Biological evaluation of surface-modified magnetic nanoparticles as a platform for colon cancer cell theranostics

Colloids and Surfaces B: Biointerfaces 161 (2018) 35–41

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Biological evaluation of surface-modified magnetic nanoparticles as a platform for colon cancer cell theranostics Maksym Moskvin a , Michal Babiˇc a , Salette Reis b , M. Margarida Cruz c , Liliana P. Ferreira c,d , Maria Deus Carvalho e , Sofia A. Costa Lima b,∗ , Daniel Horák a,∗ a

Institute of Macromolecular Chemistry, Academy of Sciences of the Czech Republic, Heyrovského Sq. 2, 162 06 Prague 6, Czech Republic UCIBIO-REQUIMTE, Departamento de Ciências Químicas, Faculdade de Farmácia, Universidade do Porto, Rua de Jorge Viterbo Ferreira, 228, 4050-313 Porto, Portugal c BioISI, Biosystems and Integrative Sciences, Faculdade de Ciências, Universidade de Lisboa, Campo Grande, 1749-016 Lisboa, Portugal d Department of Physics, University of Coimbra, 3004-516 Coimbra, Portugal e CQB, Centro de Química e Bioquímica, Faculdade de Ciências, Universidade de Lisboa, Campo Grande, 1749-016 Lisboa, Portugal b

a r t i c l e

i n f o

Article history: Received 16 June 2017 Received in revised form 1 September 2017 Accepted 10 October 2017 Available online 12 October 2017 Keywords: Iron oxide nanoparticles Carbohydrates Poly(N,N-dimethylacrylamide) Cell cycle Cellular uptake Apoptosis

a b s t r a c t Magnetic nanoparticles offer multiple possibilities for biomedical applications. Besides their physicochemical properties, nanoparticle-cellular interactions are determinant for biological safety. In this work, magnetic nanoparticles were synthesized by one-shot precipitation or two-step reaction and coated with biocompatible polymers, such as poly(l-lysine) and poly(N,N-dimethylacrylamide-co-acrylic acid), and carbohydrates, like l-ascorbic acid, d-galactose, d-mannose, and sucrose. The resulting magnetic nanoparticles were characterized by dynamic light scattering, FT-Raman spectroscopy, transmission electron microscopy, SQUID magnetometry, and Mössbauer spectroscopy. Ability of the nanoparticles to be used in theranostic applications was also evaluated, showing that coating with biocompatible polymers increased the heating efficiency. Nanoparticles synthesized by one-shot precipitation were 50% larger (∼13 nm) than those obtained by a two-step reaction (∼8 nm). Magnetic nanoparticles at concentrations up to 500 ␮g mL−1 were non-cytotoxic to L929 fibroblasts. Particles synthesized by one-shot precipitation had little effect on viability, cell cycle and apoptosis of the three human colon cancer cell lines used: Caco-2, HT-29, and SW-480. At the same concentration (500 ␮g mL−1 ), magnetic particles prepared by a two-step reaction reduced colon cancer cell viability by 20%, affecting cell cycle and inducing cell apoptosis. Uptake of surface-coated magnetic nanoparticles by colon cancer cells was dependent on particle synthesis, surface coating and incubation time. © 2017 Elsevier B.V. All rights reserved.

1. Introduction Magnetic nanoparticles, due to their magnetic field-responsive properties, have attracted considerable attention for several biomedical applications, such as biosensors, drug delivery systems, hyperthermia (cancer therapy), and magnetic resonance imaging (MRI) [1,2]. These applications require intravenous administration of the particles and their guidance to the targeted tissue is done either passively, through the enhanced permeation and retention effect, or actively, by a ligand molecule [3]. Uncoated nanoparticles exhibit inferior colloidal stability and are rapidly removed from the blood circulation through the

∗ Corresponding authors. E-mail addresses: [email protected] (S.A.C. Lima), [email protected] (D. Horák). https://doi.org/10.1016/j.colsurfb.2017.10.034 0927-7765/© 2017 Elsevier B.V. All rights reserved.

reticuloendothelial system [4]. Surface modifications can render the magnetic particles biocompatible, colloidally stable [5], and allow drug delivery [6]. Poly(ethylene glycol) (PEG) is commonly used as a non-ionogenic and hydrophilic agent to modify the nanoparticle surface through covalent binding, improving particle biocompatibility, and reducing immunogenicity [7,8]. Also, ionogenic polymers containing amino- or carboxyl groups can be anchored on the particle surface enhancing the particle uptake by cells due to increased surface charge. Optionally, poly- and monosaccharides, including sucrose, d-galactose, d-mannose, and l-ascorbic acid, have been demonstrated to be good particle stabilizers due to complex formation with iron oxides [9,10]. There are many other types of coatings reported, including polyanhydride, dendrimers, citrate, or dimercaptosuccinic acid [11–13]. Though magnetic nanoparticles have proved to be safe in experimental models [14], the development of new modified particles

