Biodegradable Hollow Mesoporous Organosilica Nanotheranostics for

Jan 31, 2018 - School of Radiation Medicine and Protection & School for Radiological and Interdisciplinary Sciences (RAD-X), Medical College of Soocho...
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Biodegradable Hollow Mesoporous Organosilica Nanotheranostics for Mild Hyperthermia-Induced Bubble-Enhanced Oxygen-Sensitized Radiotherapy Nan Lu,†,‡,§ Wenpei Fan,*,§ Xuan Yi,⊥ Sheng Wang,§ Zhantong Wang,§ Rui Tian,§ Orit Jacobson,§ Yijing Liu,§ Bryant C. Yung,§ Guofeng Zhang,∥ Zhaogang Teng,† Kai Yang,⊥ Minming Zhang,‡ Gang Niu,§ Guangming Lu,*,† and Xiaoyuan Chen*,§ †

Department of Medical Imaging, Jinling Hospital, Medical School of Nanjing University, Nanjing, Jiangsu 210002, China Department of Radiology, the Second Affiliated Hospital, Zhejiang University School of Medicine, Hangzhou, Zhejiang 310009, China § Laboratory of Molecular Imaging and Nanomedicine (LOMIN), National Institute of Biomedical Imaging and Bioengineering (NIBIB), National Institutes of Health (NIH), Bethesda, Maryland 20892, United States ⊥ School of Radiation Medicine and Protection & School for Radiological and Interdisciplinary Sciences (RAD-X), Medical College of Soochow University, Suzhou, Jiangsu 215123, China ∥ Laboratory of Cellular Imaging and Macromolecular Biophysics, National Institute of Biomedical Imaging and Bioengineering (NIBIB), National Institutes of Health (NIH), Bethesda, Maryland 20892, United States ‡

S Supporting Information *

ABSTRACT: Alleviation of tumor hypoxia has been the premise for improving the effectiveness of radiotherapy, which hinges upon the advanced delivery and rapid release of oxygen within the tumor region. Herein, we propose a “bubbleenhanced oxygen diffusion” strategy to achieve whole tumor oxygenation for significant radiation enhancement based on the “bystander effect”. Toward this end, sub-50 nm CuS-modified and 64Cu-labeled hollow mesoporous organosilica nanoparticles were constructed for tumor-specific delivery of O2-saturated perfluoropentane (PFP). Through the aid of PFP gasification arising from NIR laser-triggered mild hyperthermia, simultaneous PET/PA/US multimodality imaging and rapid oxygen diffusion across the tumor can be achieved for remarkable hypoxic radiosensitization. Furthermore, the multifunctional oxygencarrying nanotheranostics also allow for other oxygen-dependent treatments, thus greatly advancing the development of bubble-enhanced synergistic therapy platforms. KEYWORDS: mesoporous organosilica, bubble, tumor oxygenation, radiosensitization, multimodal imaging

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compared with the recently reported method of in situ oxygen production, which relies heavily on the tumor microenvironment,11,12 the tumor-specific oxygen delivery may serve as a more effective and versatile strategy for oxygenation of various hypoxic tumors, which thus underscores the importance of the pursuit of high-performance oxygen-carrying compounds. Owing to the high affinity toward oxygen molecules, perfluorocarbons (PFCs) have been used as preferential

eaturing high body penetration length and precise positioning, radiotherapy (RT) has been widely used for noninvasive treatment of deep-seated tumors inside the body with minimal side effects.1,2 However, owing to the insufficient oxygen supply in blood vessels during rapid tumor growth, and the increasd oxygen diffusion distance as well as decreased oxygen transport capacity with tumor propagation/ expansion, there exists severe hypoxia within most solid tumors, which causes strong resistance to RT.3−5 In view of the indispensable role of oxygen in increasing the sensitivity of hypoxic cancer cells to X-ray radiation and fixing the RTinduced DNA damage,6 multiple strategies have been proposed to enhance the RT efficacy via radiosensitization.7−10 However, © 2018 American Chemical Society

Received: November 17, 2017 Accepted: January 30, 2018 Published: January 31, 2018 1580

DOI: 10.1021/acsnano.7b08103 ACS Nano 2018, 12, 1580−1591

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Scheme 1. Schematic of the Construction of O2−PFP@ HMCP for PET/PA/US Imaging, Mild-HyperthermiaInduced PFP Bubble Release, and Oxygen-Sensitized Radiotherapy

oxygen-carrying candidates.13,14 The particular interest in PFCs stems from their ability to provide a strong mechanical driving force for the fast release and distribution of oxygen throughout the whole tumor via hyperthermia-induced liquid−gas transformation,15 which contributes greatly to the improved tumor oxygenation. Considering the hydrophobicity of PFC, a biocompatible nanocarrier with a hollow cavity is necessary to encapsulate the PFC liquid.16 Importantly, the size of the hollow-structured nanocarrier should be restricted below 50 nm to avoid rapid uptake by the reticuloendothelial system (RES) and simultaneously achieve rich tumor accumulation through the enhanced permeability and retention (EPR) effect.17,18 For clinical purposes, the nanocarrier must be biodegradable to avoid long-term toxicity.19 In addition, facile surface modification is an additional requisite, allowing for structural multifunctionalization of the nanocarrier. To satisfy the above multifaceted criteria, herein, biocompatible and biodegradable sub-50 nm hollow mesoporous organosilica nanoparticles (HMONs) have been successfully constructed for efficient storage of hydrophobic perfluoropentane (PFP, one type of PFC with low boiling point of 29 °C)20 liquid. By taking advantage of thiol group modification, ultrasmall CuS nanoparticles and 64Cu can be attached firmly and chelated stably on the surface of HMONs to realize photoacoustic (PA) imaging and positron emission tomography (PET) imaging, respectively. Benefiting from the nonhazardous mild hyperthermia upon low-power near-infrared (NIR) laser irradiationof CuS, the liquid PFP can be gasified into bubbles to not only enhance the tumor cell uptake of HMONs but also intensify the ultrasound (US) imaging signal. It is hypothesized that the generated bubbles can greatly promote the free diffusion of oxygen from cell to cell inside tumors. The “bystander effect” results in homogeneous tumor oxygenation for remarkably enhanced radiosensitization. Prospectively, the developed bubble-enhanced oxygen-sensitized RT along with judiciously designed oxygen-carrying nanotheranostics in this study will open up new dimensions for multimodal PET/PA/US image-guided radiosensitization of various tumors (Scheme 1) without endogenous oxygen or depth limitations.

