Fluorescent Biotin Analogues for Microstructure Patterning and

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Fluorescent Biotin Analogues for Microstructure Patterning and Selective Protein Immobilization K. Vijaya Krishna,† Subhadip Ghosh,§ Bikramjit Sharma,† Leeju Singh,‡ Saptarshi Mukherjee,*,§ and Sandeep Verma*,†,‡ †

Department of Chemistry and ‡DST Unit of Excellence on Soft Nanofabrication, Center for Environment Science and Engineering, Indian Institute of Technology-Kanpur, Kanpur-208016 Uttar Pradesh, India § Department of Chemistry, IISER-Bhopal, Bhopal-462066 Madhya Pradesh, India S Supporting Information *

ABSTRACT: Benzyl substitution on ureido nitrogens of biotin led to manifestation of aggregation-induced emission, which was studied by steady-state fluorescence, microscopy, and TD-DFT, providing a rationale into the observed photophysical behavior. Besides exhibiting solvatochromism, the biotin derivatives revealed emission peaks centered at ∼430 and 545 nm, which has been attributed to the π−π stacking interactions. Our TD-DFT results also correlate the spectroscopic data and quantify the nature of transitions involved. The isothermal titration calorimetry data substantiates that the binding of the biotin derivatives with avidin are pretty strong. These derivatives on lithographic patterning present a platform for site specific strept(avidin) immobilization, thus opening avenues for potential applications exploiting these interactions. The fluorescent biotin derivatives can thus find applications in cellular biology and imaging.

1. INTRODUCTION Molecular fluorescence greatly aids in exploring and unraveling structural and functional properties of biological systems, with excellent sensitivity.1 Certain biomolecules exhibit intrinsic fluorescence due to natural fluorophores,2 or conjugation of synthetic fluorophores is used to gain desired photophysical properties. Occasionally, fluorescence-imparting prosthetic groups emerge due to intramolecular reactions in vivo: imidazolidinone chromophore in green fluorescent protein is an excellent example of a gene-expressible natural fluorescent label.3 Chemical modification of biomolecules is an effective synthetic strategy, as a number of activated commercial fluorescent labels are available for bioconjugation.4 A recent review by Hamachi and co-workers has dealt with protein modification strategies for elucidation and tracking of protein function in vitro and in live-cell systems.5 These strategies ensure site selectivity, optimal emission profiles and prevention of signal loss during biochemical reactions. Therefore, discovery of robust fluorophores with enhanced photostability, low cytotoxicity, high quantum yields, increased fluorescence lifetimes and tunable optical fluorescence, are important for biophysical studies and chemical biology.6 High affinity biotin−avidin interaction is well explored due to its femtomolar dissociation constant, despite variations in pH, temperature, organic solvents and denaturing agents.7 Therefore, this noncovalent interaction is considered a powerful tool in affinity chromatography, affinity cytochemistry, and immunoassays.8 However, strength of this complexation also © 2015 American Chemical Society

poses problems as it is often difficult to separate pure molecules during postpurification. Thus, novel biotin analogues with compromised binding affinity are expected to serve as effective ligands for affinity purifications. Design of different biotin derivatives with lower binding affinities necessitates identification of critical sites for biotin−avidin interaction. In this context, the importance of hydrogen bonding and van der Waals contacts, through four tryptophan residues residing in the binding pocket, among other possible interactions, is reported.9

2. EXPERIMENTAL METHODS 2.1. Optical Microscopy (OM) and Fluorescence Microscopy (FM). For all the three microscopic experiments, a 10 μL aliquot of ureido substituted biotins (10 mM in MeOH) was placed on a glass slide at room temperature and allowed to dry by slow evaporation. For OM, micrographs were recorded by (Leica DM2500M) microscope. For FM, micrographs were recorded by (Leica DM 5000 B) microscope using a DAPI filter (UV excitation) with a band-pass of 356−425 nm. 2.2. Field Emission Scanning Electron Microscopy (FE SEM). A 10 μL aliquot of ureido-substituted biotins (10 mM in MeOH) was placed on a glass slide at room temperature and allowed to dry by slow evaporation and subsequently coated with gold for 45 s. FE SEM images were acquired on FEI QUANTA 200 microscope, equipped with a tungsten filament gun, operating at WD 10.6 mm and 20 kV. Received: September 16, 2015 Revised: November 5, 2015 Published: November 11, 2015 12573

