Ultrasound Triggered Tumor Oxygenation with Oxygen-Shuttle

Sep 13, 2016 - Herein, taking advantages of the great biocompatibility and high oxygen dissolving ability of perfluorocarbon (PFC), we develop an inno...
0 downloads 5 Views 8MB Size
Download PDF
Letter pubs.acs.org/NanoLett

Ultrasound Triggered Tumor Oxygenation with Oxygen-Shuttle Nanoperfluorocarbon to Overcome Hypoxia-Associated Resistance in Cancer Therapies Xuejiao Song,† Liangzhu Feng,† Chao Liang,† Kai Yang,‡ and Zhuang Liu*,† †

Institute of Functional Nano & Soft Materials (FUNSOM), Jiangsu Key Laboratory for Carbon-Based Functional Materials & Devices, Soochow University, Suzhou 215123, China ‡ School of Radiation Medicine and Protection & School for Radiological and Interdisciplinary Sciences (RAD-X), Collaborative Innovation Center of Radiation Medicine of Jiangsu Higher Education Institutions Medical College of Soochow University, Suzhou, Jiangsu 21513, China S Supporting Information *

ABSTRACT: Tumor hypoxia is known to be one of critical reasons that limit the efficacy of cancer therapies, particularly photodynamic therapy (PDT) and radiotherapy (RT) in which oxygen is needed in the process of cancer cell destruction. Herein, taking advantages of the great biocompatibility and high oxygen dissolving ability of perfluorocarbon (PFC), we develop an innovative strategy to modulate the tumor hypoxic microenvironment using nano-PFC as an oxygen shuttle for ultrasound triggered tumor-specific delivery of oxygen. In our experiment, nanodroplets of PFC stabilized by albumin are intravenously injected into tumor-bearing mice under hyperoxic breathing. With a low-power clinically adapted ultrasound transducer applied on their tumor, PFC nanodroplets that adsorb oxygen in the lung would rapidly release oxygen in the tumor under ultrasound stimulation, and then circulate back into the lung for reoxygenation. Such repeated cycles would result in dramatically enhanced tumor oxygenation and thus remarkably improved therapeutic outcomes in both PDT and RT treatment of tumors. Importantly, our strategy may be applied for different types of tumor models. Hence, this work presents a simple strategy to promote tumor oxygenation with great efficiency using agents and instruments readily available in the clinic, so as to overcome the hypoxia-associated resistance in cancer treatment. KEYWORDS: photodynamic therapy, radiotherapy, perfluorocarbon nanoemulsion, tumor hypoxia, ultrasound

O

onidase) would lead to relieved tumor hypoxia via improving intratumor blood flow.12,13 However, the tumor reoxygenation levels achieved by those methods are still limited by either the amount of available H2O2 inside the tumor microenvironment, or the oxygen delivery capability of red blood cells reaching tumor blood vessels. Perfluorocarbon (PFC), a category of chemically inert synthetic molecules composed of carbon and fluorine atoms, has many diverse applications in biomedicine.14,15 In addition to being utilized as the contrast enhancement agent of ultrasonography in the clinic,16−18 different formulations of liquid PFC have been extensively explored as an artificial blood substitute for decades owing to its high affinity to oxygen and great biocompatibility. Such perfluorocarbon molecules could be excreted by exhalation from lungs or skin pores with elimination half-time to be 3−4 days for perfluorooctyl bromide

riginating from the irregular cell proliferation, abnormal vasculature, and dysfunctional lymphatic system, tumor hypoxia has been recognized to be one of the hostile characteristics of solid tumors.1,2 According to previous studies, tumor hypoxia has been shown to be a revulsive factor of tumor metastasis and resistance to different types of therapies.3−6 In particular, for radiotherapy (RT) and photodynamic therapy (PDT) in which oxygen is an essential element for cancer cell killing, hypoxia-associate resistance often occurs for treatment of large solid tumors. Recently, different strategies have been proposed to oxygenate the tumor microenvironment in order to achieve better therapeutic outcomes.7−9 Several different groups including ours have uncovered that the tumor hypoxia could be relieved by in situ producing O2 from endogenous hydrogen peroxide (H2O2) existing inside the tumor with either catalyst or manganese dioxide (MnO2) nanoparticles to trigger the decomposition of H2O2, realizing significantly promoted treatment outcomes of RT and PDT.10,11 On the hand other, it has also been demonstrated that normalizing tumor vasculatures with either drug molecules or enzymes (e.g., hyalur© XXXX American Chemical Society

Received: June 9, 2016 Revised: September 1, 2016

A

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX

Letter

Nano Letters

Figure 1. Preparation of PFC nanoemulsion and US-triggered oxygen release from nano-PFC. (a) Scheme illustrating the fabrication of PFC nanoemulsion and the oxygen release triggered under US waves. (b) Freeze-etching replication TEM image of as-made HSA-stabilized PFC nanodroplets. (c) Size distributions of PFC nanoemuslsion before and after US treatment for 30 min. (d) Time-dependent changes of dissolved oxygen concentrations in deoxygenated pure water without or with addition of oxygen-loaded PFC nanoemlusion (PFC@O2). A US treatment was applied on these solutions within the indicated period.

