ISSN 10681620, Russian Journal of Bioorganic Chemistry, 2013, Vol. 39, No. 5, pp. 504–509. © Pleiades Publishing, Ltd., 2013. Original Russian Text © O.Yu. Kochetkova, L.I. Kazakova, D.A. Moshkov, M.G. Vinokurov, L.I. Shabarchina, 2013, published in Bioorganicheskaya Khimiya, 2013, Vol. 39, No. 5, pp. 565–571.
Adapted from a report presented at the VI Russian Symposium “Proteins and Peptides” (June 11–15, 2013)
Incorporation of Proteins into Polyelectrolyte Microcapsules by Coprecipitation and Adsorption O. Yu. Kochetkovaa, b, 1, L. I. Kazakovaa, D. A. Moshkova, M. G. Vinokurovb, and L. I. Shabarchinaa a
Institute of Theoretical and Experimental Biophysics, Russian Academy of Sciences, Pushchino, Moscow region, 142290 Russia b Institute of Cell Biophysics, Russian Academy of Sciences, Pushchino, Moscow region, 142290 Russia Received March 29, 2013; in final form, April 22, 2013
Abstract—Microcapsules composed of synthetic (sodium polystyrene sulfonate and polyallylamine hydro chloride) and biodegradable polyelectrolytes (dextran sulfate and polyarginine hydrochloride) deposited on carbonate microparticles have been obtained. The ultrastructural organization of biodegradable microcap sules has been studied by transmission electron microscopy. The shell of biodegradable microcapsules is well formed even after the deposition of six polyelectrolyte layers and has an average thickness of 44 ± 3.0 nm; their inner polyelectrolyte matrix is less branched than that of synthetic microcapsules. By using spectroscopy, the efficiency of the encapsulation of FITClabeled BSA by adsorption depending on the number of PE layers in the capsule has been estimated. It has been shown that the maximum amount of the protein is incorporated into capsules comprising six and seven polyelectrolyte layers (4 and 2 pg/capsule, respectively). It has been concluded that the adsorption of proteins into preformed polyelectrolyte capsules enables one to avoid pro tein losses that occur with the method in which biomineral cores obtained by coprecipitation are used for encapsulation. Keywords: polyelectrolyte microcapsules, polyelectrolytes, coprecipitation, electron microscopy, encapsulation DOI: 10.1134/S1068162013050087 1
INTRODUCTION Since the publication of the first papers devoted to the fabrication of polyelectrolyte microcapsules (1998) up to the present day, the study of their struc ture and physicochemical properties has remained a topic of great interest [1–3]. PEMCs are prepared by the layerbylayer adsorption of oppositely charged PEs onto charged nano and microsized template colloidal particles (cores) followed by the removal of template particles [4, 6]. Depending on the morphol ogy and composition of microparticles used as a tem plate for the deposition of PE layers, hollow PEMCs or matrixtype microcapsules filled with a spatially complicatedly organized PE complex resembling the internal structure of template particles can be formed [6, 7]. Abbreviations: BSA, bovine serum albumin; DS, dextran sul fate; EDTA, ethylenediaminetetraacetic acid; FITC, fluores cein isothiocyanate; IPEC, interpolyelectrolyte complex; PAH, polyallylamine hydrochloride; PAr, polyarginine hydrochloride; PE, polyelectrolyte; PEMC, polyelectrolyte microcapsule. 1 Corresponding author: phone: +7 (4967) 739205; email:
[email protected].
