Design of Double Stimuli-Responsive Polyelectrolyte Microcontainers

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Biomacromolecules 2010, 11, 806–814

Design of Double Stimuli-Responsive Polyelectrolyte Microcontainers for Protein Soft Encapsulation Christian Basset,† Christophe Harder,† Claude Vidaud,† and Christophe De´jugnat*,‡,§ DSV/iBEB/SBTN/LEPC, ICSM, UMR 5257 CEACNRS-UM2-ENSCM, CEA Marcoule, BP 17171, 30207 Bagnols-sur-Ce`ze cedex, France Received December 15, 2009; Revised Manuscript Received January 19, 2010

We have designed new double stimuli-responsive polyelectrolyte microcapsules to be useable under physiological conditions to handle biomacromolecules while avoiding the risk of denaturation. They are made of poly(4-vinylpyridine hydrochloride) (PVP) and poly(sodium styrene sulfonate) (PSS). The microcontainers are sensitive to temperature variation, as they irreversibly shrink under heating. In addition, the capsules reversibly swell at pH > 6, making it possible to encapsulate human serum proteins by diffusion through the polymer membrane. Encapsulation efficiency is quantified by fluorescence techniques.

Introduction The impact of potentially toxic agents such as metals or chemotherapeutic drugs is related to their interaction with biological species (proteins, oligonucleotides, etc.) by forming stable associative complexes. It is of the utmost importance to determine their specificity, their stoichiometry, and the thermodynamic constants of the complexes to understand these interactions and prevent related toxicity. However, biological studies are often limited by the availability of highly purified biological material, numerous experimental conditions, and sometimes by legal restrictions when harmfulness has to be considered. One of the major challenges is to determine these parameters related to affinity by quantifying free and bound ligands while avoiding changes to equilibrium. Classical biophysical methods (such as calorimetry, spectroscopic studies, etc.) are inappropriate for small quantities of proteins. Equilibrium dialysis is the reference method to perform this type of analysis, but it remains also restricted by the rare availability of highly purified biomacromolecules in massive quantities. Moreover, as in the field of nuclear toxicology, the use of highly toxic and radioactive substances (for example, transuranium actinides) is usually limited to very small quantities and restricted to specially equipped laboratories, where space optimization and easy handling remain major prerogatives. To this end, development of dialysis microsystems is crucial for laboratories studying noncovalent interactions between biomacromolecules (proteins, oligonucleotides) and small ligands such as metal cations. Taking all these constraints into account, we are focusing on polyelectrolyte microcapsules. They will be used as microcontainers and microreactors for the microencapsulation of biomacromolecules and the study of their interaction with small ligands based on equilibrium dialysis. Polyelectrolyte microcapsules have gained a great deal of interest in the past decade since Sukhorukov et al. applied the “layer-by-layer” technique to colloidal particles used as sacrificial templates.1-3 These * To whom correspondence should be addressed. E-mail: dejugnat@ chimie.ups-tlse.fr. † DSV/iBEB/SBTN/LEPC. ‡ ICSM, UMR 5257 CEACNRS-UM2-ENSCM. § Present address: Laboratoire des IMRCP, UMR 5623, 118, route de Narbonne - Baˆtiment 2R1, 31062 Toulouse, France.

microcapsules are now being increasingly developed in the fields of biotechnology, drug delivery, and for biomedical applications.4-8 Considered as free-standing polyelectrolyte multilayers, capsules present remarkable properties, with the most important being the selective permeability of membranes: small species (such as ions and organics) easily diffuse through the wall, whereas larger macromolecules (synthetic polymers, proteins, and DNA) do not.3 With the aim of using such microcontainers for dialysis experiments, the main constraints are, therefore, the following: (i) polyelectrolyte microcapsules should be easy to prepare and they must be monodisperse in size to allow easy quantitative measurements; (ii) encapsulation must be performed under conditions close to physiological ones (regarding pH and ionic strength) to limit stress and consequent aggregation or denaturation of the biological material, which is to be kept in as free a “natural” state as possible; (iii) handling should be as easy as possible and quantities of materials as small as possible. Biomacromolecule encapsulation in polyelectrolyte microcapsules is not basically new and has been largely investigated.4 However, the different protocols described must be considered in light of the above-mentioned constraints. A first set of studies referred to the use of calcium carbonate cores acting as both sacrificial templates and porous materials for protein accumulation (by diffusion or coprecipitation).9-11 The main advantage of this technique is that the experimental conditions are usually flexible enough to prevent protein denaturation. The resulting loaded microcapsules, however, present quite a broad size distribution and their shapes are only roughly spherical at best. Other techniques have been reported in which the capsules are formed around immobilized proteins. Although protein entrapment is efficient, the capsules obtained are highly polydisperse or solvent contacts damage the proteins.12-14 Another approach consists of tuning the membrane permeability of preformed hollow polyelectrolyte microcapsules and this has led to the development of stimuli-responsive capsules.15-18 However, a major concern is about the experimental conditions applied to switch the capsule permeability that often appear to be too extreme regarding protein stability: high or low pH, high temperature, high ionic strength, or the use of pure organic solvents.19,20 The limitation here is to work in a buffered system containing salt and having a pH close to physiological: proteins must be incorporated into the capsules very gently.

