Fuchs, H.; Koeber, S.; et al. Densely Packed Microgoblet Laser Pairs for


27


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 28 of 33

Cross-Referenced Biomolecular Detection. Adv. Sci. 2015, 2, 1500066. (14)

Sivaraman, B.; Latour, R. A. The Adherence of Platelets to Adsorbed Albumin by Receptor-Mediated Recognition of Binding Sites Exposed by Adsorption-Induced Unfolding. Biomaterials 2010, 31, 1036–1044.

(15)

Thompson, M.; Blaszykowski, C.; Sheikh, S.; Rodriguez-Emmenegger, C.; de los Santos Pereira, A. Biological Fluid-Surface Interactions in Detection and Medical Devices; Royal Society of Chemistry: Cambridge, 2016.

(16)

Kuzmyn, A. R.; de los Santos Pereira, A.; Pop-Georgievski, O.; Bruns, M.; Brynda, E.; Rodriguez-Emmenegger, C. Exploiting End Group Functionalization for the Design of Antifouling Bioactive Brushes. Polym. Chem. 2014, 5, 4124.

(17)

Vaisocherová, H.; Brynda, E.; Homola, J. Functionalizable Low-Fouling Coatings for Label-Free Biosensing in Complex Biological Media: Advances and Applications. Anal. Bioanal. Chem. 2015, 407, 3927–3953.

(18)

Blaszykowski, C.; Sheikh, S.; Thompson, M. A Survey of State-of-the-Art Surface Chemistries to Minimize Fouling from Human and Animal Biofluids. Biomater. Sci. 2015, 3, 1335–1370.

(19)

Rodriguez-Emmenegger, C.; Preuss, C. M.; Yameen, B.; Pop-Georgievski, O.; Bachmann, M.; Mueller, J. O.; Bruns, M.; Goldmann, A. S.; Bastmeyer, M.; Barner-Kowollik, C. Controlled Cell Adhesion on Poly(dopamine) Interfaces Photopatterned with Non-Fouling Brushes. Adv. Mater. 2013, 25, 6123–6127.


28

Page 29 of 33

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


(20)

Riedel, T.; Rodriguez-Emmenegger, C.; de los Santos Pereira, A.; Bědajánková, A.; Jinoch, P.; Boltovets, P. M.; Brynda, E. Diagnosis of Epstein-Barr Virus Infection in Clinical Serum Samples by an SPR Biosensor Assay. Biosens. Bioelectron. 2014, 55, 278–284.

(21)

Li, J.; Li, L.; Du, X.; Feng, W.; Welle, A.; Trapp, O.; Grunze, M.; Hirtz, M.; Levkin, P. A. Reactive Superhydrophobic Surface and Its Photoinduced Disulfide-Ene and Thiol-Ene (Bio)functionalization. Nano Lett. 2015, 15, 675–681.

(22)

de los Santos Pereira, A.; Riedel, T.; Brynda, E.; Rodriguez-Emmenegger, C. Hierarchical Antifouling Brushes for Biosensing Applications. Sensors Actuators B Chem. 2014, 202, 1313–1321.

(23)

de los Santos Pereira, A.; Kostina, N. Y.; Bruns, M.; Rodriguez-Emmenegger, C.; BarnerKowollik, C. Phototriggered Functionalization of Hierarchically Structured Polymer Brushes. Langmuir 2015, 31, 5899–5907.

(24)

Huo, F.; Zheng, Z.; Zheng, G.; Giam, L. R.; Zhang, H.; Mirkin, C. A. Polymer Pen Lithography. Science 2008, 321, 1658–1660.

(25)

Piner, R. D.; Zhu, J.; Xu, F.; Hong, S.; Mirkin, C. A. “Dip-Pen” Nanolithography. Science 1999, 283, 661–663.

(26)

Brinkmann, F.; Hirtz, M.; Greiner, A. M.; Weschenfelder, M.; Waterkotte, B.; Bastmeyer, M.; Fuchs, H. Interdigitated Multicolored Bioink Micropatterns by Multiplexed Polymer Pen Lithography. Small 2013, 9, 3266–3275.


29


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

(27)

Page 30 of 33

Liao, X.; Braunschweig, A. B.; Zheng, Z.; Mirkin, C. A. Force- and Time-Dependent Feature Size and Shape Control in Molecular Printing via Polymer-Pen Lithography. Small 2010, 6, 1082–1086.

(28)

Bian, S.; He, J.; Schesing, K. B.; Braunschweig, A. B. Polymer Pen Lithography (PPL)Induced Site-Specific Click Chemistry for the Formation of Functional Glycan Arrays. Small 2012, 8, 2000–2005.

(29)

Kumar, R.; Bonicelli, A.; Sekula-Neuner, S.; Cato, A. C. B.; Hirtz, M.; Fuchs, H. ClickChemistry Based Allergen Arrays Generated by Polymer Pen Lithography for Mast Cell Activation Studies. Small 2016, 12, 5330–5338.

(30)

Bian, S.; Scott, A. M.; Cao, Y.; Liang, Y.; Osuna, S.; Houk, K. N.; Braunschweig, A. B. Covalently Patterned Graphene Surfaces by a Force-Accelerated Diels-Alder Reaction. J. Am. Chem. Soc. 2013, 135, 9240–9243.

(31)

Kolb, H. C.; Finn, M. G.; Sharpless, K. B. Click Chemistry: Diverse Chemical Function from a Few Good Reactions. Angew. Chemie Int. Ed. 2001, 40, 2004–2021.

(32)

Rostovtsev, V. V.; Green, L. G.; Fokin, V. V.; Sharpless, K. B. A Stepwise Huisgen Cycloaddition Process: copper(I)-Catalyzed Regioselective “ligation” of Azides and Terminal Alkynes. Angew. Chemie Int. Ed. 2002, 41, 2596–2599.

