Cellular Interactions with Photo-Cross-Linked and pH-Sensitive

Nov 9, 2012 - Irene Canton,. §. Dietmar ... Department of Biomedical Science, The University of Sheffield, Western Bank, S10 2TN Sheffield, United Ki...
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Cellular Interactions with Photo-Cross-Linked and pH-Sensitive Polymersomes: Biocompatibility and Uptake Studies Jens Gaitzsch,†,‡ Irene Canton,§ Dietmar Appelhans,† Giuseppe Battaglia,*,§ and Brigitte Voit*,†,‡ †

Leibniz-Institut für Polymerforschung Dresden e.V., Hohe Straße 6, 01067 Dresden, Germany Organic Chemistry of Polymers, Technische Universität Dresden, 01062 Dresden, Germany § Department of Biomedical Science, The University of Sheffield, Western Bank, S10 2TN Sheffield, United Kingdom ‡

S Supporting Information *

ABSTRACT: Polymeric nanoparticles, specifically polymersomes, are at the leading edge of the rapidly developing field of nanotechnology. However, their use for biological applications is primarily limited by the biocompatibility of the components. Hence, optimization of polymersome synthesis protocols should carefully consider aspects of cellular toxicity. In this work, we investigate the viability of HDF and HeLa cells treated with photo-cross-linked and pH-sensitive polymersomes. We demonstrate how aspects of polymersome preparation conditions such as cross-linking density and UV irradiation time may affect their cytotoxic properties. Additionally, we also study the cellular uptake of our polymersomes into the cell types mentioned.

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INTRODUCTION Mimicking biological structures at the nanoscale by chemical means has gained increased interest among scientists within the past decade, particularly to create functional biological tools out of wholly synthetic materials.1,2 In this perspective, studies comprising the use of cell-like synthetic membranes allow for a better understanding of biological processes.3,4 These membranes are also important to design novel synthetic bionanotools, for example, polymeric vesicles (polymersomes) made of amphiphilic block copolymers.5−9 Due to their polymeric nature, they can be adjusted easily for specific applications by choosing monomers with specific sensitivities10,11 and allow for transmembrane proteins to be incorporated.6,12 This results in more control over the membrane properties, in particular, membrane stability, permeability, and controlled diffusion processes.13−15 Especially for drug-delivery systems, the use of sensitive monomers is a crucial point because they allow for a triggered disintegration of the polymersome and an efficient delivery process.10,16−18 Recently, polymersomes have attracted growing interest based on their virus-mimicking physical properties19,20 which emphasize their potential use in medicine, especially as drug delivery system and potential platform to mimic organelles as biological nanoreactors.10,21−24 In particular, the use of block copolymers that promote biocompatibility as well as pHsensitivity has produced polymersomes with membranes able to disassemble and escape the endolysosomal compartments, enhancing intracellular delivery of cargoes.21,25 In contrast, designs for artificial organelles23,26 often use biologically stable membranes with transmembrane proteins to control diffusion across the polymersome membrane.23,27 Our approach uses a © XXXX American Chemical Society

combination of pH-sensitivity and stability (photo-crosslinking) to produce vesicles with a defined and reproducible swelling−deswelling behavior.24 While the membrane remains intact in this nanoreactor, complete control over the swelling is obtained via pH-sensitivity. However, the biological compatibility of these promising candidates for synthetic biology has not been studied previously. Due to the pH-sensitivity, positive charges are likely to occur in our system. Unfortunately, the use of positively charged nanoparticle carriers for intracellular delivery has extensively been reported to produce cytotoxic and pro-inflammatory effects.28−30 Additionally, the cross-linking process using UV light may induce the formation of toxic particles due to radical formation. Here, we tested the effect of such polymersomes on different cell types and finely tuned the protocols of synthesis to allow for cellular uptake with low cell toxicity.

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MATERIALS AND METHODS

Materials. If not stated otherwise, all chemicals were used as received. Rhodamine B isocyanate, fluorescein isocyanate, MTT (3(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl-2H-tetrazolium bromide), sodium hydroxide, and hydrochloric acid were purchased from SigmaAldrich (U.K.). Phosphate buffered saline (PBS) tablets were purchased from Oxoid Ltd., U.K.; Ultrapure type 1 water was prepared using the Millipore Synergy purification System (MerckMillipore, U.K.). Methods. All polymers (Figure 1a) were synthesized as described previously.13,24 The cross-linker was synthesized within two steps to Received: September 19, 2012 Revised: November 6, 2012

