Polymerizable Biomimetic Vesicles with Controlled ... - ACS Publications

Mar 18, 2010 - (16) Zhao, H.; Robertson, N. B.; Jewhurst, S. A.; Waite, J. H. J. Biol. .... (39) Statz, A. R.; Meagher, R. J.; Barron, A. E.; Messersm...
0 downloads 0 Views 3MB Size
Download PDF
pubs.acs.org/Langmuir © 2010 American Chemical Society

Polymerizable Biomimetic Vesicles with Controlled Local Presentation of Adhesive Functional DOPA Groups Pieter Samyn,‡ J€urgen R€uhe,‡ and Markus Biesalski*,†,‡ †

Technical University of Darmstadt, Department of Chemistry Macromolecular Chemistry and Paper Chemistry, Petersenstrasse 22, D-64278 Darmstadt, Germany, and ‡University of Freiburg, Department for Microsystems Engineering (IMTEK), Chemistry and Physics of Interfaces, Georges-K€ ohler-Allee 103, D-79110 Freiburg, Germany Received December 9, 2009. Revised Manuscript Received February 7, 2010 Inspired by strong adhesive properties of mussel footprint proteins, which are largely governed by the presence of dihydroxy-phenylalanine (DOPA) amino acid moieties, we present a novel approach for presenting DOPA groups in a very defined way in order to modulate the adhesion between artificial interfaces. To this end, linear peptide amphiphiles are synthesized with attached DOPA functional groups and a polymerizable diacetylenic tail. The obtained amphiphiles can be coassembled with matrix amphiphiles into vesicles, which can be subsequently stabilized through UV-lightinduced solid-state polymerization. Depending on the molar ratio of matrix and adhesive amphiphiles, the vesicles selfassemble into spherical, fibrilar, or planar nanostructures. The adhesive properties of the surface-adsorbed vesicles are evaluated by drop casting them onto a planar solid substrate and performing macroscopic shear tests in contact with a similar substrate. The shear forces are investigated as a function of substrate chemistry, vesicle polymerization conditions, vesicle concentration, and number of adhesive DOPA groups in the interface. Substrate adhesion is enhanced by surfaceconfined vesicles and greatly depends on the presentation of DOPA groups in the adhesive interface, either as a mono- or multilayer conformation. Because the adhesive structures can be transferred onto substrates from low-viscosity aqueous solution, they may serve as interesting nanoscale gluing pads in future applications, where the high viscosity of polymerbased glues renders the controlled formation of nanoscale adhesion pads difficult.

1. Introduction The production of functional microdevices requires the parallel development of successful methods for their assembly and packaging. At present, the production of small adhesive pads with locally high bonding strength is a technical challenge. Advanced techniques under investigation are typically based on van der Waals surface interactions, using patterned carbon nanotubes,1 nanostructured or porous silica,2 needle interfaces,3 or soft fibrillar surfaces.4,5 Although satisfactory adhesive strength can be achieved, it requires intensive surface modification and can be applied only to specific substrate types. Gluing is a more versatile technique, but it is limited by the minimum amount of adhesive that can locally be deposited because of the typically high viscosities of commercially available polymer-based glues. Besides the commonly used dispensing techniques for glue deposition, a series of lithographical techniques have recently been developed for the organization of UV-curable polymer materials into micrometer-sized patterns (30 to 10 μm).6 Theoretically, the smallest local gluing pad consists of a single-molecule interaction7 *Corresponding author. E-mail: [email protected]. Phone: þ49 (0)6151 16 7475. Fax: þ49 (0)6151 16 2479.

(1) Ge, L.; Sethi, S.; Gi, L.; Ajayan, P. M.; Dhinojwala, A. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 10792–10795. (2) Northen, M. T.; Turner, K. L. Nanotechnology 2005, 16, 1159–1166. (3) Stubenrauch, M.; Fischer, M.; Kremin, C.; Hoffmann, M.; Muller, J. IET Micro Nano Lett. 2007, 2, 6–8. (4) Glassmaker, N. J.; Jagota, A.; Hui, C. Y.; Kim, J. J. R. Soc., Interface 2004, 1, 23–33. (5) Geim, A. K.; Dubonos, S. V.; Grigorieva, I. V.; Novoselov, K. S.; Zhukov, A. A.; Shapoval, S. Y. Nat. Mater. 2003, 2, 481–483. (6) Samyn, P.; Biesalski, M.; R€uhe, J. To be submitted for publication. (7) Evans, E.; Ritchie, K. Biophys. J. 1997, 72, 1541–1555. (8) Geleshuber, I. C.; Thompson, J. B.; Del Amo, Y.; Stachelberger, H.; Kindt, J. H. Mater. Sci. Technol. 2002, 18, 763–766.