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requires the identification of potential risks and toxicity effects as the particle ability to interact with cells remains unknown [15,16]. In vitro models are reliable tools to unravel the effects of particles in the human body, allowing to estimate their toxicity and biocompatibility [17]. The high surface/volume ratio of the particles enables cell and tissue accumulation but makes the particles quite reactive [18], which may result in toxicity. Among several factors, magnetic nanoparticle cellular toxicity depends on the particle size, shape, porosity, surface charge, chemical composition, and colloidal stability [19]. Moreover, the particle/cellular interactions and responses differ, depending on the cell type [20–22]. Hence, it is important to analyze the interactions of the nanoparticles with a specific cell line. Magnetic nanoparticles with potential biomedical applications may then proceed towards in vivo validation and biodistribution assays. In the present work, a systematic study was conducted to assess the in vitro magnetic nanoparticle toxicity on colon carcinoma cell lines for theranostic applications. The effect of surface coating was evaluated by synthetizing uncoated and polymer- and saccharide-coated ␥-Fe2 O3 nanoparticles. As a polymer coating, poly(l-lysine) (PLL) and poly(N,N-dimethylacrylamide-co-acrylic acid) (PDM) were selected, while saccharides included sucrose (SC), d-galactose (GA), d-mannose (MAN), and l-ascorbic acid (ASA). The magnetic properties of the nanoparticles were characterized and their ability to operate as nanoheaters for magnetic hyperthermia was evaluated. To characterize the biocompatibility of the nanoparticles obtained by both procedures, cell viability of three mammalian colon cancer cell lines was assayed. The cells included human epithelial colorectal adenocarcinoma Caco-2, HT-29, and SW-480 cells, and mouse fibroblasts (L929). The cells were incubated with the various ␥-Fe2 O3 nanoparticles, and the particle cell uptake kinetics, as well as the effects on cell viability, cell cycle, and apoptosis were quantitatively evaluated. According to our results, the biological effect of the particles was influenced by both synthesis procedure and surface coating. Surface-coated magnetic nanoparticles displayed different behaviour in terms of cell uptake and low toxicity, which is of great interest for further biomedical applications in colon cancer theranostics. 2. Materials and methods 2.1. Materials FeCl2 ·4H2 O (98%), FeCl3 ·6H2 O (98%), l-ascorbic acid (ASA; 98%), poly(l-lysine) hydrobromide (PLL; Mw = 70,000–150,000), N,Ndimethylacrylamide (DMA), acrylic acid (AA), propidium iodide (PI), and 2,2 -azobis(2-methylpropionitrile) (AIBN; recrystallized from ethanol) were obtained from Sigma-Aldrich (St. Louis, USA). Sodium hypochlorite was obtained from Bochemie (Bohumín, Czech Republic). Hydrochloric acid (35%), ammonium hydroxide (25%), hydrogen peroxide (30%), tetrahydrofuran, toluene, crystalline sucrose (SC; 98%), and d-galactose (GA; 97%) were obtained from Lach-Ner (Neratovice, Czech Republic). D(+)-mannose (99%) was obtained from Acros Organics (Geel, Belgium). Ultrapure Qwater from a Milli-Q Gradient A10 system (Millipore, Molsheim, France) was used throughout the experiments. All other reagent grade chemicals were purchased from Sigma-Aldrich and used as received. 2.2. Preparation of poly(N,N-dimethylacrylamide-co-acrylic acid) AA (0.3 g) and DMA (3 g) were dissolved in tetrahydrofuran/toluene mixture (3.5 mL/3.5 mL). After addition of AIBN (10 mg), the mixture was purged with nitrogen for 10 min

and the polymerization was started by heating at 70 ◦ C for 8 h. Resulting poly(N,N-dimethylacrylamide-co-acrylic acid) (PDM) was dissolved in ethanol and precipitated three times in diethyl ether. Dried polymer was dissolved in water (8.8 mg mL−1 ) and filtered through syringe with poly(vinylidene fluoride) membrane (Millipore; 0.22 ␮m pores).

2.3. Preparation of -Fe2 O3 nanoparticles and their surface modification The first set of saccharide-modified particles (Run I/2-4; Table 1), such as SC-, GA-, and ASA-coated nanoparticles, was prepared by one-shot precipitation and modification of the primary ␥-Fe2 O3 colloid (Run I/1) synthesized according to the following protocol. FeCl3 ·6H2 O (40 mmol) and FeCl2 ·4H2 O (20 mmol) were dissolved in water (470 mL) and charged in a 500 mL glass reactor equipped with a turbine impeller and the mixture was heated at 70 ◦ C with stirring (600 rpm). 25% NH4 OH (30 mL) was added at once to the mixture, black magnetite (Fe3 O4 ) was precipitated and the reaction mixture was heated at 90 ◦ C for 1 h with stirring (600 rpm). After cooling to room temperature (RT), 35% HCl was added to adjust pH to 5–6. Resulting Fe3 O4 was oxidized to maghemite (␥Fe2 O3 ) by addition of 30% hydrogen peroxide (5 mL) under slow heating from 20 to 90 ◦ C for 1 h with stirring (600 rpm). Finally, the ␥-Fe2 O3 particles were magnetically separated, washed with water (100 mL), and redispersed using a Branson S-450D Sonicator (Danbury, USA; 10 mm sonotrode) at 10% output for 5 min. After triple washing with water, a dispersion (100 mL) containing 45 mg of the particles per mL was obtained. Consequently, aqueous ␥Fe2 O3 colloid (containing 100 mg of dry ␥-Fe2 O3 ) was diluted with water to 15 mL, charged in a 25-mL glass reactor equipped with a turbine impeller, and the mixture was heated at 40 ◦ C with stirring (700 rpm). After dropwise addition of aqueous saccharide (40 mg; 5 mL), the reaction continued at RT for 2 h with stirring (500 rpm) under purging with argon. Resulting ␥-Fe2 O3 @SC, ␥-Fe2 O3 @GA, and ␥-Fe2 O3 @ASA nanoparticles were washed with water (30 mL), magnetically separated, and redispersed in water (20 mL) under sonication at 10% output for 3 min to concentration of 4.4 mg of ␥-Fe2 O3 per mL. The second set of surface-modified ␥-Fe2 O3 nanoparticles (Run II/2-4; Table 1) was prepared from primary ␥-Fe2 O3 colloid (Run II/1) by two-step precipitation method according to the following protocol. Aqueous 0.2 M FeCl3 (100 mL) was mixed with 0.5 M NH4 OH (95 mL; less than an equimolar amount) with sonication (12.7 mm sonotrode) at RT for 2 min to form Fe(OH)3 colloid. Aqueous 0.2 M FeCl2 (50 mL) was then added with sonication and the mixture poured into aqueous 0.5 M NH4 OH (350 mL). The resulting Fe3 O4 coagulate was left to grow for 15 min, magnetically separated and repeatedly (7–10×) washed (peptized) with water to remove all impurities (including NH4 Cl) remaining after the synthesis. Finally, 0.1 M trisodium citrate (12.5 mL) was added with sonication, and Fe3 O4 was oxidized by slow addition of 5% sodium hypochlorite solution (10 mL). The above-described washing procedure was repeated to yield the primary ␥-Fe2 O3 colloid that was filtered via syringe with mixed cellulose ester membrane (0.45 ␮m pores). To prepare ␥-Fe2 O3 @MAN particles (Table 1), aqueous Dmannose (2 mL; 128 mg mL−1 ) was added dropwise to the primary ␥-Fe2 O3 colloid (8 mL; 44 mg of iron oxide) with sonication (1 mm sonotrode) for 5 min. Similarly, aqueous PLL (0.2 mL; 1 mg mL−1 ) and aqueous PDM (5 mL; 8.8 mg mL−1 ) were added to the same primary colloid (9.8 or 5 mL) to yield ␥-Fe2 O3 @PLL and ␥-Fe2 O3 @PDM, respectively.