Ultrasmall CuS nanoparticles are well-known for strong absorbance in the NIR region and high photothermal conversion efficiency,23,24 so the direct conjugation of CuS onto the surface of HMON (HMON@CuS) through thiol groups may endow excellent PA and photothermal performance. The white to dark green color change (Figure 1c) and the energy-dispersed X-ray (EDX) spectrum (Figure S2) confirm the successful formation of HMON@CuS with 40 wt % Cu, as measured by inductively coupled plasma (ICP) analysis. However, CuS decoration did not have a notable effect on the hollow structure and particle size of HMON (Figures 1c and S1c). Finally, PEG 2000 was modified onto the surface of HMON@CuS (HMON@CuS-PEG, denoted as HMCP) to further improve the dispersity and stability, as shown by the narrow dynamic light scattering size distribution (Figure S3). The Fourier transform infrared (FT-IR) spectra and ζ-potential change also confirm the successful conjugation of CuS and PEG (Figure 1d,e). Moreover, HMON@CuS exhibits a large surface area of 282.4 m2/g (Figure S4a) and mesopore size of around 3.88 nm (Figure S4b), which allows for sufficient encapsulation of a large variety of payloads, including hydrophobic cargoes. As the disulfide bonds in the framework of HMON are inclined to be cleaved in the reductive tumor microenvironment,25,26 HMON was found to be gradually degraded in simulated glutathione (GSH) solutions (Figure S5). Accordingly, HMCP exhibits a similar time-dependent biodegradable behavior (Figures 1 and S6). Meanwhile, the dissociated ultrasmall CuS nanoparticles arising from the biodegradation of HMCP can be eliminated through feces and urine,27 which faclitates the in vivo excretion of CuS and further improves the biosafety of HMCP, thus promising for biological applications. Owing to the hollow cavity, HMON is an ideal nanocarrier for encapsulating PFP via vacuum impregnation.15,28 In contrast to the apparent phase separation between free PFP and phosphate-buffered saline (PBS) (Figure S7a), PFP-loaded

RESULTS AND DISCUSSION Distinguished from the preparation of large HMONs by using strong acid and alkali as etching agents,21 herein, a mild “ammonia-assisted selective etching” method was developed to make sub-50 nm HMONs. In brief, according to the chemical homology principle, the core/shell structured mesoporous SiO2/organosilica nanoparticles (denoted as MSN@MON, Figure 1a) were fabricated through the co-hydrolysis and cocondensation of tetraethoxysilane and bis[3-(triethoxysilyl)propyl]tetrasulfide by using cetyltrimethylammonium chloride as the pore-forming agent. It should be emphasized that the addition of 0.1 g of triethanolamine22 as the catalyst is the key to restrict the size of MSN@MON below 50 nm. Considering that the Si−C bonds within the MON shell are more stable and stronger than the Si−O bonds within the MSN core, we strategically used ammonia to selectively etch away the inner MSN core, thus yielding disulfide-bridged HMON (Figure 1b and Figure S1a,b). By using mild ammonia instead of strong HF/NaOH as the etching agent, the obtained HMON demonstrated a uniform hollow spherical morphology with an average diameter of around 40 nm. 1581

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Figure 1. TEM images of (a) MSN@MON, (b) HMON, and (c) HMON@CuS. Inset of (b): Photograph of HMON aqueous solution. Inset of (c): Photographs of HMON@CuS aqueous solutions. (d) FT-IR spectra of HMON and HMCP. The new bands at 2878 and 1464 cm−1 in HMCP indicate C−H stretching in PEG. (e) ζ-Potentials of HMON, HMON@CuS, and HMCP. The changes in ζ-potentials demonstrate successful conjugation of CuS and PEG. TEM images of biodegradable HMCP immersed in 10 mM GSH aqueous solution for (f) 3 days, (g) 2 weeks, and (h) 3 weeks.

Figure 2. (a) UV−vis spectra of HMON and HMCP. (b) Photothermal stability of HMCP within five cycles of NIR laser irradiation. (c) Temperature increase profiles of HMCP aqueous solution with different concentrations upon NIR laser irradiation (power density: 0.5 W/ cm2) for 3 min. (d) Temperature increase profiles of HMCP aqueous solution upon NIR laser irradiation with different power densities (concentration of HMCP: 100 μg/mL) for 3 min. (e) Photothermal images of HMCP aqueous solution with different concentrations upon NIR laser irradiation (laser power density: 0.5 W/cm2, upper row) for 3 min and 100 μg/mL of HMCP aqueous solution upon NIR laser irradiation with various power densities (bottom row) for 3 min.

nm) laser irradiation (Figure 2b). According to the linear regression curve between the cooling stage and negative natural logarithm of driving force temperature of HMCP (Figure S8), the photothermal conversion efficiency of HMCP was calculated to be 73.9%. Meanwhile, the photothermal effect of HMCP is dependent on its concentration as well as NIR laser power density (Figure 2c−e), which allows for temper-

HMON (PFP@HMON) can be well-dispersed in PBS (Figure S7b), indicating the efficient encapsulation of PFP into HMON without leakage. Moreover, by loading PFP into the cavity of HMCP (PFP@HMCP), the strong NIR absorption of HMCP (Figure 2a) makes the liquid−gas phase transformation of PFP possible, as HMCP demonstrates excellent yet stable photothermal performance within at least five cycles of NIR (808 1582

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Figure 3. Optical microscopy images of PFP@HMCP aqueous solution (a) before and after NIR laser irradiation for (b) 10 and (c) 20 s. Optical microscopy images of O2−PFP@HMCP aqueous solution (d) before and after NIR laser irradiation for (e) 10 s and (f) 20 s. (g) O2 concentration changes of water after adding PFP@HMCP with or without NIR laser irradiation under a N2 atmosphere. (h) O2 concentration changes of water after adding O2−PFP@HMCP with or without laser irradiation under a N2 atmosphere.

