Intracellular Processing of Proteins Mediated by Biodegradable

Multilayer polyelectrolyte capsules made by layer-by-layer assembly of oppositely charged biodegradable polyelectrolytes were filled with a model of a...
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NANO LETTERS

Intracellular Processing of Proteins Mediated by Biodegradable Polyelectrolyte Capsules

2009 Vol. 9, No. 12 4398-4402

Pilar Rivera-Gil,† Stefaan De Koker,‡ Bruno G. De Geest,*,§ and Wolfgang J. Parak*,† Fachbereich Physik und Wissenschaftliches Zentrum fu¨r Materialwissenschaften (WZMW), Philipps UniVersita¨t Marburg, Renthof 7, 35037 Marburg, Germany, and Department of Biomedical Molecular Biology and Laboratory of Pharmaceutical Technology, Ghent UniVersity, 9000 Ghent, Belgium Received August 18, 2009; Revised Manuscript Received October 9, 2009

ABSTRACT Multilayer polyelectrolyte capsules made by layer-by-layer assembly of oppositely charged biodegradable polyelectrolytes were filled with a model of a nonactive prodrug, a self-quenched fluorescence-labeled protein. After capsule uptake by living cells, the walls of the capsules were actively degraded and digested by intracellular proteases. Upon capsule wall degradation, intracellular proteases could reach the protein cargo in the cavity of the capsules. Enzymatic fragmentation of the self-quenched fluorescence-labeled protein by proteases led to individual fluorescence-labeled peptides and thus revoked self-quenching of the dye. In this way nonactive (nonfluorescent) molecules were converted into active (fluorescent) molecules. The data demonstrates that biodegradable capsules are able to convert nonactive molecules (prodrugs) to active molecules (drugs) specifically only inside cells where appropriate enzymes are at hand. In this way only cargo inside the capsules reaching cells is activated, but not the cargo in capsules which remain extracellular. The peptide fragments undergo further processing inside the cells, leading ultimately to exocytosis.

Introduction. The synthesis of multifunctional polyelectrolyte capsules has attracted enormous interest in the last years for a variety of different applications as is highlighted in a series of recent review and perspectives articles.1-7 They are synthesized by layer-by-layer assembly.8,9 This technique is based on the consecutive adsorption of oppositely charged polyelectrolytes around a preformed charged template core, which at the end of the synthesis is chemically dissolved.10,11 In the field of life sciences, applications of polyelectrolyte capsules are ranging from drug delivery,12 targeted gene therapy,13,14 molecular sensing,15-18 and vaccination19 to biosensor devices.20 The key point of the technology hereby is the possibility to load as well the walls of the capsules as their cavity with functional molecules or nanoparticles. Whereas embedding of functionalities in the capsule walls is more or less established, it still remains a challenge to efficiently host bioactive macromolecules within their cavity.21-23 Ultimately controlled release of the molecular cargo from the capsule cavity should be possible.24,25 This has been * Corresponding authors, [email protected] and [email protected]. † Fachbereich Physik und Wissenschaftliches Zentrum fu¨r Materialwissenschaften (WZMW), Philipps Universita¨t Marburg. ‡ Department of Molecular Biomedical Research, Ghent University. § Laboratory of Pharmaceutical Technology, Ghent University. 10.1021/nl902697j CCC: $40.75 Published on Web 10/27/2009

 2009 American Chemical Society

demonstrated by disintegrating the capsule walls with heat,26-28 ultrasound,29 and biodegradation.30-36 Polyelectrolyte capsules are known to be incorporated by most cells37 and do not cause acute cytotoxic effects,38 which makes them in principle suitable for intracellular applications. The idea to use the cavity of capsules for a spatially confined reaction inside the capsules has been proposed several years ago.39-43 In this way, precursor molecules (which we term here model prodrug) are locally reacted inside the capsules to products (which we term model drug). This led us to the idea to use capsules loaded with inactive molecules (as prodrug) whose conversion to active molecules (as drugs) is triggered inside cells, as has been already demonstrated with similar systems.44-46 This would ensure the presence of active molecules (drugs as cytostatica) only inside, but not outside cells, where the molecules would reside in their inactive form (as a prodrug). We loaded inactive molecules inside the cavities of biodegradable capsules. Once incorporated into cells these capsules were enzymatically degraded which in turn enabled access of intracellular enzymes into the capsule cavity. The inactive capsule cargo could then be converted to an active form by these enzymes. In this way activation exclusively takes place intracellularly.

