Luminescent mesoporous nanoreservoirs for the effective loading and intracellular delivery of therapeutic drugs

Acta Biomaterialia 10 (2014) 1431–1442

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Luminescent mesoporous nanoreservoirs for the effective loading and intracellular delivery of therapeutic drugs Sooyeon Kwon a,1, Rajendra K. Singh b,c,1, Tae-Hyun Kim b,c,1, Kapil D. Patel b,c, Jung-Ju Kim a,b,c,d, Wojciech Chrzanowski a, Hae-Won Kim b,c,d,⇑ a

The Faculty of Pharmacy, The University of Sydney, NSW 2006, Australia Institute of Tissue Regeneration Engineering (ITREN), Dankook University, Cheonan 330-714, South Korea Biomaterials and Tissue Engineering Laboratory, Department of Nanobiomedical Science & BK21 PLUS NBM Global Research Center for Regenerative Medicine, Dankook University, Cheonan 330-714, South Korea d Department of Biomaterials Science, College of Dentistry, Dankook University, Cheonan 330-714, South Korea b c

a r t i c l e

i n f o

Article history: Received 14 July 2013 Received in revised form 29 September 2013 Accepted 24 October 2013 Available online 15 November 2013 Keywords: Nanocarrier Hollow silica Luminescent Multifunctional Drug delivery

a b s t r a c t Development of biocompatible and multifunctional nanocarriers is important for the therapeutic efficacy of drug molecules in the treatment of disease and tissue repair. A novel nanocarrier of luminescent hollowed mesoporous silica (L-hMS) was explored for the loading and controlled delivery of drugs. For the synthesis of L-hMS, self-activated luminescence hydroxyapatite (LHA) was used as a template. Different thicknesses (7–62 nm) of mesoporous silica shell were obtained by varying the volume of silica precursor and the subsequent removal of the LHA core, which resulted in hollow-cored (size of 40 nm  10 nm) mesoporous silica nanoreservoirs, L-hMS. While the silica shell provided a highly mesoporous structure, enabling an effective loading of drug molecules, the luminescent property of LHA was also well preserved in both the silica-shelled and the hollow-cored nanocarriers. Doxorubicin (DOX), used as a model drug, was shown to be effectively loaded onto the mesopore structure and within the hollow space of the nanoreservoir. The DOX release was fairly pH-dependent, occurring more rapidly at pH 5.3 than at pH 7.4, and a long-term sustainable delivery over the test period of 2 weeks was observed. The nanoreservoir exhibited favorable cell compatibility with low cytotoxicity and excellent cell uptake efficiency (over 90%). Treatment of HeLa cells with DOX-loaded L-hMS elicited a sufficient degree of biological efficacy of DOX, as confirmed in the DOX-induced apoptotic behaviors, including stimulation in caspase-3 expression, and was even more effective than the direct DOX treatment. Overall, the newly developed L-hMS nanoreservoirs may be potentially useful as a multifunctional (luminescent, mesoporous and biocompatible) carrier system to effectively load and sustainably deliver small molecules, including anticancer drugs. Ó 2013 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved.

1. Introduction Nanoparticulates are considered promising candidates for diagnosis and therapeutic applications [1–3]. Inorganic nanoparticles have relatively low toxicity, and well-controlled synthesis methods allow for the fine tuning of their characteristics and surface functionalization. Hence, inorganic nanoparticles are considered powerful biocompatible material platforms for the delivery of a wide range of drugs [4–7]. Currently, drug delivery systems are being designed to deliver drug molecules in a more controlled manner, with high specificity and lower dosages, which is ⇑ Corresponding author at: Institute of Tissue Regeneration Engineering (ITREN), Dankook University, Cheonan 330-714, South Korea. Tel.: +82 41 550 3081; fax: +82 41 550 3085. E-mail address: [email protected] (H.-W. Kim). 1 These authors contributed equally to this work.

ultimately aimed at enabling interactions of the drug solely with target tissues. Mesoporous silica nanomaterials in particular have emerged as excellent candidates in drug delivery [8,9]. They have distinct advantages, including large surface area and volume, for the uptake of large quantities of drugs, and have easily adjustable surface properties, which may be adapted for specific types of drug molecules [8–12]. Moreover, the release of drugs can be tuned by modifiying the mesopore architecture and surface chemistry [8–13]. Substantial efforts are continuously being made to optimize their pore parameters, such as enhancement of pore volume or pore size, with the aim of improving the storage capacity of drug molecules and/or facilitating host-specific interactions [14–17]. Another important feature of nanoparticles, including mesoporous silica, is their ability to provide multifunctionality, e.g. fluorescent, photodynamic, electrical and magnetic properties [18,19]. This is mainly associated with the ability of the therapeutic

1742-7061/$ - see front matter Ó 2013 Acta Materialia Inc. Published by Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.actbio.2013.10.028

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systems to be used for diagnostic purposes. Organic dyes and quantum dots (QDs) have been widely employed for biological labelling and diagnosis due to their outstanding fluorescent properties [20,21]. However, the fluorescence properties of organic dyes have been shown to easily decay or be quenched, and heavy metal ions such as Cd2+ and Pb2+ from QDs are severely cytotoxic, limiting their biomedical applications [22,23]. Therefore, exploring different material sources for imaging, which are biologically safe while retaining proper imaging performance, is considered important. Herein, we report on the development of a novel mesoporous silica (MS)-based nanocarrier system which is truly multifunctional, possessing well-controlled drug loading/release ability as well as fluorescence for imaging purposes. To achieve the aim of the research we utilize luminescent hydroxyapatite (LHA). HA is biological material that is a common constituent of hard tissues, including bone and tooth, and thus has proven biocompatibility [24,25]. In particular, the HA crystal structure can exhibit self-activated luminescence, depending on the defects or impurities present in the host lattice [26–30]. LHA is considered to be a promising candidate for a luminescence source, due to its good optical properties and biocompatibility. LHA was used here as the core part of a system obtained by layering MS onto LHA; as a consequence, an LHA@MS core–shell structure was obtained. The outer mesoporous structure of LHA@MS allows drug loading and controls the drug release. Furthermore, because the core LHA part can be removed, it can provide a hollow-core MS (hMS) that is an addition drug depot. This significantly increases the capacity for drug loading when compared to other silica-based systems. Interestingly, we have observed that hMS also preserves the luminescent property of LHA, resulting in a novel luminescent, nanocarrier system hMS (L-hMS). Although very recent reports have described hollowed MS formed using a template replication approach [31,32], to the best of our knowledge, luminescent hollow MS is presented here for the first time. For applications of L-hMS in an intracellular drug delivery system, we use the drug doxorubicin (DOX). As the common side effects of DOX therapy (such as tissue necrosis and cardiotoxicity) are related to the high dosage of the drug with uncontrolled release and the lack of drug specificity [32,33], the hollow MS system is believed to provide a versatile carrier that is characterized with a high loading capacity, coupled with a controlled and sustainable release profile of drug molecules, including DOX. The intracellular penetration of the DOX-loaded nanoparticles can be imaged in situ due to the self-fluorescence property of the drug carrier. This paper presents a detailed description of the synthesis of the L-hMS, and a thorough analysis of the key physico-chemical properties of the system, including mesoporosity and optical fluorescence. The loading capacity of DOX and its release profile from the L-hMS nanoreservoir are investigated, and the in vitro biological efficacy of the intracellular delivery system is confirmed.

