Pickering Stabilized Peptide Gel Particles as ... - ACS Publications

Article pubs.acs.org/Langmuir

Pickering Stabilized Peptide Gel Particles as Tunable Microenvironments for Biocatalysis Gary Scott, Sangita Roy, Yousef M. Abul-Haija, Scott Fleming, Shuo Bai,* and Rein V. Ulijn* WestCHEM, Department of Pure and Applied Chemistry, University of Strathclyde, Glasgow G1 1XL, U.K. S Supporting Information *

ABSTRACT: We demonstrate the preparation of peptide gel microparticles that are emulsified and stabilized by SiO2 nanoparticles. The gels are composed of aromatic peptide amphiphiles 9fluorenylmethoxycarbonyldiphenylalanine (Fmoc-FF) coassembled with Fmoc-amino acids with different functional groups (S: serine; D: aspartic acid; K: lysine; and Y: tyrosine). The gel phase provides a highly hydrated matrix, and peptide self-assembly endows the matrix with tunable chemical environments which may be exploited to support and stabilize proteins. The use of Pickering emulsion to stabilize these gel particles is advantageous through avoidance of surfactants that may denature proteins. The performance of enzyme lipase B immobilized in pickering/gel microparticles with different chemical functionalities is investigated by studying transesterification in heptane. We show that the use of Pickering particles enhances the performance of the enzyme, which is further improved in gel-phase systems, with hydrophilic environment provided by Fmoc-FF/S giving rise to the best catalytic performance. The combination of a tunable chemical environment in gel phase and Pickering stabilization described here is expected to prove useful for areas where proteins are to be exploited in technological contexts such as biocatalysis and also in other areas where protein performance and activity are important, such as biosensors and bioinspired solar fuel devices.

■

INTRODUCTION Peptides are versatile building blocks for the production, via molecular self-assembly, of nanostructures for a range of applications, such as biosensors, controlled release systems, cell scaffolds, biocatalysis, and nanotemplating.1−4 Intermolecular noncovalent interactions such as hydrogen bonding, ionic interactions, and π−π stacking between peptide components can lead to the formation of fibers, which in turn entangle to form 3D gel networks. The porous structure and aqueous interior of these hydrogels may provide a suitable environment for encapsulation and immobilization of proteins for technological applications as it mimics aspects of the protein’s native surroundings.5−8 Aromatic peptide amphiphiles have become increasingly popular as building blocks for the production of functional fibrous gels.9−14 It has been shown that coassembly building blocks may be used to incorporate functional (e.g., bioactive or energy transfer) peptide components within hydrogel structures.15−19 As previously demonstrated, simple chemical functionality (e.g., OH, NH2, COOH) can be incorporated by coassembly of a structural peptide derivative (e.g., Fmoc-FF) with amino acid derivatives (Fmoc-X) with variable side chains using ionic induced20 or biocatalytic self-assembly.21 These coassembled materials combine functional and structural components and are potentially suited as tunable matrices for protein immobilization that may be exploited in biocatalysis, which is the subject of the current study. © 2013 American Chemical Society

In order to produce stable coassembled gel microparticles, we investigated the possibility to combine supramolecular selfassembly with Pickering emulsion. Pickering emulsions, stabilized by solid particles adsorbed at the liquid−liquid interface, provide an emulsion technique which gives rise to particles that are more stable than those produced by using surfactants.22−24 In addition, the absence of amphiphilic surfactants, which have surface-active properties and potential destabilizing effects on proteins, could be beneficial to the encapsulation and immobilization of biocatalysts.25−30 More generally, the use of microparticles as protein immobilization matrices enhances (i) the ratio of surface and volume, (ii) contact space between biocatalysts and reactants, and (iii) product transfer, which contribute to the activity, stability, and reusability of biocatalysts.31−35 In this work, we first combined the advantages of peptide self-assembly and Pickering emulsion to make gel microparticles with chemically tunable environments to immobilize lipase enzymes as a model system for biocatalysis in organic media. We used hydrophobic SiO2 nanoparticles (NPs) to emulsify the peptide precursor solution. A series of hydrogel compositions were prepared by combining 9-fluorenylmethoxycarbonyl (Fmoc)-diphenylalanine (FF) and n-protected Fmoc Received: September 6, 2013 Revised: October 21, 2013 Published: October 22, 2013 14321

