Biocompatible mesoporous silica-coated superparamagnetic manganese ferrite nanoparticles for targeted drug delivery and MR imaging applications

Biocompatible mesoporous silica-coated superparamagnetic manganese ferrite nanoparticles for targeted drug delivery and MR imaging applications

Journal of Colloid and Interface Science 431 (2014) 31–41 Contents lists available at ScienceDirect Journal of Colloid and Interface Science www.els...

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Journal of Colloid and Interface Science 431 (2014) 31–41

Contents lists available at ScienceDirect

Journal of Colloid and Interface Science www.elsevier.com/locate/jcis

Biocompatible mesoporous silica-coated superparamagnetic manganese ferrite nanoparticles for targeted drug delivery and MR imaging applications Banalata Sahoo a, K. Sanjana P. Devi b, Sujan Dutta a, Tapas K. Maiti b, Panchanan Pramanik a,⇑, Dibakar Dhara a,⇑ a b

Department of Chemistry, Indian Institute of Technology Kharagpur, West Bengal 721302, India Department of Biotechnology, Indian Institute of Technology Kharagpur, West Bengal 721302, India

a r t i c l e

i n f o

Article history: Received 1 April 2014 Accepted 2 June 2014 Available online 11 June 2014 Keywords: Multifunctional nanoparticles Mesoporous silica Folic acid Cancer specificity Doxorubicin Fluorescence imaging

a b s t r a c t Multifunctional mesoporous silica-coated superparamagnetic manganese ferrite (MnFe2O4) nanoparticles (M-MSN) were synthesized and evaluated for targeted drug delivery and magnetic resonance imaging (MRI) applications. MnFe2O4 nanoparticles were prepared by solvothermal route and were silica-coated by surface silylation using sol–gel reactions. Subsequently, silylation was done using (3aminopropyl)triethoxysilane in presence of a surfactant (CTAB), followed by selective etching of the surfactant molecules that resulted in amine-functionalized superparamagnetic nanoparticles (NH2-MSN). Further modification of the surface of the NH2-MSN with targeting (folate) or fluorescent (RITC) molecules resulted in M-MSN. The formation of the M-MSN was proved by several characterization techniques viz. XRD, XPS, HRTEM, FESEM, VSM, BET surface area measurement, FTIR, and UV–Vis spectroscopy. The M-MSN were loaded with anticancer drug Doxorubicin and the efficacy of the DOX loaded M-MSN was evaluated through in vitro cytotoxicity, fluorescence microscopy, and apoptosis studies. The in vivo biocompatibility of the M-MSN was demonstrated in a mice-model system. Moreover, the M-MSN also acted as superior MRI contrast agent owing to a high magnetization value as well as superparamagnetic behavior at room temperature. These folate-conjugated nanoparticles (FA-MSN) exhibited stronger T2-weighted MRI contrast towards HeLa cells as compared to the nanoparticles without folate conjugation, justifying their potential importance in MRI based diagnosis of cancer. Such M-MSN with a magnetic core required for MRI imaging, a porous shell for carrying drug molecules, a targeting moeity for cancer cell specificity and a fluorescent molecule for imaging, all integrated into a single system, may potentially serve as an excellent material in biomedical applications. Ó 2014 Elsevier Inc. All rights reserved.

1. Introduction The emergence of nanoparticle-mediated targeted drug delivery systems as an alternative to the traditional dosage forms have made a great impact leading to the development of new types of therapeutics and diagnostic tools [1]. A variety of carrier materials have been developed as vehicles for drug delivery applications e.g. polymers [2], polymeric micelles [3], liposomes [4], magnetic nanoparticles [5] and mesoporous silica [6]. Amongst the wide spectrum of carriers, mesoporous silica holds a unique position due to its potential applications in the field of catalysis [7], protein encapsulation [8], myoglobin separation [9], in addition to drug and gene delivery [10,11]. The rising interest towards mesoporous ⇑ Corresponding authors. Fax: +91 3222 282252. E-mail address: [email protected] (D. Dhara). http://dx.doi.org/10.1016/j.jcis.2014.06.003 0021-9797/Ó 2014 Elsevier Inc. All rights reserved.

silica as drug delivery vehicles is a consequence of their high surface area, well-defined pore structures with large pore volume, tunable particle size, easily modifiable surface (due to presence of numerous silanol groups), besides their excellent water dispersibility, biocompatibility, and high biomolecules loading capacity [12,13]. Magnetic resonance imaging (MRI) has emerged as one of the best non-invasive imaging techniques that has necessitated the designing and development of robust and biocompatible nanoparticles as efficient MR contrast agent with much higher transverse relaxivities [14]. The combination of mesoporous silica with magnetic nanoparticles and other functional molecules culminating in new types of hybrid nanocarrier platforms for cell imaging, diagnosis and therapy, has gained tremendous research interest among scientists in the last few years [15–20]. Magnetic nanoparticles (MNPs) have been explored for numerous biomedical applications

