Biopolymer-Encapsulated Protein Microcapsules Spontaneously

Article pubs.acs.org/Biomac

Biopolymer-Encapsulated Protein Microcapsules Spontaneously Formed at the Ionic Liquid−Water Interface Masa-aki Morikawa,*,†,§ Aki Takano,† Shuichi Tao,† and Nobuo Kimizuka*,†,‡,§ †

Department of Chemistry and Biochemistry, Graduate School of Engineering and ‡Center for Molecular Systems (CMS), Kyushu University § JST CREST, 744 Moto-oka, Nishi-ku, Fukuoka 819-0395, Japan S Supporting Information *

ABSTRACT: Aqueous microdroplets introduced in ionic liquids (ILs) provide unique interfaces where surface-modified protein microcapsules are spontaneously formed at systemic temperature. The susceptibility of proteins to form microcapsules at the water-IL microinterface depends on protein species and is related to the number of charged residues exhibited on protein surfaces. When both of the capsuleforming (host) proteins and guests biopolymers such as nucleic acids or enzymes are introduced in the aqueous microdroplets, microcapsules are formed selectively from host proteins while the guest biopolymers remain encapsulated in the aqueous pool. Microcapsules formed in the IL phase are facilely extracted to aqueous phase after consecutive cross-linking and surface modification reactions, and the whole processes can be done in one pot. Enzymes confined in the inner water phase of aqueous protein microcapsules showed innate activity, as visualized by site-specific fluorescence detection using confocal laser scanning microscopy (CLSM). The present IL-water interfacial synthesis of protein microcapsules eliminates the use of volatile organic solvents or solid colloid templates, which creates a much-coveted solution to existing technologies.

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INTRODUCTION Microcapsules formed from proteins are important class of biodegradable vehicles that can be applied to various fields such as bioreactors, controlled release technology for pharmaceuticals, and therapy.1 Because of the current strong demands in biomacromolecular drugs, the development of methodologies to quantitatively encapsulate guest biomolecules in protein microcapsules holds the key to make a leap forward in these areas. Although encapsulation of genome nucleic acids in supramolecular shells of viral capsid proteins has been developed for the purpose of viral cell transformation,2 it suffers from toxicity and difficulty in large-scale production. To make up for such shortcomings, layer-by-layer adsorption of proteins on sacrificial colloid templates has been devised,3−5 yet it requires time-consuming multistep adsorption procedures and removal of sacrificial colloidal templates, although calcium carbonate microparticles could be etched under mild condition by treating with EDTA.3 Self-assembly of amphiphiles or amphiphilic block copolymers give vesicle structures,6−8 where an essential drawback of these molecular assemblies lies in the poor efficiency in encapsulating guest molecules. Meanwhile, microemulsions provide liquid−liquid interfaces that can be utilized as templates to prepare microcapsules. Protein microcapsules have been prepared by using oil-in-water emulsions;9−12 however, inner oil droplets are not suitable to dissolve water-soluble guest biopolymers. Although it is © 2012 American Chemical Society

basically possible to introduce water-soluble guest biomacromolecules in the water pool of surfactant-based reverse micelles,13 there exist no emulsion templating methods to prepare guest-encapsulated protein microcapsules that can be successively extracted to the aqueous phase.14 Obviously, a more simple and rational methodology is required. A straightforward solution to this issue is to form protein microcapsules selectively from binary aqueous mixtures of capsule-forming (host) proteins and guest biopolymers so that guest-encapsulated protein microcapsules are obtained in one pot. We show herein that the interface formed between ionic liquids (ILs) and water provides solution to these issues. ILs have been attracting broad interests because of their unique properties such as negligible vapor pressure, low combustibility, and ionic conductivity.15 They have been applied in many disciplines, as exemplified by organic and polymer synthesis,16 inorganic synthesis,17,18 enzymatic reactions,19 and formation of self-assemblies.20−22 One of the most notable features of ILs is their limited miscibility with water and organic solvents, which is controllable depending on the chemical structure of cationic and anionic constituents. Although interfaces formed between Received: August 31, 2012 Revised: October 31, 2012 Published: November 2, 2012 4075