10.1021/bm901429q  2010 American Chemical Society Published on Web 02/04/2010

Microcontainers for Protein Soft Encapsulation

Tuning membrane permeability therefore appears to be the key point for encapsulation, transport, and release of (bio)macromolecules, bearing in mind that the biological material should be maintained close to its natural state. Moreover, once encapsulated, it is expected to potentially interact with small ligands present in the surrounding medium and equilibrium dialysis may potentially take place. Here we present stimuliresponsive capsules, potentially useable as microdialysis systems, for studying protein/metal interactions. We show that these microcontainers can be filled with proteins, although encapsulation efficiency could be optimized. Both encapsulation and conditions for use are compatible with protein stability, providing a general tool for quantifying biomacromolecule/diffusible ligand interactions.

Experimental Section Materials. PVP (MW ) 60 kDa), PSS (MW ) 70 kDa), FITC-dextran (MW ) 70 kDa), TRITC-dextran (MW ) 70 kDa), 2-morpholinoethanesulfonic acid monohydrate (MES), N-(2-hydroxyethyl)piperazineN′-2-ethanesulfonic acid (HEPES), sodium hydrogencarbonate (NaHCO3), N-methyl pyrrolidone (NMP), dimethylsulfoxide (DMSO), ethanol, sodium chloride (NaCl), sodium hydroxide (NaOH), and hydrochloric acid (HCl) were purchased from Sigma-Aldrich (France). Monodisperse polystyrene (PS) particles, diameter ) 9.61 ( 0.10 µm, were obtained from Microparticles GmbH (Berlin). Model proteins were purchased from Sigma-Aldrich (fetuin (MW ) 48 kDa), human serum albumin (MW ) 67 kDa), apo-transferrin (MW ) 80 kDa), glucose oxidase (MW ) 160 kDa)) or GE Healthcare (Rchymotrypsinogen (MW ) 25 kDa) and thyroglobulin (MW ) 670 kDa)). Fluorescent labels were obtained from Invitrogen in N-succinimidyl ester reactive form: Alexa 405 (λex ) 401 nm, λem ) 421 nm), Alexa 488 (λex ) 495 nm, λem ) 519 nm), and Alexa 633 (λex ) 632 nm, λem ) 647 nm). The water used in all the experiments was produced by a Millipore Milli-Q Plus 185 purification system and has a resistivity level higher than 18.2 mΩ · cm. Protein Labeling and Purification. Various Alexa-type fluorescent labels were coupled to proteins with different molecular weights for further use in the determination of capsule permeability by fluorescence microscopy. The following general procedure was applied in each case: 1 mg of fluorescent probe was dissolved in 100 µL of dry DMSO (stored over 4 Å molecular sieves) to give a 10 mg/mL solution. Then part of this solution was added in three steps over 15 min to 375-400 µL of a protein solubilized in 0.1 M NaHCO3 buffer at pH ) 8.3 with magnetic stirring. The volume of Alexa solution added was adjusted depending on the protein concentration and the targeted final molar ratio between the protein and the label. For each protein, the ratio was 1 label per 100 kDa. Alexa-labeled proteins were purified by size exclusion chromatography using an AKTA Purifier 10 system coupled to a size exclusion column type HR 10.10 (volume ) 8 mL) containing a Sephadex G-25 Superfine gel (all these elements were purchased from GE Healthcare). Mobile phase was a 50 mM HEPES buffer in 150 mM NaCl at pH ) 7.4. UV-Vis Absorption Spectrophotometry. UV-vis spectra were recorded using a Cary 50 spectrophotometer from Varian (France). The samples were placed into low volume quartz cuvettes from Hellma (France). Confocal Laser Scanning Microscopy (CLSM) and Fluorescence Microscopy. Optical images of polyelectrolyte capsules in solution were obtained using an Eclipse TE 2000-E confocal scanning system (Nikon, France) equipped with a 100×/1.4-0.7-oil immersion objective working in either confocal mode, transmission fluorescence mode, or transmission phase contrast mode. Scanning Electron Microscopy (SEM). Polyelectrolyte capsules were imaged under low water pressure using a FEI QUANTA 200 ESEM FEG scanning electron microscope working under environmental