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such as magnetofection for facile gene delivery [21], MRI contrast agents [22,23], to induce local hyperthermia in response to an external alternating magnetic field, to selectively destroy cancer cells [24] and for magnetically targeted carrier system in drug delivery applications [25]. In fact, superparamagnetic iron oxide nanoparticles have emerged as an efficient MRI contrast agent compared to gadolinium diethylene triaminopentaacetic acid (Gd-DTPA), due to their biocompatibility, good blood retention and increased contrast enhancement [26]. Recent studies have revealed that metal doped mixed ferrite, such as MnFe2O4 nanoparticles, possessed much higher magnetization values than magnetite and other mixed ferrite nanoparticles [27,28] So far, many reports have described MnFe2O4 nanoparticles as an efficient and the strongest MR image-contrast agent with much higher T2 relaxivity [29–31]. However, most of the MnFe2O4 nanoparticles have been synthesized in organic solvents, which results in poor aqueous dispersion stability of the particles, making them unsuitable for biological applications. Thus, the preparation of water dispersible and superparamgnetic MnFe2O4 nanoparticles is important for their biological usefulness. Meanwhile, magnetic cores with mesoporous silica shell attached with fluorescent moeities have been continuously explored for diverse applications, with most of them consisting of iron oxide nanoparticles as central core. In the present work, we have prepared a new nanocomposite system consisting of porous silica-encapsulated MnFe2O4 nanoparticles and investigated their efficacy as anticancer drug-carriers and as magnetic resonance imaging agent. In this work, MnFe2O4 nanoparticles of size 100–150 nm were prepared by solvothermal method and coated with silica through the hydrolysis of tetraethylorthosilicate (TEOS). Amine functionalities were introduced by co-condensation with (3-aminopropyl)triethoxysilane (APTES) in presence of poredirecting agent cetyl trimethyl ammonium bromide (CTAB). In the following step, CTAB was removed to produce mesoporous silica-encapsulated MnFe2O4 nanoparticles (NH2-MSN). The aminefunctionalized hybrid nanoparticles were further conjugated with folic acid (FA) for cancer-cell targeting and successively with RITC that endowed fluorescent properties to them. These so-prepared M-MSN were thoroughly characterized at each step of preparation using various techniques such as viz. XRD, HRTEM, FESEM, FTIR, VSM, XPS, BET surface area measurement, and UV–Vis spectroscopy. Finally, M-MSN were loaded with anticancer drug Doxorubicin (DOX) and their effectiveness towards targeted drug delivery was investigated through in vitro cytotoxicity (MTT assay), fluorescence microscopy, apoptosis study and in vivo histopathological analysis in mice-model system. These M-MSN were further evaluated for their effectiveness in MR imaging application. 2. Materials and methods 2.1. Materials Anhydrous ferric chloride (FeCl3), manganese chloride (MnCl2 4H2O), cetyl trimethyl ammonium bromide (CTAB), folic acid (FA), ammonia, concentrated HCl, dimethyl sulphoxide (DMSO) and ethanol were purchased from Merck, Germany. N-hydroxysuccinamide (NHS), 1-[3-(dimethylamino)propyl]-3-ethylcarbodiimide hydrochloride (EDC) were purchased from Spectrochem. Tetraethyl orthosilicate (TEOS), (3-aminopropyl)triethoxysilane (APTES), and doxorubicin hydrochloride (DOX), rhodamine b isothiocyanate (RITC), trinitrobenzene sulfonic acid (TNBS), 40 -6diamidino-2-phenylindole (DAPI), propidium iodide (PI), RNase and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) were obtained from Sigma–Aldrich Chemicals, USA. Fetal bovine serum and Minimum Essential Medium (MEM) were obtained from Hyclone, USA and Himedia, India, respectively. All

the chemicals were of reagent grade and used without further purification. 2.2. Methods 2.2.1. Synthesis of MnFe2O4 nanoparticles (MN, Scheme 1) Magnetic MnFe2O4 nanoparticles were prepared by solvothermal technique as reported earlier with some modification [32]. For the synthesis of MnFe2O4 nanoparticles, a mixture of MnCl2 (0.416 g, 2.1 mmol), FeCl3 (0.683 g, 4.2 mmol), and sodium acetate (3.0 g) were dissolved in 30 ml ethylene glycol and stirred vigorously for 1 h at room temperature to obtain a homogenous solution. This solution was then transferred to a Teflon-lined stainless steel autoclave (90.0 ml capacity), sealed, and heated to 160 °C for 24 h. After cooling the autoclave to room temperature, the MnFe2O4 nanoparticles were repeatedly washed with ethanol and distilled water, and then dried under vacuum at 60 °C for 12 h. 2.2.2. Preparation of amine-functionalized mesoporous silica coated MnFe2O4 nanoparticles (NH2-MSN, Scheme 1) Amine functionalized mesoporous silica coated MnFe2O4 nanoparticles (NH2-MSN) were prepared by reported procedure with some modifications [33]. 0.1 g of MnFe2O4 nanoparticles were first ultrasonicated with 50 ml of 0.1 M HCl aqueous solution for 10 min. These magnetic nanoparticles were then separated, washed with deionized water and homogeneously dispersed in a mixture of 80 ml ethanol, 20 ml deionized water and 1 ml concentrated ammonia aqueous solution (28%) for 30 min. To the above dispersed mixture, 0.03 g tetraethyl orthosilicate (TEOS) was added dropwise to get silica modified nanoparticles. The above solution was stirred at room temperature for 6 h. Thereafter, the product was separated and washed with ethanol and deionized water. The above silica coated nanoparticles (SiO2-MN) were homogenously dispersed in a solution containing 0.3 g cetyl trimethyl ammonium bromide (CTAB), 80 ml deionized water, 70 ml absolute ethanol and 1.2 ml concentrated ammonia solution (28%) for 30 min. To the above dispersion was added 0.3 g TEOS and then 0.5 mL APTES, the reaction mixture was stirred continuously for 12 h. Thereafter, the product was recovered by magnetic decantation and repeatedly washed with ethanol and deionized water. In order to remove the pore-directing agent CTAB, the above nanoparticles were dispersed in ethanolic solution containing 0.3 g NH4NO3 and the mixture was stirred at 60 °C and the process was repeated thrice. The products were recovered by magnetic separation, washed with ethanol followed by drying at 60 °C. 2.2.3. Folic acid conjugation on NH2-MSN (FA-MSN, Scheme 1) Folate targeted magnetic mesoporous silica nanoparticles were prepared by two-step chemical route according to our reported procedure [34]. At first, the carboxylic group of folic acid (FA) was activated using standard EDC/NHS method. 0.05 g FA was dissolved in 1:1 (v/v) water – DMSO mixture. To this, EDC and NHS were added to activate the carboxylic groups of FA and the pH of the resulting solution was made 7.0–8.0 with 0.1 M NaOH. The FA activation reaction was continued for 4 h under dark condition at room temperature. To the activated FA solution, 20 mg (1 mg/ ml) of aqueous dispersed NH2-MMSN suspension was added and the resultant solution was stirred at 25 °C for 12 h in the dark. The folate-conjugated nanoparticles were recovered through centrifugation, washed 4–5 times with DMSO and water, and finally dried under vacuum at room temperature for 24 h. 2.2.4. Preparation of RITC labeled FA-MSN (RITC-FA-MSN, Scheme 1) and RITC labeled MSN (RITC-MSN, Scheme 1) A highly fluorescent moeity, RITC was covalently linked to the residual amine groups of the FA-MSN to form magneto-fluorescent

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nanoparticles that help to detect their internalization into cells. For this, 1 mg RITC was dissolved in 2 ml of in 1:1 volume ratio of DMSO-water, and pH of this solution was adjusted to 8.0 with dilute NaOH. Then, an aqueous suspension of 10 mg of functionalized FA-MSN was added and stirred for 12 h in dark. The RITC labeled FA-MSN (RITC-FA-MSN) was then washed in water repeatedly through ultrasonication in order to remove any physically adsorbed RITC molecules on nanoparticles. Finally, the RITC-FAMSN was recovered and suspended in PBS buffer for further studies. Similarly, RITC-MSN was also prepared by covalent linking of RITC to NH2-MSN (without FA) and preserved in a refrigerator as a control for further studies.

