Biomaterials 35 (2014) 1572e1583

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Inhibition of tumor growth and vasculature and fluorescence imaging using functionalized ruthenium-thiol protected selenium nanoparticles Dongdong Sun 1, Yanan Liu 1, Qianqian Yu 1, Xiuying Qin 1, Licong Yang 1, Yanhui Zhou 1, Lanmei Chen 1, Jie Liu* Department of Chemistry, Jinan University, Guangzhou 510632, China

a r t i c l e i n f o

a b s t r a c t

Article history: Received 3 October 2013 Accepted 2 November 2013 Available online 20 November 2013

Here we reported the high tumor targeting efficacy of luminescent Ru(II)-thiols protected selenium nanoparticles (Ru-MUA@Se). We have shown that a dual-target inhibitor Ru-MUA@Se directly suppress the tumor growth but also block blood-vessel growth. We also determined that the nanoparticles entered the cells via clathrin-mediated endocytosis pathway. In a xenograft HepG2 tumor model, we found that Ru-MUA@Se effectively inhibited tumor angiogenesis and suppressed tumor growth with low side effects using metronomic chemotherapy with Ru-MUA@Se. In vivo investigation of nanoparticles on nude mice bearing HepG2 cancer xenografts confirmed that Ru-MUA@Se nanoparticles possessed high tumor-targeted fluorescence imaging, exhibited enhanced antitumor efficacy and decreased systemic toxicity. Moreover, Ru-MUA@Se not only significantly induced dose-dependent disruption of mitochondrial membrane potential in HepG2 cells after 24 h treatment, but it also enhanced reactive oxygen species (ROS) generation. Our results suggest that the potential application of these Ru-MUA@Se nanoparticles in targeting cancer imaging and chemotherapy. Ó 2013 Elsevier Ltd. All rights reserved.

Keywords: Selenium nanoparticles Angiogenesis Fluorescence imaging In vivo distribution Targeted delivery

1. Introduction Because cancer cells in these tumors require access to blood vessels for growth and metastasis, inhibiting vessel formation offers hope for reducing the mortality and morbidity from these tumors [1]. The widely held view is that these antiangiogenic therapies should destroy the tumor vasculature, thereby depriving the tumor of oxygen and nutrients. Bevacizumab (Avastin), the first drug to block blood-vessel growth, was approved in 2004, but it and other ’angiogenesis inhibitors’ have proved disappointing in the clinic, extending patients’ lives for at best a few months. The main causes of this phenomenon are tumors don’t just rely on their host’s blood vessels for nourishment d they can make their own vasculature [2,3]. The findings offer an explanation for why a class of drug once heralded as a game-changer in cancer treatment is proving less effective than had been hoped. So Avastin and drugs like it may be targeting a pathway that is not present in those tumor-made vessels. Therefore, we need to identify either more

* Corresponding author. Tel.: þ86 20 85220223; fax: þ86 20 85221263. E-mail address: [email protected] (J. Liu). 1 Tel.: þ86 20 85220223; fax: þ86 20 85221263. 0142-9612/$ e see front matter Ó 2013 Elsevier Ltd. All rights reserved. http://dx.doi.org/10.1016/j.biomaterials.2013.11.007

than one drug to shut down blood vessels of tumor and non-tumor origin, or a common molecular mechanism affecting both the tumor and the vasculature. The emergence of bionanomaterials gave a new prospect to the researchers who have always been looking for methods to solve these problems. Cancer nanotechnology is an interdisciplinary area of research in science, engineering, and medicine with broad applications for molecular imaging, molecular diagnosis, and targeted therapy. Nanotechnology offers important new tools to detect and modulate a variety of biomedical processes that occur at the nanoparticles (NPs) entering and leaving the cells and is expected to have a revolutionary impact on biology and medicine. [4e7] For biological and clinical applications, the ability to control and manipulate the accumulation of NPs for an extended period of time inside a cell can lead to improvements in diagnostic sensitivity and therapeutic efficiency [8e10]. The main problems of much NPs lack the characteristics of fluorescence, the precise real-time imaging and the understanding of these processes are still challenging. However, current studies in this research area have focused on coating nonluminescent biorecognition molecules on the surface of NPs to mediate cellular accumulation [11], and only limited progress has been made in developing luminescent nanoparticles for molecular imaging inside living cells. So the search for fluorescent