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peroxidase (HRP) in BSA microcapsules, these biomacromolecules were added to aqueous BSA solution prior to the emulsification. General Procedures. Scanning electron microscopy (SEM) was performed by Hitachi S-5000 (acceleration voltage, 25 kV). A drop of the capsule suspension was placed on a freshly cleaved mica surface and allowed to dry under ordinary pressure at room temperature. The capsules were then sputter-coated with platinum to enable imaging with SEM. Confocal laser scanning microscopy (CLSM) images were taken with a Carl Zeiss LSM510META apparatus equipped with a 63× water-immersion objective. The excitation wavelength was chosen to be 488 nm for FITC-labeled compounds and sodium fluorescein and 543 nm for rhodamine B and resorufin. ζ-potentials of β-lactogloblin (β-Lg) microcapsules suspended in citrate buffer solution (10 mM, pH 2.9−6.3) were measured on a Malvern Zetasizer Nano-ZS apparatus. UV−vis absorption and emission spectra for solution samples were recorded on a JASCO V-560 and a Perkin-Elmer LS55 spectrophotometer, respectively.

ILs and organic solvents have been studied in purposes of separation or extraction,23 their application in materials chemistry has still been in its infancy.17,24,25 We have developed an interfacial sol−gel synthesis of hollow metal oxide particles by employing toluene microdroplets as template in ILs.17 In the light of environmental inoffensiveness, the microinterface formed between ILs and water is more promising; however, the interfacial materials synthesis at the water−IL interface has been largely unexplored. Here we show a novel methodology to prepare protein microcapsules at the ILs−water microinterface (Scheme 1). It allows simultaneous and quantitative Scheme 1. Schematic Illustration for the Interfacial Synthesis of Protein Microcapsules and Their One-Pot Extraction into the Aqueous Phase

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RESULTS AND DISCUSSION Self-Assembly of Proteins at the Water/IL Microinterface. 1-Butyl-3-methylimidazolium bis(trifluoromethanesulfonyl)amide (bmimTFSA) was used as IL. As proteins, bovine serum albumin (BSA), human serum albumin (HSA), and β-lactogloblin (β-Lg) were employed. Aqueous solutions of proteins (0.1 mL) were added to bmimTFSA (1.9 mL) and the mixture was rigorously stirred for 20 min at 35 °C. Figure 1a

encapsulation of guest biopolymers such as enzymes and nucleic acids into protein microcapsules in the IL phase, which is followed by surface modification and extraction into the aqueous phase. The whole process proceeds in one pot under mild conditions, and their application as enzyme-confined microreactor is demonstrated.

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EXPERIMENTAL SECTION

Materials. 1-Butyl-3-methylimidazolium bis(trifluoromethanesulfonyl)amide (bmimTFSA) was synthesized according to the reported procedure,26 and was used as ionic liquid (IL). The IL was freeze-dried in vacuum and its water content was determined as 0.06 wt % by Karl Fischer titration using a Metrohm 831 KF Coulometer. 3′-Fluorescein-isothiocyanate-modified oligonucleotide (5′-GAA AGG TGT CTT AAA GCA TT-3′-FITC) was purchased from Fasmac. The concentration was determined by measuring the absorbance at 260 nm in a quartz cuvette of 1 mm path length. All other chemicals were purchased from Invitrogen, Sigma, and Wako Pure Chemical Industries and used as received. The water used in all experiments was purified with a Direct-Q system (Millipore) and had a resistivity higher than 18.2 MΩcm. Preparation of Protein Microcapsules. Protein microcapsules were basically prepared according to the following procedure. To 1.9 mL of 1-butyl-3-methylimidazolium bis-(trifluoromethanesulfonyl)amide (bmimTFSA) in a round-bottomed flask was added aqueous protein solution (10 mg mL−1, 0.1 mL) under vigorous stirring at 35 °C. After 20 min, aqueous glutaraldehyde (25 wt %, 0.04 mL) was added to the mixture, and stirring was further continued for 30 min. Upon centrifugation of the mixture (10 000 rpm, 15 min), protein microcapsules were accumulated at the surface of IL layer. These microcapsules were facilely extracted to aqueous solution of 2aminoethanol (0.1 M), which was washed with pure water (1 mL) and collected by centrifugation (10 000 rpm, 5 s). This washing procedure was repeated several times to remove excess 2-aminoethanol. The obtained microcapsules were resuspended in Tris-HCl buffer solution (50 mM, pH 7.4), and the whole synthesis and extraction procedures can be done in one-pot. To encapsulate DNA and horseradish

