Biocompatible Polymeric Nanoparticles as Promising Candidates for

May 27, 2015 - (4, 5) Therefore, the interactions of NPs with the immune system, which constitutes the first line of defense against foreign assault, ...
0 downloads 0 Views 9MB Size
Download PDF
Article pubs.acs.org/Langmuir

Biocompatible Polymeric Nanoparticles as Promising Candidates for Drug Delivery Sylwia Łukasiewicz,*,† Krzysztof Szczepanowicz,‡ Ewa Błasiak,† and Marta Dziedzicka-Wasylewska† †

Department of Physical Biochemistry, Faculty of Biochemistry, Biophysics and Biotechnology, Jagiellonian University, 30-387 Krakow, Poland ‡ Jerzy Haber Institute of Catalysis and Surface Chemistry, PAS, 30-239 Krakow, Poland S Supporting Information *

ABSTRACT: The use of polymeric nanoparticles (NPs) in pharmacology provides many benefits because this approach can increase the efficacy and selectivity of active compounds. However, development of new nanocarriers requires better understanding of the interactions between NPs and the immune system, allowing for the optimization of NP properties for effective drug delivery. Therefore, in the present study, we focused on the investigation of the interactions between biocompatible polymeric NPs and a murine macrophage cell line (RAW 264.7) and a human monocytic leukemia cell line (THP-1). NPs based on a liquid core with polyelectrolyte shells were prepared by sequential adsorption of polyelectrolytes (LbL) using AOT (docusate sodium salt) as the emulsifier and the biocompatible polyelectrolytes polyanion PGA (poly-L-glutamic acid sodium salt) and polycation PLL (poly L-lysine). The average size of the obtained NPs was 80 nm. Pegylated external layers were prepared using PGA-g-PEG (PGA grafted by PEG poly(ethylene glycol)). The influence of the physicochemical properties of the NPs (charge, size, surface modification) on viability, phagocytosis potential, and endocytosis was studied. Internalization of NPs was determined by flow cytometry and confocal microscopy. Moreover, we evaluated whether addition of PEG chains downregulates particle uptake by phagocytic cells. The presented results confirm that the obtained PEGgrafted NPs are promising candidates for drug delivery.

1. INTRODUCTION Biodegradable polymeric nanoparticles (NPs) are promising carriers for many active compounds. Their high application potential in pharmacology is related to improved therapeutic efficiency, enhanced protection of active molecules from the destructive influence of the external environment, maintenance of drug concentration within acceptable therapeutic limits, and reduction of side effects.1−3 It is well-known that physicochemical parameters of NPs determine their influence on biological systems.4,5 Therefore, the interactions of NPs with the immune system, which constitutes the first line of defense against foreign assault, should be considered in the design of new nanoparticle carriers, especially for drug delivery. Macrophages, the major cells of the immune system, are engaged in inflammatory processes. They perform three major functions: phagocytosis, antigen presentation, and immunomodulation via production of various cytokines and growth factors as well as initiation, maintenance, and resolution of inflammation.6 Macrophages react against pathogenic particles, acting as scavengers and internalizing foreign invaders. They can also be activated by artificial nanoparticles used as drug nanocarriers. Studies have shown rapid elimination of NPs from the bloodstream following injection.7 The opsonization process enhances phagocytosis by increasing the visibility of foreign © 2015 American Chemical Society

materials to phagocytic cells. Adsorption of plasma proteins on the surface of particles allows macrophages of the mononuclear phagocytic system (MPS) to recognize and remove nanoparticles before they reach their desired location.8 The process reduces the circulating half-life of a drug, thus limiting the ability of nanoparticles to act as effective nanocarriers in the drug delivery platform. Therefore, the design of nanoparticles invisible to phagocytic cells is important. On the other hand, macrophages can be considered as potential therapeutic targets. The cells can migrate into solid tumors; thus, they are postulated to act as “Trojan horses” carrying lethal doses of chemotherapeutics into tumors.9 NPs are mainly internalized into phagocytic cells via fluid phase endocytosis, receptor-mediated endocytosis, or phagocytosis.7 Nonspecific hydrophobic or electrostatic interactions as well as specific recognition by membrane-anchored receptor proteins determine the initial contact between NPs and the targeted cells.10 The nonspecific binding of NPs depends on certain factors, such as particle charge and shape10,11 or diffusion speed.12 NPs’ ability to bind to macrophages is dependent on Received: November 27, 2014 Revised: May 25, 2015 Published: May 27, 2015 6415

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir

without further purification. Distilled water used in all of the experiments was obtained with the three-stage Millipore Direct-Q 5UV purification system. Materials for Cell Culture Assays. The mouse murine macrophage cell line RAW 264.7 was obtained from Sigma-Aldrich, and the human leukemic monocyte cell line THP-1 was obtained from ATCC. All of the cell culture materials, including DMEM, RPMI 1640, and F12 media, heat-inactivated fetal bovine serum (FBS), MTT (3-[4,5dimethylthiazol-2-yl]-2,5 diphenyl tetrazolium bromide), endocytosis inhibitors (chlorpromazine, amiloride, filipin III), sulforhodamine B, and PMA (phorbol 12-myristate 13-acetate), were purchased from Sigma-Aldrich. The Lactate dehydrogenase (LDH) Cytotoxicity Detection Kit was obtained from Clontech Laboratories. LysoTracerRed and the Vybrant Phagocytosis Kit were obtained from Molecular Probes (Life Technologies). Methods. NP Preparation. The LbL methodology of capsule formation was originally proposed by Sukhorukov in 1998.21 He described the formation of a polyelectrolyte multilayer shell on latex microparticles. Later on, the method was used to create a polyelectrolyte multilayer shell on a liquid nanocore. In the study, nanocapsules were synthesized according to the method described by Szczepanowicz et al. (2010),22 i.e., the direct encapsulation of emulsion drops in polyelectrolyte multilayer shells. Briefly, the oil phase was prepared by dissolution of 340 g/dm3 AOT in chloroform. Polyelectrolytes were dissolved in a NaCl solution with an ionic strength of 0.015 M to obtain a concentration of 2 g/dm3 without pH adjustment. NP cores were formed by the addition of the oil phase (AOT in chloroform) to the polycation (PLL) solution while mixing with the magnetic stirrer. The polyelectrolyte multilayer shells were prepared on the emulsion drops stabilized by the AOT/PLL interfacial complex. Consecutive layers of polyelectrolytes were formed by the layer-by-layer technique using the saturation method. For the preparation of fluorescently labeled nanocapsules, FITC-PAH was used instead of one PLL layer. Because the amount of chloroform should be limited in pharmaceutical products due to its inherent toxicity, it was removed from the suspension using a rotary evaporator. The final concentration of chloroform determined by GC-ECD analysis was 0.0397 mg/dm3, which meets the requirements of the U.S. Department of Health and Human Services Food and Drug Administration, the Center for Drug Evaluation and Research (CDER), and the Center for Biologics Evaluation and Research (CBER) as published in the Guidance for Industry.23 To prepare pegylated NPs, PGA-g-PEG with various grafting ratios was used as the external layer in a multilayer shell. The pegylated NPs were synthesized according to the strategy described by Boulmedais et al. (2004)24 and Szczepanowicz et al. (2010).25 Briefly, 0.1 g of PGA, 0.46 g (1.3 and 2.1 g) of Me-PEG-NH2 (MW 5000), and 0.014 g of NHS were dissolved in 10 mL of water. Then, 0.16 g of EDC was added to the mixture while stirring. The reaction was allowed to proceed for 6 h at room temperature. After filtration, the reaction mixture was dialyzed three times (cut off at MW 12400) for 24 h with deionized water (2 L). Using 1H-NMR, the areas of the glutamic side-chain peaks representing, in this case, one monomer of glutamic acid (4.16 ppm I ∼ 1) were compared with the PEG area (3.55 ppm I ∼ 64, 177, and 278) to determine the graft ratio of the copolymer, assuming that one molecule of PEG (MW = 5000) possesses 456H hydrogen nuclei. The grafting percentage was equal to a ratio of 177/456. The achieved coupling rate was approximately 14, 39, and 61%, and these polymers are referred to as PGA-g(14)-PEG, PGA-g(39)-PEG, and PGA-g(61)-PEG, respectively. NP Size Analysis. The size distribution (the hydrodynamic diameter) of the NPs was determined using a Zetasizer Nano Series from Malvern Instruments by dynamic light scattering (DLS). Each value was obtained as an average from three runs with at least 10 measurements. All of the measurements were performed at 25 °C. Zeta Potential Measurements. Zeta potential was measured by the microelectrophoretic method using a Malvern Zetasizer Nano ZS apparatus. Each value was obtained as an average from three parallel measurements. The zeta potential of the NPs, as well as those of polyelectrolytes in solution, was measured in a 0.015 M NaCl solution. NP Concentration Measurements. NP concentration was determined by nanoparticle tracking analysis (NTA) using an NS500