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Table 1 Properties of surface-modified ␥-Fe2 O3 nanoparticles. Designation

Dn (nm)

Ð

Dh (nm)

PDI

Coatinga (wt.%)

␨-potentialb (mV)

Run I

␥-Fe2 O3 ␥-Fe2 O3 @SC ␥-Fe2 O3 @GA ␥-Fe2 O3 @ASA

13 12 14 15

1.22 1.31 1.26 1.24

110 157 168 164

0.12 0.22 0.24 0.26

– 4.6 5.3 7.0

44 ± 3 36 ± 2 27 ± 2 −25 ± 2

Run II

␥-Fe2 O3 ␥-Fe2 O3 @MAN ␥-Fe2 O3 @PLL ␥-Fe2 O3 @PDM

8 7 6 10

1.28 1.34 1.33 1.27

63 73 90 173

0.30 0.25 0.33 0.12

– 54 0.5 50

−52 ± 1 −46 ± 1 −31 ± 1 −25 ± 4

Dn − number-average particle diameter (TEM); Ð − dispersity (TEM); Dh − hydrodynamic diameter (DLS); PDI − polydispersity index (DLS); a amount of coating determined from elemental analysis; b ␨-potential measured at pH 6.5.

2.4. Characterization of the nanoparticles Size and dispersity (Ð = Dw /Dn , where Dw and Dn are weight- and number-average particle diameter, respectively) were determined from microphotographs (using at least 300 particles) obtained by a Tecnai Spirit transmission electron microscope (TEM; FEI; Brno, Czech Republic) and analyzed with Atlas image analysis software (Tescan, Brno, Czech Republic) with a standard error of 0.6 nm. The samples for TEM analysis were prepared by spraying aqueous iron oxide dispersion on a grid with carbon membrane and drying. Surface ␨-potential, hydrodynamic diameter (Dh ), and polydispersity index (PDI) were measured by dynamic light scattering (DLS) using a ZEN3600 Nano-ZS Zetasizer (Malvern Instruments; Malvern, Worcestershire, UK). FT-Raman spectra were measured on a Thermo Nicolet 6700 FT-IR spectrometer with attached NIR FT-Raman module (Thermo Fisher Scientific; Waltham, USA). An excitation laser with the 180◦ reflecting sample geometry and 1064 nm wavelength was used in pair with an air-cooled In-Ga-As detector. Content of the modifier on the surface was calculated from elemental analysis (Perkin-Elmer 2400 CHN; Beaconsfield, UK). 57 Fe Mössbauer spectra were collected at 78 K in transmission mode using a conventional constant acceleration spectrometer and a 50 mCi 57 Co source in a Rh matrix. The ferrofluids were frozen in adequate Mössbauer sample-holders and introduced in the cryostat at temperatures <200 K. The velocity scale was calibrated using ␣-Fe foil. The spectra were fitted using distributions of the magnetic field with the WinNormos program. Magnetic properties of the nanoparticle dispersions were characterized using a QD-MPMS SQUID magnetometer as a function of temperature and applied magnetic field. A small volume of each dispersion (0.5–1 mL) was placed in a quartz tube and purged with nitrogen to remove oxygen before introducing it in the magnetometer. The temperature dependence of the magnetic moment was measured from 10 to 250 K to keep the dispersion frozen under an applied magnetic field of 5 mT, after cooling from 250 K in zero magnetic field (zero field cooled − ZFC) and after cooling under the measurement field (field cooled − FC). Hysteresis curves were obtained at 250 K for magnetic fields up to 5.5 T. Induction heating of the nanoparticles was performed using an experimental set-up based on an Easy Heat 0224 device (Ambrell) described previously [23], for AC magnetic field with 13.9 kA m−1 amplitude and 274 kHz frequency.