A

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Figure 1. Use of amphiphilic block copolymers C2−10, C2−20, C4−10, and C4−20 in polymersomes formation by photo-cross-linking approach ([2 + 2] cycloaddition) in time-dependent manner (a) resulting in different polymersomes properties, especially, for C4−20 (b); s = seconds; (c) Development of particle diameter determined via DLS (volume intensity plot), the sharp rise indicates cross-linked polymersomes and the consecutive decrease in diameter showing a rise in cross-linking density. give the methacrylate derivative used (details in SI). Within the crosslinker, either an ethanol (C2) or butanol (C4) spacer was used. The final polymer was then synthesized starting from the PEG-Br macroinitiator and the monomers of the second block in the corresponding ratio using standard ATRP conditions (details in SI). Properties of block copolymers and polymersomes are also presented in SI. All polymers differed in their cross-linker (amount or spacer). While the spacer in the cross-linker used (C2 or C4) gave the first part of the polymer name, their cross-linker content within the hydrophobic block (in mol %) gave the second part of the polymer name. Here, the polymers C2−10, C2−20, C4−10, and C4−20 were used. Gel permeation chromatography (GPC) was done using a PL GPC 50+ system, equipped with a PL-AS-RT autosampler (both Polymer Laboratories, UK). The measurements were carried out at a concentration of 1 mg/mL in THF, which was also the solvent in the GPC, at 40 °C and a flow of 1 mL/min. The results were recorded using an RI detector and calculated against a polystyrene standard. DLS studies of 2 g/L aqueous vesicle solutions were carried out over a range of pH at 25 °C using a ZETASIZER Nano series instrument (Malvern Instruments, UK) equipped with a multipurpose autotitrator (MPT-2) and Dispersion Technology Software (version 5.00). The data were collected by the NIBS (noninvasive backscatter) method using a helium−neon laser (4 mW, λ = 632.8 nm) and a fixed angle of 173°. All data were obtained using vol % evaluation, assuming an RI of 1.5 for the polymer. The peak size given is the z-average within the measurements, except for radiation-dependent measure-

ments (Figure 1c), where the peak maximum was used to ensure the detection of disassembled polymersomes. The UV irradiation was carried out within an EXFO Omnicure 1000 (Lumen Dynamics Group Inc., Canada) equipped with a highpressure mercury lamp as the UV source. UV−vis measurements were done at Cary 100 scan (Varian Inc., U.K.), V-630 (Jasco, Japan). They were carried out in a range from 700 to 300 nm using 1 nm steps at a scan speed of 400 nm/min. The hollow fiber filtration (HFF) was performed using a KrosFlo Research IIi (SpectrumLabs, U.S.A.), equipped with a polysulfonebased separation module (MWCO: 500 kDa, Spectrum Laboratories, U.S.A.). Staining of maltose-modified hyperbranched poly(ethylene imine) (PEI-Mal): PEI-Mal (possessing structure B with ⌀ of about 10 nm) were synthesized as described previously (SI).27 PEI core has a molecular weight of about 25000 g/mol. Furthermore, PEI-Mal possesses enough secondary amino groups for staining with dye molecules.27 A total of 40 mg of PEI-Mal was dissolved in 1 mL of H2O. Then, 0.8 mg of rhodamine B isothiocyanate or fluorescein isocyanate was dissolved in 0.2 mL of DMSO and both solutions are mixed and stirred for 14 h. To remove any free fluorescein or rhodamine B, the solution was purified using a KrosFlo research IIi system with an MWCO of 100 kD until no free dye was detected in the filtrate. Freeze-drying was used to remove residual solvent. Preparation of polymersomes in buffer-free water: A solution of 0.2 wt % polymer C2−10, C2−20, C4−10, or C4−20 in acid (pH = 2) B

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rhodamine B fluorescent molecules were subsequently energized with a green (532 nm) laser and their signals, along with particle-scattered light, were detected (yellow parameter 564 nm), and then processed, by BD FACSArray electronics. Flow data were collected from N = 3 independent experiments and population mean values ± SEM were represented. Imaging cellular uptake of PEI-Mal polymersomes. Live fluorescence microscopy: Cells were seeded at a density of 5 × 103 cells/well in BD Falcon 96-well imaging plates and grown until 50% confluence. Cells were treated with 1 mg/mL of PEI-Mal encapsulated C4−20 polymersomes overnight (typically 16 h). The cells were finally washed three times with PBS and, finally, Hoechst 33342 solution (Thermo Scientific, U.K.) was added at 1 μg/mL for 5 min as a live imaging nuclear counterstain. Stained cells were imaged with the optical spinning disk (obj. 20×).

