Cytotoxicity and Internalization of Polymer Hydrogel Capsules by

Jul 9, 2010 - Capsule cytotoxity, an essential aspect of a drug delivery vehicle, was ..... The authors would like to thank Dr. Brigitte Städler, Dr...
0 downloads 0 Views 2MB Size
Download PDF
Biomacromolecules 2010, 11, 2123–2129

2123

Cytotoxicity and Internalization of Polymer Hydrogel Capsules by Mammalian Cells Alexander N. Zelikin,*,§,† Kerry Breheney,‡ Remy Robert,‡ Elvira Tjipto,† and Kim Wark*,‡ Centre for Nanoscience and Nanotechnology, Department of Chemical and Biomolecular Engineering, The University of Melbourne, Parkville, Victoria 3010, Australia, and CSIRO, Molecular and Health Technologies, Parkville, Victoria 3052, Australia Received May 7, 2010; Revised Manuscript Received June 28, 2010

Polymer hydrogel capsules comprised of poly(methacrylic acid) chains and cross-linked via disulfide linkages were investigated for their cytotoxicity and mechanism of internalization in a variety of mammalian cells. The capsules were internalized by all the tested cell lines which differed in their morphology and function and over short to medium term (24 h) revealed no reduction in viability and metabolic activity of cells. The mechanism of capsule uptake was analyzed using inhibitors of various cellular entry pathways. Of these, blocking the clathrinmediated endocytotic pathway resulted in a statistically significant reduction in capsule uptake, suggesting this was the predominant pathway of capsule entry in these cell lines. The uptake of solid particles with similar surface chemistry was not significantly decreased by the inhibitor of the clathrin-mediated pathway, which suggested that softness and concomitant flexibility of the hydrogel capsules were factors governing the entry mechanism. This work represents the first systematic study of the interaction of polymer hydrogel capsules with mammalian cells and provides essential information for the application of these capsules in biomedicine.

Introduction Supramolecular polymer1,2 and lipid3 assemblies are attracting increased attention as successful candidates for diverse biomedical applications such as drug delivery, creation of synthetic biomimetic microreactors4 and assembly of artificial cells.5 Among these, polymer capsules obtained via sequential (layerby-layer, LbL) deposition of polymers onto sacrificial template particles have shown potential as a versatile, tunable platform. This technology allows capsules with defined sizes and diverse structural components, including biodegradable polymers, to be assembled.6,7 Significant advances have also been made in drug and reagent loading techniques, as well as subcompartmentalization of capsules.5 However, the interaction between such capsules and live cells, tissues, and organs remains largely unexplored. Herein, we used LbL-derived poly(methacrylic acid) hydrogel capsules (PMA HC) with sizes 500 nm and 1 µm and investigated their cytotoxicity, interaction with proteins, and internalization by mammalian cells. We chose these capsules based on their recent success in a multitude of biomedical applications, from creation of prototype artificial organelles8 and subcompartmentalized microreactors9,10 to delivery of anticancer drugs11 and oligopeptide vaccines12 in vitro and in vivo vaccination.13 More recently, we used these capsules to demonstrate the first example of a microreactor that also serves as a delivery vehicle for cellular internalization of the de novo synthesized RNA.14 These successful applications rely on the drug loading techniques we have developed for encapsulation of protein,15 oligopeptides,12 single16 and double-stranded DNA,17 and small drugs11 and are also facilitated by the * To whom correspondence should be addressed. [email protected] (K.W.); [email protected] (A.N.Z.). § Current address: Department of Chemistry, Aarhus University, Aarhus 8000 Denmark. † The University of Melbourne. ‡ CSIRO.

outstanding colloidal stability of PMA HC,18 which is currently unmatched by other LbL-derived systems. While the material science aspects of the capsule formation and drug loading techniques are rather well established, capsule interactions with live cells largely remains uncharacterized. In this work, we investigated the interaction of PMA HC with a range of mammalian cells and demonstrated that (i) capsules of all sizes studied were effectively internalized by all mammalian cells lines, both adherent and nonadherent; (ii) regardless of their size, over short to medium term observation (24 h), hydrogel capsules did not exhibit cytotoxicity in any of the mammalian cell lines tested, (iii) of the cell entry pathways tested, internalization of capsules was significantly reduced only in the presence of an inhibitor of clathrin mediated pathway, a feature not observed for the parent core particles, and (iv) the mechanism of entry into cells was not dependent on the capsule size. These results clearly validate PMA HCs as suitable materials for drug delivery to cells with diverse morphology and function. It also provides an understanding of the predominant uptake mechanism and enables opportunities to further engineer the capsules to manipulate this interaction and subsequent cellular trafficking of these drug carriers. This is the first systematic investigation of the interactions between polymer hydrogel capsules and mammalian cells.

