Mussel-Inspired Chemistry for Robust and Surface-Modifiable

ARTICLE pubs.acs.org/Langmuir

Mussel-Inspired Chemistry for Robust and Surface-Modifiable Multilayer Films Junjie Wu,†,‡ Liang Zhang,†,‡ Yongxin Wang,†,‡ Yuhua Long,†,‡ Huan Gao,§ Xiaoli Zhang,† Ning Zhao,*,† Yuanli Cai,§ and Jian Xu*,† †

Beijing National Laboratory for Molecular Sciences, Laboratory of Polymer Physics and Chemistry, Institute of Chemistry, Chinese Academy of Sciences, Beijing 100190, PR China ‡ Graduate University of the Chinese Academy of Sciences, Beijing 100049, PR China § Jiangsu Key Laboratory of Advanced Functional Polymer Design and Application, Department of Polymer Science and Engineering, College of Chemistry, Chemical Engineering and Materials Science, Soochow University, Suzhou 215123, China

bS Supporting Information ABSTRACT: In this article, we report a bioinspired approach to preparing stable, functional multilayer films by the integration of mussel-inspired catechol oxidative chemistry into a layer-by-layer (LbL) assembly. A polyanion of poly(acrylic acid-g-dopamine) (PAA-dopamine) bearing catechol groups, a mussel adhesive protein-mimetic polymer, was synthesized as the building block for LbL assembly with poly(allylamine hydrochloride) (PAH). The oxidization of the incorporated catechol group under mild oxidative condition yields o-quinone, which exhibits high reactivity with amine and catechol, thus endowing the chemical covalence and retaining the assembled morphology of multilayer films. The cross-linked films showed excellent stability even in extremely acidic, basic, and highly concentrated aqueous salt solutions. The efficient chemical cross-linking allows for the production of intact free-standing films without using a sacrificial layer. Moreover, thiol-modified multilayer films with good stability were exploited by a combination of thiols-catechol addition and then oxidative cross-linking. The outstanding stability under harsh conditions and the facile functionalization of the PAA-dopamine/PAH multilayer films make them attractive for barriers, separation, and biomedical devices.

’ INTRODUCTION Layer-by-layer (LbL) assembly is a facilely and highly versatile method of constructing nanoscale multilayer films on a variety of surfaces.1
3 This method affords the creation of functional thin films with variable components, structures, and properties, which have been applied in the fields of biomimetic coatings, chemical sensors, biomedical engineering, and so on.4
6 Commonly, the driving forces of LbL assembly are noncovalent intermolecular interactions such as electrostatic forces,1,2 hydrogen bonding,7
9 and charge-transfer interactions.10,11 It is obvious that these noncovalently bonded multilayer films are weak and unstable, especially when used in harsh environments, as in the case of corrosion barriers,12,13 separation membranes,14,15 and durable devices such as medical implants.2,6 Therefore, covalent cross-linking of noncovalently assembled multilayer films is usually utilized to improve their stability. To this end, various strategies, including UV irradiation,16,17 temperature elevation,18,19 and others,20
24 have been extensively exploited. However, with the development of multilayer films for specific applications, stable multilayer films with chemical or biological functionality (e.g., the incorporation of fluorescent small molecules, ligands, drugs, peptides, and proteins) are gratifying. An exciting example of a r 2011 American Chemical Society

stable, functional multilayer film is combination of LbL assembly with the “click chemistry” of the cycloaddition reaction between alkynes and azides introduced by Caruso and coworkers.25 Robust multilayer films, that are coupled with antifouling,26 fluorscent27 and responsive28 functions, respectively, have been successfully prepared using this tactic. In addition, stable, functional multilayer films have also been exploited on the basis of other click-type reactions such as those for azlactones-amines,29 thiol-ene chemistry,30 and activated esters-amines.31 Other than the approaches mentioned above, the strategy for the stabilization and functionalization of noncovalent multilayer films has rarely been reported. Herein, we propose a biomimic route inspired by marine mussels to prepare robust surface-modifiable LbL-assembled multilayer films. Recently, mussels have attracted much attention because of their strong underwater adhesion to almost all types of surfaces.32 It has been confirmed that 3,4-dihydroxy-phenylalanine (DOPA), a particular amino acid in secreted mussel adhesive proteins (MAPs), is responsible for the strong adhesion.33,34 Received: July 15, 2011 Revised: September 29, 2011 Published: October 06, 2011 13684