water (previously deionized) was prepared and stirred until the whole polymer is dissolved. The final solution was passed through a 0.2 μm nylon filter to remove any remaining particles, including dust. Now, 1 M NaOH was added through a 0.2 μm nylon filter until pH 10 was reached. When polymersomes were formed, the solution became turbid. The vesicles were characterized using DLS. Preparation of polymersomes in PBS: A solution of 0.4 weight-% polymer C2−10, C2−20, C4−10 or C4−20 in acid (pH = 2) water (previously deionized) was prepared and stirred until the whole polymer was dissolved. The final solution was passed through a 0.2 μm nylon filter to remove any remaining particles, including dust. This solution was mixed with an equal amount of 0.01 M PBS buffer solution. Following this, 1 M NaOH was added through a 0.2 μm nylon filter until pH 10 was reached. When vesicles were formed, the solution appeared turbid. The vesicles were characterized using DLS. Cross-linking of polymersomes: Polymersome dispersions were placed in a UV chamber and irradiated for either 30, 120, or 600 s. Encapsulation of PEI-Mal into polymersomes and consecutive cleaning was done as described previously.24 The PEI-Mal was premixed with the dissolved polymer and encapsulated during the selfassembly process. Any nonincluded cargo was separated. Cell culture: Primary human dermal fibroblasts (HDFs) were obtained from LGC standards (Teddington, U.K.). Cells were maintained in DMEM (Biosera, U.K.) supplemented with 10% (v/ v) fetal calf serum, 2 mM L-glutamine, 100 IU/mL penicillin, 100 mg/ mL streptomycin, and 0.625 μg/mL amphotericin B (all from SigmaAldrich, U.K.). Cells were subcultured routinely using 0.02% (w/v) trypsin-EDTA (Sigma-Aldrich, U.K.) and were used for experimentation between passages 4 and 8. HeLa cells (ATCC CCL2) were also maintained in the same culture conditions. Polymersomes dispersions were sterilized by filtration through 0.2 μm nylon filters prior to adding to cell cultures. The material was not held back in a substantial amount (verified by visual and DLS analysis) Flow cytometry was carried out using a BD FACS Array bioanalyzer (BD, U.S.A.). Cellular imaging was performed using a spinning disk confocal microscopy (BD, U.S.A.) at 20× objective magnification. MTT-ESTA: The MTT (3-(4,5-dimethyl-2-thiazolyl)-2,5-diphenyl2H-tetrazolium bromide) assay was used to measure cellular metabolic potential as a eluted stain bioassay (ESTA) of treated cells after polymersome treatment. In brief, 3−4 × 104 HDF or HeLa cells were cultured per well in 24-well plates until 70% confluence (typically 48 h). Cells were incubated for 24 h with varying concentrations of the polymersome solutions. Polymersome solutions were prepared according to the different protocols described previously. Then they were tested to determine the effect on cellular viability. After treatment, cell cultures were thoroughly washed in PBS and then incubated with MTT solution (0.5 mg/mL MTT in PBS, 1 mL per well of 24-well plate) for 45 min at 37 °C and in a 95% air/5% CO2 environment. Intracellular dehydrogenase activity reduces MTT to a purple-colored formazan salt. After 45 min, the solution was aspirated and the insoluble intracellular formazan product was solubilized and released from cells by adding acidified iso-propanol (0.3 mL per well of 24-well plate or 1 mL/cm2 cultured tissue) and was incubated for 10 min. The optical density at 570 nm was then measured (with a reference filter at 630 nm) using a plate reading spectrophotometer. For statistical analysis (paired Student’s t-test), experiments were performed in triplicate wells with a total of N = 3 independent experiments. Polymersomes uptake kinetics. Flow cytometry: HDFs cells were seeded at a density of 3−4 × 104 and cultured in 24 wells for two days until 70% confluence. Cells were then treated with rhodamine B labeled polymersomes at a concentration of 1 mg/mL in normal cell medium. Cellular uptake kinetics of polymersomes was then monitored over 24h at different time points by flow cytometry. Briefly, cells were washed twice with PBS following the treatments and then detached using trypsin-EDTA solution. Afterward, cells were pelleted, resuspended and plated in 96-well plates in cold PBS in preparation for automated sampler flow analysis using BD FACSArrayTM bioanalyzer. A total number of 10000 events (cells) per sample were assayed for rhodamine B detection. Intracellular

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RESULTS AND DISCUSSION Inspired by the fact that polymersomes have been reported to be promising tools in many biological applications7,16,22,31,32 and our recently published data showing that pH-sensitive, photo-cross-linkable polymersomes allow for mechanical and chemical control of the membrane permeability, 24 we postulated that these polymersomes would be excellent candidates to create nanoreactors,13,24 providing that they are nontoxic to the target cells. For these studies we used our previously synthesized amphiphilic block copolymers (Figure 1).13,24 The hydrophilic part of them consists of the well-known biocompatible and nonfouling polyethyleneglycol (PEG).7,33 The hydrophobic part is a statistical mixture of the pH-sensitive poly(diethylaminoethylmethacrylate) (PDEAEM) 34,35 and a photo-cross-linking unit [3,4-dimethyl maleic imido ethyl methacrylate (DMIEM; Figure 1) with a C2-spacer in C2−10 and C2−20; 3,4-dimethyl maleic imido butyl methacrylate (DMIBM; Figure 1) with a C4-spacer in C4−10 and C4−20]. We synthesized polymers with 10 mol % (C2−10 and C4−10) as well as 20 mol % (C2−20 and C4−20) cross-linker content within the hydrophobic part. The data of C2−10, C2−20, C4− 10, and C4−20 polymers are comprised in Table 1 and all polymers were used for the formation of polymersomes. We anticipated three potential sources of toxicity to be taken into consideration during the development of the synthesis protocols of nanoreactors from the polymers used: (I) residual Table 1. Properties of the Different Polymers C2−10, C2− 20, C4−10, and C4−20 and Their Polymersomes name C2− 10 C2− 20 C4− 10 C4− 20

a

molar mass (Mn;b kg/mol)

polymersome sizec (nm)

pKa,swd (in PBS)

pKa,swd (in H2O)

time to cross-linke (s)

19.0

125

7.7

7.3

180

21.5

120

7.6

7.0

120

20.0

100

7.8

7.1

120

22.5

125

7.4

6.8

30

a

Name of polymer used (structure shown in Figure 1). bMn is determined from signal intensities in the 1H NMR spectra; the PDI (determined via GPC) of the copolymers is 1.3 (1.4 for C4−10). c Polymersome diameter determined via DLS, shown as peak diameter of intensity; PDI of polymersomes always 0.2. dpKa,sw value of polymersomes dependent on cross-linking time. eShortest crosslinking time for fabricating stable polymersomes determined via DLS (peak diameter), see Figure 1. C