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were acquired from Gibco (Invitrogen; Carsbad, CA, USA). The cells were grown and maintained in an incubator at 37 ◦ C under 5% CO2 atmosphere. 2.5.2. In vitro cell viability assay Fibroblasts (L929) and human colon cancer cells (Caco-2, HT29, and SW-480) were seeded on 96-well plates at 10,000 cells per well in supplemented DMEM medium (100 ␮L). The medium containing various concentrations of uncoated ␥-Fe2 O3 (Run I/1 and II/1), ␥-Fe2 O3 @SC (Run I/2), ␥-Fe2 O3 @GA (Run I/3), ␥-Fe2 O3 @ASA (Run I/4), ␥-Fe2 O3 @MAN (Run II/2), ␥-Fe2 O3 @PLL (Run II/3), and ␥Fe2 O3 @PDM (Run II/4) nanoparticles (0–500 ␮g mL−1 ) was added to the wells. Control cells were incubated under the same conditions in the absence of the nanoparticles. After 24 h of treatment, the excess of particles was removed by washing the cells with phosphate-buffered saline (PBS) three times and the cell viability was assessed. For the colorimetric assay, the treated cells were washed with PBS twice, incubated with 3-(4,5-dimethylthiazol2-yl)-2,5-diphenyltetrazolium bromide (MTT; 0.5 mg mL−1 ) for 3 h, the medium was removed, and formazan crystals were solubilized by a 10 min treatment with dimethyl sulfoxide (100 ␮L). The absorbance in each well was read on a SynergyTM HT Multimode microplate reader (BioTek Instruments; Winooski, VT, USA) at 545 nm and cell viability was calculated according to the following equation [24]: Cell viability (%) = OD545(sample) /OD545(control) × 100, where OD545(sample) and OD545(control) are optical density of solution containing magnetic nanoparticles and control without particles, respectively. For the propidium iodide (PI) viability assay, cells treated with the particles for 24 h were washed with PBS twice, recovered using 0.25% (w/v) trypsin, PI (20 ␮g mL−1 ) was added, and the cells analyzed using an Accuri C6 flow cytometer (Ann Arbor, MI, USA). 2.5.3. Cell cycle analysis Colon cancer cell lines were incubated with the nanoparticles at a concentration 500 ␮g mL−1 for 24 h, the cells were harvested, resuspended, and incubated in ice-cold 70% ethanol for 5 min. After washing with cold PBS, staining with PI (20 ␮g mL−1 ), and ribonuclease (100 ␮g mL−1 ) at RT for 45 min, the cells were analyzed using an Accuri C6 flow cytometer.

2.5. In vitro cellular assays 2.5.1. Cell culture Primary adherent mouse fibroblast connective tissue cells (L929) and human colon cancer cells (Caco-2, HT-29, and SW480) obtained from American Type Cell Culture (ATCC-LGC Standards; Barcelona, Spain) were cultured in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% (v/v) fetal bovine serum (FBS) and 1% (v/v) penicillin/streptomycin. All culture reagents

2.5.4. Analysis of cell apoptosis Flow cytometric analysis of external phosphatidylserine exposure for cell apoptosis assessment was described previously [25]. Briefly, the cells were analyzed for annexin V binding and PI incorporation to distinguish between apoptotic and necrotic cells. Analysis was performed on an Apoptosis Assay Kit (Molecular Probes; Eugene, OR, USA) according to the manufacturer’s instructions and CellQuest software (BD Biosciences; San Jose, CA, USA).

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2.5.5. Cellular uptake assay To compare the cellular uptake of uncoated and coated magnetic nanoparticles, 100,000 colon cancer cells (Caco-2, HT-29, and SW480) were seeded in DMEM medium (400 ␮L) in 24-well plates. Upon cell adhesion, the medium was replaced with 25–100 ␮g of medium containing magnetic nanoparticles. After 4, 10, and 24 h, the cells were washed with PBS twice, trypsinized, collected in medium (200 ␮L), and analyzed in the flow cytometer to determine the side-scattering height (SSCH) fluorescence levels in fluorescence laser 1 channel. Standard deviations were calculated from three replicates. 2.5.6. Statistical analysis The in vitro results were presented as means ± standard deviations from at least three independent experiments and compared by one-way ANOVA, followed by Dunnet’s test. P <0.05 was taken as statistically significant and P <0.01 as highly significant. 3. Results and discussion 3.1. Iron oxide nanoparticles: synthesis, coating and characterization Several surface-modified maghemite (␥-Fe2 O3 ) nanoparticles intended for colon carcinoma theranostics were designed, synthesized, and characterized. ␥-Fe2 O3 was selected as a preferable form of iron oxide, because Fe3 O4 is easily oxidized in air and consequently less stable over a long period of time [26]. To find an optimal method of ␥-Fe2 O3 particle preparation in terms of future biological applications and minimal cell toxicity, two slightly different approaches were chosen. While in the first approach (Run I) the nanoparticles were prepared by one-shot precipitation of FeCl2 and FeCl3 with ammonia at 90 ◦ C, stabilized by the addition of HCl, and oxidizing the resulting Fe3 O4 to ␥-Fe2 O3 with H2 O2 , the second method (Run II) consisted in two-step precipitation at RT, stabilization with trisodium citrate and oxidation with NaOCl. Another distinction between both types of the particles consisted in the selection of the surface modifier. While the surface of the Run I particles was modified with a carbohydrate (sucrose, SC or d-galactose, GA), or its derivative (l-ascorbic acid, ASA) to enhance the particle stability and antioxidative properties, those of Run II were modified with d-mannose (MAN), as well as with polymers, such as poly(l-lysine) (PLL) and poly(N,N-dimethylacrylamide-coacrylic acid) (PDM) (Fig. 1). The particles synthesized by method Run I were carefully washed with water to remove any residual (unbound) carbohydrate. It was shown earlier that amounts of PLL, PDM, and MAN used in the preparation of the Run II nanoparticles were found suitable for mesenchymal stem cell labeling without necessity of additional washing [27,28]. According to TEM (Supplementary Material, Fig. S1), dry particles prepared in one-shot precipitation (Run I) were larger (Dn ∼13 nm) than those obtained by two-step reaction (Dn ∼8 nm; Run II). Due to the low electron density of organic coatings, they were not noticeable in the TEM micrographs; as a result, Dn values of coated and uncoated particles were similar. Particle size distributions assessed by TEM and DLS using the dispersity (Ð) and the polydispersity index (PDI), respectively, were moderately broad and approximately the same for all the magnetic particles (Table 1). Hydrodynamic size (Dh , obtained from DLS) of the surface-modified Run I magnetic nanoparticles (∼160 nm) was larger in comparison with Run II particles (73–173 nm), probably due to slight cluster formation. It should be noted that due to the presence of the hydration layer, Dh of the particles in water was larger than Dn of the dried particles. Moreover, the mean value obtained from TEM is proportional to Di while DLS provides the intensity dis-