The diverse variants of the design of PE capsules make possible their wide use in applied fields [8–10]. Depending on the purpose of the application, the shell of microcapsules can be formed from nondegradable PEs (PAH, sodium polystyrene sulfonate, polydial lyldimethylammonium chloride), referred below to as synthetic, or biodegradable PEs (DS, sodium alginate, PAr, and others). Capsules formed from biopolymers are used for the delivery of biologically active compo nents to cells and tissues and in the design of pro longedaction drugs [11–15]. The greatest number of publications in this field are concerned with the encapsulation of highmolecularweight compounds, in particular proteins [16–18]. In the literature, sev eral methods of successful encapsulation of proteins into PEMCs are described [19–21]. The most extensively employed method for the fab rication of proteincontaining PEMCs is based on the use of a biomineral core as a template for the adsorp tion of PE layers [22, 23]. Biomineral cores represent colloidal, usually micronsized particles, which result from the coprecipitation of proteins with the salts of alkalineearth metals, in most cases calcium carbon
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ate [22–24]. After the formation of a PE shell on these particles and the decomposition of the mineral template (by medium acidification and dissolution in EDTA), PE capsules with a content of the protein of about 10 pg per microcapsule are obtained [6, 16, 25, 26]. Along with the coprecipitation method, the encap sulation of bioactive compounds into preformed PEMCs by adsorption was described [6, 16, 26]. Bala bushevich et al. reported on the efficient encapsula tion of proteins of different molecular weights and charges into PEMCs formed with the use of melamine formaldehyde cores. The authors believe that the incorporation of a protein into microcapsules is due to the presence inside their cavity of a gellike structure, which results from the core decomposition [26]. Volod’kin et al. described the adsorption of pro teins in PEMCs formed with the use of the calcium carbonate mineral core [6]. The authors explained the possibility of the encapsulation of proteins by their interaction with charged uncompensated regions of the IPEC formed due to the adsorption of PE in the cavities of porous calcium carbonate microparticles. In addition to purely logical conclusions about the role of electrostatic interactions in this process, the authors presented, as evidence for the existence of these frag ments, the Xray diffraction data indicating that cap sules fabricated from synthetic PEs PSS and PAH contain free charged functional groups of polystyrene sulfonate. At present, there are no direct instrumental methods for evaluating the amount of free charged groups of PE inside complexes. In our opinion, the analysis of the internal ultra structural organization of PEMC may provide a picto rial idea of the protein adsorption process in PEMC, the distribution of protein molecules in capsules, and their potential amount. Using the method of transmis sion electron microscopy, we have analyzed earlier the ultrathin sections of PEMCs formed on porous min eral CaCO3 particles by means of synthetic PEs, PAH and PSS [6]. We have shown that the sequential adsorption of oppositely charged PEs during the prep aration of PEMCs leads first to the formation of a branched internal PE matrix. As the cavities of a CaCO3 microparticle are filled with the polymers, the formation of the PE layers of the capsule shell occurs. In the case of 4.5–5µm capsules prepared from the above indicated pair of PEs, a uniform wellformed shell results after the deposition of nine PE layers. The same number of PE layers provides a reliable retention of proteins inside capsules when biomineral particles are used as a template for the deposition of PEs [6, 7]. In this connection, it is of interest to establish a relationship between the number of PE layers of the capsule shell and the amount of the protein that can be encapsulated by adsorption. These data would enable one to rule out the losses of a protein at the stage of its encapsulation. In addition, as distinct from the encap RUSSIAN JOURNAL OF BIOORGANIC CHEMISTRY
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sulation method involving the use of coprecipitants, the method based on adsorption makes it possible to encapsulate strictly specified small amounts of com pounds into PEMC. Therefore, one of the main goals of this study was to determine the architecture of PE capsules that is optimal for the protein encapsulation by adsorption. An understanding of this problem is of particular importance for the encapsulation of pro teins and peptides that possess a high biological activ ity and are often very expensive. In the present work, we solved these problems using the fluorescently labeled BSA as a model protein; its encapsulation was controlled spectrophotometrically. Considering the possibility of further practical application of the adsorption method for the encapsu lation of biologically active compounds and the use of the resulting capsules in biomedicine and pharmaceu tics, it seemed reasonable to study the encapsulation of proteins into PEMCs formed from natural biopoly mers. The ultrastructure of biodegradable capsules formed from DS and PAr was determined by transmis sion electron microscopy. For adsorption, the