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nanoparticles that can improve the efficiency bioanalysis and clinical diagnostics constitutes an urgent priority. The essential trace mineral, selenium, is of fundamental importance to human health. Many studies have shown that the potential for utilization of Se in a new way, to overt cancer through a combination with well-established chemotherapeutic and hormonal agents, and Se nanoparticles (SeNPs) attract increasing attention due to their excellent anticancer activities and low toxicity [12,13]. Moreover, Ru(II)-polypyridyl complexes have been intensively studied due to their photophysical properties, higher antitumor activity and more acceptable toxicity profiles [14,15]. Thiol-protected nanoparticles based on other noble metals, such as Pd, Pt, or Ir are expected to have very different properties, besides showing important applications in electron transfer processes and in photochemical and electronic devices [16]. Under this concept, we have developed a simple method to prepare bright and photostable thiols modified selenium nanoparticles with attached photosensitizers Ru(II)-polypyridyl complexes. In this manuscript, we investigated and captured the uptake process of the Ru(II)-thiols coated NP entering cells by electron microscopy, established the mechanism of uptake, and investigated intracellular apoptosis mechanism. We also provided a simple and efficient protocol for the preparation of Se nanoparticles stabilized by Ru(II)-thiols. The luminescent Ru(II)-thiols was coated on the surface of SeNPs during the reduction reaction of Na2SeO3 with NaBH4. 2. Materials and methods 2.1. Materials and reagents All reagents and solvents were purchased commercially and used without further purification unless specially noted, and Ultrapure MilliQ water (18.2 MW) was used in all experiments. Na2SeO3 and 11-Mercaptoundecanoic acid (MUA, 95%) were purchased from SigmaeAldrich Chemical Co. Fetal bovine serum (FBS) was purchased from Gibco (Life Technologies AG, Switzerland). 3-(4, 5-dimethylthiazol2-yl)-2, 5-diphenyltetrazolium bromide (MTT), N-acetylcysteine (NAC), 20 , 70 dichlorofluorescein diacetate (DCF-DA), Hoechst 33342, 5, 50 , 6, 60 -tetraethylimidacarbocyanine iodide (JC-1), LysoTracker Green and endocytosis inhibitor were from Sigma (St. Louis, MO, U.S.). Caspase-3 substrate (Ac-DEVD-AMC) was purchased from Biomol (Germany). Caspase-9 substrate (Ac-LEHD-AFC) and caspase-8 substrate (Ac-IETD-AFC) were purchased from Calbiochem. 2.2. Synthesis of ligands and complex Ruthenium(III) chloride hydrate was purchased from Alfa Aesar; 1,10phenanthroline and other materials were obtained from Sigma. The full synthesis details of 2-(4-11-mercaptoamide -N- phenyl)-imidazo [4,5f] [1,10] phenanthroline and Ru(2-(4-11-mercaptoamide-N-phenyl)-imidazo [4,5f] [1,10] phenanthroline) (phenanthroline)2(PF6)2$2H2O (Ru-MUA) [17] were according to methods in the supporting information. 2.3. Synthesis of Ru-MUA@Se Na2SeO3 (20.0 mg) is first dissolved in 2 mL of water, and then 10 mL of acetone are added. To the red solution, H3PO4 (300 mL) is added, followed by Ru-MUA (80.0 mg). Ru-MUA addition caused the solution color to change from red to redbrownish. Temperature is adjusted to 0  C by means of an ice bath, and then a freshly prepared aqueous solution (1 mL) of NaBH4(26.0 mg) is added within 5 s, causing the solution to turn immediately black. Stirring is continued at 0  C for 5 min and at RT for other 5 min, and then solvents are evaporated under vacuum and the particles washed three times with dichloromethane (10 mL portions) and three times with water (10 mL portions), with sonication and centrifugation (4500 rpm, 10 min) after each washing cycle. Briefly, TEM samples were prepared by dispersing the powder particles onto a holey carbonfilm on copper grids. The micrographs were obtained on Hitachi (H-7650) for TEM operated at an accelerating voltage at 80 kV. SEM-EDX analysis was carried out on an EX-250 system (Horiba) and employed to examine the elemental composition of Ru-MUA@Se. The size distribution of the nanoparticles was measured by PCS on a Nano-ZS instrument (MalvernInstruments Limited). 2.4. Cell culture Human umbilical vein endothelial cells (HUVECs) were isolated from freshly delivered umbilical cords using type II collagenase as previously described by Baudin et al. [18] HUVECs were characterized using von Willebrand Factor VIII and CD31. HUVECs were cultured in DMEM/F12 containing 20% fetal bovine serum (FBS,

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Gibco), 10 mg/mL heparin, 5 ng/mL bFGF and 10 ng/mL EGF at 37  C in a humidified atmosphere containing 5% CO2. Human liver hepatocellular carcinoma cell line (HepG2) were purchased from American Type Culture Collection (Manassas, VA) and maintained in RPMI 1640 or DMEM media supplemented with fetal bovine serum (10%), penicillin (100 units/ml) and streptomycin (50 units/ml) at 37  C in a humidified incubator with 5% CO2 atmosphere. 2.5. Animals Male C57/BL/6 mice (6 weeks old) were purchased from Guangdong Medical Laboratory Animal Center (Guangzhou, China). The animals were kept in an environmentally controlled breeding room (temperature: 25  1  C, relative humidity: 50  5%, 12 h dark/light cycle from 6:00 a.m. to 6:00 p.m.), with free access to sterilized tap water and commercial laboratory rodent chow. All animal experiment procedures were conducted in accordance with institutional and Chinese government guidelines for the care and use of experimental animals. 2.6. MTT assay Cell viability was determined by measuring the ability of cells to transform MTT to a purple formazan dye. (See full details according to methods in the literature) [19]. 2.7. Flow cytometric analysis Apoptosis was detected with an annexin V-FITC kit purchased from TOYOBO (Japan) according to the manufacturer’s instructions. Cells were seeded in 35 mm culture dishes and allowed to attach overnight. The cells were treated with RuMUA@Se for 24 h, collected, and washed twice with PBS. To detect early and late apoptosis, both adherent and floating cells were harvested together and resuspended in annexin V binding buffer (10 mM HEPES/NaOH pH 7.4, 140 mM NaCl, 2.5 mM CaCl2) at a concentration of 106 cells/mL. Subsequently, 5 mL of FITCconjugated annexin V and 5 mL of propidium iodide were added to 100 mL of the cell suspension (105 cells). The cells were incubated for 15 min at room temperature in the dark. Finally, 400 mL of annexin V binding buffer was added to each tube, and cells were analyzed using a FACS Calibur (BD Biosciences). 2.8. Live cell confocal microscopy 2.8.1. Live cell imaging HepG2 and HUVEC cells were grown on chamber slides to 70% confluence. RuMUA@Se (20 mg/mL) was added to the culture medium (final DMSO concentration, 0.1% v/v) and incubated for varying amounts of time at 37  C and nuclear staining was performed by Hoechst 33342 for 10 min. The cells were then washed with PBS (2  200 mL) and photographed with a Leica TCS SP5 confocal microscope (Leica Microsystems, Wetzlar, Germany) using a planapochromate 63  /NA 1.4 oil immersion objective. The confocal microscope was equipped with an ArKr laser which was used to excite RuII (488 nm excitation, detection at 560e615 nm (green) and 625e754 nm (red)). 2.8.2. Live cell imaging after treatment with LysoTracker green Cells were rinsed with PBS and incubated with 500 nM LysoTracker Green for 10 min before laser scanning confocal microscopy imaging. (LysoTracker Green channel: EX 488 nm, EM 505e525 nm bandpass. Gray channel: optical images. Yellow: colocalization of red and green fluorescence.) 2.8.3. Live cell imaging after treatment with endocytic inhibitors Inhibitor treatment required serum-free conditions throughout. Cells were treated with chloroquine, NH4Cl and colchicine at the stated concentrations for 30 min, then with Ru-MUA@Se (20 mg/mL) plus the inhibitor for 1 h before imaging. Temperature dependence studies used cells that had been cooled at 4  C for 30 min then incubated with 20 mg/mL Ru-MUA@Se (10% PBS: 90% serum-free media) at 4  C for 1 h. 2.9. TEM imaging of nanoconstructs and cancer cells For TEM, cells were incubated with Ru-MUA@Se (20 mg/mL, 12 h and 24 h) then fixed using 3% glutaraldehyde and dehydrated using ethanol. TEM samples were sectioned in Araldite resin by microtome and examined on a FEI Tecnai instrument operating at 80 kV equipped with a Gatan 1 k CCD Camera. 2.10. In vivo and Ex vivo fluorescence imaging All animal experiments (NO. SCXK2008-0002) were approved by the Institutional Animal Care and Use Committee of Jinan University. HepG2 cells (1  107) were injected subcutaneously into the right fore of the nude mice ages 5e6 weeks old. When the tumor volume reached 200e500 mm3 (about 3 weeks after inoculation), 10 mg Ru-MUA@Se were intravenously injected into the mouse models with liver cancer via the tail vein. These mice were imaged at 0.5 h, 2 h, 4 h, 8 h, and 24 h post-injection by using IVIS Lumina imaging system (Xenogen (Caliper Life Sciences), Hopkinton, MA, USA). The fluorescence signals were acquired at a lateral position on the condition of 465-nm excitation filter and DsRed emission filter. Then,

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these mice were killed. Then major organs, such as the liver, heart, lung, spleen, and kidney and the tumor, were collected to perform further fluorescence imaging observation.