Figure 4. (a) Flow cytometry analyses of U87MG cells incubated with HMCP-FITC and PFP@HMCP-FITC for 15 min without or with NIR laser irradiation. Inset of (a): Corresponding relative mean fluorescence intensity. (b) Confocal luminescence imaging of U87MG cells after incubated with FITC-conjugated HMCP and PFP@HMCP for 15 min without or with NIR laser irradiation. Scale bar: 50 μm. (c) Bio-TEM images of U87MG cells incubated with HMCP and PFP@HMCP for 15 min without or with NIR laser irradiation. The yellow arrows refer to internalized HMCP nanoparticles upon NIR laser irradiation. The red arrows refer to internalized PFP@HMCP nanoparticles upon NIR laser irradiation.

environment. As shown in Figure 3g,h, when O2−PFP@ HMCP is added instead of PFP@HMCP, the dissolved oxygen concentration was rapidly increased and could persist for several minutes, suggesting the stable encapsulation of oxygensaturated PFP in HMCP. After exposure to NIR irradiation, a sharp increase of dissolved oxygen concentration was observed due to the burst release of O2 from PFP gasification. Moreover, the dissolved oxygen concentration remained at a relatively

ature-responsive gasification of PFP, as shown by the gradually emerging PFP bubbles with prolonged NIR laser irradiation (Figure 3a−c). Furthermore, many more bubbles were observed for oxygen-saturated PFP@HMCP (O2−PFP@ HMCP) upon NIR irradiation (Figure 3d−f), which should be attributed to the release of both PFP and O2. The above temperature-responsive O2 release is expected to increase the dissolved oxygen concentration in the hypoxic 1583

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Figure 5. (a) Relative viabilities of hypoxic U87MG cells incubated with HMCP, PFP@HMCP, and O2−PFP@HMCP for 24, 48, and 72 h after exposure to 6 Gy of X-ray irradiation. (b) Flow cytometry analysis of O2 in hypoxic U87MG cells incubated with HMCP, PFP@HMCP, and O2−PFP@HMCP for 15 min without or with NIR laser irradiation. Inset of (b): Corresponding relative mean fluorescence intensity. (c) Confocal luminescence images of hypoxic U87MG cells after incubated with HMCP, PFP@HMCP, and O2−PFP@HMCP without or with NIR laser irradiation. Scale bar: 50 μm. ***P < 0.001.

high level within 15 min postirradiation, indicating that O2− PFP@HMCP serves as an excellent oxygen reservoir for alleviating hypoxia. It has been reported that bubbles can create permeable defects in lipid bilayers,29 which may contribute to enhancing the cellular uptake of nanoparticles. In order to verify this hypothesis, our designed HMCP was labeled with fluorescein isothiocyanate (FITC) to allow for quantitative evaluation and visual observation of intracellular transport via flow cytometry analysis and confocal luminescence imaging, respectively. The NIR laser power density was tuned to maintain the temperature around 42 °C in cells, so that the generated mild hyperthermia would not damage the cells. No matter whether exposed to NIR laser irradiation, both HMON and HMCP were biocompatible (Figure S9). As shown in Figure 4a, although HMCP-FITC combined with NIR laser irradiation enhanced the U87MG cell uptake to some extent owing to the mild temperature increase,30 the cellular uptake of PFP@HMCPFITC could be further increased upon NIR irradiation. This may be ascribed to the generated PFP bubbles driving more nanoparticles into the cytoplasm, as exhibited by the stronger green fluorescence intensity of PFP@HMCP-FITC around the blue nuclei than that of HMCP-FITC upon NIR laser irradiation (Figure 4b). Moreover, the bio-TEM images provided visual evidence of a greater number of PFP@ HMCP nanoparticles in U87MG cells subjected to NIR laser irradiation (Figure 4c, the red arrows). By contrast, only a few nanoparticles were endocytosed without any treatment, and the intracellular transport efficiency of HMCP was also suboptimal even with exposure to NIR laser irradiation (Figure 4c, the yellow arrows), which further confirmed the bubble-enhanced tumor cell uptake of nanoparticles. Therefore, it can be concluded that the mild hyperthermia-induced liquid−gas transformation of PFP enhances the tumor celluptake of

nanoparticles in addition to promoting the fast intracellular/ intercellular delivery and release of oxygen, which may substantially alleviate the tumor cell hypoxia. In contrast to normal U87MG cells, hypoxic U87MG cells were cultured in a low-oxygen atmosphere of 1% O2/5% CO2/ 94% N2. When exposed to X-ray radiation, hypoxic cells exhibited a much higher viability than normal cells, which confirmed the stronger resistance of hypoxic cells to RT (Figure S10). We then examined the potential of our designed O2−PFP@HMCP in overcoming the hypoxic resistance and improving the RT effects. As the temperature of an incubator at 37 °C is higher than the boiling point of PFP, NIR laser irradiation is not required for inducing PFP gasification. Figure S11 shows that O2−PFP@HMCP exhibited low cytotoxicity at 37 °C, which indicates that the generated PFP and oxygen bubbles did not result in permanent damage to the cells. However, O2−PFP@HMCP gave rise to a significant radiation enhancement effect, much better than HMCP and PFP@ HMCP, which may be attributed to the rapid PFP bubbleinduced O2 release for greatly increased sensitivity of hypoxic U87MG cells to X-ray radiation (Figure 5a and Figure S12). Moreover, the calcein AM/propidium iodide stained images also demonstrate a larger population of dead cells following treatment with O2−PFP@HMCP + RT (reflected by the stronger red fluorescence intensity in comparison to other treatments) as well as an increased hypoxic cell death rate with increasing incubation time and X-ray doses (Figures S13 and S14). The corresponding mechanism of hypoxic radiosensitization by O2−PFP@HMCP was explored by probing the intracellular O2 level using a [Ru(dpp)3]Cl2 indicator. It can be observed that the red fluorescence of [Ru(dpp)3]Cl2 was quenched in the cells treated by O2−PFP@HMCP and NIR laser irradiation for 15 min, which indicated the greatly diminished hypoxia 1584

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Figure 6. (a) Flow cytometry analysis of ROS in hypoxic U87MG cells incubated with fresh medium, HMCP, PFP@HMCP, and O2−PFP@ HMCP without or with X-ray irradiation. Inset of (a): Corresponding relative mean fluorescence intensity. (b) Single cell gel electrophoresis (comet) assay of hypoxic U87MG cells incubated with fresh medium, PFP@HMCP, and O2−PFP@HMCP before and after exposure to X-ray irradiation. (c) Quantitative analysis of DNA damage of hypoxic U87MG cells incubated with fresh medium, PFP@HMCP, and O2−PFP@ HMCP before and after exposure to X-ray irradiation. *P < 0.05, **P < 0.01.