dx.doi.org/10.1021/la403448s | Langmuir 2013, 29, 14321−14327

Langmuir

Article

Scheme 1. Cartoon of Pickering Emulsion Droplet Stabilized by SiO2 NPs and Gelled by Functionalized Peptide Fibers for Protein Immobilization

methane (DCM) and trifluoroacetic acid (TFA) overnight. The excess TFA was removed by the crystallization of the products from ether. Final traces of TFA were removed by continuous washing with ether. Preparation of Hydrophobic SiO2 NPs. SiO2 NPs were synthesized by the Stöber method.36 In 1 L flask, 345 mL of ethanol, 26 mL of water, and 9 mL of ammonia were mixed by a magnetic stirrer at room temperature. 20 mL of TEOS was added rapidly to above solution while stirring continued. The reaction mixture was then stirred at room temperature overnight. After that the solution became turbid. The resulting SiO2 NPs were collected by centrifugation at 8000 rcf for 5 min. After decanting the supernatants, the SiO2 NPs were dispersed in ethanol. The washing cycle was repeated three times to remove the excess TEOS and ammonia. For hydrophobization of NPs surface, TMODS (40 μL) was added into SiO2 NPs dispersion in ethanol (2 mL, 3 wt %), followed by heating at 60 °C overnight. After washing with ethanol three times, the resulting SiO2 NPs were transferred from ethanol to THF and finally heptane. Preparation of Peptide Hydrogels in Bulk. Fmoc-FF (10 mM) and Fmoc-X (S, D, K or Y, 10 mM) powders were dispersed in 0.8 mL of water. After adding 90 μL of NaOH (0.5 M), the powders were completely dissolved by alternate sonicating and vortexing. 0.5 M HCl was added drop by drop to adjust the pH value to 7.8. 100 μL CaCl2 solutions (0.1 M) were added into the above solution. After several hours, the peptide hydrogels were formed. Immobilization of Enzymes in Peptide Pickering Emulsion Droplets. Buffer solution (pH 8) was prepared by dissolving 94 mg of NaH2PO4·H2O and 2.5 g of Na2HPO4·7H2O in 100 mL of water. 100 μL CalB buffer solutions (125 mg/mL) were mixed with the gelator precursor solution mentioned above. 200 μL of the resulting enzyme/ gelator mixture was added to dispersion of hydrophobic SiO2 NPs in heptane (4 mL), followed by homogenization using a homogenizer (VWR VDI 12 S2, 125 W) for 10 s. The emulsions were stored in fridge at 2−8 °C overnight to form the gel particles. Catalytic Performance of Free and Immobilized Enzymes. First, 200 μL of 1-octanol and octanoic acid were added into a 4 mL Pickering emulsion solution to initiate the esterification reaction, which was carried out on a magnetic stirrer with 200 rpm at 30 °C. Every 5 min, 10 μL of supernatants separated from the emulsions by centrifugation at 1000 rcf for 1 min was diluted within 990 μL of chloroform and analyzed by gas chromatography (GC) to determine the product concentration and reaction rate. Second, to determine the specific activity, the same reaction mentioned above was carried out for 10 min. After 10 min, the emulsions were removed by centrifugation at

amino acids, serine (S, with side chain R = CH2OH), aspartic acid (D, with side chain R = CH2COOH), lysine (K, with side chain R = (CH2)4NH2) and tyrosine (Y, with side chain R = CH2C6H4OH) to gelate the microparticles and introduce chemical functionality into the networks via coassembly (Scheme 1).20 The enzyme lipase B from Candida Antarctica (CalB) is then immobilized into these structures, and the catalysis of the esterification reaction of octanol and octanoic acid in heptane is used to investigate how the catalytic performance is influenced by the gel matrix.