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Langmuir The ortho-dihydroxyphenyl (catechol) moiety of DOPA plays multiple roles in mussel adhesion. The dismutation reaction of oxidized catechol (o-quinone) with phenol, the Michael addition or Schiff base reaction of o-quinone with amine, and the Michael addition reaction of o-qunione with thiol are the chemical foundation of the strong affinity for organic surfaces, whereas the strong coordination ability of catechol with many metal/metal oxides results in the high adhesion to inorganic surfaces.35
37 The oxidative chemistry and coordination interactions also lead to the rapid solidification of adhesive proteins in seawater38,39 and reinforce the adhesive materials of mussels as well.40,41 This biomimetic concept has been introduced into the current LbL assembly. On the basis of the strong interfacial binding property of catechol, Messersmith and co-workers42 exploited substrateindependent LbL assembly, even on a challenging substrate such as poly(tetrafluoroethylene) without prior surface modification by using catechol-modified poly(ethylenimine) and hyaluronic acid. Kotov et al.43 fabricated a strong nanocomposite by the LbL assembly of clay and polymer containing DOPA groups. DOPA served as an anchor for polymer attachment to clay and a bridge between polymer chains to forming a 3D network by Fe3+ coordination. A small amount of DOPA dramatically improved the toughness of the nanocomposite. These studies focused on the universal and strong interfacial adhesion (either between a multilayer film and substrates42 or between two building blocks43) of polymers containing catechol groups. Caruso et al.44 prepared biodegradable capsules using dopamine-modified poly(L-glutamic acid) (PGA) by a one-step or multistep technique of dopamine assembly. This method may be used to construct stable multilayer films on planar substrates. The assembly is based on the long-time (12 h) polymerization of conjugated dopamine moieties similar to the formation of pure polydopamine (PDA).34 Catechol oxidative chemistry has also been widely used in other areas, including surface modification,34,45,46 interface enhancement,47 and hydrogel formation;48,49 however, this chemistry has not been employed to improve the bulk stability and functionalize multilayer films. In this article, a mussel adhesive protein-mimetic polyanion was synthesized by the conjugation of dopamine, an analogue of DOPA, onto poly(acrylic acid) (PAA). Multilayer films were prepared by the alternate deposition of dopamine-modified PAA (PAA-dopamine) with a polycation of poly(allylamine hydrochloride) (PAH). The assembled films can be cross-linked under a mild oxidative condition. The chemistry and morphology evolution during cross-linking were studied by UV
visible, XPS, FTIR spectra, SEM, and AFM. The stability of the crosslinked film was assessed by exposing the film to an aqueous solution with an extreme pH value or a high ionic strength. The cross-linked multilayer films can be released from the substrate to produce intact free-standing films. Moreover, the feasibility of preparation of stable multilayer films with surface modification by thiols was illustrated.

’ EXPERIMENTAL SECTION Materials. Poly(acrylic acid) (PAA, Mw ≈ 8000, 35 wt % aqueous solution), dopamine hydrochloride, N-(3-dimethylaminopropyl)-N0 ethylcarbodiimide hydrochloride (EDC 3 HCl), poly(ethylene glycol acrylate) (PEGA, Mw = 454), and (3-aminopropyl)triethoxysilane (APTS) were purchased from Sigma-Aldrich. Poly(allylamine hydrochloride) (PAH, Mw ≈ 70 000) and 1-dodecanethiol (CH3(CH2)11SH) were purchased from Alfa Aesar. Thiol-terminated poly(poly(ethylene

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glycol acrylate)) (polyPEGA-SH, Mw ≈ 30 kDa and Mw/Mn < 1.2), was synthesized according to a previously reported method by reversible addition
fragmentation chain transfer (RAFT) polymerization of PEGA in aqueous solution under visible light radiation, followed by ammonolysis of the chain-transfer agent.50 Other reagents were analytical grade and used as received without further purification. For all experiments, deionized water was used. Instruments and Measurements. 1H nuclear magnetic resonance (NMR) measurements were carried out on a Bruker DMX-400 NMR spectrometer. UV
visible characterization was performed on a Shimadzu UV-1601PC spectrophotometer using a 1 cm quartz cuvette or quartz slides. FTIR spectra were recorded on a Bruker Equinox 55 FT-IR/FAR 106 spectrophotometer. The test specimens were prepared by the KBr disk method.The thickness of the multilayer film deposited on silicon was determined by EC-400 ellipsometry in air (J. A. Woollam. Co., Inc.). The amplitude (Ψ) and phase (Δ) of the polarized light were measured at incident angles of 65 and 70°. The film thickness was obtained by fitting the Ψ and Δ curves employing multilayer model Si/SiO2/polymer/air. The thickness of the SiO2 layer was measured for each silicon substrate before LbL assembly (generally ∼2.3 nm). The polymer layer was considered to be a Cauchy layer with an assumed refractive index of nfilm(λ) = 1.45 + (0.01/λ2).51,52 The data were the average of five different measurements. Atomic force microscopy (AFM) measurements were carried out in air at room temperature on a Nanoscopy IIIA (Digital Instruments, Inc.) in tapping mode employing a silicon slide substrate. A commercial silicon probe (model TESP100) with a typical resonance frequency of about 300 kHz was used to obtain the image. The mean surface roughness was calculated using nanoscope software. Scanning electron microscopy (SEM) images were obtained on a field emission scanning electron microscope (JSM 6700F JEOL, Japan) with an acceleration voltage of 5 kV. X-ray photoelectron spectrometry (XPS) spectra were recorded on an ESCALab220I-XL spectrometer (VG Scientific) with an Al Kα X-ray source (1486.6 eV). Static water contact angles were measured on a homemade instrument by a sessile drop method with 5 μL of water. The data were the average of five measurements on different places. Synthesis of PAA-Dopamine. A PAA aqueous solution (617.1 mg of a 35 wt % solution containing 3 mmol of acrylic acid units) was diluted with a 0.1 M phosphate buffer solution (30 mL), and then 173 mg (0.9 mmol) of EDC 3 HCl and 171 mg (0.9 mmol) of dopamine hydrochloride were added. The reaction solution was adjusted to 6.0 with 1.0 M NaOH and HCl and stirred for 24 h under N2 gas protection. The reaction mixture was dialyzed using a regenerated cellulose dialysis membrane (cutoff molecular weight 3500) in deionized water until dopamine in the washing solution was undetectable by UV
visible spectroscopy. The dialyzed polymer (PAA-dopamine) was lyophilized to afford a fluffy white solid. 1H NMR (D2O, water residue at 4.79 ppm as reference): δ 6.95
6.80 (d, br, 2H, C6HH2(OH)2
), 6.75 (s, br, 1H, C6H2H(OH)2
), 4.79 (D2O), 3.21 (s, br, 2H, C6H3(OH)2
CH2
CH2(NH)
C(dO)
), 2.89 (s, br, 2H, C6H3(OH)2
CH2
CH2(NH)
C(dO)
), 2.28 (s, br 1H,
NH
C(dO)
CH(CH2
)
), 1.20
2.0 (d, br, 2H,
NH
C(dO)
CH(CH2
)
). Substrate Preparation. Quartz, silicon, and glass slides were treated in boiling piranha solution (30% H2O2/98% H2SO4 1:3 v/v) for 2 h, rinsed with copious amounts of deionized water, and finally dried with a stream of N2. After this process, the substrates were hydrophilized. (Caution! Piranha solution is a very aggressive, corrosive solution, and appropriate safety precautions should be utilized.) To obtain NH2-terminated substrates, the hydrophilized substrates were immersed in a 1  10
5 M APTS/toluene solution for 6 h. The physically adsorbed APTS on the surface was removed by ultrasonic treatment in pure toluene three times. LbL Assembly of Multilayer Films. The LbL deposition process on hydrophilized or NH2-terminated substrates was controlled with an 13685