and 8 days for perfluorodichlorooctane.19−21 For instance, Oxypherol (Fluosol-43) has obtained FDA approval in improving myocardial oxygenation and preventing abnormalities in ventricular function.22−24 On the other hand, Oxygent has been approved in Russia as a temporary intravascular oxygen carrier for hemorrhagic shock and perfusion of human organs.24,25 In a recent work, it was proposed that photosensitizer-coupled PFC nanoparticles could act as an adjuvant treatment to increase the therapeutic outcome of PDT via dragging oxygen molecules close to photosensitizers.26 However, the efficiency of this strategy could still be limited due to the low tumor endogenous oxygen concentration. Unlike hemoglobin that show a sigmoidal oxygen-dissociation curve to allow efficient binding of oxygen under high oxygen pressure (in the lung) and rapid oxygen release under hypoxic environment (tissues), PFCthough having a high oxygen solubilityreleases oxygen simply by diffusion through the oxygen concentration gradient with a low efficiency.27−29 We recently found that near-infrared (NIR) light mediated photothermal effect could be utilized to trigger the timely release of oxygen from PFC loaded inside hollow nanostructures, subsequently contributing to remarkably enhanced radiotherapeutic efficacy.30 However, NIR light has a short tissue penetration depth, whereas local administration of nanoagent was applied in that work only as a proof-of-concept experiment with limited value for clinical translation. Ultrasound (US) has been widely used in diagnostic imaging due to its safety, noninvasiveness, real-time visualization ability, excellent tissue penetration depth, and relatively low instrumentation cost.31,32 High intensity of ultrasound could also be utilized to trigger drug release or induce ablation

therapy of tumors.33,34 Herein, we develop a US-triggered tumor oxygenation strategy by using nano-PFC as the oxygen carrier to modify the tumor hypoxia for enhanced cancer PDT and RT. In our work, it is found that PFC nanodroplets stabilized by human serum albumin (HSA) could dissolve a large amount of oxygen, whose release could be readily triggered by an external low-frequency/low-power US treatment. After intravenous (i.v.) injection of nano-PFC, mice are going through a 30 min hyperoxic breathing with ultrasound treatment simultaneously applied at the tumor site. Via the blood circulation system, PFC nanodroplets with high oxygen loading after passing the lung would circulate into the tumor, in which oxygen is efficiently released from nano-PFC under US stimulation, resulting in dramatically enhanced tumor oxygenation as revealed by both in vivo photoacoustic (PA) imaging and ex vivo immunofluorescence staining. Finally, in vivo treatment experiments by using different tumor models demonstrate that our developed strategy could significantly improve the therapeutic efficacies of PDT and RT for cancer treatment. Our work presents an effective strategy to relieve the hypoxia-associated resistance in oxygen-involving cancer therapies using ultrasound triggered local oxygen release inside the tumor from nano-PFC, which acts as an artificial blood oxygen shuttle. This technique shows great promise for future clinical transition since the agents and instruments employed by this approach are readily available in the clinic. Results and Discussion. Owning to its immiscibility with both aqueous and lipophilic solutions, perfluorocarbons need to be emulsified for intravenous administration. By adopting the well-developed methods for preparation of liquid PFC nanoemusion, human serum albumin (HSA) was utilized as B

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX

Letter

Nano Letters

Figure 2. US-induced tumor oxygenation using PFC nanoemulsion. (a) Scheme showing the mechanism of US-triggered local oxygenation in the tumor using nano-PFC as the oxygen shuttle. (b) PA imaging of 4T1 solid tumors to determine tumor oxygenation status by measuring the ratios of oxygenated hemoglobin (λ = 850 nm) and deoxygenated hemoglobin (λ = 750 nm) before and 30 min after indicated treatments. (c) Quantification of the oxyhemoglobin saturation in the tumor. (d) Representative immunofluorescence images of tumor slices stained by the Hypoxyprobe. The blood vessels and hypoxia areas were stained with anti-CD31 antibody (red), and antipimonidazole antibody (green), respectively. (e) Quantification of tumor hypoxia and blood vessel densities for different groups shown in (d).

burst-like oxygen release was observed as indicated by its rapid increase of dissolved oxygen concentration (Figure 1d). Those results demonstrate that the oxygen release from PFC is sensitive to the applied external ultrasound waves. Inspired by the high oxygen loading ability and USresponsive oxygen release behavior of nano-PFC, we then designed an innovative strategy to realize highly efficient oxygen delivery into tumors with nano-PFC under the assistance of locally applied US (Figure 2a). Upon intravenous injection, PFC nanodroplets with long blood circulation time (Figure S3) would load oxygen within lung capillaries. After entering the solid tumor via blood circulation, there would be a burst release of oxygen from those oxygen-loading PFC nanodroplets triggered by US waves applied on the tumor. Those PFC nanodroplets could then be reoxygenated after they are circulated back into the lung. Those cycles can be repeated as long as the mouse is placed under hyperoxic inhalation with a US transducer placed onto its tumor. In order to figure out the feasibility of the above strategy in improving tumor oxygenation, in vivo PA imaging and ex vivo immunofluorescence staining were carried out for tumors. PA imaging relying on the photoacoustic effect is an emerging noninvasive biomedical imaging modality and has been shown to be effective for structural, functional, and molecular imaging.37,38 Because the near-infrared (NIR) optical absorbance peak of hemoglobin would change from its deoxygenated status (750 nm) to the oxygenated state (850 nm), the absorbance of hemoglobin at 750 and 850 nm are commonly