DOI: 10.1021/acs.langmuir.5b03476 Langmuir 2015, 31, 12573−12578

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Langmuir 2.3. Atomic Force Microscopy (AFM). AFM samples were imaged with an atomic force microscope (Molecular Imaging, USA) operating under the Acoustic AC mode (AAC), with the aid of a cantilever (NSC 12(c) from MikroMasch). The force constant was 0.6 N/m, while the resonant frequency was 150 kHz. The images were taken at room temperaure, with the scan speed of 1.5−2.2 lines/sec. The data acquisition was done using PicoView 1.8.2 software, while the data analysis was done using PicoView. A 10 μL aliquot of ureidosubstituted biotins (10 mM in MeOH) was placed on a desired surface at room temperature and allowed to dry by slow evaporation. The sample coated surface was further dried in vacuum before imaging with AFM. 2.4. Isothermal Titration Calorimetry Analysis (ITC). Isothermal titration calorimetry experiments were performed with a Nano ITC, TA Instruments. All the solutions used for ITC analysis were degassed before the experiments. The sample cell was loaded with avidin solution and was continuously stirred at 300 rpm by the injection syringe containing the biotin derivatives solutions (having 10−20 times greater concentration than avidin concentration). Titrations were carried out for 25 injections over a period of 75 min, and the temperature was maintained at 25 °C. Both reactants were dissolved in the same solution containing 100 mM sodium phosphate buffer (pH 7.4). The respective control experiments were carried out by injecting the titrant into the buffer solution in the absence of the avidin to correct for the heat change due to mixing and dilution. The data were analyzed by NanoAnalyze software v2.4.1 and fitted with an independent site binding model. 2.5. UV Spectrophotometer. Absorbance spectra were recorded on Varian Cary 100 BIO with 10 mm quartz cell at 25 ± 0.1 °C. 2.6. Fluorescence Spectrophotometer. Steady-state fluorescence measurements were recorded on a Horiba Jobin Yvon Fluorolog 3-111. The fluorescence spectra were measured with a 10 mm path length quartz cuvette. The emission and the excitation slits were kept at 5 and 2 nm, respectively. 2.7. Nanoimprint Lithography (NIL). Nanoimprint Lithography (NIL) system from Obducat equipped with full area thermal imprint, using the patented Soft Press technology was used. The system was capable of uniform heating over a wide range of temperature and pressure settings, makes imprinting possible over a wide range of polymers. 2.8. Sample Preparation for NIL. Substrates used for NIL (silicone, glass and PET sheet) were washed with acetone and methanol and dried under an air stream to remove any dust particles. A 1 mg/100 μL solution of 1 (Figure 1) was spin coated on these substrates at 900 rpm for 90 s at room temperature and used for NIL.