elec trondense protein ferritin was used, which enables one to locate the protein in capsules after adsorption without the additional contrasting of capsule sections. RESULTS AND DISCUSSION Using transmission electron microscopy, we obtained images of sections of PEMCs formed on СаСО3 microparticles by means of biodegradable PEs DS and PAr and compared them with capsules formed on the same carbonate cores using synthetic PEs PSS and PAH (Fig. 1). Both types of capsules were con trasted with an aqueous solution of uranyl acetate and lead citrate to visualize the PE shell. Microcapsules composed of PSS and PAH deposited on СаСО3 microparticles had no regular external shell and con sisted of threadlike linear and closed nanosized ele ments. The internal structure of these capsules repre sented a complicatedly organized spongelike matrix consisting of spatially organized threadlike and closed molecules of PEs and their complexes (Fig. 1a). The regular shell of these microcapsules was formed after the deposition of nine PE layers, which we previ ously described in more detail [7]. It is seen from Fig. 1b that, in biodegradable micro capsules, one can recognize a shell and an internal branched matrix. It is important to note that the shell of these microcapsules is well formed even after the deposition of six PE layers; its thickness is on the aver age 44 ± 3.0 nm, and the internal PE matrix is less branched than in synthetic PEMCs. Presumably, these structural features of biodegradable microcapsules are due to the larger sizes of DS molecules as compared with PSS and PAH and, as a consequence, the diffi culty of their penetration into the cavities of porous СаСО3 microparticles. Vol. 39
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(a)
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Fig. 1. Electron microphotographs of ultrathin sections of PEMCs formed on carbonate cores from (a) synthetic PEs having the architecture (PAH/PSS)3 and (b) biodegradable PEs with the architecture (DS/PAr)3.
The presence of the IPEC in biodegradable micro capsules formed on СаСО3 cores suggests that they can be effectively loaded with charged and amphiphilic molecules, including proteins, using the adsorption method. For encapsulation, we chose ferritin, an ironcontaining electrondense protein (Fig. 2a). Ferritin made it possible to visualize the ultrastructural organization of these microcapsules without contrast ing the samples for transmission electron microscopy. Figure 2b presents an electron micrograph of the ultrathin section of a biodegradable microcapsule with the incorporated protein. It is seen from the figure that ferritin was adsorbed onto all structural elements of the microcapsule by interacting with the IPEC of the internal matrix and the outer capsule shell. Using the spectroscopy method, we obtained data that enabled us to assess the efficiency of protein inclusion into synthetic microcapsules depending on the number of PE layers (Fig. 3). FITClabeled BSA was used as an object of encapsulation. FITCBSA was incorporated into microcapsules formed on carbonate cores by adsorption. An analysis indicated that the amount of the pro tein adsorbed into a microcapsule depends on the number of PE layers in its shell (Fig. 3). The maximum amount of BSA was contained in capsules with the number of PE layers 6 and 7 (4 and 2 pg/capsule, respectively). A further increase in the number of PE layers led to a decrease in the amount of the encapsu lated protein. Probably, this is related to a decrease in the permeability of the shell for protein molecules. The upper “terminal” layer also played an impor tant role in the adsorption process. We showed that,
with the polyanion PSS as the upper layer, almost no protein encapsulation occurred. It should be noted that the key points in this method were the use of a porous carbonate microparticle, which contributes to the formation of a branched interpolyelectrolyte matrix of the microcapsule, as well as the architecture and structure of PEs the external shell is formed of. It was shown that, by using the coprecipitation method, it is possible to encapsulate about 10 pg pro tein/capsule [6], which accounts for more than 60% of the total amount of the protein being encapsulated; i.e., the losses with this approach are about 40%. The use of adsorption enables one to almost completely avoid these losses. This is primarily achieved by the application of microcapsules formed on porous car bonate microparticles. In addition, the adsorption method enables one to preliminarily calculate the capacity of a microcapsule by the choice of the appro priate architecture and the composition of PEs. Thus, the adsorption method can be considered as a universal approach that can be applied for the encap sulation of various polar and amphiphilic molecules. It is hoped that the use of biodegradable microcapsules may become in future a new chapter in biotechnology and nanomedicine. EXPERIMENTAL The following PEs were used: PSS (MW 70 kDa), PAH (MW 70 kDa), DS (MW 15 kDa), and PAr (MW 70 kDa; all Sigma–Aldrich, Germany). BSA was from Sigma (United States), and ferritin was from MP Bio medicals (France). The following reagents were used:
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Fig. 2. Electron microphotographs of (a) ferritin and (b) a biodegradable (DS/PAr)3 microcapsule formed on a carbonate core after the adsorption of ferritin.