Figure 7. (a) In vivo blood circulation half-life of HMCP. (b) Representative coronal PET images of a U87MG tumor-bearing mouse at 1, 6, 24, and 48 h postinjection of 100 μCi 64Cu-labeled HMCP. (c) PET quantification of biodistribution of 64Cu-HMCP at different time points postinjection. In vivo PA imaging of a U87MG tumor (d) before injection of HMCP and (e) at 24 h postinjection. In vivo B-mode US imaging of a U87MG tumor injected with O2−PFP@HMCP (f) before and (g) 4 h after NIR laser irradiation 24 h postinjection. Overlay of PA oxygen saturation mapping and US imaging of a U87MG tumor injected with O2−PFP@HMCP (h) before and (i) 4 h after NIR laser irradiation 24 h postinjection.

destruction, while gasified PFP bubbles play an indispensable role in driving the fast intracellular O2 release and intercellular O2 diffusion, which further improves oxygenation and enhances hypoxic radiosensitization based on the so-called “bystander effect”. Owing to the small size and PEG modification, the blood circulation half-life of HMCP in vivo (Figure 7a) is much longer than that of other reported silica nanoparticles.33,34 Instead of using conventional labor-intensive and inefficient procedures for labeling silica nanoparticles with 64Cu through the conjugation of NOTA,35 herein, a facile, yet stable radiolabeling method was developed based on the strong chelating ability of thiol groups on the silica surface toward 64Cu. According to the quantitative PET imaging, HMCP quickly accumulated at the U87MG tumor site within 1 h, and the tumor uptake reached a peak (more than 7%ID/g) at 24 h and then decreased slightly at 48 h (Figure 7b,c). Accordingly, due to the PA ability of CuS

arising from the rapid O2 release and diffusion through cells. Contrastingly, other nanoparticles without O2 did not improve the oxygenation, as seen by the unchanged red fluorescence of [Ru(dpp)3]Cl2 (Figure 5b,c). Furthermore, the intensified oxygenation was accompanied by the enhanced X-ray-induced reactive oxygen species (ROS) generation, as evidenced by the recurrent green fluorescence of DCFH-DA (a probe for detecting ROS) in the cells treated by O2−PFP@HMCP + RT in Figure 6a and Figure S15. DNA damage by ROS was assessed by single cell gel electrophoresis (comet) assay (Figure 6b). In general, the longer tail of fluorescent DNA stain signifies greater DNA damage.31,32 Although the DNA damage of hypoxic cells is not highly dependent on the X-ray dosage, the addition of O 2 −PFP@HMCP caused much more significant DNA damage than that of PFP@HMCP (Figure 6c). Therefore, it is O2 rather than PFP that brings about significant DNA damage for X-ray-triggered hypoxic cell 1585

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Figure 8. (a) Immunofluorescence staining analysis of HIF-1α expression in tumors from mice injected with PBS and O2−PFP@HMCP without and with NIR laser irradiation. Scale bar: 50 μm. (b) Western blot analysis of HIF-1α expression in tumors from mice injected with PBS, and O2−PFP@HMCP with NIR laser irradiation. (c) Photothermal imaging of mice bearing a U87MG tumor injected with PBS and O2−PFP@HMCP upon NIR laser irradiation for 10 min. (d) Temperature change curves of mice bearing U87MG tumors injected with PBS and O2−PFP@HMCP upon NIR laser irradiation for 10 min. (e) Tumor growth curves of mice bearing U87MG tumors subjected to various treatments. (f) Western blot analysis of p53 and caspase3 expression in tumors from mice injected with PBS and O2−PFP@HMCP after NIR laser irradiation and radiotherapy. (g) TUNEL stained images of tumors in the six groups. Scale bar: 200 μm. (h) H&E stained images of tumors in the six groups. Scale bar: 200 μm. ***P < 0.001.

O2−PFP@HMCP to tumor hypoxia alleviation through mild hyperthermia-triggered PFP bubble-driven oxygen release/ supply. The enhanced tumor oxygenation along with the PET/PA/US triple modality imaging based on O2−PFP@ HMCP paved the way for the following imaging-guided oxygen-elevated RT. Before starting the treatment, the PA/US imaging and immunofluorescence staining analysis (Figure S21) were performed to confirm that the tumor had developed enough hypoxia when its volume reached around 60 mm3. In order to facilitate the liquid−gas transformation of PFP while avoiding possible thermal degradation, the temperature of tumors treated with O2−PFP@HMCP + NIR was maintained around 42 °C (recorded using an IR thermal camera, Figure 8c,d), which had a marginal effect on tumor growth, similar to the control group and O2−PFP@HMCP group (Figures 8e and S22). Owing to the strong resistance of hypoxic tumors to Xray irradiation, RT only produced a limited inhibitory effect on tumor growth. The therapeutic effect of O2−PFP@HMCP + RT was also not satisfactory because of the limited O2 release without NIR laser irradiation. Notably, mild hyperthermia promoted O2 release and diffusion, thus alleviating tumor hypoxia to a great extent for enhanced hypoxic radiosensitization, which substantially improved the antitumor effect of X-ray radiation. As a result, tumors treated by O2−PFP@ HMCP + NIR + RT gradually diminished and were finally eliminated in 20 days. The Western blot result (Figure 8f) clearly demonstrated the remarkable up-regulation of p53 and caspase3 expression in the O2−PFP@HMCP + NIR + RT treated tumor, which directly led to much larger scale tumor