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Figure 2. MTT-ESTA viability test on HeLa cells and HDFs after 24 h incubation with C4−20 polymersomes (c = 0.02 mg/mL, cross-linked 30 s) produced under different treatment conditions (PBS: UV irradiation in buffer; PBS, HFF: photo-cross-linking in buffer followed by exchange of buffer by nonirradiated buffer; No PBS: photo-cross-linking in buffer-free water; No PBS, HFF: exchange of solvent by nonirradiated H2O after UV irradiation; PEI-Mal: encapsulated PEI-Mal in C4−20 polymersomes, including photo-cross-linking in buffer-free water).

needed the shortest irradiation time (30 s) to reach full membrane stabilization. As shown in Figure 1c, the longer the irradiation time and, hence, the higher the cross-linking density, the less polymersomes swell. Achieving large cross-linking density completely hinders the polymersomes to swell. An important aspect to be considered in the design of protocols for polymersomes production suited for biological use is the pH and ionic strength of the solution. Typical physiological pH and ionic strength ranges for mammalian cell culture are pH between 7 and the optimal 7.4 and ionic strength between 0.1 and 0.2 (corresponding to 100−200 mmol/L of sodium chloride in phosphate-buffered saline (PBS).38,39 However, it is well-known that cells contain subcellular compartments specialized in the digestion of exogenous materials upon endocytosis where the pH is considerably lower than the physiological pH.16,40 These compartments, known as endosomes and lysosomes (depending on the stage of the endocytic process), are characterized by an acidic lumen,41 with early endosomes having a pH of 6.5 and lysosomes with a pH of 4.5.40,42 The real challenge of creating a nanoreactor that is responsive to pH is to maintain the cargo shielded and protected in the unfavorable intracellular environment until controlled release occurs under optimal conditions for stability.16 We therefore carefully monitored the pHdependent swelling behavior of each type of cross-linked polymersomes by evaluating their pKa,sw values in a phosphatefree environment and in PBS (DLS study, SI). The pKa,sw is defined as the degree of 50% of polymersome swelling. Furthermore, the pKa,sw is the starting point for polymersome permeability when decreasing the pH, which is accompanied by the protonation of the amino groups in PDEAEM.13 For efficient biological use as nanoreactors, swelling of the polymersomes should start between the mild acidic pH of the early endosomes (6.5) and the physiological pH of the cell media (7.4). Hence, the targeted range of the pKa,sw of the polymersomes is between 6.5 and 7.4. A thorough DLS study (Figure 5-SI) of all the polymersome formulations yielded a characteristic pH-dependent swelling behavior with the corresponding pKa,sw values listed in Table 1. If measured in deionized water, we observed pKa,sw values of all polymersomes

chemicals from the polymer synthesis; (II) positive charges arising from the PDEAEM protonation; (III) free radicals that induce possible formation of toxic byproducts during the UVirradiation of the photo-cross-linking process. We started by finely tuning the photo-cross-linking process in order to produce a stable and defined swelling behavior of the membrane with the minimum irradiation time to avoid unwanted byproducts of the UV-irradiation process. In comparison to chemical cross-linking,36 photo-crosslinking with UV light is considerably faster. To avoid generation of free radical side-reactions that may compromise cellular viability,37 we kept the UV-irradiation time of polymersomes to the minimum possible in order to produce stable cross-linking. Thus, we tested the effect of the irradiation time on the crosslinking efficiency in our different polymers C2−10, C2−20, C4−10, and C4−20 (Figure 1). Table 1 also collects the different properties of polymersomes obtained from the corresponding polymers. In our polymersomes, UV irradiation leads to the formation of a cyclobutane ring between the polymer chains forming the membrane (SI).13 Stable cross-linking is reached, once the bonds prevent polymersome disintegration upon acidification. Experimentally, this was determined by DLS, measuring particle diameter of polymersomes at pH 4 (SI). When enough cross-linking bonds are established within the polymersome membrane, no disassembly is detected at pH 4, but the overall diameter increases.24 Disassembly is due to the positive charges of the pH-sensitive PDEAEM parts evolving upon acidification. Once cross-linked, these groups still repel each other upon acidification but are stopped at a certain point by the crosslinking bonds, which results in the swelling observed. Therefore, a sudden increase in diameter was visible, once UV irradiation time was enough to create a sufficient number of cross-linked units in the polymersomes to withstand the low pH condition (Figure 1c). Our results revealed, as expected, that polymersomes with higher cross-linker density require shorter irradiation times (Table 1 and Figure 1c). Similarly, longer cross-linking units (C4 polymers in comparison to C2 polymers) resulted in shorter cross-linking times (Table 1). Consequently, polymersomes made of the C4−20 polymers D

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Figure 3. MTT-ESTA viability test of C4−20 polymersomes at various concentrations on HeLas (a) HDFs (b) combined with the influence of UV irradiation time on polymersomes cross-linking state (see also Figure 1); pH dependency of the zeta potential of cross-linked C4−20 polymersomes in dependence of UV irradiation time (c); Changes in charge (blue, d) development upon increasing cross-linking density. Non-cross-linked polymersomes disassemble into positively charged polymers, semi-cross-linked ones swell and evolve a surface charge, while densely cross-linked polymersomes are only charged inside the membrane (d): s = seconds (a−d).