■

EXPERIMENTAL SECTION

Materials. Tetraethyl orthosilicate (TEOS) (≥99.0%), trimethoxy(octadecyl)silane (TMODS) (90%), ammonium hydroxide solution (28.0−30.0%), phenylalanine tert-butyl ester hydrochloride (≥99.0%), 1-octanol (≥99.0%), octanoic acid (≥98.0%), lipase B from Candida Antarctica (CalB) (10.8 U/mg), fluorescein isothiocyanate−dextran (average Mw 70 000, FITC:glucose = 1:250), sodium phosphate monobasic monohydrate (ACS reagent, 98.0%−102.0%), and sodium phosphate dibasic heptahydrate (ACS reagent, 98.0%−102.0%) were purchased from Sigma-Aldrich and used as received. Fmoc-Ser-OH, Fmoc-Asp-OH, Fmoc-Lys-OH, Fmoc-Tyr-OH, and Fmoc-Phe-OH were purchased from BACHEM. 2-(1H-Benzotriazole-1-yl)-1,1,3,3tetramethyluronium hexafluorophosphate (HBTU) was purchased from Novabiochem. All other chemicals and solvents were purchased from Sigma-Aldrich and used without further purification. Synthesis of Fmoc-FF. To a stirred solution of Fmoc protected Lphenylalanine (1 equiv) in dimethylformamide (DMF), coupling agent HBTU (1.2 equiv) was added and stirred for 15 min. Hydrochloride salt of the C-terminal protected tert-butyl ester of L-phenylalanine (1.2 equiv) was added to the reaction mixture followed by addition of an organic base, diisopropylethylamine (2.5 equiv). The coupling reaction was stirred overnight. Solvent was evaporated, and the remains were dissolved in ethyl acetate. The ethyl acetate layer was washed with 1 N sodium bicarbonate solution as well as 1 N hydrochloric acid solution to remove excess acid or base, followed by saturated brine solution. The organic layer was dried over magnesium sulfate, and the solvent was evaporated. The final products were purified by column chromatography using 230−400 mesh silica gels as stationary phase and chloroform−methanol mixture as eluent. Then, deprotection of the tert-butyl group was carried out in 50:50 mixtures of dichloro14322

dx.doi.org/10.1021/la403448s | Langmuir 2013, 29, 14321−14327

Langmuir

Article

1000 rcf for 1 min. The product concentration was determined by diluting 10 μL of supernatants with 990 μL of chloroform and analyzed by GC. Characterization. Fluorescence emission spectra were measured on a Jasco FP-6500 spectrofluorometer with light measured orthogonally to the excitation light, at a scanning speed of 100 nm min−1. The excitation wavelength was 280 nm, and emission data were recorded in the range between 300 and 600 nm. The spectra were measured with a bandwidth of 3 nm with a medium response and a 3 nm data pitch. The mechanical properties of hydrogels were measured out by dynamic frequency sweep experiments on strain-controlled rheometer (Kinexus Pro rheometer) using parallel-plate geometry (20 mm diameter) with a 0.50 cm gap. The experiments were performed at 25 °C, and this temperature was controlled throughout the experiment using an integrated electrical heater. Additional precautions were taken to minimize solvent evaporation and to keep the sample hydrated: a solvent trap was used, and the internal atmosphere was kept saturated. To ensure that the measurements were made in the linear viscoelastic regime, an amplitude sweep was performed and the results showed no variation in elastic modulus (G′) and viscous modulus (G″) up to a strain of 1%. The dynamic modulus of the hydrogel was measured as a frequency function, where the frequency sweeps were carried out between 0.1 and 100 Hz. The measurements were repeated three times to ensure reproducibility. The atomic force microscopy (AFM) images were obtained by scanning the mica surface in air under ambient conditions using a Veeco diINNOVA scanning probe microscope (VEECO/BRUKER, Santa Barbara, CA) operated in tapping mode. 20 μL of gels prepared using the standard procedure described above was diluted to a total volume of 100 μL solution in deionized water. Then it was placed on a trimmed and freshly cleaved mica sheet (G250-2 mica sheets 1 in. × 1 in. × 0.006 in.; Agar Scientific Ltd., Essex, UK) attached to an AFM support stub and left to air-dry overnight in a dust-free environment. The AFM scans were taken at 512 × 512 pixels resolution. Typical scanning parameters were as follows: tapping frequency 308 kHz, integral and proportional gains 0.3 and 0.5, respectively, set point 0.5− 0.8 V, and scanning speed 1.0 Hz. The silica nanoparticles were characterized by transmission electron microscopy (TEM, LEO 912 energy filtering transmission electron microscope with 14 bit/2 Proscan CCD camera). The structure of Pickering emulsion droplets was determined by a scanning electron microscope (SEM, Hitachi S-3000N). The peptide microparticles were characterized by fluorescence microscopy. The devices were mounted on an inverted microscope (AXIO Observer A1, Zeiss), and images were acquired using an EMCCD LucaR camera (Andor Technologies). Images (using brightfield and fluorescence microscopy excited at 365 nm) were acquired using Zeiss ×5, ×10, ×20 dry objective and the appropriate filter set (filter set 10, Zeiss) for the fluorophore being imaged. Data were processed and analyzed using Matlab and ImageJ. The catalysis experiment was carried out by gas chromatography (GC, Agilent Technologies 7890A GC system).