2.2.5. Nanoparticle characterization The formation of MnFe2O4 nanoparticles and MSN were assessed by Phillips PW 1710 X-ray diffractometer (XRD) with Ni-filtered Cu Ka radiation (k = 1.54 Å). The presence of amorphous silica was analyzed from low angle XRD pattern in the range of (2h = 0.5– 8°). The surface composition of the nanoparticles at each step of surface modification was determined from FTIR spectra. Samples for FTIR spectra were prepared in KBr in the range 400– 4000 cm1. Surface functional groups and composition of FA-MSN were further confirmed from XPS analysis using Al Ka excitation source in ESCA-2000 Multilab apparatus (VG microtech) with a model Nexus-870, Thermo Nicolet Corporation, Wisconsin, USA. The size and shape of the individual nanoparticles were observed by high-resolution transmission electron microscopy (HRTEM, JEOL 3010, Japan) operated at 300 kV. For HRTEM study, the nanoparticles were thoroughly dispersed in water by ultrasonication. A drop of the solution was placed on a carbon coated copper grid, air-dried and the images were taken. The surface morphology of the nanoparticles was analyzed by field emission scanning electron microscopy (FESEM) with Phillips CM 200 microscope. The average particle size from HRTEM and FESEM micrographs was analyzed using image J software. The conjugation of folic acid on NH2-MSN was revealed by UV–Vis spectrophotometric analysis. The surface area, pore size and pore distribution were determined by using a N2 adsorption–desorption instrument (Quantachrome Corporation, Quantachrome Autosorb Automated Gas Sorption System).

2.2.6. Drug loading and release study 10 mg of FA-MSN were incubated with 10 ml of DOX aqueous solution (0.18 mg mL1) and the mixture was kept on a shaker for 48 h under dark conditions. After 48 h, the DOX-loaded nancomposites (DOX-MSN) were recovered by magnetic separation followed by centrifugation. DOX-MSN were washed twice to remove unbound drug molecules. After loading, the supernatant and the washings were mixed and its absorbance noted at 481 nm. Thereafter, the amount of DOX was loaded from the decrease in the absorbance value with respect to that of the original DOX solution. The drug loading content and drug encapsulation efficiency (EE) was determined from following formula

Drug loading contents ð%Þ ¼

Weight of drug in nanoparticles  100 Weight of nanoparticles taken

Drug entrapment efficiency ð%Þ ¼

Weight of drug in nanoparticles  100 Weight of drug injected

The release of DOX from DOX-MSN was carried out both at physiological pH (7.4) and at lysosomal pH of cancer cells (5.6) at 25 °C. The amount of DOX released was monitored spectrophotometrically at 481 nm and the amount of the released drug was calculated from a standard curve of free DOX solution.

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2.2.7. Cell lines and cytotoxicity assay The cytotoxicity was evaluated with standard MTT (3-(4,5dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay. For in vitro cytotoxicity experiments, two types of cells namely human cervical adeno carcinoma (HeLa) and normal fibroblast (L929) cell lines were acquired from the National Centre for Cell Sciences (NCCS), Pune, India. The cells were cultured in Minimum Essential Medium (MEM) or Dulbecco’s Modified Eagle’s medium (DMEM) supplemented with 10% fetal bovine serum, penicillin (100 units/ml), streptomycin (100 mg/ml), and 4 mM l-glutamine at 37 °C in tissue culture flasks with 5% CO2 and 95% air humidified atmosphere. For experimental purpose, trypsinized cells were adjusted to a concentration of 1  105 cells/ml and plated in a 96 well flat bottom culture plates (180 ll/well). For biocompatible studies, cells were then incubated with different concentrations of pristine MnFe2O4 nanoparticles (MN), FA-MSN, and DOX-MSN at 37 °C in a humidified 5% CO2 incubator for 24 h. The percentage of cell viability was finally estimated by MTT assay. 2.2.8. Intracellular uptake studies The internalization of nanoparticles into cells was observed by fluorescence microscope imaging technique. For intracellular uptake studies, 10 lg/ml RITC labeled FA-MSN were incubated in HeLa and L929 cells for 1 h, 2 h and 4 h respectively in order to determine the time-dependent uptake of NPs inside the HeLa cells. Again, the receptor specificity of FA for the HeLa cells was checked by incubating RITC-MSN and RITC-FA-MSN for 4 h. After incubation, the cells were fixed with 4% paraformaldehyde for 15 min and stained with DAPI (1 mg/ml) for 10 min at 37 °C. Thereafter, cells were washed with PBS thrice to remove any non-internalized nanoparticles from the medium. Finally, the above-incubated cells were observed under Olympus IX 70 fluorescence microscopy. 2.2.9. DAPI staining for nuclear morphology study After incubation with FA-MSN and DOX-MSN, the morphology of the HeLa cells was visualized by staining the cells with DAPI. HeLa cells were treated with FA-MSN (control nanoparticles) as well as 5, 10, 15, and 25 lg mL1 DOX-MSN for 24 h at 37 °C. The cells were fixed with 4% formaldehyde for 15 min, permeabilized with 0.1% Triton X-100 and stained with 1 mg/ml DAPI for 10 min. The cells were then rinsed with PBS and examined under fluorescence microscopy (Olympus IX 70). Apoptotic cells were detected by the hypochromic subdiploid staining analysis. 2.2.10. In vivo cytotoxicity studies through histopathology experiment In vivo toxicity studies of the FA-MSN in mice model system were conducted according to the norms of local ethics committee. For in vivo toxicity study, two groups of healthy swiss albino mice (each group consisting of five males and five females) weighing 20–22 g were taken. FA-MSN were suspended in millipore water and sonicated for 20 min. 100 ll of aqueous dispersed FA-MSN (300 lg mL1) were intravenously injected through the tail vein of the mice. After 7 days of treatment, these mice were sacrificed and their major organs such as spleen, liver, kidney, stomach, heart and brain were dissected for histopathology analysis. The small piece of tissue sections of these organs were removed and then fixed with 10% formalin solution, embedded in paraffin and stained with hematoxylin and eosin (H&E), according to standard clinical pathology technique. The histological sections were observed under light microscope. One control set (mice treated with millipore water only) of experiments was performed for comparison with the above nanoparticle-treated mice. 2.2.11. In vitro cellular MR imaging studies of FA-MSN Samples for MR phantom imaging were prepared by incubating 106 numbers of cells with different nanoparticles (FA-MSN and