Langmuir 2010, 26(11), 8573–8581

or diatomic interactions.8 Such systems have been scientifically proven but are technically difficult to control. An impressive example of controlling adhesion on the nanoscale is found in nature. Living cells adhere to various substrates by means of a hierarchical structure of nano- to microintegrated attachment mechanisms with local adhesive forces of about 100 pN to 1 μN.9,10 Another mechanism is used by certain molluscs (e.g., blue mussels (M. edulis)) that adhere to various substrates by means of a byssal thread and foot plaque.11 Mussel foot proteins (Mefp) are in direct contact with the substrates to which mussels adhere. Their primary structure consists, among further sequences, of a regularly found decapeptide (Ala-Lys-Pro-SerTyr-Hyp*-diHyp*-Thr-DOPA*-Lys).12-17 Mefp’s are remarkable because their adhesive properties are nonspecific, strong, and irreversible: because of locally high adhesive strength, mussels can adhere strongly, even when the contact areas are very small. The structure and adhesive properties of the Mefp’s have been investigated by many researchers.18-21 On the basis of their (9) Benoit, M.; Gaub, M. E. Cell Tiss. Org. 2002, 175, 174–179. (10) Tsang, P. H.; Li, G.; Brun, Y. V.; Freund, L. B.; Tang, J. X. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 5764–5768. (11) Deming, T. J. Curr. Opin. Chem. Biol. 1999, 3, 100–105. (12) Zhao, H.; Waite, J. H. J. Biol. Chem. 2006, 281, 26150–26158. (13) Silverman, H. G.; Roberto, F. F. Mar. Biotechnol. 2007, 9, 661–681. (14) Wiegemann, M. Aquat. Sci. 2005, 67, 166–176. (15) Moeser, G. M.; Carrington, E. J. Exp. Biol. 2006, 209, 1996–2003. (16) Zhao, H.; Robertson, N. B.; Jewhurst, S. A.; Waite, J. H. J. Biol. Chem. 2006, 281, 11090–11096. (17) Lin, Q.; Gourdon, D.; Sun, C.; Andersen, N. H.; Anderson, T. H.; Waite, J. H.; Israelachvili, J. N. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 3782–3786. (18) Waite, J. H. Int. J. Biol. Macromol. 1990, 12, 139–144. (19) Papov, V. V.; Diamond, T. V.; Biemann, K.; Waite, J. H. J. Biol. Chem. 1995, 270, 20183–20192. (20) Ninan, L.; Monahan, J.; Stroshine, R. L.; Wilker, J. J.; Shi, R. Biomaterials 2003, 24, 4091–4099.

Published on Web 03/18/2010

DOI: 10.1021/la904629a

8573

Article

properties, a variety of functional synthetic materials have been developed22-24 that take advantage of the key role of DOPA (3,4dihydroxy-L-phenylalanine) catechol groups, which are enzymatic modifications of a tyrosine amino acid.25 The amount of DOPA varies from 10 to 15 mol % along the byssal thread toward 30 mol % near the adhesive plaque appearing as a foamlike substance.26 The extraction of the adhesive directly from mussels has proven to be inefficient because approximately 10 000 mussels were required to yield 1 g of protein.27 In-depth investigations of the adhesive properties of the protein materials were limited by the inefficient enzymatically triggered conversion of the tyrosine amino acids in the precursor protein to the respective adhesive DOPA sites. Taylor et al.28 described an improved method for hydroxylating tyrosine-containing sequences in polypeptides into peptidyl DOPA using mushroom tyrosinase at relatively high enzyme-to-substrate ratios. Finally, direct chemical synthesis methods have been used to mimic the cohesive and adhesive properties of Mefp’s by incorporating catechol functional groups into synthetic polymers. Interesting examples progress from hydrogel-forming polymers that statistically carry catechol functional groups29-35 to polymers that are end functionalized with DOPA groups36-38 to DOPA-modified oligomeric peptides that were developed to generate stable antifouling surface coatings.39 DOPA functional materials have been investigated as novel affinity fusion tags (e.g., poly-(DOPA)4 tags) for immobilizing proteins on biomaterials.40,41 The many functions of DOPA highlight the importance of being able to control its content and location when designing a bioinspired adhesive interface.42 However, a number of major challenges remain. First, DOPA-functional synthetic polymers expose the adhesive groups in a rather statistical manner, once applied as a “bioinspired glue” on a surface. To understand the applicability of catechol-functional polymers and polymeric materials as novel adhesives, the presentation of the respective (21) Baty, A. M.; Suci, P. A.; Tyler, B. J.; Geesy, G. G. J. Colloid Interface Sci. 1996, 177, 307–315. (22) Kanyalkar, M.; Srivastava, S.; Coutinho, E. Biomaterials 2002, 23, 389– 396. (23) Waite, J. H.; Holten-Andersen, N.; Jewhurst, S.; Sun, C. J. J. Adhes. 2005, 81, 297–317. (24) Kitamura, M.; Kawakama, K.; Nakamura, N.; Tsumoto, K.; Uchiyama, H.; Ueda, Y.; Kumagai, I.; Nakaya, T. J. Polym. Sci., Part A: Polym. Chem. 1999, 37, 729–736. (25) Waite, J. H. J. Biol. Chem. 1983, 258, 2911–2915. (26) Cha, H. J.; Hwang, D. S.; Lim, S. Biotechnol. J. 2008, 3, 1–8. (27) Hwang, D. S.; Yoo, H. Y.; Jun, J. H.; Moon, W. K.; Cha, H. J. Appl. Environ. Microbiol. 2004, 70, 3352–3359. (28) Taylor, S. W. Anal. Biochem. 2002, 302, 70–74. (29) Loizou, E.; Weisser, J. T.; Dundigalla, A.; Procar, L.; Schmidt, G.; Wilker, J. J. Macromol. Biosci. 2006, 6, 711–718. (30) Haemers, S.; van der Leeden, M. C.; Koper, G. J. M.; Frens, G. Langmuir 2002, 18, 4903–4907. (31) Burzio, L. A.; Waite, J. H. Protein Sci. 2001, 10, 735–740. (32) Yu, M. E.; Hwang, J. Y.; Deming, T. J. J. Am. Chem. Soc. 1999, 121, 5825– 5826. (33) Guvendiren, M.; Messersmith, P. B.; Shull, K. R. Biomacromolecules 2008, 9, 122–128. (34) Lee, H.; Lee, B. P.; Messersmith, P. B. Nature 2007, 448, 338–342. (35) Lee, B. P.; Chao, C. Y.; Nunalee, F. N.; Motan, E.; Shull, K. R.; Messersmith, P. B. Macromolecules 2006, 39, 1740–1748. (36) Dalsin, J. L.; Lin, L.; Tosatti, S.; V€or€os, J.; Textor, M.; Messersmith, P. B. Langmuir 2005, 21, 640–646. (37) Catron, N. D.; Lee, H.; Messersmith, P. B. Biointerfaces 2006, 1, 134–141. (38) Doraiswamy, A.; Dinu, C.; Cristescu, R.; Messersmith, P. B.; Chisholm, B. J.; Stafslien, S. J.; Chrisey, D. B.; Narayan, R. J. J. Adhes. Sci. Technol. 2007, 13, 1–13. (39) Statz, A. R.; Meagher, R. J.; Barron, A. E.; Messersmith, P. B. J. Am. Chem. Soc. 2005, 127, 7972–7973. (40) Jennissen, H. P.; Laub, M. Mater-wiss Werkstofftech. 2007, 38, 1035–1039. (41) Burke, S. A.; Jones, M. R.; Lee, B. P.; Messersmith, P. B. Biomed. Mater. 2007, 2, 203–210. (42) Alfonta, L.; Zhang, Z.; Uryu, S.; Loo, J. A.; Schultz, P. G. J. Am. Chem. Soc. 2003, 125, 14662–14663.