Experimental Section Materials. Unless stated otherwise, all chemicals and materials were purchased from Sigma-Aldrich and used as received without purification. SiO2 particles of 300 nm, 500 nm, and 1 µm diameter were purchased from MicroParticles GmbH (Berlin, Germany). Poly(methacrylic acid), MW 15 KDa, was purchased from Polysciences (U.S.A.). Alexa Fluor 488 and 633 maleimides, RPMI-1640 + L-glutamine, DMEM, L-glutamine, Opti-MEM 1 reduced serum medium, MEM nonessential amino acids, sodium pyruvate, trypsin, penicillin, and streptomycin were purchased from Invitrogen. Fetal bovine serum (FBS) was from JRH.

10.1021/bm100500v  2010 American Chemical Society Published on Web 07/09/2010

2124

Biomacromolecules, Vol. 11, No. 8, 2010

Methods. Flow cytometry measurements were performed using Partec CyFlow Space flow cytometer with an absolute volume counting capability using 488 and 633 nm excitation wavelengths. The capsules and cells were imaged using an Olympus IX71 Digital Wide field fluorescence microscope and a Leica confocal laser scanning microscope. Assembly of PMA HC was performed as described elsewhere.18 In brief, monodisperse silica template particles were alternately incubated in a 1 g/L solution of poly(vinylpyrrolidone) (PVPON, 10 KDa) and a 1 g/L solution of thiolated PMA (12 mol % of carboxyl groups converted into thiol groups) in 20 mM sodium acetate buffer, pH 4, with intermediate washing via centrifugation/redispersion cycles. A total of 11 polymer layers were deposited (six layers of PVPON and five layers of thiolated PMA), after which time the thiol groups were oxidized using chloramine T. The template particles were removed using dilute hydrofluoric acid, and the resulting capsules were washed using 20 mM sodium acetate buffer, pH 4. When transferred into a buffer with pH > 6.5, ionization of PMA caused the release of PVPON from the capsules walls and yielded single component PMA HC stabilized via disulfide linkages. The capsules exhibit a characteristic hydrogel swelling in response to a change in pH, presence of low molecular weight solutes, and so on, and in cell culture medium, the capsules templated on 500 nm SiO2 were ∼600 nm in diameter and the capsules templated on 1 µm SiO2 attained a size of 1.7 µm in diameter, as determined by dynamic light scattering characterization. For cell uptake experiments, the PMA HC were fluorescently labeled with Alexa Fluor 633 using a 1 g/L solution of Alexa Fluor 633 maleimide in DMSO, which was added to a dispersion of PMA HCs in 50 mM MES buffer, pH 6, to a final DMSO content of 5-10 vol %. In a typical labeling experiment, approximately 1010-1011 500 nm PMA HC were labeled using 10-50 µg of the dye. The reaction between maleimide dye molecules and residual nonoxidized thiol groups within the PMA HC capsules wall was allowed to proceed for at least 2 h, after which time the capsules were washed with fresh buffer via centrifugation and redispersion cycles until the supernatant became nonfluorescent. For capsule counting, we used a sample of PVPON synthesized via a reversible addition-fragmentation chain transfer (RAFT) polymerization technique to obtain a polymer sample labeled with Alexa Fluor 488.19 Briefly, to convert the polymer terminal thioester group into a thiol, an aliquot of the synthesized polymer was dissolved and incubated in a 1 M sodium borohydride solution for 1 h. After this time, excess borohydride was quenched with concentrated hydrochloric acid, and the solution pH was adjusted to ∼7. The resulting solution was supplemented with Tris-EDTA buffer to 10 mM and charged with Alexa Fluor 488 maleimide solution (typically 0.1% to the weight of the polymer). The reaction was allowed to proceed overnight, after which time the polymer was recovered via gel filtration and freeze-drying. An aliquot of PMA HC was mixed with 10 volumes of PBS, pH 7.4, to release PVPON. An aliquot of the resulting single component capsules was subsequently mixed with an equal volume of the Alexa Fluor 488 labeled PVPON solution in 50 mM sodium acetate buffer, pH 4. The fluorescent polymer, being in excess to that released from the capsules, effectively infiltrates into the PMA HC hydrogel to give rise to highly fluorescent capsules. This procedure has proven effective for labeling of the PMA HC with sizes as low as 300 nm and allowed for reliable and reproducible counting of the capsules on a CyFlow Partec Space flow cytometer, with absolute volume counting triggering on Alexa Fluor 488 fluorescence. Cell Culture. The CHO-K1 WT (ATCC: CCL61), NR6 (a 3T3 derivative), HEK293T (ATCC: CRL 11268), M17 (ATCC: CRL-2267), P388D1 (ATCC: CCL-46), and RAMOS (ATCC: CRL-1596) cell lines were maintained according to the American Type Culture Collection (ATCC) protocols. RAMOS cells were grown in suspension at 37 °C and 5% CO2. All other cells were cultured as monolayers at 37 °C and 5% CO2. CHO and HEK293T cells were grown in RPMI-1640 + L-glutamine supplemented with 5% FBS and antibiotics (penicillin 500 U/mL, streptomycin 50 µg/mL). P388D1 and RAMOS cells were grown