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conditions (therefore, no metallization processes was required to image the samples). Acceleration voltage was between 9 and 30 kV, depending on the sample density. Capsule Preparation. Capsule assembly was based on the wellknown layer-by-layer technique. Because the PS cores bear a negative charge, the first layer was made of PVP. (PVP/PSS)8 multilayers were prepared by alternating adsorption of PVP and PSS onto the surface of PS microparticles from 2 mg · mL-1 polyelectrolyte solutions in 0.2 M NaCl. The pH of the PVP dipping solution was adjusted to 4 to ensure almost complete protonation of the pyridiniums groups. Nonadsorbed polymers were washed out twice with water at each adsorption step after centrifugation at 2000 g for 1 min. After deposition of the final layer, the PS core was removed by dissolution in NMP followed by three washes in NMP and two washes in water. For this procedure, the centrifugation steps were conducted at a reduced acceleration (200 g) but for a longer time (20 min) to prevent hollow capsule deformation. Capsules Counting. Capsule concentration was determined using a Malassez hematocymeter, composed of two 1 mm3 chambers. Each chamber is itself divided into compartments with a volume of 0.5 × 10-3 mm3. Typically, 12 µL of a diluted capsule suspension are deposited in one chamber, and after 10 min sedimentation the capsules are manually counted using a microscope. Capsule Dissolution for Mass Determination. Capsule mass was determined by global quantification of polymers in a sample with a known number of capsules. Measurement was made by UV-vis spectrophotometry. First a calibration curve was determined using equimolar mixtures of PSS and PVP at concentrations ranging between 10 and 100 µg/mL in a solvent composed of 33% water and 67% 1 mM NaOH in ethanol. Under these conditions, polyelectrolytes fully dissociate and remain perfectly soluble. A linear regression relating absorbance at 257 nm as a function of concentration gave the massic absorption coefficient: ε ) 9.6 × 10-3 mg-1 · L · cm-1. Then the sample containing the capsules could be analyzed: typically, 60 µL of capsule aqueous suspension was added to 120 µL of 1 mM NaOH in ethanol. After 20 min, the capsules were totally dissolved and absorbance at 257 nm was recorded. Knowing the capsule concentration, one could then calculate the mean mass of the capsule. Temperature Treatment. The tubes containing the capsule suspension are heated while shaking using a Thermomixer at 300 rpm for 30 min and at 70 °C (or other temperature when studying the temperature effect). Protein Encapsulation. Typically, 20 µL of resealed capsules (heated at 70 °C for 30 min) suspended in water (concentration is 35 millions capsules per mL) are mixed with 35 µL of Hepes buffer (40 mM) containing 50 mM NaCl at pH ) 6. A total of 10 µL of labeled protein (3 to 16 mg/mL) stored in Hepes buffer (50 mM) containing 150 mM NaCl at pH ) 7.4 is added to this mixture. In the global mixture, the total buffer concentration is 29 mM, NaCl concentration is 50 mM, and pH ) 6.7 (this is the highest value that can be used before capsule dissolution). The protein concentration is 0.5-2.5 mg/ mL in the mixture. The mixture is shaken at 300 rpm for 20 min at 20 °C. Finally, the “closing” solution is added: 15 µL of 200 mM MES containing 50 mM NaCl at pH ) 3. This allows the pH to fall to 5.1 without significantly changing ionic strength. The final mixture is shaken for 5 min. Then the capsules are centrifuged at 200 g for 20 min at 20 °C, the supernatant is removed and replaced by a 20 mM MES solution in 50 mM NaCl at pH ) 5. This operation is performed twice to ensure complete washing of nonencapsulated protein. Encapsulated Protein Concentration Determination. CLSM was used for in situ determination of protein concentration inside the capsules. A calibration curve was produced using labeled protein solutions and measuring fluorescence intensity at different concentrations (concentrations were low enough to obtain a linear relationship between intensity and concentration). Then the filled capsules were imaged under exactly the same conditions so that the fluorescence emitted by the labeled encapsulated protein could be directly related to its concentration inside the capsule.

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Figure 1. (a) Structures of the polyelectrolytes used in this study. (b) pH-induced encapsulation protocol of biomacromolecules: initially, the capsules are not permeable to the proteins, which remain outside (A), then the permeability is reversibly tuned by pH variations (B, C), and finally, the nonencapsulated remaining protein is washed out (D).

Results and Discussion 1. Preparation and Characterization of the Microcapsules. Choice of the Sacrificial Template and ConstitutiVe Polyelectrolytes. With a view to controlling and performing accurate quantitative characterization of protein encapsulation and later to using the loaded microreactors as microdialysis systems, the capsules must be regular in size and spherical. Of the available spherical monodisperse cores, melamine-formaldehyde (MF) particles were not selected because their dissolution is not complete and non-negligible amounts of oligomers remain as a gel inside the capsule.21 Even though they might be considered as the templates of choice for our study, spherical silica microparticles were also avoided because they require the use of hydrofluoric acid aqueous solutions for dissolution, which imposes special equipment and severe security restrictions. Our choice, then, was to use PS latexes as sacrificial templates, as they have been shown to produce really hollow spherical capsules after complete elimination of the core by dissolution with organic solvents.22 Protein stability is an essential parameter that restricts experimental conditions during encapsulation: as we plan to work with human serum proteins and keep their natural structure, limiting any risk of denaturation or aggregation, ionic strength should be maintained between 50 and 150 mM, the pH should not reach values outside a 5-8 window, the temperature should be maintained below 40 °C, and the medium must remain 100% aqueous. These specific conditions exclude almost all the stimuli-responsive-based encapsulation protocols described and drastically reduce the choice of constituent polyelectrolyte. We therefore decided to focus on a pH-sensitive system,23-25 which can be used under physiological conditions, based on a strong polyanion and a weak polycation, namely, PSS and PVP