8574 DOI: 10.1021/la904629a

Samyn et al.

concentration and location of DOPA (i.e., catechol) groups at the interface needs to be controlled. If catechol-functionalized polymers are considered, then nonspecific surface interaction may largely affect the latter parameters. To better exploit the properties of nonoxidized DOPA (i.e., catechol)-functionalized materials, it is important to have control over the presentation of the functional adhesive sites relatively to the surface and to provide a stable matrix incorporating the adhesive groups. An interesting way to present functional groups in a well-controlled manner at planar or spherical interfaces has been introduced by the use of so-called peptide amphiphiles containing specific peptide sequences.43,44 The peptide amphiphiles consist of amphiphilic molecules such as lipids or simple fatty acids that are modified with a peptide sequence at the polar headgroup. They are versatile building blocks and have the ability to self-assemble spontaneously into organized 2D (i.e., monolayers and bilayers)45,46 or 3D structures (i.e., vesicles and rodlike micelles)47 presenting the active headgroup at the outer surface. To enhance the stability of such self-organized structures, a polymerizable amphiphile was recently used to achieve stable monolayer assemblies48 as well as vesicle-based peptidefunctional nanoparticles.49 For the latter, diacetylenic fatty acids were used and polymerized under ultraviolet radiation involving a 1,4 addition reaction between the unsaturated carbon-carbon links. Since the pioneering work of Wegner50 in the 1970s, the polymerization of diacetylenes and diacetylenic amphiphiles has been recognized as solid-state polymerization and has been further detailed in the literature.51-53 The polymerization reaction enhances the vesicle stability and produces an electron-rich polymer backbone consisting of conjugated double and triple bonds that exhibit interesting chromatic properties, such as the absorption of visible light. In particular, the successful polymerization of diacetylenic vesicles can be detected by the appearance of a blue or red color upon UV light irradiation. In previous work, we have shown that polymerized vesicles exposing a defined number of functional peptide groups at their surface are welldefined in size (about 100 to 200 nm in diameter) and can be dispersed in a low-viscosity aqueous environment.49 Here we describe the synthesis of polymerizable peptide amphiphiles based on diacetylenic fatty acids, which are modified with DOPA functional groups. We study the self-assembly of such molecules into vesicles and their stabilization through UV-light-induced polymerization. Vesicles with a defined number of DOPA groups are prepared by mixing a DOPA-functional amphiphile with a polymerizable matrix peptide amphiphile. To study the adhesion properties of the obtained systems, polymerized versicles were adsorbed onto test materials such as silicon, glass, and PMMA substrates and characterized with respect to their morphology and adhesive properties. (43) Berndt, P.; Fields, G. B.; Tirrell, M. J. Am. Chem. Soc. 1995, 117, 9515– 9522. (44) Yu, Y. C.; Berndt, P.; Tirrell, M.; Fields, G. B. J. Am. Chem. Soc. 1996, 118, 12515–12520. (45) Fields, G. B.; Lauer, J. L.; Dori, Y.; Forms, P.; Yu, Y. C.; Tirrell, M. Biopolymers 1998, 47, 143–151. (46) Dori, Y.; Bianco, H.; Satija, S. K.; Fields, G. B.; McCarthy, J. B.; Tirrell, M. J. Biomed. Mater. Res. 2000, 50, 75–81. (47) Hartgerink, J. D.; Beniash, E.; Stupp, S. I. Science 2001, 294, 1684–1688. (48) Biesalski, M.; Knaebel, A.; Tirrell, M.; Tu, R. Biomaterials 2006, 27, 1259– 1269. (49) Biesalski, M.; Tu, R.; Tirrell, M. Langmuir 2005, 21, 5663–5666. (50) Wegner, G. Pure Appl. Chem. 1977, 49, 443–454. (51) Bloor, D.; Chance, R. R. Polydiacetylenes: Synthesis, Structure, And Electronic Properties; M. Nijhoff: Dordrecht, The Netherlands, 1985. (52) Okada, S.; Peng, S.; Spevak, W.; Charych, D. Acc. Chem. Res. 1998, 31, 229–239. (53) Huo, Q.; Russell, K. C.; Leblanc, R. M. Langmuir 1999, 15, 3972–3980.

Langmuir 2010, 26(11), 8573–8581

Samyn et al.

Article

Figure 1. Peptide amphiphiles: (a) TDA-Gly-OMe, matrix amphiphile and (b) TDA-(EO)2-DOPA-C2-OH, adhesive amphiphile.

2. Material Preparation and Characterization Materials. All chemicals and solvents were received from Sigma-Aldrich or Roth KG (p.a. grade or higher) and used without further purification, if not otherwise stated. 10,12-Tricosadiynoic acid (TDA) was received from GFS Chemicals (Powell, OH), glycine methyl ester (H-Gly-OMe 3 HCl) was received from Advanced ChemTech (Louisville, KY), and 2-hydroxyethylamine-2-chlorotrityl resin (Novasyn TGT), 1-hydroxy-benzotriazole (HOBT), 2-(1-hydroxy-benzotriazole-1-yl)-1,1,3,3-tetramethyluronium hexafluorophosphate (HBTU), Fmoc-DOPA (acetonid)OH,54 and Fmoc-NH-(PEG)2-OH were received from Merck Chemicals Ltd. (Nottingham, U.K.). Monomeric TDA was received as a white-blue powder that was purified by dissolving the solid in chloroform and filtering through a 0.45 μm nylon filter. The recrystallized TDA was finally obtained as a white powder by evaporating the solvent in vacuum. TDA was stored under dark conditions and at 4 °C until further use. The used D.I.-grade water (pH 7) was purified by passing it through a Milli-Q system (Millipore, Eschborn, Germany) prior to use.