(Figure S16), the strongest PA signal of HMCP in the tumor was detected at 24 h postinjection (Figures 7d,e and S17), which further confirmed optimal tumor accumulation of HMCP at the 24 h time point. Following PFP encapsulation and O2 adsorption, O2−PFP@ HMCP can be employed as an US imaging contrast agent (Figure S18). It should be noted that the phase change temperature of PFP in vivo was increased to over 37 °C because of the additional blood pressure, so there was no US signal in the tumor at 24 h postinjection of O2−PFP@HMCP without NIR laser irradiation. Excitingly, when the tumor temperature was increased to 42 °C after mild NIR laser irradiation, the ultrasound signal quickly intensified, which may be attributed to both the generated PFP and released O2 bubbles, as evidenced by the much stronger US signal appearing in the tumor treated by O2−PFP@HMCP versus PFP@HMCP (Figures 7f,g, and S19). Subsequently, the co-registration of multiwavelength PA/ US imaging was used to measure vascular saturated O2 (sO2).36 As expected, the sO2 was substantially increased at 24 h postinjection of O2−PFP@HMCP upon mild NIR laser irradiation (Figures 7h,i and S20), indicating the diminished hypoxia and enhanced oxygenation of the tumor. The hypoxiainducible factor 1α (HIF-1α) expression has been considered as a reliable indicator of tumor hypoxia level, which was assessed by immunofluorescence staining and Western blot. As exhibited in Figure 8a, the red fluorescence of anti-HIF-1α antibody was largely quenched throughout the tumor cells treated by O2− PFP@HMCP upon mild NIR laser irradiation, which coincided well with the substantial down-regulation of HIF-1α expression (Figure 8b), further indicating the significant contribution of 1586

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EXPERIMENTAL SECTION

cell apoptosis/necrosis in sharp contrast to the other treatments (as exhibited by the TUNEL and H&E stained images in Figure 8g,h). As calculated by the Image-pro plus 6.0 software (Media Cybernetics, Inc., MD, USA), the apoptosis/ necrosis percentages in the TUNEL staining images (Figure 8g) were determined to be 16.2, 28.8, 36.4, 47.8, and 64.5% in the groups of control, O2−PFP@HMCP, O2−PFP@HMCP + NIR, RT, and O2−PFP@HMCP + RT, respectively. However, the value in the O2−PFP@HMCP + NIR + RT group was very difficult to obtain owing to the extremely large apoptosis/ necrosis area. The above excellent anticancer effect of O2−PFP@HMCP + NIR + RT results from the combination of the mild hyperthermia arising from NIR laser irradiation, triggering the generation of PFP bubbles, which are responsible for distributing released O2 throughout the tumor, thus leading to a “domino effect”, including improved tumor oxygenation, elevated ROS production, increased DNA damage, and enhanced tumor cell death. Encouragingly, the body weight of all the mice experienced negligible change within 20 days (Figure S23), suggesting the biosafety of these treatments. In addition, the major organs from mice injected with HMCP were collectedat day 30 postinjection for H&E staining, and no obvious damage or long-term adverse toxicity was found in these tissues (Figure S24). The well-designed gas generation system O2−PFP@HMCP in this study has shown excellent advantages in mild hyperthermia-induced bubble-enhanced oxygen-sensitized radiotherapy. Upon NIR laser irradiation, the generated PFP bubbles arising from mild hyperthermia (Figure 3a−f) not only enhance the tumor cell uptake of nanoparticles (Figure 4) but also assist in oxygen diffusion throughout the tumor (Figures 5b,c and 8a). The considerable oxygen supply helps to increase the sensitivity of hypoxic cells to X-ray radiation and elevate the intracellular ROS level (Figures 6a and S15), thus intensifying the X-ray-induced DNA damage (Figure 6b,c) for remarkable radiation enhancement (Figures 5a and 8e). By increasing the power density of NIR laser irradiation, O2−PFP@HMCP is able to increase the temperature to 60 °C for photothermal therapy. By taking into account that the gas generation system can also be used as a deep-penetrating drug delivery system for improved chemotherapy,37 the resulting combination of photothermal therapy, chemotherapy, and radiotherapy is expected to produce synergistic therapeutic effects for substantially enhanced treatment efficacy.

Chemicals and Reagents. Tetraethoxysilane (TEOS), bis[3(triethoxysilyl)propyl]tetrasulfide (BTES), cetyltrimethylammonium chloride solution (CTAC), triethanolamine (TEA), ethanol, concentrated HCl (37%), ammonia aqueous solution (NH3·H2O, 25 wt %), sodium sulfide nonahydrate (Na2S·9H2O), copper chloride dihydrate (CuCl2·2H2 O), sodium citrate (3-mercaptopropyl)trimethoxysilane (MPTES), (3-aminopropyl)triethoxysilane (APTES), and 2′,7′-dichlorofluorescein diacetate (DCFH-DA) were obtained from Sigma-Aldrich (MO, USA). Perfluoropentane was bought from VWR (PA, USA). PEG2000-SH was purchased from Laysan Bio Inc. (AL, USA). Deionized water (Millipore) with a resistivity of 18 MΩ·cm was employed in all of the experiments. Minimum essential medium (MEM), fetal bovine serum (FBS), dimethyl sulfoxide (DMSO), 0.05% trypsin-EDTA, PBS, and penicillin−streptomycin solution were acquired from Gibco Laboratories (NY, USA). 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), FITC, calcein AM, and propidium iodide (PI) were bought from Thermo Fisher Scientific Inc. (WI, USA). Tris(4,7diphenyl-1,10-phenanthroline)ruthenium(II) dichloride complex ([Ru(dpp)3]Cl2) was purchased from Santa Cruz Biotechnology (TX, USA). The U87MG human glioblastoma cell line was acquired from American Type Culture Collection (ATCC). Characterization. Transmission electron microscopy (TEM) images were acquired on a Tecnai T12 microscope (FEI Company, OR, USA). Scanning electron microscope images were obtained on a Hitachi SU-70 Schottky field emission gun scanning electron microscope (Hitachi, IL, USA). EDX was performed by using a JEM-2100F transmission electron microscope with an EDX detector system (JEOL USA, Inc., MA, USA). Ultraviolet−visible−nearinfrared (UV−vis−NIR) absorption spectra were performed on a Genesys 10S UV−vis spectrophotometer (Thermo Fisher Scientific Inc., WI, USA). The ζ-potential and dynamic light scattering measurement were tested on a Horiba SZ-100 nanoparticle analyzer (Horiba Instruments Inc., NY, USA). FT-IR spectra were measured on a Thermo Nicolet Nexus870 spectrometer (Nicolet Instruments Inc., WI, USA). Student’s two-tailed t test was applied for statistical analysis. Synthesis of MSN@MON. Typically, 20 g of CTAC aqueous solution (10 wt %) and 1 g TEA aqueous solution (10 wt %) were first mixed and stirred at 95 °C, and then TEOS (1 mL) was added dropwise. After reacting for 1 h, a mixture of TEOS (0.5 mL) and BTES (1 mL) was added for another 4 h of reaction. The resultant MSN@MON products were collected after centrifugation and then washed with ethanol several times. The as-prepared MSN@MON was dispersed in ethanol (100 mL) containing concentrated HCl (37%) (10 mL) and heated at 78 °C for 12 h to remove the template CTAC. The extraction procedure was repeated for several times until CTAC was thoroughly removed, and the CTAC-free MSN@MON products were redispersed in 20 mL of water. Synthesis of HMON. An ammonia-assisted selective etching strategy was used to selectively etch away the inner MSN core, leaving the outer MON shell with a hollow structure. A certain amount of ammonia solution was added into the obtained solution of MSN@ MON, which was allowed to react for 3 h at 95 °C. After the solution was cooled to room temperature, the final HMON products were obtained after centrifugation and washing with water for several times. Synthesis of HMCP. First, copper sulfide (CuS) nanoparticles were synthesized via a reported method.38 Two milliliters of 0.05 M Na2S·9H2O aqueous solution was added into 100 mL of mixed 1 mM CuCl2·2H2O and 0.68 mM sodium citrate aqueous solution. After being stirred at room temperature for 5 min, the mixture was heated to 80 °C for another 10 min of reaction. Then, the dark-green CuS solution was immediately transferred to an ice bath. Afterward, the asprepared HMON (15 mg) was dispersed in ethanol (40 mL), followed by addition of a mixture of MPTES (0.15 mL) and ammonia (0.2 mL, 25 wt %) aqueous solution. Then the obtained solution was stirred overnight, and the resultant HMON-SH was collected after centrifugation and washing with ethanol for several times. To obtain HMON@CuS, a mixture of CuS solution (60 mL) and HMON-SH