Figure 1. Enzymatic cleavage of protein cargo. Embryonic NIH/3T3 fibroblasts were incubated with (a) nondegradable PSS/PAH or (b) degradable DEXS/pARG capsules filled with the fluorogenic protein cargo, DQ-OVA. Images were taken immediately after addition of the capsules (t ) 0 h) over time up to 120 h with a confocal microscope in different channels, green, red, and transmission (for a detailed description including movies, we refer readers to the Supporting Information). An overlay of the different channels is presented in the figures.

We chose the DQ-ovalbumine (DQ-OVA) conjugate as model of an inactive prodrug. DQ-OVA is composed out of the protein ovalbumine which is chemically saturated with BODIPY dyes. Due to the close proximity of dye molecules, their green fluorescence is almost completely self-quenched.47 Enzymatic degradation of DQ-OVA by proteases leads to dye-labeled peptide fragments; see Figure 1. This revokes the close proximity of the dye molecules and therefore relieves the quenching. In this way a nonfluorescent and thus inactive complex can be enzymaticly converted into fluorescent and thus active molecules. Conversion of DQ-OVA, as model for a prodrug, to fluorescent fragments, as model for an active drug, is restricted in two ways to the intracellular space. Only the wall of those capsules internalized by cells is degraded by intracellular proteases and only in this case intracellular proteases can also convert the nonfluorescent DQ-OVA into fluorescent fragments. Results. As already published, both sodium poly(styrenesulfonate)/poly(allylamine hydrochloride) (PSS/PAH) and dextran sulfate/poly-L-arginine (DEXS/pARG) capsules were taken up by living cells.30 Capsule incorporation, cellular division, growth, and movement were followed with confocal microscopy. According to cellular requirements, cells engulf the capsules located in their proximity. Upon cellular Nano Lett., Vol. 9, No. 12, 2009

division, each daughter cell takes part of the capsules internalized by their mother cell. These time-resolved measurements in living cells demonstrated that the engulfment of both types of capsules did not interfere acutely with the proliferation and viability of the cells. In agreement with other published results for PSS/PAH capsules37 capsule deformation was also observed for DEXS/pARG capsules. Upon incorporation of capsules by living cells, a strong deformation of the capsules occurs. Despite the mechanical stress suffered by the internalized capsules, there was no evidence of rupture of the capsule wall that could lead to an uncontrolled release of cargo release. The DQ-OVA remained trapped in the capsule cavity. When DQ-OVA is concentrated in a small volume, like the capsule cavity, the DQ fluorophore molecules form red fluorescent excimers. Therefore, capsules loaded with DQOVA which are located in the extracellular medium or which have been just incorporated by cells exhibit weak red fluorescence. In the case of the nondegradable PSS/PAH capsules this did not change over incubation periods up to several days; see Figure 1a. Cells proliferated and passed capsules to their respective daughter cells, and the red fluorescence of capsules did not alter. This was the expected behavior, as synthetic PSS/PAH capsules are not subject to 4399

