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Tumor Microenvironment-Responsive Nanoparticle Delivery of Chemotherapy for Enhanced Selective Cellular Uptake and Transportation within Tumor Wen-Chia Huang, Shih-Hong Chen, Wen-Hsuan Chiang, Chu-Wei Huang, Chun-Liang Lo, Chorng-Shyan Chern, and Hsin-Cheng Chiu Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.6b00956 • Publication Date (Web): 02 Nov 2016 Downloaded from http://pubs.acs.org on November 3, 2016

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Tumor Microenvironment-Responsive Nanoparticle Delivery of Chemotherapy for Enhanced Selective Cellular Uptake and Transportation within Tumor Wen-Chia Huang,† Shih-Hong Chen,†,‡ Wen-Hsuan Chiang,† Chu-Wei Huang,† Chun-Liang Lo,§ Chorng-Shyan Chern⁋ and Hsin-Cheng Chiu*,†

Department of Biomedical Engineering and Environmental Sciences, National Tsing Hua

University, Hsinchu 300, Taiwan

Department of Anesthesiology, National Taiwan University Hospital-Hsinchu Branch, Hsinchu

300, Taiwan §

Department of Biomedical Engineering, National Yang-Ming University, Taipei, 112, Taiwan

⁋

Department of Chemical Engineering, National Taiwan University of Science and Technology,

Taipei 106, Taiwan *To whom correspondence should be addressed. Fax: 886-35718649. Tel: 886-35750829. Email: [email protected] (H.-C. Chiu)

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KEYWORDS. Cellular Delivery, Chemotherapy, Nanoparticles, Tumor Hypoxia, PEG Detachment.

ABSTRACT. A novel drug delivery strategy featured with enhanced uptake of nanoparticles (NPs) by targeted tumor cells and subsequent intratumoral cellular hitchhiking of chemotherapy to deep tumor regions was described. The NP delivery system was obtained from assembly of poly(lactic acid-co-glycolic acid)-grafted hyaluronic acid (HA-g-PLGA) together with an anticancer drug, SN38, in aqueous phase, followed by implementing the NP surface with a layer of methoxypoly(ethylene glycol)-b-poly(histamine methacrylamide) (mPEG-b-PHMA) via hydrophobic association to improve the colloidal stability both in vitro and in vivo. Upon arrival of these PEGylated NPs at the acidic tumor site through the EPR effect, mPEG-b-PHMA became detached from the NP surface by the charge transition of the PHMA blocks from neutral (hydrophobic) to positively charged (hydrophilic) state via acid-induced protonation of their imidazole groups in tumor microenvironment. The exposure of HA shell on the naked NP thus resulted in enhanced uptake of NPs by CD44-expressed tumor cells, including cancer cells and tumor-associated macrophages (TAMs). Along with the TAMs being further chemotactically recruited by hypoxia cells, the engulfed nanotherapeutics was thus transported into the avascular area in which the anticancer action of chemotherapy occurred by virtue of the drug release alongside PLGA degradation, similar to those arising in other tumor non-hypoxia regions.

1. INTRODUCTION Malignant tumors, characterized by immoderately growing and invading normal tissues alongside fast cell proliferation, severely threaten human life. To enhance the efficacy of chemotherapeutics and reduce their side effects, various nanoparticle (NP)-based drug delivery


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strategies have been extensively studied.1-5 Targeting of NP-based chemotherapeutics to tumors relies largely on the enhanced permeability and retention (EPR) effect and the implementation of NP surfaces with pertinent ligands which enable NPs to specifically bind with cancer cells.1,5 For instance, the NPs outlayered with hyaluronic acid (HA) have demonstrated their enhanced tumor uptake by targeting to the cellular surface HA receptor, CD44, on cancer cells and tumorassociated macrophages (TAMs).6-9 Obviously, the presence of CD44 on these cancer and tumor associated cells is required in order to serve as an effective HA docking receptor. Owing to CD44 molecules on mammalian cell surfaces playing a pivotal role in mediating cell–cell interactions and cell adhesion and migration, it is not surprising that the CD44 molecules were found largely on invasive and metastatic cancer cells and wandering TAM cells.10,11 Although the HA-based nanotherapeutics has shown significant accumulation in tumors, the rapid clearance of the NPs by reticuloendothelial system (RES) in liver and spleen remains a major concern. In order to reduce recognition and phagocytosis by RES and concomitantly prolong residence in blood circulation, the nanomedicine is often elaborated with poly(ethylene glycol) (PEG) on the outer surfaces.12 Nevertheless, the transportation of chemotherapy into deep tumor tissues, particularly the hypoxia regions, is still often impeded even though the nanomedicine approaches are exploited. This is because hypoxic cancer cells in solid tumors are far away from nearby blood vessels (a distance of ca 70-100 µm), leading to rather limited access of therapeutics to these avascular regions.13-15 Unfortunately, under the circumstances after chemo/radio therapeutic treatments, the remnant tumor cells localized in hypoxia are closely related to local recurrence and metastasis of tumors.16,17 To overcome this problem, an effective drug delivery approach capable of delivering therapeutics into tumor avascular tissues is in great demand.