hydrogen bonding pattern. This modification was expected to alter its binding affinity and possibly exert effect on solution properties and morphology, through aggregative mechanism (Figure 1). Binding affinities of 1 (4.88 × 105 M−1) and 2 (1.3 × 106 M−1) for avidin were determined by ITC analysis (Figure S1, SI). 1 and 2 (10−4 M in methanol) exhibited λabs max at 209 nm and a hump at 265 nm, corresponding to ureido and benzylic groups, respectively (Figure 2a). When explored further, we were surprised to observe fluorescence in 1 and 2 (having quantum yield of 2.9% and 2.1% for 1 and 2 respectively, taking 8anilino-1-napthaenesulfonic acid as a reference), when their solutions were illuminated by a hand-held UV lamp (inset Figure 2a). Thus, it was clear that benzylation confers fluorescence with emission maxima at 310 nm (Figure 2a, blue and magenta) with corresponding excitation peaks at 210 nm and ∼265 nm, for both 1 and 2 (Figure 2a, green and red). In the excitation spectra, the higher intensity of peak at 265 nm indicates that the fluorescence of these derivatives arises due to the excited benzene moiety rather than the excited ureido groups. A prominent shoulder at around 355 nm, in addition to the base peak (λex = 260 nm), for 2 (Figure 2a, magenta) can be attributed to an additional benzyl group. Although sparingly soluble in water, 1 and 2 were readily soluble in ethanol and methanol. A red shift for the emission peaks was observed with increasing solvent polarity (Figure 2b) and importantly, their fluorescence was conserved in solvents of varying polarities. Emission profiles of 1 in 1,4-dioxane, a nonpolar aprotic solvent and in DMF, a polar aprotic solvent also revealed a solvent dependent bathochromic shift with increasing polarity (Figure S2, SI). This trend may suggest a possible intramolecular charge transfer (ICT) from the carbonyl group to the π* orbitals of the benzyl group for these derivatives. Excitation of 1 and 2 at 350 nm (corresponding to the wavelength of the UV lamp), revealed emission peaks centered at ∼430 and 545 nm, presenting a clear signature of bluish emission (Figure 2c). Samples were screened over different excitation wavelengths to establish that observed emission was independent of excitation wavelength and appear exclusively from samples of biotin derivatives, and to rule out genesis of these peaks due to higher order scattering (Figure S3, SI). A distinct peak at ∼545 nm corresponding to a green emission was also observed (Figure 2c), which is perhaps masked by stronger blue fluorescence. A time-dependent fluorescence spectra of 1 and 2 revealed that fluorescence intensities increased gradually over a week without any shift in emission wavelength, suggesting a favorable hydrophobic microenvironment, which is generated due to π−π stacking of benzylic groups present in the molecule10 (Figure 2d,e). These observations collectively suggest that fluorescence in 1 and 2 emerges perhaps due to favorable hydrophobic and stacking interactions of the benzyl ring, leading to a situation similar to aggregation-induced emission (AIE).11 The latter is a photophysical phenomenon that occurs as a direct consequence of molecular aggregation, followed by restriction of intramolecular rotations in molecules having aggregative propensity. It redirects the nonluminescent decay process of excited states through a radiative pathway.12 In a seminal work, Zhao and coworkers have reported the AIE observed in triphenyl benzene derivatives and obtained red-shifted emission peaks, which have been attributed to the excimer formation and cofacial stacking of the cores.11 A number of systems involving hydrocarbons,13 heteroatoms,14 organometallics,15 are known to exhibit AIE

Figure 1. Chemical modifications at the ureido group of biotin: N′1,N′3-bis benzyl biotin, 1. N′1,N′3-bis benzyl biotin benzyl ester, 2. 2.9. Synthesis of Ureido-Substituted Biotin Conjugates. Monomethylation and benzylation were carried out by nucleophilic substitution reactions with methyl iodide and benzyl bromide. A strong base like NaH was required for proton abstraction from the N′ nitrogen of the ureido group. However, for bismethylation, strong reflux conditions in aq. HCHO and aq. HCOOH were employed followed by methylation of free acid group (Schemes S1 and S2, Supporting Information (SI)). All reactions were carried out under inert atmosphere in dry solvents, and products were characterized by NMR and HRMS analysis.

3. RESULTS AND DISCUSSION We decided to substitute the −NH hydrogens of the ureido group with benzyl groups, to disrupt donor−acceptor−donor 12574

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Figure 2. (a) Spectral profiles of substituted biotin derivatives (10−4 M in methanol): Absorption spectra: 1 (orange), 2 (black); excitation spectra: 1 (green), 2 (red); emission spectra: 1 (blue), 2 (magenta). Inset: Solutions of 1 and 2 (10 mM in MeOH) illuminated by a UV light at 365 nm. (b) Solvent dependent emission profiles of 1 (λex at 209 nm). (c) Emission spectra of 1 and 2 (λex at 350 nm). Time-dependent fluorescence spectra showing changes in emission intensity over 7 days: (d) 1; (e) 2.

Figure 3. DFT-optimized geometries: (a) 1 and 2 using B3LYP/6-31G(d,p) (hydrogens are not shown for clarity); (b) Molecular orbital diagrams of 1; (c) Molecular orbital diagrams of 2.

effect. We propose that the molecular structure of benzylsubstituted chiral biotin derivatives, where freely rotatable aromatic benzylic groups are substituted in the ureido group, makes it a strong candidate for AIE which results in the redshifted emission peaks centered ∼430 and 545 nm. Thus, in the present case, the π−π stacking of benzylic groups most probably results in the AIE, which has also been substantiated by our AFM studies (Figure S4g,h, SI). We decided to apply time-dependent density functional theory (TD-DFT) to gain insight into spectral properties of 1 and 2. More recently, it has also been shown as a powerful tool in determining electronic excited states (EES) and to interrogate photoabsorption features.16,17 Hence, DFT-aided geometry optimizations were carried out for 1 and 2, by using B3LYP exchange-correlation functional for these calculations,18,19 with 6-31G(d,p) as the basis set. The normal-mode analysis of vibrational frequencies was performed, and the absence of negative frequencies confirmed the structures to be at minima (at least local) (Figure 3a; Table S1, SI). Optimized structures of 1 and 2 were used for TD-DFT studies (B3LYP/6-31G(d,p)) to calculate vertical excitation energies, which characterize their photophysical properties.