2. Materials and methods 2.1. Preparation of LHA LHA was fabricated by the precipitated reaction of the supersaturated solutions of A and B. Solution A was prepared by mixing 0.6 g of hexadecytrimetyl ammonium bromide (CTAB) with 1.3743 g of calcium nitrate tetrahydrate and 3% of zinc nitrate hexahydrate dissolved in 60 ml of deionized water. The zinc ions were added to decrease the LHA particle size to a few tens of nanometers [34]. A small amount (3 ml) of ammonium hydroxide solution (28%) was added to solution A to adjust the pH to 9–10. Solution B was prepared by mixing 1.7646 g of trisodium citrate

and 0.4752 g of ammonium hydrogen phosphate dissolved in 15 ml of deionized water. After vigorously stirring for 10 min, solution B was introduced into solution A. A ratio of 1:1 was selected from trisodium citric acid/calcium nitrate molar proportion (Cit/ Ca). After additional agitation for 60 min, the solution was transferred to a bottle held in a stainless steel autoclave, which was then sealed and allowed to react at 185 °C for 24 h. When the autoclave had cooled to room temperature, the LHA was precipitated out of solution by centrifugation at 8000 rpm. Next, the precipitated LHA nanoparticles were washed with deionized water and ethanol, then dried at 80 °C in air for 24 h to obtain LHA nanoparticles for further use. 2.2. Preparation of LHA@MS MS coatings on the LHA template were obtained by mixing the silica precursor tetraethyl orthosilicate (TEOS) with LHA. Before adding the TEOS, 30 mg of freshly synthesized LHA in 200 ml of ethanol was sonicated for 30 min in an ultrasonic bath and then stirred at 500 rpm to make sure all of the LHA particles were well dispersed for the uniform silica coating. Next, 5 g of CTAB in 8 ml of ammonium hydroxide and 22 ml of deionized water was added to the LHA solution and mixed for 30 min under high-power ultrasound using a Sonoreactor (Ulsso Hitech) operating at 20 kHz and 700 W. Different amounts of TEOS (50, 100, 200, 300 and 400 ll) were then added to the mixture, which was further sonicated for 2 h to prevent agglomeration. The mixture was then stirred overnight at 1200 rpm to obtain well-dispersed mesoporous silica-coated LHA (LHA@MS). The solution was subsequently centrifuged and thoroughly washed with absolute ethanol and deionized water to ensure CTAB removal, then dried overnight under a vacuum. 2.3. Preparation of L-hMS To obtain L-hMS, the cored LHA in the LHA@MS was dissolved by an etching process. LHA@MS200 was chosen as the representative sample for this hollow structuring. For this, 0.15 g of LHA@MS200 was dispersed in 30 ml of deionized water and then mixed with 20 ml of 1 N HCl solution. The mixture was sonicated for 2 h and then stirred at 1200 rpm to prevent agglomeration and ensure thorough mixing. The mixture was next treated with 20 mg of ammonium nitrate solution in 40 ml of deionized water at 60 °C overnight. The nanoparticle solution was centrifuged and washed with ethanol and deionized water, then dried overnight under a vacuum to obtain the L-hMS. The nanocarrier samples prepared in this study are coded as summarized in Table 1. 2.4. Sample characterizations To assess the crystalline structure of the prepared nanoparticles, we employed X-ray diffraction (XRD) using a diffractometer (Rigaku) operating at 40 kV and 40 mA, with Cu Ka radiation (k = 1.5418 Å). Scanning was made with diffraction angles of 2h = 0–60°, a scanning rate of 2 °min1 and step size of 0.02°. Fourier transform infrared spectroscopy (FTIR; Varian 640-IR) was used to characterize the functional groups and the chemical bonding of the samples. For each spectrum, 20 scans in the range of 400–2000 cm1 wave numbers were recorded in transmission mode, using the KBr pellet method. Nanoparticle size and morphology were determined using transmission electron microscopy (TEM; JEOL-7100). The samples for TEM were prepared by dispersing the nanoparticles in ethanol and placing a drop of the suspension onto a carbon-coated copper grid. The thickness of silica shells formed on the nanoparticles was measured from arbitrarily

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S. Kwon et al. / Acta Biomaterialia 10 (2014) 1431–1442 Table 1 Nanoparticle sample code and the concentration of materials used for the preparation. Sample code

LHA LHA@MS

Materials used

MS50 MS100 MS200 MS300 MS400

L-hMS

Note

LHA (mg)

TEOS (ll)

