Preparation of hybrid fluorescent–magnetic nanoparticles for application to cellular imaging by self-assembly

Colloids and Surfaces A: Physicochem. Eng. Aspects 392 (2011) 103–109

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Colloids and Surfaces A: Physicochemical and Engineering Aspects journal homepage: www.elsevier.com/locate/colsurfa

Preparation of hybrid fluorescent–magnetic nanoparticles for application to cellular imaging by self-assembly Xiaoyu Wang, Fang He ∗ , Fu Tang, Ning Ma, Lidong Li ∗ School of Materials Science and Engineering, University of Science and Technology Beijing, Beijing 100083, PR China

a r t i c l e

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Article history: Received 8 June 2011 Received in revised form 15 September 2011 Accepted 30 September 2011 Available online 8 October 2011 Keywords: Self-assembly Nanoparticle Polyelectrolyte Conjugated polymer

a b s t r a c t A unique fluorescent–magnetic hybrid bimodal nanocomposite was prepared by the layer-by-layer selfassembly (LbL) technique fabrication of water-soluble conjugated polymers (CPs) onto the CoFe2 O4 @SiO2 core–shell nanoparticles (NPs). First, magnetic CoFe2 O4 nanoparticles were prepared as the magnetic core and coated with a SiO2 shell to obtain a good dispersion in aqueous solution. Then the polyelectrolytes and cationic conjugated polymer PFV was assembled onto the surface of core–shell nanoparticles by the LbL technique. The prepared nanocomposites were magnetically responsive and fluorescent, simultaneously. Finally, the biomacromolecule heparin sodium (HS) was then assembled on the outer layer of the nanocomposite to provide a cytocompatible surface. The nanocomposites show monodispersity, good fluorescence and good biocompatibility that are useful for efficient cellular imaging. Moreover, the colloidal stability and the cellular uptake ability of the nanocomposition with HS layer were efficiently improved. © 2011 Elsevier B.V. All rights reserved.

1. Introduction Bifunctional and multifunctional nanocomposites have obtained great attention for increased efficiency and versatility in the numerous applications of nanomaterials in complex systems [1]. Among these multimodal nanocomposites, fluorescent–magnetic hybrid nanoparticles, which can provide luminescence property and contrast enhancement in magnetic resonance imaging (MRI) at the same time, offer great potential for applications in complex biotechnology, sensor, and quality inspection [2]. Therefore, several approaches have been taken to develop fluorescent–magnetic nanocomposition (FMN) [3–9]. These approaches can be concluded into two main procedures. One is the core–shell design, where the magnetic core is coated by the inorganic or organic shells [6–8]. In another approach, silica or polymers containing the luminescent molecules were combined the agglomerated large magnetic nanoparticles together and formed a FMN [9]. However, systems with small organic fluorophore are easy to leak toxic components from the compositions, which will hinder their application in biological system, and also suffer wide emission lines and photobleaching problem. Conjugated polymers (CPs) have proven to be attractive materials for various optoelectronic applications including light-emitting

∗ Corresponding authors. Tel.: +86 10 82377202; fax: +86 10 82375712. E-mail addresses: [email protected] (F. He), [email protected] (L. Li). 0927-7757/$ – see front matter © 2011 Elsevier B.V. All rights reserved. doi:10.1016/j.colsurfa.2011.09.040