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Scheme 1. (a) Catechol Oxidative Chemistry Explored from Mussel Adhesive Proteins and (b) a Stable, Functional Multilayer Film Prepared by the Integration of Catechol Oxidative Chemistry into the Layer-by-Layer Assembly

Figure 1. 1H NMR spectrum of PAA-dopamine. polyPEGA-SH was dissolved in methanol (MeOH) to modify the asassembled multilayer films. The solvents were pre-equilibrated by bubbling with N2 to prevent the formation of disulfide bonds between thiol groups. Nine bilayer multilayer films were immersed in solutions, followed by the addition of triethylamine (final concentration of 10 mM). Here, triethylamine was used to create a basic environment for the oxidation of catechol. After 18 h of reaction, the multilayer films were rinsed completely with solvents and dried with N2. The crosslinking of thiol-modified multilayer films followed the same procedure as mentioned above.

automated device (North Tianfu Ltd., Beijing, China). The concentrations of PAA-dopamine and PAH aqueous solutions were 0.5 and 1.0 mg/mL, respectively. The pH values of PAA-dopamine and PAH solutions were adjusted to 3.5 and 7.5 with 1 M HCl and NaOH, respectively. The pH conditions were chosen because the majority of AA groups in PAA-dopamine and the amine in PAH are neutralized under these conditions. The slightly charged polymers tend to diffuse in and out, which leads to fast exponential growth during film assembly.53,54 The substrates were immersed in PAA-dopamine and PAH solutions alternately, with three rinses in water to remove the excess polymers. The deposition and rinse times were 15 and 1 min, respectively. During LbL assembly, the PAH solution was the first deposition solution for hydrophilized substrates and PAA-dopamine solution was the first deposition solution for NH2-terminated substrates. No drying step was performed in the film-deposition procedure. The final films were dried with N2 after assembly. Stabilization of Multilayer Films. The obtained multilayer films were immersed in a 1 mM NaIO4 aqueous solution for oxidizationinduced cross-linking. To cross-link the multilayer films completely, the immersion time was 2 h for 9 bilayers and 12 h for 20 bilayers. The crosslinked films were rinsed thoroughly with water and dried with N2. To investigate the stability of the resultant film, the multilayer film-covered substrates were immersed in 0.1 M NaOH or HCl or 5 M NaCl aqueous solution for UV
visible and SEM measurements. The free-standing multilayer films were obtained by the exfoliation of cross-linked 20bilayer films in a 0.1 M NaOH aqueous solution. Thiol Modification of Multilayer Films. 1-Dodecanethiol (5 mM) was dissolved in dichloromethane (DCM), and 5 mg/mL