their charge and surface moieties. Because macrophages display negatively charged sialic acids on their surface, positively charged NPs are more likely to affect macrophages and induce inflammation than the anionic or neutral NPs.6 However, it is postulated that anionic NPs can bind to scavenger receptors of macrophages13 or attach to cells nonspecifically.14 When NPs are not bound to the plasma membrane but are located close to it, they can be internalized coincidentally by passive engulfment into membrane invaginations.10 Additionally, it has been shown that the particle size contributes to the generation of macrophage response.15 Despite the lack of clear evidence that the NP size influences cellular attachment, it appears that it affects the cellular uptake rate and the mechanism of internalization.10 Particles of approximately 2 μm and those that have a hydrophobic surface are most effectively phagocytosed.11 The internalization rates of particles smaller or larger than 2 μm and those with a hydrophilic surface are lower.11 The internalization process strongly depends on the surface properties of NPs, which influence interactions with plasma proteins, consequently affecting the NP biodistribution in the body.16,17 Functionalization of NPs by adding specific moieties capable of selective recognition of receptors expressed only on target cells plays a key role in NP specific delivery.18 Additionally, decorating the surface area with neutral hydrophilic polymers, e.g., polyethylene glycol (PEG-grafting), increases the circulation time of the NPs in the bloodstream through minimizing or eliminating protein adsorption by blocking electrostatic and hydrophobic interactions that enable opsonin binding. This effect is related to the PEG properties. Uncharged hydrophilic residues and high surface mobility lead to high steric exclusion.19 Creating a hydrophilic layer around the particles also protects them by suppressing their uptake by macrophages. However, pegylation can improve NP internalization by other cell types (for example, cancer cells).7 Recently, we showed that polymeric NPs formed by the layerby-layer (LbL) method using a polycation PLL (poly-L-lysine) and a polyanion PGA (poly-L-glutamic acid sodium salt) were promising candidates as nanocarriers in pharmacology due to their relatively low toxicity and biocompatibility.20 This method is essentially dedicated to hydrophobic drug encapsulation and hydrophobic phase-soluble materials. However, knowledge about the interactions of the NPs with the immune system remains limited. Therefore, in the present work, we focused on the studies of the interactions between biocompatible polymeric PLL−PGA NPs and phagocytic cells (mouse murine macrophage cell line RAW 264.7 and human leukemic monocyte cell line THP-1). To eliminate nonspecific binding and to improve biocompatibility of the NPs, pegylated external layers were prepared using PGA grafted by PEG (PGA-g-PEG). The influence of the physicochemical properties of the obtained NPs (charge, size, surface modification) on viability, phagocytosis potential, and endocytosis was elucidated.

2. EXPERIMENTAL SECTION Materials. NP Preparation. The polycations poly-L-lysine hydrobromide (PLL, MW ∼15000−30000) and poly(fluorescein isothiocyanate allylamine hydrochloride) (FITC-PAH, MW ∼70000), the polyanions poly-L-glutamic acid sodium salt (PGA, MW ∼15000− 50000) and methoxypolyethylene glycol amine (Me-PEG-NH2, MW ∼5000), N-hydroxysulfo succinimide sodium salt (NHS, ≥98.5%), N(3-(dimethylamino)propyl)-N′-ethylcarbodiimide (EDC, ≥97%), docusate sodium salt (AOT), chloroform, and sodium chloride were obtained from Sigma-Aldrich. All of the materials were used as received 6416