within the desired pH range (6.8 − 7.3) (Table 1 and SI). However, when measured in the more physiologically relevant PBS only the pKa,sw of polymersomes made of C4−20 polymer (7.4) was suitable (Table 1 and SI), while all other pKa,sw were too high (pKa,sw > 7.4). Thus, C4−20 polymersomes combine the shortest UV irradiation time (30 s) and the optimal pKa,sw of all the polymers tested in PBS (pKa,sw = 7.4). Therefore, vesicles consisting of the C4−20 polymer were the choice for cell biocompatibility testing. To study the effect of C4−20 polymersomes on cellular viability under different production protocols (Figure 2) we used HeLa cells and primary HDF cells. Toxicity studies were carried out using the MTT assay8 based on the metabolic activity of intracellular reductases (mainly mitochondrial enzymes).43 Using this assay, we observed a higher toxic effect of polymersomes, prepared and diluted in PBS compared to those produced in DI water as a starting solution (Figure 2). This was very likely due to the formation of phosphate radicals associated with toxic subproducts during UV irradiation in PBS.44,45 To remove any small molecules inducing cellular toxicity, an additional purification step using hollow fiber filtration (HFF, MWCO = 500 kDa) was introduced. Here, the irradiated PBS is exchanged by nonirradiated PBS buffer solution, without affecting the polymersomes stability.24 This

procedure slightly improved the biocompatibility of the polymersomes (Figure 2a). Interestingly, when the same purification process was applied on polymersomes prepared in a buffer-free solution, no effect on cell viability was observed, indicating that the toxic subproducts are linked to the PBS buffer (Figure 2). Furthermore, this proves that the majority of the toxic compounds are bound to the polymer, because the HFF purification leads to replacement of the irradiated buffer against a fresh one. Any remaining toxicity must therefore result from toxic compounds bound to the polymer chains. To include more than just an empty lumen into the cells, a large enzyme-sized cargo was internalized into our vesicles. Here, the soft maltose decorated hyperbranched poly(ethylene imine) (PEI-Mal; ⌀ ∼ 10 nm)46 nanoparticles are a suitable candidate due their supreme biological properties as well as their ability to load drugs. To remove any free PEI-Mal, the solution needs to be HFF-cleaned, which is a known method to retain PEI-Mal molecules within the polymersomes.24 Due to the results just discussed, we could be sure that this cleaning would not significantly change the toxicity levels. Surprisingly, their enclosure lead to cell viability levels of 90% and above for both cell types tested, meaning the loss of all toxicity (Figure 2). Even the viability for HeLa cells, which could not be pushed above 60% previously, now topped 90% (Figure 2). Here, it can E

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potential. This is consequently leading to the positive charges being confined in the inner part of the polymersome membrane. Our data (Figure 3c) confirmed that the fully cross-linked status on our polymers was achieved with an irradiation time of 600 s, because reduced cationic zeta potential values are maintained at acidic pH values. We then tested the effect of UV-irradiation time (and thus surface charge) on cellular viability via MTT (Figure 3a,b). For non-cross-linked and less densely cross-linked polymersomes (0, 30, and 120 s of UV-irradiation), no clear dependency of cellular toxicity and irradiation time was visible. While the positive charges decreased for increasingly cross-linked vesicles, they were now concentrated at the polymersome surface and not diluted throughout the solution. This lack of dilution opposes the decreasing overall charge and results in an unchanged cell viability observed. Interestingly, densely crosslinked polymersomes (600 s of UV-irradiation) yielded much more biocompatible vesicles in both primary HDFs and HeLa cells (Figure 3a,b). For these vesicles, the charges are covered and not even present and concentrated on the surface as for less dense cross-linked polymersomes and cannot influence cell viability. Despite the reduced swelling caused by the dense cross-linking, these vesicles are still feasible for drug release, although at a lower level than less dense cross-linked ones.52 After fine-tuning several processing parameters, we were able to produce highly biocompatible nanoparticles. These nanoparticles could be easily adapted to perform as drug delivery devices or nanoreactors, depending on their ability to encapsulate and deliver cargo. In order to test encapsulation efficiency of polymersomes (preferably with cargo) and to get first insights into cell uptake kinetics the polymersome needed be tracked using optical methods, for example, due to fluorescence. A way to reach this without altering the polymer was to enclose a fluorescent-labeled cargo. As previously discussed and comprised in Figure 2, polymersomes hosting PEI-Mal yield very high levels of cell viability, making this cargo a virtually perfect candidate for our purpose, especially because PEI-Mal can be labeled with various molecules, also dyes, very easily in one step.24,46 Thus, studies using live fluorescence imaging and flow cytometry (Figure 4) for cellular uptake, including their time-scale of this process, were now conducted using dye-marked PEI-Mal loaded into polymersomes. First of all, we conducted live fluorescence micrographs of HeLa cells and HDFs (Figure 4). For both cell types analyzed, an intracellular accumulation of the fluorescent PEI-Mal cargo could be determined. The PEI-Mal, used in this study, does not leave the polymersomes, unless a pressure is applied at an acidic pH.23 While an acidic pH occurs during endolysosomal uptake into the cells,40 no transmembrane pressure as high as applied with the HFF system is present in the cells, because they do not suffer any mechanical stress during microscopy. Hence, we could take the detected cargo as detected polymersomes. The confined space, in which the polymersomes could be detected, not only proofs cellular uptake, but is also another proof for low toxicity values. Because all cargo is nicely agglomerated around the stained nuclei, the cell membranes do not show increased permeability, meaning that the cells can be seen as intact and alive. This result strengthens the results comprised in Figure 2 from the initial MTT study. A powerful way to reach time-resolved quantitative result of the cellular uptake process is the so-called fluorescence activated cell sorting (FACS analysis). Due to autofluorescence, the control experiment already shows an uptake of 0.8% of the