– 30 30 30 30 30 –

– 50 100 200 300 400 –

selected TEM images (>50 nanoparticles). The atomic composition of the samples was analyzed by energy-dispersive spectroscopy (EDS; Bruker). The zeta (f) potential of the samples was measured using a Malvern Zetasizer (ZEN3600; Malvern). The f-potential was measured in triplicate at 25 °C in deionized water. Specific surface area, pore volume and pore size were determined by nitrogen gas adsorption/desorption isotherm at 77 K using a Quadrasorb SI automated surface area and pore size analyzer (2SI-MP-9 Quantachrome). Samples were degassed under a vacuum prior to analysis, with a degassing temperature of 100 °C and an outgas time of 4 h. The nanoparticle size was analyzed with a particle size analyzer (Zetasizer Nano ZS, Malvern Instruments, UK), based on a dynamic light scattering (DLS) measurement. The fluorescence properties of the nanoparticle samples were observed by photoluminescence (PL) using a Jasco FP-6500spectrofluorometer. Fluorescence images of the samples were also obtained using a UV transilluminator (Vilber Lourmat), with deionized water as a reference solution. The electron paramagnetic resonance (EPR) properties of the nanoparticles were characterized using a JEOL spectrometer (model JES-FA200) operated at an Xband frequency (m) of 9.4 GHz with 100 kHz magnetic field modulation. 2.5. DOX loading and release studies DOX was dissolved in PBS at pH 7.4 to produce a stock solution at a concentration of 120 lg ml1. A standard curve was obtained by measuring optical intensities of serial dilutions of the stock solution (20, 30, 40, 50, 60, 80, 100 and 120 lg ml1) using a UV–vis spectrophotometer at an absorbance of 483 nm. For the DOX loading test, 1 mg of nanoparticle samples (LHA@MS and LhMS) was ultrasonically dispersed in each DOX solution for 5 min, which were then kept in a 37 °C water bath for 4 h. To quantify the drug loading amount, the nanoparticles were centrifuged at 10,000 rpm for 5 min and the supernatant was gathered for the assay using a UV–vis spectrophotometer (A483 nm). To examine the release profile of DOX from the nanocarrier samples, the DOX-loaded nanocarriers (LHA@MS and L-hMS), which were prepared in a 120 lg ml1 initial DOX concentration, were used. To test the DOX release kinetics, 2 mg of each sample was dispersed in 2 ml of PBS prepared with different pH values (5.5 and 7.4), and then incubated at 37 °C. At different time points (up to 2 weeks), the amount of drug released into the medium was measured. After centrifugation of the nanoparticle–drug complex at 10,000 rpm for 5 min, the supernatant was gathered and then assessed using a UV–vis spectrophotometer (A483 nm). The medium was refreshed at each testing. 2.6. In vitro cell toxicity assay The in vitro cellular toxicity of the prepared nanocarriers was examined using the human cervical cancer cell line HeLa cells (American Type Culture Collection, Rockville). Cells were

LHA only MS-shelled LHA

MS200 after removal of LHA

maintained at 37 °C in an atmosphere of 5% CO2 in a-minimum essential medium (a-MEM; Gibco) with 10% fetal bovine serum (Gibco) and 1% penicillin–streptomycin. The medium was replaced two or three times per week, and the cells were passaged to subconfluency. For the MTS assays, HeLa cells were seeded in 96-well plates at a density of 104 cells per well and allowed to attach for 24 h. The attached cells were then treated with L-hMS, DOX or DOX-loaded L-hMS with various doses in culture medium for 24 h. After each treatment, the cells were incubated with mixtures of 100 ll of aMEM and 20 ll of MTS solution (CellTiter 96 Aqueous One Solution; Promega) for 4 h in the dark at 37 °C. At the end of the MTS treatment, the OD values at 490 nm were read using an iMark microplate reader (BioRad). Cell viability was determined based on the percentage of the OD value of the experimental group over that of the control group. 2.7. Fluorescence imaging for DOX-loaded nanocarrier in HeLa cells HeLa cells, seeded at 105 cells in each well of 6-well culture plates treated with L-hMS, DOX or DOX-loaded L-hMS for 4 h, were harvested and fixed with 4% paraformaldehyde solution for 30 min. The fixed cells were washed with cold PBS (4 °C) and then plated on glass slides. The fluorescent images were observed and analyzed under a Zeiss LSM 510 laser-scanning confocal microscope (Zeiss). Cells were counterstained with 40 ,6-diamidino-2-phenylindole (DAPI; Invitrogen, USA) to observe the nucleus. 2.8. Fluorescein isothiocyanate (FITC)–Annexin V and propidium iodide (PI) double staining for apoptosis detection FITC–Annexin V and PI double stain was used to detect the apoptosis induced by DOX-loaded L-hMS. The HeLa cells treated with L-hMS, DOX or DOX-loaded L-hMS for 24 h were harvested and washed with cold PBS. The washed cells were then stained using an FITC–Annexin V Apoptosis Detection Kit (BD Pharmingen). Briefly, the cells were resuspended in 1 ml of 1 binding buffer at a concentration of 1  106 cells ml1. Next, 5 ll of FITC– Annexin V and 5 ll of PI were added per 100 ll of cell suspension (1  105 cells). After gentle vortexing, the cells were incubated for 15 min at room temperature in the dark. Subsequently, 400 ll of 1 binding buffer was added to each tube prior to analysis using a FACSCalibur flow cytometer (BD Biosciences). The data acquired for 10,000 cells in each sample were analyzed using the CellQuest Pro software (BD Biosciences). 2.9. Analysis of caspase-3 expression A Caspase 3 (active) FITC Staining Kit (BD Pharmingen) was used to detect the apoptosis induced by the DOX-loaded L-hMS. HeLa cells treated with L-hMS, DOX or DOX-loaded L-hMS were cultured for 24 h and then harvested. The collected cells were washed with cold PBS and then fixed with BD Cytofix/Cytoperm™

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Fig. 1. Schematic showing the development of luminescent nanoreservoirs for the loading and sustained delivery of drug DOX. LHA nanocrystals were covered with a thin layer of mesoporous silica (LHA@MS) which was subsequently hollow-cored by removal of the LHA (L-hMS). DOX molecules can be loaded onto the mesopores of the LHA@MS or inner hollow space of the L-hMS. The MS layer thickness was controlled by varying the amount of added TEOS.

solution for 30 min on ice. The fixed cells were washed twice with BD Perm/Wash™ buffer and then resuspended in the BD Perm/ Wash™ buffer plus FITC–Caspase 3 antibody for 30 min at room temperature in the dark. The stained cells were then washed with BD Perm/Wash™ buffer and analyzed using a FACSCalibur flow cytometer. The data acquired for 10,000 cells in each sample were analyzed using the CellQuest Pro software. 3. Results and discussion 3.1. Preparation and characteristics of nanocarriers As shown in the schematic view in Fig. 1, the luminescent HA (LHA) used as the core template was layered with mesoporous silica (LHA@MS) by the sol–gel reaction using TEOS. The shell thickness was controlled by varying the TEOS concentration (from 50 to 400 ll TEOS per 30 mg of LHA). Prior to adding TEOS, the LHA solution was ultrasonicated to aid even dispersion, as well as uniform sol–gel reaction and coating processes. More specifically, partially hydrolyzed TEOS molecules adsorb and nucleate heterogeneously onto the surface of LHA nanoparticles, and this is accompanied by a condensation reaction under alkaline conditions (pH 9–10 used herein) to form silanol groups, enabling stable deposition of a silica layer onto the LHA nanoparticles [17]. Subsequently, the core LHA was removed to create hollow-core MS nanoparticles, which are also luminescent (L-hMS). Drug molecules are then loaded both onto the mesopores of the silica shell and within the hollow space for subsequent release, thus making a proper delivery system. Table 1 summarizes the compositions and characteristics of the different luminescent nanocarriers prepared in this study. The morphology of the nanocarriers was examined by TEM, as shown in Fig. 2. LHA, used as the core template (Fig. 2a), showed a rod-shaped morphology, with dimensions of 30–40 nm length  10 nm width (an average of 37 nm  11 nm). Fig. 2b–e shows the TEM images of the LHA@MS nanocarriers produced with different TEOS concentrations. The silica shell was mesoporous and covered the LHA nanoparticles to a uniform thickness. The thickness of the shell was controlled by the fabrication conditions, i.e. the TEOS concentration (Fig. 2g). There was a gradual increase in shell thickness from 7 to 62 nm as the TEOS amount increased from 50 to 400 ll. There was a strong linear relationship between the thickness and the TEOS content (R2 = 0.99), which indicates that the shell thickness can be easily tailored by controlling the TEOS concentration. As the shell part is required to play a key role in taking up drug molecules, the precise control of the shell