Zelikin et al. in RPMI-1640 + L-glutamine supplemented with 10% FBS, penicillin 50 U/mL, and streptomycin 50 µg/mL. NR6 fibroblasts were grown in DMEM supplemented with 2 mM L-glutamine, 5% FBS, penicillin 500 U/mL, and streptomycin 50 µg/mL. M17 cells were grown in OptiMEM 1 reduced serum medium supplemented with nonessential amino acids, sodium pyruvate, 10% FBS, penicillin 500 U/mL, and streptomycin 50 µg/mL. PMA HC Uptake. Cells were seeded into 96-well plates (Nunc) at a density of 4 × 104 cells per well in complete growth medium and allowed to attach overnight at 37 °C and 5% CO2. After this time, PMA HCs in PBS were added to the cells to give a final well volume of 290 µL and incubated for 24 h at 37 °C, 5% CO2, with shaking at 90 rpm. Prior to analysis, the cells were washed twice with prewarmed PBS, trypsinized, and stored on ice. Cells were then analyzed using a CyFlow Partec flow cytometer. Raw data were analyzed using Flow Jo software with a built-in population comparison algorithm. CellTiter-Glo Cell Viability Assay. Cells were seeded into 96-well opaque tissue culture plates (PerkinElmer) at a density of 4 × 104 cells/ well in 100 µL growth medium and allowed to attach overnight at 37 °C and 5% CO2. The cells were then exposed to 500 nm PMA HC at various capsule: cell ratios and incubated for 24 h. The viability of the cells was assessed using CellTiter-Glo luminescent cell viability kit from Promega Corporation (Madison, WI) according to the manufacturer’s instructions. The assay detected the amount of bioluminescent ATP present, which was directly proportional to the number of viable cells. Luminescence was measured on a FLUOstar Optima (BMG LABTECH). Inhibition of Uptake. Cells were seeded into 96-well plates (Nunc) at a density of 4 × 104 cells per well in complete growth medium and allowed to attach overnight at 37 °C and 5% CO2. The cell culture medium was replaced with fresh growth medium without antibiotics, supplemented with either 10 µM chlorpromazine, 1 µg/mL filipin, or 50 µM amiloride for 30 min prior to the addition of Alexa Fluor 633 (AF-633) labeled PMA HC. Cells were incubated a further 24 h, with shaking at 90 rpm, washed twice with PBS, trypsinized, and analyzed by flow cytometry, as described above. The presented data are mean ( SD of at least two independent runs, six replicates in each run, typically counting at least 1000 cells per replicate; date were analyzed using a one way ANOVA test (***p < 0.001; **p < 0.01; *p < 0.05; nonsignificant unless marked otherwise).

Results and Discussion Assembly of PMA HCs relies on the hydrogen bondingassisted sequential deposition of PMA and poly(vinylpyrrolidone) onto sacrificial template particles. We have previously shown that this assembly proceeds without aggregation of the particles, which makes it a convenient starting platform to obtain polymer capsules of different compositions (single component PMA18 or PVPON20 hydrogel capsules). Single component disulfide stabilized PMA HCs,18,21 as used in this work were obtained using thiol modified PMA, conversion of thiols into bridging disulfide linkages within the assembled multilayered film, removal of the core particles, and finally increasing pH to release PVPON. This assembly was routinely performed using 500 nm and 1 µm sized particles without aggregation which ensures a high yield (i.e., minimal particle loss during assembly) and produces a population of capsules monodisperse in size (Figure 1). To assemble PMA HCs, we used 300 nm, 500 nm, and 1 µm commercial monodisperse silica template particles. The resulting single component hydrogel capsules were larger in size compared to the parent core particles due to the characteristic hydrogel swelling.18 Throughout the text, capsule sizes are quoted as those of the respective template particles. To ensure accurate and reproducible dosage, the capsules were counted using a flow cytometer with an absolute volume

Cytotoxicity of Polymer Hydrogel Capsules

Biomacromolecules, Vol. 11, No. 8, 2010

2125

Figure 1. Confocal laser scanning microscopy images of 1 µm (top row) and 500 nm (bottom row) PMA HC.