the stabilizer to form nanoemulsion of perfluoro-15-crown-5ether under ultrasonication (Figure 1a). The fabricated PFC nanodroplets showed uniform size distribution under transmission electron microscopy (TEM) (Figure 1b). By utilizing dynamic light scattering (DLS), it was found that the mean size of the as-prepared nano-PFC was about 160 nm, which is consistent with the TEM observation. Moreover, DLS measurement and TEM imaging indicated that the as-prepared PFC nanodroplets had a great stability as their size distribution showed negligible fluctuation even after 30 min of US treatment (Figure 1c and Figure S1). In addition, cell viability tested revealed that such HSA-stabilized PFC nanodroplets exhibited little cytotoxicity to different types of cells (Figure S2). Then, the oxygen loading and US-responsive oxygen release of nano-PFC was studied using an oxygen meter to measure dissolved oxygen concentrations in aqueous solutions. The saturated oxygen loading within our nano-PFC at 25 °C under 1 atm of pure oxygen was measured to be ∼1.45 ± 0.08 mg/ mL, consistent with the results reported in literature.35,36 Upon adding oxygen saturated PFC nanoemusion into deoxygenated water (prepared by boiling under nitrogen atmosphere), the dissolved oxygen concentration in water rapidly increased and saturated within the first 100 s, afterward it showed a slow decrease with the extending of incubation time. This result demonstrates the excellent oxygen loading ability but a gradual release profile of PFC. When a low-power/low frequency US treatment (3.5 W, 1.0 MHz) was applied onto this solution, a C

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX

Letter

Nano Letters

Figure 3. In vivo photodynamic therapy enhanced by US-triggered tumor oxygenation with nano-PFC. (a) Scheme showing our experimental process. (b) Tumor growth curves of different groups of mice (five mice per group) after various treatments including: untreated control (Ctr), PDT, PDT + O2, PDT + O2 + US, PDT + PFC + O2, PDT + PFC + O2 + US. The detailed experimental parameters may be found in the Materials and Methods section. (c) Average tumor weights of different groups after various treatments as indicated in (b). (d) H&E-stained and TUNELstained tumor slices collected from different groups of mice 1 day after various treatments. P values: *P < 0.05, **P < 0.01, ***P < 0.001, ANOVA.

under hyperoxic breathing but no US treatment (PFC + O2), large hypoxia areas remain exist in their tumors. By semiquantitative statistical analysis of the hypoxia signals in those tumors, the percentage of hypoxia positive area dropped from ∼56% for untreated group to ∼13% for the PFC + O2 + US group, while the percentage of hypoxia positive area in tumors for other control groups showed no significant or a slight decrease. Note that the blood vessel densities in different groups appeared to largely unchanged, suggesting such treatment would not notably affect the tumor blood vasculature (Figure 2e). In addition to Pimonidazole-based hypoxia staining, antihypoxia inducible factor 1 alpha (HIF-1α), an oxygen regulated subunit of HIF-1, was also examined to evaluate the tumor hypoxia status. The HIF-1 α expression level in tumors treated by PFC + O2 + US remarkably decreased in comparison with all other control groups (Figure S4). Those ex vivo immunofluorescence staining results were consistent with PA imaging data, and further confirmed that our local US-triggered tumor oxygenation strategy with nanoPFC as the US-responsive oxygen carrier could be a rather effective method to overcome tumor hypoxia. Next, we wondered whether such improved tumor oxygenation with PFC plus local US treatment would bring any advantage for cancer treatment. Photodynamic therapy is clinically adapted light-trigger therapy in which photosensitizing molecules in the presence of light would excite surrounding oxygen molecules from their ground triplet state into highenergy singlet state to destruct diseased cells.40,41 For PDT

adopted for PA imaging to evaluate the blood oxygenation status. In our experiment, 4T1 tumor-bearing mice breathing pure oxygen were i.v. injected with 200 μL nano-PFC (containing 30 μL of PFC). Right after PFC injection, an ultrasound transducer (1 MHz, 3.5 W) was placed on the tumor for 30 min. PA imaging was conducted for the same mouse before treatment and right after 30 min of such US treatment (Figure 2b). It was found that oxygenation levels over the entire tumor area (sO2 Tot) showed a fast increase from ∼17% for untreated tumors to ∼49% for those on PFCinjected mice treated with 30 min hyperoxic breathing plus tumor-localized US treatment (PFC + O2 + US). However, much less significant tumor oxygenation enhancement was observed for those 4T1 tumors on control mice received only simultaneous hyperoxic breathing and US treatment without PFC injection (Ctr + O2 + US), or PFC injection plus hyperoxic breathing without US treatment (PFC + O2) (Figure 2c). Then, the tumor oxygenation enhancement effect by our method was further confirmed by examining the tumor slices of different treated groups using Hypoxyprobe (pimonidazole hydrochloride) for immunofluorescence staining following the standard protocol.39 Consistent to PA imaging results, it was found that PFC + O2 + US treatment could dramatically decrease the tumor hypoxia signals (Figure 2d). In contrast, for mice under hyperoxic breathing with tumors in absence (Ctr + O2) or presence of tumor-focused US treatment (Ctr + O2 + US) but without PFC injection, as well as PFC injected mice D

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX

Letter

Nano Letters

Figure 4. In vivo radiotherapy enhanced by US-triggered tumor oxygenation with nano-PFC. (a) Scheme showing our experimental process. (b) Tumor growth curves of different groups of mice (five mice per group) after various treatments including: untreated control (Ctr), RT, RT + O2, RT + O2 + US, RT + PFC + O2 and RT + PFC + O2 + US. (c) Average tumor weights of different groups as indicated in (b) after treatments. (d) H&Estained and TUNEL-stained tumor slices collected from different groups of mice 1 day after various treatments. P values: *P < 0.05, **P < 0.01, ***P < 0.001, ANOVA.