10,12-Tricosadiynoic-glycidyl-methyl-ester (TDA-Gly-OMe) Amphiphile. Amphiphile TDA-Gly-OMe (matrix amphiphile, Figure 1a) was synthesized by conventional solution-phase organic chemistry means. In brief, 15 g (0.043 mol) of TDA was dissolved in 30 mL of DMF and stirred with 9 mL (0.052 mol) of diisopropyl ethylamine (DIPEA) under cooling at about 5 °C. For the activation of the carboxylic acid function, HBTU (16.3 g, 0.043 mol) and anhydrous HOBt (5.8 g, 0.043 mol) were added and the solution was stirred until all solid material was dissolved (about 1 h). Eleven grams (0.086 mol) of H-Gly-OMe 3 HCl was separately dissolved in 30 mL of DMF, stirred for about 1 h, and added dropwise to the solution of activated TDA. The reaction solution was then stirred for another 15 h at room temperature. Precipitation of the product was facilitated by dropping the DMF solution into ice-cold water, and the product was collected by filtration and further purified on an alumina column using a 1:1 v/v chloroform/methanol mixture. Drying of the product in vacuum yielded a photosensitive white solid (final yield: 78% of theoretical yield). The solid was stored in the dark at -20 °C until further use. The product was characterized with 1H/13C NMR (CDCl3, 250 MHz, Bruker) and FTIR (Excalibur FTS 3000, BioRad Laboratories, Cambridge, MA). 1 H NMR (CDCl3, δ): 0.88 (t, 3H, CH3); 1.2-1.7 (m, 28H, CH2); 2.2 (m, 6H, CH2CO and CH2CtC); 3.8 (s, 3H, OCH3), 4.1 (d, 2H, NHCOCH2); 6 (b, 1H, NH). 13C NMR (CDCl3, δ): 172.1, 168.6, 68.8, 62.3, 51.1, 40.6, 36.0, 31.7, 29.6, 29.3, 28.8, 28.6, 28.5, 28.1, 25.8, 22.6, 17.9, 14.1. FTIR (in KBr; ν in cm-1): 3310 (NH), 2922 (CH), 1733 (COOR), 1678 (CONH), 1575 (CONH, CH).

10,12-Tricosadiynoic-amido-bis(ethylenoxide)-dihydroxiphenylalanine-2-hydroxyethyl Amide (TDA-(EO)2-DOPAC2-OH) Amphiphile. Amphiphile TDA-(EO)2-DOPA-C2-OH (adhesive amphiphile, Figure 1b) was synthesized according to solid-phase organic chemistry protocols using a Fmoc protecting group strategy.49 In brief, 2-hydroxyethylamine-2-chlorotrityl (54) Hu, B.; Messersmith, P. B. Tetrahedron Lett. 2000, 41, 5795–5798.

Langmuir 2010, 26(11), 8573–8581

resin (3.8 g, 0.65 mmol, load: 0.17 mmol/g) was swollen for 30 min in 20 mL of DMF, and 10 equiv of Fmoc-DOPA (acetonid)-OH (3 g, 6.5 mmol), together with diisopropyl carbodiimide (DIC, 0.47 mL) and dimethyl aminopyridine (DMAP, 8 mg), was added. The resin was shaken for 20 min at room temperature. After subsequent filtration and repetitive washing steps with DMF, methanol, and methylene chloride, the coupling step was repeated three times in order to increase the yield. The Fmoc protection was then removed by shaking the resin in piperidine (20% in DMF) three times for 10 min, followed by repetitive DMF washing. The coupling of 5 equiv of an ethylene oxide spacer (1.5 g, 3.25 mmol) and 5 equiv of TDA (1.3 g, 3.25 mmol) in 1:1 v/v DCM/DMF was carried out accordingly, each activated by 5.5 equiv of HOBt (0.48 g, 3.57 mmol), 5.5 equiv of HBTU (1.35 g, 3.57 mmol), and 10 equiv of DIPEA (1.13 mL, 6.5 mmol). Cleavage of the peptide amphiphile from the resin and removal of the acetonid side chain protecting group was done by treating the resin with 95% TFA/water for 1 h at room temperature. The crude product was precipitated from ice-cold methyl t-butyl ether, decanted, and freeze dried in vacuum. The peptide amphiphile was purified by HPLC on a reversed-phase C4 column with a 0.1% TFA gradient of acetonitrile in water. The final product was characterized with 1H/13C NMR (DMSO-d6, 250 MHz, Bruker), FTIR (Nicolet Magna 850), and electron spray ionization mass spectrometry (ESI)-MS. The presence of unprotected catechol groups offering adhesive properties was confirmed by FTIR analysis (unprotected catechol: νC-O = 1285 cm-1) and NMR spectra of TDA-(EO)2-DOPA-C2-OH and Fmoc-DOPA (acetonid)-OH: the 13C NMR spectrum did not show a signal at δ = 117.8, which is typical of the quaternary carbon of an acetonide protecting group of catechol, whereas a peak appeared at δ = 19.8 for the unprotected catechol. 1 H NMR (DMSO-d6, δ): 0.88 (t, 3H, CH3); 1.00 (d, 2H, CH2 on a DOPA side chain); 1.20-1.30 (m, 28H, CH2); 1.40-1.65 (s, 1H, CH on a DOPA side chain); 1.84 (s, 4H, CH2CHOH on a DOPA side chain); 2.10 (m, 6H, CH2CO and 4H, CH2CtC); 2.35 (s, 2H, CHOH in DOPA catechol); 2.89 (d, 2H, CH2 on a DOPA side chain); 3.18 (d, 1H, DOPA); 3.45-3.47 (m, DOPA); 4.12 (m, 2H, CH2CO in PEG); 5.25 - 5.45 (m, 2H, CHOH in DOPA catechol); 6.61-6.55 (m, 3H, DOPA phenyl ring); 7.24 (d, 2H, CHOH in DOPA); 8.18 (d, CONH); 8.3 (d, CONH). 13C NMR (DMSO-d6, δ): 173.2, 168.6, 144.0, 121.5, 134.5, 84.6, 80.0, 67.5, 64.2, 60.0, 49.46, 45.0, 41.80, 41.36, 41.02, 40.69, 40.35, 40.02, 39.69, 35.0, 32.5, 31.53, 30.0, 29.6, 25.0, 22.5, 19.8, 17.9, 14.6. FTIR (in KBr; ν in cm-1): 3310 (NH), 2922 (CH), 1678 (CONH) 1575 (CONH, CH), 1285 (CO, catechol). ESI-MS: Mcalc = 751.04 g/mol (calculated) Mexp = 751.2 g/mol (experimental). Vesicle Preparation and Polymerization. The synthesized amphiles were dissolved in chloroform, and 5 mg/mL stock solutions were prepared. The solutions were mixed in concentrations containing 0, 10, 20, 30, 35, 50, 70, 90, and 100 mol % adhesive amphiphiles and corresponding amounts of matrix amphiphiles to add up to 100%. After drying under vacuum for chloroform removal, the amphiphilic films were hydrated in a small vial with water for 3 h above the melting temperature (Tm = 67 °C). The solutions were sonicated above the melting DOI: 10.1021/la904629a