(0.5 mL), and analyzed on a flow cytometer. The percentages of red and green fluorescence were quantified by Expo32TM software (Beckman CoulterÔ). The percentage of green fluorescence from JC-1 monomer was used to represent the cells that lost 6jm.

2.11. Xenograft mouse model The 5-week-old to 6-week-old severe combined immune deficiency (SCID) male mice (ordered from NIH) weighing w 20 g were divided into groups with five mice per group. HepG2 cells were s.c. injected (1  107 cells per mouse) into the mice. [20] After the tumors had become established (w 50 mm 3), the mice were s.c. injected with or without 3 mg/kg Ru-MUA@Se everyday. The mice body weights and tumor sizes were recorded everyday, and the tumor sizes were determined by Vernier caliper measurements and calculated as length  width  height. After two weeks, mice with s.c. tumors not greater than 1.5 cm in diameter were sacrificed [21].

2.18. Measurement of intracellular reactive oxygen species (ROS) generation The effects of Ru-MUA@Se on intracellular ROS generation in HUVEC and HepG2 cells were monitored by DCF-DA assay. Briefly, the cells were harvested by centrifugation, washed twice with PBS, and incubated with 10 mM DCF-DA in PBS at 37  C for 30 min. The cells were then incubated with different concentration of RuMUA@Se at 37  C for 24 h. The intracellular ROS level was examined under a fluorescence microscope with the excitation and emission wavelengths at 495 nm and 525 nm, respectively (Nikon Eclipse 80i).

2.12. Aortic ring assay The aortic ring assay was performed as described previously with some modifications [22]. Aortas isolated from SpragueeDawley rats were cleaned off periadventitial fat and cut into 1e1.5 mm long rings. Aortas were rinsed, placed on the matrigel pre-coated wells, and covered with another 100 mL matrigel. Medium containing VEGF (300 ng/mL), with or without Ru-MUA@Se, was added to the wells. After 6 days, the microvessel growth was recorded with inverted microscope and the number of branching sites was quantified with Image-Pro Plus software (Media Cybernetics, USA). The results were the means calculated from five replicates of each experiment. Three independent experiments were carried out. 2.13. Tube formation assay Matrigel was dissolved at 4  C for overnight, and each well of prechilled 24-well plates was coated with 100 mL Matrigel and incubated at 37  C for 45 min. HUVECs (4  104) were added in 1 mL endothelial cell growth medium with various concentration of complexes. After 24 h of incubation at 37  C, 5% CO2, endothelial cell tube formation was assessed with an inverted photo-microscope. Tubular structures were quantified by manual counting of low-power fields and percent inhibition was expressed using untreated wells as 100%. 2.14. Invasion assay The invasion assay was performed in Transwell (8 mm pore; Corning, Lowell, MA) pre-coated with matrigel for 8 h at 37  C. The bottom chambers were filled with 600 mL DMEM/F12 with 1% FBS supplemented with bFGF (20 ng/mL). HUVECs (5  104 cells per chamber) suspended in 100 mL DMEM/F12 with 1% FBS were seeded in the top chambers. Both top and bottom chambers contained the same concentrations of Ru-MUA@Se. Cells were allowed to invade for 24 h. Non-invaded cells were scraped with cotton swab on the top surface of the membrane and invaded cells were fixed with methanol and stained with Griess solution. The membrane was left to dry in the air. Images were taken using an Olympus IX70 inverted microscope; the invaded cells were counted in five independent areas per membrane. The results were the means calculated from five replicates of each experiment. Three independent experiments were performed.

3. Results and discussion 3.1. Preparation and characterization of Ru-MUA@Se nanoparticles A simple and efficient protocol for the preparation of Se nanoparticles stabilized by alkyl thiols containing Ru(II)-polypyridyl complex with 11-mercaptoundecanoic acid functional end-groups (Ru-MUA@Se) is proposed. Se nanoparticles were capped with Ru-MUA molecules to form more compact and stable globular nanocomposites (Scheme S1, in Supporting Information). The morphology of the Ru-MUA@Se was characterized by TEM. Fig. S1A (see in Supporting Information) clearly revealed that the assynthesized Ru-MUA@Se presented monodisperse and homogeneous spherical structure with an average diameter < 100 nm. Stability of nanomaterials is an important factor that affects its medicinal application. Therefore, the size distribution and stability of Ru-MUA@Se was investigated by a Zetasizer Nano-ZS particle analyzer. Fig. S1B (see in Supporting Information) showed that the average particles size of Ru-MUA@Se was 80.7 nm, which was similar to that of the TEM. An elemental composition analysis employing EDX showed the presence of a strong signal from the Se atoms (11.29%), together with S atom signal (7.18%), C (33.43%), O (27.41.4%) and Ru (16.25), respectively (Fig. S1C, in Supporting Information). The Cu atom signal (4.75%) was due to the copper-mesh matrix that was used to suspend the particles before loading on to the machine. 3.2. Ru-MUA@Se inhibits cell viability