The intensity decrease of the emission peak at 325 nm (representing emission of free Fmoc groups) was assigned to the formation of the fluorenyl π−π interactions in the formed fibres (Figure S1). Atomic force microscopy (AFM) was used to investigate the different fibrous morphologies of these materials including the diameter, the length, and the topography (Figure 1). The fibers

Figure 1. AFM images of Fmoc-FF/S, Fmoc-FF/D, Fmoc-FF/K, and Fmoc-FF/Y fibers after gelation and photographs of gels (insets). Scale bar is 1 μm.

in all the gels were densely packed to three-dimensional networks. Differences in average fiber length were apparent with Fmoc-FF/S fibers being substantially longer and FmocFF/K being wider compared to the others. Figure 2 and Figure S2 show the linear viscoelastic responses of the materials. The average storage moduli (G′) over the

■

Figure 2. Oscillatory rheology of the elastic modulus (G′) of FmocFF/S, Fmoc-FF/D, Fmoc-FF/K, and Fmoc-FF/Y hydrogels.

RESULTS AND DISCUSSION First, coassembled gels were formed and characterized in bulk by combination of 10 mM gelator Fmoc-FF and 10 mM functional components Fmoc-X in solution at pH 7.8, similar to the approach published previously.20 Upon addition of a 0.1 mM solution of CaCl2, gelation occurred, giving rise to opaque gels of Fmoc-FF/S (presenting CH2−OH functionality), FmocFF/Y (presenting a hydrophobic environment through CH2C6H4OH functionality), negative charged Fmoc-FF/D (CH 2 −COO − ), and positive charged Fmoc-FF/K ((CH2)4NH3+) (Scheme 1) within 1 h. Fluorescence spectroscopy was used to observe a spectroscopic change during gelation due to changes in fluorenyl environment with the excitation wavelength at 280 nm.21,37

frequency range of 0.1−20 Hz were 10.9 kPa (Fmoc-FF/D) > 10.0 kPa (Fmoc-FF/S) > 7.7 kPa (Fmoc-FF/Y) > 6.4 kPa (Fmoc-FF/K). The storage modulus (G′) exceed the loss modulus (G″) by a factor of 24.3 (Fmoc-FF/D), 11.9 (FmocFF/S), 8.0 (Fmoc-FF/K), and 7.6 (Fmoc-FF/Y), demonstrating that these materials formed viscoelastic gels. Next, we moved onto formulating these functional gels into Pickering emulsions. SiO2 NPs were synthesized by the Stöber method.36 Transmission electron microscopy (TEM) (Figure S3) and dynamic light scattering (DLS) showed that the average diameter was around 120 nm. 14323

dx.doi.org/10.1021/la403448s | Langmuir 2013, 29, 14321−14327

Langmuir

Article

Figure 3. (a) Fluorescence microscopy images of gelled Fmoc-FF/S, Fmoc-FF/D, Fmoc-FF/K, and Fmoc-FF/Y droplets containing FITC-labeled dextran, stabilized by SiO2 NPs in heptane (scale bar is 50 μm). The insets are histograms of the size distribution of the gelled Fmoc-FF/S, FmocFF/D, Fmoc-FF/K, and Fmoc-FF/Y droplets from fluorescence microscopy images. (b) SEM micrograph of gelled Fmoc-FF/S droplets stabilized by SiO2 NPs (scale bar is 2 μm).