Fig. 1. Chemical structure of the surface modifiers. (A) Sucrose, (B) galactose, (C) ascorbic acid, (D) d-mannose, (E) poly(l-lysine), and (F) poly(N,Ndimethylacrylamide-co-acrylic acid).

tribution characterized by Z-average, which is proportional to Di 6 and sensitive to large particles. As expected, Dh of the surfacemodified particles was larger than that of primary ␥-Fe2 O3 due to the presence of the surface shell. Absolute value of the ␨-potential (pH 6.5) for the magnetic nanoparticles obtained in Run I and II was typically >25 mV suggesting their good colloidal stability (Table 1). Neat ␥-Fe2 O3 , ␥-Fe2 O3 @SC, and ␥-Fe2 O3 @GA Run I particles bore positive charge due to the stabilization with HCl, while the trisodium citrate-stabilized Run II particles were negatively charged. The amount of coating on the nanoparticles, determined from elemental analysis, differed depending on the way of the synthesis. Magnetic Run I nanoparticles contained ∼6 wt.% of coating, while more than half of ␥-Fe2 O3 @MAN and ␥-Fe2 O3 @PDM weight consisted of the surface modifier. This discrepancy can be ascribed to rather rigorous washing of Run I particles. In a previous work describing ␥-Fe2 O3 @PLL nanoparticles with 0.45 wt.% of polymer coating, the particles exhibited high cell labeling efficiency and no toxicity [29]. Run I and II magnetic nanoparticles were thoroughly characterized and compared with published results [27–29]. FT-Raman spectra of ␥-Fe2 O3 @SC and ␥-Fe2 O3 @GA nanoparticles had certain similarities (Supplementary Material, Fig. S2A,B). The peaks at 642, 661, and 704 cm−1 were attributed to ␦(C O) wagging vibrations of CH OH groups. Two pairs of bands were observed at (i) 1039 and 1068 cm−1 , ascribed to stretching (C O) of primary hydroxyl groups, and (ii) 1126 and 1137 cm−1 , ascribed to stretching (C O) of secondary hydroxyls. The peaks at 1238 and 1248 cm−1 documented the presence of as (C O C) stretching vibrations of ether oxygen bonds in the saccharide rings, and those at 1462 and 1488 cm−1 were attributed to methylene group scissoring vibrations ␦(H C H). The main difference in the spectra of the ␥-Fe2 O3 @SC and ␥-Fe2 O3 @GA nanoparticles consisted in the presence of stretching (C O) vibrations of ether oxygen connecting monosaccharide rings at 1348 cm−1 (Supplementary Material, Fig. S2A). The FT-Raman spectrum of ␥-Fe2 O3 @ASA nanoparticles exhibited several characteristic stretching and deformation vibrations: ring (C C) at 1667 cm−1 , ring (C O C) at 1128 cm−1 , ring (C C) at 819 cm−1 , ␦OOP (–OH) at 628 cm−1 , bending ␦ring (–OH)

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at 1028 cm−1 , twisting ␦(C O H) at 1255 cm−1 , and wagging ␦(C H) at 1321 cm−1 (Fig. S2C). The FT-Raman spectroscopy thus confirmed the presence of SC, GA, and ASA on the ␥-Fe2 O3 surface.

Table 2 Specific loss power (SLP) calculated from the induction heating measurements. Designation

SLP (W g−1 )