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir

of 15 μL per well (ca. 0.15 × 106 NPs/cell). Following incubation, the medium containing NPs was removed, and cells were incubated for the next 2 h at 37 °C while resuspended in HBSS buffer with fluoresceinlabeled E. coli particles. The negative control was free medium, and the positive control consisted of untreated cells. The background fluorescence, resulting from noninternalized E. coli, was quenched by adding Trypan Blue. The results were acquired using a fluorescence plate reader (TECAN Infinitive200Pro) (488 nm excitation, 520 nm emission wavelength). Five replicates for each experimental condition were performed. The obtained results represent the average from three independent experiments. Flow Cytometry Experiments. The quantitative cellular uptake of different types of NPs in RAW 264.7 and THP-1 cells was measured by flow cytometry. Before the experiment, the cultured medium was replaced with fresh medium with 10% FBS containing various types of NPs that were fluorescently labeled (FITC-labeled) at a volume of 30 μL per well (ca. 0.6 × 105 NPs/cell). After 0.5 or 2 h of incubation under standard culture conditions, cells were washed five times with cold PBS (phosphate buffer saline, pH 7.4) to remove unincorporated NPs and were eventually suspended in 1 mL of cold PBS for analysis. To determine the degree of surface-bound NPs that may interfere with the analysis of cellular uptake, corresponding experiments were performed under cold conditions (4 °C). To estimate the influence of different endocytosis inhibitors on the NP cellular uptake, RAW 264.7 and THP1 cells were preincubated for 1 h under standard culture conditions with the following inhibitors: chlorpromazine (CPZ 8 μg/mL), filipin III (1 μg/mL), or amiloride (50 μM). The final inhibitor concentrations that were not toxic to the cells were evaluated using an MTT assay (data not shown). After preincubation, NPs were added to the culture medium containing inhibitors. Their uptake was analyzed using a BD FACSCalibur flow cytometer and CellQuestPro software. Ten thousand events were acquired for each sample. The results for cells incubated with 0.015 M NaCl as well as nonfluorescent NPs indicated the background fluorescence (autofluorescence of the cells). The obtained results represent the average from three independent experiments. Confocal Microscopy Imaging. Confocal microscopy was used to analyze the cellular localization and uptake of the fluorescently labeled NPs in RAW 264.7 and THP-1 cells. Images were acquired using a Leica LSC SP5 laser scanning confocal microscope (Leica) equipped with a 63× HCX PL APO NA 1.4 oil immersion lens (Leica). Data were acquired in a sequential mode. FITC excitation was performed at 488 nm (Ar laser), and emission was measured at 500−550 nm; sulforhodamine B and LysoTracker Red excitation was performed at 561 nm (DPSS laser), and emission was measured at 580−650 nm. Cells were incubated with 60 μL of NPs under standard culture conditions. For endocytosis pathway studies, preincubation with inhibitors was performed, as described for the flow cytometry assay. Cells were washed with warm PBS to remove unincorporated NPs. Prior to the microscopic examination, aqueous sulforhodamine B solution was added to the cell culture at a final concentration of 1.73 μM. Sulforhodamine B does not enter into cells if the cell membrane is intact, thus enabling visualization of the cell exterior.26 To visualize lysosomes, cells were incubated for 30 min with LysoTracker Red (50 nM) (fluorescent acidotropic probes for labeling acidic organelles in live cells) added to the culture medium. The images of living cells were registered at 37 °C in F12 medium supplemented with 2% FBS. Morphological Studies of THP-1 Cell Differentiation. Cells cultured (standard condition) in a 12-well plate were incubated with various types of synthesized NPs (180 μL per well, ca. 0.15 × 106 NPs/cell) for 3 days. Confocal microscopy was used to acquire images that showed the morphology of the cells. Acquisition was performed at 37 °C. Cells stimulated with PMA were used as a positive control. Statistical Analysis. Data are presented as the mean ± standard error (SEM). The statistical significance was evaluated using the Mann− Whitney U-test. *p < 0.05, **p < 0.01.

instrument from NanoSight. All of the measurements were performed in a 0.015 M NaCl solution at 25 °C. Stability Studies (Colloidal and Biogolical). To evaluate the colloidal and biological stability of the NPs, the size distribution (hydrodynamic diameter) and zeta potential of the NPs were monitored with time. Freshly prepared suspensions of the NPs were stored in 0.015 M NaCl and FBS solutions. The aggregation process was also monitored during cell culturing. Cell Cullture. RAW 264.7 cells were grown in DMEM supplemented with 1% L-glutamine, high glucose, and 10% heat-inactivated fetal bovine serum (FBS). THP-1 cells were grown in RPMI medium supplemented with 10% FBS. Both of the cell lines were cultured at 37 °C in a humidified incubator with a 5% CO2 atmosphere. Two days before the experiment, cells were seeded into the appropriate 96-well plates at a density of 3 × 104 cells per well for the MTT test and 1 × 104 cells per well for the LDH test. Cells were seeded into 6-well plates at a density of 2 × 105 cells per well for flow cytometry measurements or on 15 mm diameter glass coverslips placed in 40 mm plates at a density of 2 × 105 cells for confocal microscopy imaging. For the Vybrant Phagocytosis Assay, RAW 264.7 cells were seeded at a density of 1 × 105 per well (96well plate) 2 h before the experiment. In the case of THP-1 cells stimulation, for the Vybrant Phagocytosis Assay (0.5 × 105 cells per well, 96-well plate) and differentiation studies (4 × 105 cells per well, 12-well plate), cells were cultured in a medium containing 100 ng/mL PMA 3 days before the experiment. Cell Viability and Cytotoxicity Assays. MTT Reduction Test. The evaluation of cell viability (RAW 264.7 and THP-1) was performed using an MTT reduction test. NPs with various numbers of polyelectrolyte layers and surface modifications were resuspended in 0.015 M NaCl, added in variable doses to the fresh medium, and transferred into each well. Doses of 100, 50, and 20 μL of NPs per well were used, which was equivalent to approximately 0.85 × 106, 0.4 × 106, and 0.15 × 106 NPs per cell, respectively. After a 24 h treatment, the medium with the NPs was removed, and the cells were incubated with 0.5 mg/mL MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) resuspended in a serum-free medium for 4 h at 37 °C in a 5% CO2 atmosphere. After incubation, the medium was removed, and 100 μL of DMSO was added to each well. Yellow tetrazolium salt was converted by viable cells to purple formazan, which was dissolved in dimethyl sulfoxide. The absorbance of a product was detected at 570 nm using a microplate reader (TECAN Infinitive M200 Pro). Untreated cells as well as cells incubated with 0.015 M NaCl served as a control. Six replicates for each experimental condition were performed. The obtained results represent the average cell viability from three independent experiments. For the THP-1 cells, the tests were performed with and without PMA stimulation. LDH Cytotoxicity Detection Kit. A cytotoxicity assay was performed in RAW 264.7 cells after 4 h of incubation with various types and doses of NPs (analogous to the MTT assay). LDH (lactate dehydrogenase) released into the cell culture medium correlates with cell membrane damage, which was determined using a Cytotoxicity Detection Kit (LDH) according to the manufacturer’s protocol. Plates were centrifuged at 250 g for 10 min, and 100 μL of culture supernatant was removed and incubated with 100 μL of the reaction mixture for 30 min in the dark at room temperature. The LDH level was determined using a colorimetric assay. The absorbance was measured at 490 nm (the reference wavelength was 610 nm) (TECAN Infinitive200). The two following controls were performed: low control, which measured the activity of spontaneous LDH release from untreated cells and cells incubated with 0.015 M NaCl, and high control, which indicated the maximum LDH level released in response to Triton X-100, causing cell death. Six replicates for each of the experimental conditions were performed. The results represent the cytotoxic effect from three independent experiments. Vybrant Phagocytosis Assay. A phagocytosis assay was performed using a Vybrant Phagocytosis Assay Kit according to the manufacturer’s instructions. Because THP-1 cells are not adherent, initially they were activated with PMA. Briefly, cells (RAW 264.7 and THP-1) cultured in 96-well microplates were incubated for 2 h under standard culture conditions in the presence of various types of synthesized NPs at a dose