PFC) plus 30 min hyperoxic breathing before 660 nm laser irradiation (PDT + PFC + O2), and i.v. injection of 200 μL PFC nanoemulsion (30 μL of PFC) plus 30 min simultaneous hyperoxic breathing and tumor-focused US treatment before 660 nm laser irradiation (PDT + PFC + O2 + US). After various treatments, tumor volumes of each group were monitored (Figure 3b,c). It was found that the tumor growth rate of those treated with PDT + PFC + O2 + US was remarkably inhibited compared to the control group, while those of PDT, PDT + O2, PDT + O2 + US, and PDT + PFC + O2 treated groups exhibited moderate growth delaying effect without distinct differences among these groups. In addition, to evaluate whether the generated oxygen may have the potential to produce harmful gas embolization in the tumor, mice were treated by i.v. injection of 200 μL PFC nanoemulsion (30 μL of PFC) plus 30 min simultaneous hyperoxic breathing and tumor-focused US treatment (PFC + O2 + US) (Figure S6). No appreciable delay of tumor growth was found for mice treated by PFC + O2 + US, indicating that the US triggered oxygen release from nano-PFC would not directly induce harmful effect to the tumor. Besides, histological and pathological changes of tumors 1 day after different treatments were evaluated by hematoxylin and eosin (H&E) staining and terminal deoxynucleotidyl transferase-mediated dUTP-biotin nick end labeling (TUNEL) assay, respectively (Figure 3d). From H&E staining results, we found that PDT + PFC + O2 + US treated tumors exhibited

treatment of tumors, chlorin e6 (Ce6), a commonly used photosensitizer, was modified with three hexylamine via amide bonds to increase its hydrophobicity, and then loaded into the hydrophobic lipid bilayers of polyethylene glycol (PEG) modified liposomes (Figure S5a). The singlet oxygen (1O2) production under the excitation of 660 nm light was determined by singlet oxygen sensor green (SOSG) based on its fluorescence recovery in the presence of 1O2. Liposome@ Ce6 showed high efficiency in light-triggered 1O2 generation, proving the potential use of Liposome@Ce6 as an effective agent for PDT (Figure S5b). Such liposomal Ce6 exhibited excellent tumor passive homing via the enhanced permeability and retention (EPR) effect (Figure S5c,d) to enable effective tumor-targeted PDT. In our experiment, when the tumor size reached ∼100 mm3, those 4T1 tumor bearing mice were randomly divided into six groups (5 mice per group), with one group received an i.v. injection of 200 μL of saline, whereas the other five groups received i.v. injection of 200 μL of liposomal Ce6 (0.8 mg/mL in terms of Ce6) (Figure 3a). At 24 h post injection (p.i.), those 5 group mice with liposomal Ce6 injection received different treatments as follows: 30 min 660 nm laser irradiation on the tumor at 20 mW cm−2 (PDT), 30 min hyperoxic breathing followed by 30 min 660 nm laser irradiation (PDT + O2), 30 min simultaneous hyperoxic breathing and tumor-focused US treatment followed by 30 min 660 nm laser irradiation (PDT + O2 + US), i.v. injection of 200 μL PFC nanoemulsion (30 μL of E

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX

Letter

Nano Letters

Figure 5. CT 26 tumor model experiments. (a) Representative PA images of CT26 solid tumors to determine tumor oxygenation status by measuring the ratios of oxygenated hemoglobin (λ = 850 nm) and deoxygenated hemoglobin (λ = 750 nm) before and 30 min after indicated treatments. (b) Quantification of the oxyhemoglobin saturation in the tumor. (c) Representative immunofluorescence images of tumor slices stained by the Hypoxyprobe. (d) Quantification of tumor hypoxia and blood vessel densities for different groups shown in (c). (e) Tumor growth curves of different groups of mice (five mice per group) after various treatments including: untreated control (Ctr), PDT + PFC + O2, PDT + PFC + O2 + US, RT + PFC + O2, RT + PFC + O2 + US. (f) Average tumor weights of different groups as indicated in (e) after various treatments. P values: *P < 0.05, **P < 0.01, ***P < 0.001, ANOVA.

tumor growth. In contrast, RT alone, RT + O2, RT + O2 + US, RT + PFC + O2 all showed moderate inhibition effects on tumor growth (Figure 4b,c). As revealed by H&E and TUNEL staining, the most severe cell morphology change and cell apoptosis were observed in tumors from mice treated with RT + PFC + O2 + US compared to other groups. Our results collectively evidenced that the enhanced tumor oxygenation by nano-PFC triggered by tumor-focused US would be greatly helpful in overcoming hypoxia-associated resistance in both PDT and RT, resulting in remarkably improved therapeutic outcomes for both types of therapies (Figure 4d). At last, to prove the universality of this strategy for other types of solid tumors, a CT-26 mouse colon cancer model was chosen to repeat the above experiments with 4T1 tumors. Similar to the effect for 4T1 tumors, mice bearing CT-26 tumors with sizes of ∼100 mm3 showed a fast tumor oxygenation when treated with PFC + O2 + US as uncovered by PA imaging, and its sO2 Tot increased from 17% to 49% post such treatment, whereas those treated with O2 + US and PFC + O2 did not show obvious tumor oxygenation increase

most severe damage, whereas moderate damages were observed for tumors on mice received treatments of PDT, PDT + O2, PDT + O2 + US and PDT + PFC + O2. Moreover, TUNEL assay showed consistent results and indicated that PDT + PFC + O2 + US was the most effective in inducing tumor cell apoptosis. Radiotherapy (RT), an important first-line treatment modality in oncology, utilizes the ionizing radiation (e.g., X-ray, γ-ray) to induce DNA damage and thus kills cancer cells.42,43 The surrounding oxygen is known to be able to react with DNA breaks induced during RT so as to prevent DNA repairing by tumor cells and enhance RT-induced cell killing.44 Again, six groups of mice were used in our experiment: untreated control, RT, RT + O2, RT + O2 + US, RT + PFC + O2, and RT + PFC + O2 + US. US treatment was applied on the tumor for 30 min when mice were breathing pure oxygen (Figure 4a). The radiation dose used for RT was 6 Gy. Similar to the results of PDT treatment, RT treatment of tumors on PFC-injected mice breathing pure oxygen and exposed to tumor-focused US (RT + PFC + O2 + US) was also the most effective in inhibiting the F