cellular degradation.48 Therefore the capsule walls impose a barrier to intracellular enzymes which thus cannot reach the capsule cavities with the DQ-OVA cargo. The structural integrity of DQ-OVA remained unharmed and its green fluorescence remained quenched. The situation is changed in the case of biodegradable DEXS/pARG capsules; see Figure 1b. The quenching of the green fluorescence DQ-OVA is partially relieved after a few hours for internalized capsules. This can be explained by degradation of the capsule walls by intracellular enzymes, as shown before.30 Degradation of capsule walls opens a way for intracellular enzymes such as proteases to reach the DQOVA cargo. By enzymatic cleavage of DQ-OVA to single dye-labeled peptides, self-quenching of the DQ dye is relieved and a bright green fluorescent signal appeared, whereas the red fluorescence signal originating from excimers diminished. As a result, enzymatic processing of DQ-OVA took place intracellularly and could be measured by the turnover of red fluorescence (close proximity of adjacent DQ dyes) to green fluorescence (large distance between DQ dyes). The cavity of the capsules located extracellularly, i.e., the ones which had not been incorporated by the cells until this time, was not visible in the green channel but rather retained its original red fluorescence. These results demonstrate that DQ-OVA was intracellulary digested by enzymes, as DEXS/pARG capsules are susceptible for intracellular degradation. PSS/PAH capsules on the other hand were resistant against enzymatic degradation and, therefore, proteases could not reach the encapsulated DQ-OVA. According to these results only biodegradable DEXS/pARG but not nondegradable PSS/PAH capsules could trigger enzymatic cleavage of the encapsulated protein. For a more detailed understanding we tried to emulate enzymatic degradation and cleavage of DQ-OVA. For this purpose DQ-OVA, DEXS/pARG capsules with DQ-OVA as cargo in their cavity, and PSS/PAH capsules with DQOVA as cargo in their cavity were incubated in microplates at room temperature with proteases and the time dependence of the green fluorescence was monitored with a spectrofluorometer. As can be seen in Figure 2 the green fluorescence increased over time for all samples. However, increase was much higher in the case of the free DQ-OVA and DQ-OVA encapsulated with DEXS/pARG. As it was hard to adjust the different samples to have the same DQ-OVA concentration, we fitted the time-dependent fluorescence traces I(t) with the following function I(t) ) I0 + Imax(1 - exp(-t/τ)). From the three fit parameters I0, Imax, and τ, the first two are concentration related and thus were disregarded. The third parameter τ describes the time constant of the enzymatic reaction, including both, eventual degradation of the encapsulation shell around DQ-OVA and cleavage of DQ-OVA. We found time values of τ ) 30 ( 8 h, 80 ( 30 h, and 33 ( 10 h for DQ-OVA, DQ-OVA encapsulated with PSS/PAH, and DQ-OVA encapsulated with DEXS/pARG, respectively. DQ-OVA encapsulated within PSS/PAH is much less efficiently digested into peptide fragments than free DQ-OVA or DQ-OVA encapsulated with DEXS/pARG. Whereas proteases can directly access DQ-OVA and sufficiently fast 4400

Figure 2. Kinetics of degradation. Free and encapsulated DQ-OVA were incubated with a mixture of different proteases, called Pronase, whose proteolytic activity extends for a broad range of proteins including ovalbumin as described in the Supporting Information. The fluorescence intensity was measured at different time points with a spectrofluorometer. The fluorescence versus time for each system, free DQ-OVA, PSS/PAH(DQ-OVA), and DEXS/pARG(OVA) was plotted. The continuous lines are fits to the data using function I(t) ) I0 + Imax(1 - exp(-t/τ)). The increase in the fluorescence signal and thus degradation of DQ-OVA reached saturation after 50-60 h of incubation.

degrade the walls of DEXS/pARG capsules to access DQOVA, the wall of PSS/PAH capsules imposes a barrier hindering access of proteases to the DQ-OVA cargo. These spectral data are in good agreement with the data obtained with cell cultures. However, the question arises why in the PSS/PAH capsules green fluorescence is found at all. Charge density and thickness of capsule walls and pH and ionic strength of the medium are factors that strongly influence the permeability of polyelectrolyte capsules.49,50 In general PSS/PAH capsules are known for their enhanced mechanical stability and low permeability.51 Nevertheless, our experiments demonstrate that the cargo inside PSS/PAH capsules is not fully protected, though its enzymatic degradation is significantly slowed down by protection by the capsule walls. Although degradation of DEXS/pARG capsules inside cells was visible from the first day, the green fluorescence of cleaved DQ-OVA kept rising in the following days of incubation. After around 3-5 days green fluorescence was observed not only in the cavity of the capsules but also in small spots all over the cytoplasm of the cell; see Figure 3. This indicated that whereas the capsules remained in the intracellular vesicular compartments where they were originally internalized, the single dye-labeled peptide fragments which originated from the enzymatic cleavage of the encapsulated DQ-OVA could leave these compartments to be distributed along the entire cell body. Spots of fluorescentemitting peptide fragments were mostly found at the leading edges of the cells, which suggest that they might be transported with the actin cytoskeleton of the cells.47 These results indicate that after processing of encapsulated DQOVA, the resulting peptides could leave the intracellular vesicle in which the capsule is located. As most likely capsules undergo an endocytosis-like uptake process upon incorporation by cells,37 we speculated that the spots of peptide fragments visible in the cytoplasm might be attributed Nano Lett., Vol. 9, No. 12, 2009