be suspected that the PEI-Mal molecules might enclose and, therefore, cover toxic molecules, which do not leave the polymersomes upon HFF treatment, as PEI-Mal is a known system to load small molecules.46−48 Besides their enclosure of toxic material, this result now allows for PEI-Mal containing polymersomes to be used later in cellular uptake studies. However, treatment with increasing concentrations of the polymersomes on our cells revealed another source of toxicity independent to the filtration process (Figure 3a,b). We anticipated that cell viability could have been compromised due to the positive charge of the C4−20 polymer in acidic pH caused by PDEAEM protonation. Such protonation effects are known to show a concentration-dependent toxic behavior because cationic charge is produced.3 The use of cationic particles in biological systems is controversial. On the one hand, cationic particles often display higher internalization rates compared to anionic or neutral formulations as they interact with the anionic cell membrane.49 This is the key for success of cationic polymeric nanoparticles (for example, chitosan) for intracellular delivery.50 On the other hand, the use of cationic nanoparticle carriers for intracellular delivery has extensively been reported to produce cytotoxic and pro-inflammatory effects.28,29 To avoid toxicity from the positive charges in our systems, we UV-irradiated the nanoparticles for increased periods of time. As the UVirradiation time increases, so the increasing cross-linking density leads to a reduced swelling (Figure 1), assumingly combined with a lower overall surface charge on the nanoparticles. The surface charge should decrease because a lower swelling would allow less-charged PDEAEM groups to reach the polymersome surface. A convenient way to detect positive surface charge was to measure the zeta potential, which was monitored for C4−20 polymersomes at several crosslinking densities (0, 30, 120, and 600 s UV-irradiation applied). With increasing UV-irradiation time, a decrease in zeta potential was observed at physiological pH (Figure 3a). When samples were not irradiated, a sharp rise in surface charge was visible between pH 7.4 and 7.0, at the pH range where non-cross-linked polymersomes disassemble. Free positively charged polymer chains present in the solution after disassembly consequently caused a highly positive zeta potential (Figure 3c,d).51 For cross-linked polymersomes, the sharp rise upon pH switch shrinks considerably until it is virtually absent for densely cross-linked polymersomes (i.e., after 600 s of UV-irradiation). As previously mentioned, our cross-linked polymersomes show a characteristic swelling− deswelling behavior upon repeated pH changes. At acidic conditions, all polymer chains are protonated, leading to positive charges repelling each other in order to get the largest distance possible in between them. A combination of both effects, meaning low pH and low cross-linking density in polymersomes, may cause loosely cross-linked chain patches to flip around. Because the repelling force between the positively charged PDEAEM units cannot be decreased by disassembly in the cross-linked state, the polymer chains disorder and mix formerly hydrophobic and hydrophilic parts to increase entropy as much as possible. As a consequence, they reveal the positive charge to the outside of the vesicle and cause an increased zeta potential (Figure 3c,d). Logically, an increased cross-linking density means that the polymer chains are forced to remain in their ordered state and less loosely cross-linked patches are present in the membrane. A decreased number of patches leads to a decreased surface charge and, thus, also to a lower zeta F

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in PBS proved to be toxic, probably due to phosphate-radicalassociated toxic subproducts from PBS formed during the UV irradiation, the ones prepared in buffer-free water were highly biocompatible. In addition, their pH-dependent cationic surface charge resulted in increased toxicity. Dense cross-linking by long UV irradiation aided to exhibit the PEG corona as dominant outer component, which resulted in shielding of the positive charge and a greater biocompatibility. With nontoxic material in hand, it was logical to investigate the cellular uptake behavior as well, this time including cargo. According to the results presented, our pH-sensitive and crosslinked polymersomes with a hyperbranched polymer as sample cargo are internalized rapidly. Optical images of both a cell-line and primary cells revealed a good internalization of our loaded vesicles after 24 h. Additionally, a kinetic study on the primary cells used showed a continuous uptake behavior. These promising results show the great potential of our system as drug delivery system or synthetic and internalized organelle. We would like to point out that our work shows, in a very obvious manner, how side reactions and preparation conditions can turn a material from toxic at low concentrations into a nontoxic material, which transports cargo. According to the results presented, also minor details, like the use of a phosphate buffer, can have a great impact on cellular toxicity levels. We are confident that these results motivate many in the community of scientist to closely analyze every synthetic detail in order to make their system work out nicely in the end. In further studies, we aim to replace our cargo by enzymes and to trigger enzymatic reactions within polymersomes once they entered the cell. This may allow for synthetic vesicles without transmembrane proteins to be used in synthetic biology.

Figure 4. Live fluorescence micrographs (×20) of stained polymersomes uptake on HDFs. The nucleus is stained blue (Hoechst). Intracellular delivered cargo (PEI-Mal within polymersomes) is shown in green color FACS analysis (shown below) of the uptake kinetics (images for 0, 9, and 24 h, data only for 3 and 6 h) of C4−20 polymersomes with rhodamine-labeled PEI-Mal by HDFs. The number represents the fluorescence intensity detected and the percentage given represents the total percentage of cells positive for uptake.