thickness is of particular importance in the performance of the nanocarriers. We subsequently removed the LHA core part by treatment with acid and the hollow-core morphology of the LhMS was obtained (Fig. 2f). The mesoporous structure of the silica shell was shown to be intact after the removal process. This hollow-core mesoporous nanoreservoir has a high drug loading capacity due to the volumes available in both the core and pores in the shell. The shell part is considered to act as a barrier against the diffusion of drug molecules, i.e. increasing the shell thickness would retard the escape of drug molecules through the mesopore channels as soon as the drug molecules were incorporated within the hollow space. The LHA@MS nanoparticles sizes range from 49 nm x 23 nm to 161 nm x 135 nm, based on the TEM images. The hydrodynamic particle size was also examined by means of DLS measurement. LHA@MS-200 and L-hMS were used for the representative samples. The DLS assay showed particle sizes of 120 and 102 nm for LHA@MS-200 and L-hMS, respectively. Considering the size of LHA@MS-200 (93 nm  67 nm) taken from the TEM images, the DLS-measured size showed a somewhat increased value, reflecting the hydrodynamic effects of nanoparticles in solution. Such a small increase generally means that the nanoparticles are pretty well dispersed and not severely agglomerated [35]. The phase and crystalline structure of the developed nanocarriers were analyzed using XRD. LHA showed peaks typical for crystalline HA (Fig. 3a). When the MS shell was formed, one dominant broad peak at 2h = 23°, which corresponds to a typical amorphous phase of silica, was detected. The relative intensity of the silica peak increased with increasing shell thickness. No other peaks except those associated with HA and silica were observed in any of the LHA@MS samples. When the LHA core was removed (LhMS sample), only a broad amorphous silica peak was detected. This finding confirmed the complete removal of the LHA core phase. The chemical structure of the samples was analyzed by FTIR spectroscopy (Fig. 3b). The LHA core showed characteristic bands at 1096, 1020, 960, 600 and 565 cm1, which are associated with the typical vibrations of P–O in PO34  ions [26]. Additional bands were observed at 1577 and 1375 cm1, which correspond to CO23 . The stretching vibration of OH was seen at 630 cm1. In the LHA@MS samples, silica-related bands were also present. Si–O–Si (symmetric stretching at 1220 and 1095 cm1, and asymmetric stretching at 801 cm1), Si–OH (symmetric stretching at 954 cm1) and Si–O (bending at 465 cm1) are the major bands [17], and the intensities increased as the silica shell thickened. For the L-hMS, only silica-related bands were revealed. The FTIR results were in agreement with the XRD patterns.

S. Kwon et al. / Acta Biomaterialia 10 (2014) 1431–1442

Fig. 2. TEM images of the luminescent nanocarriers: (a) LHA, (b) MS50, (c) MS100, (d) MS200, (e) MS400 and (f) L-hMS. (g) The thickness of the mesoporous silica shell for various LHA@MS compositions, showing a steady increase in the thickness with increasing concentration of TEOS, used as the silica precursor.

Interestingly, the bands in the range of 1400–1800 cm1 were evidenced in all of the samples (LHA, LHA@MS and particularly L-hMS), demonstrating a common chemical bond in all of the nanocarriers. This is related to the presence of carbon-related impurities. In fact, in pure LHA the key luminescent factor is the CO 2 radical ion, and the bands were also ascribed to this. Importantly, this radical group also exists in the LHA-removed hollow MS sample, which suggests the retention of the radical ions within the hollow space after the LHA removal process. EDS analyses of the carriers further confirmed the chemical composition of each nanocarrier system (Fig. 3c). While Ca and P were clearly seen in LHA, an additional Si peak appeared as the primary signal in LHA@MS samples, and there was only an Si peak (no Ca and P) in the L-hMS nanoreservoirs. This elemental composition

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result confirmed the complete removal of the core apatite crystalline phase, and was in agreement with the FTIR results. The nanocarriers were dispersed in distilled water to examine the surface charge property through f-potential measurement (Fig. 3d). Negative f-potentials were observed for all samples. In particular, the f-potential of LHA became more negative with the coverage of the MS shell, and increased with increasing shell thickness (18.1 mV for LHA to 26 mV for LHA@MS400). The f-potential of L-hMS (21.8 mV) was comparable to that of LHA@MS200 (22.4 mV), reflecting the effect of same silica composition. While LHA could be negatively charged due to the presence of OH and PO34  ions in the crystal structure, the MS shell is liable to hydrolyse in aqueous medium, forming abundant negative Si–OH groups on the surface. The silica shell properties were analyzed in greater depth by observing the mesopore structure, including the surface area, pore volume and pore size, which are important parameters in determining the capacity to load drug molecules. First, the nitrogen gas adsorption/desorption isotherm of the nanocarrier samples was plotted (Fig. 4a). All of the LHA@MS samples showed typical type IV isotherms, representing narrow hysteresis loop areas. The volume adsorbed increased as the MS shell thickness increased at a given pressure region. The hollowed MS showed the highest volume adsorbed. In particular, the isotherm curve showed a notable hysteresis loop in the relative pressure region of 0.5 < p/p0 < 0.95, which is characteristic of a cavity networked nanostructure, as has been similarly observed elsewhere [36,37]. Based on the isotherms, the specific surface area and pore volume were measured. The surface area of the samples increased as the MS shell thickness increased from 110 to 510 m2 g1 (Fig. 4b), and the pore volume also increased (from 0.15 to 0.450 cm3 g1, Fig. 4c). Furthermore, the hollow MS showed a substantial increase in both surface area and pore volume (1012 m2 g and 0.818 cm3 g1, respectively). This finding suggests that the hollow core is a depot for a large quantity of drug molecules, along with the mesopores of the shell. In the LHA@MS samples, the mesopores were 3.6 nm in size, whereas, for the hollowed MS, the average mesopore size was 4.3 nm. This small change in size can be related to the removal process of the LHA from the core. Otherwise, the BJH model used for the mesopore size estimation might not accurately discriminate the mesopores from the inner hollow space. However, based on the typical pore size distribution curves (inset in Fig. 4d), which show a narrow distribution of pore size with a sharp peak at the average value (4.3 nm), the pore enlargement effect is considered to be more reasonable. After confirming the efficacy of silica shells in producing a highly mesoporous structure, as well as the large capacity of the hollowed space in L-hMS, we next sought to examine the luminescent property of the nanocarriers. It has been reported previously that, when citric acid is used as a chelating agent for the sol–gel preparation of HA, the R–C–COO (Cit3) group is readily adsorbed onto the HA surface to form a Ca–citrate chelating complex. During further processing, the Cit3 group may cleave to R–C and CO2  , and some CO2  radicals are trapped in the HA lattice or interstitial positions, which consequently result in the formation of luminescent centers. The typical fluorescence emission spectrum of the luminescent HA over the wide excitation range is shown in Fig. 5a. A broad spectrum at 300–400 nm, with a maximum at 340 nm, was noticed, and the corresponding emission spectrum was observed at 427 nm. The emission spectrum bands of all of the nanocarrier samples are presented in Fig. 5b. Spectra comprising two bands centered at 427 and 466 nm were similarly observed for all nanocarriers, including the MS-shelled LHA and LHA-hollowed MS samples. The PL emission spectra of LHA@MS samples is understandable, as this originates from the cored LHA, and the lowered intensity in the emission band reflects the shielding