’ RESULTS AND DISCUSSION Catechol chemistry is responsible for the strong affinity, rapid solidification, and mechanical enhancement of mussel adhesive proteins. Scheme 1a illustrates the mechanism of catechol oxidative chemistry. Catechol is readily oxidized by dissolved oxygen in the presence of oxidase or in alkaline seawater to yield o-quinone.33,34 Highly reactive o-quinone prefers to react with catechol by quinone-phenol dismutation, with amine by Michael addition and Schiff base reaction, or with thiol by Michael addition.35
37 Here, this bioinspired chemistry is integrated into a layer-by-layer assembly (Scheme 1b). Catechol is incorporated into multilayer films by the electrostatic assembly of dopaminemodified PAA (PAA-dopamine) with the polycation of PAH. The reactions of o-quinone (oxidation of catechol) with amine or residual unoxidized catechol are expected to generate strong chemical covalent bonds that improve the stability of the multilayer film. The reaction between thiol and o-quinone is designed to tailor the surface composition of the multilayer film. PAA-dopamine is synthesized by the conjugation of dopamine to PAA using carbodiimide coupling protocols. Figure 1 shows the 1H NMR spectrum of PAA-dopamine. The catechol content (mol %) in PAA-dopamine is determined from f = A/A0. Here, A is the integral area of the peaks at δ 6.95
6.60 corresponding to the amount of H in the aromatic rings of grafted catechol moieties. A0 is the integral area of the peaks at δ 2.5
1.2 representing the amount of H in the polymeric backbone. The ratio of dopamine conjugated in the resulting polymer is about 12% according to this method. The appearance of an amide peak in the FTIR spectrum of PAA-dopamine indicates the formation of an amide bond between PAA and dopamine (Supporting Information, Figure S1). The UV
visible spectrum of PAAdopamine shows a single characteristic absorbance peak of catechol at 280 nm (Figure 2a). No observation of other absorptions for the quinone oxidized form shows that conjugated catechol has not been oxidized during conjugation. LbL assembly was performed by alternating the deposition of PAH and PAA-dopamine and was monitored by UV
visible 13686

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Figure 2. (a) UV
visible spectra of a 0.2 mg/mL PAA-dopamine aqueous solution during oxidization by equimolar NaIO4. The inset shows the corresponding photographs of PAA-dopamine solution in a quartz cuvette. (b) UV
visible spectra of the oxidization process of (PAA-dopamine/ PAH)9 films in a 1 mM NaIO4 aqueous solution. The insets are the corresponding photographs of films deposited on glass slides.

Figure 3. N 1s XPS spectra of (PAA-dopamine/PAH)9 films (a) before and (b) after oxidation by 1 mM NaIO4 for 2 h.

spectroscopy and ellipsometry. Here, the deposition pH of the PAA-dopamine aqueous solution was adjusted to 3.5. Different from the dopamine-mediated continuous assembly of dopaminemodified PGA at alkaline pH,44 the dopamine groups are stable in the acidic environment and cannot self-polymerize to lead to the continuous growth of PAA-dopamine during LbL assembly. The characteristic absorbance of catechol at 280 nm and the thickness of the film increase exponentially as a function of the assembly cycle (Supporting Information, Figure S2). The assembly behavior of the PAA-dopamine/PAH multilayer film is similar to the assembly of PAA/PAH under the same pH conditions,53,54 indicating that the conjugation of dopamine on PAA did not significantly affect the LbL assembly process. Because the catechol groups have been incorporated into the multilayer film, catechol oxidative chemistry was utilized for film stabilization. NaIO4 was selected as an oxidant for the benefit of diffusion into multilayer films. Before performing the oxidation reaction in the multilayer film, we first studied the NaIO4mediated oxidation of PAA-dopamine in aqueous solution by UV
visible spectrophotometry (Figure 2a). Equimolar NaIO4 (NaIO4/catechol 1:1) was added to the PAA-dopamine aqueous solution to oxidize catechol. When NaIO4 was added to the solution, the transparent solution turned pink immediately (inset of Figure 2a) and the characteristic peak of catechol shifted from 280 to 300 nm with a new peak appearing near 470 nm (Figure 2a), indicating the oxidization of catechol to o-quinone.51 As the oxidization proceeds, the intensity of the two newly appearing peaks decreases with time (Figure 2a). The peak near

470 nm disappears after 24 h of oxidization, and a yellow/orange solution is finally obtained (inset of Figure 2a). Although the reason for the decay of the quinone characteristic peak with NaIO4 oxidization has been unclear until now, similar phenomena have been observed during the periodate-mediated oxidation of catechol derivatives.55,56 Additionally, the peak at 300 nm shifts to around 270 nm after 24 h of incubation (Figure 2a), implying the formation of some di-DOPA by quinone-phenol dismutation (Scheme 1a).57,58 The shoulder emerges at around 330 nm maybe because of the rearrangement of o-quinone.56,57 The as-assembled (PAA-dopamine/PAH)9 films were immersed in 100 mL of a 1 mM NaIO4 aqueous solution for oxidative cross-linking, and the UV
visible spectra are presented in Figure 2b. The initial oxidization of the film is similar to that of the PAA-dopamine solution. The characteristic peak of catechol shifts from 280 to 300 nm with the appearance of a new peak at 470 nm (Figure 2b), and the film turns pink immediately (inset of Figure 2b), indicating the formation of o-quinone in the multilayer film. The peaks of o-quinone disappeared, and the film color faded after 2 h of immersion (Figure 2b). According to the mechanism of catechol oxidative chemistry, both catechol groups in PAA-dopamine and amines in PAH may react with o-quinone. Here, however, we assume that the quinone-amine addition is the dominant reaction because of the excess of amine groups in films. However, the formation of di-DOPA structure in the cross-linked film cannot be absolutely excluded. These reactions result in the variations of UV
visible spectra and the cross-linking of the film. 13687