CONCLUSIONS In summary, we have introduced a mild ammonia-assisted selective etching strategy for the fabrication of sub-50 nm biocompatible and biodegradable HMON for encapsulation and efficient delivery of oxygen-saturated hydrophobic PFP. Through surface modification of ultrasmall CuS nanoparticles and PEGylation, the generated mild hyperthermia upon NIR laser irradiation can induce the liquid−gas phase transformation of PFP, which not only greatly intensifies the US imaging signal owing to the burst release of PFP and O2 bubbles but also forcefully drives the oxygen diffusion throughout the tumor for considerable hypoxia alleviation and remarkable X-ray radiation enhancement. The silica-based nanotheranostics developed in this study for cancer therapy are generalizable and may open up new dimensions for other types of gas-enhanced synergistic therapy under significant multimodal imaging guidance. 1587

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ACS Nano

In Vitro Biocompatibility Assessment of HMON, HMCP, PFP@HMCP, and O2−PFP@HMCP. U87MG human glioblastoma cells were cultured in 96-well plates with a density of 5 × 103 cells per well under a humidified atmosphere with 1% O2/5% CO2/94% N2 for 24 h at 37 °C. For HMON, the medium was replaced with various concentrations of HMON in 100 μL of MEM. For HMCP, the medium was replaced with HMCP in 100 μL of MEM at different concentrations, and in laser groups, the cells were subjected to 808 nm NIR laser irradiation for 15 min, during which time the temperature was maintained around 42 °C by adjusting the laser power density. Then the cells were cultured for another 24 h. For long-term cell toxicity, cells were treated with 100 μg/mL of HMCP, PFP@HMCP, and O2−PFP@HMCP separately for 24, 48, and 72 h. The cells incubated with MEM were set as a control group. The cytotoxicity of the materials was measured by a standard MTT assay. Typically, medium from each well was replaced with 10% MTT in 100 μL of PBS solution, and cells were incubated for another 4 h at 37 °C. After the medium was removed, DMSO (150 μL) was used to dissolve the formazan crystals. The absorbance at 570 nm was determined by a microplate reader (BioTek Instruments, VT, USA). In Vitro NIR Laser and Bubble-Enhanced Cell Uptake. First, HMCP and PFP@HMCP were modified with FITC (denoted as HMCP-FITC and PFP@HMCP-FITC, respectively). FITC (15 mg) and APTES (100 μL) were stirred in ethanol (5 mL) for 24 h protected from light, and then 0.5 mL of the above aminofunctionalized FITC precursor was mixed with 20 mg of HMCP (or PFP@HMCP) in ethanol (20 mL) for 24 h with stirring. The obtained HMCP-FITC and PFP@HMCP-FITC were centrifuged and washed with PBS for several times. For confocal laser scanning microscopy (CLSM), U87MG cells were seeded in a Lab-Tek chamber slide system (Thermo Fisher Scientific, WI, USA) with 5 × 103 cells per well in MEM (200 μL) containing 10% FBS and cultured in a 1% O2/ 5% CO2/94% N2 humidified atmosphere for 24 h at 37 °C. Then cells were incubated with HMCP-FITC or PFP@HMCP-FITC, without or with NIR laser irradiation for 15 min. For NIR laser irradiation groups, the temperature was kept around 42 °C by adjusting the laser power density. For groups without NIR laser irradiation, cells were incubated at room temperature. After the cells were washed with PBS several times, they were stained with UltraCruz mounting medium (Santa Cruz Biotechnology, TX, USA). Cellular uptake of HMCP-FITC (or PFP@HMCP-FITC) was measured using CLSM (LSM 780, Carl Zeiss, Germany) by detecting the fluorescence of FITC (λex = 495 nm, λem = 519 nm). For flow cytometry, U87MG cells at a density of 5 × 104 cells/mL were cultured in 6-well plates in 2.5 mL of medium under humidified 1% O2/5% CO2/94% N2 atmosphere at 37 °C. After 24 h of incubation, cells were treated with the above-mentioned method, harvested, and washed with PBS. Then a BD Accuri C6 (BD, NJ, USA) was employed for flow cytometry. Cells subjected to various treatments were also observed by using bio-TEM. In Vitro Cellular Hypoxia Detection by Optical Probe. For CLSM, U87MG cells were seeded into a Lab-Tek chamber slide system with a density of 5 × 103 cells in each well cultured in 200 μL of MEM containing 10% FBS in a humidified 1% O2/5% CO2/94% N2 atmosphere at 37 °C. After 24 h of incubation, cells were incubated with 10 μg/mL [Ru(dpp)3]Cl2 for 12 h and washed with PBS several times. Then the cells were treated with 100 μg/mL of HMCP, PFP@ HMCP, and O2−PFP@HMCP without or with NIR laser irradiation for 15 min, and the temperature was kept at 42 °C in the NIR laser irradiation groups by adjusting the laser power density. For groups without NIR laser irradiation, cells were placed at room temperature. After the cells were washed using PBS for several times and stained with UltraCruz mounting medium, intracellular oxygen level was measured using CLSM by detecting the fluorescence of [Ru(dpp)3]2+Cl2 (λex = 450 nm, λem = 610 nm). For flow cytometry, U87MG cells (5 × 104 cells/mL) were cultured in 6-well plates in 2.5 mL of medium under a humidified atmosphere of 1% O2/5% CO2/ 94% N2 for 24 hat 37 °C, and then cells were treated as described above. After the cells were harvested and washed with PBS, a BD Accuri C6 was used for flow cytometry.