by directly introducing the substrates into the emulsions (Scheme 2). We compared this system (“Native CalB”) with a Pickering stabilized solution of CalB, which in turn was compared to Pickering gel emulsions with different chemical functionality. The control “Native CalB” experiment was carried out by directly introduced CalB buffer solution into heptane containing reactants on a magnetic stirrer with 200 rpm at 30 °C. High-performance liquid chromatography (HPLC) was used to check whether the gels were chemically inert with respect to hydrolysis by CalB. A solution of Fmoc-FF (10 mM) containing CalB (12 mg/mL) was shown to result in 3.6% hydrolysis after 24 h. Within the time scale of the experiments Fmoc-FF was degraded, which is not expected to affect the gel matrix and functionality within the time scale of the experiments. First, we measured the concentration of products catalyzed by each of these biocatalytic Pickering/peptide gel microparticles preparations by GC every 5 min to determine the reaction rates. Figure 4 (inset) shows a linear correlation within

To stabilize water/oil emulsions, SiO2 NPs were functionalized with hydrophobic trimethoxy(octadecyl)silane (TMODS) and transferred from water to ethanol, THF, and finally heptane stepwise. The resulting hydrophobic SiO2 NPs were used to emulsify the droplets of aqueous solution containing peptide precursors and CalB in heptane. By dissolving 1 mg/mL fluorescein isothiocyanate (FITC)-labeled dextran in aqueous solution, the gelled peptide microparticles were subsequently visualized by fluorescent microscopy (Figure 3a). The size of microparticles was in the range of 4−15 μm, and the average sizes were 7.6 μm (Fmoc-FF/S), 8.6 μm (Fmoc-FF/D), 8.4 μm (Fmoc-FF/K), and 7.7 μm (Fmoc-FF/ Y). Scanning electron microscopy (SEM) showed that the SiO2 NPs were close packed onto the surface of the microparticles (Figure 3b). It should be noted that due to the drying process for SEM sample preparation, the gelled droplets collapsed and could not maintain their rounded shape. Finally, the catalytic performance of CalB was investigated using the esterification of octanol and octanoic acid in heptane 14324

dx.doi.org/10.1021/la403448s | Langmuir 2013, 29, 14321−14327

Langmuir

Article

(Fmoc-FF/Y, 402 μmol min−1 mg−1) times compared to native CalB in a liquid biphasic system. No clear correlation exists between enzyme activity and G′ or size of Pickering particles while there is a correlation between activity and polarity of functional groups, which indicated that the gel network and functionality of peptide sequences could obviously affect the catalytic performance of CalB. Compared with the buffer Pickering droplets without peptide gelators, Fmoc-FF/S and Fmoc-FF/D peptide networks enhanced the activity of CalB, but Fmoc-FF/K and Fmoc-FF/Y networks reduced the activity. The hydrophobicity parameters log P (partition coefficient) are 2.01 (Fmoc-FF/S) < 2.08 (FmocFF/D) < 2.6 (Fmoc-FF/K) < 4.15 (Fmoc-FF/Y)the same order of decreasing catalytic rates. These results imply that the dominant effect is the hydrophilic/hydrophobic interactions between the enzymes and hydrogel networks. Since the esterification of hydrophobic reactants catalyzed by hydrophilic CalB at the oil/liquid interface, the catalytic performance is likely to depend on the amount of enzyme adsorbed on the interfaces. Since CalB has a largely hydrophilic surface, we propose that this enables it to more freely diffuse through the hydrophilic Fmoc-FF/S and Fmoc-FF/D peptide networks to the interface, leading to higher activity compared with hydrophobic Fmoc-FF/K and Fmoc-FF/Y networks. The exact nature of these stabilizing effects will be addressed in the future, which will involve a systematic study of proteins with different ionic and hydrophobic properties in gel matrices. To determine the reusability of enzymes immobilized within peptide particles, the same catalytic reaction was carried out for 10 min. After that, the emulsions were removed by centrifugation and redispersed in fresh heptane solution containing reactants. The reaction cycle was repeated for four times. Figure 5 showed the reusability of CalB immobilized