3. RESULTS AND DISCUSSION Preparation and Characterization of Surface-Modified Polymeric NPs. Unmodified NPs were obtained as described 6417

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir previously1,20 (Figure 1). Briefly, for the preparation of the nanoemulsion as particle cores, 0.1 mL of the oil phase (AOT in

stabilization. Tails should reduce or even eliminate the electric or hydrophobic unspecific interactions of the NPs with plasma proteins. The final concentration of the obtained NPs (all types) measured by the NS500 NanoSight instrument was ∼1 × 1012 NPs/mL. Cell Viability Analysis. Quantifying cell viability is crucial for understanding the interaction between NPs and the targeted cell. Studies indicate the changes in NP toxicity are dependent on the surface charge of NPs. It is known that positively charged particles are more toxic than negatively charged particles.27 Therefore, the viability of cells exposed to various types of obtained NPs was determined. Using the MTT test, the cellular metabolic activity was measured. The presented results (Figure 2) indicate a dependence of the macrophage cell viability on NP dose, number of polyelectrolyte layers in the capsule shell, and surface charge. The observed effect is concentration-dependent. Negatively charged NPs, consisting of a higher number of polyelectrolyte layers, exhibit a less unfavorable influence on cell viability. Another good marker present in all metabolically active cells is ATP, a key energy metabolite. Its concentration declines rapidly during apoptosis or necrosis. Therefore, to assess cell viability, we also performed tests based on ATP measurements (ATPlite 1step assay). The obtained results were consistent with the MTT test (data not shown). The mechanism of the cytotoxic effect is probably similar for both of the investigated cell lines and may be related to the more efficient adsorption of positively charged nanoparticles on the cell membrane, the tendency toward the reduction of lipid density, and, eventually, the plasma membrane disruptions. Therefore, to evaluate the effects of the NPs on cell membrane integrity, measurements of the LDH (stable cytoplasmic enzyme) release into the cell culture medium were performed. The obtained results indicate a correlation between the NP surface charge and cell membrane destabilization. Positively charged NPs exhibit a higher cytotoxic effect (Figure 2). Parallel trends were described and explained in our previous work20 for HEK 293 cells. Additional cell viability experiments using modified NPs with six layers of the PEGgrafted shell were conducted. The results indicated that pegylation, independent of the grafting ratio, did not cause toxicity because we did not observe any LDH release or reduction of cell metabolic activity. A summary of the experiments is presented in Figure 2. On the basis of the results described above, we decided to focus on studies of NPs consisting of five and six polyelectrolyte layers in doses that do not affect cell vitality. Vybrant Phagocytosis Assay. One of the major properties of macrophages is their phagocytic potential. Therefore, we studied whether the synthesized PLL−PGA NPs can modulate macrophage phagocytosis. As shown in Figure 3, phagocytosis was significantly up-regulated by unmodified NPs, especially positively charged NPs in the case of THP-1 cells. This result suggests that the protein adsorption phenomenon on the NP surface plays a role in phagocytosis. As expected, the phagocytic activity of pegylated NPs remains at the same level, independent of the grafting ratio. The PEG attachment neutralizes the effect resulting from the charged component of the particle. Analysis of in Vitro Uptake by Macrophages. To verify the hypothesis that synthesized PLL−PGA NPs are suitable candidates for drug delivery carriers, the interaction with phagocytic cells was investigated. Two methodologies (flow cytometry and confocal microscopy) were adopted to study cellular uptake. To evaluate the level of NP internalization, RAW

Figure 1. Schematic illustration of the synthesized NP. The dark green lines represent the positively charged PLL, the red line represents the negatively charged PGA, and the blue and green lines represent PEG chains.

chloroform) was added to the aqueous PLL solution (c = 0.1 g/ dm3) under continuous mixing. The optimal ratio of surfactant (AOT) to polycation PLL was determined by measuring the zeta potential of the emulsion droplets and examining their stability. The optimal ratio AOT/PLL was found when the zeta potential of the emulsion droplets reached a value close to the zeta potential of the PLL in solution immediately after overcharging. Using this approach, the amount of unadsorbed polycation PLL was minimized because most of it was consumed to form the AOT/PLL interfacial complex. The average drop size measured by DLS was approximately 80 nm (PDI < 0.2). Emulsions were stable up to 1 year because we did not observe any significant changes in their size and zeta potential. Polyelectrolyte multilayer shells were constructed on the prepared nanoemulsion droplet using the layer-by-layer method (the saturation technique). A fixed volume of nanoemulsion was added to the PGA solution while mixing with a magnetic stirrer. The volume of PGA solution used to form the saturated anionic layer was chosen empirically by analyzing simultaneous zeta potential measurements. For PLL, the optimal volume of the PGA solution was found when the zeta potential of the NPs reached a value close to the zeta potential of the PGA in solution immediately after overcharging. The procedure of sequential deposition of PLL and PGA layers described above was repeated until an appropriate number of polyelectrolyte layers in a shell was formed. For preparation of pegylated NPs, PLL-terminated NPs with five polyelectrolyte layers were additionally coated with synthesized pegylated polyelectrolytes (PGA-g(14)-PEG, PGAg(39)-PEG, and PGA-g(61)-PEG) using the same procedure as that for the normal polyelectrolyte layer. The optimal volumes of PGA-g-PEGs used to form the stable layers were achieved when the zeta potential of NPs reached a value similar to pegylated PGA in solution, ca. 0. Using this method, we prepared pegylated NPs with various densities of PEG chains at their surface. Figure 2A illustrates a typical saw-like dependence of the NP zeta potential on the adsorption of subsequent layers. These results can be considered as evidence for the formation of consecutive layers of particle shells. The average sizes of the prepared NPs are shown in Table 1. To determine whether the NPs would aggregate in biosystems, the stability of pegylated NPs in fetal bovine serum (FBS) was tested. We found that all types of pegylated NPs retained their size without significant changes for at least 48 h. Moreover, we did not observe any aggregation process of pegylated NPs in the coculture system, indicating that PEG tails at the particle surface provide sufficient steric 6418

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir

Figure 2. Biocompatibility studies performed in RAW 264.7 and THP-1 cells after incubation with different types (various numbers of NP layers) and doses of NPs. (A) Nanoparticle preparation - dependence of NP zeta potential on the adsorption of subsequent polyelectrolyte layers. (B, C) MTT tests after a 24 h incubation with NPs. (D) MTT tests after 24 h of incubation with PEG-grafted (various PEG ratios) NPs at dose 3. (E) Cytotoxicity assay LDH release, after 4 h of incubation with NPs. Dose 3 did not cause any measurable LDH release. Dose 1 - 0.85 × 106 NPs/cell, dose 2 - 0.45 × 106 NPs/ cell, and dose 3 - 0.15 × 106 NPs/cell.