Figure 2. Cytotoxicity data obtained using 500 nm PMA HC, CellTiterGlo cell viability assay, and a representative panel of cells. Experiments were conducted in 96-well plates with an initial seeding density of 4 × 104 cells/well and a 24 h incubation time of cells with capsules. The numbers in the figure legend indicate the administered capsules to cell ratio.

counting function using fluorescence parameter as a trigger (see Experimental Section). Capsule cytotoxity, an essential aspect of a drug delivery vehicle, was assessed in five different cells lines including kidney (HEK), muscle (NR6), ovary (CHO), neuron (M17), and macrophage (P388D1) over a 24 h period using the CellTiterGlo cell viability assay. A range of capsule to cell ratios was tested using 500 nm PMA HC (Figure 2). Cell viability was not significantly affected in any of the cell lines, even at a ratio of 1000 capsules to 1 cell (Figure 2), whereas all cell lines treated with 25% DMSO viability were reduced to less than 20% (data not shown). The morphology of CHO cells was not affected following treatment with 500 nm capsules, as observed by fluorescence microscopy (Figure 3a,b). Confocal laser

Figure 3. Bright field microscopy (A) and confocal laser scanning microscopy (B-D) images of CHO cells incubated with 500 nm (A-C) and 1 µm PMA HC (D): bright field image (A), an overlay of bright field and fluorescent images (B), and fluorescence images of cells with internalized 500 nm (C) and 1 µm (D) PMA HC. The capsules were fluorescently labeled with Alexa Fluor 488; cell membrane staining was performed using FM-64 dye. Images A and B demonstrate that incubation with capsules does not lead to a change in morphology of CHO cells, thus supporting the conclusion that PMA HC are nontoxic to mammalian cells in the chosen incubation conditions. For confocal imaging (C,D), after incubation with capsules, the cells were washed twice with PBS, trypsinized, and transferred into PBS buffer. Presented images reveal that most of the capsules were within the cell cytoplasm and not associated with the cell surface, thus demonstrating that the outlined washing procedure was sufficient to remove the majority of surface bound or free capsules.

scanning micrographs confirmed that 500 nm and 1 µm capsules were internalized by mammalian cells (Figure 3c,d). Together, the data in Figures 2 and 3 imply that the PMA HCs do not contain or release toxic products that affect cell viability upon

2126

Biomacromolecules, Vol. 11, No. 8, 2010

binding and internalization into cells, at least over the period of 24 h. This puts PMA HC in stark contrast with other LbLderived capsules, which were shown to cause cellular toxicity.22–24 Indeed, the number of viable C6 glioma and 3T3 fibroblast cells was decreased upon incubation with capsules assembled using either synthetic or natural polyelectrolytes (poly(sodium 4-styrenesulfonate) (PSS)/poly(allylamine hydrochloride) (PAH); bovine serum albumin (BSA)/PAH; alginate/polylysine, etc.) with sizes from 1 to 10 µm, even at a moderate (10:1) capsule to cell ratio.22 Similarly, there was a reduction in the viability of denditric cells when treated with 3 µm capsules assembled from dextran sulfate and poly(arginine).23 We note that other reports demonstrate that polyelectrolyte capsules can be nontoxic to cells (e.g., PSS/PAH capsules taken in a 100:1 ratio to dendritic cells and macrophages),25 which may be reflective of different lengths of incubation with cells, capsule concentration, cell specific effects, and possibly batch-to-batch variation in the capsules. However, PMA HC have consistently proven to be nontoxic at all concentrations and to all cell types tested. We emphasize that in these experiments we specifically considered a short to medium term effect of capsules on the cells’ viability and metabolic activity, namely, a 24 h observation time, which has relevance to applications such as vaccine delivery. For other applications (e.g., delivery of RNAi therapeutics), a longer term toxicity study and a concurrent investigation of the intracellular capsules’ residence time is warranted, which is the subject of ongoing research. To investigate the interaction of capsules with mammalian cells, we used flow cytometric analysis and Alexa Fluor 633 as a fluorescent dye to label the capsules, which was chosen to minimize the background signal resulting from cell autofluorescence. The model CHO cell line was used to develop a protocol for measuring capsule uptake. In optimizing the protocol, the following was observed: (i) a higher ratio of capsules to cells was desirable to obtain results in a 24 h time period, (ii) in the presence of excess capsules, the concentration of cells was a far less important parameter than the number of capsules, and (iii) no significant differences were observed between rocking the assay plate or keeping it idle during the incubation period. In all further experiments, unless stated otherwise, a 96-well format, with seeding 4 × 104 cells per well and 100 capsules to 1 cell ratio, was used. After incubation with capsules, the cells were washed twice with PBS, trypsinized, and transferred into PBS buffer for subsequent analysis by flow cytometry. The raw fluorescence histograms (Figure 4) were compared to the negative population using the Flow Jo analysis software and the built-in population comparison algorithm to obtain a numerical value of the fraction of cells with associated fluorescence, that is, PMA HC. While a fluorescence read out does not distinguish between adsorbed and internalized capsules, confocal laser scanning photographs (Figure 3c,d) show that the employed washing protocol effectively removes the capsules bound to the outer surface of the cells, and therefore, the fluorescent signal obtained by flow cytometry corresponds to internalized capsules. Different cell types were assessed for their ability to internalize PMA HCs after 3 and 24 h post-incubation (Figure 5). Both phagocytic and nonphagocytic cells internalized 500 nm and 1 µm diameter capsules, indicating that PMA HCs are amenable for drug delivery applications for all of the cell lines tested. In all but one cell line, an increased incubation time resulted in a greater fraction of cells with internalized capsules (Figure 5). A notable exception was the RAMOS B cells, which exhibited a greater uptake level at the 3 h time point compared with 24 h

Zelikin et al.