diodes, field-effect transistors, and photovoltaic devices due to their excellent photo- and electroluminescent properties [10–12]. Recently, CPs have also been developed as a promising class of functional materials in the production of fluorescent nanoparticles [13–28]. The obtained fluorescent nanoparticles have shown normal to high brightness, good photostability, and low cytotoxicity that are ideal for cellular imaging. However, the development of hybrid FMN based on CPs are still at a relatively early stage [25–27]. LbL self-assembly technique, which can direct the formation of thin films of functional polyelectrolytes or biomacromolecule onto nanoparticles substrate, is a versatile and simple for nanoparticles surface modifications [29–32]. Many fluorescent nanoparticles and FMN have been produced based on the simple LbL self-assembly technique [28,33–38]. Recently, we have developed fluorescent nanoparticles via a simple electrostatic self-assembly technique based on cationic water-soluble conjugated polymers and Ag@SiO2 core–shell nanoparticles [28]. These NPs are shown to exhibit monodispersity, high stability, bright fluorescence, and good biocompatibility. Therefore they are successfully used for cellular imaging. In this paper, we prepare a novel fluorescent–magnetic nanocomposite based on small magnetic nanoparticles by a simply and efficient LbL self-assembly of ionic water-soluble conjugated polymer. The nanocomposites also show monodispersity, good fluorescence and good biocompatibility that are especially suitable for efficient cellular imaging. Moreover, by using the LbL technique it is easy to envisage many other applications evolving for multifunctional nanoparticles through simply assembling functionalized

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polyelectrolyte on the particle surface. Upon assembling the cytocompatible heparin sodium can be introduced onto the surface as the outmost layer of the nanocomposite, the colloidal stability and the cellular uptake ability for the nanocomposite are efficiently improved. This system provides a new method for the preparation of the multimodal nanoparticles for the complex biologic systems. 2. Experimental 2.1. Materials Cobalt(II) acetylacetonate, iron(III) acetylacetonate, oleylamine, oleic acid, 1,2-hexadecanediol, benzylether, polyoxyethylene (5)nonylphenyl ether (Igepal co-520), poly(diallyldimethylammonium chloride) (PDDA, Mw = 200 000–350 000), poly(sodium styrenesulfonate) (PSS, Mw = 70 000) were obtained from Sigma–Aldrich. Heparin sodium was obtained from Beijing BIODEE Biotechnology Co., Ltd. Poly[9,9 -bis(6 -(N, N,N-trimethylammonium)-hexyl)fluorene-2,7-ylenevinylene-coalt-1,4-phenylene dibromide] (PFV) was synthesized in our lab, the procedure has been reported in our previous publication [28]. All the chemicals were used as received without any further purification. 2.2. Measurements The morphology of the nanoparticles was recorded by JEM 2100 transmission electron microscope with an accelerating voltage of 120 kV. TEM samples were prepared by dropping the nanoparticles suspension onto carbon-coated copper grids and allowed to dry in air. Stained specimens were prepared by depositing the sample solutions on the grid and staining with phosphotungstic acid before TEM observations. Magnetic measurements were carried out with a vibrating sample magnetometer (Lakeshore, Inc.). The photograph of the nanoparticles aqueous suspensions was taken using a digital color camera under a UV lamp with max = 365 nm. The zeta potential and average hydrodynamic diameters of the nanoparticles were measured using Nano ZS90 (Malvern Instrument Ltd.) in dilute solutions at room temperature. Absorption spectra were collected on a Hitachi U3900 spectrophotometer. Photoluminescence (PL) spectra data were obtained from a Hitachi F-7000 fluorescence spectrometer at room temperature with an excitation at 437 nm. The confocal microscopy of the nanoparticles was recorded by Olympus FV1000-IX81 with 405 nm laser excitation. The cellular images were taken with Olympus 1X71 fluorescence microscopy with a 100 W mercury lamp as the light source. 2.3. Synthesis of CoFe2 O4 NPs The synthesis of CoFe2 O4 NPs was divided into two procedures as the literature. Firstly, CoFe2 O4 NPs (6 ± 1 nm) were fabricated as the seeds using reported procedure [39]. In a typical procedure, iron(III) acetylacetonate (706 mg, 2.0 mmol), cobalt(II) acetylacetonate (257 mg, 1.0 mmol), 1,2-hexadecanediol (2.87 g, 10.0 mmol), oleic acid (2.1 mL, 6.0 mmol), oleylamine (2.2 mL, 6.0 mmol), and phenyl ether (20 mL) were combined and mechanically stirred under a flow of nitrogen. The mixture was heated to 200 ◦ C for 2 h, and then under a flow of nitrogen, heated to reflux (∼300 ◦ C) for 1 h. The dark-brown mixture was cooled to room temperature by removing the heat source. Under ambient condition, methanol (40 mL) was added to the mixture, and a black material was precipitated and separated via centrifugation (8000 rpm, 15 min). The product, CoFe2 O4 NPs, was repeatedly washed three times with methanol and distilled water, finally dried under a vacuum overnight. The resulting dry black power was dissolved in hexane