Results show that maximum oscillator strength (f) of 0.1335 was obtained for 1 at 219.61 nm, which is close to its experimental absorption peak at 209 nm. This peak corresponds to S0−S1 transition with maximum contribution from HOMO−6 to LUMO transition (∼61%), with additional contributions coming from HOMO−5 to LUMO (∼21.5%) and HOMO−7 to LUMO+1 (∼9.5%) transitions. In HOMO− 6 (Figure 3b), the electron density is located on the carbonyl group of the ureido moiety and on one of the benzyl groups, while the electron density resides only over the benzyl group in LUMO (Figure 3b). The disappearance of electron density from the ureido group and increased electron density over the benzyl group in LUMO suggests that absorption occurs perhaps due to intramolecular charge transfer (ICT) from the carbonyl group to π* orbitals of the benzyl group. Further, absorption at 266.09 nm was also observed with f = 0.0124, with maximum contribution from HOMO−2 to LUMO (∼61%) (Figure 3b). The oscillator strength, which is an indicator of the intensity of a transition, is markedly reduced, suggesting that these calculations match well with experimental observations. Similarly, TD-DFT results of 2 show an absorption peak at 219.65 nm (f = 0.1416) with maximum 12575

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Figure 4. Schematic illustrates the self-assembly and subsequent AIE observed for benzyl biotins owning to CH···π interaction as observed by DFT studies (scale bar corresponds to 10 μm).

Figure 5. Micropatterning of 1 by NIL using a soft PDMS stamp on a silicon substrate as visualized by fluorescence microscope: (a) blue fluorescence of patterned biotin derivative under DAPI filter, (b) red emission due to immobilized Cy3-Streptavidin under TRITC filter, and (c) composite images capturing both the emissions as prepared from ImageJ 1.48 V.

bilayers on graphene sheets,28 for cell alignment on nanoparticle arrays29 and for selective reorientation of prion protein,30 to name a few. With this background, we devised a NIL study with 1 owing to its solubility features and possibility of further functionalization. A polydimethylsiloxane (PDMS) stamp was used to imprint patterns on substrates spin coated with uniform layer of 1, and the patterning was optimized over a range of pressure, temperature, and concentration gradients. Ordered patterns of the stamp could be readily translated onto substrates coated with 1 (1 mg/100 μL) at room temperature and low pressure conditions (Figure S6, Table S2, SI). Intrinsic fluorescence of 1 was conserved on NIL patterning, where different morphological features of the stamp were successfully transferred on coated substrate as revealed by SEM and nanoprofiler analysis (Figure S7, SI). Having determined the binding affinity of 1 for avidin by ITC analysis, we then decided to probe surface selective decoration via high affinity 1strept(avidin) complexation. We chose Cy3-streptavidin, a dye with strong emission in red region, for visualization, without any interference emission from 1. The patterned substrates on Cy3-streptavidin immobilization revealed blue and red fluorescence, respectively, when viewed under DAPI and TRITC filter, without any visible distortion in patterns despite exposure to protein solution (Figure 5; Figure S8, SI).

contribution coming from HOMO−8 to LUMO transition (∼62%) (Figure 3c). This peak can be attributed to ICT from carbonyl group of the ureido moiety to the benzyl group of 2 in LUMO (Figure 3c). Further, existence of a peak at 265.63 nm, having f = 0.0122, appears with maximum contribution from HOMO−2 to LUMO transition (Figure 3c). DFT studies were then carried out for two molecules of 1 and also of 2 to probe into the nature of mutual interactions of its benzyl groups. Geometric optimizations were carried out using B97D functional with TZVP basis set. The optimized structures revealed the presence of intermolecular CH···π interaction (2.84 Å), via hydrogen from the benzyl ring of one molecule pointing toward the centroid of the benzyl ring of another molecule, within acceptable interacting distance with a C−H− X angle of 145.26°, where X is the centroid of the phenyl ring (Figure S5, SI).20 Given the possibility of aggregation in these systems and our prior interest in solution-phase self-assembly,21 we were also curious to determine the possible morphology of these derivatives in solution-phase. Studies with 1 and 2 revealed formation of polydispersed, well-defined spherical aggregates (Figure 4 and Figure S4, SI). Thus, we decided to assess the possibility of using 1 for microscale surface patterning using nanoimprint lithography (NIL), which allows patterning by using a soft stamp forged on a master.22 Lithography coupled with biomolecular interactions, such as Au-thiol,23 biotin−streptavidin,24 and His-Tag-Ni,25 when implemented with AFM and fluorescence microscopy, offer a great tool in nanofabrication to create preconceived designs. This technique has been used for generating artificial neuronal assemblies,26 for fluorescent detection of DNA on a photonic crystal platform,27 for creating functionalized lipid