Figure 3. Release of digestion product of DQ-OVA out of the capsule. 3T3 cells were incubated with biodegradable DEXS/pARG(OVA) capsules and were imaged with confocal microscopy. The left picture shows a group of cells that have incorporated several capsules. A fluorescence signal can be seen from the degraded capsules but also from outside the compartment where the capsules are located. These spots correspond to an accumulation of the digestion products of DQ-OVA that are further processed intracellularly. In this regard, it seems that the peptide fragments are transported to different compartments inside the cells, since the spots are of different sizes. A zoomed image of one area of the cell culture is presented in the right picture.

to exocytotic vesicles. In order to verify this hypothesis, we analyzed the fluorescence of the cell medium. Indeed, after cells had incorporated capsules for several days, the green fluorescence signal was found in the extracellular medium of the cultivated cells. A series of control experiments involving capsules only and cells only gave further evidence that the fluorescence found in the cell medium after several days originated from die-labeled peptide fragments which had been produced from DQ-OVA by enzymatic cleavage inside cells and had been subsequently exocytosed. Thus, by encapsulating DQ-OVA, which serves as nonfluorescent precursor, fluorescent peptide fragments were specifically synthesized in the cavity of biodegradable polyelectrolyte capsules upon enzymatic degradation of the capsule walls and enzymatic cleavage of the DQ-OVA. In this way enzymatic activity inside living cells was used to trigger the conversion of nonfluorescent molecule to a fluorescent one. Discussion. One of the great challenges in capsule technology is the construction of a capsule that can regulate some cellular effect. In this study, the ability of biodegradable polyelectrolyte microcapsules to trigger the enzymatic cleavage of encapsulated material was demonstrated. The presented data show that after enzymatic degradation of internalized biodegradable DEXS/pARG capsules, encapsulated proteins are available for proteolytic cleavage. This led to fluorescent peptide fragments which are released to presumably smaller vesicles inside the cytoplasm and which are finally exocytosed. On the other hand, PSS/PAH capsules are known for their enhanced mechanical stability and low permeability,51 properties that make these capsules presumably resistant to enzymatic digestion. Nevertheless, in general polyelectrolyte capsules exhibit a strong increase of permeability with increasing ionic strength and decreasing pH values.49,50 Since the capsules are transported for intracellular processing to intracellular acidic compartments where the pH values are low,37 some diffusion of the enzyme through the capsule wall of both capsules types as a consequence of increased permeability cannot be excluded. Regardless the slight increase in the green signal and its distribution in the cavity of the capsules probably due to DQ-OVA loosely bound to the capsule shell where it could be more accessible to Nano Lett., Vol. 9, No. 12, 2009

proteases, DQ-OVA from PSS/PAH capsules could not be significantly digested, the structural integrity of DQ-OVA was maintained and, therefore, was only visible in the red channel for the entire measurement. This fact indicates that DQ-OVA remains encapsulated inside the capsules as a consequence of failed degradation of PSS/PAH capsules. On the contrary, DQ-OVA from intracellular DEXS/pARG capsules exhibited a significant increase in the green fluorescent signal in a time-dependent manner with an absence of a red signal after several hours of cellular incubation. These experiments demonstrate that it is possible to convert nonactive precursor molecules to active molecules exclusively inside cells upon harnessing intracellular enzymes. In this way it might be possible in the future to introduce nonactive prodrugs into cells, which only inside cells would be activated to a drug with cytostatic effect. This would help to reduce side-effects outside cells. It has to be pointed out that our system in principle involves two enzymatic steps. After capsule incorporation first the walls of the capsules have to be enzymatically degraded by specific proteases. Upon degradation of the capsule walls other enzymes can enter the capsule cavity and process the cargo inside. In our kinetic experiments the enzymes used for capsule degradation and cleavage of the cargo were not distinguishable, since a mixture of proteases was used. However, one may think of cargo molecules which are only specifically activated by special enzymes only present in certain types of cells. This could allow for even higher specifity, which would activate administered prodrugs only in target cells. In combination with targeting mechanisms52 for specific cellular uptake of capsules this might be a valuable concept for specific drug targeting and activation. Experimental Section. Materials. Poly(sodium 4-styrenesulfonate) (PSS; Mw ∼ 70 kDa), poly(allylamine hydrochloride) (PAH Mw ∼ 70 kDa), dextran sulfate sodium salt (DEXS Mw ∼ 10 kDa), poly-L-arginine hydrochloride (PARG Mw > 70 kDa), calcium chloride, sodium carbonate, and EDTA were purchased from Sigma-Aldrich. DQ-OVA was purchased from Invitrogen. Capsule Synthesis. DQ-OVA loaded calcium carbonate (CaCO3) microparticles with an average diameter of 2 µm 4401