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Langmuir The N 1s XPS results also show evidence of the multilayer film cross-linking (Figure 3). The photopeak of N 1s around 399.4 and 401.6 eV corresponds to the binding energy for free and protonated primary amine groups of PAH, respectively. The peak at around 400.6 eV can be attributed to the secondary amine in PAA-dopamine (amide bond formed during dopamine conjugation, Figure 3a). In contrast to the un-cross-linked film, the proportion of the free primary amine in the cross-linked film decreased (from 32.3 to 19.7%) whereas that of the secondary amine increased (from 16.2 to 17.8%, Figure 3b). This result implies that some free primary amines are converted to secondary amines during catechol oxidative chemistry.59 In addition, the oxidative cross-linking is performed in NaIO4 aqueous solution, which is a weak acid solution (pH ∼5.3 measured by a pH monitor). Under this condition, some primary amine groups of PAH are protonated, resulting in the increase in the peak area of protonated primary amine groups. Catechol-amine and diDOPA structures are also detected in cross-linked multilayer films by FTIR spectra measurements (Supporting Information, Figure S3), which further support the cross-linking reaction in the film. Combined with the results of UV
visible, XPS, and

Figure 4. (a, c) SEM images and (b, d) AFM height images (5 μm  5 μm) of un-cross-linked and cross-linked (PAA-dopamine/PAH)9 films deposited on silicon wafers. The scale bar in the SEM images is 1 μm.

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FTIR spectra, it can be confirmed that the building blocks of PAA-dopamine and PAH in the multilayer film have been covalently coupled by oxidatively induced cross-linking. However, the obtained information is not enough to distinguish all of the chemical reactions during cross-linking because of the complication of catechol oxidative chemistry. To understand more about this chemistry, another investigation such as the application of model compounds is preferable for further study. The SEM and AFM images in Figure 4 show the morphology of multilayer films before and after cross-linking. Both films show relatively rough morphology with abundant nanoprotuberances on the surface. There is no obvious change in surface morphology during cross-linking except that the size of the nanoprotuberances slightly increased (Figure 4c,d). The root-meansquare (rms) roughness of the cross-linked film is 17.7 nm, which is almost the same as that for the un-cross-linked film (17.8 nm). The morphology maintained during cross-linking demonstrates that the catechol oxidative chemistry is soft and gentle. Taken in this sense, the method proposed here is superior to thermally induced cross-linking, which usually introduces microscale wavelike profiles on multilayer films because of the expansion and contraction of film along with temperature alteration.18,54 The potency of cross-linking for the improvement of film stability is assessed by immersing the as-assembled and crosslinked films in 0.1 M NaOH and HCl and 5 M NaCl aqueous solutions separately. To avoid the whole film being exfoliated from the substrate during the test, an APTS -modified substrate was used. This silane coupling agent can stick to the substrate by chemisorption, and its amine group can assemble with PAAdopamine and also react with the catechol group during oxidative cross-linking. The morphology change in the films after various solution etchings confirms the stability enhancement by crosslinking. The un-cross-linked films are obviously disrupted after immersion in etching solutions for 10 min (Figure 5a
c), whereas the cross-linked film maintains its integrity and no detectable morphology change is observed after treatment with etching solutions for 2 h (Figure 5d
f). In addition, UV
visible spectra were also used to quantify the mass residual of the films during solution etching. Although the characteristic peak of

Figure 5. SEM images of the un-cross-linked (top row) and cross-linked (PAA-dopamine/PAH)9 films (bottom row) after immersion in (a, d) 0.1 M NaOH, (b, e) 0.1 M HCl, and (c, f) 5 M NaCl solution. The immersion times for the un-cross-linked and cross-linked films are 10 min and 2 h, respectively. The scale bar is 1 μm. 13688

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Langmuir catechol groups at 280 nm disappeared after cross-linking (Figure 2b), the absorption of the curve still can be used to characterize the quantity of the film. This method has been used to characterize the linear growth of PVA/clay multilayer films that have smooth, monotonous absorption curves.60 In comparison to the less than 30% mass residuals of un-cross-linked films etching in acid, base, or salt aqueous solutions for 10 min, the mass residuals of the cross-linked films are higher than 94% even after 2 h (Supporting Information, Figure S4). The results demonstrate that the stability of the film in harsh environments has been dramatically enhanced after oxidative cross-linking. Stable multilayer films are often fabricated to get free-standing films without using an additional sacrificial layer. Here, crosslinked multilayer films are released from a hydrophilized substrate easily by immersion in a 0.1 M NaOH solution for 5 min. The fast liberation of the cross-linked films is due to the weakening of the interaction of PAH with the support in basic solution. The free-standing multilayer film remains intact in water (Figure 6a) and can be handled by tweezers in air after drying (inset in Figure 6a), suggesting the excellent stability of the cross-linked films. The surface and cross-sectional SEM images of the film (Figure 6b) show defect-free morphology

Figure 6. (a) Photograph of a cross-linked (PAA-dopamine/PAH)20 free-standing film in water. The inset in a is a free-standing film (2  1 cm2) in air. (b) SEM image of the free-standing (PAA-dopamine/ PAH)20 film. The insert in b is a cross-sectional image of the film. The samples for SEM measurements were exfoliated in 0.1 M NaOH solution and then transferred to silicon slides. The scale bar is 1 μm.