Scheme 2. Cartoon of Pickering Emulsion Droplet Stabilized by SiO2 NPs and Gelled by Peptide Fibers for Enzymatic Catalysis of Organic Reactions in Organic Media

Figure 4. Chart of the normalized activity of native CalB, CalB in Pickering emulsion buffer droplets, and CalB immobilized in gelled Fmoc-FF/S, Fmoc-FF/D, Fmoc-FF/K, and Fmoc-FF/Y Pickering emulsion droplets stabilized by SiO2 NPs. The normalized activity values are included. The inset is a plot of the reaction rates of esterification catalyzed by native CalB and CalB immobilized in FmocFF/S, Fmoc-FF/D, Fmoc-FF/K, and Fmoc-FF/Y hydrogel microparticles in 30 min. Figure 5. Plot of the activity of CalB immobilized in Fmoc-FF/S (black) and Fmoc-FF/D (red) hydrogel microparticles in 10 min versus the number of reuses.

the first 30 min (up to 70% conversion). Subsequently, the catalytic performances of native and immobilized enzymes in 10 min were carried out and compared. Figure 4 shows that, compared with native CalB, the activity of CalB immobilized in buffer (i.e., no gel) containing Pickering emulsion droplets increased 2.36 times (from 220 to 520 μmol min−1 mg−1). In the native CalB experiment, the aqueous CalB containing droplets tend to coalesce,38 leading to small interface area for enzymatic catalysis. This coalescence was avoided in the emulsion system, which is thought to contribute to the observed rate enhancement. Upon comparison with the gelled Pickering emulsions, the catalytic activity of CalB increased considerably. The enhancement in activity was 3.9 (Fmoc-FF/ S, 866 μmol min−1 mg−1), 2.5 (Fmoc-FF/D, 559 μmol min−1 mg−1), 1.9 (Fmoc-FF/K, 427 μmol min−1 mg−1), and 1.8

within the best performing matrices, Fmoc-FF/S and FmocFF/D. The results indicated that there was good retention of activity over four cycles, which may be of interest for the practical use of these systems.

■

CONCLUSIONS In conclusion, we demonstrated a novel approach using SiO2 NPs to emulsify the peptide gel microparticles which provides chemically tunable environments which may be used for protein immobilization and biocatalysis. This approach combines the advantages of peptides self-assembly and Pickering emulsions. Pickering emulsion droplets significantly enhanced the interfacial area of the hydrophobic substrates in 14325