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX

Letter

Nano Letters

was redissolved in PBS for future use. The morphology and size distribution of nano-PFC was characterized by freeze-etching replication transmission electron microscopy and Malvern zetazizer. Preparation of Hydrophobic Ce6 and Liposome@Ce6. For the fabrication of hydrophobic Ce6, Ce6, hexylamine, EDC, NHS, and trimethylamine (TEA) (molar ratio = 1:4:4:4) were dissolved in anhydrous dichloromethane and stirred at room temperature for 24 h. Then, the reaction mixture was condensed by rotary evaporation and purified by thin layer chromatography (TLC) using dichloromethane:ethyl acetate (1:2, v/v) as the solvent system (RF, 0.7). Afterward, the individual band was scraped, dispersed with methanol, and centrifuged to collect the supernatant. Finally, the hydrophobic Ce6 was obtained by rotary evaporation. To prepare Liposome@Ce6, DPPC, cholesterol, DSPE-PEG (5k), and hydrophobic Ce6 at a 6:4:0.5:0.5 molar ratio were dissolved in chloroform. After dissolution, the solution was blow-dried by N2 to form a lipid film. Then, 2 mL of PBS solution was added and stirred at 65 °C for 30 min. The suspension was extruded through 200 nm filters at 50 °C for 40 times then purified by using G-50 sephadex column to obtain Liposome@Ce6. Measurement of O2 Release from Nano-PFC Triggered by US. First, 4 mL of as-prepared PFC solution in a 50 mL sample tube was stored in an aseptic oxygen chamber (O2 flow rate = 5 L/min) for 1 min for PFC oxygenation. The oxygen concentrations in aqueous solutions pretreated with N2 bubble were measured using a portable dissolved oxygen meter (Rex, JPBJ-608, China) before and after adding 4 mL of oxygenated PFC solution. To explore the effect of US treatment on the oxygen release from PFC, both aqueous solutions with and without adding of PFC solutions were first recorded as abovementioned. Then, 350 s later, an external US treatment was applied onto this tube for 500 s and oxygen concentration was recorded for about 1200 s in total (1 MHz, 3.5 W). Cellular Experiments. Murine breast cancer 4T1 cells and murine colon cancer CT-26 cells were obtained from American Type Culture Collection (ATCC) and cultured at 37 °C in a humidified atmosphere containing 5% CO2. All cells were cultured in normal RPMI-1640 culture medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/ streptomycin. For in vitro methyl thiazolyltetrazolium (MTT, Sigma-Aldrich) assay, both 4T1 and CT-26 cells were seeded into 96-well plates at a density of 1 × 104 cells per well. Then, 24 h post cell seeding, both types of cells were incubated with various amounts of PFC for additional 24 h. Then, 25 μL MTT stock solution (5 mg/mL) was added into each well and incubated for another 4 h before measuring it absorbance at 570 nm using a microplate reader (Bio-Rad, model 680). Blood Circulation Measurement of Nano-PFC. The blood circulation profile of nano-PFC was measured by gas chromatography−mass spectrometry following the literature protocol with slight modifications.49 Blood samples were obtained by drawing approximately 25 μL of blood from female nude mice at different time points after i.v. injection of 200 μL of PFC nanoemulsion (containing 30 μL of perfluoro15-crown-5-ether). The blood samples were dissolved in 20 μL lysis buffer (1% Triton X-100) to dissolve the blood. 1,1,1,3,3Pentafluorobutane (500 μL) was added to each sample and the mixture was vortex-mixed for 60 s. Afterward, 50 μL of ethanol was added into each sample, which was placed in an ice-cold ultrasonic bath for 30 s and then vortex-mixed for 300 s. The mixture was centrifuged (4 °C) at 1500 g for 2 min and frozen