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Fig. 3. (a) XRD pattern and (b) FTIR spectrum of the nanocarrier samples. (c) EDS peaks showing the atomic composition of each sample (Ca and P in LHA vs. Ca, P and Si in LHA@MS vs. only Si in L-hMS). (d) f-potential of the nanoparticles measured at pH 7.0, showing a gradual increase in the negative values as the shell thickness increases.

Fig. 4. Mesopore structure analyzed by the BET method: (a) N2 adsorption/desorption isotherm, (b) specific surface area, (c) specific pore volume and (d) mesopore size of each nanocarrier sample. The mesopore size distribution of two representative samples is also included in the inset of (d). While the N2 adsorption/desorption curves were observed to be similar in all the LHA@MS samples, the curve of L-hMS differed, showing a large area in the hysteresis loop in a relatively high pressure region, which is ascribed to the hollowed space. The surface area and pore volume increased gradually as the MS shell thickness increased, and much higher values were recorded in the LhMS. The mesopore size of all the LHA@MS samples were comparable (3.5 nm) and a slightly bigger size was observed for the L-hMS particles (4.4 nm).

effects of the existing MS shell. However, the high emission spectrum seen in the hollow MS sample is somewhat surprising as the LHA core had been removed. It is thus reasoned that the CO2  radical, which is believed to play a key role in the lumines-

cent behavior of LHA, remains in the hollow core even after the removal process. While some part of the CO2  radical may be lost during the removal process, most is considered to remain, as only a negligible decrease in the luminescence of L-hMS compared to

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Fig. 5. PL properties of the nanocarriers: (a) a typical PL excitation spectrum seen at 340 nm upon emission at 427 nm; (b) PL emission spectra of all nanocarrier samples, representing a maximal emission peak at the same position (427 nm) while the intensity is reduced as the MS shell thickness is increased; (c) fluorescence images of the samples dissolved in deionized water visualized using a UV transilluminator; and (d) EPR spectra of the nanocarrier samples measured under an applied magnetic field.

that of LHA@MS200 (the sample before the hollowing process) was observed. The FTIR results in the preceding section indicate that common bands around 1400–1800 cm1 were observed for all samples, including LHA, LHA@MS and L-hMS, which also supports the universal existence of CO 2 radicals. The results observed from PL measurement were consistently seen in the images generated by the UV transilluminator (Fig. 5c). The PL property of the LhMS was also quite stable, as the PL intensity of the nanoparticles was observed to be preserved after storage for a month in ambient conditions. In order to confirm the luminescence mechanism, EPR spectroscopy was performed on the nanocarrier samples (Fig. 5d). The LHA sample exhibits EPR bands typically at g = 2.1177 and g = 1.9805 (where g is the Lande g-factor), indicating that there indeed exist paramagnetic defects in LHA nanoparticles. Since the EPR signal cannot be caused by P5+, Ca2+ or O2 (there is no single electron in these ions), it must arise from some radical-related defect, such as peroxyl radicals or carbon dioxide radical anions (CO2  ) [26,29,38,39]. Similarly, EPR signals were seen in other nanocarrier samples (LHA@MS and L-hMS), while the signal intensity was decreased when the MS shell thickness increased, and also when the nanocarrier was hollowed. The EPR result being in good agreement with the PL measurements demonstrates that all the nanocarriers have effective paramagnetic defects, which are most likely CO2  radical ions. 3.2. Drug loading and release profiles In order to examine the capacity of the developed nanocarriers for the loading and delivery of therapeutic molecules, DOX anticancer drug was selected. DOX has a pKa of 8.3, and is thus positively charged at pH 7.4 [40], which enables it to be attracted to

the negatively charged nanocarriers without additional surface functionalization. Furthermore, DOX is a small molecule and can thus possibly be incorporated within the mesopore channels (3– 4 nm herein) and even within the hollow space. The DOX loading study was first carried out using two representative nanocarriers (LHA@MS200 and hollow L-hMS). The DOX loading was recorded at a given nanocarrier content (1 mg) while varying the DOX concentrations from 20 to 120 lg. For the LHA@MS200 sample, the DOX loading amount increased gradually with increasing concentration of DOX used, and a maximum loading of 30 lg was recorded for 60 lg/ml1 of sample (Fig. 6a). In the case of L-hMS, the DOX loading amount also increased linearly with increasing DOX content, up to 80 lg/ml1, at which level the maximum loading amount was as high as 70 lg. The loading capacity of both nanocarriers was thus significantly different, i.e. the amount of DOX loaded onto L-hMS (70 lg) was more than double that loaded onto LHA@MS200 (30 lg). These results clearly show the effects of the hollow core in the loading of DOX molecules. As DOX is oppositely charged to the MS surface and is small in size, its incorporation into the mesopore channels should occur easily, and even penetration through the channels into the hollow space should be possible. The DOX drug molecule is considered to be well chosen for demonstrating the loading capability of the currently developed hollowed nanocarriers. The loaded DOX was subsequently released into PBS with different pH values. While the pH 7.4 represents normal physiological conditions, the acidic pH 5.3 is representative of the extracellular tissues of tumor cells (pH  5–6) [41]. The initial 48 h of release patterns from either LHA@MS200 or L-hMS are presented first (Fig. 6b). The release pattern clearly shows a two-stage behavior, i.e. an initial rapid release up to 10 h followed by a substantially sustained release. It is considered that the initial rapid release