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NH2-MSN) of concentrations 0.01–0.05 mg/mL in 12 well plates for 4 h. Cells were then washed with PBS for three times and fixed with paraformaldehyde. After that, 5 ml of 2% low-melting agarose gel was added to each well containing nanoparticle-internalized cells in order to avoid air susceptibility artifacts. Cell suspensions were allowed to solidify at 4 °C. MRI was performed keeping the samples under a 3 T clinical MRI scanner (GE Medical systems, Milwaukee) using a pre-fabricated sample holder. A spin-echo multisection pulse sequence was applied to acquire the MR phantom images. In order to determine the transverse relaxation (T2) for each sample, coronal images were acquired by variable echo times (TE) of 13.3–212.8 ms with repetition time (TR) of 1770 ms. Other parameters were set as follows: an acquisition matrix of 256 mm  256 mm, field of view of 240 mm  240 mm, section thickness of 8 mm. The MRI signal intensity (SI) and visualization of the phantom images were analyzed using the standard software provided by the manufacturer. Transverse relaxivity (T2) values were calculated by plotting the SI of each sample for a range of TE values [35]. Transverse relaxation rates R2 (1/T2) were calculated from the data acquired from MRI instrument and then plotted against concentration of the nanoparticles in the gels. The slope of the line gives the relaxation time and relaxivity value.

3. Results and discussion 3.1. Preparation of multifunctional magnetic mesoporous silica nanoparticles (M-MSN) Multifunctional MnFe2O4 nanoparticles encompassing the superparamagnetic properties, porosity, targeting and imaging ability in a single entity was designed as a potential carrier for therapeutics and as a magnetic resonance (MR) imaging probe. The superparamagnetic nature of the said nanoparticles renders them suitable as a probe for MR based diagnosis to locate the affected organ or tissue prior to the delivery of a chemotherapeutic agent. The detailed procedure for preparation of M-MSN is presented in Scheme 1. Aqueous dispersible MnFe2O4 nanoparticles (MN) of diameter 100–150 nm were synthesized through solvothermal route using ethylene glycol as reducing agent. Further surface functionalization of the MN involved their modification with TEOS to produce silica coated MnFe2O4 nanoparticles (SiO2-MSN). In the next step, APTES was used as silane precursor to introduce amine functionalities on the nanoparticles, and CTAB was used as the pore-forming agent. Subsequently, CTAB was removed from amine functionalized mesoporous silica-coated magnetic MnFe2O4 nanoparticles (NH2-MSN). The present approach demonstrates a relatively simple in situ method for the formation of porous and amine functionalized silica nanoparticles as compared to multistep modification procedure reported earlier [33,36,37]. Thereafter, folic acid (for cancer-specific targeting) was linked with a part of the amine groups on NH2-MSN through formation of amide bond. The fluorescent moiety, RITC, was then conjugated to the nanoparticles to provide them with fluorescent properties. Finally, doxorubicin (DOX), an anticancer drug, was loaded into the porous nanoparticles and successively, the drug release study was carried out under two different pH conditions The DOX loaded nanoparticles were tested on both cancer and normal cell lines. The surface chemistry and conjugation of different functional moieties were elucidated from FTIR spectra. FTIR spectra of MnFe2O4 nanoparticles (MN), NH2-MSN, FA-MSN are shown in Fig. 1. As shown in FTIR spectrum of free MnFe2O4 nanoparticles, the peaks at 3400 cm1 and 585 cm1 corresponds to O–H bond vibration and M–O bond vibration of the magnetic Mn-ferrite core respectively. In the spectrum of NH2-MSN, peaks at 1057 cm1,

1647 cm1 and 1458 cm1 corresponds to Si–O–Si, N–H bending and C–N stretching vibrations respectively suggesting a successful grafting of APTES in a single, in situ approach. Additionally, two peaks at 2936 cm1 (asymmetric CH2 stretching) and 2856 cm1 (symmetric CH2 stretching) were seen (Fig. 1b) that may be attributed to the presence of APTES on the nanoparticles. Further modification by folic acid led to disappearance of the peak at 1647 cm1 corresponding to the amine group bending, while the appearance of two new peaks at 1654 cm1 and 1595 cm1 correspond to amide bond formation. Conjugation of FA on the NH2-MSN was also confirmed from simple UV–Vis spectroscopic analysis, as shown in Fig. S1 (electronic supporting information, ESI). Pure FA exhibited two bands at 280 nm and 360 nm owing to the p–pand n–p transitions respectively, corresponding to the enone moeity of FA molecule [38]. For NH2-MSN, no detectable peaks were observed (shown in Fig. S1a), whereas FA-MSN revealed two peaks corresponding to free FA, suggesting the attachment of FA molecules to the NH2-MSN. 3.2. Physiochemical properties of the synthesized nanoparticles 3.2.1. Wide-angle and low-angle XRD study The successful formation of pristine MnFe2O4 nanoparticles (MN) and NH2-MSN was confirmed from XRD analysis. The wideangle and low-angle XRD patterns are shown in Fig. 2. The XRD patterns showed that all the peaks of the pristine nanoparticles matched well with the standard XRD peaks in MnFe2O4 (JCPDS standard data, Card No. 10-0319). In addition to the abovementioned peaks, the NH2-MSN exhibited a broad peak between 20° and 30° indicating the presence of silica in the nanoparticles. The small-angle X-ray diffraction patterns of NH2-MSN (Fig. 2b) clearly showed a distinct peak at 2h 1.8–2.8° suggesting the presence of amorphous silica in the composites. The well-resolved diffraction peaks at 2.4° and a small peak at 3.4° correspond to (1 0 0) and (1 1 0) plane of amorphous silica shell. respectively, revealing the two-dimensional hexagonal mesoporous structure of the nanoparticles [39]. 3.2.2. XPS analysis XPS is a highly useful technique for the determination of elements present in a compound, besides their oxidation state and chemical composition. The XPS spectra for the FA-MSN (Fig 3a) revealed the spectrum for Fe 2p in the nanoparticles and two peaks are obtained for Fe at 710.516 eV and 724.081 eV for Fe 2p3/2 and Fe 2p1/2 respectively that is consistent with the literature values [40]. The peaks are obtained in the ratio of 2:1. Fig. 3b reveals the high-resolution XPS spectra for Mn 2p. For Mn 2p, two peaks are assigned to 639.677 eV and 653.629 eV with a satellite peak at 646.087 eV for Mn 2p3/2, Mn 2p1/2 and Mn2+ respectively. The peaks matched well with the standard XPS peaks of MnFe2O4 nanoparticles described in literature [41]. Hence, from XPS analysis, it is evident that the core of the FA-MSN is indeed composed of MnFe2O4. Further analysis revealed the Si peak at 152.549 eV corresponding to Si 2s (shown in inset of Fig. 3c). Taking account of C 1s peak binding energy at 284.45, all other peaks were deconvoluted accordingly. The Si 2p (Shown in Fig 3c.) can be deconvoluted into peaks at binding energy values of 102.67 eV and 104.7 eV for Si–C and Si–O bonds respectively, in agreement with literature [42]. Additionally, C 1s peak can be deconvoluted into 4 peaks (Fig. 3d) corresponding to the binding energies values of 284.45 eV, 285.54 eV, 286.415, eV and 292.384 eV for C–C, C–O, C–O–C, and NHCO respectively, in accordance to the presence of O atom in four different environments. Since the FA molecule was attached to NH2-MSN via amide bond, so the peak for amide bond unequivocally confirmed the conjugation of folic acid to NH2-MSN. Three distinct peaks for O 1s element (Fig. 3e) were