(Figure 5a,b). Furthermore, the ex vivo immunofluorescence staining analysis also demonstrated that PFC + O2 + US treatment could efficiently relieve the tumor hypoxia. The positive hypoxia area remarkably decreased from ∼88% for control group to ∼74% and ∼20% for PFC + O2 and PFC + O2 + US groups, respectively (Figure 5c,d). In vivo PDT and RT were further carried out on CT-26 tumor bearing mice, which were randomly divided into five groups for different treatments: saline, PDT + PFC + O2, PDT + PFC + O2 + US, RT + PFC + O2, and PDT + PFC + O2 + US. PDT and RT treatments were conducted following exact the same parameters as we used for the treatment of 4T1 tumor model. As expected, we found that the enhanced tumor oxygenation resulted from treatment of “PFC + O2 + US” could also remarkably enhance the therapeutic efficacies of PDT and RT for CT-26 tumors (Figure 5e,f). All these results strongly demonstrate that our PFC + O2 + US strategy could be a universal approach in improving the tumor oxygenation and enhancing oxygen involved therapies such as PDT and RT. Conclusions. In this work, PFC nanoemulsion was prepared by using HSA as the stabilizer to coat PFC nanodroplets. Such PFC nanodroplets, though being biocompatible and nontoxic, showed a high oxygen carrying ability and a US-response burst-like oxygen release profile. In vivo PA imaging and ex vivo immunofluorescence staining collectively evidenced that our developed US-triggered local tumor oxygenation strategy using nano-PFC as the oxygen-shuttle could be an effective approach to dramatically relieve hypoxia for different types of solid tumor models. Owing to the reversed hypoxia-associated resistances by this method, remarkably improved therapeutic efficacies of PDT and RT treatments of both 4T1 and CT-26 tumor models have been achieved over the conventional treatment protocols. The tumor oxygenation strategy developed here could be superior to other hypoxia relieving methods as its efficiency is not limited by the inherent oxygen-binding ability of hemoglobin in red blood cells (for tumor vasculature normalization method), or the amount of available endogenous H2O2 (10−50 μM) inside the tumor (the in situ oxygen production method).45,46 Importantly, because several formulations of PFC emulsions have been either clinically approved or in late-phase clinical trials as blood substitutes, whereas ultrasound is a noninvasive clinically adapted instrument, our proposed US-triggered tumor oxygenation method with nano-PFC may shows great potential for future clinical translation. Such a technique, if successful translated, may not only be useful to improve PDT and RT as demonstrated in this work but also bring therapeutic benefits to other types of mainstream cancer therapies such as chemotherapy and immunotherapy.9,47,48 Materials and Methods. Materials. Dipalmitoyl-sn-glycero-3-phosphocholine (DPPC) was purchased from NOF America Corporation. 1,2-Distearoyl-sn-glycero-3-phosphoethanolamine-N-[methoxy(poly(ethylene glycol))-5000] (DSPEPEG) was purchased from Laysan Bio Inc. Cholesterol (CH) was purchased from J&K Co. Ltd. Human serum albumin (HSA) was purchased from Sigma-Aldrich. Perfluoro-15crown-5-ether was purchased from J&K Scientific Ltd. Preparation of PFC Nanoemulsion. First, 300 μL of perfluoro-15-crown-5-ether (PFC) was added to 4 mL of PBS solution containing 40 mg of HSA. The mixture was oscillated slightly and then emulsified in an ice bath using an ultrasonic probe for 200 s. The obtained emulsion was centrifuged at 8000 rmp/min for 3 min and sedimentation G

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX

Letter

Nano Letters at −30 °C for 2 h. Just before analysis by gas chromatography− mass spectrometry (GC−MS), the upper ethanolic phase was withdrawn and the lower phase was put into another 2 mL tube. Them, 1 mL of this solution was injected into the chromatograph for GC-MS measurement to determine the concentrations of perfluoro-15-crown-5-ether in each sample. All experiments were performed under 4 °C. Animal Tumor Models. Female nude mice were purchased from Peng Sheng Biological Technology Co. Ltd. and operated under protocols approved by Soochow University Laboratory Animal Center. Then, 2 ×106 4T1 cells or CT-26 cells suspended in 50 μL of PBS were subcutaneously injected into the back of each mouse to build the tumor model. In Vivo Photoacoustic Imaging. Mice bearing 4T1 or CT26 tumors with sizes of ∼100 mm3 were randomly divided into three groups including control group injected with 200 μL of saline, mice i.v. injected with 200 μL of PFC nanoemulsion (containing 30 μL of PFC) plus simultaneous hyperoxic breathing for 30 min, and mice i.v. injected with 200 μL of PFC nanoemulsion (30 μL of PFC) plus simultaneous hyperoxic breathing and tumor focused US treatment for 30 min. Then, their tumor oxygenations before and after relative treatments were recorded by using a PA imaging system (FujiFilm VisualSonics Inc.) utilizing the Oxy-hem mode (750 and 850 nm). Ex Vivo Immunofluorescence Staining. Mice bearing 4T1 tumors were randomly divided into five groups including: 1, control group (with saline injection); 2, O2 group (saline injection plus 30 min hyperoxic breathing); 3, O2 + US group (simultaneous hyperoxic breathing and tumor focused US treatment for 30 min); 4, PFC + O2 group (i.v. injection of PFC plus hyperoxic breathing); 5, PFC + O2 + US group (i.v. injection of PFC plus simultaneous hyperoxic breathing and tumor focused US treatment for 30 min). After receiving various treatments, the tumors were surgically excised at 90 min after intraperitoneal injection of Pimonidazole hydrochloride (0.6 mg/mouse) (Hypoxyprobe-1 plus kit, Hypoxyprobe Inc.), mounted with optimum cutting temperature (OCT) compound (Sakura Finetek) and then sliced into 8 μm slices for further staining. For detection of Pimonidazole, antipimonidazole mouse monoclonal antibody C (dilution 1:200, Hypoxyprobe Inc.) was used as primary antibody, whereas Alex 488conjugated goat antimouse secondary antibody (dilution 1:200, Jackson Inc.) was exploited for further staining and fluorescence imaging. For CD31 staining, rat-antimouse CD31 antibody and rhodamine-conjugated donkey−antirat antibody were used as the primary and secondary antibody, respectively, to indicate the tumor blood vessels. For HIF-1α staining, mouse anti-HIF-1α monoclonal antibody and goat− antimouse IgG antibody conjugated with FITC were used as primary and secondary antibody, respectively, according to recommended procedure by the manufacturers. In Vivo PDT and RT. Mice bearing 4T1 tumors were randomly divided into six groups including: 1, control group (with saline injection);2, PDT group; 3, PDT + O2 group; 4, PDT + US + O2 group; 5, PDT + PFC + O2 group; and 6, PDT + PFC + O2 + US group. For PDT treatment, mice were first i.v. injected with 200 μL of liposomal Ce6 (Ce6 dose = 0.8 mg/kg) according to our previously developed methods 24 h in advance. Then, mice of group 5 and 6 were i.v. injected with 200 μL of PFC nanoemulsion (30 μL of PFC). For hyperoxic breathing, mice were equipped with oxygen masks flowed with pure oxygen for 30 min. For US treatment, a US probe was