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Fig. 6. DOX drug loading and release study using representative nanocarrier samples of LHA@MS200 and L-hMS. (a) The DOX loading curve on each nanocarrier sample, obtained by measuring the amount of DOX loaded onto the nanocarrier while varying the initial DOX concentration (20–120 lg/ml1). The DOX loading was saturated at 60 lg/ml1 for LHA@MS200 and at 80 lg ml1 for L-hMS, and the maximal loading attained was significantly higher on the L-hMS (70 mg) vs. LHA@MS200 (30 lg). DOX release from the nanocarrier samples was observed at two different pH values (5.3 and 7.4) at 37 °C. (b) DOX release from the two different nanocarrier samples examined over the initial period of 48 h and (c) longterm release of DOX from the L-hMS sample over 300 h. DOX was released pHdependently, i.e. more rapidly at pH 5.3 than at pH 7.4, and the pH-dependence was more noticeable in L-hMS than in LHS@MS. DOX was continuously released over the long term, with a release profile which was largely pH-dependent (higher rate at low pH) in the L-hMS sample. The data fitted a combined zero-order model (initial stage) and Ritger–Peppas model (later stage) well. The parameters are summarized in Table 2.

primarily resulted from the DOX molecules present at the outer surface or the outermost region of the mesopore channels, where the water molecules permeate rapidly to dissociate the DOX molecules, thereby enabling their release. The enhanced protonation of DOX molecules associated with acidic pH thus easily and rapidly

occurs in this region, resulting in a significant pH-dependent release. On the other hand, the second stage is considered to reflect the release of the DOX that is present inside the mesopore channels (for the case of LHA@MS200) and/or within the hollow space (for the case of L-hMS), and is mainly dominated by a diffusion mechanism. The higher initial release (up to 10 h) of DOX from L-hMS than from LHA@MS200 is presumably due to the greater quantity of DOX loaded in the L-hMS. With regard to the effect of pH, initially, DOX was released more rapidly under acidic conditions than under neutral conditions from both nanocarriers, indicating a pHdependent release pattern. A possible explanation for this pH-sensitive phenomenon is that DOX hydrophilicity and solubility could increase in acidic environments due to the stronger protonation of the –NH2 groups in the DOX molecules [42,43]. Of note, the L-hMS nanoreservoir was highly sensitive to pH. The resultant DOX release from L-hMS at pH 5.3 was 38% at 10 h, 42% at 24 h and 53% at 48 h, which were almost twice the percentages released at pH 7.4 (19% at 10 h, 23% at 24 h and 34% at 48 h). After confirming the pH-dependent DOX release behavior, we further examined the DOX release profile from the hollow nanocarriers over the longer term (Fig. 6c). As was demonstrated, the whole release pattern consists of two stages, i.e. an initial rapid release stage up to 10 h and a later sustained release region to the end (300 h), with gradually curved patterns for both pH values. Interestingly, the initially generated pH-dependent difference was shown to be preserved in the second stage without any narrowing of the gap between pH 7.4 and 5.3. As was demonstrated, the later sustained profile results from the DOX diffusion out through the mesopore channels. Although the water molecules (and protons in acidic pH) are freely diffused so can dissociate DOX molecules and engage in ion exchange (as ionic bonds are also possibly engaged between DOX and the MS surface), the process of DOX diffusion takes time, and results in the release being sustained over time. We next sought to find a release mechanism of DOX for this two-stage behavior. While a zero-order model was applied for the first linear stage up to 10 h (Mt/M1 = K0t), the Ritger–Peppas empirical equation was employed for the later stage, which follows the power law [44] Mt/M1 = Ktn, where Mt and M1 are the absolute amounts of drug released at time t and at infinite time (1), respectively, K0 and K are the release rate constant for each equation, incorporating the structural and geometric characteristics of the drug delivery device, and n is the release exponent, indicative of the drug release mechanism. Based on the model, fittings were also presented in the graphs, which showed fairly good fits to the original data. Table 2 summarizes the obtained values of the parameters (K0, K and n) that determine the drug release kinetics equations. At the first stage, the rate constant was higher in an acidic condition, suggesting the higher protonation of DOX molecules associated with acidic pH and the consequently more rapid dissociation and release. Furthermore, the rate constant at the second stage was substantially higher in an acidic condition, reflecting a more rapid initial increase in the behavior curve. The power constants obtained for the second stage were 0.32 (at pH 5.3) and 0.46

Table 2 Summary of the obtained kinetic parameters (K0, K, and n) of curve fittings of the DOX release graph in Fig. 6c. Linear release with zero-order kinetics: Mt/M1 = K0t at 1st stage, and then 2nd stage with a power law relationship provided empirically by Ritger–Peppas: Mt/M1 = Ktn. Model Zero-order (1st stage) Ritger–Peppas Model(2nd stage)

Parameter

hMS (pH7.4)

hMS (pH5.3)

K0 K n

2.33 6.08 0.46

4.62 16.8 0.32

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(at pH 7.4), which demonstrates that the release kinetics is controlled by diffusion to a greater degree [44,45]. Moreover, the slightly lower power constant in the acidic medium was due to the more rapid depletion of drug molecules. Based on the DOX loading and release study, the hollow MS is considered to be an effective carrier for drug delivery. It is important to highlight both the large quantity of DOX molecules loaded within the hollow space and the long-term sustained delivery over 12 days by means of a diffusion-controlled mechanism. Silica nanoparticles have been previously studied for drug delivery purposes, but burst release of the drug from the nanostructure and the rapid depletion of the drug are reported to be common problems [18,46,47]. Some of these limitations have been addressed by a number of complex steps, including the formation of a poly(ethylene glycol) layer on the nanoparticles, which slowed down the drug release through pore narrowing [48]. The fabrication of the hollow core presented here demonstrates the feasibility of controlling the drug release profile by modifying the structure of the carrier. However, the efficacy of the hollow MS as demonstrated here for DOX will not necessarily be applicable to all other types of drugs, as the mesopore structure (pore size and porosity) and the surface functional groups must be properly adjusted for the specific drug that is being applied.