4. OUTLOOK In conclusion, interesting molecular properties of ureidomodified biotin derivatives brought forth by tuning their noncovalent interactions, leading to self-assembled aggregates, is demonstrated. Stable interacting units of these ensembles leading to aggregation-induced emission and ensuing fluo12576

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(9) Freitag, S.; Chu, V.; Penzotti, J. E.; Klumb, L. A.; To, R.; Hyre, D.; Le Trong, I.; Lybrand, T. P.; Stenkamp, R. E.; Stayton, P. A Structural Snapshot of an Intermediate on the Streptavidin-Biotin Dissociation Pathway. Proc. Natl. Acad. Sci. U. S. A. 1999, 96, 8384− 8389. (10) Anand, U.; Mukherjee, M. Exploring the Self-Assembly of a Short Aromatic αβ(16−24) Peptide. Langmuir 2013, 29, 2713−2721. (11) Bao, C.; Lu, R.; Jin, M.; Xue, P.; Tan, C.; Xu, T.; Liu, G.; Zhao, Y. Helical Stacking Tuned by Alkoxy Side Chains in π-Conjugated Triphenylbenzene Discotic Derivatives. Chem. - Eur. J. 2006, 12, 3287−3294. (12) Hong, Y.; Lam, J. W. Y.; Tang, B. Z. Aggregation-Induced Emission. Chem. Soc. Rev. 2011, 40, 5361−5388. (13) Zhao, Q.; Li, K.; Chen, S.; Qin, A.; Ding, D.; Zhang, S.; Liu, Y.; Liu, B.; Sun, J. Z.; Tang, B. Z. Aggregation-Induced Red-NIR Emission Organic Nanoparticles as Effective and Photostable Fluorescent Probes for Bioimaging. J. Mater. Chem. 2012, 22, 15128−15135. (14) Yang, Z.; Qin, W.; Lam, J. W. Y.; Chen, S.; Sung, H. H. Y.; Williams, I. D.; Tang, B. Z. Fluorescent pH Sensor Constructed from a Heteroatom-Containing Luminogen with Tunable AIE and ICT Characteristics. Chem. Sci. 2013, 4, 3725−3730. (15) Jia, X.; Yang, X.; Li, J.; Li, D.; Wang, E. Stable Cu Nanoclusters: From an Aggregation-Induced Emission Mechanism to Biosensing and Catalytic Applications. Chem. Commun. 2014, 50, 237−239. (16) Pieslinger, G. E.; Albores, P.; Slep, L. D.; Baraldo, L. M. Class III Delocalization in a Cyanide-Bridged Trimetallic Mixed-Valence Complex. Angew. Chem., Int. Ed. 2014, 53, 1293−1296. (17) Vlasov, I. I.; Shiryaev, A. A.; Rendler, T.; Steinert, S.; Lee, S.-Y.; Antonov, D.; Vörös, M.; Jelezko, F.; Fisenko, A. V.; Semjonova, L. F.; Biskupek, J.; Kaiser, U.; Lebedev, O. I.; Sildos, I.; Hemmer, R. P.; Konov, V. I.; Gali, A.; Wrachtrup, J. Molecular-Sized Fluorescent Nanodiamonds. Nat. Nanotechnol. 2014, 9, 54−58. (18) Stephens, P. J.; Devlin, F. J.; Chabalowski, C. F.; Frisch, M. J. Ab Initio Calculation of Vibrational Absorption and Circular Dichroism Spectra Using Density Functional Force Fields. J. Phys. Chem. 1994, 98, 11623−11627. (19) Grimme, S. Do Special Noncovalent π-π Stacking Interactions Really Exist? Angew. Chem., Int. Ed. 2008, 47, 3430−3434. (20) Nishio, M. The CH/π Hydrogen Bond in Chemistry. Conformation, Supramolecules, Optical Resolution and Interactions Involving Carbohydrates. Phys. Chem. Chem. Phys. 2011, 13, 13873− 13900. (21) Joshi, K. B.; Verma, S. Ditryptophan Conjugation Triggers Conversion of Biotin Fibers into Soft Spherical Structures. Angew. Chem., Int. Ed. 2008, 47, 2860−2863. (22) Mujahid, A.; Iqbal, N.; Afzal, A. Bioimprinting Strategies: From Soft Lithography to Biomimetic Sensors and Beyond. Biotechnol. Adv. 2013, 31, 1435−1447. (23) Chai, J.; Wong, L. S.; Giam, L.; Mirkin, C. A. Single-Molecule Protein Arrays Enabled by Scanning Probe Block Copolymer Lithography. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 19521−19525. (24) Palankar, R.; Medvedev, N.; Rong, A.; Delcea, M. Fabrication of Quantum Dot Microarrays Using Electron Beam Lithography for Applications in Analyte Sensing and Cellular Dynamics. ACS Nano 2013, 7, 4617−4628. (25) Wu, C.-C.; Reinhoudt, D. N.; Otto, C.; Velders, A. H.; Subramaniam, V. Protein Immobilization on Ni(II) Ion Patterns Prepared by Microcontact Printing and Dip-Pen Nanolithography. ACS Nano 2010, 4, 1083−1091. (26) Staii, C.; Viesselmann, C.; Ballweg, J.; Shi, L.; Liu, G.-Y.; Williams, J. C.; Dent, E. W.; Coppersmith, S. N.; Eriksson, M. A. Positioning and Guidance of Neurons on Gold Surfaces by Directed Assembly of Proteins Using Atomic Force Microscopy. Biomaterials 2009, 30, 3397−3404. (27) Endo, T.; Ueda, C.; Kajita, H.; Okuda, N.; Tanaka, S.; Hisamoto, H. Enhancement of the Fluorescence Intensity of DNA Intercalators Using Nano-Imprinted 2-Dimensional Photonic Crystal. Microchim. Acta 2013, 180, 929−934.