applied to the surface of the tumor site for 30 min during hyperoxic breathing. Afterward, to conduct the PDT treatment, mice were irradiated under a 660 nm laser at the power density of 0.02 W/cm2 for 30 min. For RT treatment, mice bearing 4T1 tumors were randomly divided into six groups similar to that for PDT except that RT was applied in this experiment: 1, control group; 2, RT group; 3, RT + O2 group; 4, RT + US + O2 group; 5, RT + PFC + O2 group; and 6, RT + PFC + O2 + US group. Mice in groups 2, 3, 4, 5, and 6 received an X-ray irradiation with a dose of 6 Gy. The other experiment methods and parameters were identical to that applied in PDT as above-mentioned. After various treatments, the lengths and widths of the tumors were measured by a digital caliper every 2 days for a period of 14 days. The tumor volume was calculated according to the following formula: volume = width2 × (length/2). At 14 day post-treatment, the mice from each group were scarified and each tumor was excised for weighting. On the other hand, at 1 day post various treatments, tumors from different groups were collected, fixed using 4% formaldehyde, mounted with paraffine, sliced, and stained with H&E and TUNEL following standard protocols. Then, each slice was examined by a digital microscope (Leica QWin).

■

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.nanolett.6b02365. TEM image of nano-PFC; in vitro cytotoxicity assay for nano-PFC; blood circulation profile of nano-PFC; HIF1α stained immunofluorescence imaging of tumor slices; characterization and tumor uptake of Liposome@Ce6; tumor growth on nano-PFC injected control mice with US treatment but no other treatment. (PDF)

■

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This work was partially supported by the National Basic Research Programs of China (973 Program) (2012CB932600), the National Natural Science Foundation of China (51525203, 51132006), a Juangsu Natural Science Fund for Distinguished Young Scholars (BK20130005), Collaborative Innovation Center of Suzhou Nano Science and Technology, and a Project Funded by the Priority Academic Program Development (PAPD) of Jiangsu Higher Education Institutions.

■

REFERENCES

(1) Brown, J. M.; Wilson, W. R. Nat. Rev. Cancer 2004, 4, 437−447. (2) Höckel, M.; Vaupel, P. Semin. Oncol. 2001, 28, 36−41. (3) Yang, Z. F.; Poon, R. T.; To, J.; Ho, D. W.; Fan, S. T. Cancer Res. 2004, 64, 5496−5503. (4) DeClerck, K.; Elble, R. C. Front. Biosci., Landmark Ed. 2010, 15, 213−225. (5) Matsuo, M.; Matsumoto, S.; Mitchell, J. B.; Krishna, M. C.; Camphausen, K. Semin. Radiat. Oncol. 2014, 24, 210−217. H