Table 1. Name, Abbreviation, and Average Size of Synthesized NPs Just after Preparation, after 1 Year of Storage in 0.015 M NaCl NPs

abbreviation

size (nm)

size (after 1 year) (nm)

AOT/PLL AOT(PLL/PGA)2,5 AOT(PLL/PGA)3 AOT(PLL/PGA)3-g(14)-PEG AOT(PLL/PGA)3-g(39)-PEG AOT(PLL/PGA)3-g(61)-PEG

NP I-PLL NP V-PLL NP VI-PGA NPs VI PGA-g(14)-PEG NPs VI PGA-g(39)-PEG NPs VI PGA-g(61)-PEG

60 ± 4 68 ± 6 75 ± 3 85 ± 5 85 ± 7 85 ± 6

64 ± 5 72 ± 4 77 ± 7 84 ± 4 88 ± 6 90 ± 5

affect the internalization process.28−34 Therefore, in this study, experiments were conducted using differently charged NPs. The NP dose used in the uptake experiments was safe and did not cause membrane damage. In Figure 2E, the cytotoxicity data

264.7 and THP-1 cells were incubated with various types of fluorescently labeled PLL−PGA NPs. It is well established that physicochemical properties of NPs, such as size, shape, solubility, surface charge, and architecture, 6419

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir

Figure 3. Vybrant phagocytosis assaythe influence of NPs (various types) on the phagocytic potential of (A) RAW 264.7 and (B) THP-1 cells.

Figure 4. Flow cytometry analysis of NP (various types) internalization: (A) RAW 264.7 cells - 0.5 h incubation with NPs, (B) RAW 264.7 cells - 2 h incubation with NPs, and (C) THP-1 cells - 0.5 h incubation with NPs. To establish the mechanism of NP uptake, cells were treated with specific inhibitors of endocytosis (CPZ, filipin III, or amiloride) prior to incubation with particles. At low temperatures (4 °C), the process was inhibited.

design of effective drug delivery systems requires addressing the following challenges: avoiding opsonin adsorption, increasing circulation time, and enabling NPs to reach the desired site of action. Thus, for in vivo medical applications, nonspecific NP binding to patients’ blood elements should be minimized. A higher protein adsorbability of a hydrophobic surface compared with a hydrophilic surface has been shown to enhance the uptake of more hydrophobic particles by phagocytes in vitro and rapid removal of hydrophobic particles in vivo.35 A widely used technique for masking NPs relies on adsorbing or grafting shielding groups to hinder the hydrophobic and electrostatic interactions between plasma proteins that bind the particle surface.35−38 Therefore, to minimize nonspecific binding, the selected NPs consisting of six polyelectrolyte layers were coated with a hydrophilic, flexible, and nonionic polymer, such as PEG. Strongly hydrated PEG chains mask the original surface and

confirm that, for the chosen dose, possible membrane damage caused by positively charged NPs did not occur. The observed results did not indicate any spectacular changes in the level of internalization when we compared positively and negatively charged NPs (Figure 4). The situation was similar for the RAW 264.7 and THP-1 cell lines. The slightly accelerated uptake in the case of THP-1 cells may result from better accessibility of NPs to the cells, which were not adherent, and the tests were performed in the cell suspension. Thus, the contact surface volume was higher in the case of THP-1 cells than in RAW 264.7 cells. Cho et al. (2011) demonstrated that, in the absence of particle aggregation, the cellular uptake of NPs depends on the ratio of sedimentation to diffusion velocities,12 which concurs with our findings. As mentioned above, opsonization is the key process by which NPs can be recognized and cleared by phagocytic cells. The 6420

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir

Figure 5. Confocal microscopy analysis of internalization of NPs (various types) in RAW 264.7 cells. To establish the mechanism of the NP uptake, cells were treated with specific inhibitors of endocytosis prior to incubation with particles. White bars indicate 25 μm. Green - NPs, red - sulforhodamine B. In panel A, the X−Y section is shown together with the corresponding X−Z (bottom) and Y−Z (right) sections. (A, B) Internalization of NPs without inhibitor. (C) Internalization of NPs preceded by incubation with CPZ. Filipin III and amiloride did not cause any significant changes in the internalization (data not shown). The images are representative of three independent experiments.

Figure 6. Confocal microscopy analysis of internalization in THP-1 cells. To establish the mechanism of the NP uptake, cells were treated with specific inhibitors of endocytosis prior to incubation with particles. White bars indicate 25 μm. Green - NPs, red - sulforhodamine B. (A) Internalization of nanoparticles without inhibitor. (B) Internalization of NPs preceded by incubation with CPZ. Filipin III and amiloride did not cause any significant changes in the internalization (data not shown). The images are representative of three independent experiments.

remarkably differed compared with unmodified NPs (Figure 4). The observed effect was similar for all of the pegylated NPs with various grafting ratios, suggesting that the addition of PEG chains to the surface area of a particle modifies its properties, making it more “invisible” to phagocytic cells. Moreover, the adsorption of PGA-g(14)-PEG (the lowest PEG density) on the NP surface is sufficient for its proper modification. Parallel trends were observed in the RAW 264.7 and THP-1 cell lines. Moreover, the NP uptake was inhibited at low (4 °C) temperatures (data

provide steric hindrance, which plays an important role in preventing protein adsorption and cell membrane disruption. Due to the presence of PEG chains at the particle surface, the zeta potential decreased to a value close to zero (Figure 2A), reducing or even eliminating the electric, unspecific interactions of NPs while simultaneously preventing the aggregation of NPs. This effect is either due to the ionic strength of the physiological media or due to chemical reactions with components of the cell media and protein adsorption. Internalization of PEG-modified NPs 6421

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir

Figure 7. Intracellular localization of NPs (various types) after a 2 h incubation period. Green - fluorescently labeled NPs, red - lysosomes stained with LysoTracker Red. (A) Experiments performed in RAW 264.7 cells. (B) Experiments performed in THP-1 cells. Co-localization of NPs and lysosomes appearing in yellow pixels indicates that NPs are present in the lysosomes. White bars indicate 10 μm. The images are representative of three independent experiments.