Run I

␥-Fe2 O3 ␥-Fe2 O3 @ASA

81 ± 2 85 ± 2

Run II

␥-Fe2 O3 ␥-Fe2 O3 @MAN ␥-Fe2 O3 @PLL ␥-Fe2 O3 @PDM

65 ± 1 87 ± 2 75 ± 2 73 ± 2

3.2. Magnetic characterization and hyperthermia measurements The Mössbauer spectra of the uncoated Run I and II ␥-Fe2 O3 nanoparticles showed that the hyperfine parameters obtained from the fitting analysis (Supplementary Material, Fig. S3A) were quite similar and typical for ␥-Fe2 O3 nanoparticles at 78 K: isomer shift of 0.43(1) mm s−1 , quadrupole splitting of 0.00(1) mm s−1 , and a distribution of hyperfine magnetic fields with mean values of 48.1 ± 5.6 T and 48.7 ± 4.5 T for Run I and II, respectively. The magnetization hysteresis curve of ␥-Fe2 O3 @ASA nanoparticles at 250 K resembled those for all the other particles displaying the expected behaviour of ferrimagnetic ␥-Fe2 O3 with a high slope at low fields and reaching saturation for external fields >1 T (Supplementary Material, Fig. S3B). The amount of ␥-Fe2 O3 in the colloids was obtained from the corresponding saturation magnetizations at 250 K, assuming that the saturation magnetization of bulk ␥-Fe2 O3 was 75 A m2 kg−1 [30]. Inhomogeneous distribution of the ␥-Fe2 O3 particles in the measured samples added 5% uncertainty to the determination of ␥-Fe2 O3 concentrations in the nanoparticle dispersions. While concentrations of ␥-Fe2 O3 in the Run I nanoparticles determined from the saturation magnetization were in the range of 2–3.9 mg mL−1 , those in the Run II were higher (6.6–7.6 mg mL−1 ). Zero field cooled (ZFC) and field cooled (FC) magnetization curves of Run II ␥-Fe2 O3 nanoparticles showed the typical temperature dependence for superparamagnetic nanoparticles [31,32]. Broad peak with a maximum at ∼200 K appeared in the spectrum of ␥-Fe2 O3 @MAN and ␥-Fe2 O3 @PDM and at ∼160 K in the spectrum of ␥-Fe2 O3 @PLL particles (Supplementary Material, Fig. S3C). The peak of uncoated Run II ␥-Fe2 O3 nanoparticles was broader than that of coated particles. Considering that the maximum defines the blocking temperature (TB ) of the particle distribution and that the particle size was similar (Dn = 6–10 nm), the difference observed between coated and uncoated particles can be explained by aggregation of the latter. Aggregation implies the existence of magnetic dipolar interactions that usually shift TB to a higher value. Variances in ZFC curves observed between the coated nanoparticles indicate that the specific coatings affect the nanoparticle magnetic moment differently. ZFC-FC curves of Run I ␥-Fe2 O3 and ␥-Fe2 O3 @ASA nanoparticles showed a continuous increase of magnetization with temperature up to 250 K indicating that the associated TB was above this temperature (Supplementary Material, Fig. S3D). This is consistent with the larger size of the Run I magnetic nanoparticles (∼14 nm) compared with that of Run II (∼8 nm). The temporal evolution of the temperature in the particle dispersion was measured during 100 s under the alternating magnetic field and was fitted considering a constant magnetic heating power supplied by the nanoparticles and linear energy exchanges between the particles and the environment [23]. During the hyperthermia measurements, ␥-Fe2 O3 @GA and ␥-Fe2 O3 @SC nanoparticles were deposited on tube walls and the heating efficiency, which requires both temperature equilibrium and good particle dispersion at the beginning of the measurement, could not be evaluated. To determine the heating power from the temperature variation, the glass container contribution was carefully evaluated in the measurements, lasting for 100 s at similar temperatures. The heating efficiency of the nanoparticles was evaluated by the specific loss power (SLP; Table 2). The Run II surface-modified particles had an increased SLP as documented by ␥-Fe2 O3 @MAN nanoparticles displaying 33% higher SLP than the neat ␥-Fe2 O3 particles, while ␥-

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Fe2 O3 @PLL and ␥-Fe2 O3 @PDM had only ∼12% higher SLP. Coating of ␥-Fe2 O3 nanoparticles with polymers was thus a useful tool to increase efficiency of the nanoparticles in magnetic hyperthermia. SLP of ␥-Fe2 O3 @ASA nanoparticles was slightly increased (by 6%) compared to that of neat ␥-Fe2 O3 . Higher heat release efficiency of the coated nanoparticles can be explained by their improved dispersibility in water, because coating reduced the attractive interactions between magnetic nanoparticles and prevented their consequent aggregation. Run I ␥-Fe2 O3 nanoparticles, due to their larger size, had higher heating efficiency than those of Run II. This is in agreement with earlier reports describing importance of the iron oxide nanoparticle size on the heating efficiency in an alternating magnetic field [31]. 3.3. In vitro cytotoxicity The nanoparticle toxicity was studied on three colon cancer cell lines (Caco-2, HT-29, and SW-480) and L929 fibroblasts, taken as a reference for the safety assessment according to ISO 10993-1:2009 on “Biological evaluation of medical devices”. In vitro toxicity assays are usually based on colorimetric or fluorimetric detection. Data from dye-based assays may vary according to the type of magnetic nanoparticles and their interaction with dye. It is highly recommended to apply more than one assay to determine the nanoparticle toxicity for risk assessment [33]. Cell viability was detected by MTT and PI incorporation assay upon exposure to uncoated ␥-Fe2 O3 (Run I and II), ␥-Fe2 O3 @SC, ␥-Fe2 O3 @GA, ␥-Fe2 O3 @ASA, ␥-Fe2 O3 @MAN, ␥-Fe2 O3 @PLL, and ␥-Fe2 O3 @PDM nanoparticles (Supplementary Material, Fig. S4). Both viability assays showed similar cytotoxic profiles in terms of flow cytometric analysis upon PI incorporation and all nanoparticles seemed to be well tolerated by L929 fibroblasts (Supplementary Material, Fig. S4D,H). Some differences were observed in the Caco-2 and SW480 cell response towards the Run I and II particles. The Run I nanoparticles at concentrations ≤500 ␮g mL−1 did not have statistically significant effect on Caco-2, HT-29, and SW-480 cell viability after 24-h treatment (Supplementary Material, Fig. S4A–C). Highest concentration of Run I nanoparticles (500 ␮g mL−1 ) resulted in cell viabilities >90% but the same concentration of Run II particles induced statistically significant decrease (20%; P <0.01) in Caco2 and SW-480 cell viability, whereas HT-29 cell viability did not significantly decrease (Supplementary Material, Fig. S4E–G). Nanoparticle interaction with the cell surface might induce other cellular responses, such as programmed cell death or changed cell cycle [11]. As described earlier, the nanoparticle uptake by the cells is strongly influenced by their cell cycle phase [34]. The cell cycle of all three colon cancer lines incubated with the magnetic nanoparticles was investigated by flow cytometry. Consistently with the aforementioned results, the Run I particles, at a concentration of 500 ␮g mL−1 , did not statistically influence the Caco-2, HT-29, and SW-480 cell cycle after 24-h treatment (Supplementary Material, Fig. S5A–C). In contrast, Run II particles, at the same concentration, increased the number of damaged cells in the subG1 population (up to 35–40%) and decreased the G0 –G1 (up to 25–30%), as well as S cycle phases (to ∼15%), when compared to