(33)

Gierlich, J.; Burley, G. a.; Gramlich, P. M. E.; Hammond, D. M.; Carell, T. Click Chemistry as a Reliable Method for the High-Density Functionalisation of AlkyneModified Oligodeoxyribonucleotides. Org. Lett. 2006, 8, 3639–3642.


30

Page 31 of 33

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


(34)

Wang, Q.; Chan, T. R.; Hilgraf, R.; Fokin, V. V.; Sharpless, K. B.; Finn, M. G. Bioconjugation by copper(I)-Catalyzed Azide-Alkyne [3 + 2] Cycloaddition. J. Am. Chem. Soc. 2003, 125, 3192–3193.

(35)

Sletten, E. M.; Bertozzi, C. R. Bioorthogonal Chemistry: Fishing for Selectivity in a Sea of Functionality. Angew. Chemie - Int. Ed. 2009, 48, 6974–6998.

(36)

Davydova, M.; de los Santos Pereira, A.; Bruns, M.; Kromka, A.; Ukraintsev, E.; Hirtz, M.; Rodriguez-Emmenegger, C. Catalyst-Free Site-Specific Surface Modifications of Nanocrystalline Diamond Films via Microchannel Cantilever Spotting. RSC Adv. 2016, 6, 57820–57827.

(37)

Jones, D. M.; Brown, A. A.; Huck, W. T. S. Surface-Initiated Polymerizations in Aqueous Media: Effect of Initiator Density. Langmuir 2002, 18, 1265–1269.

(38)

Eichelsdoerfer, D. J.; Liao, X.; Cabezas, M. D.; Morris, W.; Radha, B.; Brown, K. A.; Giam, L. R.; Braunschweig, A. B.; Mirkin, C. A. Large-Area Molecular Patterning with Polymer Pen Lithography. Nat. Protoc. 2013, 8, 2548–2560.

(39)

Scofield, J. H. Hartree-Slater Subshell Photoionization Cross-Sections at 1254 and 1487 eV. J. Electron Spectros. Relat. Phenomena 1976, 8, 129–137.

(40)

Tanuma, S.; Powell, C. J.; Penn, D. R. Calculations of Electron Inelastic Mean Free Paths. V. Data for 14 Organic Compounds over the 50-2000 eV Range. Surf. Interface Anal. 1994, 21, 165–176.

(41)

Szleifer, I. Polymers and Proteins: Interactions at Interfaces. Curr. Opin. Solid State


31


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 32 of 33

Mater. Sci. 1997, 2, 337–344. (42)

Eisenblaetter, J.; Bruns, M.; Fehrenbacher, U.; Barner, L.; Barner-Kowollik, C. Synthesis of Polymers with Phosphorus Containing Side Chains via Modular Conjugation. Polym. Chem. 2013, 4, 2406.

(43)

Collman, J. P.; Devaraj, N. K.; Eberspacher, T. P. A.; Chidsey, C. E. D. Mixed AzideTerminated Monolayers: A Platform for Modifying Electrode Surfaces. Langmuir 2006, 22, 2457–2464.

(44)

Zorn, G.; Liu, L.; Árnadóttir, L.; Wang, H.; Gamble, L. J.; Castner, D. G.; Yan, M. X-Ray Photoelectron Spectroscopy Investigation of the Nitrogen Species in Photoactive Perfluorophenylazide-Modified Surfaces. J. Phys. Chem. C 2014, 118, 376–383.

(45)

Poloukhtine, A. A.; Mbua, N. E.; Wolfert, M. A.; Boons, G. J.; Popik, V. V. Selective Labeling of Living Cells by a Photo-Triggered Click Reaction. J. Am. Chem. Soc. 2009, 131, 15769–15776.

(46)

Chang, P. V; Prescher, J. A.; Sletten, E. M.; Baskin, J. M.; Miller, I. A.; Agard, N. J.; Lo, A.; Bertozzi, C. R. Copper-Free Click Chemistry in Living Animals. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 1821–1826.

(47)

Urtizberea, A.; Hirtz, M.; Fuchs, H. Ink Transport Modelling in Dip-Pen Nanolithography and Polymer Pen Lithography. Nanofabrication 2015, 2, 43–53.

(48)

Zheng, Z.; Daniel, W. L.; Giam, L. R.; Huo, F.; Senesi, A. J.; Zheng, G.; Mirkin, C. A. Multiplexed Protein Arrays Enabled by Polymer Pen Lithography: Addressing the Inking


32

Page 33 of 33

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


Challenge. Angew. Chemie Int. Ed. 2009, 48, 7626–7629. (49)

Green, N. M. Avidin. Adv. Protein Chem. 1975, 29, 85–133.

(50)

Rodriguez-Emmenegger, C.; Houska, M.; Alles, A. B.; Brynda, E. Surfaces Resistant to Fouling from Biological Fluids: Towards Bioactive Surfaces for Real Applications. Macromol. Biosci. 2012, 12, 1413–1422.

(51)

Rodriguez Emmenegger, C.; Brynda, E.; Riedel, T.; Sedlakova, Z.; Houska, M.; Alles, A. B. Interaction of Blood Plasma with Antifouling Surfaces. Langmuir 2009, 25, 6328– 6333.

(52)

Thompson, M.; Sheikh, S.; Blaszykowski, C.; Romaschin, A. CHAPTER 1: Biosensor Technology and the Clinical Biochemistry Laboratory - Issue of Signal Interference from the Biological Matrix. In Detection Challenges in Clinical Diagnostics; 2013; pp. 1–34.

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