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on a large scale, indicating that the cross-linking occurred uniformly over the whole film. This uniform cross-linking is attributed to the homogeneous distribution and high reactivity of catechol groups with the facile diffusion of oxidant. According to the previous report on the kinetics of catechol oxidative chemistry, the reaction between thiol and o-quinone is one of the most favorable reactions, which is about 50 000 times faster than quinone-amine addition and 30 times faster than quinone-phenol coupling.37 Thus, it is possible to prepare multilayer films with both tailorable function and stability. For this purpose, the as-assembled multilayer film was first modified by thiols and then oxidatively cross-linked. This treatment order is very important. If the oxidative cross-linking is carried out in advance, then the catechol groups are more likely consumed during the oxidation process and lead to unsuccessful modification (Supporting Information, Figure S5). Two model thiol molecules, hydrophobic 1-dedecanethiol and hydrophilic polyPEGA-SH, were selected. In addition, the thiol-catechol reactions were performed in organic solvent, and an organic weak base of triethylamine was utilized to initiate catechol oxidization. We expect that most catechol-thiol addition reactions occur on surfaces rather than inside the film and that residual catechol can be used for further film cross-linking. The XPS spectra (Figure 7a,b) reveal the presence of sulfur (S 2p = 168 eV) on the modified films.34 The static water contact angle of the asassembled film is 37 ( 1°, but it increases to 60 ( 2° after modification by 1-dodecanethiol and decreases to 10 ( 1° after decoration with polyPEGA-SH (Figure 7c
e). These results indicate the successful wettability control of the multilayer films by grafting functional molecules. Functional multilayer films with good stability can be obtained by cross-linking thiol-modified films. After thiol modification, the absorbance of catechol at 280 nm in the film is almost the same (Supporting Information, Figure S6a,b), implying that most of the catechol inside the films is retained. These residual catechol groups can undergo a cross-linking reaction that endows thiol-modified films with good stability in harsh environments (Supporting Information, Figure S6c,d). Furthermore,

Figure 7. XPS spectra of the (PAA-dopamine/PAH)9 film modified with (a) 1-dodecanethiol and (b) polyPEGA-SH. (c
e) Photographs showing the wettability of the original (PAA-dopamine/PAH)9 film (c, CA = 37 ( 1°) and films modified with 1-dodecanethiol (d, CA = 60 ( 2°) and polyPEGASH (e, CA = 10 ( 1°). 13689

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Langmuir the influence of the cross-linking treatment on the morphology and wettability of thiol-modified multilayer films is examined. The morphology and surface wettability of films are not obviously different from those of the un-cross-linked ones (Supporting Information, Figure S7). These results prove that catechol oxidative chemistry is a mild cross-linking method and can effectively preserve the endowed thiol functions. Given the vast number of commercially available and facilely synthesized thiols and biomolecules containing thiols, versatile functional and stable multilayer films can be expected by this approach. Nevertheless, it should be noted that the low grafting rate of catechol may lead to unqualified thiol layers for practical applications. Therefore, building blocks containing adequate amounts of catechol are indispensable to achieving available functional, stable multilayer films.

’ CONCLUSIONS In summary, mussel adhesive protein-inspired catechol oxidative chemistry has been integrated into the layer-by-layer assembly of PAA-dopamine and PAH to improve the performance of the electrostatic multilayer films. On the basis of the high reactivity of the catechol moiety, chemical cross-linked multilayer films maintaining the initial morphology were obtained under mild oxidization conditions. UV
visible spectra, FTIR, and XPS analyses confirmed the chemical covalence between building blocks. The cross-linked multilayer films exhibit excellent stability even in extremely acidic, basic, and highly concentrated aqueous salt solutions. The cross-linked multilayer films can be released from substrates facilely by simple treatment with alkaline solution to produce intact free-standing films. More significantly, multilayer films with both tailored function and stability can be fabricated by a combination of thiol modification and oxidative cross-linking. Given the modular nature of the LbL assembly technique, the extensive building blocks, and the facile incorporation of the multifunctional catechol group, this bioinspired approach can be used to construct a range of multilayer films for advanced engineering and biomedical materials. ’ ASSOCIATED CONTENT

bS

Supporting Information. Figures S1
S7 and their related analyses. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*Fax/Tel: (+86) 10-82619667. E-mail: [email protected]; [email protected].