8575

Article

Samyn et al.

Figure 2. Self-organization of the matrix and adhesive amphiphiles and stabilization by UV-induced solid-state polymerization: (a) vesicle formation and (b) polymerization mechanism of the TDA tails (edited from Wegner et al.),50 with R1 being the DOPA-functionalized peptide sequence or matrix sequence and R2 being the alkyl chain. temperature of the molecules, using a tip sonifier (Bandelin Sonopuls HD 2070, power 12 W, time 5 min). After filtration and a minimum incubation period of 12 h at 4 °C, the vesicle solutions were exposed to ultraviolet irradiation (4W TLC-lamp, 250 nm) for 90 s in order to perform a polymerization reaction. The solutions had a final vesicle concentration of 0.125 mg/mL and were further diluted to 0.0125 mg/mL by adding water (pH 7). The polymerized vesicles were found to be stable for about 5 days in aqueous solution at this pH when stored at 4 °C. The chemical composition of the nonpolymerized and polymerized vesicles was characterized by FTIR and UV-vis spectroscopy. The morphology of the vesicles was characterized by TEM and ESEM.

Characterization of Surface-Adsorbed Amphiphiles and Vesicles. For the investigation of the morphology of surfaceconfined vesicles as well as adhesive properties of the vesicle layers prepared, a general procedure was followed for the adsorption of vesicles from solution onto their respective substrates: a volume of 50 μL of dispersed amphiphiles or vesicles in aqueous solution (pH 7) with concentrations between 0 and 0.125 mg/mL was drop cast onto PMMA, glass, or silicon substrates. The substrates were subsequently rinsed with ethanol and water prior to use. First, the morphology of the adsorbed, dried adhesive was characterized by atomic force microscopy (tapping mode). Second, an adhesive joint was formed by immediately placing an identical substrate onto the wet substrate. The substrates were allowed to rest for about 10 min in order to build the adhesive bond in the overlap region (surface area 25  50 mm2). The samples were vertically mounted into a Zwick tensile tester and clamped over a minimum length of 20 mm into the pulling heads. A load cell with a maximum capacity of 1 kN and an accuracy of 0.1 N at a speed of 5 mm/min was used during shear experiments. Although the adhesive strength on the microscale is usually expressed in MPa (N/mm2), we will report data in terms of shear forces (N) using a nanostructured adhesive material, where adhesive contacts are established only in a discrete number of nanoscopic points and hence the real contact area is smaller than the apparent macroscopic contact area. The adhesive strength was determined to be the maximum shear force at the beginning of the experiment, and the evolution of the shear force over a sliding distance of 25 mm was further measured. The statistical variation of a single value of the adhesive strength is about 5 to 7%, as determined from three repetitive measurements. All data apply to amphiphiles or vesicles contained in an aqueous film between the substrates and can be used to evalute the efficiency of the adhesive vesicles in modifying the adhesive properties.

3. Results Preparation and Characterization of Polymerized Vesicles Exposing DOPA Functional Groups. A polymerizable, DOPA-functional amphiphile and a matrix amphiphile were 8576 DOI: 10.1021/la904629a