dx.doi.org/10.1021/la403448s | Langmuir 2013, 29, 14321−14327

Langmuir

Article

(12) Hirst, A. R.; Roy, S.; Arora, M.; Das, A. K.; Hodson, N.; Murray, P.; Marshall, S.; Javid, N.; Sefcik, J.; Boekhoven, J.; van Esch, J. H.; Santabarbara, S.; Hunt, N. T.; Ulijn, R. V. Biocatalytic induction of supramolecular order. Nat. Chem. 2010, 2, 1089−1094. (13) Yang, Z.; Gu, H.; Fu, D.; Gao, P.; Lam, J. K.; Xu, B. Enzymatic formation of supramolecular hydrogels. Adv. Mater. 2004, 16, 1440− 1444. (14) Cheng, G.; Castelletto, V.; Jones, R. R.; Connon, C. J.; Hamley, I. W. Hydrogelation of self-assembling RGD-based peptides. Soft Matter 2011, 7, 1326−1333. (15) Zhou, M.; Smith, A. M.; Das, A. K.; Hodson, N. W.; Collins, R. F.; Ulijn, R. V.; Gough, J. E. Self-assembled peptide-based hydrogels as scaffolds for anchorage-dependent cells. Biomaterials 2009, 30, 2523− 2530. (16) Hirst, A. R.; Smith, D. K. Two-component gel-phase materials Highly tunable self-assembling systems. Chem.Eur. J. 2005, 11, 5496−5508. (17) Wang, C.; Guo, Y.; Wang, Z.; Zhang, X. Superamphiphiles based on charge transfer complex: Controllable hierarchical self-assembly of nanoribbons. Langmuir 2010, 26, 14509−14511. (18) Hartgerink, J. D.; Beniash, E.; Stupp, S. I. Molecular selfassembly into one-dimensional nanostructures. Self-assembly and mineralization of peptide-amphiphile nanofibers. Science 2001, 294, 1684−1688. (19) Nalluri, S. K. M.; Ulijn, R. V. Discovery of energy transfer nanostructures using gelation-driven dynamic combinatorial libraries. Chem. Sci. 2013, 4, 3699−3705. (20) Jayawarna, V.; Richardson, S. M.; Hirst, A. R.; Hodson, N. W.; Saiani, A.; Gough, J. E.; Ulijn, R. V. Introducing chemical functionality in Fmoc-peptide gels for cell culture. Acta Biomater. 2009, 5, 934−943. (21) Abul-Haija, Y. M.; Roy, S.; Frederix, P. W. J. M.; Javid, N.; Jayawarna, V.; Ulijn, R. V. Biocatalytically triggered co-assembly of two-component core/shell nanofibers. Small 2013, DOI: 10.1002/ smll.201301668. (22) Pickering, S. U. CXCVI-Emulsions. J. Chem. Soc. Trans. 1907, 91, 2001−2021. (23) Binks, B. P. Particles as surfactants - similarities and differences. Curr. Opin. Colloid Interface Sci. 2002, 7, 21−41. (24) Chen, T.; Colver, P. J.; Bon, S. A. F. Organic-inorganic hybrid hollow spheres prepared from TiO2-stabilized Pickering emulsion polymerization. Adv. Mater. 2007, 19, 2286−2289. (25) Khmelnitsky, Y. L.; Hilhorst, R.; Visser, A.; Veeger, C. Enzyme inactivation and protection during entrapment in reversed micelles. Eur. J. Biochem. 1993, 211, 73−77. (26) Sheldon, R. A.; Van Pelt, S. Enzyme immobilisation in biocatalysis: Why, what and how. Chem. Soc. Rev. 2013, 42, 6223− 6235. (27) Carrea, G.; Riva, S. Properties and synthetic applications of enzymes in organic solvents. Angew. Chem., Int. Ed. 2000, 39, 2226− 2254. (28) Klibanov, A. M. Improving enzymes by using them in organic solvents. Nature 2001, 409, 241−246. (29) Schmid, A.; Dordick, J. S.; Hauer, B.; Kiener, A.; Wubbolts, M.; Witholt, B. Industrial biocatalysis today and tomorrow. Nature 2001, 409, 258−268. (30) Oldfield, C.; Freedman, R. B.; Robinson, B. H. Enzyme hyperactivity in AOT water-in-oil microemulsions is induced by “lone” sodium counterions in the water-pool. Faraday Discuss. 2005, 129, 247−263. (31) Kumar, R. K.; Li, M.; Olof, S. N.; Patil, A. J.; Mann, S. Artificial cytoskeletal structures within enzymatically active bio-inorganic protocells. Small 2013, 9, 357−362. (32) Wang, Z.; van Oers, M. C. M.; Rutjes, F. P. J. T.; van Hest, J. C. M. Polymersome colloidosomes for enzyme catalysis in a biphasic system. Angew. Chem., Int. Ed. 2012, 51, 10746−10750. (33) Bai, S.; Wu, C.; Gawlitza, K.; von Klitzing, R.; AnsorgeSchumacher, M. B.; Wang, D. Y. Using hydrogel microparticles to transfer hydrophilic nanoparticles and enzymes to organic media via stepwise solvent exchange. Langmuir 2010, 26, 12980−12987.

contact with enzymes, the mass transfer of the products, and accessibility of the catalyst in reaction system. Self-assembly of peptides endows the hydrogel networks with biocompatible environment and different functional groups which interact with enzymes. A combination of these advantages considerably enhances the catalytic performance of enzymes, providing an approach to facilitate the use of proteins in a variety of technological applications.

■

ASSOCIATED CONTENT

S Supporting Information *

Fluorescence spectra of peptides before and after gelation, rheology spectra of elastic modulus (G′) and viscous modulus (G″) of peptides hydrogels, and TEM image of silica nanoparticles. This material is available free of charge via the Internet at http://pubs.acs.org.