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Fig. 2. Cellular interaction with the magnetic nanoparticles. Dependence of Caco-2, HT-29, and SW-480 cellular uptake on type of synthesis and particle coating measured as side-scattered fluorescence height (SSCH); the nanoparticles (250 ␮g mL−1 ) were internalized by the cells for 24 h. Values represent the mean ± standard deviation (n = 3). * P <0.05, ** P <0.01, and *** P <0.001.

untreated controls (Supplementary Material, Fig. S5D–F). Identical results were observed for Caco-2, HT-29, and SW-480 cell lines exposed to the particles during cell apoptosis assessment (Supplementary Material, Fig. S5G). Compared to those from Run I, higher percentage of apoptotic cells exposed to the Run II particles reflected strong cellular response and induction of cell death (Supplementary Material, Fig. S5G). 3.4. Cellular uptake kinetics The physico-chemical characteristics of magnetic nanoparticles that provide optimal internalization in non-phagocytic cells remain uncertain. To date, few studies dealt with the cellular uptake kinetics of magnetic nanoparticles in colon cancer cells [22,35]. Cells can be morphologically analyzed based on their size and granularity by flow cytometry, assessed by the forwardand side-scattering, respectively. As internalized nanoparticles also scatter light, the side-scatter analysis in flow cytometry can assess cellular uptake of the particles [10]. Because cellular uptake depends on the particle incubation time [36], colon cancer cell lines were exposed to non-toxic concentrations of the Run I and II magnetic nanoparticles for 4, 10, and 24 h. Uptake of the magnetic nanoparticles by cancer cells shifted the sidescattered height in the flow cytometric analysis. The uptake kinetics for viable Caco-2, HT-29, and SW-480 cells treated with 250 ␮g of uncoated and coated nanoparticles per mL was determined (Supplementary Material, Figure S6), as dead cells were excluded by selecting phenotypically live cells (negative PI staining). The results clearly indicated that all the magnetic nanoparticles were internalized by the three human colon cell lines. A statistically significant (P <0.05) increase of surface-coated Run I nanoparticle (␥-Fe2 O3 @SC, ␥-Fe2 O3 @GA, and ␥-Fe2 O3 @ASA) uptake was noticed compared to uncoated Run I nanoparticles (Supplementary Material, Fig. S6A,C,E). ␥-Fe2 O3 @MAN and ␥-Fe2 O3 @PLL nanoparticles exhibited significantly (P <0.01) increased uptake by Caco-2, HT-29, and SW-480 cell lines, in comparison to uncoated Run II magnetic nanoparticles (Supplementary Material, Fig. S6B,D,F). In contrast, PDM coating on the ␥-Fe2 O3 nanoparticles hampered internalization of the nanoparticles by the colon cancer cell lines. This can be ascribed to hydrophilicity of PDM, which is analogous to PEG coating [37,38], and smaller absolute value of ␨-potential of ␥-Fe2 O3 @PDM particles. The cellular uptake of Run II magnetic nanoparticles was time- and coating-dependent. Qualitative estimation of the particle uptake by various cancer cells demonstrated higher uptake of the Run I nanoparticles in cancer cells compared to the Run II ones (Fig. 2). This could be explained by recognition of carbohydrate-modified particles by lectins, such

as mannose-binding protein in the cell membrane [39]. The order of magnetic nanoparticles uptake by colon cancer cells was Caco2 > HT-29 > SW-480.

4. Conclusions Biomedical applications of magnetic nanoparticles depend on several inherent properties, but particularly on their cellular interactions. Lower internalization into cancer cells has hampered therapeutic efficacy in cancer treatment and diagnosis. The present work illustrates that preparation method and surface coating of ␥Fe2 O3 nanoparticles influence cellular interactions with selected colon cancer cells. Carbohydrate- and polymer-coated nanoparticles provided satisfactory cell viability over a biomedically relevant time period of 24 h. Carbohydrate coating of the Run I nanoparticles enhanced both biocompatibility and uptake by Caco-2, HT-29, and SW-480 epithelial colorectal adenocarcinoma cell lines. However, modification of the particles with relatively low amounts of sucrose and d-galactose, in comparison with Run II d-mannosemodified particles, induced deposition of the particles on tube walls during magnetic measurements. The Run II uncoated and polymer- or d-mannose-coated particles at a concentration of 500 ␮g mL−1 reduced colon cancer cell viability by ca. 20%. Cellular uptake studies demonstrated that all the magnetic nanoparticles were internalized by the cancer cells. In agreement with previous studies, the mechanism of nanoparticle uptake into the cells is different depending on the type of coating [40]. While carbohydrate-coated iron oxide nanoparticles were taken up via the lectine-mediated membrane transport mechanism present in the majority of mammalian cells, PLL served as a transfection agent. As a result, PLL-coated nanoparticles entered the cells via electrostatic interaction between positive charges of PLL and negative cell surface. In contrast, PDM-coated nanoparticles were presumably engulfed by endocytosis [40]. Thus, surface coatings, with the exception of PDM, enhanced particle internalization in three human colon cancer cell lines. One of the possible applications for the developed surfacemodified magnetic nanoparticles is the hyperthermia cancer treatment. For this purpose, the heating released by magnetic induction was also evaluated. It was shown that the coating, especially with biocompatible polymers, increased the heating efficiency compared to the uncoated nanoparticles, which is a promising result regarding magnetic hyperthermia applications. In summary, the results described here demonstrate that the synthesized surface-modified magnetic nanoparticles can be prospectively used for in vivo colon cancer theranostics.