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Scheme 1. Schematic illustration of the steps for the fabrication of multifunctional mesoporous silica-coated superparamagnetic manganese ferrite (MnFe2O4) nanoparticles (M-MSN).

observed at binding energy values of 529.536 eV, 532.09 eV, and 532.28 eV corresponding to M–O, C–O and NHCO respectively. The M–O peak corroborates the presence of MnFe2O4 in the composition. Fig. 3f shows two peaks for deconvoluted spectra of N 1s element, one at 399. 5 eV for –NH2 groups and another peak at 400.5 eV for –NHCO group, suggesting the presence of both amine and amide groups in the nanoparticles.

distributions (Fig. 4b) for NH2-MSN show unequal pore size distribution in the range 2 nm - 50 nm, with the maximum around 4–5 nm. The average pore volume and pore diameter of NH2-MSN were found to be 0.197 cm3/g and 2.1 nm respectively. Such formation of silica layer with multiple pores increased the surface area of the resultant nanoparticles manifold, which makes it useful for carrier of drug molecules.

3.2.3. BET Measurements The porous nature of the prepared NH2-MSN was confirmed from nitrogen adsorption–desorption isotherms plot. The Brunauer–Emmett–Teller (BET) surface areas and BJH pore size distribution for NH2-MSN are shown in the Fig. 4. The BET curve for NH2-MSN exhibited type-IV adsorption–desorption isotherm pattern according to IUPAC nomenclature for porous samples. For the isotherm patterns, the leap start at relative pressure of 0.15 clearly indicated the presence of pores in the nanoparticles. The BET surface area for NH2-MSN was found to be 372 m2/g. BJH pore

3.2.4. Morphology and size of the nanoparticles The porous nanostructure was also observed from TEM analysis, as shown in Fig. 5. MnFe2O4 nanoparticle cores were found to be almost spherical in shape, having size in the range of 100– 150 nm (Fig 5a). The average size of NH2-MSN was found to be 200–300 nm, with of an outer porous silica shell of thickness 20–25 nm and inner black magnetic core. TEM image of one nanoparticle in high magnification is displayed in the inset of Fig. 5b. The selected area diffraction pattern (SAED) and EDX for NH2-MSN are given in ESI (Fig. S2). The SAED pattern suggests

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the polycrystalline nature of the pure MnFe2O4 nanoparticles (MN) and their corresponding silica-coated nanoparticles. The energy dispersive X-ray spectroscopy (EDX) confirmed the presence of Mn, Fe, O, Si, C and N elements in the sample, which corroborated with the elemental composition obtained from XPS analysis. The FESEM image of NH2-MSN is shown in Fig. 6 that indicated spherical morphology of the prepared nanoparticles with an average size of 200–300 nm. After the preparation of MN by solvothermal method, the individual small sized nanocrystals aggregate to form a big nanocluster in order to reduce the interfacial energy between the individual nanocrystals [43].

Fig. 1. FTIR spectra of (a) MN, (b) NH2-MSN, and (c) FA-MSN.

3.2.5. Magnetic measurements Retention of magnetic properties and achievement of higher magnetization values is of utmost importance for application in MRI studies. The magnetic nature of MN, NH2-MSN and FA-MSN were analyzed by VSM-SQUID measurements, as presented in Fig. 7. The saturated magnetization value (Ms) for pristine

Fig. 2. (a) Wide angle XRD pattern of MN, SiO2-MN, NH2-MSN and (b) low angle XRD pattern of NH2-MSN.

Fig. 3. High resolution XPS spectra of FA-MSN showing the binding energy of different elements (a) Fe 2p, (b) Mn 2p, (c) Si 2p (Si 2s in inset), (d) C 1s, (e) O 1s, and (f) N 1s.

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Fig. 4. (a) Nitrogen adsorption–desorption isotherm pattern and (b) BJH pore distribution of NH2-MSN.

Fig. 5. TEM image (a) MN and (b) NH2-MSN.

Fig. 6. FESEM image of NH2-MSN. Fig. 7. VSM curve for (a) MN, (b) NH2-MSN, and (c) FA-MSN.

MnFe2O4 nanoparticles was 95–99 emu/g which was much higher compared to magnetic iron-oxide nanoparticles (60– 65 emu/g). Enhancement in the magnetization value of Fe3O4 crystal lattice due to the incorporation of Mn2+ was also observed in this study [44,45]. MnFe2O4 nanoparticles and NH2-MSN showed no coercivity and both the curves showed superparamagnetic behavior at room temperature. No hystersis loop and remanence for MN and NH2-MSN were observed, supporting the superparamagnetic properties of both the materials. The Ms values for NH2-MSN was 73–75 emu/g which was lower compared to the free nanoparticles (99 emu/g). The silica coating on the nanoparticles lowered the magnetization value. Further folate modification

on NH2-MSN reduced the Ms value to 60–63 emu/g. Nevertheless, the retention of higher Ms value of the nanoparticles facilitates them to serve as a good MRI contrast agent. 3.3. Drug loading and release study The doxorubicin (DOX) loading and entrapment efficiency were evaluated using standard calibration curve obtained for a series of DOX solution. The DOX loading content and encapsulation efficiency for FA-MSN were calculated to be 11.3% and 60.4%. We believe that the loading of DOX in the nanoparticles occurred