Fig. 7. (a) Effect of L-hMS, DOX and DOX-loaded L-hMS on HeLa cell viability. HeLa cells were treated with various concentrations of DOX (0, 0.1, 0.5, 1, 5, 10, 20, 40, 60 lg ml1), cells were treated for 24 h and their viability was determined by MTS assay. The corresponding concentrations of L-hMS and DOX-loaded L-hMS were 0, 1.43, 7.15, 14.28, 71.4, 142.8, 285.6, 571.2, 856.8 lg ml1. (b) Confocal microscopic images of the HeLa cells incubated with 1 lg ml1 DOX solution or 1 lg ml1 DOXloaded L-hMS for 4 h. The nucleus of the HeLa cell was counter-stained with DAPI.

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3.3. In vitro drug delivery of DOX from nanoreservoir First, the effects of the DOX-loaded L-hMS nanoreservoirs on cell viability using representative tumor cells (HeLa cell line) were examined. For the positive reference group, we treated the HeLa cells with varying doses of plain DOX molecules, in what is generally known to be an effective dose range. The viability of HeLa cells treated with DOX for 24 h was substantially reduced in a DOX dose-dependent manner, clearly illustrating the cytotoxic effects of DOX (Fig. 7a). Next, we utilized the DOX-loaded L-hMS nanoreservoir. In this case, the DOX loading quantities matched the quantities of DOX used for the reference group (0, 0.1, 0.5, 1, 5, 10, 20, 40 and 60 lg ml1). As a result, the corresponding concentrations of the L-hMS nanoreservoirs were determined as 0, 1.43, 7.15, 14.28, 71.4, 142.8, 285.6, 571.2 and 856.8 lg ml1, respectively. When the HeLa cells were treated with the DOX-loaded L-hMS, there was also a substantial reduction in cell viability, which occurred in a fairly dose-dependent manner, and with a similar pattern to that observed for the direct treatment of DOX. In order to confirm whether this effect was from the L-hMS nanoreservoir (and not from the DOX), we also treated the HeLa cells with just L-hMS (without DOX), at the same concentrations as used for the DOX-loaded L-hMS. In this case, the cell viability was preserved well. Even at a concentration of L-hMS nanoreservoirs as high as 285.6 lg ml1, the survival fraction of HeLa cells was 90.4%, indicating that L-hMS has acceptable biocompatibility. This good cell viability of the L-hMS nanoreservoir is considered highly beneficial when utilizing the nanocarrier system more profoundly to deliver therapeutic molecules through the intracellular uptake process. This result also supports the hypothesis that the cytotoxic effects exerted in the DOX-loaded L-hMS were not from the L-hMS nanoreservoirs but from the DOX molecules loaded onto the carrier. More significantly, the cell viability level caused by the DOX-loaded L-hMS nanoreservoirs is just slightly lower than that of the DOX-only treatment, even though not all of the DOX is released from the L-hMS nanoreservoirs during the initial 24 h. This result is attributed to the different cellular uptake mechanisms between naked DOX and nanoreservoir-assisted DOX [43]. The DOX-loaded L-hMS nanoreservoirs involve a possible endocytosis mechanism, which is more effective at entering the cells than naked DOX. Based on these results, the L-hMS nanoreservoirs can potentially be used as a vehicle to contain and deliver anticancer drugs, particularly DOX, to target cancer cells, to enhance the efficacy of anticancer drug delivery due to its pH sensitivity. The cellular uptake of the DOX-loaded L-hMS was subsequently examined. HeLa cells were treated with 1 lg of DOX or the corresponding concentration of DOX-loaded L-hMS for 4 h, and the location of the DOX-loaded L-hMS in the HeLa cells was verified by confocal laser scanning microscopy (Fig. 7b). The red regions represent the fluorescence signal of DOX itself and the blue signals are from the DAPI-stained nucleus. The red fluorescence signals of DOX can thus be observed in the cytoplasm (around the blue signals) or within the nucleus (coincidence with blue signals) of the HeLa cells when treated with DOX. When the cells were treated with naked DOX for 4 h, most red signals were found in the cytoplasm in a diffused pattern. Likewise, in the case of 4 h of treatment with DOX-loaded L-hMS, similar red signals were clearly noticeable within cells. In particular, more intense red signals, presumably due to the collection of DOX molecules within the L-hMS nanoreservoir, were profoundly noticeable. It was thus clear that just 4 h of treatment resulted in a rapid uptake of the DOX-loaded L-hMS nanoreservoirs by the cells. Furthermore, some diffuse red signals in the cytoplasm were ascribed to the DOX molecules possibly having been released from the L-hMS nanoreservoirs. The pHsensitive DOX molecules would be released to some extent and localized primarily in the cytoplasm as the tumor cells have weak

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Fig. 8. Induction of apoptosis in HeLa cells by the treatment of DOX-loaded L-hMS. (a) Representative dot plots of Annexin V/PI staining of HeLa cells untreated (Ctrl), and treatment with either 14.28 lg ml1 L-hMS, 1 lg ml1 DOX or 14.28 lg ml1 DOX-loaded L-hMS. The lower left quadrant contains the vital (double-negative) population. The lower right quadrant contains the early apoptotic (Annexin V positive/PI negative) population. The upper left quadrant contains the necrotic cell (Annexin V negative/PI positive) population, and the upper right contains the late apoptotic population. (b) FACS analysis of the cells positive for FITC showing significant expression of caspase-3 in the DOX-loaded L-hMS (58.61%), in contrast to the DOX (7.05%), L-hMS (0.83) and control groups (0.82%).

acidic environments [33,34,36–40]. Based on the DOX release profiles (as presented in Fig. 6), in the first few hours around 10–20% of the loaded DOX was released. Recalling the cell viability result (in Fig. 7a) effected by the L-hMS-assisted DOX treatment, the significant efficacy was considered to result from the DOX molecules released inside the cellular compartment during the first 24 h. As the DOX release profile indicated that approximately 50% of the quantity loaded was released during the first 24 h at pH 5.3, this biological outcome was confirmed. Although this amount cannot be directly extrapolated to the living cellular environment, a fairly similar range of DOX concentration, which would consequently have a biological effect, is considered to be released in the intracellular area. It should also be noted that, as the loaded DOX is released continuously and sustainably from the L-hMS system within the intracellular compartment, possible long-term efficacy is also envisaged. This is one of the great merits of the L-hMS nanoreservoir delivery system, which enables no repeated dosage treatment of DOX. The confocal fluorescence microscope images and cell viability results demonstrate that the DOX-loaded L-hMS nanoreservoirs were rapidly internalized, and that DOX molecules were then released in a high quantity, resulting in localization mainly within the cytoplasm, which in turn elicited biological efficacy (reduced cellular viability). We further investigated the cellular-uptake characteristics of DOX-loaded L-hMS nanoreservoirs by using flow cytometry. First, the cellular uptake level of the DOX-loaded L-hMS was quantified with flow cytometry by determining the cellular signal of the DOX originating from the DOX-loaded L-hMS. To assess the apoptosis induced by the L-hMS-assisted DOX, the treated cells were double-stained with FITC–Annexin V and PI, and then analyzed by flow cytometry. Externalization of phosphatidylserine (PS) from the inner to the outer cell membrane is an early indicator of apoptosis. Therefore, Annexin V, a phospholipid-binding protein with a high affinity for PS, can be used as an effective marker for