Figure 1. Aqueous protein microdroplets formed in hydrophobic ILs. (a) Bright-field optical microscopic image of the emulsion prepared from the mixture of bmimTFSA (1.9 mL) and aqueous BSA (10 mg mL−1, 0.1 mL). (b) Confocal fluorescence image of a water microdroplet formed in bmimTFSA. Aqueous solution of FITClabeled BSA (10 mg mL−1) was emulsified in bmimTFSA. (c) Fluorescence intensity profile along with the red line in CLSM image.

shows a bright-field optical microscopic image of the IL-water emulsion obtained by mixing bmimTFSA and aqueous BSA (10 mg mL−1). Aqueous microdroplets with diameters below 10 μm are abundantly seen. Similar microdroplets were also formed from the other water-immiscible imidazolium- or ammonium-based IL with TFSA or PF6− counterions. The ILs play indispensable role for the formation of microcapsules, since such stable emulsions were not obtainable with common organic solvents. When aqueous BSA and chloroform or toluene were mixed rigorously, they separated into each liquid phases immediately after stopping of the stirring. Although an emulsion-like mixture was obtained by mixing aqueous BSA and viscous liquid paraffin, micrometer-sized water droplets were not produced as confirmed by optical microscopy. These 4076

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as mentioned above. Very interestingly, luminescent hollow microspheres (diameter, 10−20 μm) are abundantly seen. Such hollow microcapsules were similarly obtained from the other proteins such as HSA and β-Lg. In contrast with the case of BSA, HSA, and β-Lg, microcapsules were not formed when glycoproteins such as glucose oxidase (GOD) and HRP were employed. The formation of protein microcapsules thus depends on the molecular structure, which can be related to the proportion of lysine residues contained in these proteins (BSA, 59 Lys per 583 residues;27 HSA, 59 Lys per 584 residues;27 β-Lg, 15 Lys per 162 residues;28 GOD, 15 Lys per 583 residues;29 HRP, 6 Lys per 308 residues30). The lower content of lysine residues in GOD and HRP and the presence of surface sugar chains on the surface of these glycoproteins seem to be disadvantageous for the cross-linking reaction. SEM images of BSA and β-Lg microcapsules are shown in Figure 2b and c, respectively. Withered microcapsules with diameters of 7−15 μm were observed for BSA, whereas red-blood-cell-like flattened microcapsules (diameter, 2−7 μm) were observed for β-Lg. The deformation from spherical structures is attributable to the drying process, and maintenance of these morphologies reveals stability and elasticity of these microcapsules. Permeability of Protein Microcapsules. Permeation property of protein microcapsules was then investigated. Fluorescent dye molecules (fluorescein, Flu; rhodamine B, RhB) were employed as permeants. Microcapsules consisting of β-Lg were employed, and their ζ potential was determined at varied pH (Figure S1, Supporting Information). It showed negative surface charge above pH 4.8, which is consistent with the isoelectric point of β-Lg (pI, 5.2) and decreased number of Lys residues caused by modification with glutaraldehyde. To βLg microcapsules suspended in Tris-HCl buffer (50 mM, pH 7.4), equimolar mixture of Flu and RhB in Tris-HCl buffer ([Flu] = [RhB] = 33 μM) was added. After 30 min, the crosssectional CLSM image of β-Lg microcapsules in aqueous dye solution was observed, as shown in Figure 3a. The green color indicates the luminescence of Flu in the 505−530 nm region (excitation wavelength, λex = 488 nm), whereas the red color visualizes the luminescence of RhB observed at a wavelength beyond 560 nm (λex = 543 nm). Figure 3b shows the fluorescence intensity profile along with the broken line shown in Figure 3a. The green luminescence of Flu is observed for both the outside and the interior of β-Lg microcapsules, whereas the luminescence intensity is obviously weaker on the microcapsule wall (Figure 3b, green line). Apparently, anionic Flu molecules permeate anionic microcapsules without being strongly adsorbed to the β-Lg layer. In contrast, intense luminescence of RhB is observed preferentially on the microcapsule wall, revealing strong tendency of zwitter ionic RhB to be adsorbed on β-Lg. To investigate the permeant diffusivity, FITC-labeled dextran (FITC-dextran) with varied average molecular weights (4k, 10k, 40k and 150k Da) were employed. Figure 3c shows CLSM images of β-Lg microcapsules in Tris-HCl buffer 30 min after the addition of aqueous FTIC-dextran (λex = 488 nm). When aqueous FITC-dextran (4k Da) was added, its fluorescence was also observed from the inner volume of β-Lg microcapsules (Figure 3c, left). In contrast, FITC-dextran with higher average molecular weight of 40k did not show permeability, as can be seen from the dark inner volume (Figure3c, right). Figure 3d compares time courses of relative fluorescence intensity (Iinside/ Ioutside of a microcapsule) observed after adding FITC-dextrans