Fluorescently labeled BSA was obtained by incu bating a solution of the protein with an alcohol FITC solution (Sigma–Aldrich, Germany) for 2 h at room temperature in 0.1 M borate buffer (pH 9.0) in the [dye]/[protein] ratio 3 : 1. After the termination of incubation, the mixture was dialyzed against deion ized water. The fixation and contrasting of preparations for the transmission electron microscopy were performed using glutaraldehyde (Polaron, United States); osmium tetraoxide, lead uranyl acetate and lead cit rate (Sigma, United States); and epoxide resins (Schuhard, Germany). In the spectrophotometric study, deionized water with a resistance of above 18.2 MOhm/cm was used, which was prepared by the threestage purification on a MilliQ Plus 185 device. In other experiments, bidistilled water was used. Preparation of CaCO3 microparticles. To a 1 M aqueous CaCl2 solution, an equivalent volume of 1 M aqueous Na2CO3 solution was quickly added under stirring (350 rpm). After stirring for 30 s, the suspen sion was left for 5—7 min until the complete clarifica tion of the supernatant. The resulting CaCO3 particles were centrifuged for 30 s at 4000 g, washed three times with bidistilled water, and used for the preparation of PEMCs. The growth of microparticles was controlled RUSSIAN JOURNAL OF BIOORGANIC CHEMISTRY
by a Nikon Eclipse E200 light microscope (Japan); the size distribution of particles was 4.5–5.0 µm. PEMCs were fabricated by layerbylayer adsorp tion of oppositely charged PEs onto mineral cores. The adsorption of polymers onto the surface of CaCO3 microparticles was carried out in 0.5 M NaCl solutions 5 pg BSA/microcapsule
EDTA (Sigma, United States); and calcium chloride, sodium carbonate, sodium chloride, and sodium tet raborate (Reakhim, Russia; all of chemical purity grade).
4 3 2 1 0 6
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8 9 10 Number of PE layers
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Fig. 3. Dependence of the incorporation of BSA into syn thetic microcapsules by adsorption on the number of PE layers in the capsule shell. Microcapsules with the number of PE layers 6, 8, and 10 had the architecture (PSS/PAH)3,4,5, and those with the number of PE layers 7, 9, and 11 had the architecture (PAH/PSS)3,4,5 PAH. Vol. 39
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containing the PEs PSS and PAH or DS and PAr at a concentration of 2 mg/mL. After each adsorption step, particles were washed three times from unbound polymers. Microparticles were separated from the supernatant by centrifugation. After the deposition of the appropriate number of layers, the carbonate core was dissolved in 0.2 M EDTA. The size of ready microcapsules correlated with the size of microparti cles used and was 4.5–5.0 µm. Using synthetic PEs, we obtained microcapsules containing no proteins and having the following architecture of PE shells: 1) (PSS/PAH)n and (PAH/PSS)n, where n = 3–5; 2) (PAH/PSS)n PAH, where n = 3–5. With the use of biodegradable PEs, microcapsules with the architecture (DS/PAr)3 were obtained. The resulting microcapsules were washed addition ally three