apoptosis. PI, a nonspecific DNA intercalating agent, can enter necrotic or damaged cells, but is excluded by the membranes of living cells or early apoptotic cells. As shown in Fig 8a, almost no apoptotic cells were detected in the untreated control or L-hMS (w/o DOX) treated HeLa cells. However, cell apoptosis was induced substantially in the groups treated with DOX and DOX-loaded L-hMS, to levels of approximately 19.6 and 57.4%, respectively. Interestingly, greater apoptosis resulted from the treatment of the DOXloaded L-hMS over the DOX-only treatment. In other words, in the group treated with DOX only, as little as 19.6% of the population exhibited apoptotic cells, whilst, in the DOX-loaded L-hMS group, as much as 57.4% was apoptotic. The non-apoptotic cell fraction is considered to be necrotic, dying from a cause other than apoptosis. At this point, the cell death mechanism of HeLa cells treated with DOX is considered different to that of cells treated with DOX-loaded L-hMS: it was more apoptotic when the DOX was delivered by the L-hMS nanoreservoir system. To confirm the apoptotic behaviors of cells, caspase-3 expression of cells was further examined. Caspase-3 is a key protease that is activated during the early stages of apoptosis and, like other members of the caspase family, plays a key role in apoptosis and inflammation. As shown in Fig. 8b, virtually no caspase-3-positive cells were detected in the untreated control and L-hMS (w/o DOX) treated HeLa cells. The DOX-treated HeLa cells showed about 7.1% of caspase-3 positive cells. However, there was a significant increase to about 58.6% of caspase-3 positive cells in the L-hMS-assisted DOX-treated HeLa cells. These results on caspase-3 expression demonstrate that the HeLa cells exhibited greater apoptotic activity due to L-hMS-delivery of DOX than that due to DOX alone, which is in agreement with the FACS analyses. With regard to the specific stimulatory apoptotic efficacy of the DOX delivery system via the L-hMS nanoreservoir in direct comparison to the DOX-only treatment, no clear mechanism has yet been determined. It is anticipated that the highly effective endocytosis

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process into cancer cells, including the HeLa cell line [49], as well as the ability of silica nanoparticles with a mesoporous surface structure to escape endosomes, might have played critical roles in the intracellular distribution and apoptotic signaling fate of the DOX molecules. While we have also not yet clarified the enhanced long-term anticancer effects of the delivery systems, the role of the L-hMS nanoreservoir in retaining DOX molecules for continuous release within the intracellular compartments over a period of weeks to a month is clearly envisaged, and future studies on this are warranted. 4. Conclusions We have demonstrated the potential efficacy of the novel luminescent mesoporous nanocarrier L-hMS as a delivery system for therapeutic drugs. The anticancer drug DOX was used to illustrate the nanocarrier’s ability to deliver the drug in a controlled fashion. It was demonstrated that the developed system could effectively load and sustainably release the drug in a pH-dependent manner over a period of weeks. Highly efficient intracellular penetration of the nanoreservoirs was evidenced and enhanced apoptotic effects in tumor cells were observed by the DOX delivered through the L-hMS nanoreservoirs. Together with the self-activating luminescent property and excellent cell compatibility, the newly developed L-hMS nanoreservoirs are considered to hold great promise for therapeutics and imaging in tissue repair and disease treatments. Acknowledgement This study was supported by a grant from Priority Research Centers Program (2009-0093829), National Research Foundation, South Korea. Appendix A. Figures with essential color discrimination Certain figures in this article, particularly Figs. 1, and 3–8, are difficult to interpret in black and white. The full color images can be found in the on-line version, at http://dx.doi.org/10.1016/ j.actbio.2013.10.028. References [1] Cho K, Wang X, Nie SM, Chen Z, Shin MD. Therapeutic nanoparticles for drug delivery in cancer. Clin Cancer Res 2008;14:1310–6. [2] Kim J, Piao YZ, Hyeon T. Multifunctional nanostructured materials for multimodal imaging, and simultaneous imaging and therapy. Chem Soc Rev 2009;38:372–90. [3] Chatterjee DK, Fong LS, Zhang Y. Nanoparticles in photodynamic therapy: an emerging paradigm. Adv Drug Deliv Rev 2008;60:1627–37. [4] Guo Y, Shi DL, Cho H, Dong ZY, Kulkarni A, Pauletti GM, et al. In vivo imaging and drug storage by quantum-dot-conjugated carbon nanotubes. Adv Funct Mater 2008;18:2489–97. [5] You J, Zhang GD, Li C. Exceptionally high payload of doxorubicin in hollow gold nanospheres for near-infrared light-triggered drug release. ACS Nano 2010;4:1033–41. [6] Tang S, Huang XQ, Chen XL, Zheng NF. Hollow mesoporous zirconia nanocapsules for drug delivery. Adv Funct Mater 2010;20:2442–7. [7] Lin MM, Kim HH, Kim H, Dobson J, Kim DK. Surface activation and targeting strategies of superparamagnetic iron oxide nanoparticles in cancer-oriented diagnosis and therapy. Nanomedicine 2010;5:109–33. [8] Manzanoab M, Vallet-Regi M. New developments in ordered mesoporous materials for drug delivery. J Mater Chem 2010;20:5593–604. [9] Yang P, Gaib S, Lin J. Functionalized mesoporous silica materials for controlled drug delivery. Chem Soc Rev 2012;41:3679–98. [10] Li Z, Barnes JC, Bosoy A, Stoddart JF, Zink JI. Mesoporous silica nanoparticles in biomedical applications. Chem Soc Rev 2012;41:2590–605. [11] Ariga K, Vinu A, Yamauchi Y, Ji Q, Hill JP. Nanoarchitectonics for mesoporous materials. Bull Chem Soc Jpn 2012;85:1–32. [12] Coll C, Bernardos A, Martinez-Manez R, Sancenon F. Gated silica mesoporous supports for controlled release and signaling applications. Acc Chem Res 2013;46:339–49.

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