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through physical interactions. Fig. 8 presents the drug release profile of DOX from DOX-MSN. It can be clearly seen that the DOXMSN released DOX more favorably at lysosomal pH condition as compared to physiological pH. The initial burst release followed by slow release of DOX was observed at both the pH values. The drug release percentage increased with time and finally became constant after certain time interval. The drug release rate is slow as it is evident from the drug-release profile, as presented in Fig. 8. The drug release rate was found to depend on the pore size of the MSN. Larger the pore size, faster the release of drug takes place. The slow drug release at both pH conditions (pH 7.4 and pH 5.6) from DOX-MSN is possibly due to the small pore size of MSN. Nonetheless, the fact that a higher amount of DOX was released at pH 5.6, in comparison to that at pH 7.4, makes this system potentially useful. At low pH (pH 5.6), DOX molecules got protonated and were released into the aqueous medium. The cumulative and higher drug release rate at lysosomal pH condition of cancer cells (pH 5.6) increased the efficacy of destroying the cancer cells, which indeed needed for drug delivery applications. 3.4. Biological assay 3.4.1. MTT assay The in vitro cytotoxicity of MN and FA-MSN was tested on both HeLa cancerous cells lines and normal L929 cell lines, as shown in Fig 9. As observed by MTT assay, both types of cells were about 95% viable after incubation with MN and FA-MSN up to concentrations of 1–20 lg mL1. This confirmed that the nanoparticles were nontoxic to both the cells. On the other hand, DOX-MSN induced toxicity on both the cells types due to the action of the loaded DOX as shown in Fig 9c. However, DOX loaded nanoparticles caused more toxicity to the HeLa cells than the L929 cells. 15 lg mL1 of DOX-MSN induced 11% death of the L929 cells whereas the same amount of DOX-MSN caused 35% death of the HeLa cells. Again, upon treatment with the maximum studied dose of DOX-MSN (50 lg mL1), 65% HeLa cell death was observed in contrast to only 20% L929 cells cell death. Therefore, from cell viability assay it can be elucidated that DOX-MSN induced more toxicity to cancer cells than normal fibroblast cells. As folate receptors are present in higher number on the surface of the HeLa cells than the normal L929 cells, higher amounts of FA-MSN selectively entered into the HeLa cancer cells than normal L929 cells, as confirmed from intracellular uptake behavior (shown in supporting figure, Fig. S5). Hence, there is enhanced cytotoxicity of DOXMSN to HeLa cells than normal fibroblast cells.

3.4.1.1. Inracellular uptake by fluorescence microscopy analysis. The cellular internalization of MSN was checked by fluorescence microscopy analysis. The uptake efficiency of folate-conjugated nanoparticles (RITC-FA-MSN) was compared with nanoparticles having no FA (RITC-MSN). As shown by the uptake images in Fig. 10, the time dependent uptake of RITC-FA-MSN into the HeLa cells revealed that the nanoparticles gradually internalized into cells presumably through receptor mediated endocytosis pathway and produced bright red fluorescence, as observed by fluorescence microscopy. The fluorescent signal intensity was increased 2–3fold with increase in time of incubation of nanoparticles with cells. The cellular uptake of FA-MSN into L929 cells was compared with the uptake into HeLa cells and it was found that the uptake in normal cells was significantly less in comparison with HeLa cells (see ESI, Fig. S3). Further, the targeting efficacy of the RITC-FA-MSN towards the HeLa cells was compared to RITC-MSN (control MSN without FA) by measuring the extent of uptake (see ESI, Fig. S4). The internalization of MSN into HeLa cancer cells was more for RITC-FA-MSN as compared to RITC-MSN. It is known that folic acid has a high affinity to bind with folate receptors that are overexpressed in several human tumors including kidney, ovarian, and endometrial cancer, and serve as promising targeting agent in biomedical applications [46,47]. Due to receptor specificity of the targeting moeity on the nanocarriers, the cellular internalization of FA conjugated MSN in HeLa cells was enhanced as compared to normal cells, which in turn is expected to reduced the undesired side-effects of the anticancer drug to the normal cells. 3.4.2. Nanoparticles induced apoptosis study It has been reported that DOX causes malfunctioning of the mitochondria by non-specific oxidative damage to the outer and the inner membranes, or by direct interaction with the mitochondrial DNA or enzymes involved in cell respiration [48]. To demonstrate the efficacy of DOX-MSN towards cancer cells, HeLa cells were cultured with 10 lg mL1 FA-MSN (control set), and with 5, 10, 15, 25 lg mL1 of DOX-MSN. The apoptosis images are shown in Fig. 11. It was observed that apoptosis, which is a significant feature of a DNA fragment, was absent in the control set (FA-MSN, Fig. 11), as evident from the nuclear morphology (shown by yellow colored arrow) that remained intact. On the other hand, significant DNA cleavage of the HeLa cells has been noticed on treatment with DOX-MSN, as visible from the nuclei fragments with apoptotic nuclei (Fig. 11b–e) in a dose dependent manner. The formation of these apoptotic bodies was observed in a dose-dependent manner, with no such morphology changes observed in the control cells. The percentage of these cells exhibiting apoptosis have been counted, calculated and reported in ESI (Fig. S5). This confirms that DOX-MSN induced apoptosis of HeLa cells. 3.5. MRI study

Fig. 8. DOX release profile at different pH from DOX-MSN.

To test the applicability of these MSN as MRI contrast agent, an in vitro MRI study has been carried out for NH2-MSN and FA-MSN in HeLa cells. Fig. 12 represents phantom image of the nanoparticles after internalization into HeLa cells. A concentration dependent change in signal intensities was observed from MRI images. As the concentration increased, there was gradual increase in signal darkening. Again, the FA-MSN exhibited significantly more darkening of HeLa cells compared to NH2-MSN indicating FA-MSN as a good MRI contrast agent. Relaxivity plot (R2 = 1/T2) (ESI Fig. S6) indicates linear relationship with nanoparticle concentration in both the cases i.e., NH2-MSN and FA-MSN. Due to receptor specificity of FA-MSN towards cancer cells, larger amount of FA-MSN accumulated inside HeLa cells as compared to NH2-MSN and resulted in decreased relaxation time (increasing the relaxivity) [35]. Superparamagnetic nanoparticles darkened the tissue by

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Fig. 9. MTT assay of (a) MN, (b) FA-MSN, and (c) DOX-MSN treated on HeLa cells and L929 cells.

Fig. 10. Fluorescence microscopy images of HeLa cells after (a) 1 h, (b) 2 h and (c) 4 h of incubation with RITC-FA-MSN.