(chemical structures are shown in Figure 1a). The principle is to tune the reversible permeability of the polymeric film to allow the protein diffusion from the outside of the capsule to the inside before closing the membrane and then keeping the material entrapped,23 as illustrated in Figure 1b. The pH sensitivity should be set by PVP, which has a pKa reported to be close to 5. Its degree of ionization will therefore depend on pH being in the neutral range.25-27 Preparation of PSS/PVP Hollow Microcapsules. PS template removal was not possible using classic tetrahydrofuran treatment22 because core dissolution was accompanied by a collapse of the multilayers, leading to micrometer-sized compact polymer globules. A gentler dissolution process used N-methyl pyrrolidone (which is much less toxic than N,N-dimethylformamide). After several washes, it produced spherical hollow microcapsules, as evidenced by microscopy (Figure 2). The solvent effect is a first sign that the PSS/PVP membrane is mechanically weaker than the reference PSS/PAH film. This could be due to a better charge mismatch in the PSS/PAH system than in PSS/ PVP, in which steric hindrance of aromatic phenyl and pyridiniums could induce defects in charge pairing. This is supported by the fact that the minimal number of bilayers required to produce stable capsules 10 µm in diameter was six in the case of PSS/PVP compared to four when using PSS/PAH (the comparison is based on the use of polyelectrolytes of similar chain lengths).22 Although, with smaller PS templates, fewer layers were needed to ensure film stability (data not shown), we decided to focus on 10 µm capsules made of eight polyelectrolyte bilayers, namely, (PSS/PVP)8. Capsule Weight Determination. Bearing in mind that (PSS/ PVP)8 capsules will be used in microdialysis experiments, they will be immersed in solutions of the ligands with which they



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Figure 2. (PSS/PVP)8 microcapsules obtained after core dissolution, observed by optical microscopy (left) or confocal laser scanning microscopy (right, lower scale), evidencing hollow structure and size monodispersity. Scale bars represent 10 µm.

might interact more or less. It is therefore important to determine the amount of polymeric material present in the membrane of each individual capsule. To measure this weight, a known quantity of microcapsules was dissolved in an alkaline alcoholic solution to ensure both complete separation and solubilization of polyelectrolytes. Sodium hydroxide has been used to neutralize the pyridinium form of PVP: the acid-base reaction produced the uncharged form of PVP, leading to its separation with PSS. Ethanol is a suitable solvent for this uncharged form of PVP, preventing its precipitation in the solution. The absorption spectrum of this resulting solution was recorded and compared to a reference titration curve of equimolar amounts of PSS and PVP under the same conditions. The hypothesis that (PSS/PVP) multilayers are made of equimolar amounts of PSS and PVP is supported by the fact that the molar ratio between the polycation and the polyanion in various types of polyelectrolyte complexes and microcapsules has been reported to be close to 1.28-30 This simple analysis led to a mass of 10 pg per (PSS/PVP)8 capsule, which is consistent with other values reported in the literature: for 5 µm (PSS/PDADMAC)4 (PDADMAC: poly(diallyldimethylammonium chloride)) microcapsules (half the bilayer number and a quarter of the covered surface, overall eight times less polymer, compared with the present case), capsule weights were determined to be 1.25 pg.31 It should be noted here that this indirect weighing technique is very simple and could become one of the only suitable methods when working under special conditions such as glove boxes in nuclear installations. Capsule Permeability and Resealing Attempts. Confocal microscope observations of (PSS/PVP)8 capsules immersed in aqueous solution of fluorescently labeled protein revealed that up to 70% of the microcapsules were permeable to various proteins (fetuin, human serum albumin, transferrin) and labeled 70 kDa dextran, although about 30% remained impermeable (Figure 3, left). This observation shows that the capsules might be partially defective during core dissolution and cannot be used in that form for encapsulation experiments as described in Figure 1b. This is another proof that PSS/PVP multilayers are less compact and more porous than the reference PSS/PAH system. To repair the defects in the membrane, we tried to deposit additional polyelectrolyte layers on the top of the hollow