■

AUTHOR INFORMATION

Corresponding Authors

*E-mail [email protected] (S.B.). *E-mail [email protected] (R.V.U.). Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We thank the Leverhulme Trust, ERC (Starting Grant EMERgE) and Marie Curie initial training network ReAd (289723). We thank Dr. Jugal Sahoo and Dr. Nadeem Javid for TEM and DLS measurements and Dr. Michele Zagnoni and Barbara Schlicht for microscopy measurements.

■

REFERENCES

(1) Yan, X.; Li, J.; Möhwald, H. Self-assembly of hexagonal peptide microtubes and their optical waveguiding. Adv. Mater. 2011, 23, 2796− 2801. (2) Palmer, L. C.; Stupp, S. I. Molecular self-assembly into onedimensional nanostructures. Acc. Chem. Res. 2008, 41, 1674−1684. (3) Yang, Z.; Liang, G.; Xu, B. Enzymatic hydrogelation of small molecules. Acc. Chem. Res. 2008, 41, 315−326. (4) Boyle, A. L.; Woolfson, D. N. De novo designed peptides for biological applications. Chem. Soc. Rev. 2011, 40, 4295−4306. (5) Gao, Y.; Zhao, F.; Wang, Q.; Zhang, Y.; Xu, B. Small peptide nanofibers as the matrices of molecular hydrogels for mimicking enzymes and enhancing the activity of enzymes. Chem. Soc. Rev. 2010, 39, 3425−3433. (6) Kiyonaka, S.; Sada, K.; Yoshimura, I.; Shinkai, S.; Kato, N.; Hamachi, I. Semi-wet peptide/protein array using supramolecular hydrogel. Nat. Mater. 2004, 3, 58−64. (7) Hoffman, A. Hydrogels for biomedical applications. Adv. Drug Delivery Rev. 2002, 54, 3−12. (8) Peppas, N.; Huang, Y.; Torres-Lugo, M.; Ward, J.; Zhang, J. Physicochemical, foundations and structural design of hydrogels in medicine and biology. Annu. Rev. Biomed. Eng. 2000, 2, 9−29. (9) Adams, D. J.; Topham, P. D. Peptide conjugate hydrogelators. Soft Matter 2010, 6, 3707−3721. (10) Chen, L.; Morris, K.; Laybourn, A.; Elias, D.; Hicks, M. R.; Rodger, A.; Serpell, L.; Adams, D. J. Self-assembly mechanism for a naphthalene-dipeptide leading to hydrogelation. Langmuir 2010, 26, 5232−5242. (11) Orbach, R.; Mironi-Harpaz, I.; Adler-Abramovich, L.; Mossou, E.; Mitchell, E. P.; Forsyth, V. T.; Gazit, E.; Seliktar, D. The rheological and structural properties of fmoc-peptide-based hydrogels: The effect of aromatic molecular architecture on self-assembly and physical characteristics. Langmuir 2012, 28, 2015−2022. 14326

dx.doi.org/10.1021/la403448s | Langmuir 2013, 29, 14321−14327

Langmuir

Article

(34) Wu, C.; Bai, S.; Ansorge-Schumacher, M. B.; Wang, D. Y. Nanoparticle cages for enzyme catalysis in organic media. Adv. Mater. 2011, 23, 5694−5699. (35) Bai, S.; Debnath, S.; Gibson, K.; Schlicht, B.; Bayne, L.; Zagnoni, M.; Ulijn, R. V. Biocatalytic self-assembly of nanostructured peptide microparticles using droplet microfluidics. Small 2013, DOI: 10.1002/ smll.201301333. (36) Stö ber, W.; Fink, A.; Bohn, E. Controlled growth of monodisperse silica spheres in micron size range. J. Colloid Interface Sci. 1968, 26, 62−69. (37) Sadownik, J. W.; Leckie, J.; Ulijn, R. V. Micelle to fibre biocatalytic supramolecular transformation of an aromatic peptide amphiphile. Chem. Commun. 2011, 47, 728−730. (38) Ansorge-Schumacher, M. B. Two-phase systems with solidified water phases - Tools for technical use of sensitive catalysts. Mini-Rev. Org. Chem. 2007, 4, 243−245.

14327

dx.doi.org/10.1021/la403448s | Langmuir 2013, 29, 14321−14327