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX

Letter

Nano Letters (6) Zannella, V. E.; Dal Pra, A.; Muaddi, H.; McKee, T. D.; Stapleton, S.; Sykes, J.; Glicksman, R.; Chaib, S.; Zamiara, P.; Milosevic, M. Clin. Cancer Res. 2013, 19, 6741−6750. (7) Jordan, B.; Sonveaux, P. Front. Pharmacol. 2012, 3, 94. (8) Gordijo, C. R.; Abbasi, A. Z.; Amini, M. A.; Lip, H. Y.; Maeda, A.; Cai, P.; O’Brien, P. J.; DaCosta, R. S.; Rauth, A. M.; Wu, X. Y. Adv. Funct. Mater. 2015, 25, 1857−1587. (9) Huang, C.-C.; Chia, W.-T.; Chung, M.-F.; Lin, K.-J.; Hsiao, C.W.; Jin, C.; Lim, W.-H.; Chen, C.-C.; Sung, H.-W. J. Am. Chem. Soc. 2016, 138, 5222−5225. (10) Fan, W.; Bu, W.; Shen, B.; He, Q.; Cui, Z.; Liu, Y.; Zheng, X.; Zhao, K.; Shi, J. Adv. Mater. 2015, 27, 4155−4161. (11) Prasad, P.; Gordijo, C. R.; Abbasi, A. Z.; Maeda, A.; Ip, A.; Rauth, A. M.; DaCosta, R. S.; Wu, X. Y. ACS Nano 2014, 8, 3202− 3212. (12) Gong, H.; Chao, Y.; Xiang, J.; Han, X.; Song, G.; Feng, L.; Liu, J.; Yang, G.; Chen, Q.; Liu, Z. Nano Lett. 2016, 16, 2512−2521. (13) Yoon, H. Y.; Koo, H.; Choi, K. Y.; Lee, S. J.; Kim, K.; Kwon, I. C.; Leary, J. F.; Park, K.; Yuk, S. H.; Park, J. H.; Choi, K. Biomaterials 2012, 33, 3980−3989. (14) Winter, P. M.; Cai, K.; Caruthers, S. D.; Wickline, S. A.; Lanza, G. M. Expert Rev. Med. Devices 2007, 4, 137−145. (15) Kaneda, M. M.; Caruthers, S.; Lanza, G. M.; Wickline, S. A. Ann. Biomed. Eng. 2009, 37, 1922−1933. (16) Mason, R. P.; Antich, P. P.; Babcock, E. E.; Gerberich, J. L.; Nunnally, R. L. Magn. Reson. Imaging 1989, 7, 475−485. (17) Rapoport, N.; Nam, K.-H.; Gupta, R.; Gao, Z.; Mohan, P.; Payne, A.; Todd, N.; Liu, X.; Kim, T.; Shea, J.; et al. J. Controlled Release 2011, 153, 4−15. (18) Tran, T. D.; Caruthers, S. D.; Hughes, M.; Marsh, J. N.; Cyrus, T.; Winter, P. M.; Neubauer, A. M.; Wickline, S. A.; Lanza, G. M. Int. J. Nanomed. 2007, 2, 515−526. (19) Cohn, C. S.; Cushing, M. M. Crit. Care Clin. 2009, 25, 399−414. (20) Keipert, P. E. Artif. Cells Blood Subst. Biotechnol. 1995, 23, 381− 394. (21) Kim, H. W.; Greenburg, A. G. Artif. Organs 2004, 28, 813−828. (22) Young, L. H.; Jaffe, C. C.; Revkin, J. H.; McNulty, P. H.; Cleman, M. Am. J. Cardiol. 1990, 65, 986−990. (23) von der Hardt, K.; Kandler, M. A.; Brenn, G.; Scheuerer, K.; Schoof, E.; Dötsch, J.; Rascher, W. Crit. Care Med. 2004, 32, 1200− 1206. (24) Castro, C. I.; Briceno, J. C. Artif. Organs 2010, 34, 622−634. (25) Lowe, K. Blood Rev. 1999, 13, 171−184. (26) Cheng, Y.; Cheng, H.; Jiang, C.; Qiu, X.; Wang, K.; Huan, W.; Yuan, A.; Wu, J.; Hu, Y. Nat. Commun. 2015, 6, 8785. (27) Ma, Z.; Monk, T. G.; Goodnough, L. T.; McClellan, A.; Gawryl, M.; Clark, T.; Moreira, P.; Keipert, P. E.; Scott, M. G. Clin. Chem. 1997, 43, 1732−1737. (28) Riess, J. G. Artif. Cells Blood Subst. Biotechnol. 2006, 34, 567− 580. (29) Riess, J. G. Artif. Cells Blood Subst. Biotechnol. 2005, 33, 47−63. (30) Song, G.; Liang, C.; Yi, X.; Zhao, Q.; Cheng, L.; Yang, K.; Liu, Z. Adv. Mater. 2016, 28, 2716−2723. (31) Rapoport, N. Y.; Kennedy, A. M.; Shea, J. E.; Scaife, C. L.; Nam, K.-H. J. Controlled Release 2009, 138, 268−276. (32) Kim, H. J.; Matsuda, H.; Zhou, H.; Honma, I. Adv. Mater. 2006, 18, 3083−3088. (33) Yohe, S. T.; Kopechek, J. A.; Porter, T. M.; Colson, Y. L.; Grinstaff, M. W. Adv. Healthcare Mater. 2013, 2, 1204−1208. (34) Ma, M.; Xu, H.; Chen, H.; Jia, X.; Zhang, K.; Wang, Q.; Zheng, S.; Wu, R.; Yao, M.; Cai, X.; et al. Adv. Mater. 2014, 26, 7378−7385. (35) Dias, A.; Freire, M.; Coutinho, J. A.; Marrucho, I. Fluid Phase Equilib. 2004, 222, 325−330. (36) Lowe, K. C. J. Fluorine Chem. 2001, 109, 59−65. (37) Beard, P. Interface focus 2011, 602. (38) Shao, Q.; Morgounova, E.; Jiang, C.; Choi, J.; Bischof, J.; Ashkenazi, S. J. Biomed. Opt. 2013, 18, 076019−076019.

(39) Westbury, C.; Pearson, A.; Nerurkar, A.; Reis-Filho, J.; Steele, D.; Peckitt, C.; Sharp, G.; Yarnold, J. The British journal of radiology 2014, 25046649. (40) Henderson, B. W.; Gollnick, S. O.; Snyder, J. W.; Busch, T. M.; Kousis, P. C.; Cheney, R. T.; Morgan, J. Cancer Res. 2004, 64, 2120− 2126. (41) Dolmans, D. E.; Fukumura, D.; Jain, R. K. Nat. Rev. Cancer 2003, 3, 380−387. (42) Meijer, T. W.; Kaanders, J. H.; Span, P. N.; Bussink, J. Clin. Cancer Res. 2012, 18, 5585−5594. (43) Song, G.; Liang, C.; Yi, X.; Zhao, Q.; Cheng, L.; Yang, K.; Liu, Z. Adv. Mater. 2016, 28, 2654−2654. (44) Antonovic, L.; Lindblom, E.; Dasu, A.; Bassler, N.; Furusawa, Y.; Toma-Dasu, I. J. Radiat. Res. 2014, 55, 902−911. (45) Kuang, Y.; Balakrishnan, K.; Gandhi, V.; Peng, X. J. Am. Chem. Soc. 2011, 133, 19278−19281. (46) Chen, H.; Tian, J.; He, W.; Guo, Z. J. Am. Chem. Soc. 2015, 137, 1539−1547. (47) Malmberg, K.-J. Cancer Immunol. Immunother. 2004, 53, 879− 892. (48) Hansen-Algenstaedt, N.; Stoll, B. R.; Padera, T. P.; Dolmans, D. E.; Hicklin, D. J.; Fukumura, D.; Jain, R. K. Cancer Res. 2000, 60, 4556−4560. (49) Audran, M.; Krafft, M. P.; De Ceaurriz, J.; Maturin, J. C.; Sicart, M. T.; Marion, B.; Bougard, G.; Bressolle, F. J. Chromatogr., Biomed. Appl. 2000, 745, 333−343.

I

DOI: 10.1021/acs.nanolett.6b02365 Nano Lett. XXXX, XXX, XXX−XXX