times with water to remove the degradation products of the carbonate core and stored in the form of suspension at 4°C. The titer of capsules in the sus pension was determined using a Goryaev chamber. Encapsulation of a protein into PEMCs formed on CaCO3 microparticles through adsorption. For the encapsulation of a protein through adsorption, a known amount of a protein (BSA or ferritin) was added to a suspension of PEMCs in water and incu bated for 10 min at 4°C under stirring. Then the mix ture was centrifuged for 30 s at 4000 g and washed three times with cold deionized water. Microcapsules with the adsorbed protein were separated from the superna tant by centrifugation and resuspended in the appro priate amount of water. Transmission electron microscopy. Preparation of ultrathin sections of PEMCs. Sections about 70 nm thick were prepared by a conventional method on an LKB3 ultratome (Sweden) [27]. To do this, micro capsules were fixed for 1 h in a 5% aqueous glutaralde hyde solution. The resulting suspensions were centri fuged for 5 min at 3000 rpm, and the sediments were resuspended for complete fixation in a 1% aqueous osmium tetroxide solution. After the fixation for 1 h, the suspension was sedimented again, and the sedi ments were dehydrated by successive washing in 30, 50, 75, and 90% mixtures of acetone with water and then in 100% waterfree acetone, after which they were soaked in acetone–epoxide resin Epon 812 in the ratios 1 : 1 and 1 : 3 for 12 h in each mixture, and embedded into fresh resin. After the completion of polymerization, samples were cut by glass knives. Sec tions were trapped onto supporting nets, contrasted with an aqueous solution of uranyl acetate (1.5 h) and lead citrate (30 min), and examined in a Tesla BS500 electron microscope (Czech Republic) at an acceler ating voltage of 90 kV and a magnification of x18 000. Negatives were subjected to a morphometrical analysis using a Mikrofot photomultiplier (GDR) at a fixed 6.5 and 9fold magnification.
ACKNOWLEDGMENTS The authors would like to thank Dr. R.Sh. Shtan chaev (Institute of Theoretical and Experimental Bio physics, Russian Academy of Sciences) for the help in the morphometry of negatives of electron microscope images. This work was supported by the Russian Founda tion for Basic Research (project no. 130401507 A). REFERENCES 1. Sukhorukov, G.B., Donath, E., Davis, S., Lichtenfeld, H., Caruso, F., Popov, V.I., and Mohwald, H., Polym. Adv. Technol., 1998, vol. 9, pp. 759–767. 2. Inozemtseva, O.A., Portnov, S.A., Kolesnikova, T.A., and Gorin, D.A., Ross. Nanotekhnol., 2007, vol. 2, pp. 68–80. 3. Dubrovskii, A.V., Shabarchina, L.I., Kim, Yu.A., and Sukhorukov, B.I., Zh. Fiz. Khim., 2006, vol. 80, pp. 1914–1919. 4. Voigt, A., Lichtenfeld, G.B., Sukhorukov, G.B., Zas trov, H., Donath, E., Baumler, H., and Mohwald, H., Ind. Eng. Chem. Res., 1999, vol. 38, pp. 4037–4043. 5. Donath, E., Sukhorukov, G.B., Caruso, F., Davis, S.A., and Mohwald, H., Angew. Chem., Int. Ed. Engl., 1998, vol. 37, pp. 2202–2205. 6. Volodkin, D.V., Petrov, A.I., Petrov, M., and Sukho rukov, G.B., Langmuir, 2004, vol. 20, pp. 3398–3406. 