with 0.1% oleic acid to obtain the CoFe2 O4 NPs hexane solution (15 mg/mL). Then, the prepared CoFe2 O4 NPs seeds were used to grow larger particles. In brief, iron(III) acetylacetonate (706 mg, 2.0 mmol), cobalt(II) acetylacetonate (257 mg, 1.0 mmol), 1,2-hexadecanediol (2.87 g, 10.0 mmol), oleic acid (700 ␮L, 2.0 mmol), oleylamine (730 ␮L, 2.0 mmol), and phenyl ether (20 mL) were mixed and mechanically stirred under a flow of nitrogen. 6 mL of the assynthesized 6 ± 1 nm CoFe2 O4 NPs hexane solution (15 mg/mL) was added to the reaction. The mixture was first heated to 100 ◦ C for 30 min to remove hexane, and then increased to 200 ◦ C for 1 h. Under a blanket of nitrogen, the mixture was further heated to 300 ◦ C for 30 min to reflux. Following the workup procedures described as above, the monodispersed 12 ± 1 nm CoFe2 O4 NPs were obtained. 2.4. Preparation of CoFe2 O4 /SiO2 core/shell NPs (CS NPs) The silica-coated CoFe2 O4 NPs was performed through the formation of reverse microemulsion based on the base-catalyzed hydrolysis of tetraethylorthosilicate (TEOS) [40]. Briefly, the nonionic surfactant Igepal co-520 (1 mL, 2.24 mmol) was dispersed in 16 mL of cyclohexane under sonication. Next, 1 mL of CoFe2 O4 NPs solution (1 mg/mL in cyclohexane) was added, and the mixture was sonicated at room temperature for 30 min to yield a transparent, brown solution of reverse microemulsion. 150 ␮L of 30% ammonium hydroxide was added, the resulting mixture sonicated for 15 min, and then 80 ␮L of TEOS was added. After 30 min, the final mixture was transferred to mechanically stirring at room temperature for 24 h. When methanol was added into the reaction solution, the core/shell NPs were destabilized from the microemulsion and precipitated. The resultant precipitate was washed three times with methanol and distilled water. Finally, the wet powder was dispersed in distilled water. The thickness of SiO2 shell achieves to 8.5 ± 1.5 nm. 2.5. Fabrication of the CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposite by LbL self-assembly approach The adsorption of the first layer of cationic PDDA was performed by adding 100 ␮L of PDDA (1 mg/mL) solution to 1 mL of the CS NPs with occasional shaking. After 20 min, the sample was washed twice by centrifugation in deionized water. Finally, they were redispersed in 1 mL of deionized water. Next, negatively charged PSS was then deposited onto the coated nanoparticles in the same conditions and procedures. The above processes were repeated until two PDDA/PSS bilayers were fabricated. After that, 100 ␮L of PFV (1 mM) solution was added into the (PDDA/PSS)2 -modified CS NPs suspension with negatively charged PSS as the outermost layer. After adsorption for an hour under gentle shaking, the dispersion was centrifuged and purified in the same method. Subsequently, 100 ␮L of heparin sodium (1 mg/mL) solution was mixed into the particle suspension to deposit a layer of heparin sodium. As outlined above for the prime layer, the desired functional coating of CS NPs/(PDDA/PSS)2 /PFV/HS layers was prepared. 2.6. Cellular imaging for nanocomposites 100 ␮L of CS NPs/(PDDA/PSS)2 /PFV nanocomposition and CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposition was added into 1 mL of DMEM medium containing A549 cells in 35 mm × 35 mm plate ([PFV] = 1 × 10−5 M), respectively. After incubation at 37 ◦ C for 10 h, the medium was removed, and the cells were washed twice with phosphate saline buffer solution (PBS, pH 7.4). Then the fluorescence images and phase contrast bright-field images were recorded on fluorescence microscopy (Olympus 1X71), without other