prepared by means of solid-phase organic chemistry and solution-phase synthesis, respectively, as described in the Material Preparation and Characterization section. The amphiphiles were self-organized into vesicular structures (Figure 2a) and subsequently polymerized (Figure 2b) following a protocol published elsewhere.49 The molar composition of the vesicles was varied by adjusting the respective concentrations of the amphiphiles in the stock solutions. After the preparation of the vesicles, the chemical composition was proven by FTIR (Supporting Information 1). UV-vis spectroscopy showed that vesicles containing up to 35 mol % TDA-(EO)2-DOPA-C2-OH have strong absorption bands with a maximum absorbance at about 640 (blue) to 550 nm (red) (Supporting Information 2). This wavelength is characteristic of a well-ordered polymer system, indicating that the vesicles were successfully polymerized. Vesicles with 35-100 mol % TDA-(EO)2-DOPA-C2-OH exhibit no absorption band at any visible wavelength upon UV-light irradiation, suggesting that the amphiphiles do not polymerize. It is known that the polymerization of diacetylenic groups requires monomeric ordering with typical intermolecular distances of 5 A˚ at an angle of approximately 45°.50 Thus, the polymerization is likely affected by the size or by molecular interactions of the polar headgroups of the amphiphiles and is successful only at low concentrations of the DOPA-functional amphiphile. The morphology of polymerized vesicles was studied by cryogenic TEM (Figure 3a) and shows well-defined spherical or rod shapes for polymerized assemblies containg up to 35 mol % of the DOPA-functional amphiphile and filmlike or fiberlike structures at higher molar percentages of the adhesive component. The morphology of polymerized vesicles that were adsorbed onto solid surfaces was characterized by AFM analysis (Figure 3b), also indicating well-defined morphologies that depend on the molar percentage of adhesive amphiphiles. The vesicles containing 10 to 20 mol % TDA-(EO)2-DOPA-C2-OH have a spherical shape with a diameter of about 200-400 nm and a height of about 100 nm. Vesicles containing more than 35 mol % TDA-(EO)2-DOPA-C2-OH typically form rather planar structures with a length of 100 to 150 nm and a height of about 10 nm. The static contact angles of water droplets on a dried film of vesicles adsorbed on PMMA varied from 78 ( 3° (pure PMMA slide) to 95 ( 3° (vesicles containing 10 mol % TDA-(EO)2DOPA-C2-OH), 82 ( 1° (50 mol % TDA-(EO)2-DOPA-C2-OH), and 79 ( 2° (100 mol % TDA-(EO)2-DOPA-C2-OH), respectively (Supporting Information 3). These studies suggest that the substrate locally contains adsorbed nanoscale vesicles with morphologies and topologies that depend on the amphiphilic Langmuir 2010, 26(11), 8573–8581

Samyn et al.

Article

Figure 3. Morphological characterization of polymerized vesicles containing different amounts (mol %) of TDA-(EO)2-DOPA-C2-OH amphiphiles: (a) TEM images and (b) AFM height images of vesicles deposited on PMMA substrates.

Figure 4. Shear tests on adhesive interlayers with (i) disordered and nonpolymerized amphiphiles or (ii) polymerized vesicles containing 20 mol % TDA-(EO)2-DOPA-C2-OH in contact with SiO2: online measurement of shear force in the tensile tester over a 25 mm sliding distance and optical microscopy of ruptured interfaces.

composition and polymerization of the vesicles. Comparing the surface characterization by environmental AFM and cryogenic TEM studies of the polymerized vesicles illustrates that the adsorption of vesicles onto planar substrates does not alter the morphology of the nano-objects. Adhesive Properties of Surface-Adsorbed Vesicles. To study the adhesion properties, we performed shear tests on surfaces decorated with adsorbed vesicles using a standard tensile tester. First, reference shear tests were performed using disordered and nonpolymerized TDA-Gly-OMe (matrix) amphiphiles and TDA-(EO)2-DOPA-C2-OH (adhesive) amphiphiles deposited from a chloroform solution between two chemically identical surfaces with different molar percentages. Second, shear tests were performed with assembled amphiphiles that were deposited from an aqueous solution with a fixed concentration of 0.125 mg/mL. Here we followed different routes: (i) we adsorbed assembled, Langmuir 2010, 26(11), 8573–8581

nonpolymerized vesicles onto a solid substrate, (ii) we performed UV-light-induced polymerization of the adsorbed vesicles in contact with the substrate, or (iii) we first polymerized the vesicles in solution and subsequently drop cast these materials onto the substrate. An example of recorded force-displacement curves and ruptured interfaces for adhesive interlayers with nonpolymerized and polymerized vesicles is shown in Figure 4. For disordered and nonpolymerized amphiphiles, the adhesive strength at bond rupture is low (e.g., 2.0 N, Figure 4) and the performance over longer sliding times is dynamically unstable, as seen by a progressive increase in shear force over the sliding path. The shape of disordered and nonpolymerized amphiphiles is obviously not controlled, and the assemblies may spontaneously reorganize under mechanical stress, causing instabilites. Although the amphiphiles are initially randomly oriented in the interface, microscopic images show that they tend to align along the DOI: 10.1021/la904629a

8577

Article

Samyn et al.

Figure 5. Adhesive strength of nonassembled and nonpolymerized amphiphilic mixtures containing different amounts (mol %) of TDA-(EO)2-DOPA-C2-OH amphiphiles deposited on various substrates.

direction of highest shear and form fibrous structures. This (re)organization may consequently increase the interfacial shear resistance. The fibrous amphiphilic structures form inhomogeneous, noncoherent domains, and the cracks in between those domains lead to premature failure of the adhesive bond between both substrates (Figure 4). Bond rupture is typically initiated by delaminations that propagate over the interface. The adhesive strength of the disordered amphiphilic monolayers is presented in Figure 5 as a function of the amphiphilic concentration deposited on PMMA and SiO2 substrates. The concentration of the amphiphile mixtures is progressively diluted from 5 to 0.001 mg/mL in water. The shear resistance of amphiphiles at the lowest concentrations approximately equals the value of a pure interfacial water film. A slight increase in adhesive strength is noticed for 0 mol % TDA-(EO)2-DOPA-C2-OH (i.e., pure matrix amphiphile), likely caused by the slightly higher viscosity of the interfacial film. It is evident that higher molar percentages of TDA-(EO)2-DOPA-C2-OH amphiphiles within the vesicles yield better adhesive strength. For assembled and polymerized vesicles, the adhesive strength at bond rupture is higher (e.g., 3.7 N, Figure 4) compared to that for disordered and nonpolymerized assemblies. It is also dynamically more stable as a function of the sliding path, as observed from a relatively flat force-displacement diagram. The morphological stability of polymerized vesicles and the formation of macroscopic vesicle patterns in the adhesive interface are confirmed by a microscopic image (Figure 4), where the vesicles do not form a homogeneous film. The adhesive strength of polymerized vesicles relative to the strength of nonpolymerized (but assembled) vesicles is quantified with deposition on glass substrates (Figure 6a) or PMMA substrates (Figure 6b) as a function of the molar composition of matrix and adhesive amphiphiles. The adhesive strength of polymerized vesicles is higher than that of the nonpolymerized vesicles and is even further increased if polymerization is carried out in contact with the substrate. Those tendencies are reproducible for all molar compositions of the vesicles and are more clear for glass substrates than PMMA substrates because the latter surfaces have a higher roughness (Ra,PMMA = 1.7 nm, Ra,glass = 1.0 nm) disfavoring intense contact between the substrate and adhesive vesicles. A large increase in adhesive strength after the polymerization of the amphiphiles in contact with PMMA might be caused by additional chemical activation of PMMA under UV radiation. The surface activation of PMMA is confirmed by an increase in total surface energy from 42 ( 2 to 51 ( 2 mJ/m2 after UV radiation (contact angle data not 8578 DOI: 10.1021/la904629a