not shown), as indicated by the absence of any fluorescent NPs inside the cells. Additionally, this result suggests the contribution of endocytosis in the process. Figures 5 and 6 show representative images of cells preincubated with NPs, revealing a qualitative illustration for the flow cytometry experiments. Effects of Different Endocytosis Inhibitors on NP Uptake. Recently published data have shown that multiple pathways can be involved in endocytosis of NPs.39 Therefore, in the present study, we also focused on the identification of the endocytosis pathways involved in the internalization of polymeric PLL−PGA NPs. To establish the mechanism of NP uptake, the RAW 264.7 and THP-1 cell lines were treated with specific inhibitors of endocytosis prior to incubation with the particles. The obtained results are similar for both of the investigated cell lines. Moreover, similar trends were observed for each NP type. In phagocytic cells, three major endocytosis pathways occur, clathrin-mediated endocytosis, caveolae-mediated endocytosis, and macropinocytosis.40 Clathrin-mediated endocytosis is a type of receptor-dependent, GTPase- and dynamin-required endocytosis.41 Chlorpromazine (CPZ) is a cationic amphiphilic agent, which perturbs clathrin-coated pit formation.42 For all of the used NPs (different charge and surface modifications), the strongest inhibition of the internalization process was found by blocking the clathrin-mediated endocytosis pathway with CPZ. However, the uptake of unmodified NPs decreased greatly compared with PEG-grafted NPs (Figure 4). The slight difference observed after 30 min of incubation in the case of pegylated NPs may be due to passive membrane penetration; however, this phenomenon typically occurs for NPs much smaller than 100 nm.43 Another type of endocytic pathway that occurs in macrophages is mediated through caveolae and is a cholesterol- and dynamin-dependent, receptor-mediated pathway.44,45 In the present study, preincubation of RAW 264.7 and THP-1 cells with filipin III (an inhibitor that binds to cholesterol and distorts the structure and functions of cholesterol-rich membrane domains40) did not cause any significant variations in the endocytosis level. It has been previously described40 that caveolae-dependent particle internalization occurs when the

nanoparticle size ranges from 200 to 500 nm or when smaller particles aggregate; this most likely explains the results described above (the average size of our NPs was ca. 80 nm). Moreover, NP internalization may also be inhibited by amiloride, which influences macropinocytosis. This type of endocytosis is distinct from clathrin- and caveolae-mediated endocytosis because there are no apparent coat structures.46 Amiloride activity is related to the blockade of the Na+/H+ exchanger and the prevention of membrane ruffling.40,47 The experiments performed show a slight decrease in the internalization level following cell incubation with amiloride. In conclusion, we postulate that clathrin-mediated endocytosis is the major contributor to the uptake of synthesized polymeric PLL−PGA NPs in both of the investigated phagocytic cell lines. We did not record any difference in the uptake level for different pegylated NPs. A summary of the corresponding experiments performed using confocal microscopy is shown in Figures 5 and 6. As observed, all of the used endocytosis inhibitors could decrease the cellular uptake of the investigated NPs to some extent, with CPZ having the strongest effect; however, the measurement was not quantitative. Intracellular Localization of NPs after Macrophage Internalization. Confocal microscopy was used to determine the localization of PLL−PGA NPs inside of RAW 264.7 and THP-1 cells. The colocalization studies were performed with the contribution of the fluorescently labeled NPs and fluorescent probes specific for lysosomes. All of the investigated NPs were found within lysosome compartments. In Figure 7, the colocalization images are shown. The yellow pixels, which result from overlaying the red fluorescent vesicles and green fluorescent NPs, suggest that NPs were trapped inside of the lysosomes following internalization. Morphological Studies of THP-1 Cell Differentiation. Monocytes are highly plastic and heterogeneous, and they change their functional phenotype in response to environmental stimulation.48 A model system for monocyte maturation is the differentiation of monocytic cell lines.49 During infection, circulating blood monocytes differentiate into macrophages. This maturation process prepares the cell to actively participate 6422

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Langmuir

■

in the inflammatory and immune response.50,51 In addition to the morphological changes, functional changes occur, such as the loss of proliferation ability and antigenic phenotype changes. Therefore, in this study, we determined whether synthesized NPs stimulate THP-1 cell differentiation. The effect on THP-1 cell morphology caused by PMA, a macrophage-like differentiationinducing compound, was compared with NPs. The cells were incubated under the same conditions as those used with PMA and NPs. Figure 8 presents a summary of the obtained results.

Article

ASSOCIATED CONTENT

S Supporting Information *

The size distribution (the hydrodynamic diameter) of the NPs was determined using a Zetasizer Nano Series from Malvern Instruments by dynamic light scattering (DLS). The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.langmuir.5b01226.

■

AUTHOR INFORMATION

Corresponding Author

*Fax: (+48) 012 664 69 02. Phone: (+48) 012 664 61 34. E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS The authors wish to thank Prof P. Warszynski for his assistance and Aud. M. Bouzga for NMR characterization of pegylated polyelectrolytes and Dr K. Stalinska and Dr M. Bzowska for providing us with the THP-1 cell line. This work was supported by grants from the Ministry of Science and Higher Education JUVENTUS IP2011031571 and N N401 009640 project. The Faculty of Biochemistry, Biophysics and Biotechnology is a partner of the Leading National Research Center (KNOW) supported by the Ministry of Science and Higher Education.

■

REFERENCES

(1) Antipov, A. A.; Sukhorukov, G. B. Polyelectrolyte Multilayer Capsules as Vehicles with Tunable Permeability. Adv. Colloid Interface Sci. 2004, 111, 49−61. (2) Ariga, K.; Lvov, Y. M.; Kawakami, K.; Ji, Q.; Hill, J. P. Layer-byLayer Self-Assembled Shells for Drug Delivery. Adv. Drug Delivery Rev. 2011, 63, 762−771. (3) Borodina, T. N.; Rumsh, L. D.; Kunizhev, S. M.; Sukhorukov, G. B.; Vorozhtsov, G. N.; Feldman, B. M.; et al. Polyelectrolyte Microcapsules as Systems for Delivery of Biologically Active Substances. Biomed. Khim. 2007, 53, 557−565. (4) Bhaskar, S.; Tian, F.; Stoeger, T.; Kreyling, W.; de la Fuente, J. M.; Grazú, V.; Borm, P.; Estrada, G.; Ntziachristos, V.; Razansky, D. Multifunctional Nanocarriers for Diagnostics, Drug Delivery and Targeted Treatment Across Blood-Brain Barrier: Perspectives on Tracking and Neuroimaging. Part. Fibre Toxicol. 2010, 7, 1−25. (5) Roy, R.; Kumar, S.; Tripathi, A.; Das, M.; Dwivedi, P. D. Interactive Threats of Nanoparticles to the Biological System. Immunol. Lett. 2014, 158, 79−87. (6) Dwivedi, P. D.; Misra, A.; Shanker, R.; Das, M. Are Nanomaterials a Threat to the Immune System? Nanotoxicology 2009, 3, 19−26. (7) Zhang, Y.; Kohler, N.; Zhang, M. Surface Modification of Superparamagnetic Magnetite Nanoparticles and their Intracellular Uptake. Biomaterials 2002, 23, 1553−1561. (8) Owens, D. E., III; Peppas, N. A. Opsonization, Biodistribution, and Pharmacokinetics of Polymeric Nanoparticles. Int. J. Pharm. 2006, 307, 93−102. (9) Soto, E. R.; Caras, A. C.; Kut, L. C.; Castle, M. K.; Ostroff, G. R. Glucan Particles for Macrophage Targeted Delivery of Nanoparticles. J. Drug Delivery 2012, 2012, 143524. (10) Vercauteren, D.; Rejman, J.; Martens, T. F.; Demeester, J.; De Smedt, S. C.; Braeckmans, K. On the Cellular Processing of Non-Viral Nanomedicines for Nucleic Acid Delivery: Mechanisms and Methods. J. Controlled Release 2012, 161, 566−581. (11) Sharma, G.; Valenta, D. T.; Altman, Y.; Harvey, S.; Xie, H.; Mitragotri, S.; Smith, J. W. Polymer Particle Shape Independently Influences Binding and Internalization by Macrophages. J. Controlled Release 2010, 147, 408−412.