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Fig. 1. TEM micrographs of (a) CoFe2 O4 NPs and (b) CoFe2 O4 @SiO2 NPs, the scale bars in the images are 20 nm; (c) field-dependent magnetization hysteresis loop of CoFe2 O4 nanoparticles.

treatment using a 455/70 nm excitation filter with 600 ms exposure time. 2.7. Cell viability assay by MTT method A549 cells were seeded in 96-well tissue culture plates and maintained overnight in DMEM medium. Cells were then

treated with various concentrations of CS NPs/(PDDA/PSS)2 /PFV and CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposition (0–20 ␮M), respectively. After 24 h incubation at 37 ◦ C, the medium was poured out. Then 100 ␮L of 3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide (MTT, 1 mg/mL in PBS) was added into each well. The MTT medium was removed after 4 h. The cells were lysed by adding 150 ␮L of DMSO, and the absorbance of the

Scheme 1. The procedure of the preparation of the nanocomposites and the chemical structures of the polymeric components.

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Fig. 2. (a) TEM micrographs of CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposition, the TEM sample was stained with phosphotungstic acid; (b) confocal microscopy of CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposition in solution, obtained with 405 nm laser excitation. The scale bars in the images are 10 nm and 10 ␮m, respectively.

purple formazan at 520 nm was monitored by a Spectra MAX 340PC plate reader.

3. Results and discussion 3.1. Synthesis and characterization of core–shell CoFe2 O4 @SiO2 NPs Cobalt ferrite (CoFe2 O4 ) nanoparticles are chosen as the magnetic core in our system for their ease of synthesis, remarkable chemical stability, high magnetocrystalline anisotropy, and moderate saturation magnetization [41]. To avoid magnetic agglomeration of the nanoparticles, the magnetic cores should be designed to superparamagnetic [42]. It is reported that through controlling the size of the CoFe2 O4 nanoparticles less than 16 nm, the magnetism of the particles can be stabilized to superparamagnetic at room temperature [43]. Therefore, in this system CoFe2 O4 nanoparticles with average size 12 ± 1 nm were firstly synthesized using the reported procedures [39]. The nanoparticles remained monodisperse in size with no obvious aggregation from the TEM image (Fig. 1a). The magnetization curve of the nanoparticles showed no hysteresis, indicating the CoFe2 O4 nanoparticles were superparamagnetic (Fig. 1c). Then silica shells were synthesized by a reverse microemulsion approach to modify the NPs for further assembly experiments. Fig. 1b is the typical TEM images of the core shell CoFe2 O4 @SiO2 NPs which shows that almost all of the NPs was uniformly coated and the average thickness of the silica shell was 8.5 ± 1.5 nm.

3.2. Fabrication of CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposite by LbL technique. Scheme 1 shows the assembly layers design and the molecular structures of the polymeric components. The cationic conjugated polymer PFV is chosen as luminescent polymer for its good light stability and good self-assembling ability. Before the deposition of PFV, four layers of flexible polyelectrolytes PDDA and PSS are firstly assembled by the standard LbL technique on the negative silica shell, to enhance the adsorption stability of PFV layer. Then the luminescent PFV is deposited through electrostatic interaction. In a final step, a layer of biomacromolecule heparin sodium is assembled to increase the cellular uptake ability and biocompatibility of the nanocomposite for their potential application in bioimaging. In addition, heparin sodium can also interact with a variety of growth factors with heparin binding domains and gives potential application to label the growth factors. 3.3. Characterization of CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposite. Fig. 2a illustrates the TEM images of CS NPs after deposition of polyelectrolytes. The sample was stained with phosphotungstic acid because the polyelectrolyte layers cannot be easily observed in TEM images without staining. The TEM images showed that a thin polymer shell was fabricated on the surface of the CS NPs, but the thickness cannot be measured. Laser-scanning confocal fluorescence microscopy (LSCM) image in Fig. 2b confirmed that

Fig. 3. (a) Zeta-potential values as a function of polyelectrolyte layer number during the fabrication of the nanocomposites; (b) average hydrodynamic diameters of nanoparticles with different bilayers of polyelectrolytes.