’ ACKNOWLEDGMENT We thank Prof. Shuguang Yang at Donghua University for helpful discussion. This work was supported by the National Natural Science Foundation of China (50821062, 51173194), the 973 Project (2007CB936400), and ICCAS (CMS-LX200913). ’ REFERENCES (1) Decher, G. Science 1997, 277, 1232–1237. (2) Tang, Z. Y.; Wang, Y.; Podsiadlo, P.; Kotov, N. A. Adv. Mater. 2006, 18, 3203–3224. (3) Zhang, X.; Chen, H.; Zhang, H. Y. Chem. Commun. 2007, 1395–1405.

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(4) Ji, J.; Fu, J. H.; Shen, J. C. Adv. Mater. 2006, 18, 1441–1444. (5) Wang, Y. D.; Joshi, P. P.; Hobbs, K. L.; Johnson, M. B.; Schmidtke, D. W. Langmuir 2006, 22, 9776–9783. (6) Boudou, T.; Crouzier, T.; Ren, K. F.; Blin, G.; Picart, C. Adv. Mater. 2009, 22, 441–467. (7) Stockton, W. B.; Rubner, M. F. Macromolecules 1997, 30, 2717–2725. (8) Yang, S. G.; Zhang, Y. J.; Zhang, X. L.; Xu, J. Soft Matter 2007, 3, 463–469. (9) Huo, F. W.; Xu, H. P.; Zhang, L.; Fu, Y.; Wang, Z. Q.; Zhang, X. Chem. Commun. 2003, 874–875. (10) Zhang, Y. J.; Cao, W. X. Langmuir 2001, 17, 5021–5024. (11) Shimazaki, Y.; Mitsuishi, M.; Ito, S.; Yamamoto, M. Langmuir 1998, 14, 2768–2773. (12) Andreeva, D. V.; Fix, D.; Mohwald, H.; Shchukin, D. G. Adv. Mater. 2008, 20, 2789–2794. (13) Andreeva, D. V.; Fix, D.; Mohwald, H.; Shchukin, D. G. J. Mater. Chem. 2008, 18, 1738–1740. (14) Li, X. F.; De Feyter, S.; Chen, D. J.; Aldea, S.; Vandezande, P.; Du Prez, F.; Vankelecom, I. F. J. Chem. Mater. 2008, 20, 3876–3883. (15) Bruening, M. L.; Dotzauer, D. M.; Jain, P.; Ouyang, L.; Baker, G. L. Langmuir 2008, 24, 7663–7673. (16) Wu, G. L.; Shi, F.; Wang, Z. Q.; Liu, Z.; Zhang, X. Langmuir 2009, 25, 2949–2955. (17) Sun, J. Q.; Wu, T.; Sun, Y. P.; Wang, Z. Q.; Zhang, X.; Shen, J. C.; Cao, W. X. Chem. Commun. 1998, 1853–1854. (18) Harris, J. J.; DeRose, P. M.; Bruening, M. L. J. Am. Chem. Soc. 1999, 121, 1978–1979. (19) Yang, S. Y.; Rubner, M. F. J. Am. Chem. Soc. 2002, 124, 2100–2101. (20) Zhang, Y. J.; Guan, Y.; Zhou, S. Q. Biomacromolecules 2005, 6, 2365–2369. (21) Schuetz, P.; Caruso, F. Adv. Funct. Mater. 2003, 13, 929–937. (22) Schneider, A.; Vodouhe, C.; Richert, L.; Francius, G.; Le Guen, E.; Schaaf, P.; Voegel, J. C.; Frisch, B.; Picart, C. Biomacromolecules 2007, 8, 139–145. (23) Dragan, E. S.; Bucatariu, F. Macromol. Rapid Commun. 2010, 31, 317–322. (24) Yu, Y.; Zhang, H.; Zhang, C. H.; Cui, S. X. Chem. Commun. 2011, 47, 929–931. (25) Such, G. K.; Quinn, J. F.; Quinn, A.; Tjipto, E.; Caruso, F. J. Am. Chem. Soc. 2006, 128, 9318–9319. (26) Kinnane, C. R.; Such, G. K.; Antequera-Garcia, G.; Yan, Y.; Dodds, S. J.; Liz-Marzan, L. M.; Caruso, F. Biomacromolecules 2009, 10, 2839–2846. (27) Such, G. K.; Tjipto, E.; Postma, A.; Johnston, A. P. R.; Caruso, F. Nano Lett. 2007, 7, 1706–1710. (28) Huang, C. J.; Chang, F. C. Macromolecules 2009, 42, 5155–5166. (29) Buck, M. E.; Zhang, J.; Lynn, D. M. Adv. Mater. 2007, 19, 3951–3955. (30) Connal, L. A.; Kinnane, C. R.; Zelikin, A. N.; Caruso, F. Chem. Mater. 2009, 21, 576–578. (31) Seo, J.; Schattling, P.; Lang, T.; Jochum, F.; Nilles, K.; Theato, P.; Char, K. Langmuir 2010, 26, 1830–1836. (32) Waite, J. H.; Tanzer, M. L. Science 1981, 212, 1038–1040. (33) Lin, Q.; Gourdon, D.; Sun, C. J.; Holten-Andersen, N.; Anderson, T. H.; Waite, J. H.; Israelachvili, J. N. Proc. Natl. Acad. Sci. U.S.A. 2007, 104, 3782–3786. (34) Lee, H.; Dellatore, S. M.; Miller, W. M.; Messersmith, P. B. Science 2007, 318, 426–430. (35) Lee, H.; Scherer, N. F.; Messersmith, P. B. Proc. Natl. Acad. Sci. U.S.A. 2006, 103, 12999–13003. (36) Xu, L. Q.; Yang, W. J.; Neoh, K.-G.; Kang, E.-T.; Fu, G. D. Macromolecules 2010, 43, 8336–8339. (37) Sagert, J.; Sun, C.; Waite, J. H. Chemical Subtleties of Mussel and Polychaete Holdfasts; Springer-Verlag: Berlin, 2006; pp 125
143. (38) Zeng, H. B.; Hwang, D. S.; Israelachvili, J. N.; Waite, J. H. Proc. Natl. Acad. Sci. U.S.A. 2010, 107, 12850–12853. 13690