Figure 6. Adhesive strength of coassembled amphiphiles and polymerized vesicles on (a) glass substrates and (b) PMMA substrates: (i) coassembled, nonpolymerized amphiphiles, (ii) coassembled vesicles polymerized before surface adsorption, (iii) coassembled vesicles polymerized after surface adsorption, and (iv) coassembled and polymerized vesicles applied to a UV-activated surface (120 s at 250 nm).

shown). This effect was also observed by Wei et al.55 and was explained by the transformation of ester groups into carboxylic end-groups, which in the present case may enhance vesicle adsorption. After the polymerization of the vesicles in contact with the surface, they arrange in fiberlike patterns on glass and in dendritic patterns on PMMA (Figure 7). Next, we were interested in understanding the influence of the number of adsorbed, adhesion-mediating vesicles on the adhesive properties of the prepared surfaces. Therefore, the concentration of dispersed polymerized vesicles containing 10 20, and 100 mol % TDA-(EO)2-DOPA-C2-OH amphiphiles was varied between 0.125 and 0.0125 mg/mL by progressive dilution in water; subsequently, a constant volume of 50 μL of adhesive solution was deposited onto a fixed surface area. Because the counterface substrate is immediatly applied, the vesicle concentration in solution directly relates to the number of vesicles deposited in the interface and a small concentration of vesicles in solution yields a small number of vesciles in the contact area. The adhesive strength of the joint PMMA, glass, and SiO2 substrates is shown in Figure 8 as a function of the vesicle (55) Wei, S.; Vaidya, B.; Patel, A. B.; Soper, S. A.; McCarley, R. L. J. Phys. Chem. 2005, 109, 16988–16996.

Langmuir 2010, 26(11), 8573–8581

Samyn et al.

Article

Figure 7. Macroscopic morphology of polymerized vesicles containing 20 mol % TDA-(EO)2-DOPA-C2-OH amphiphiles in contact with (a) glass substrates and (b) PMMA substrates (optical microscopy).

concentration and the molar percentage of TDA-(EO)2-DOPAC2-OH amphiphiles within the vesicles. For every vesicle concentration, higher molar percentages of adhesive amphiphiles in the vesicles do not necessarily imply higher adhesive strength. The adhesive strength of vesicles containing 10 and 20 mol % TDA-(EO)2-DOPA-C2-OH amphiphiles first increases from about 2 N (shear force for a pure water film between both substrates) to about 5, 20, and 35 N or 7, 35, and 45 N at 0.03 mg/mL vesicle concentration, respectively. If solutions are deposited with vesicle concentrations larger than 0.03 mg/mL, then the adhesive strength decreases again and at a vesicle concentration of 0.125 mg/mL it becomes similar to that of a pure water film. Thus, a maximum in shear strength is observed at intermediate vesicle concentrations and is independent of the molar percentage. (Similar trends are observed in Figure 8a,b.) This behavior is reproducible for every substrate type and indicates that the properties of the adhesive interlayer uniquely determine the adhesive properties. For vesicles containing 100 mol % TDA-(EO)2-DOPA-C2-OH (Figure 8c), the adhesive strength for all samples studied is much weaker than that for the 10 and 20 mol % vesicles and increases linearly with vesicle concentration from about 2 N for 0 mg/mL to only about 4 and 14 N for 0.125 mg/mL solutions for glass and SiO2 substrates, respectively (Figure 8c). For PMMA substrates, we observed only a marginal increase in the shear strength. At first it seems Langmuir 2010, 26(11), 8573–8581

Figure 8. Influence of the concentration of polymerized vesicles on adhesive strength for vesicles of varying composition of TDA-(EO)2DOPA-C2-OH amphiphiles ((a) 10 (b) 20, and (c) 100 mol %) and different substrate types ((i) PMMA, (ii) glass, and (iii) SiO2).

surprising that vesicles that contain more adhesion moieties exhibit drastically different adhesion behavior. The observed influence of vesicle concentration on adhesive strength illustrates the role of the morphology of the nanostructured adhesive interlayer. For vesicles containing 100 mol % TDA-(EO)2-DOPA-C2-OH amphiphiles, the lower adhesive strength with lower concentration agrees with expectations from continuous adhesive films because a smaller number of adhesive DOI: 10.1021/la904629a

8579

Article

Samyn et al.

Figure 9. Topography of the adhesive interlayer, depending on the concentration of surface-adsorbed vesicles: (a) AFM height images and profiles of vesicles deposited on silicon substrates and (b) schematic representation of the adhesive interface.