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Fig. 4. (a) Absorption and (b) emission spectra of CS NPs/(PDDA/PSS)2 , CS NPs/(PDDA/PSS)2 /PFV and CS NPs/(PDDA/PSS)2 /PFV/HS nanocompositions. The emission intensity was normalized by the absorbance at 437 nm. The excitation wavelength is 437 nm for emission measurements.

luminescent PFV was adsorbed on the nanocomposite and no obvious aggregation of NPs was observed after assembling. At the same time, the deposition of the each polyelectrolyte layer was demonstrated by zeta potential measurements and dynamic light scattering (DLS). Zeta potential values indicated a clear alternation of surface charge after each layer deposition, ranging from strongly positive to strongly negative (Fig. 3a). The results of DLS in Fig. 3b illustrated that the average hydrodynamic diameters of the nanoparticles were gradually increased with the deposition of polyelectrolytes and no abrupt changes were observed, which indicated that the nanocomposition have little aggregation. The deposition of PFV layer on the surface of the CS NPs was further verified from the absorption and emission spectra. Fig. 4a shows the UV–vis absorption spectra of the CS NPs after the adsorption of polyelectrolytes. PDDA and PSS were found to have no obvious absorption peak in the region from 250 to 600 nm. Therefore, the absorption of CS NPs/(PDDA/PSS)2 nanocomposition was similar with core–shell NPs. After adding 400 ␮L of conjugated polymer PFV solution ([PFV] = 1 × 10−3 M) into 600 ␮L of CS NPs/(PDDA/PSS)2 nanocomposite and then separation, obvious new absorption peaks were observed at 437 nm and 466 nm, which are in accordance with the absorption of PFV in aqueous solution [39]. This indicates that the PFV has adsorbed on the surface of the nanocomposite and deposition percentage of PFV is estimated as ∼16.5% from the absorbance of the PFV. Fig. 4b gives the emission spectra of nanocomposite before and after the assembly of heparin sodium. The integrated emission intensity was normalized by the absorbance at the excitation wavelength of 437 nm. The emission spectra of the nanocomposite gave the emission peak of PFV at 484 nm and the shoulder peak at 512 nm. Both the absorption and emission peaks did not shift compared with PFV in aqueous solution, which illustrated that the PFV were less aggregated on the surface due to the small size of the nanocomposite. The fluorescence quantum yields (QY) of the nanocomposition was 4.5%, measured using rhodamine 6G as the standard. The QY of the nanocomposition was only 10% less than PFV solution in water. Therefore, the self-quenching of PFV was greatly inhibited in our nanocomposite system. The spectra also showed that the deposition of heparin sodium had little effect to the optical properties of the nanocomposite. Fig. 5 shows the images of the self-assembled nanocomposite aqueous solutions before and after exposure to a magnetic field. As shown the image in Fig. 5a, the nanocomposite solution in the cuvette was yellow solution. Upon irradiated by a 365 nm UV light, the solution illustrated brightly green fluorescence of PFV. While the solution was exposed to an external magnetic field, the yellow magnetic nanoparticles were aggregated to the magnetic side of the