capsules, as had already been successfully shown in the case of (PSS/PAH) capsules.22 However, this attempt only led to the destruction of the capsules, even after the first additional layer. Another way to fill pores in polyelectrolyte multilayers is to increase the ionic strength of the surrounding solution. This can be used to slightly decrease electrostatic interactions between oppositely charged polyelectrolytes to make them more fluid; in this way, polymer chains can diffuse and partially fill the pores. However, the addition of salt (0.2 M) had a very similar effect in the case of additional layers: most of the capsules rapidly collapsed. The ultimate alternative was to heat the capsules to bring sufficient transient fluidity to the system. In the case of PSS/PDADMAC capsules, it was shown that annealing such systems led to more compact multilayers with decreased permeability, even for small molecules.32 For PSS/ PVP capsules, the effect of temperature was similar but not so dramatic. Indeed, permeability was slightly reduced compared with the initial capsules, but it never led to 100% impermeable capsules: at best, heating at 70 °C for 30 min afforded about 30% resealed capsules, as visualized by CLSM (Figure 3, right). However, this annealing was chosen as a preliminary treatment of the capsules before encapsulation, as described later in the paper. 2. Capsule Response to External Stimuli. Temperature Effect. Heat-induced shrinking appears to be irreversible, as cooling back the samples to room temperature (or even at 4 °C) did not induce any swelling of the capsules to the initial size. This observation shows that the phenomenon is thermodynamically driven and that the initial state is not at thermodynamic equilibrium. In a first experiment, the capsule diameter was measured after 30 min annealing the capsules at different temperatures from 25 up to 95 °C (Figure 4a,b). In a first regime capsules start to shrink when the temperature increases above 55 °C. This threshold temperature should correspond to a glass transition (this has been observed to occur at about 35-40 °C for the PSS/PDADMAC system).33 The heating-induced size reduction and permeability decrease can be correlated to the well described PSS/PDADMAC system.30,32,34 Capsules made of an even number of PSS and PDADMAC layers irreversibly shrink upon heating, and this has been attributed to competition between electrostatic and hydrophobic forces. In our case, the

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Figure 3. Confocal laser scanning microscopy observations of (PSS/PVP)8 microcapsules in the presence of fluorescently labeled 70 kDa dextran. Most of the initial microcapsules are permeable (left), whereas heating at 70 °C for 30 min leads to about 30% impermeable shrunk capsules (right). Scale bars represent 10 µm.

Figure 4. (a) Evolution of capsule size after heating for 30 min at different temperatures and the corresponding CLSM (b) and SEM (c) micrographs. (d) Evolution of capsule size when heating at 70 °C as a function of time. Scale bars in (b) represent 10 µm.

same evolution is primarily observed but both the size profile and the microscope observations have evidenced a phase transition at about 82 °C. Above this temperature, the capsules collapse in a kind of “implosion”: although they globally maintain a globular shape, new features can be observed inside the capsules. Moreover, capsule diameter seems to reach a minimum size (3.5 µm). In this second regime, hydrophobic interactions as well as dehydration could imply PVP precipita-

tion (this polymer is highly hydrophobic). This could be attributed to partial desorption of the inner layers that rearrange in a kind of polymer matrix in the capsule lumen. Scanning electron microscopy was performed on dry samples and clearly shows the mechanical resistance acquired by capsules upon heating. Initially nonheated capsules collapse completely upon drying, and capsules heated to 70 °C also collapse, but the membrane is clearly thicker and more resistant; globules


obtained after heating the capsules to 90 °C do not collapse and present a rougher surface (Figure 4c). The capsules’ temperature sensitivity could have been used to reseal the capsules in the presence of proteins and achieve direct heatinginduced encapsulation, but it is clearly too high to avoid biomacromolecule denaturation. We tried to decrease this glass transition temperature by heating in the presence of 0.1-0.2 M NaCl, with a view to achieving heat-induced direct protein encapsulation. In that case, ionic strength was low enough to prevent protein denaturation but assumed to enhance polymer fluidity by screening interpolyelectrolyte interactions. However, we observed partial collapse and deformation of some capsules. The salt effect leads to more constrained capsules, and therefore, annealing in pure water will be preferred. Then in a second set of experiments the decrease in size was recorded at 70 °C as a function of incubation time (Figure 4d). The shrinking is fast during the first 10 min then stabilizes after 3 h, leading to capsules with a final size of about 6 µm. This prolonged heating induced noticeable capsule deformation. We can use the heating effect to temporarily soften the polymeric membrane that increases fluidity to make the polymer chains partially fill the pores in the multilayer film. However, to use these capsules as microcontainers, we have to stop the annealing before the irreversible phase transition. For that reason, the partial “repairing” of capsules was chosen, achieved by heating at 70 °C for 30 min. pH effect. Capsules in which at least one of the constituents is a weak electrolyte are sensitive to pH changes. In the present system, PSS is a strong polyelectrolyte, whereas PVP is a weak polyelectrolyte that can be partially, then fully, deprotonated upon increasing pH. Figure 5a shows capsule size variation as a function of pH. In fact, PSS/PVP capsules, whether heated or not, swell at pH values higher than 6, and maximum swelling is observed at pH ) 6.7 in a 50 mM buffer solution, with a final size of about 11 µm. Above this threshold the capsules dissolve. The swelling remains limited to 120% for the initial capsules, whereas heated capsules can swell up to 165% (Figure 5b). This phenomenon is driven by an electrostatic imbalance due to variations of charge density (Figure 5c). It usually leads to swelling of the polyelectrolyte multilayers, and in the case of capsules, the mean diameter increases when pH reaches a critical value. Due to the electrostatic nature of the polyelectrolyte complex, heat-shrunk capsules also swell and the relative swelling is constant for capsules that have been heated for different times. It shows that heating above the glass transition has an effect on the relative swelling, but the driving force (electrostatics) leads to a common stable swollen state whether the capsules were heated or not. This pH effect has already been described for other microcapsules containing at least one weak polyelectrolyte, such as PAH,22 poly(methacrylic acid) (PMA),35,36 or PVP.25 In the pH-induced swollen state, the capsules are much more permeable to macromolecules, and this kind of system has previously been successfully used for macromolecule encapsulation in (PSS/PAH) capsules.23 As the swelling of (PSS/PVP)8 capsules is reversible, permeability is assumed to decrease when reducing the pH to 6. Because this system is reversibly sensitive in the neutral pH range, it therefore appears to be a potentially good system for biomacromolecule encapsulation under physiologically compatible conditions. 3. Protein Encapsulation. During protein encapsulation it was necessary to work in a narrow pH range and maintain reasonable ionic strength. It is a compromise between changes in environmental conditions, required for permeability change,