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ASSOCIATED CONTENT

S Supporting Information *

Polymer synthesis, as well as characterization of polymersome properties, is included. This material is available free of charge via the Internet at http://pubs.acs.org.

whole population, while the fluorescence intensity is automatically set to “1.0” (Figure 4, 0 h). For the PEI-Mal-loaded polymersomes investigated, efficient uptake kinetics could be monitored for the HDFs (cell type investigated here).16 Rising continuously over 19% after 3 h of incubation to 40% after 6 h, almost two-thirds of the cell population (63%) already show internalized polymersomes after only 9 h of incubation. Besides the number of cells, the fluorescence intensity has also risen considerably over 5.0 (3 h) and 7.3 (6 h) to 9.3 (9 h, Figure 4) during this time of incubation. After 1 day of cellular treatment, almost the whole cell population (86%) is positive for polymersomes uptake. Although the number of cells only increased by one-third from 9 to 24 h of incubation time, the amount of polymersomes, meaning the fluorescence intensity, almost doubled to about 17 (Figure 4). This result mainly evidenced that polymersome uptake continues after an initial amount entered the cells. Hence, FACS proofs not only a fast, but also continuous polymersome uptake kinetics into HDFs.

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AUTHOR INFORMATION

Corresponding Author

*Fax: +493514658565 (B.V.); +441142225945 (G.B.). Tel.: +493514658590 (B.V.); +441142222305 (G.B.). E-mail: voit@ ipfdd.de (B.V.); g.battaglia@sheffield.ac.uk (G.B.). Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS Financial support of the Rosa-Luxemburg Foundation and the Dresden International Graduate School for Biomedicine and Bioengineering (DIGS-BB) is gratefully acknowledged.

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REFERENCES

(1) Szostak, J. W.; Bartel, D. P.; Luisi, P. L. Nature 2001, 409, 387. (2) Schwille, P. Science 2011, 333, 1252. (3) Hoffmann, F.; Cinatl, J.; Kabickova, H.; Cinatl, J.; Kreuter, J.; Stieneker, F. Int. J. Pharm. 1997, 157, 189. (4) Luisi, P. L.; Walde, P.; Oberholzer, T. Curr. Opin. Colloid Interface Sci. 1999, 4, 33. (5) Discher, D. E.; Discher, B. M.; Won, Y. Y.; Ege, D. S.; Lee, J. C. M.; Bates, F. S.; Hammer, D. A. Science 1999, 284, 1143. (6) Kim, K. T.; Meeuwissen, S. A.; Nolte, R. J. M.; van Hest, J. C. M. Nanoscale 2010, 2, 844.

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CONCLUSIONS In summary, we demonstrated the fabrication of polymersomes with a definite swelling, depending on their cross-linking state given by the UV irradiation time. This is in large contrast to other polymersomes that disassemble upon a pH switch.10,31 A main goal of our work was to create nontoxic swelling polymersomes. While polymersomes prepared and cross-linked G