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Disclosure of interest The authors report no conflicts of interest. Acknowledgements This work was supported by the Czech Science Foundation (No. 16-01128J) and the European Union and Portuguese Foundation for Science and Technology under the partnership agreement PT2020 UID/MULTI/04378/2013, POCI/01/0145/FEDER/007728, UID/MULTI/04046/2013, and UID/MULTI/00612/2013. S. Lima thanks Operac¸ão NORTE-01-0145-FEDER-000011 for researcher contract. This paper is also based upon work from COST Action RADIOMAG (TD1402) supported by COST (European Cooperation in Science and Technology). Opportunity for M. Moskvin’s doctoral studies at the Charles University is acknowledged. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at https://doi.org/10.1016/j.colsurfb.2017.10. 034. References [1] Y.W. Jun, J.W. Seo, J. Cheon, Nanoscaling laws of magnetic nanoparticles and their applicabilities in biomedical sciences, Accounts Chem. Res. 41 (2008) 179–189. [2] G. Liu, J. Gao, H. Ai, X. Chen, Applications and potential toxicity of magnetic iron oxide nanoparticles, Small 9 (2013) 1533–1545. [3] T. Lam, P. Pouliot, P.K. Avti, F. Lesage, A.K. Kakkar, Superparamagnetic iron oxide based nanoprobes for imaging and theranostics, Adv. Colloid Interface Sci. 199 (2013) 95–113. [4] W. Wu, Q. He, C. Jiang, Magnetic iron oxide nanoparticles: Synthesis and surface functionalization strategies, Nanoscale Res. Lett. 3 (2008) 397–415. [5] M. Mahmoudi, S. Sant, B. Wang, S. Laurent, T. Sen, Superparamagnetic iron oxide nanoparticles (SPIONs): Development, surface modification and applications in chemotherapy, Adv. Drug Deliv. Rev. 63 (2011) 24–46. [6] V. Mulens, M.P. Morales, D.F. Barber, Development of magnetic nanoparticles for cancer gene therapy: A comprehensive review, ISRN Nanomater. (2013) (2013), Article ID 646284. [7] A. Al Faraj, A.P. Shaik, A.S. Shaik, Effect of surface coating on the biocompatibility and in vivo MRI detection of iron oxide nanoparticles after intrapulmonary administration, Nanotoxicology 9 (2015) 825–834. [8] L. Maurizi, A.L. Papa, L. Dumont, F. Bouyer, P. Walker, D. Vandroux, N. Millot, Influence of surface charge and polymer coating on internalization and biodistribution of polyethylene glycol-modified iron oxide nanoparticles, J. Biomed. Nanotechnol. 11 (2015) 126–136. [9] C.C. Berry, S. Wells, S. Charles, G. Aitchison, A.S.G. Curds, Cell response to dextran-derivatised iron oxide nanoparticles post internalisation, Biomaterials 25 (2004) 5405–5413. [10] A.M. Dias, A. Hussain, A.S. Marcos, A.C. Roque, A biotechnological perspective on the application of iron oxide magnetic colloids modified with polysaccharides, Biotechnol. Adv. 29 (2011) 142–155. [11] Q.M. Kainz, O. Reiser, Polymer- and dendrimer-coated magnetic nanoparticles as versatile supports for catalysts scavengers, and reagents, Acc. Chem. Res. 47 (2014) 667–677. [12] K. Andreas, R. Georgieva, M. Ladwig, S. Mueller, M. Notter, M. Sittinger, J. Ringe, Highly efficient magnetic stem cell labeling with citrate-coated superparamagnetic iron oxide nanoparticles for MRI tracking, Biomaterials 33 (2012) 4515–4525. [13] R. Mejías, L. Gutiérrez, G. Salas, S. Pérez-Yagüe, T.M. Zotes, F.J. Lázaro, M.P. Morales, D.F. Barber, Long term biotransformation and toxicity of dimercaptosuccinic acid-coated magnetic nanoparticles support their use in biomedical applications, J. Control. Release 171 (2013) 225–233. [14] X. Zhu, S. Tian, Z. Cai, Toxicity assessment of iron oxide nanoparticles in Zebrafish (Danio rerio) early life stages, PLoS One 7 (2012) e46286. [15] X. Wu, Y. Tan, H. Mao, M. Zhang, Toxic effects of iron oxide nanoparticles on human umbilical vein endothelial cells, Int. J. Nanomed. 5 (2010) 385–399. [16] M. Lévy, F. Lagarde, V.A. Maraloiu, M.G. Blanchin, F. Gendron, C. Wilhelm, F. Gazeau, Degradability of superparamagnetic nanoparticles in a model of intracellular environment: Follow-up of magnetic, structural and chemical properties, Nanotechnology 21 (2010) 395103.

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