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and protein stability. To achieve such encapsulation, pH variations were controlled using buffer mixtures, maintaining relatively stable ionic strength. With the same objective, several final washes were performed in a buffer solution, which is the final stocking solution. Encapsulation experiments were conducted using various fluorescently labeled proteins: R-chimotrypsinogen (Chymo, 25 kDa), fetuin (Fet, 48 kDa), human serum albumin (HSA, 67 kDa), transferrin (Tf, 80 kDa), glucose oxidase (Gox, 160 kDa), and thyroglobulin (TG, 670 kDa). Changing the molecular weight affects encapsulation efficiency: Chymo was never observed within the capsules (after washes), which proves that encapsulation failed, in contrast to all other larger probed proteins (up to 670 kDa). In the case of small proteins, fast exchanges between the inner compartment and the outer medium cannot be excluded. From this point of view, the membrane of (PSS/PVP)8 capsules presents a molecular weight cut-off value of between 25 and 48 kDa. Moreover, the larger proteins (Tf and larger) were often observed as precipitates inside the capsules. Fet and HSA remained as free solutions (homogeneous fluorescence). Visually, confocal microscope observations using labeled proteins revealed that at best 25% of the capsules contained the proteins (Figure 6). This can be related to the initial permeability of the heated capsules, that was about 30%. So we can conclude that, once repaired, almost 80% of the capsules can be filled with proteins using this pH-driven encapsulation. Another major observation is the strong fluorescence of the capsule walls. This can reveal either strong nonspecific protein adsorption on the polyelectrolyte multilayers or entrapment of the proteins within the polymer film (proteins are also polyelectrolyte and certainly compete with PVP and PSS in the multilayered assembly). Moreover, it appears that capsules do not shrink back to their initial size upon reducing pH in the presence of proteins: the final diameter is about 9-10 µm. This confirms that proteins act as competitive polyelectrolyte and some of them “repair” defective capsule membranes by interacting with PSS and PVP and filling the pores. Regarding isoelectric points, no effect was observed and this might be related to unsymmetrical charge distribution on the protein surface: whatever the pH, there are different protein parts with opposite charges and biomacromolecules always have the chance to interact with the polyelectrolyte multilayers by electrostatic interaction. Fluorescence quantitative analysis was conducted to estimate the amount of encapsulated material. We decided to rupture the capsules in order to release the encapsulated labeled proteins and then measure the fluorescence of the supernatants. Although native unheated capsules are easily ruptured by simply freezing them at -20 °C, capsules treated at 70 °C appeared to be much more resistant: seven freeze/thaw cycles (-80 to 37 °C) were required. Such enhanced mechanical properties are very interesting for further separation processes such as centrifugation. The opened capsules were then centrifuged and the supernatant analyzed by fluorescence. However, this technique was not reliable, probably due to the nonspecific adsorption of very small amounts of proteins onto the surface of the microtubes, and the results were never reproducible. Moreover, nonspecifically adsorbed proteins on capsules may also interfere with released proteins. To overcome this problem, we made direct semiquantitative measurements using a confocal laser scanning microscope, measuring the fluorescence emitted from the inner part of the capsules. This fluorescence was then compared with standard protein solutions used for calibration. The results show that about 0.5 pg of protein was encapsulated in each filled

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Figure 5. (a) Evolution of size as a function of pH for nonheated capsules (filled circles) and capsules heated at 70 °C for 30 min (filled squares) or for 180 min (filled triangles). (b) Relative size evolution of the same capsules, taking the initial size at pH ) 5.5 as the 100% reference level (circles, squares, and triangles refer to the same samples depicted in (a)). (c) Schematic representation of electrostatic repulsions that destabilize LbL assembly upon increasing pH.