dx.doi.org/10.1021/bm3014704 | Biomacromolecules XXXX, XXX, XXX−XXX

Biomacromolecules

Article

(7) Blanazs, A.; Armes, S. P.; Ryan, A. J. Macromol. Rapid Commun. 2009, 30, 267. (8) Battaglia, G.; Ryan, A. J.; Tomas, S. Langmuir 2006, 22, 4910. (9) Zhang, X. Y.; Tanner, P.; Graff, A.; Palivan, C. G.; Meier, W. J. Polym. Sci., Part A: Polym. Chem. 2012, 50, 2293. (10) Onaca, O.; Enea, R.; Hughes, D. W.; Meier, W. Macromol. Biosci. 2009, 9, 129. (11) Le Meins, J. F.; Sandre, O.; Lecommandoux, S. Eur. Phys. J. E 2011, 34. (12) Meier, W.; Egli, S.; Nussbaumer, M. G.; Balasubramanian, V.; Chami, M.; Bruns, N.; Palivan, C. J. Am. Chem. Soc. 2011, 133, 4476. (13) Gaitzsch, J.; Appelhans, D.; Grafe, D.; Schwille, P.; Voit, B. Chem. Commun. 2011, 47, 3466. (14) Lensen, D.; Vriezema, D. M.; van Hest, J. C. M. Macromol. Biosci. 2008, 8, 991. (15) Christian, D. A.; Cai, S.; Bowen, D. M.; Kim, Y.; Pajerowski, J. D.; Discher, D. E. Eur. J. Pharm. Biopharm. 2009, 71, 463. (16) Canton, I.; Battaglia, G. Chem. Soc. Rev. 2012, 41, 2718. (17) Lee, J. S.; Feijen, J. J. Controlled Release 2012, 161, 473. (18) De Oliveira, H.; Thevenot, J.; Lecommandoux, S. Wiley Interdiscip. Rev.-Nanomed. Nanobiotechnol. 2012, 4, 525. (19) Lewis, A.; Battaglia, G.; Canton, I.; Stratford, P. A61K, 47/48 ed.; Biocompatibles UK Ltd.: United Kingdom, 2009. (20) Lomas, H.; Canton, I.; MacNeil, S.; Du, J.; Armes, S. P.; Ryan, A. J.; Lewis, A. L.; Battaglia, G. Adv. Mater. 2007, 19, 4238. (21) Lomas, H.; Du, J. Z.; Canton, I.; Madsen, J.; Warren, N.; Armes, S. P.; Lewis, A. L.; Battaglia, G. Macromol. Biosci. 2010, 10, 513. (22) Renggli, K.; Baumann, P.; Langowska, K.; Onaca, O.; Bruns, N.; Meier, W. Adv. Funct. Mater. 2011, 21, 1241. (23) Meier, W.; Tanner, P.; Egli, S.; Balasubramanian, V.; Onaca, O.; Palivan, C. G. FEBS J. 2011, 278, 32. (24) Gaitzsch, J.; Appelhans, D.; Wang, L. G.; Battaglia, G.; Voit, B. Angew. Chem., Int. Ed. 2012, 51, 4448. (25) Blanazs, A.; Massignani, M.; Battaglia, G.; Armes, S. P.; Ryan, A. J. Adv. Funct. Mater. 2009, 19, 2906. (26) van Dongen, S. F. M.; Verdurmen, W. P. R.; Peters, R. J. R. W.; Nolte, R. J. M.; Brock, R.; van Hest, J. C. M. Angew. Chem., Int. Ed. 2010, 49, 7213. (27) Broz, P.; Driamov, S.; Ziegler, J.; Ben-Haim, N.; Marsch, S.; Meier, W.; Hunziker, P. Nano Lett. 2006, 6, 2349. (28) Jacobsen, N. R.; Moller, P.; Jensen, K. A.; Vogel, U.; Ladefoged, O.; Loft, S.; Wallin, H. Part. Fibre Toxicol. 2009, 6. (29) Omidi, Y.; Barar, J.; Akhtar, S. Curr. Drug Delivery 2005, 2, 429. (30) Lonez, C.; Vandenbranden, M.; Ruysschaert, J. M. Prog. Lipid Res. 2008, 47, 340. (31) Brinkhuis, R. P.; Rutjes, F. P. J. T.; van Hest, J. C. M. Polym. Chem. 2011, 2, 1449. (32) Pegoraro, C.; MacNeil, S.; Battaglia, G. Nanoscale 2012, 4, 1881. (33) Hillmyer, M. A.; Petersen, M. A.; Yin, L. G.; Kokkoli, E. Polym. Chem. 2010, 1, 1281. (34) Adams, D. J.; Butler, M. F.; Weaver, A. C. Langmuir 2006, 22, 4534. (35) Letchford, K.; Burt, H. Eur. J. Pharm. Biopharm. 2007, 65, 259. (36) Chambon, P.; Blanazs, A.; Battaglia, G.; Armest, S. P. Langmuir 2012, 28, 1196. (37) Lin, F.; Wu, J. L.; Liu, J. G. J. Photochem. Photobiol., A 2000, 131, 49. (38) Tipton, K. F.; Dixon, H. B. Methods Enzymol. 1979, 63, 183. (39) Phelan, M. C. In Current Protocols in Cell Biology; Bonifacino, J. S., et al., Eds.; Wiley: New York, 2007; Chapter 1, Unit 1 1. (40) Lee, R. J.; Wang, S.; Low, P. S. Biochim. Biophys. Acta 1996, 1312, 237. (41) van Dongen, S. F. M.; Nallani, M.; Cornelissen, J. L. L. M.; Nolte, R. J. M.; van Hest, J. C. M. Chem.−Eur. J. 2009, 15, 1107. (42) Murphy, R. F.; Powers, S.; Cantor, C. R. J. Cell Biol. 1984, 98, 1757. (43) Mosmann, T. J. Immunol. Methods 1983, 65, 55. (44) Huang, P.; Feng, L.; Oldham, E. A.; Keating, M. J.; Plunkett, W. Nature 2000, 407, 390.

(45) Passi, S.; Picardo, M.; Zompetta, C.; Deluca, C.; Breathnach, A. S.; Nazzaroporro, M. Free Radical Res. Commun. 1991, 15, 17. (46) Appelhans, D.; Komber, H.; Quadir, M. A.; Richter, S.; Schwarz, S.; van der Vlist, J.; Aigner, A.; Muller, M.; Loos, K.; Seidel, J.; Arndt, K. F.; Haag, R.; Voit, B. Biomacromolecules 2009, 10, 1114. (47) Hobel, S.; Loos, A.; Appelhans, D.; Schwarz, S.; Seidel, J.; Voit, B.; Aigner, A. J. Controlled Release 2011, 149, 146. (48) Polikarpov, N.; Appelhans, D.; Welzel, P.; Kaufmann, A.; Dhanapal, P.; Bellmann, C.; Voit, B. New J. Chem. 2012, 36, 438. (49) Harush-Frenkel, O.; Debotton, N.; Benita, S.; Altschuler, Y. Biochem. Biophys. Res. Commun. 2007, 353, 26. (50) Kim, S. H.; Jeong, J. H.; Chun, K. W.; Park, T. G. Langmuir 2005, 21, 8852. (51) Agarwal, A.; Unfer, R.; Mallapragada, S. K. J. Biomed. Mater. Res., Part A 2007, 81A, 24. (52) Yassin, M.; Appelhans, D.; Mendes, R.; Rümmeli, M.; Voit, B. Chem.Eur. J. 2012, 18, 12227.

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