Figure 4. Typical flow cytometric histograms obtained for the CHO cells incubated with the fluorescently labeled 1 µm PMASH capsules (dark) and the negative population of the cells (light). Increase in the cell fluorescence is attributed to the association of the capsules with the cells followed by their internalization.

Figure 5. Efficiency of uptake of PMA HC, 1 µm (left) and 500 nm (right) in size, exhibited by mammalian cells. Efficiency of uptake is expressed as percent of cells with associated capsules, which was estimated using Alexa Fluor 633 labeled capsules and flow cytometry as means of analysis. The raw histograms were analyzed using Flow Jo analysis software to yield numerical values corresponding to the fraction of cells with internalized capsules.

for both capsule sizes. The latter observation suggests either that RAMOS has a different uptake mechanism/s or physiological conditions specific to its cell membrane, which differs from the other epithelial cell lines tested. While the M17 cells exhibited a lower level of uptake for the 1 µm capsules, it does not imply that the uptake of PMA HCs by these cells is restricted to smaller capsules, as these cells successfully internalized 1 µm PMA HC at higher capsule to cell ratios (data not shown). To determine if the capsules used a passive or active entry pathway, the uptake assay was performed at 37 and at 4 °C. At lower temperatures, cellular metabolic activity is greatly diminished and internalization processes are slowed down due to the solidification of the membrane lipid bilayer. The uptake of the capsules was lower at 4 °C compared to 37 °C (Figure 6), which suggests that capsule internalization is an energydependent process. Studies in the field have suggested that the charge on the outer surface of the nanoparticle is responsible for its interaction with the cell membrane either directly or via adsorption of serum proteins assisting cell entry. In the case of PMA HCs, the level of capsule uptake was not dependent on the presence of serum in the assay medium. Likewise, incubating

Cytotoxicity of Polymer Hydrogel Capsules

Figure 6. Efficiency of uptake of 500 nm PMA HC exhibited by CHO cells upon incubation with capsules in cell culture media at 4 °C (B) and 37 °C (C-E) in the presence (B,C) or absence (D,E) of serum proteins. Sample (A) corresponds to cells only in the absence of capsules; for experiment (E), the capsules were preincubated with serum proteins, isolated from serum, and administered onto cells cultured in a serum-free medium.

capsules with serum prior to performing the assay did not affect capsule uptake levels compared to those that were not pretreated. These observations suggest that the capsules exhibit a minor level of interaction with serum proteins, that is, low fouling behavior. To elucidate the cell entry mechanism(s) used by PMA HCs, we investigated the uptake of the capsules by mammalian cells in the presence of chlorpromazine, filipin, and amiloride, which inhibit clathrin-mediated endocytosis (CME), caveolae-mediated endocytosis (CvME), and macropinocytosis (MP), respectively.26 While the majority of inhibitor studies investigating endocytosis remove the inhibitors prior to the addition of test material, it has been previously reported that the effect of inhibitors is reversible, and in a particular case of chlorpromazine, the cells can restore their normal metabolic pathways within 60 min.27,28 Together with the fact that this time frame is too short to allow a detectable number of capsules to enter into cells, this led us to conduct incubation of capsules with cells in the presence of inhibitors (10 µM chlorpromazine, 1 µg/mL filipin, 50 µM amiloride)29–31 for the entire duration of the experiment, 24 h. In our preliminary experiments we verified that these conditions had a minimal to negligible effect on cell viability (see Supporting Information). Throughout these experiments, the same preparation of capsules was used to allow a direct comparison of the data. Both the fraction of cells with internalized capsules and the mean fluorescence of the population (i.e., an average number of capsules internalized by each cell) were analyzed. While the latter parameter produced a greater response to the presence of inhibitors, similar conclusions could be drawn from both analyses. For 500 nm PMA HCs, the uptake level in CHO cells was not significantly affected by filipin and amiloride. In contrast, CHO cells treated with chlorpromazine showed a statistically significant reduction in capsule uptake (p < 0.001; Figure 7). Similarly, capsule uptake was significantly reduced in the presence of chlorpromazine in neuronal cells (M17) and macrophages (P338D1). The CME inhibitor significantly decreased capsule uptake levels and mean fluorescence intensity (p < 0.01 and p < 0.001 for M17 and P388D1, respectively),