(30 mg) was stirred for 40 min. The product was obtained after centrifugation and redispersed in 100 mL of ethanol. For PEGylation, 25 mg of PEG2000-SH was added into the above solution, which was refluxed for 24 h at 78 °C. Then the final HMCP products were obtained after centrifugation and washing with water several times. Synthesis of O2−PFP@HMCP. Ten milligrams of HMCP was placed in a 15 mL centrifuge tube with a rubber stopper. After the air in the tube was evacuated by a vacuum pump, PFP (50 μL) was quickly injected into the tube. PFP loaded HMCPs (PFP@HMCP) were obtained after brief sonication in ice−water for 1 min, followed by being redispersed in PBS solution (5 mL) for further use. For oxygen saturation, the above solution (1 mL) was put in an aseptic oxygen chamber for 10 min at an oxygen flow rate of 5 L/min according to a reported method.39 The obtained product was denoted as O2−PFP@HMCP. Measurement of Photothermal Performance of HMCP. Different concentrations of HMCP in PBS solution (200 μL) were exposed to an 808 nm NIR laser at 0.5 W/cm2 for 3 min, or a certain concentration of 100 μg/mL HMCP was irradiated by different power densities of the NIR laser. Later, 100 μg/mL of HMCP PBS solution (200 μL) was irradiated by a 0.5 W/cm2 NIR laser for five cycles to examine its photothermal stability. A laser energy meter (Coherent Inc., CA, USA) was employed to determine the NIR laser power density. The real-time temperatures of the solutions were recorded using an infrared thermal imaging system. Calculation of Photothermal Conversion Efficiency of HMCP at 808 nm. The photothermal conversion efficiency of HMCP was calculated by the following equation:

η=

hS(Tmax − Tamb) − Q 0 I(1 − 10−A)

where h is the heat transfer coefficient, S is the surface area, Tmax is equilibrium temperature, Tamb is surrounding ambient temperature, Q0 refers to heat absorption of the quartz cell, I is the laser power, and A is the absorbance of HMCP at 808 nm. If the heat input is equal to the heat output

hS =

∑i miCp , i τs

≈

mH2OC H2O τs

where mH2O refers to the weight of water, CH2O stands for the specific heat capacity of water, and τs is time constant of the sample. At the cooling stage

t = − τs ln θ = − τs ln

T − Tamb Tmax − Tamb

Therefore, τs can be calculated by using the linear regression curve between cooling stage and negative natural logarithm of driving force temperature of HMCP. Measurement of Bubble Release. One milliliter of PBS solution of PFP@HMCP or O2−PFP@HMCP (5 mg/mL) was dropped onto a silanated slide (K-D Medical, MD, USA) and covered with a coverslip (Thermo Fisher Scientific Inc., WI, USA). We made sure that there were no bubbles in the solution after the coverslip was placed. Then the samples were exposed to NIR laser irradiation (0.5 W/cm2) for 10 and 20 s. The regions of interest at each time point were recorded by an IX81 microscopy (Olympus, Tokyo, Japan). Measurement of Oxygen Release. The oxygen concentrations in PBS solution were measured by using a MW600 standard portable dissolved oxygen meter (Milwaukee Instruments, Inc., NC, USA). Deoxygenated PBS (15 mL) was put in a 50 mL centrifuge tube filled with nitrogen and sealed by a rubber stopper. The probe of the oxygen meter was inserted into the tube through the rubber stopper to measure the real-time change in oxygen concentrations. Afterward, 5 mL of PBS solution of O2−PFP@HMCP or PFP@HMCP (5 mg/ mL) was injected into the tube. For NIR laser irradiation groups, the solutions were irradiated by a 0.5 W/cm2 808 nm NIR laser. The oxygen concentrations were recorded every 30 s for a total of 15 min. 1588