Fig. 11. Apoptosis study of HeLa cells (a) treated with nanoparticles without DOX (FA-MSN); treated with nanoparticles containing DOX (DOX-MSN), (b) 5 lg/ml and (c) 10 lg/ml, (d) 15 lg/ml, and (e) 25 lg/ml.

decreasing the relaxation time of the water protons of the specific tissue. Folate conjugated nanoparticles improved the intracellular uptake manifold as revealed by MRI signal intensities (shown in phantom image). In addition to drug delivery, FA-MSN caused more darkening of HeLa cells than only MSN (without folate).

Due to receptor-mediated endocytosis, FA-MSN penetrated into the HeLa cells easily and produced more dark images than nanoparticles those do not contain folic acid. Due to the cancer specific targeting behavior of FA-MSN to the HeLa cells, it decrease T2 relaxation time with good contrast ability.

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Fig. 12. MRI phatom image of HeLa cells treated with FA-MSN (left panel) and (b) NH2-MSN (right panel).

of the prepared MSN towards the major organs of a mice model was assessed from histopathology analysis to check whether FAMSN caused any sort of tissue inflammations and damage. For this purpose, aqueous dispersed solution of FA-MSN was injected through tail vein of mice and these mice were sacrificed after seven days of post treatment. One control set (mice treated with only water) was analyzed in similar manner as with FA-MSN treated mice. The mice were healthy after intravenous injection of FAMSN and exhibited regular body weight gain similar to the control mice. The histological sections of the major organs of the treated mice and the control mice are presented in Fig. 13. As from the figures, no apparent toxicity was observed for the treated mice, and the tissue sections of different organs were found to be normal, similar to the control one. The histological sections of other major organs such as liver, spleen, and stomach of both FA-MSN treated mice and control mice are presented in ESI (Fig. S7). Liver and spleen were the target organs during intravenous injection due to abundant blood supply and presence of RES within these organs. However, no acute inflammation, splenic disorder or any damage or distortions in the liver tissues were noted. All the other major organs such as heart, lung, brain, and kidney were normal with respect to control. Hence, in vivo biological study suggested that the mesoporous silica encapsulated MnFe2O4 were biocompatible in nature and could well be used for establishing a successful targeted drug delivery system.

4. Conclusions Multifunctional mesoporous silica-coated superparamagnetic manganese ferrite (MnFe2O4) nanoparticles attached with fluorescent and targeting moieties was prepared by a facile method. These were found to be potentially capable of drug delivery specifically to cancer cells, cancer cell imaging and useful as MRI contrast agent in diagnosis of cancer. HRTEM images clearly showed magnetic nanoparticles core coated with mesoporous shell of thickness 20–25 nm with the overall dimension of the resulting nanoparticles 200–300 nm which is in desired size-range for drug delivery application. It was shown that these nanoparticles were taken up specifically by HeLa cancer cells in comparison to normal cells. Drug loading studies indicated that the outer mesoporous silica shell encapsulated significant amount of anticancer drug, DOX, and the release of DOX preferably at lysosomal pH. In vitro biological studies revealed that the DOX-loaded folate-targeted nanoparticles achieved excellent efficacy for simultaneous targeting and destroying cancer cells. Their ability to specifically accumulate in cancer cells along with their supepararmagnetic behavior make them useful as magnetic resonance contrast agent. In vivo studies in mice-model system confirmed the biocompatibility of the nanoparticles. From all the biological studies, it is envisioned that these M-MSN synthesized by us is an excellent candidate that may serve as a vehicle in cancer-specific targeting, imaging and MRI application in a single entity.

Acknowledgments

Fig. 13. Histopathological analysis of major organs such as brain, kidney, lung, heart of control mice (left panel), and mice treated with FA-MSN (right panel). FAMSN were intravenously injected into mice and major organs of mice were dissected for analysis after 7 days of post injection.

3.6. In vivo toxicity assay Introduction of foreign particle with toxic effect may alter the normal and regular morphology of the major organs. The toxicity

Financial support from the Department of Science and Technology, Government of India, New Delhi is acknowledged. B. Sahoo and K. Sanjana P. Devi acknowledge CSIR and DBT, New Delhi respectively for their research fellowships. Authors are grateful to Shouvik Mitra, Prasun Patra and Prof. Arunava Goswami of AERU, Biological sciences division, Indian Statistical Institute, Kolkata for their assistance with in vivo toxicity assay; department of Physics, IIT Kharagpur for XPS analysis; Prof. Biswajit Chowdhury of department of Applied Chemistry, ISM Dhanbad for BET surface

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area measurement; Dr. S.K. Sharma of Eko MRI Center, Kolkata for executing MRI studies. Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.jcis.2014.06.003. References [1] Y. Liu, H. Miyoshi, M. Nakamura, Colloids Surf. B 58 (2007) 180–187. [2] J. Chen, J. Ouyang, J. Kong, W. Zhong, M.M. Xing, ACS Appl. Mater. Interf. 5 (2013) 3108–3117. [3] J. Gong, M. Chen, Y. Zheng, S. Wang, Y. Wang, J. Controlled Release 159 (2012) 312–323. [4] T. Wang, S. Yang, V.A. Petrenko, V.P. Torchilin, Mol. Pharm. 7 (2010) 1149– 1158. [5] B. Chertok, A.E. David, V.C. Yang, Biomaterials 31 (2010) 6317–6324. [6] X. Mei, D. Chen, N. Li, Q. Xu, J. Ge, H. Li, B. Yang, Y. Xu, J. Lu, Soft Matter 8 (2012) 5309–5316. [7] G. Gupta, M.N. Patel, D. Ferrer, A.T. Heitsch, B.A. Korgel, M. Jose-Yacaman, K.P. Johnston, Chem. Mater. 20 (2008) 5005–5015. [8] T. Itoh, R. Ishii, T. Ebina, T. Hanaoka, Y. Fukushima, F. Mizukami, Bioconjugate Chem. 17 (2006) 236–240. [9] K.-S. Jang, H.-J. Kim, J.R. Johnson, W.-G. Kim, W.J. Koros, C.W. Jones, S. Nair, Chem. Mater. 23 (2011) 3025–3028. [10] H. Tang, J. Guo, Y. Sun, B. Chang, Q. Ren, W. Yang, Int. J. Pharm. 421 (2011) 388– 396. [11] M.-H. Kim, H.-K. Na, Y.-K. Kim, S.-R. Ryoo, H.S. Cho, K.E. Lee, H. Jeon, R. Ryoo, A.D.-H. Min, ACS Nano 5 (2011) 3568–3576. [12] F. Tang, L. Li, D. Chen, Adv. Mater. 24 (2012) 1504–1534. [13] L. Bau, B. Bartova, M. Arduinia, F. Mancin, Chem. Commun. 45 (2009) 7584– 7586. [14] H.M. Joshi, Y.P. Lin, M. Aslam, P.V. Prasad, E.A. Schultz-Sikma, R. Edelman, T. Meade, V.P. Dravid, J. Phys. Chem. C 113 (2009) 17761–17767. [15] S.-H. Xuan, S.-F. Lee, J.T.-F. Lau, X. Zhu, Y.-X.J. Wang, F. Wang, J.M.Y. Lai, K.W.Y. Sham, P.-C. Lo, J.C. Yu, C.H.K. Cheng, K.C.-F. Leung, ACS Appl. Mater. Interf. 4 (2012) 2033–2040. [16] Y. Chen, H. Chen, D. Zeng, Y. Tian, F. Chen, J. Feng, A.J. Sh, ACS Nano 4 (2010) 6001–6013. [17] Y. Deng, D. Qi, C. Deng, Xiangmin. Zhang, D. Zhao, J. Am. Chem. Soc. 130 (2008) 28–29. [18] Z. Teng, C. Sun, X. Su, Y. Liu, Y. Tang, Y. Zhao, G. Chen, F. Yan, N. Yang, C. Wang, G. Lu, J. Mater. Chem. B 1 (2013) 4684–4691. [19] J. Lee, S.Y. Lee, S.H. Park, H.S. Lee, J.H. Lee, B.-Y. Jeong, S.-E. Park, J.H.J. Chang, Mater. Chem. B 1 (2013) 610–616. [20] C.-X. Zhao, L. Yu, A.P.J.J. Middelberg, Mater. Chem. B 1 (2013) 4828–4833.