2.15. Chicken chorioallantoic membrane (CAM) assay The contribution of the test compounds to angiogenesis was investigated ex vivo using the chick embryo CAM assay. Briefly, fertilized chicken eggs (10 eggs/group) were incubated at 37  C and 80% humidity. On the sixth day of incubation, a square window was opened in the shell, and CAMs were injected with different test compounds by insulin syringe. The window was sealed with a transpore tape after injection, and eggs were returned to the incubator. After 48 h of incubation, CAM arterious branches in each treatment group were photographed and counted by using a Nikon digital camera system (Chiyoda-ku, Tokyo, Japan). The antiangiogenic effect of the test compounds was presented as relative number of arterious branches. 2.16. Matrigel plug assay Matrigel (0.5 mL/plug) containing 300 ng VEGF and 150 units heparin with RuMUA@Se (20 mg/mL) was injected (S.C.) into the ventral area of the 6 weeks old male C57/BL/6 mice (five mice per group). Matrigel mixed with medium alone was used as a negative control. After 14 days of implantation, the matrigel plugs were removed and the surrounding tissues were trimmed. The matrigel plugs were fixed and embedded with paraffin. Five-micron sections were stained by hematoxylin-eosin (H&E) stain. The number of erythrocyte-filled blood vessels in high power field (HPF; 200.) was counted (plug number, 4e5; t test, P < 0.005). Three independent experiments were performed. 2.17. Determination of mitochondrial membrane potential (6jm) An aggregate-forming lipophilic dye JC-1 was used to assess the status of 6jm in cells exposed to Ru-MUA@Se. Cells cultured in 6-well plates were trypsinized and suspended in PBS buffer containing 10 mg/mL of JC-1 (0.5 mL), and then incubated at 37  C for 10 min. The cells were then collected by centrifugation, resuspended in PBS

The inhibitory effect of Ru-MUA@Se on bFGF-induced endothelial cells and HepG2 cells viability was first evaluated by MTT assay. As shown in Fig. 1A, endothelial cells were seeded in 5% FBScontaining DMEM/F12 medium without growth factors, and the viability in the presence of Ru-MUA@Se for 24 was determined after stimulation with VEGF. VEGF stimulation for 24 h increased the number of HUVECs to about 1.5 fold; Ru-MUA@Se significantly suppressed VEGF-induced HUVECs viability in a dose-dependent manner. We also investigated the effects of Ru-MUA@Se on HepG2 cells, in contrast to control, Ru-MUA@Se at concentration of 40 mg/mL significantly decreased the cancer cell viability to 19.7%, and the Ru-MUA@Se significantly inhibited the growth of HepG2 cells in a dose-dependent manner (IC50 ¼ 18.5 mg/mL, Fig. 1B). Our data suggested that Ru-MUA@Se was a potent inhibitor of VEGFinduced endothelial cell viability and the higher cytotoxic effects of Ru-MUA@Se on cancer cells HepG2. 3.3. In vitro cellular uptake of Ru-MUA@Se Flow cytometry enables the rapid measurement of ruthenium luminescence intensity for multiple cell populations. Using primarily flow cytometry, we can explore the cellular uptake mechanism of Ru-MUA and Ru-MUA@Se by comparing the ruthenium

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Fig. 1. Ru-MUA@Se inhibited cell viability. (A) Ru-MUA@Se inhibited VFGF-induced cell viability in a dose-dependent manner. HUVECs were treated with Ru-MUA@Se with or without 20 ng/mL VFGF for 24 h, and viability was measured by MTT assay. (##P < 0.01, VFGF-treated group vs. no VFGF-treated group; **P < 0.01, VFGF and Ru-MUA@Se-treated group vs. VFGF-treated group). (B) Ru-MUA@Se inhibited HepG2 cells viability in a dose-dependent manner. (xxP < 0.01).

luminescence following different incubation conditions [23,24]. As shown in Fig. 2A and B, HUVEC cells were incubated with the RuMUA and Ru-MUA@Se under different time conditions, respectively. The mean Ru-MUA luminescence increases with incubation time, from 87  7 after 2 h to 726  71 after 8 h. Interestingly, RuMUA functionalized selenium nanoparticles (Ru-MUA@Se) was significantly more sensitive to time, with the mean fluorescence increasing from 115  17 after 0.5 h to 1171  127 after 4 h, suggesting that the most abundant components of complex Ru-MUA increase uptake when they are bound strongly to selenium. As expected, similar results were observed in an assay carried out on RuMUA@Se after incubation with HepG2 cells. After 0.5, 2.0 and 4.0 h incubation with 10 mg/mL Ru-MUA loaded nanoparticles, the intracellular luminescence of Ru-MUA@Se increased to 108  7, 880  87, 1280  95, respectively, which were about 2e4 times of 10 mg/mL Ru-MUA without selenium (Fig. 2D and E). These results revealed that Ru-MUA -modified selenium nanoparticles efficiently enhances the cellular uptake of Ru-MUA without selenium decoration.

3.4. Lysosomes escape for Ru-MUA@Se to the nucleus Lysosomes are major intracellular degradation organelles. Material to be catabolized or processed is delivered to the lysosomes either by direct engulfment or by fusion of digestive vacuoles with the lysosomes [25]. Since lysosomes are the physiological barriers in organelle-targeted delivery process, visualization of the ability to locate in or escape from lysosomes is very important to evaluate drug delivery system and will provide a significant direction to improve therapy efficacy [26e28]. The interaction of Ru-MUA@Se with HepG2 cells and HUVECs has been investigated by colocalization study using LysoTracker Green. As shown in Fig. 3A, with the incubation time prolonged, Ru-MUA@Se showed well-defined time-dependent increase in fluorescence intensity. Simultaneously, the observed yellow colocalization areas of the red fluorescence from Ru-MUA@Se and the green fluorescence from LysoTracker Green also exhibited an increasing trend, which implied a gradual accumulation of Ru-MUA@Se into the lysosomes

Fig. 2. Time-dependent cellular uptake profiles of Ru-MUA and Ru-MUA@Se. Flow cytometric results of (A) HUVEC cells incubated with Ru-MUA (10 mg/mL); (B) HUVEC cells incubated with Ru-MUA@Se (10 mg/mL); (D) HepG2 cells incubated with Ru-MUA (10 mg/mL) and (E) HepG2 cells incubated with Ru-MUA@Se (10 mg/mL). Luminescence data were obtained by excitation at 488 nm with emission at 600e620 nm. The fluorescence intensity was determined by flow cytometry as described in Materials and Methods. Values expressed are means  SD of triplicates.