rescence properties of benzyl biotins were also studied in detail with the help of TD-DFT calculations. Their inherent fluorescence emission and propensity to form soft spherical structures was combined for micropatterning using NIL and demonstrated to immobilize Cy3-Streptavidin in a site specific manner by exploiting Biotin-strept(avidin) interactions. As a future endeavor we are exploring the possibility of enhancing water solubility and tuning fluorescence emission of these derivatives, and we believe that the results presented here could provide a new direction toward generation of biomoleculedecorated monolayers and their label-free fluorescence detection.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir.5b03476. Several spectroscopic analyses including 1H and 13C NMR, HR-mass data, ITC, UV−vis, steady-state fluorescence, FE-SEM, AFM, OM, FM, NIL data, and DFT analysis of the biotin derivates (PDF)

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. *Email: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS S.V. acknowledges MHRD supported Centre for Excellence in Chemical Biology, DST for a J C Bose Fellowship, and DAESRC for an Outstanding Investigator Award. S.M. acknowledges IISER Bhopal and the DST-Fast track scheme from SERB, for financial support. DFT calculations were performed using computer cluster installed at the Department of Chemistry, IIT Kanpur through DST-FIST program. We also thank Prof. Amalendu Chandra, IIT Kanpur, for his input on DFT calculations.

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REFERENCES

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Langmuir (28) Hirtz, M.; Oikonomou, A.; Georgiou, T.; Fuchs, H.; Vijayaraghavan, A. Multiplexed Biomimetic Lipid Membranes on Graphene by Dip-Pen Nanolithography. Nat. Commun. 2013, 4, 2591. (29) Subramani, C.; Saha, K.; Creran, B.; Bajaj, A.; Moyano, D. F.; Wang, H.; Rotello, V. M. Cell Alignment using Patterned Biocompatible Gold Nanoparticle Templates. Small 2012, 8, 1209− 1213. (30) Sanavio, B.; Scaini, D.; Grunwald, C.; Legname, G.; Scoles, G.; Casalis, L. Oriented Immobilization of Prion Protein Demonstrated via Precise Interfacial Nanostructure Measurements. ACS Nano 2010, 4, 6607−6616.

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DOI: 10.1021/acs.langmuir.5b03476 Langmuir 2015, 31, 12573−12578