41

[21] L. Qi, L. Wu, S. Zheng, Y. Wang, H. Fu, D. Cui, Biomacromolecules 13 (2012) 2723–2730. [22] M.-H. Lee, J.L. Thomas, M.-H. Ho, C. Yuan, H.-Y. Lin, ACS Appl. Mater. Interf. 2 (2010) 1729–1736. [23] S. Narayanan, B.N. Sathy, U. Mony, M. Koyakutty, S.V. Nair, D. Menon, ACS Appl. Mater. Interf. 4 (2012) 251–260. [24] K.H. Bae, M. Park, M.J. Do, N. Lee, J.H. Ryu, G.W. Kim, C. Kim, T.G. Park, T. Hyeon, ACS Nano 6 (2012) 5266–5273. [25] M. Talelli, C.J.F. Rijcken, T. Lammers, P.R. Seevinck, G. Storm, C.F.V. Nostrum, W.E. Hennink, Langmuir 25 (2009) 2060–2067. [26] J.-H. Ke, J.-J. Lin, J.R. Carey, J.-S. Chen, C.-Y. Chen, L.-F. Wang, Biomaterials 31 (2010) 1707–1715. [27] J. Lu, S. Ma, J. Sun, C. Xia, C. Liu, Z.W. A, X. Zhao, F. Gao, Q. Gong, B. Song, X. Shuai, H. Ai, Z. Gu, Biomaterials 30 (2009) 2919–2928. [28] H. Yang, C. Zhang, X. Shi, H. Hua, X. Du, Y. Fang, Y. Mad, H. Wu, S. Yang, Biomaterials 31 (2010) 3667–3673. [29] H.M. Kim, H. Lee, K.S. Hong, M.Y. Cho, M.-H. Sung, H. Poo, Y.T. Lim, ACS Nano 5 (2011) 8230–8240. [30] J. Yang, E.-K. Lim, H.J. Lee, J. Park, S.C. Lee, K. Lee, H.-G. Yoon, J.-S. Suh, Y.-M. Huh, S. Haam, Biomaterials 29 (2008) 2548–2555. [31] E.-K. Lim, J. Yang, C.P.N. Dinney, J.-S. Suh, Y.-M. Huh, S. Haama, Biomaterials 31 (2010) 9310–9319. [32] B. Sahoo, S.K. Sahu, S. Nayak, D. Dhara, P. Pramanik, Catal. Sci. Technol. 2 (2012) 1367–1374. [33] C. Wang, S. Tao, W. Wei, C. Meng, F. Liua, M. Han, J. Mater. Chem. 20 (2010) 4635–4641. [34] B. Sahoo, K.S.P. Devi, R. Banerjee, T.K. Maiti, P. Pramanik, D. Dhara, ACS Appl. Mater. Interf. 5 (2013) 3884–3893. [35] S. Gandhi, S. Venkatesh, U. Sharma, N.R. Jagannathan, S. Sethuramana, U.M. Krishnan, J. Mater. Chem. 21 (2011) 15698–15707. [36] Y. Wang, B. Li, L. Zhang, P. Li, L. Wang, J. Zhang, Langmuir 28 (2012) 1657– 1662. [37] X.L. Zhang, H.-Y. Niu, W.-H. Li, Y.-L. Shi, Y.-Q. Cai, Chem. Commun. 47 (2011) 4454–4456. [38] B. Sahoo, K.S.P. Devi, S.K. Sahu, S. Nayak, T.K. Maiti, D. Dhara, P. Pramanik, Biomater. Sci. 1 (2013) 647–657. [39] S. Liu, H. Chen, X. Lu, C. Deng, X. Zhang, P. Yang, Angew. Chem. Int. Ed. 49 (2010) 7557–7561. [40] T. Fan, D. Pan, H. Zhang, Ind. Eng. Chem. Res. 50 (2011) 9009–9018. [41] Y. Fu, P. Xiong, H. Chen, X. Sun, X. Wang, Ind. Eng. Chem. Res. 51 (2012) 725– 731. [42] Z. Xu, D. Wang, M. Guan, X. Liu, Y. Yang, D. Wei, C. Zhao, H. Zhang, ACS Appl. Mater. Interf. 4 (2012) 3424–3431. [43] B.Y. Yu, S.-Y. Kwak, Dalton Trans. 40 (2011) 9989–9998. [44] Y.-W. Jun, J.-W. Seo, J. Cheon, Acc. Chem. Res. 41 (2008) 179–189. [45] B.H.B. Na, I.C. Song, T. Hyeon, Adv. Mater. 21 (2009) 2133–2148. [46] X. Wang, A.R. Morales, T. Urakami, L. Zhang, M.V. Bondar, M. Komatsu, A.K.D. Belfield, Bioconjugate Chem. 22 (2011) 1438–1450. [47] Y.-L. Liu, J.-L. Zhang, G. Cheng, G.-Y. Hong, J.-Z. Ni, Nanotechnology 23 (425702) (2012) 8. [48] P. Mohan, N. Rapoport, Mol. Pharm. 7 (2010) 1959–1973.