Figure 8. Morphological studies of THP-1 cell differentiation.

We observed a different morphology of the cells depending on the stimulating agent. The PMA-treated cells exhibit an adherent growth phenotype and a loss of proliferation. The investigated PLL−PGA NPs did not cause THP-1 differentiation in PEGgrafted or unmodified particles.

4. CONCLUSIONS In the present study, we examined the effects of various polyelectrolyte PLL−PGA NPs (different charge, shell thickness, shell modification) obtained using the LbL method on their interactions with phagocytic cells (RAW 264.7 and THP-1 cell lines). The obtained results indicate that the negatively charged unmodified NPs consisting of higher numbers of polyelectrolyte layers exhibit a lower unfavorable effect on cell viability. Negatively and positively charged NPs as well as pegylated NPs do not lead to differentiation of THP-1 cells. In contrast to pegylated NPs, unmodified (especially positively charged) NPs modulate macrophage phagocytosis. Unmodified NPs are preferentially internalized by both of the investigated cell lines compared with PEG-grafted NPs. The strongest uptake inhibition was evaluated by blocking the clathrin-mediated endocytosis pathway. After cellular uptake, all types of investigated NPs were found to localize in the lysosomes. On the basis of our observations, we can conclude that the obtained polymeric NPs can be successfully modified (by PEG-grafting) in a way that enables them to be invisible to phagocytic cells, confirming the previously postulated theory20 that PLL−PGA NPs are promising candidates for drug delivery. 6423

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir

Nanoparticles Influence their Biodistribution and Uptake by Macrophages. Eur. J. Pharm. Biopharm. 2011, 77, 417−423. (33) Alexis, F.; Pridgen, E.; Molnar, L. K.; Farokhzad, O. C. Factors Affecting the Clearance and Biodistribution of Polymeric Nanoparticles. Mol. Pharmaceutics 2008, 5, 505−515. (34) He, C.; Hu, Y.; Yin, L.; Tang, C.; Yin, C. Effects of Particle Size and Surface Charge on Cellular Uptake and Biodistribution of Polymeric Nanoparticles. Biomaterials 2010, 31, 3657−3666. (35) Li, F.; Zhu, A.; Song, X.; Ji, L.; Wang, J. The Internalization of Fluorescence-Labeled PLA Nanoparticles by Macrophages. Int. J. Pharm. 2013, 453, 506−513. (36) Liu, Y.; Hu, Y.; Huang, L. Influence of Polyethylene Glycol Density and Surface Lipid on Pharmacokinetics and Biodistribution of Lipid-Calcium-Phosphate Nanoparticles. Biomaterials 2014, 35, 3027− 3034. (37) Pignatello, R.; Leonardi, A.; Pellitteri, R.; Carbone, C.; Caggia, S.; Graziano, A. C. E.; Cardile, V. Evaluation of New Amphiphilic PEG Derivatives for Preparing Stealth Lipid Nanoparticles. Colloids Surf., A 2013, 434, 136−144. (38) Semete, B.; Booysen, L.; Kalombo, L.; Ramalapa, B.; Hayeshi, R.; Swai, H. S. Effects of Protein Binding on the Biodistribution of PEGylated PLGA Nanoparticles Post Oral Administration. Int. J. Pharm. 2012, 424, 115−120. (39) Kastl, L.; Sasse, D.; Wulf, V.; Hartmann, R.; Mircheski, J.; Ranke, C.; Carregal-Romero, S.; Martinez-Lopez, J. A.; Fernandez-Chacon, R.; Parak, W. J.; Elsasser, H. P.; Rivera Gil, P. Multiple Internalization Pathways of Polyelectrolyte Multilayer Capsules into Mammalian Cells. ACS Nano 2013, 7, 6605−6618. (40) Roy, R.; Parashar, V.; Chauhan, L. K. S.; Shanker, R.; Das, M.; Tripathi, A.; Dwivedi, P. D. Mechanism of Uptake of ZnO Nanoparticles and Inflammatory Responses in Macrophages Require PI3K Mediated MAPKs Signaling. Toxicol. In Vitro 2014, 28, 457−467. (41) Rappoport, J. Z. Focusing on Clathrin-Mediated Endocytosis. Biochem. J. 2008, 412, 415−423. (42) Wang, L. H.; Rothberg, K. G.; Anderson, R. G. Mis-Assembly of Clathrin Lattices on Endosomes Reveals a Regulatory Switch for Coated Pit Formation. J. Cell Biol. 1993, 123, 1107−1117. (43) Nel, A. E.; Madler, L.; Velegol, D.; Xia, T.; Hoek, E. M.; Somasundaran, P.; Klaessig, F.; Castranova, V.; Thompson, M. Understanding Biophysicochemical Interactions at the Nano-Bio Interface. Nat. Mater. 2009, 8, 543−557. (44) Aa, M. A. E. M.; Huth, U. S.; HÃ ¤fele, S. Y.; Schubert, R.; Oosting, R. S.; Mastrobattista, E.; Hennink, W. E.; Peschka-SÃ 1/4ss, R.; Koning, G. A.; Crommelin, D. J. A. Cellular Uptake of Cationic Polymer-DNA Complexes Via Caveolae Plays a Pivotal Role in Gene Transfection in COS-7 Cells. Pharm. Res. 2007, 24, 1590−1598. (45) Nichols, B. Caveosomes and Endocytosis of Lipid Rafts. J. Cell Sci. 2003, 116, 4707−4714. (46) Sarkar, K.; Kruhlak, M. J.; Erlandsen, S. L.; Shaw, S. Selective Inhibition by Rottlerin of Macropinocytosis in Monocyte-Derived Dendritic Cells. Immunology 2005, 116, 513−524. (47) Sasahara, T.; Yayama, K.; Tahara, T.; Onoe, H.; Okamoto, H. Na +/H+ Exchanger Inhibitor Augments Hyperosmolarity-Induced Vasoconstriction by Enhancing Actin Polymerization. Vasc. Pharmacol. 2013, 59, 120−126. (48) Yang, J.; Zhang, L.; Yu, C.; Yang, X.; Wang, H. Monocyte and Macrophage Differentiation: Circulation Inflammatory Monocyte as Biomarker for Inflammatory Diseases. Biomarker Res. 2014, 2, 1−9. (49) Kreutz, M.; Andreesen, R. Induction of Human Monocyte to Macrophage Maturation in Vitro by 1,25-Dihydroxyvitamin D3. Blood 1990, 76, 2457−2461. (50) Takashiba, S.; Van Dyke, T. E.; Amar, S.; Murayama, Y.; Soskolne, A. W.; Shapira, L. Differentiation of Monocytes to Macrophages Primes Cells for Lipopolysaccharide Stimulation Via Accumulation of Cytoplasmic Nuclear Factor kappaB. Infect. Immun. 1999, 67, 5573− 5578. (51) Zhou, J.; Zhu, P.; Jiang, J. L.; Zhang, Q.; Wu, Z. B.; Yao, X. Y.; Tang, H.; Lu, N.; Yang, Y.; Chen, Z. N. Involvement of CD147 in