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Fig. 3. Laser scanning confocal microscopy images of HepG2 cells and HUVECs. Colocalization of Ru-MUA@Se (red fluorescence) and lysosomes (green fluorescence) in HepG2 cells (A) and HUVECs (B). The cells were treated with lysosomal marker LysoTracker Green and Ru-MUA@Se (20 mg/mL) at 37  C. (C) Confocalfluorescence images of Ru-MUA@Se (20 mg/ mL) in HepG2 cells and HUVECs stained with Hoechst 33342 (10 mM). (red emission from ruthenium complex, excited at 488 nm and emitted at 625e754 nm; green emission also from ruthenium complex, excited at 488 nm and emitted at 560e615 nm; blue emission from Hoechst 33342 excited at 405 nm and emitted at 420e480 nm). Scale bar: 10 um.

of HepG2 cells. This phenomenon has also been occurred in HUVECs, as shown in Fig. 3B. In the case of HepG2 cells, translocation of Ru-MUA@Se, however, does not stop at lysosomes. Confocal images showed that the high photoluminescence signals of the Ru-MUA@Se colocalized with the Hoechst 33342-stained nucleus as the incubation time increased from 0.5 to 2 h (Fig. 3C, rows 1 and 2). In contrast,

although 12 h incubation times, a large number of fluorescence dots of complex Ru-MUA, which not functionalized selenium accumulated particularly in the perinuclear of HepG2 cells (Fig. 3C, row 3). Surprisingly, as shown in Fig. 3C (rows 4 and 5), there were the high concentrations of particles near the nucleus of HUVECs treated with Ru-MUA@Se and Ru-MUA for 2 h and 12 h, respectively. This finding highlights the role that Ru-MUA@Se serves as a carrier of

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high, local concentrations of aptamer that can be delivered in the nucleus of HepG2 cells. The results also demonstrate that RuMUA@Se is able to escape the lysosomes to avoid any potential degradation and enter the nucleus to impart therapy with high efficacy. 3.5. Understanding the mechanisms of cellular uptake We further evaluated the mechanism of uptake of Ru-MUA@Se. First, we evaluated whether the uptake of Ru-MUA@Se were due to energy-dependent pathways. In these experiments, Ru-MUA@Se was incubated with the HepG2 cells and HUVECs at low temperature (4  C instead of 37  C) environments, respectively (Fig. 4A). If the nanoparticles entered via energy-dependent pathways, a decrease in the uptake of Ru-MUA@Se would be observed [8,29]. Indeed, this was the case. The phenomenon suggests that energy plays a very important role in the process of Ru-MUA@Se entering cells. Next, we determined if uptake was likely due to a clathrinmediated process. Chlorpromazine is an amphiphilic drug that prevents the recycling of clathrin proteins from endosomes back to the cell membrane, thus inhibiting the formation of new clathrincoated pits. In this case, Ru-MUA@Se uptake was significantly reduced (Fig. 4A); thereby suggesting that clathrin-mediated endocytosis is the main route of internalization. Furthermore, to discern any role that caveolae-mediated endocytosis may play in nanoparticle uptake, genistein, a tyrosine kinase inhibitor known to sequester and alter cholesterol-rich domains within the plasma membrane, were used. Both HepG2 cells and HUVECs treated with genestein did not affect uptake, suggesting that the caveolaemediated pathway probably did not play a significant role (Fig. 4A). Lysosomotropic agents, such as chloroquine, are nonspecific weak bases that diffuse across membranes in a concentrationdependent manner, thereby neutralizing the pH of endocytic

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vesicles [30]. Endosomal acidification is one of the hallmarks of clathrin-mediated endocytosis. Surprisingly, pretreatment of the HepG2 cells and HUVECs with chloroquine resulted in a slight decrease of fluorescence intensity, indicating that the internalized Ru-MUA@Se can escape from endocytic vesicles (endo-lysosomal vesicles) and resist the lysosomal degradation (Fig. 4A). To confirm the different activity of certain inhibitors, we used flow cytometric to reevaluate the potential redistribution of RuMUA@Se nanoparticles (Fig. 4B). Relative fluorescence intensity shows the differences between the cellular uptake of Ru-MUA@Se without and with inhibitor. We again proved a reduced cellular uptake in the same cases as observed in Fig. 4B, such as RuMUA@Se with chlorpromazine in HUVECs and HepG2 cells. On the other hand, we could show no fluorescence intensity change in cellular uptake as we previously found by confocal images, such as Ru-MUA@Se with genistein or with chloroquine in HUVECs and HepG2 cells, respectively. Altogether, we could visualize the same effects of the inhibitors by flow cytometric as observed in the confocal luminescence image experiments. Results were similar for both HepG2 cells and HUVECs, suggesting that the mechanism of uptake of Ru-MUA@Se was similar. We also probed the mechanism of uptake of Ru-MUA, the complex is taken up by live cells via temperature-dependent and non-endocytotic mechanism (data not shown). 3.6. Nanoparticle delivery to HUVEC and HepG2 cells Transmission electron microscopy (TEM) is a powerful and unique technique for structure characterization. Thus, in order to try and understand the nanoparticles uptake in more detail, we made further investigations with TEM to gain information about the uptake, fate, and localization of nanoparticles inside the cells (Fig. 5). After 12 h of incubation with HepG2 cells (Fig. 5AeC), Ru-

Fig. 4. Elucidating the mechanism of Ru-MUA@Se nanoparticle uptake. (A) Confocal luminescence image of living cells were pretreated with endocytosis inhibitors and then incubated with 20 mg/mL Ru-MUA@Se at 37  C for 1 h. (B) Relative cells uptake by flow cytometric in the presence of inhibitors were expressed as percentages compared to those without inhibitors. Error bars represent standard error of the mean (s.e.m.) with n ¼ 3.

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MUA@Se nanoparticles were found clustered in cellular compartments that appear to the nuclear membrane, and for Ru-MUA@Se nanoparticles, the nuclear membranes were more likely to fold inward in order to absorb the material from outside, suggesting the delivery of nanoparticles carrying ruthenium complexes was nuclear targeting. More importantly, at longer times (24 h), most of the Ru-MUA@Se were grouped in the cell nucleus may be by endocytosis. Fig. 5DeF showed major changes in nuclear phenotype near the site of the nanoparticles and we found that increased incubation times of the nanoparticles resulted in even deeper invaginations or multibranched folds of the nuclear envelope at 24 h, which strongly implies that the presence of the nuclear envelope produced the localized changes in the cancer cell nuclear envelope. Apparently, Ru-MUA@Se nanoparticles were able to enter HepG2 cells and escape the endosome/lysosome pathway, and were able to interact properly with the nuclear pore complex. As shown in Fig. 5GeI, Ru-MUA@Se nanoparticles accumulated around the nuclear membrane after 12 h of incubation with HUVEC cells. Not surprisingly, after 24 h Ru-MUA@Se nanoparticles were found clustered around the cytoplasmic side of the nuclear membrane with relatively few located inside the nucleus (Fig. 5JeL). These data also correlate well with the confocal microscopy results and close observations along the cell membrane suggested that the RuMUA@Se nanoparticles diffused across the cellular boundary, causing severe impairment to the intracellular organelles and therefore promoting cell death. 3.7. In vivo imaging and biodistribution Xenograft tumors in nude mice have been reported to be more invasive than in situ carcinoma and can form the construction of a