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Wang et al. have recently developed an NP-based drug carrier system from amphiphilic block copolymers of 7-ethyl-10-hydroxylcamptothecin (SN38) analogues with varying NP sizes in the range 20~300 nm in order to study the size effect on cancer drug delivery.18 It was found that the optimal NP size pertinent to the EPR-associated tumor accumulation is ca 160 nm in diameter, yet the smaller NPs (ca 30 nm) exhibit a superior performance in penetration across the dense extracellular matrix into deep tumor tissues. For this reason, Sun et al. have developed a “cluster-bomb”-like nanoassemblies for high therapeutic efficacy in cancer treatment.19 With a relatively large particle size, the cluster bombs can transport the small drug-loaded “bomblets” into tumor more efficiently by the EPR effect. Once arriving at tumor site, the cluster bombs rapidly dissociated and released small bomblets in response to the tumor environmental pH, thereby leading to the deep tissue penetration of therapeutics within tumor. Distinct from the aforementioned nanocarriers requiring a complicated procedure in materials synthesis, a nanovehicle system capable to enhance its tumor uptake and penetration was developed simply by the surface decoration of poly(lactic-co-glycolic acid) (PLGA)-based nanotherapeutics with pH-responsive N-acetyl histidine modified D-α-tocopheryl polyethylene glycol succinate.20 By virtue of the pH-inducible surface charge transition and small particle size, the drug-carrying NPs show excellent therapeutic penetration and accumulation in tumor hypoxia. Other approaches involving the use of monocytes/tumor associated macrophages (TAMs) as a cellular Trojan carrier, by taking advantages of their inherent capability of being chemotactically recruited to tumor hypoxia,21,22 have demonstrated a remarkable performance in highly active targeting to these avascular areas,14,15,23 though the application was somewhat restricted by the tedious procedures of monocytes/macrophages isolation. The solid tumor microenvironment is rather heterogeneous and comprised of various cells such as cancer cells, fibroblasts,


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lymphocytes, monocytes/TAMs, etc.24,25 Considering the monocytes/TAMs amounting to 50% of the total tumor mass, it is worthy to develop a smart drug delivery system that can accumulate within tumor via EPR effect and then be readily and selectively taken up by tumor cells including both cancer cells and TAMs, while the latter can further intratumorally deliver nantherapeutics in a hitchhiking manner into avascular regions. Herein, a novel strategy dealing with the enhanced tumor cellular uptake and penetration of nanotherapeutics in combination with the delicately designed NP-based delivery system and cellular transport of these carriers by tumor cells across the dense tumor matrices into avascular hypoxic regions was described. To validate the concept, NPs obtained from assembly of biodegradable PLGA-grafted hyaluronic acid (HA-g-PLGA) along with an anticancer drug, SN38, in aqueous phase were adopted in this work. These NPs were coated with a layer of methoxypoly(ethylene glycol)-b-poly(histamine methacrylamide) (mPEG-b-PHMA) copolymer via hydrophobic association to improve their colloidal stability. In this manner, these NPs stabilized by surface PEGylation greatly reduce the rapid systemic clearance by the RES and, consequently, increase their accumulation in tumors via the EPR effect. Upon arrival of NPs at tumors, the PEGylated polymeric layer detaches from NP surfaces due to the transition of PHMA blocks from hydrophobic to hydrophilic state by extensive protonation of the imidazole groups in weak acidic microenvironment of tumors. The PEG shedding from NP surfaces has been shown to facilitate cellular uptake and even penetration into deep tumor tissues.26-28 After PEG detachment, the HA-rich shell of NP then becomes naked. This will then further significantly promote the cellular uptake of NPs by CD44 presenting cancer cells and TAMs.22,29,30 Owing to the inherent chemotactic recruitment of TAMs by hypoxic cells, the engulfed therapeutic payloads can be further delivered into tumor avascular regions21,22 and,


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subsequently, released in these areas by virtue of PLGA degradation. A schematic illustration of the proposed drug delivery strategy is shown in Scheme 1.