(12) Cho, C.; Zhang, Q.; Xia, Y. The Effect of Sedimentation and Diffusion on Cellular Uptake of Gold Nanoparticles. Nat. Nanotechnol. 2011, 6, 385−391. (13) Rigotti, A.; Acton, L.; Krieger, M. the Class-B Scavenger Receptors Sr-Bi and Cd36 are Receptors for Anionic Phospholipids. J. Biol. Chem. 1995, 270, 16221−16224. (14) Wilhelm, C.; Billotey, C.; Roger, J.; Pons, J. N.; Bacri, J. -.; Gazeau, F. Intracellular Uptake of Anionic Superparamagnetic Nanoparticles as a Function of their Surface Coating. Biomaterials 2003, 24, 1001−1011. (15) Nicolete, R.; Santos, D. F. d.; Faccioli, L. H. The Uptake of PLGA Micro Or Nanoparticles by Macrophages Provokes Distinct in Vitro Inflammatory Response. Int. Immunopharmacol. 2011, 11, 1557−1563. (16) Tomita, Y.; Rikimaru-Kaneko, A.; Hashiguchi, K.; Shirotake, S. Effect of Anionic and Cationic n-Butylcyanoacrylate Nanoparticles on NO and Cytokine Production in Raw264.7 Cells. Immunopharmacol. Immunotoxicol. 2011, 33, 730−737. (17) Poon, Z.; Lee, J. B.; Morton, S. W.; Hammond, P. T. Controlling in Vivo Stability and Biodistribution in Electrostatically Assembled Nanoparticles for Systemic Delivery. Nano Lett. 2011, 11, 2096−2103. (18) Morton, S. W.; Poon, Z.; Hammond, P. T. The Architecture and Biological Performance of Drug-Loaded LbL Nanoparticles. Biomaterials 2013, 34, 5328−5335. (19) Zhang, M.; Desai, T.; Ferrari, M. Proteins and Cells on PEG Immobilized Silicon Surfaces. Biomaterials 1998, 19, 953−960. (20) Lukasiewicz, S.; Szczepanowicz, K. In Vitro Interaction of Polyelectrolyte Nanocapsules with Model Cells. Langmuir 2014, 30, 1100−1107. (21) Sukhorukov, G. B.; Donath, E.; Lichtenfeld, H.; Knippel, E.; Knippel, M.; Budde, A.; Möhwald, H. Layer-by-Layer Self Assembly of Polyelectrolytes on Colloidal Particles. Colloids Surf., A 1998, 137, 253− 266. (22) Szczepanowicz, K.; Dronka-Góra, D.; Para, G.; Warszyński, P. Encapsulation of Liquid Cores by Layer-by-Layer Adsorption of Polyelectrolytes. J. Microencapsulation 2010, 27, 198−204. (23) www.Fda.Gov. (24) Boulmedais, F.; Frisch, B.; Etienne, O.; Lavalle, P.; Picart, C.; Ogier, J.; Voegel, J. -.; Schaaf, P.; Egles, C. Polyelectrolyte Multilayer Films with Pegylated Polypeptides as a New Type of Anti-Microbial Protection for Biomaterials. Biomaterials 2004, 25, 2003−2011. (25) Szczepanowicz, K.; Hoel, H. J.; Szyk-Warszynska, L.; Bielanska, E.; Bouzga, A. M.; Gaudernack, G.; Simon, C.; Warszynski, P. Formation of Biocompatible Nanocapsules with Emulsion Core and Pegylated Shell by Polyelectrolyte Multilayer Adsorption. Langmuir 2010, 26, 12592−12597. (26) Chodosh, J.; Dix, R. D.; Howell, R. C.; Stroop, W. G.; Tseng, S. C. Staining Characteristics and Antiviral Activity of Sulforhodamine B and Lissamine Green B. Invest. Ophthalmol. Visual Sci. 1994, 35, 1046−1058. (27) Kolbe, A.; del Mercato, L. L.; Abbasi, A. Z.; Rivera Gil, P.; Gorzini, S. J.; Huibers, W. H.; Poolman, B.; Parak, W. J.; Herrmann, A. De Novo Design of Supercharged, Unfolded Protein Polymers, and their Assembly into Supramolecular Aggregates. Macromol. Rapid Commun. 2011, 32, 186−190. (28) Huhn, D.; Kantner, K.; Geidel, C.; Brandholt, S.; De Cock, I.; Soenen, S. J.; Rivera Gil, P.; Montenegro, J. M.; Braeckmans, K.; Mullen, K.; Nienhaus, G. U.; Klapper, M.; Parak, W. J. Polymer-Coated Nanoparticles Interacting with Proteins and Cells: Focusing on the Sign of the Net Charge. ACS Nano 2013, 7, 3253−3263. (29) Shimoni, O.; Yan, Y.; Wang, Y.; Caruso, F. Shape-Dependent Cellular Processing of Polyelectrolyte Capsules. ACS Nano 2013, 7, 522−530. (30) Xiao, K.; Li, Y.; Luo, J.; Lee, J. S.; Xiao, W.; Gonik, A. M.; Agarwal, R. G.; Lam, K. S. The Effect of Surface Charge on in Vivo Biodistribution of PEG-Oligocholic Acid Based Micellar Nanoparticles. Biomaterials 2011, 32, 3435−3446. (31) Foged, C.; Brodin, B.; Frokjaer, S.; Sundblad, A. Particle Size and Surface Charge Affect Particle Uptake by Human Dendritic Cells in an in Vitro Model. Int. J. Pharm. 2005, 298, 315−322. (32) Arnida; Janát-Amsbury, M. M.; Ray, A.; Peterson, C. M.; Ghandehari, H. Geometry and Surface Characteristics of Gold 6424

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425

Article

Langmuir Overexpression of MMP-2 and MMP-9 and Enhancement of Invasive Potential of PMA-Differentiated THP-1. BMC Cell Biol. 2005, 6, 25.

6425

DOI: 10.1021/acs.langmuir.5b01226 Langmuir 2015, 31, 6415−6425