capillary network. It would allow the enrichment of nanomedicines in tumors due to the EPR effect. Fig. 6A shows the realtime images of nanoparticles in the tumor-bearing mice, in which the whole bodies of live mice were monitored at 0.5 h, 2 h, 4 h, 8 h and 24 h after administration, respectively. During the living imaging test, there were no signals in the tumor tissues in the blank group (not injected Ru-MUA@Se), suggesting no autofluorescence in vivo images and the fluorescence signal intensity of in vivo imaging is close to a true reflection of the Ru-MUA@Se retained inside the organs. With the time prolong, the fluorescence signals were clearly accumulated in liver and kidney after intravenous administration of Ru-MUA@Se at 0.5 h, and Ru-MUA@Se signals were almost no detected in tumor tissue. However, preferential accumulation of fluorescence was obvious in the tumor site other than liver or other normal tissues at 8 h after injection. For RuMUA@Se, the signal at the tumor site increased significantly at 2 h, peaked at 4 h, and then decreased slowly within 24 h (Fig. 6A). Then, these mice were killed. Then major organs, such as the liver, heart, lung, brain, and kidney and the tumor, were collected to perform further fluorescence imaging observation. As shown in Fig. 6B, ex vivo imaging further confirmed the obvious fluorescence signal in mice (Fig. 6A). The results demonstrated that most of Ru-MUA@Se accumulated in the tumor, followed by liver and kidneys 4 h after injection. Moreover, at 24 h after nanoparticles injection, obvious decrease of Ru-MUA@Se signals in the tumor were observed, which is in agreement with previous results. This high tumor target ability of nanoparticles might be due to a combination of an EPR effect and receptor-mediated uptake of nanoparticles. Overall, in vivo biodistribution studies fully demonstrate that Ru-MUA@Se could target and migrate to the liver cancer cells in vivo.

Fig. 5. TEM images of HepG2 and HUVEC cells treated with 20 mg/mL Ru-MUA@Se for (A) 12 h, (D) 48 h, (G) 12 h, and (J) 48 h, respectively. Representative high magnification TEM micrographs of a part of HepG2 cells for 12 h (B and C), HepG2 cells for 24 h (E and F), HUVEC cells for 12 h (H and I), and HUVEC cells for 12 h (K and L), respectively. N stands for nucleus and the black arrows indicate Ru-MUA@Se nanoparticles within a cell.

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Fig. 6. (A) In vivo imaging of tumor-bearing mice after administration of Ru-MUA@Se at 0.5 h, 2 h, 4 h, 8 h and 24 h. (B) Ex vivo fluorescence images of tissues including heart, liver, spleen, lung, kidney and tumor collected at 4 h post-injection of Ru-MUA@Se (a) at 24 h post-injection of Ru-MUA@Se (b).

3.8. In vivo antitumor efficacy To observe the anti-tumor activity and systemic toxicity of RuMUA@Se, we applied different concentrations of Ru-MUA@Se in a liver tumor (HepG2) xenograft model. We found that administration of 3 mg/kg/d Ru-MUA@Se for 15 days substantially suppressed tumor volume (Fig. 7A) and reduced tumor weight (Fig. 7C), the average tumor volume in the control mice increased from 52.83  2.67 to 1165.37  72.20 mm3 after 15 days, whereas the average tumor volume in the Ru-MUA@Se-treated mice

increased only from 51.36  1.95 to 285.39  25.71 mm3. Additionally, the average tumor weight in the control group was 0.26  0.08 g, whereas the average tumor weight in Ru-MUA@Setreated group was only 0.031  0.0007 g (Fig. 7C), a significant inhibition in tumor growth. Fig. 7B showed the variation of relative body weight of the mice with time. Compared to the initial body weights of tumor-bearing mice, no significant body weight loss was observed after the administration of Ru-MUA@Se, showing that Ru-MUA@Se were well-tolerated at the tested dose level.

Fig. 7. Ru-MUA@Se inhibits tumor growth in vivo. HepG2 cells were injected s.c. (1  107 cells per mouse) into the 5-week-old to 6-week-old SCID male mice. After the tumors had established (w50 mm3), the mice were injected with or without 3 mg/kg/d Ru-MUA@Se. After two weeks, mice were sacrificed and tumors were removed and taken images by Nikon camera. A, tumors size of control group and treated with Ru-MUA@Se group; B, the average mouse body weight of control group and treated with Ru-MUA@Se group; C, tumors from the mice with Ru-MUA@Se treatment were significantly smaller than that from control group. Columns, mean; bars, SE (n ¼ 5; t test, P < 0.001).

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3.9. Nanoparticles enhanced immunity and resistibility of tumorbearing mice To further determine the antitumor effect of Ru-MUA@Se therapy in vivo, HepG2 tumor tissues were stained with hematoxylin and eosin or were detected by the TUNEL assay. As shown in Fig. 8A, the tumors treated with Ru-MUA@Se demonstrated the number of microvessels was significantly decreased in vivo. Apoptotic cell death was further confirmed by DNA fragmentation and nuclear condensation as detected by TUNEL enzymatic labeling and DAPI co staining assay. TUNEL can detect the early stage of DNA fragmentation in apoptotic cells prior to changes in morphology. According to the results of TUNEL assay in which apoptosis cells were labeled green FL (Fig. 8B), Ru-MUA@Se led to a significant increase in DNA fragmentation and nuclear condensation in the HepG2 cells. Taken together these results indicated that the cell death induced by the Ru-MUA@Se is mainly caused by apoptosis. 3.10. Ru-MUA@Se inhibits angiogenesis in vitro and in vivo To examine the inhibitory effect of Ru-MUA@Se on angiogenesis, we performed aortic ring assays using isolated aortas from mice. The 1-mm-long to 1.5-mm-long aortic rings were put on Matrigel and covered by another Matrigel layer and ECGM with or without Ru-MUA@Se. As shown in Fig. 9A, VEGF in Matrigel can dramatically induce microvessel sprouting, whereas addition of 20 mg/mL Ru-MUA@Se significantly antagonizes microvessel sprouting, suggesting that Ru-MUA@Se dramatically inhibited angiogenesis in vitro. To further verify the inhibitory effect of Ru-MUA@Se on VEGFinduced angiogenesis, we used Matrigel plug assay for antiangiogenesis effect of Ru-MUA@Se in vivo. As shown in Fig. 9B, matrigel plugs containing VEGF excised from mice were dark red and filled with blood vessels, indicating that functional vasculatures had