observations clearly demonstrate the advantage of ILs for preparing protein microcapsules. To visualize protein microcapsules by using confocal laser scanning microscopy (CLSM), fluorescein isothiocyanatelabeled bovine serum albumin (FITC-BSA) was employed. Figure 1b shows a CLSM image of aqueous FITC-BSA emulsified in bmimTFSA (λex = 488 nm, luminescence intensity above 505 nm was recorded). Spherical microstructures (diameter ∼5 μm) were found, where the cross sectional profile (Figure 1c) of green FITC-BSA fluorescence show accumulation of proteins at the IL-water microinterface. These CLSM observations clearly indicate spontaneous formation of protein microcapsules at the IL-water microinterface. Simultaneous Surface Modification and Transfer of Protein Microcapsules into the Aqueous Phase. Because BSA microcapsules formed in IL-water emulsions collapse when they are rigorously mixed with water, they were stabilized by chemical cross-linking. Aqueous glutaraldehyde (0.1 mmol) was added to the IL-water emulsion containing BSA microcapsules, and the mixture was further stirred for 30 min to complete the cross-linking reaction at 35 °C. The mixture was centrifuged (10 000 rpm, 10 min) to accumulate glutaraldehyde-treated protein microcapsules at the surface. Aqueous 2aminoethanol (0.1 M, 1 mL) was then layered on the IL-water emulsion, and upon gently shaking the mixture, the aqueous phase became luminescent, as shown in the inset in Figure 2a.

Figure 2. Images of protein microcapsules. (a) Confocal fluorescence image of FITC-BSA microcapsules in water (λex = 488 nm). Inset shows a picture of FITC-BSA microcapsules extracted into the aqueous 2-aminoethanol (0.1 M) layer on the IL phase. The specimen was illuminated by a UV lamp (λex = 365 nm). (b,c) SEM images of BSA (b) and β-Lg (c) microcapsules.

The presence of 2-aminoethanol in the aqueous phase is necessary because protein microcapsules stabilized by the glutaraldehyde-treatment are not extractable to the aqueous phase. It indicates that the reaction between glutaraldehyde and lysine residues decreased the surface charge and consequently converted the microcapsule surface hydrophobic. It was found that the unreacted aldehyde groups remain on the microcapsule surface, and their modification with hydrophilic amines enhanced the hydrophilicity of the microcapsule and allowed their facile extraction to the aqueous phase. (See the Supporting Information.) To purify the sample, we further centrifuged the aqueous layer, and collected specimens were rinsed with pure water. They were resuspended in pure water by hand-shaking, and this procedure was repeated for several times. Figure 2a shows a CLSM image of the luminescent aqueous phase obtained by resuspending the specimen purified 4077