Biomacromolecules, Vol. 11, No. 8, 2010

2127

whereas inhibitors of CvME (filipin) and MP (amiloride) did not significantly affect PMA HC uptake in both cell lines. Furthermore, the uptake mechanism in CHO cells remained the same for all three capsule sizes studied (300 nm, 500 nm and 1 µm): for all the capsule sizes internalization was inhibited by chlorpromazine but not filipin or amiloride (Figure 7), even when the latter were taken at higher concentrations (2 µg/mL filipin, 500 µM amiloride; see Supporting Information). Interestingly, uptake of PMA HCs by the RAMOS B cells was not affected by either chlorpromazine, filipin, and amiloride and, thus, may signify a different mechanism of internalization. The mechanism of cell entry is now widely accepted as an important aspect which defines the intracellular fate of a drug or gene carrier.32,33 It determines if the internalized cargo is sorted through a destructive pathway or released into the cytosol and thus affects drug efficacy. However, trends described in the literature regarding the effect of particle size on cell entry mechanisms remain inconclusive. Various research groups used particles in the same size range from 0.1 to 5 µm and have reported results with apparent discrepancies. An increase in particle size was shown to have no effect on the mechanism of particle entry (MP for all sizes)34 in one study, whereas in another it resulted in a change from predominantly CME to CvME (with a threshold at ∼200-500 nm),35 while in yet another study a shift to predominantly CME was observed.36 From a different perspective, the data on cellular internalization reported for LbL-derived polyelectrolyte capsules (sized 3-4 µm) suggest a CvME entry37 or a combination of CvME and MP,38 that is, a different mechanism to that identified for PMA HCs. Two apparent differences between other particles or capsules and the PMA HCs used in this study are first the hydrogel nature of the PMA capsules and second their surface chemistry. Indeed, the structure of PMA HC is that of a volume of water (buffer) surrounded only by a 30 nm thick, swollen polymer membrane,21 which makes them inherently soft and deformable. These characteristics of a drug carrier were previously recognized by Cohen et al.39 as important to explain a similar performance of 35 nm and 3.5 µm hydrogel vaccine carriers. These factors are also known to play an important role in interactions between cell membrane and biomaterials in tissue engineering.40 To investigate the impact of these factors on cellular uptake, PMA HCs were directly compared to solid silica particles with an adsorbed layer of PMA, that is, hydrogel capsules and solid particles with a similar surface chemistry. Internalization of solid particles was not significantly affected in the presence of chlorpromazine (Figure 8), suggesting that CME was not a major mechanism of PMASH-coated particles’ entry. This also infers that the hydrogel nature of PMA HCs and not their surface chemistry is a decisive factor in the mechanism of cell entry. We also used similar-sized PAH/PSS capsules, a two-component polyelectrolyte system that was expected to be more rigid than the highly swollen single component PMA HCs but less rigid than the solid silica particles. The uptake of these PAH/PSS capsules was significantly decreased in the presence of chlorpromazine compared to that of the solid particles (p < 0.01), but to a lesser degree than PMA HCs (p < 0.05; Figure 8), further supporting the above hypothesis. We note that, while further investigation into the mechanism of uptake of solid particles was beyond the scope of this project, we suggest that this is the first direct evidence of different internalization pathways for a hydrogel and a solid sphere with similar size and surface chemistry.

2128

Biomacromolecules, Vol. 11, No. 8, 2010

Zelikin et al.

Figure 7. Normalized uptake level (top row) and population mean fluorescence (bottom row) of cells incubated with PMA HC in the presence of specific inhibitors of cellular uptake mechanisms: chlorpromazine (10 µM; inhibitor of clathrin-mediate endocytosis), filipin (1 mg/L; inhibitor of the caveolae-mediated uptake), amiloride (50 µM, macropinocytosis inhibitor). Left: uptake and mean fluorescence for different mammalian cells incubated with 500 nm PMA HC; right: uptake and mean fluorescence of CHO cells incubated with PMA HC of differed size. In each case, the uptake level (percent of cells with associated capsules) and mean population fluorescence were normalized using the data for the respective cells/capsules in absence of inhibitors; the presented data are mean ( SD of at least two runs, six samples each run, analyzed using a one-way ANOVA test (***p < 0.001; **p < 0.01; *p < 0.05; nonsignificant unless marked otherwise).

positive controls using these and other hydrogel test materials. Also, to address concerns raised as to the specificity of inhibitors in their action,26 we are currently investigating internalization of capsules using clathrin-deficient cell lines, and these results will be reported in a subsequent publication.

Conclusions

Figure 8. Internalization of PMA HC, PSS/PAH polyelectrolyte capsules and PMASH-coated silica particles, all ∼500 nm, by CHO cells in the presence of inhibitors of endocytosis. All conditions and data analyses as in Figure 7.