formed. In contrast, the color of Matrigel plugs containing VEGF plus 20 mg/mL Ru-MUA@Se group was significantly pale, indicating less blood vessel formation. The number of neovessels was analyzed and quantified after being H&E stained (Fig. 9C). RuMUA@Se at 20 mg/mL strongly inhibited the vessel number and the formation of microvessels. Together, these results indicated that Ru-MUA@Se was capable of suppressing VEGF-induced neovessel formation in vitro and in vivo. 3.11. Ru-MUA@Se inhibits bFGF-induced HUVEC invasion and tube formation To determine the effect of Ru-MUA@Se on angiogenesis, we firstly examined how Ru-MUA@Se regulates capillary tubule formation of HUVECs. As shown in Fig. S2A (See in Supporting Information), HUVECs formed a robust and complete tube network in the presence of bFGF within 8 h post-seeding. However, 10 mg/mL Ru-MUA@Se partially abolished this process with the reduction of number and length of tube-like structures, and the capillary tube was completely disassembled in the presence of 20 mg/mL RuMUA@Se, which exhibited stronger inhibitory effect compared to Ru-MUA and positive control suramin. To further assess the antiangiogenic property of Ru-MUA@Se in vitro, we examined its inhibitory effects on the chemotactic motility of endothelial cells using transwell assay (Fig. S2B, in Supporting Information). As shown in Fig. S2B, a large number of cells migrated to the lower side of membrane in the transwell chamber after stimulation with bEGF. However, the number of invasive cells was dramatically reduced in the presence of RuMUA@Se, suggestive of a potent inhibitory effect of Ru-MUA@Se on bFGF-induced endothelial cell motility. These data were also similar to suramin. Together, these results indicated that RuMUA@Se could block bFGF-induced angiogenesis in vitro by inhibiting cell motility, and endothelial cell tubular structure formation.

Fig. 8. A, effects of Ru-MUA@Se on tumor angiogenesis in xenograft mouse tumor model. Tumors were fixed with Histochoice MB (Molecular Biology) tissue fixative (Amresco) and embedded with paraffin. The 5-mm sections were stained with H&E (magnification, 200). B, Detection of cell apoptosis in tumor tissue of each group by TUNEL dyeing method (Magnification 400).

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Fig. 9. Ru-MUA@Se inhibits microvessel sprouting ex vivo and angiogenesis in vivo. A, morelloflavone inhibits microvessel sprouting in mouse aortic ring assay. Aortic segments isolated from SpragueeDawley rats were placed in the Matrigel-covered wells and treated with VEGF in the presence or absence of Ru-MUA@Se. B and C, Ru-MUA@Se inhibits angiogenesis in Matrigel plug assay. Six-week-old C57/BL/6 mice were injected with 0.5 mL of Matrigel containing 20 mg/mL Ru-MUA@Se, 300 ng of VEGF, and 20 units of heparin into the ventral area (n ¼ 5 per group). After 6 d, representative Matrigel plugs were removed and photographed in B. The Matrigel plugs were fixed with formalin and 5-mm sections were stained with H&E (magnification, 200) staining in C. The number of vessels in HPF was counted in the presence or absence of Ru-MUA@Se at 20 mg/mL, respectively. Columns, mean (n ¼ 5); bars, SE. **, P < 0.01 vs. VEGF alone.

3.12. Assessment of CAM blood vessels The antiangiogenic effect in vivo was evaluated on chicken chorioallantoic membrane (CAM) neovascularisation model. As shown in Fig. S2C (in Supporting Information), angiogenesis of fertilized eggs was clearly observed after 48 h of treatment, for bFGF-treated CAMs (80 ng/CAM) on day 7, greater effects on angiogenesis were observed at 48 h and normalized values for rates of vascular growth relative to untreated CAMs. bFGF stimulated the growth of smaller vessels in the chicken CAM. Ru-MUA@Se started to significantly inhibit CAM angiogenesis at a low concentration of 5 mg/mL/egg, and completely inhibited the angiogenesis at 20 mg/ mL/egg. The result showed the amount of branch vessels, particularly smaller vessels, was significantly decreased. Ru-MUA@Se could largely reduce the angiogenesis on the chorio-allantoic membrane of fertilized eggs as compared to those of the control group, as well as the function of suramin in early lesion formation in the CAM model. 3.13. Analysis of apoptosis by PI staining The propidium iodide (PI) flow cytometric assay has been widely used for the evaluation of apoptosis in different experimental models. It is based on the principle that apoptotic cells, among other typical features, are characterized by DNA fragmentation and, consequently, loss of nuclear DNA content [31]. As shown in Fig. 10A, exposure of the HepG2 cells to different concentrations of Ru-MUA@Se for 48 h resulted in marked dosedependent increase in the proportion of apoptotic cells as reflected by the subdiploid peak. The values of sub-G1 varied from 2.26% to as high as 37.64% with increasing treated concentration. However, HUVEC treated with different concentrations of RuMUA@Se for 48 h, we were not able to detect a significant increase in the percentage of apoptosis. Caspases are the key effector

molecules of the physiological death process known as apoptosis, although some are involved in activation of cytokines, rather than cell death [32]. There are two well characterized activation pathways for apoptosis: one initiated by the cell surface death receptors (the so called extrinsic pathway, operating through caspase-8) and the other triggered by changes in mitochondrial integrity (known as the intrinsic pathway, utilizing caspase-9) [33,34]. To confirm the activation of both pathways by Ru-MUA@Se in HUVEC and HepG2 cells, we used specific caspase inhibitors to determine which, if any, would block Ru-MUA@Se’s effects. As shown in Fig. 9B, RuMUA@Se-induced apoptotic cell death was remarkably suppressed by pretreatment with “pan-caspase” inhibitor z-VAD-fmk, suggesting that Ru-MUA@Se-induced apoptotic cell death occurs mainly in a caspase-dependent manner. The caspase-8 specific inhibitor (z-IETD-fmk) as well as the caspase-9 specific inhibitor (zLEHD-fmk), each partially abrogated Ru-MUA@Se-induced apoptosis in HUVEC and HepG2 cells (Fig. 10B), whereas the z-IETDfmk was nearly as potent as the pan-caspase inhibitor in HepG2 cells. These data suggest that both the intrinsic and extrinsic pathways of caspase cascades contribute to Ru-MUA@Se induced apoptosis in HUVEC and HepG2 cells. 3.14. Ru-MUA@Se induce mitochondria dysfunction via ROS overproduction Mitochondria play a major role in apoptosis triggered by many stimuli; loss of mitochondrial transmembrane potential is a key event in apoptosis focus on mitochondria. HUVEC and HepG2 cells were untreated or treated with Ru-MUA@Se for 24 h, stained with JC-1, and analyzed by flow cytometry. Fig. 11A shows a clear increase in the percentage of cells (lower right quadrants) that emitted only green fluorescence after Ru-MUA@Se treatment, representing cells with depolarized mitochondrial membranes. The number of cells with loss of or reduced mitochondria membrane