dx.doi.org/10.1021/la2027237 |Langmuir 2011, 27, 13684–13691

Langmuir

ARTICLE

(39) Sever, M. J.; Weisser, J. T.; Monahan, J.; Srinivasan, S.; Wilker, J. J. Angew. Chem., Int. Ed. 2004, 43, 448–450. (40) Harrington, M. J.; Masic, A.; Holten-Andersen, N.; Waite, J. H.; Fratzl, P. Science 2010, 328, 216–220. (41) Holten-Andersen, N.; Fantner, G. E.; Hohlbauch, S.; Waite, J. H.; Zok, F. W. Nat. Mater. 2007, 6, 669–672. (42) Lee, H.; Lee, Y.; Statz, A. R.; Rho, J.; Park, T. G.; Messersmith, P. B. Adv. Mater. 2008, 20, 1619–1623. (43) Podsiadlo, P.; Liu, Z. Q.; Paterson, D.; Messersmith, P. B.; Kotov, N. A. Adv. Mater. 2007, 19, 949–955. (44) Ochs, C. J.; Hong, T.; Such, G. K.; Cui, J.; Postma, A.; Caruso, F. Chem. Mater. 2011, 23, 3141–3143. (45) Ham, H. O.; Liu, Z. Q.; Lau, K. H. A.; Lee, H.; Messersmith, P. B. Angew. Chem., Int. Ed. 2011, 50, 732–736. (46) Kang, S. M.; Rho, J.; Choi, I. S.; Messersmith, P. B.; Lee, H. J. Am. Chem. Soc. 2009, 131, 13224–13225. (47) Ryu, S.; Lee, Y.; Hwang, J. W.; Hong, S.; Kim, C.; Park, T. G.; Lee, H.; Hong, S. H. Adv. Mater. 2011, 23, 1971–1975. (48) Bilic, G.; Brubaker, C.; Messersmith, P. B.; Mallik, A. S.; Quinn, T. M.; Haller, C.; Done, E.; Gucciardo, L.; Zeisberger, S. M.; Zimmermann, R.; Deprest, J.; Zisch, A. H. Am. J. Obstet. Gynecol. 2010, 202, 85. e1
9. (49) Lee, Y.; Chung, H. J.; Yeo, S.; Ahn, C. H.; Lee, H.; Messersmith, P. B.; Park, T. G. Soft Matter 2010, 6, 977–983. (50) Shi, Y.; Gao, H.; Lu, L. C.; Cai, Y. L. Chem. Commun. 2009, 1368–1370. (51) Ott, P.; Trenkenschuh, K.; Gensel, J.; Fery, A.; Laschewsky, A. Langmuir 2010, 26, 18182–18188. (52) Ham, H. O.; Liu, Z.; Lau, K. H. A.; Lee, H.; Messersmith, P. B. Angew. Chem., Int. Ed. 2011, 50, 732–736. (53) Lu, Y. X.; Chen, X. L.; Hu, W.; Lu, N.; Sun, J. Q.; Shen, J. C. Langmuir 2007, 23, 3254–3259. (54) Ma, Y.; Sun, J. Q.; Shen, J. C. Chem. Mater. 2007, 19, 5058–5062. (55) Graham, D. G.; Jeffs, P. W. J. Biol. Chem. 1977, 252, 5729–5734. (56) Rzepecki, L. M.; Waite, J. H. Arch. Biochem. Biophys. 1991, 285, 27–36. (57) Lee, B. P.; Dalsin, J. L.; Messersmith, P. B. Biomacromolecules 2002, 3, 1038–1047. (58) Andersen, S. O.; Jacobsen, J. P.; Roepstorff, P. Tetrahedron 1980, 36, 3249–3252. (59) Beamson, G.; Briggs, D. High Resolution XPS of Organic Polymers: The Scienta ESCA300 Database; John Wiley and Sons: Chichester, U.K., 1992. (60) Podsiadlo, P.; Kaushik, A. K.; Arruda, E. M.; Waas, A. M.; Shim, B. S.; Xu, J. D.; Nandivada, H.; Pumplin, B. G.; Lahann, J.; Ramamoorthy, A.; Kotov, N. A. Science 2007, 318, 80–83.

13691

dx.doi.org/10.1021/la2027237 |Langmuir 2011, 27, 13684–13691