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ACS Nano In Vitro Cellular ROS Detection. For CLSM, U87MG cells (5 × 103 per well) in 200 μL of medium were seeded into a Lab-Tek chamber slide system in a humidified 1% O2/5% CO2/94% N2 atmosphere at 37 °C. After 24 h of incubation, cells were incubated with 10 μM DCFH-DA in fresh medium for 20 min, protected from light. After that, the cells were washed using fresh medium for several times and incubated with HMCP, PFP@HMCP, and O2−PFP@ HMCP (100 μg/mL) for 2 h at 37 °C. Then the cells were exposed to 6 Gy of X-ray radiation under a Multirad 160 kV X-ray (Faxitron Bioptics, LLC, AZ, USA), followed by CLSM observation. For flow cytometry, U87MG cells (5 × 104 cells/mL) were seeded in 6-well plates in 2.5 mL of medium in a humidified atmosphere of 1% O2/5% CO2/94% N2 for 24 h at 37 °C, and then cells were treated as mentioned above. After the cells were harvested and washed using PBS, a BD Accuri C6 was used for flow cytometry. The cells incubated with MEM were used as a control. In Vitro Tumor Cell Radiotherapy. U87MG cells were cultured in 96-well plates with a density of 5 × 103 cells per well in MEM (100 μL) containing 10% FBS. For cells incubated in normal and hypoxia conditions, the plates were put in 21% O2/5% CO2/74% N2 and 1% O2/5% CO2/94% N2 humidified atmosphere, respectively, for 24 h at 37 °C. After the cells were subjected to 2, 4, and 6 Gy of X-ray radiation and incubated for another 24, 48, and 72 h, standard MTT assay was used to test the cell viabilities. To measure the contribution of materials to radiosensitization, the cells were incubated with 100 μg/mL of HMCP, PFP@HMCP, and O2−PFP@HMCP for 2 h after incubation in a humidified atmosphere of 1% O2/5% CO2/94% N2 at 37 °C for 24 h. Afterward, 4 and 6 Gy of X-ray irradiation was imposed on the cells. Cells treated with fresh medium were set as control. At 24, 48, and 72 h postirradiation, standard MTT assay was employed to measure the cell viabilities. For differentiation between live and dead cells, U87MG cells subjected to various treatments were costained with calcein AM and PI and then observed under an IX81 fluorescence microscope. In Vivo Blood Terminal Half-Life Measurement. All animal experiments were performed in accordance with a National Institutes of Health Animal Care and Use Committee (NIHACUC) approved protocol. The FVB mice (n = 3) were intravenously injected with HMCP in PBS at the dosage of 20 mg/kg. At various time points (2 min, 5 min, 10 min, 30 min, 1 h, 2 h, 4 h, 8 h, and 24 h) postinjection, 15 μL of blood was collected and dissolved into 985 μL of physiological saline containing 10 mM EDTA anticoagulant. The concentration of silica of each sample was determined by ICP analysis. The curve of blood terminal half-life was fitted based on a singlecompartment pharmacokinetic model. On day 30 postinjection, one mouse was euthanized, and the long-term toxicity of HMCP was evaluated by hematoxylin and eosin (H&E) staining analysis of tissues from major organs. The mouse injected with PBS was set as a control. In Vitro and In Vivo Ultrasound and Photoacoustic Imaging. In vitro and in vivo photoacoustic and B-mode ultrasound imaging was conducted on a VisualSonics Vevo LAZR system (VisualSonics Inc. NY, USA). A total of 100 μL of 4 mg/mL HMCP, PFP@HMCP, and O2−PFP@HMCP in PBS was injected intravenously into U87MG tumor-bearing mice. PA imaging was conducted before injection and at various time points after injection. B-mode US imaging of tumor was completed 24 h postinjection and before laser irradiation. Then the tumors were subjected to NIR laser for 15 min, during which time the temperature was maintained around 42 °C. Afterward, US imaging was recorded at various time points. In Vivo PET Imaging. Radioactive HMON@ 64 CuS-PEG (HM64CP) was synthesized by conjugating 64Cu on the surface of HMCP through thiol groups. U87MG tumor-bearing mice were injected intravenously with about 100 μCi of HM64CP in PBS. PET scanning at different time points postinjection was conducted on a Siemens Inveon microPET scanner (Siemens Healthcare GmbH, Germany). Quantitative PET data were acquired by drawing regions of interest on the organs with decay-corrected whole-body coronal images. The percentage injected dose per gram of tissue (%ID/g) was then calculated based on the measured values.

In Vivo Radiotherapy. Thirty female athymic nude mice 4−6 weeks old were used. Xenograft tumors were prepared by subcutaneous injection of 2 × 106 U87MG cells on the right rear leg. When the tumor volume reached around 60 mm3, these 30 mice were divided into six groups: control, O2−PFP@HMCP, O2−PFP@ HMCP + NIR, RT, O2−PFP@HMCP + RT, and O2−PFP@HMCP + NIR + RT. The mice were injected intravenously with 100 μL of PBS in control and RT groups, and 100 μL of 4 mg/mL O2−PFP@HMCP in PBS solution in the other four treatment groups. After 24 h, the tumors in the O2−PFP@HMCP + NIR group were subjected to NIR laser for 10 min, during which time the temperature was maintained around 42 °C by adjusting the laser power density. For RT and O2− PFP@HMCP + RT groups, the tumors were subjected to 8 Gy of Xray radiation. For the O2−PFP@HMCP + NIR + RT group, the tumors were first irradiated with the NIR laser for 10 min and then subjected to 8 Gy of X-ray radiation. One mouse from each group was euthanized on the third day after the treatment for tumor pathological staining. Tumor dimensions and body weight were measured every 2 days post-treatment. Tumor volumes were determined according to the following formula: volume (V) = length × width2/2. The relative tumor volume (V/V0) and body weight (W/W0) were normalized to the initial tumor volume (V0) and body weight (W0).

ASSOCIATED CONTENT S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.7b08103. Additional related experimental data (PDF)

AUTHOR INFORMATION Corresponding Authors

*E-mail: [email protected]. *E-mail: [email protected]. *E-mail: [email protected]. ORCID

Zhaogang Teng: 0000-0001-8792-764X Kai Yang: 0000-0002-6670-1024 Guangming Lu: 0000-0003-4913-2314 Xiaoyuan Chen: 0000-0002-9622-0870 Notes

The authors declare no competing financial interest.

ACKNOWLEDGMENTS We greatly appreciate financial support from the National Key Basic Research Program of the PRC (2014CB744504 and 2014CB744501), the Natural Science Foundation of Jiangsu Province (BK20160017), and the National Natural Science Foundation of China (51602203, 81530054). We also acknowledge the support from the Intramural Research Program (IRP), National Institute of Biomedical Imaging and Bioengineering (NIBIB), National Institutes of Health (NIH). REFERENCES (1) Hogle, W. P. The State of the Art in Radiation Therapy. Semin. Oncol. Nurs. 2006, 22, 212−220. (2) Song, G.; Cheng, L.; Chao, Y.; Yang, K.; Liu, Z. Emerging Nanotechnology and Advanced Materials for Cancer Radiation Therapy. Adv. Mater. 2017, 29, 1700996. (3) Horsman, M. R.; Overgaard, J. Hyperthermia: a Potent Enhancer of Radiotherapy. Clin. Oncol. 2007, 19, 418−426. (4) Sun, X.; Li, X. F.; Russell, J.; Xing, L.; Urano, M.; Li, G. C.; Humm, J. L.; Ling, C. C. Changes in Tumor Hypoxia Induced by Mild Temperature Hyperthermia as Assessed by Dual-Tracer Immunohistochemistry. Radiother. Oncol. 2008, 88, 269−276. 1589

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DOI: 10.1021/acsnano.7b08103 ACS Nano 2018, 12, 1580−1591

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DOI: 10.1021/acsnano.7b08103 ACS Nano 2018, 12, 1580−1591