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average molecular weights of 10k and 40 k Da are reported to be 2.3 and 4.6 nm,31 respectively. Therefore, β-Lg microcapsules reveal barrier properties against molecules having diameter of ∼9 nm. Quantitative Encapsulation of Biopolymers in Protein Microcapsules. Encapsulation of DNA was further investigated for BSA microcapsules. A single-strand DNA labeled with FITC at the 3′-position (5′-GAA AGG TGT CTT AAA GCA TT-3′-FITC, 2.5 μM) was mixed with aqueous BSA (10 mg mL−1), and the mixture (0.1 mL) was added to bmimTFSA (1.9 mL) under rigorous stirring. After stirring at 35 °C for 20 min, microcapsules were extracted to Tris-HCl buffer, as previously described. Figure 4a shows CLSM image of microcapsules resuspended in Tris-HCl buffer (λex = 488 nm). It is seen that fluorescence of FITC is observed only for the inner volume (Figure 4b), as confirmed by fluorescence spectra obtained at positions 1 (inside microcapsule) and 2 (bulk buffer phase). Apparently, DNA exists only inside BSA microcapsules, and it is apparent that the present methodology allows simultaneous and quantitative encapsulation of water-soluble guest biopolymers inside protein microcapsules. The microcapsules also serve as reactor vessels for enzymatic reactions, as demonstrated by encapsulation of HRP. As previously described, HRP alone never forms microcapsules that can be extracted to the aqueous phase because of the small number of Lys residues existing on the protein surface; however, this is a desirable feature for guest proteins that are not incorporated in the capsule walls when they coexist together with easily capsule-forming (host) proteins such as BSA. To test this hypothesis, aqueous mixture (0.1 mL) of BSA (10 mg mL−1) and HRP (0.1 mg mL−1) were added to bmimTFSA (1.9 mL) under rigorous stirring. HRP@BSA protein microcapsules formed in the IL phase are cross-linked and extracted to the aqueous phase according to the previously

Figure 3. Permeability behavior of β-Lg microcapsules. (a) Confocal fluorescence image of β-Lg microcapsules in aqueous buffer (50 mM Tris/HCl, pH 7.4) containing sodium fluorescein (33 μM) and rhodamine B (33 μM). The fluorescence image was obtained after 30 min by double excitation of 488 nm with bandpass filter (505−530 nm, green) and 543 nm with long-pass filter (560 nm, red). (b) Fluorescence intensity profile along with the broken line in panel a. (c) CLSM images of β-Lg microcapsules obtained after 30 min in aqueous buffer (50 mM Tris/HCl, pH 7.4) containing FITC-labeled dextran (4k and 40k Da, [FITC group] = 30 μM). (d) Time courses of relative fluorescent intensity of inner and outer capsule upon the addition of aqueous FITC-dextran. The arrow indicates the time at which FITCdextran with different molecular weights (4k, 10k, 40k and 150k Da) was added.

to the aqueous β-Lg microcapsules. In the case of FITCdextrans with average molecular weights of 4k and 10k, relative luminescence intensity showed immediate increase within 10 s. In contrast, the fluorescence intensity of the microcapsule interior was very small for FITC-dextran with higher molecular weights (40k, 150k). Stokes radius of dextran molecules with

Figure 4. Encapsulation properties of BSA microcapsules. (a) Confocal fluorescence image of BSA microcapsules loaded with FITC-DNA in aqueous buffer (50 mM Tris/HCl, pH 7.4, λex = 488 nm, emission monitored from 495 to 666 nm). (b) Emission spectra of the region 1 and 2 in panel a. (c) Confocal fluorescence (FL), differential interference contrast (DIC), and the merged images of HRP-loaded BSA capsules in aqueous buffer (50 mM Tris/HCl, pH 7.4). The fluorescence image was obtained after 15 s of imaging (λex = 543 nm, visualized below 560 nm). Aqueous H2O2 was added to the mixture of HRP@BSA microcapsules and Amplex Red ([Amplex Red] = [H2O2] = 10 μM). (d) Time dependency of emission intensity upon the addition of aqueous H2O2. Regions 3 and 4 correspond to the circle in panel c. The arrow indicates the time at which aqueous H2O2 was added. 4078

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microcapsules. The one-pot quantitative encapsulation of water-soluble guest biomolecules, and scrapping volatile organic solvents, surfactants, or solid colloid templates solve the major problems with existing technology. Third, these protein microcapsules formed in the IL phase are transferred to aqueous phase by way of successive cross-linking and augmentation of the surface hydrophilicity. The enzymatic ability, that is, tertiary structure of guest enzyme, is maintained inside aqueous microcapsules, as shown by microspectrophotometry with CLSM. Moreover, the protein microcapsules show barrier property against macromolecules, as demonstrated for β-Lg. The spontaneous separation of biopolymers at the IL−water microinterface would be widely applied to synthesize hierarchical architectures in many disciplines.