capsule, corresponding to an inner protein concentration ranging from 0.5 to 1 mg/mL. The final concentration was globally dependent on the starting protein solution, but we did not observe higher values than 1 mg/mL inside the capsules. For example, an encapsulation realized with 0.65 mg/mL HSA (concentration during capsule opening) led to a 0.5 mg/mL HSA solution inside the capsules. Increasing the bulk protein concentration up to 2.5 mg/mL led to a maximal 1 mg/mL encapsulated protein solution. This threshold might be due to the internal osmotic pressure imposed by the encapsulated protein. However, these preliminary results are comparable to values reported in the literature: Mak et al. described the

encapsulation of labeled bovine serum albumin at 0.125 mg/ mL into capsules displaying an equivalent diameter;12 Li et al. indicated a lysozyme concentration of 0.6 mg/mL inside 9.8 µm capsules before optimization.11

Conclusion In this paper we have described the preparation and use of double stimuli-responsive polyelectrolyte microcapsules useable under physiological conditions for protein encapsulation and further use as microdialysis systems. Capsules assembled from PSS and PVP are temperature-sensitive as they shrink upon



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Figure 6. (a) Confocal laser scanning microscope observation of capsules after encapsulation of fluorescently labeled HSA; white arrows indicate filled capsules, which represent about 25% of the total amount of capsules. (b) Close-up of CLSM after fetuin encapsulation with fluorescence intensity profiles showing whether the protein is internalized (top capsule) or not (bottom capsule). Scale bars in both micrographs represent 10 µm.

heating. In addition, the same capsules, heated or not, are reversibly sensitive to pH changes, and their size and permeability can be increased by increasing pH. Due to the imposed constraints of mild encapsulation conditions, we focused on protein entrapment into preformed hollow capsules, but some problems have arisen, especially due to damage during core dissolution and still perfectible membrane repair. However, we succeeded in achieving protein encapsulation by applying adequate pH shifts to (PSS/PVP)8 capsules, first allowing the macromolecule to diffuse through the membrane, followed by a “closing step” that sealed the larger pores of the container, keeping the protein entrapped. The final protein concentration

inside the capsules was evaluated to be about 1 mg/mL. Moreover, the final capsules display good mechanical strength that makes them easy to handle. To the best of our knowledge, this is the first time that microencapsulation of proteins has occurred under such mild conditions using doubly stimulatable polyelectrolyte microcapsules. This approach already paves the way for new fields of study, such as characterizing interactions between proteins and metal cations or between DNA and chemotherapeutic agents. System optimization is under way, especially reduction of interactions between proteins and capsule membrane by polyelectrolyte modification (PEGylation) and reduction of protein/protein interactions by addition of well-

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known additives (trehalose or “soft” uncharged surfactants like n-octyl-β-D-glucopyranoside).37,38 The system will further be validated as a microdialysis system by quantifying protein/uranyl interactions that have already been demonstrated using chromatographic techniques.39-41 Acknowledgment. This work was supported by the “Environmental Toxicology Program” of the Commissariat a` l’Energie Atomique” (CEA). Dr. R. Podor is greatly thanked for electron microscopy observations.

References and Notes (1) Multilayer Thin FilmssSequential Assembly of Nanocomposite Materials; Decher, G., Schlenhoff, J. B., Eds.; Wiley-VCH: Weinheim, 2003. (2) Johnston, A. P. R.; Cortez, C.; Angelatos, A. S.; Caruso, F. Curr. Opin. Colloid Interface Sci. 2006, 11, 203–209. (3) Peyratout, C. S.; Da¨hne, L. Angew. Chem., Int. Ed. 2004, 43, 3762– 3783. (4) Sukhorukov, G. B.; Mo¨hwald, H. Trends Biotechnol. 2007, 25, 93– 98. (5) Liu, X. D.; Yu, W. T.; Wang, W.; Xiong, Y.; Ma, X. J.; Yuan, Q. A. Prog. Chem. 2008, 20, 126–139. (6) Matsusaki, M.; Akashi, M. Exp. Opin. Drug DeliVery 2009, 6, 1207– 1217. (7) De Geest, B. G.; Sukhorukov, G. B.; Mo¨hwald, H. Exp. Opin. Drug DeliVery 2009, 6, 613–624. (8) De Geest, B. G.; De Koker, S.; Sukhorukov, G. B.; Kreft, O.; Parak, W. J.; Skirtach, A. G.; Demesteer, J.; De Smedt, S. C.; Hennink, W. E. Soft Matter 2009, 5, 282–291. (9) Volodkin, D. V.; Larionova, N. I.; Sukhorukov, G. B. Biomacromolecules 2004, 5, 1962–1972. (10) Petrov, A. I.; Volodkin, D. V.; Sukhorukov, G. B. Biotechnol. Prog. 2005, 21, 918–925. (11) Li, X.; Li, X.; Zhang, J.; Zhao, S.; Shen, J. J. Biomed. Mater. Res. 2008, 85A, 768–776. (12) Mak, W. C.; Bai, J.; Chang, X. Y.; Trau, D. Langmuir 2009, 25, 769– 775. (13) Kim, B. S.; Oh, J. M.; Kim, K. S.; Seo, K. S.; Cho, J. S.; Khang, G.; Lee, H. B.; Park, K.; Kim, M. S. Biomaterials 2009, 30, 902–909. (14) Giteau, A.; Venier-Julienne, M. C.; Marchal, S.; Courthaudon, J. L.; Sergent, M.; Montero-Menei, C.; Verdier, J. M.; Benoit, J. P. Eur. J. Pharm. Biopharm. 2008, 70, 127–136. (15) Sukhishvili, S. Curr. Opin. Colloid Interface Sci. 2005, 10, 37–44. (16) Kozlovskaya, V.; Kharlampieva, E.; Mansfield, M. L.; Sukhishvili, S. A. Chem. Mater. 2006, 18, 328–336. (17) Tong, W. J.; Gao, C. Y. J. Mater. Chem. 2008, 18, 3799–3812.