were synthesized by coprecipitation of CaCl2 and Na2CO3 in the presence of DQ-OVA. In detail, 200 µg of DQ-OVA was dissolved in 5 mL of water followed by the addition of 0.625 mL of CaCl2 (1 M) and 0.625 mL of Na2CO3 (1 M). After 30 s of reaction the precipitate was centrifuged and the supernatant was discarded. Two bilayers, DEXS/pARG and PSS/PAH, were deposited by alternate adsorption from aqueous 1 mg/mL solutions in 0.5 M NaCl, with two washing/centrifugation steps between each polyelectrolyte deposition step. Hollow capsules were obtained by addition of 10 mL of EDTA solution (buffered at pH 5) followed by two washing/centrifugation steps with PBS. Finally the capsules were stored in 1 mL of PBS at 4 °C. Fluorescence Spectroscopy. Fluorescence images and spectra were taken with a LSM 510 META confocal microscope from Zeiss and with a spectrofluorometer from Jovin Yvon, respectively. Images comprised phase contrast and green and red fluorescence channels. Particular care was taken to maintain the same settings in the microscope during the comparison experiments with the biodegradable DEXS/ pARG and nondegradable PSS/PAH capsules and to verify that the fluorescence signals were originating from the capsule cavity. As not all capsules within one cell are in the same focal plane, the comparison of fluorescence signals derived from different capsules turned out to be complex. Therefore fluorescence scans along the z-axis were performed. More details about the experimental procedure can be seen in the Supporting Information. Acknowledgment. This work was supported by the European Union (project Nanointeract) and the German Research Foundation (DFG, SPP 1313, Grant PA 794-4). B.G.D.G. thanks the FWO Vlaanderen for a postdoctoral fellowship. Supporting Information Available: Descriptions of cell culture and incubation, confocal microscopy, and spectroscopy used. This material is available free of charge via the Internet at http://pubs.acs.org. References (1) Sukhorukov, G. B.; Fery, A.; Brumen, M.; et al. Phys. Chem. Chem. Phys. 2004, 6 (16), 4078. (2) Sukhorukov, Gleb B.; Rogach, Andrey L.; Zebli, Bernd; et al. Small 2005, 1 (2), 194. (3) Johnston, A. P. R.; Cortez, C.; Angelatos, A. S.; et al. Curr. Opin. Colloid Interface Sci. 2006, 11 (4), 203. (4) De Geest, B. G.; De Koker, S.; Sukhorukov, G. B.; et al. Soft Matter 2009, 5 (2), 282. (5) Sukhorukov, G. B.; Mohwald, H. Trends Biotechnol. 2007, 25 (3), 93. (6) Sukhorukov, Gleb B.; Rogach, Andrey L.; Garstka, Malgorzata; et al. Small 2007, 3 (6), 944. (7) Rivera Gil, P.; del Mercato, L. L.; del Pino, P.; et al. Nano Today 2008, 3 (3-4), 12. (8) Decher, Gero Science 1997, 277, 1232. (9) Schneider, G.; Decher, G. Langmuir 2008, 24 (5), 1778. (10) Donath, Edwin; Sukhorukov, Gleb B.; Caruso, Frank; et al. Angew. Chem., Int. Ed. 1998, 37 (16), 2202. (11) Sukhorukov, Gleb B.; Donath, Edwin; Davis, Sean; et al. Polym. AdV. Technol. 1998, 9 (10-11), 759. (12) Cortez, C.; Tomaskovic-Crook, E.; Johnston, A. P. R.; et al. AdV. Mater. 2006, 18 (15), 1998. (13) Zaitsev, Sergey; Cartier, Regis; Vyborov, Oleg; et al. Pharm. Res. 2004, 21 (9), 1656.

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Nano Lett., Vol. 9, No. 12, 2009