points decreases the overal interfacial strength. For vesicles containing 10-20 mol % adhesive amphiphiles, however, the maximum adhesive strength occurs around 0.03 mg/mL concentration, independently of the surface type and molar percentage of adhesive amphiphiles. The reason for the existence of the optimum concentration of surface-adsorbed vesicles can be understood if the topography of the interface is studied. Figure 9 shows the AFM analysis of vesicles deposited at low concentrations (0.01 mg/L) and high concentrations (0.06 and 0.12 mg/mL) onto a silicon substrate, together with a schematic representation of the adhesive interface. The AFM height images and profiles show that the vesicles are arranged in a monolayer configuration with a thickness of 30 nm when deposited at low concentration, whereas they form multilayer aggregates with a thickness of up to 300 nm when deposited at high concentrations. At low concentrations, the shear stresses are concentrated on a vesicular monolayer. Then, the adhesive strength of each individual vesicle contributes to the adhesion between the substrates and the adhesive strength increases with an increasing number of adhesive points. For that reason, polymerized vesicles having 20 mol % TDA-(EO)2DOPA-C2-OH amphiphiles provide a higher adhesive strength than do compositions with 10 mol % adhesive amphiphiles. At high concentrations, individual vesicles in contact with the substrate may not be able to make contact with the opposite substrate but they are in direct contact with other vesicles in the multilayer assembly. Consequently, the interfacial shear is concentrated within the plane of lowest resistance, which is located inside the multilayer adhesive film rather than at the substrate surface. Then, the interface morphology provides rather good lubricating properties if the vesicles are deposited from highly concentrated solutions. The present data indicate the benefits and unique behavior of a nanostructured adhesive monolayer, where the adhesive groups within every vesicle effectively contribute to the adhesion between substrates. At present, however, open questions remain about the statistiscial distribution and theoretical accessibility of the DOPA groups in the interface, presented either as discrete contact points 8580 DOI: 10.1021/la904629a

that depend on the local surface-concentration or as a homogeneous filmlike substance. The local presentation of DOPA groups and effects on the macroscale adhesive strength should be considered in relation to the local adhesive strength of a single vesicle. We considered a preliminary evaluation estimating the number of effective contact points (Supporting Information 4). The latter is currently under detailed investigation, and results will be discussed in the future.

4. Conclusions The synthesis, morphology, and chemical characterization of novel bioinspired adhesive nano-objects were evaluated by macroscopic adhesion experiments. Dihydroxy-phenylalanine (DOPA) functional groups were incorporated into well-defined selfassembled architectures allowing for the controlled local presentation of adhesives on the nanoscale. It was demonstrated that the adhesive groups (DOPA) can be coupled to a photopolymerizable fatty acid (TDA, tricosadiyonic acid) through standard peptide synthesis protocols to form a DOPA group containing amphiphile TDA-(EO)2-DOPA-C2-OH. The co-self-assembly of such functional amphiphiles with matrix amphiphile TDA-Gly-OMe, consisting of the same fatty acid modified with an inert amino acid headgroup, allows for the buildup of vesicles with a known concentration of adhesive sites and control over the presentation of the adhesive groups at the vesicle surface. The vesicular morphology depends on the molar amount of the adhesive amphiphile, and it changes from spherical-like (diameter ∼100 to 200 nm) to fiberlike (diameter, ∼5 to 10 nm; length, several hundred of nanometers), with higher contents of TDA-(EO)2DOPA-C2-OH. UV-light-induced polymerization of the selfassembled architectures allows for the formation of dimensionally, chemically, and mechanically stable spherical vesicles if the ratio of polymerizable groups to the other groups was chosen properly. Whereas previous work on functional diacetylenic assemblies has focused on the development of materials with interesting chromatic properties that are useful in sensing applications, the present work utilizes for the first time polymerized diacetylenic materials Langmuir 2010, 26(11), 8573–8581

Samyn et al.

for the development of nanoadhesive patches at surfaces, which enriches possible applications of these materials. Adhesive properties were tested by adsorbing polymerized and nonpolymerized vesicles onto varying solid substrates. A second substrate having the same surface chemistry was pressed onto the surface-adsorbed vesicle layer, and the shear forces required to separate both substrates were measured. The adhesive properties of nonpolymerized materials are mechanically unstable, and surface-adsorbed vesicles change their structure upon shear, arranging into islandlike domains with fibrillar structure. In contrast, polymerized assemblies show strong, reproducible adhesive properties. The molar percentage of the adhesive amphiphiles within the surface-deposited vesicles and the concentration of surface-adsorbed polymerized vesicles determine the adhesive strength in the interface, independent of the kind of substrate used. Thus, both parameters can be used to control and adjust the adhesive strength between two opposing surfaces: in particular, higher molar percentages of DOPA groups within the vesicles and higher numbers of vesicles adsorbed to the substrate strongly increase the shear forces required to separate the substrates. However, the number of vesicles deposited onto the substrate is critical. If a very large number of vesicles are deposited onto the substrate, then they form multilayers with greatly reduced adhesive strength, presumably because the vesicle-vesicle interactions can break more easily and fewer forces are required to shear vesicles against each other. Thus, the vesicle-based adhesive systems exhibit a clear optimum of the adhesion performance at some intermediate composition and vesicle concentration. The

Langmuir 2010, 26(11), 8573–8581

Article

data further clearly demonstrate that the accessibility of the DOPA-functional sites impacts the overall interfacial adhesion strongly, so this parameter needs to be carefully taken into account for the design of such bioinspired adhesives. Finally, an interesting feature of the bioinspired adhesive is that it can be applied to a low-viscous solution and that the viscosity of the solution is more or less independent of the glue/vesicle to solvent ratio. This is in strong contrast to conventional glues, where the viscosity is usually rather high and increases strongly with increasing polymer content. Such behavior could break the pathway into smaller and smaller glue pads. Acknowledgment. We thank Prof. Dr.-Ing. Wilde and Dipl.Ing. J. Dalin for fruitful and stimulating interactions within the joint Nanopad project. We greatly acknowledge the German Federal Ministry for Eduction and Research (BMBF) for funding within the program Micro- Nano Integration for Microsystems Technology (MNI-mst): Nanopad. We thank Dr. Oswald Prucker for valuable support. Dr. Ralph Thomann at the FMF of Freiburg University assisted us in taking TEM measurements. The technical support by Mr. Matthias Preisendanz and Mr. Stefan Schwarz is most appreciated. Supporting Information Available: FTIR and UV-vis of polymerized vesicles. Water contact angle determination on vesicles adsorbed onto the PMMA substrate. Illustrative statistical evaluation. This material is available free of charge via the Internet at http://pubs.acs.org.

DOI: 10.1021/la904629a

8581