mentioned procedure. After repetitive washing of the obtained microcapsules with pure water and centrifugation, they were resuspended in Tris-HCl buffer (50 mM, pH 7.4). The enzymatic activity of HRP@BSA microcapsules was monitored by Amplex Red assay (Figure S2, Supporting Information).32,33 To a Tris-HCl buffer (50 mM, 1 vol % DMSO, pH 7.4) containing Amplex Red (10 μM) and hydrogen peroxide (10 μM), HRP@BSA microcapsules were added, and the mixture was allowed to stand for 30 min. As a reference, BSA microcapsules prepared in the absence of HRP were employed. The colorless reaction mixture containing HRP@BSA microcapsules became red with the progress in time, which is ascribed to resorufin formed by HRP-catalyzed oxidation of Amplex Red. This is confirmed from absorption (λmax = 563 nm) and fluorescence spectra (λem = 586 nm) of supernatant obtained after centrifugal separation of microcapsules (Figure S2, Supporting Information). HRP is exclusively encapsulated in BSA microcapsules and not present in the bulk solution, as shown by the following measurements. First, a Tris-HCl buffer solution containing Amplex Red (10 μM) and hydrogen peroxide (10 μM) was added to the supernatant obtained after separating microcapsules by centrifugation. Absorption intensity for resorufin did not increase, even after 30 min (Figure S3, Supporting Information), indicating that HRP did not exist in the bulk phase. To visualize the progress of enzymatic reaction inside BSA microcapsules, the mixture of HRP@BSA microcapsules and Amplex Red was placed on a glass-bottomed dish. After 10 min, hydrogen peroxide was added to the mixture, and the progress in enzymatic reaction was imaged in real time by using CLSM ([Amplex Red] = [hydrogen peroxide] = 10 μM, 50 mM Tris-HCl buffer, pH 7.4). Figure 4c shows CLSM images 15 s after the addition of hydrogen peroxide (upper, fluorescence (FL) image; middle, differential interference contrast (DIC) image; bottom, merged image). As clearly seen from the merged image, fluorescence of resorufin is observed only from the inside of HRP@BSA microcapsules. Figure 4d compares the time course of fluorescence intensity changes observed inside HRP@BSA microcapsules (region 3) and in the bulk solution phase (region 4) after the addition of aqueous hydrogen peroxide. It confirms that there is no increase in fluorescence intensity in the bulk aqueous phase (region 4), whereas fluorescent resorufin was produced only inside the microcapsules. These observations clearly demonstrate that HRP is totally localized in the aqueous interior of BSA microcapsules with maintaining its enzymatic activity. Therefore, the IL−water interface allows separation and localization of coexisting biopolymers within a microcapsule structure. The simultaneous formation of microcapsules and inclusion of guest biopolymers is a feature not available in existing methodologies, and it validates the unique potential of IL−water microinterfaces.

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ASSOCIATED CONTENT

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AUTHOR INFORMATION

S Supporting Information *

ζ-potential, UV−vis absorption, fluorescence spectra, and detailed methods. This material is available free of charge via the Internet at http://pubs.acs.org.

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This research was partially supported by a Grant-in-Aid for Scientific Research (nos. 18045028 and 20031023) on Priority Area “Science on Ionic Liquids” (area no. 452), a Grant-in-Aid for Scientific Research on Innovative Areas “Emergence in Chemistry” from the Ministry of Education, Culture, Sports, Science and Technology of Japan, and by JST, CREST.

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REFERENCES

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CONCLUSIONS We have developed a novel methodology to prepare protein capsules by using the microinterface formed between ILs and water. Proteins such as BSA, HSA, and β-Lg rapidly form microcapsules at the interface under mild conditions, where guest biopolymers are quantitatively encapsulated in the inner aqueous pool. The significance of present work is summarized as follows. First, this is the first report on the formation of protein microcapsules by using IL−water interface. Second, it provides a simple and useful methodology to encapsulate guest biopolymers concomitantly with the formation of protein 4079

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