7. Kazakova, L.I., Dubrovskii, A.V., Moshkov, D.A., Sha barchina, L.I., and Sukhorukov, B.I., Biofizika, 2007, vol. 52, pp. 850–854. 8. Studer, D., Palankar, R., Bédard, M., Winterhalter, M., and Springer, S., Small, 2010, vol. 6, pp. 2412–2419. 9. Kazakova, L.I., Shabarchina, L.I., and Sukhorukov, G.B., Phys. Chem. Chem. Phys., 2011, vol. 13, pp. 11110– 11117. 10. Del Mercato, L.L., Abbasi, A.Z., Ochs, M., and Parak, W.J., ACS Nano, 2011, vol. 5, pp. 9668–9674. 11. De Koker, S., De Cock, L.J., RiveraGil, P., Parak, W.J., Velty, R.A., Vervaet, C., Remon, J.P., and De Geest, B.J., Adv. Drug Deliv. Rev., 2011, vol. 63, pp. 748–761. 12. De Geest, B.J., Dejugnat, C., Verhoeven, E., Sukho rukov, G.B., Jonas, A.V., Plain, J., Demeester, J., and De Smedt, S.C., J. Controlled Rerlease, 2006, vol. 116, pp. 159–169. 13. De Koker, S., De Geest, B.J., Singh, S.K., De Rycke, R., Naessens, T., Van Kooyk, Y., Demeester, J., and De Smedt, S.C., Angewandte Chemie (Int. Ed.), 2009, vol. 48, pp. 8485–8489. 14. Borodina, T.N., Rumsh, L.D., Kunizhev, S.M., Sukhorukov, G.B., Vorozhtsov, G.N., Fel’dman, B.M., Rusanova, A.V., Vasil’eva, T.V., Strukova, S.M., and Markvicheva, E.A., Biomed. Khim., 2007, vol. 53, pp. 662–671. 15. Borodina, T.N., Rumsh, L.D., Kunizhev, S.M., Sukhorukov, G.B., Vorozhtsov, G.N., Fel’dman, B.M., and Markvicheva, E.A., Biomed. Khim., 2007, vol. 53, pp. 557–565.
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INCORPORATION OF PROTEINS INTO POLYELECTROLYTE MICROCAPSULES 16. She, Zhen, Antipina, M.N., Li, Jun, and Sukho rukov, G.B., Biomacromolecules, 2010, vol. 11, pp. 1241–1247. 17. Volodkin, D.V., Larionova, N.I., and Sukhorukov, G.B., Biomacromolecules, 2004, vol. 5, pp. 1962–1972. 18. Qi, W., Yan, X.H., Juan, L., Cui, Y., and Li, J.B., Bio macromolecules, 2009, vol. 10, pp. 1212–1216. 19. Antipov, A.A., Sukhorukov, G.B., Donat, E., and Möh wald, H., J. Phys. Chem. B, 2001, vol. 105, pp. 2281– 2284. 20. Balabushevich, N.G., Tiourina, O.P., Volodkin, D.V., Larionova, N.I., and Sukhorukov, G.B., Biomacromo lecules, 2003, vol. 4, pp. 1191–1197. 21. Tiourina, O.P., Antipov, A.A., Sukhorukov, G.B., Lari onova, N.I., Lvov, Y., and Möhwald, H., Macromol. Biosci., 2001, vol. 1, pp. 209–214.
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22. Antipov, A.A., Shchukin, D., Fedutik, Y., Petrov, A.I., Sukhorukov, G.B., and Mohwald, H., Colloid. Surf.: Physicochem. Eng. Aspect, 2003, vol. 224, pp. 175–184. 23. Sukhorukov, G.B., Volodkin, D.V., Gunther, A.M., Petrov, A.I., Shenoy, D.V., and Möhwald, H., J. Mater. Chem., 2004, vol. 14, pp. 2073–2081. 24. Petrov, A.I., Volodkin, D.V., and Sukhorukov, G.B., Biotechnol. Prog., 2005, vol. 21, pp. 918–925. 25. Kazakova, L.I., Dubrovskii, A.V., Santalova, I.M., Moshkov, D.A., Apolonnik, N.V., and Shabarchina, L.I., Russ. J. Bioorg. Chem., 2012, vol. 38, pp. 51–55. 26. Balabushevich, N.G., Sukhorukov, G.B., and Lario nova, N.I., Vestn. Mosk. Univ. Ser. 2 Khim., 2002, vol. 43, pp. 374–377. 27. Moshkov, D.A., in Adaptatsiya i ul’trastruktura neirona (Adaptation and Neuron Ultrastructure), Moscow: Nauka, 1985.
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