The experiments presented above demonstrate that of all the inhibitors of endocytosis tested in this work, only chlorpromazine was effective in suppressing PMA HC cell entry, which strongly suggests a CME mechanism of capsules internalization. This mechanism is rather unexpected for these relatively largesized vessels and is possibly due to the inherent softness of these hydrogel capsules. It is also possible that chlorpromazine dependence as described above does not imply that PMA HC are internalized via a classic CME. Thus, while classical CME results in lysosomal trafficking of cargo and a hindered cytosolic drug delivery,33 we have previously reported that various drugs with a cytosolic or nuclear site of action were effective when delivered to cells using PMA HC.11–13 We therefore believe that the mechanism of PMA HCs uptake and trafficking is possibly more complex, involving either nonclassical CME or other pathways. We acknowledge that inhibitor studies as described above do not rule out the implication of other cell entry mechanisms, and to address this in greater detail, we are currently designing inhibitor experiments with more stringent

Herein, we presented the results of a systematic investigation of cytotoxicity and cellular internalization of the poly(methacrylic acid) hydrogel capsules, a supramolecular polymer drug carrier system. We established that the capsules are not toxic and internalized in a variety of mammalian cell lines of diverse morphology and physiology. We provided evidence that the capsules ranging in size from 300 nm to 1 µm were internalized predominantly by a clathrin-mediated endocytotic pathway. By direct comparison of the capsules and solid particles with similar surface properties, we demonstrated that a plausible explanation for this entry pathway may be due to the inherent softness and deformability of the hydrogel capsules. Acknowledgment. The authors would like to thank Dr. Brigitte Sta¨dler, Dr. Andrew Price, Dr. Katharina Ladewig (University of Melbourne), and Dr. Ben Boyd (Monash University) for valuable scientific discussions. This work was supported by the Australian Research Council Discovery Project (A.N.Z.) scheme. Supporting Information Available. Cytotoxicity characterization of inhibitors of uptake and additional data on cell entry inhibition. This material is available free of charge via the Internet at http://pubs.acs.org.

References and Notes (1) LoPresti, C.; Lomas, H.; Massignani, M.; Smart, T.; Battaglia, G. J. Mater. Chem. 2009, 19, 3576–3590. (2) Lensen, D.; Vriezema, D. M.; van Hest, J. C. M. Macromol. Biosci. 2008, 8, 991–1005. (3) Torchilin, V. P. Nat. ReV. Drug DiscoVery 2005, 4, 145–160.

Cytotoxicity of Polymer Hydrogel Capsules (4) Vriezema, D. M.; Aragones, M. C.; Elemans, J. A. A. W.; Cornelissen, J. J. L. M.; Rowan, A. E.; Nolte, R. J. M. Chem. ReV. 2005, 105, 1445–1489. (5) Stadler, B.; Price, A. D.; Chandrawati, R.; Hosta-Rigau, L.; Zelikin, A. N.; Caruso, F. Nanoscale 2009, 1, 68–73. (6) De Geest, B. G.; Sanders, N. N.; Sukhorukov, G. B.; Demeester, J.; De Smedt, S. C. Chem. Soc. ReV. 2007, 36, 636–649. (7) Quinn, J. F.; Johnston, A. P. R.; Such, G. K.; Zelikin, A. N.; Caruso, F. Chem. Soc. ReV. 2007, 36, 707–718. (8) Price, A. D.; Zelikin, A. N.; Wang, Y. J.; Caruso, F. Angew. Chem., Int. Ed. 2009, 48, 329–332. (9) Stadler, B.; Chandrawati, R.; Price, A. D.; Chong, S. F.; Breheney, K.; Postma, A.; Connal, L. A.; Zelikin, A. N.; Caruso, F. Angew. Chem., Int. Ed. 2009, 48, 4359–4362. (10) Chandrawati, R.; Stadler, B.; Postma, A.; Connal, L. A.; Chong, S. F.; Zelikin, A. N.; Caruso, F. Biomaterials 2009, 30, 5988–5998. (11) Sivakumar, S.; Bansal, V.; Cortez, C.; Chong, S. F.; Zelikin, A. N.; Caruso, F. AdV. Mater. 2009, 21, 1820–1824. (12) Chong, S. F.; Sexton, A.; De Rose, R.; Kent, S. J.; Zelikin, A. N.; Caruso, F. Biomaterials 2009, 30, 5178–5186. (13) Sexton, A.; Whitney, P. G.; Chong, S. F.; Zelikin, A. N.; Johnston, A. P. R.; De Rose, R.; Brooks, A. G.; Caruso, F.; Kent, S. J. ACS Nano 2009, 3, 3391–3400. (14) Price, A. D.; Zelikin, A. N.; Wark, K. L.; Caruso, F. AdV. Mater. 2010, 22, 720–724. (15) Zelikin, A. N.; Quinn, J. F.; Caruso, F. Biomacromolecules 2006, 7, 27–30. (16) Zelikin, A. N.; Li, Q.; Caruso, F. Angew. Chem., Int. Ed. 2006, 45, 7743–7745. (17) Zelikin, A. N.; Becker, A. L.; Johnston, A. P. R.; Wark, K. L.; Turatti, F.; Caruso, F. ACS Nano 2007, 1, 63–69. (18) Zelikin, A. N.; Li, Q.; Caruso, F. Chem. Mater. 2008, 20, 2655–2661. (19) Zelikin, A. N.; Such, G. K.; Postma, A.; Caruso, F. Biomacromolecules 2007, 8, 2950–2953. (20) Kinnane, C. R.; Such, G. K.; Antequera-Garcia, G.; Yan, Y.; Dodds, S. J.; Liz-Marzan, L. M.; Caruso, F. Biomacromolecules 2009, 10, 2839–2846. (21) Becker, A. L.; Zelikin, A. N.; Johnston, A. P. R.; Caruso, F. Langmuir 2009, 25, 14079–14085. (22) An, Z. H.; Kavanoor, K.; Choy, M. L.; Kaufman, L. J. Colloids Surf., B 2009, 70, 114–123. (23) De Koker, S.; De Geest, B. G.; Cuvelier, C.; Ferdinande, L.; Deckers, W.; Hennink, W. E.; De Smedt, S.; Mertens, N. AdV. Funct. Mater. 2007, 17, 3754–3763.