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Fig. 10. (A) Analysis of cells apoptosis by PI staining. HUVEC and HepG2 cells treated with different concentrations of Ru-MUA@Se for 48 h were collected and stained with PI after fixation by 70% ethanol. Following flow cytometry, cellular DNA histograms were analyzed by the MultiCycle software. (B) Effects of caspase inhibitors (40 mM) on apoptosis induced by Ru-MUA@Se. Cells were pretreated with z-VAD-fmk (general caspase inhibitor), z-IETD-fmk (caspase-8 inhibitor), and z-LEHD-fmk (caspase-9 inhibitor) for 2 h followed by coincubation with Ru-MUA@Se for 24 h. Apoptotic cells were determined by flow cytometry. All results were obtained from three independent experiments.

potential (6jm) increased over time after the addition of RuMUA@Se and was dose-dependent for HUVEC and HepG2 cells suggesting that Ru-MUA@Se triggered apoptotic pathways in cells through induction of mitochondria dysfunction. The mitochondrial respiratory chain is the major source of intracellular ROS generation and, at the same time, the generation of intracellular ROS may be

related to mitochondrial dysfunction and the induction of apoptosis [35]. The intracellular ROS generation in HUVEC and HepG2 cells treated by Ru-MUA@Se was measured by fluorescence method. This assay is based on the cellular uptake of a nonfluorescent probe (DCFH-DA), which is subsequently hydrolyzed by intracellular esterase to form dichlorofluorescein, DCFH. As shown

Fig. 11. Ru-MUA@Se induced the depletion of mitochondrial membrane potential (6jm) and intracellular ROS level in HUVEC and HepG2 cells, respectively. (A) Cells treated with Ru-MUA@Se for 24 h were harvested and stained with JC-1 and then analyzed by flow cytometry. The number in the down region of each dot plot represents the percentage of cells that emit green fluorescence due to the loss of 6jm. (B) Cells incubated with 10 mM DCFeDA in PBS for 30 min were treated with different concentrations of Ru-MUA@Se for 24 h at 37  C, and then imaged under a fluorescence microscope. (C) Quantitative analysis of DCF fluorescence intensity of HUVEC and HepG2 cells treated with Ru-MUA@Se, respectively. (D) Effects of NAC on ROS generation induced by Ru-MUA@Se. Cells were pretreated with 2 mM NAC for 4 h and then exposed to 10 mg/mL Ru-MUA@Se for 24 h. All results were obtained from three independent experiments.

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in Fig. 11B, Ru-MUA@Se treatment dose-dependently increased DCF fluorescence intensity in HUVEC and HepG2 cells, indicating the up-regulation of intracellular ROS levels, and Fig. 11C showed ROS generation in HepG2 cells treated with Ru-MUA@Se increased slightly stronger than that of in HUVEC cells. To further examine the important role of ROS generation in Ru-MUA@Se-induced apoptosis, we next investigate the effects of a thiol-reducing antioxidant NAC on intracellular ROS generation. The data revealed that NAC effectively prevented the increase in ROS overproduction (Fig. 11D) induced by Ru-MUA@Se. These results suggest that RuMUA@Se-induced apoptosis in HepG2 cells is ROS-dependent. 4. Conclusion In this paper we have described a simple and highly efficient procedure for the synthesis of luminescent Ru(II)-thiols protected Se nanoparticles with small average NPs diameter and narrow NPs size distribution. We have shown that a dual-target inhibitor RuMUA@Se directly suppress the tumor growth but also block blood-vessel growth. Moreover, by using fluorescence confocal microscopy and TEM imaging studies, we have demonstrated their cellular uptake and localization within the cytoplasm and nucleus. We demonstrate the efficacy of Ru-MUA@Se delivery both in vitro and in vivo to solid tumors after systemic administration in a mouse tumor-xenograft model. The nanocarrier tested exhibited biocompatibility, formed nanoparticles of optimal size and improved localization within tumor tissue promoted by the EPR effect characteristic to tumor vasculature. The results also indicate that clathrin-mediated endocytosis plays an important role in the Ru-MUA@Se uptake and contributes to the more uptake and easier enter the nucleus of Ru-MUA@Se in HepG2 cells than in HUVECs. After intracellular uptake by endocytosis, Ru-MUA@Se escapes out of the endolysosomes from degradation. Furthermore, RuMUA@Se-induced apoptosis was also found dependent on ROS generation. Although further investigations are warranted to evaluate the potential use of Ru-MUA@Se nanoparticles, these results at least suggest the potential application of these nanoparticles in tumor growth inhibiting and tumor imaging. Acknowledgments This work was supported by the National Natural Science Foundation of China (20871056, 21171070 and 21371075), the Planned Item of Science and Technology of Guangdong Province (c1011220800060), and the Fundamental Research Funds for the Central Universities. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.biomaterials.2013.11.007. References [1] Jain RK. Normalization of tumor vasculature: an emerging concept in antiangiogenic therapy. Science 2005;307:58e62. [2] Wang R, Chadalavada K, Wilshire J, Kowalik U, Hovinga KE, Geber A, et al. Glioblastoma stem-like cells give rise to tumour endothelium. Nature 2010;468:829e33. [3] Ricci-Vitiani L, Pallini R, Biffoni M, Todaro M, Invernici G, Cenci T, et al. Tumour vascularization via endothelial differentiation of glioblastoma stem-like cells. Nature 2010;468:824e8. [4] Fu CC, Lee HY, Chen K, Lim TS, Wu HY, Lin PK, et al. Characterization and application of single fluorescent nanodiamonds as cellular biomarkers. PNAS 2007;104:727e32.

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