cuvette, leaving a clear solution behind (Fig. 5b). Upon the exposure to the UV light, the bright green fluorescence can be observed from the magnetic side of the cuvette, indicating that almost all the luminescence PFV have adsorbed onto the surface of the magnetic nanocomposite and the complex were both magnetically responsive and fluorescent, simultaneously. 3.4. Cellular imaging of CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposite Conjugated polymers nanoparticles, which have high cytocompatibility, have shown good potential application in biological sciences. Therefore, to demonstrate these nanocomposites in application on bioimaging, the bimodal nanoparticles were co-cultured with A549 lung cancer cells to assess their potential as fluorescent probes and examine their cell toxicity. The fluorescence images of A549 cells were monitored after co-culturing with nanocomposites for 24 h at 37 ◦ C and washing twice with PBS buffer (pH 7.4). Fig. 6 shows the fluorescence image and an overlap image of phase contrast and fluorescence image of A549 cells without (Fig. 6a) and with (Fig. 6b and c) treated with CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposite. The nanocomposites were found to be located in the cytoplasm and around the nucleus, via endocytosis to the interior of the cells for a staining pattern. This observation indicated that the nanocomposite had potential application in fluorescent labeling and sensing in cell. More interestingly, in this system we found that the biocompatibility HS layer could efficiently improve the colloidal stability of the nanocomposite and the endocytosis to the interior of the cells. When 100 ␮L of CS NPs/(PDDA/PSS)2 /PFV

Fig. 5. Images of aqueous nanocomposition solutions before and after UV irradiation (a) in the absence and (b) in the presence of an external magnetic field.

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Fig. 6. (a) Fluorescence image of A549 cells without treatment of nanoparticles; (b) Fluorescence image and (c) the overlapped phase contrast and fluorescence image of A549 cells after co-culture with CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposition for 24 h at 37 ◦ C; (d) fluorescence images of A549 cells after co-culturing with CS NPs/(PDDA/PSS)2 /PFV nanocomposition for 24 h at 37 ◦ C. The fluorescence images were recorded with a fluorescence microscope (Olympus 1X71) using a 455/70 nm excitation filter with 600 ms exposure time.

and CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposite solution were added into 1 mL of the culture medium of A549 cells (the final concentration of PFV was 1 × 10−5 M), respectively, the images in Fig. 6c and d illustrated that most of the CS NPs/(PDDA/PSS)2 /PFV nanocomposites were destroyed and cannot be located in the cytoplasm of the cell. It indicates that the CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposites showed a higher uptake ability by than the CS NPs/(PDDA/PSS)2 /PFV due to the deposition of HS layer. The cytotoxicity is an important factor to the cellular labeling and imaging materials. For this reason, the cytotoxicity of the

nanocomposite was studied using the MTT cell-viability assays. The absorbance of MTT at 520 nm is dependent on the degree of activation of the cells. Therefore, cell viability can be calculated by the ratio of absorbance of the cells incubated with nanocomposite to that of the cells incubated with culture medium only. Fig. 7 gives the cell viability curves as a function of the concentrations of CS NPs/(PDDA/PSS)2 /PFV and CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposites. It is shown that the cell viability decreases less than 20% after 24 h incubation of A549 cells when the concentration of adsorbed PFV (in repeated units) less than 2 ␮M. The cell viability is more than 50% when the concentration of PFV up to 20 ␮M. The results indicated that both of the nanocomposites have low cytotoxicity. More importantly, the cell viability of incubated by the CS NPs/(PDDA/PSS)2 /PFV/HS nanocomposite was higher than that of incubated by the CS NPs/(PDDA/PSS)2 /PFV nanocomposite, indicating the introduction of the HS layer increased the cytocompatibility of the nanocomposites.

4. Conclusion

Fig. 7. Cell viability results after incubation of A549 cells with various concentration of PFV (0–20 ␮M) adsorbed on nanocomposite. The percentage cell viability is calculated relative to that of the cells without adding nanocomposite is defined as a viability of 1.0.

In conclusion, a bimodal fluorescent–magnetic nanocomposite based on fluorescent ionic water-soluble conjugated polymer and small magnetic nanoparticles was developed by a simply LbL fabrication. This self-assembly technique remains the monodispersity of the magnetic NPs and the bright fluorescence of PFV. Therefore, the nanocomposites show monodispersity, fluorescence and good biocompatibility that are suitable for efficient cellular imaging. In addition, the outer layer of the nanocomposite is assembled with polysaccharide heparin sodium, which efficiently increases the cellular uptake ability and biocompatibility of the nanocomposites. This line of research provides a new approach for the preparation of the multimodal nanoparticles for the complex biologic systems.

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