Basset et al. (18) Kozlovskaya, V.; Kharlampieva, E.; Erel, I.; Sukhishvili, S. A. Soft Matter 2009, 5, 4077–4087. (19) Lvov, Y.; Antipov, A. A.; Mamedov, A.; Mo¨hwald, H.; Sukhorukov, G. B. Nano Lett. 2001, 1, 125–128. (20) Balabushevich, N. G.; Tiourina, O. P.; Volodkin, D. V.; Larionova, N. I.; Sukhorukov, G. B. Biomacromolecules 2003, 4, 1191–1197. (21) Dong, W. F.; Ferri, J. K.; Adalsteinsson, T.; Scho¨nhoff, M.; Sukhorukov, G. B.; Mo¨hwald, H. Chem. Mater. 2005, 17, 2603–2611. (22) De´jugnat, C.; Sukhorukov, G. B. Langmuir 2004, 20, 7265–7269. (23) De´jugnat, C.; Halozˇan, D.; Sukhorukov, G. B. Macromol. Rapid Commun. 2005, 26, 961–967. (24) Tong, W.; Gao, C.; Mo¨hwald, H. Colloid Polym. Sci. 2008, 286, 1103– 1109. (25) Mauser, T.; De´jugnat, C.; Sukhorukov, G. B. J. Phys. Chem. B 2006, 110, 20246–20253. (26) Satoh, M.; Yoda, E.; Hayashi, T.; Komiyama, J. Macromolecules 1989, 22, 1808–1812. (27) Franck-Lacaze, L.; Sistat, P.; Hughet, P. J. Membr. Sci. 2009, 326, 650–658. (28) Petrov, A. I.; Antipov, A. A.; Sukhorukov, G. B. Macromolecules 2003, 36, 10079–10086. (29) Smith, R. N.; Reven, L.; Barrett, C. J. Macromolecules 2003, 36, 1876– 1881. (30) Ko¨hler, K.; Shchukin, D. S.; Mo¨hwald, H.; Sukhorukov, G. B. J. Phys. Chem. B 2005, 109, 18250–18259. (31) De´jugnat, C.; Ko¨hler, K.; Dubois, M.; Sukhorukov, G. B.; Mo¨hwald, H.; Zemb, T.; Guttmann, P. AdV. Mater. 2007, 19, 1331–1336. (32) Ko¨hler, K.; Sukhorukov, G. B. AdV. Funct. Mater. 2007, 17, 2053– 2061. (33) Mu¨ller, R.; Ko¨hler, K.; Weinkamer, R.; Sukhorukov, G. B.; Fery, A. Macromolecules 2005, 38, 9766–9771. (34) Ko¨hler, K.; Biesheuvel, P. M.; Weinkamer, R.; Mo¨hwald, H.; Sukhorukov, G. B. Phys. ReV. Lett. 2006, 97, 188301. (35) Mauser, T.; De´jugnat, C.; Sukhorukov, G. B. Macromol. Rapid Commun. 2004, 25, 1781–1785. (36) Mauser, T.; De´jugnat, C.; Mo¨hwald, H.; Sukhorukov, G. B. Langmuir 2006, 22, 5888–5893. (37) Maury, M.; Murphy, K.; Kumar, S.; Mauerer, A.; Lee, G. Eur. J. Pharm. Biopharm. 2005, 59, 251–261. (38) Jain, N. K.; Roy, I. Eur. J. Pharm. Biopharm. 2008, 69, 824–834. (39) Vidaud, C.; Dedieu, A.; Basset, C.; Plantevin, S.; Dany, I.; Pible, O.; Que´meńeur, E. Chem. Res. Toxicol. 2005, 18, 946–953. (40) Basset, C.; Dedieu, A.; Guerin, P.; Que´meńeur, E.; Meyer, D.; Vidaud, C. J. Chromatogr., A 2008, 1185, 233–240. (41) Dedieu, A.; Berenguer, F.; Basset, C.; Prat, O.; Que´meńeur, E.; Pible, O.; Vidaud, C. J. Chromatogr., A 2009, 1216, 5365–5376.

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