Biomacromolecules, Vol. 11, No. 8, 2010

2129

(24) Kirchner, C.; Javier, A. M.; Susha, A. S.; Rogach, A. L.; Kreft, O.; Sukhorukov, G. B.; Parak, W. J. Talanta 2005, 67, 486–491. (25) Wattendorf, U.; Kreft, O.; Textor, M.; Sukhorukov, G. B.; Merkle, H. P. Biomacromolecules 2008, 9, 100–108. (26) Ivanov, A. I., Ed. Pharmacological inhibition of endocytic pathways: Is it specific enough to be useful? In Methods in Molecular Biology; Humana Press: Totowa, NJ, 2008; Vol. 440, pp 15-33. (27) Wang, L. H.; Rothberg, K. G.; Anderson, R. G. W. J. Cell Biol. 1993, 123, 1107–1117. (28) Huth, U. S.; Schubert, R.; Peschka-Suss, R. J. Controlled Release 2006, 110, 490–504. (29) Lai, S. K.; Hida, K.; Man, S. T.; Chen, C.; Machamer, C.; Schroer, T. A.; Hanes, J. Biomaterials 2007, 28, 2876–2884. (30) Panyam, J.; Zhou, W. Z.; Prabha, S.; Sahoo, S. K.; Labhasetwar, V. FASEB J. 2002, 16, 1217–1226. (31) Walsh, M.; Tangney, M.; O’Neill, M. J.; Larkin, J. O.; Soden, D. M.; McKenna, S. L.; Darcy, R.; O’Sullivan, G. C.; O’Driscoll, C. M. Mol. Pharmaceutics 2006, 3, 644–653. (32) Doherty, G. J.; McMahon, H. T. Annu. ReV. Biochem. 2009, 78, 857– 902. (33) Khalil, I. A.; Kogure, K.; Akita, H.; Harashima, H. Pharmacol. ReV. 2006, 58, 32–45. (34) Muro, S.; Garnacho, C.; Champion, J. A.; Leferovich, J.; Gajewski, C.; Schuchman, E. H.; Mitragotri, S.; Muzykantov, V. R. Mol. Ther. 2008, 16, 1450–1458. (35) Rejman, J.; Oberle, V.; Zuhorn, I. S.; Hoekstra, D. Biochem. J. 2004, 377, 159–169. (36) Gratton, S. E. A.; Ropp, P. A.; Pohlhaus, P. D.; Luft, J. C.; Madden, V. J.; Napier, M. E.; DeSimone, J. M. Proc. Natl. Acad. Sci. U.S.A. 2008, 105, 11613–11618. (37) De Geest, B. G.; Vandenbroucke, R. E.; Guenther, A. M.; Sukhorukov, G. B.; Hennink, W. E.; Sanders, N. N.; Demeester, J.; De Smedt, S. C. AdV. Mater. 2006, 18, 1005. (38) De Koker, S.; De Geest, B. G.; Singh, S. K.; De Rycke, R.; Naessens, T.; Van Kooyk, Y.; Demeester, J.; De Smedt, S. C.; Grooten, J. Angew. Chem., Int. Ed. 2009, 48, 8485–8489. (39) Cohen, J. A.; Beaudette, T. T.; Tseng, W. W.; Bachelder, E. M.; Mende, I.; Engleman, E. G.; Frechet, J. M. J. Bioconjugate Chem. 2009, 20, 111–119. (40) Wong, J. Y.; Leach, J. B.; Brown, X. Q. Surf. Sci. 2004, 570, 119–133.

BM100500V