Scheme 1. Schematic of the strategy developed in this work to enhance uptake of nanotherapeutics by cancer cells and TAMs and penetration into deep tumor tissues by TAMs via cellular hitchhiking transport. 2. EXPERIMENTAL SECTION 2.1 Materials. Sodium hyaluronate, the sodium salt of HA, with an average molecular weight (MW) of 15 kDa was purchased from Lifecore Biomedical. Tetrabutylammonium hydroxide (TBA), SN38 and 3-(4,5-dimethyl-thiazol-2yl)-2,5-diphenyl tetrazolium bromide (MTT) were obtained from Sigma-Aldrich. Deionized water was produced by the Milli-Q Synthesis System (18 MΩ, Millipore). All chemicals were reagent grade and used as received. Tramp-C1 mouse prostate cancer cell line was purchased from American Type Culture Collection (CRL-2730). Mouse


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macrophage-like Raw 264.7 cell line and NIH 3T3 mouse embryo fibroblast cell line were obtained from Food Industry Research and Development Institute (Taiwan). RPMI-1640 medium was purchased from Gibco. The Dulbecco’s modified Eagle medium (DMEM) was acquired from Invitrogen. Cell lines were incubated at 37 °C in humidified 5% CO2/air atmosphere. Six- to 8-week-old C57BL/6JNarl male mice were purchased from National Laboratory Animal Center, Taiwan. The approved guides for the care and use of laboratory animals by the Institutional Animal Care and Use Committee (IACUC) of National Tsing Hua University, Taiwan (approved number: IACUC:10243) were followed at all time. All surgeries were performed under Zoletil/Rompun anesthesia, and all efforts were made to minimize suffering. 2.2 Synthesis of HA-g-PLGA HA (1.0 g) was dissolved in water (10 mg/mL). With the medium pH being adjusted to 3.0 by 1.0 N HCl with stirring for 1 h, the HA aqueous solution was dialyzed against deionized water to remove the NaCl. TBA in 4-fold molar excess with respect to the HA repeating unit was then added to the HA solution under magnetic stirring for 12 h. The resulting solution (10 mL) was concentrated to ca 100 mg/mL by rotary evaporation under vacuum and then mixed with the DMSO solution of PLGA (3.96 g; LA/GA 67/33; Mn 10000 g/mol) to a water/DMSO ratio of 2.0. HA-g-PLGA was prepared by the coupling reaction mediated by 1-ethyl-3-(3dimethylaminopropyl)carbodiimide (EDC) and N-hydroxysuccinimide (NHS) (both 20 mol% with respect to the HA repeating unit) at 25 oC. The crude product was subjected to dialysis against DMSO (Cellu·Sep H1 MWCO 50000) at 25 oC for 5 days, followed by dialysis against deionized water at 4 oC for 3 days. The final product was collected by lyophilization. The synthesis route and chemical structure of the copolymer is schematically illustrated in Figure


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S1a (Supporting Information). The extent of grafting PLGA in the HA backbone was determined by 1H-NMR in DMSO-d6 at ambient temperature. Based on the relative integral ratio of the feature signals of methine protons at δ 5.1 ppm from the lactic acid residues of PLGA and methyl protons at δ 1.9 ppm from HA, the PLGA content was estimated to be ca 9.5 mol% with respect to the repeating unit of HA (Figure S1b). The HA-g-PLGA (MW 5.27×104 g/mol) used in this work was thus obtained. 2.3 Synthesis of Rhodamine B-Labelled mPEG-b-PHMA Copolymer The

copolymer,

mPEG-b-PHMA,

was

prepared

by

extensive

amidation

of

methoxypoly(ethylene glycol)-b-poly(methacrylic acid) (mPEG-b-PMAA) with histamine. The polymer precursor, mPEG-b-PMAA, was first prepared by radical polymerization of methacrylic acid (6.0 mmol) using 4,4’-azobis-(4-cyanopentanoic acid)-conjugated mPEG (0.04 mmol, MW 5000) as the macromolecular initiator and cysteamine (0.28 mmol) as the chain transfer reagent in ethanol under N2 atmosphere at 70 oC over a period of 24 h. This was followed by precipitation from diethyl ether and drying under vacuum after filtration. The synthesis and characterization of the PEG macroinitiator was described in detail elsewhere.4 The mPEG-bPMAA copolymer (150 mg) was then labeled with rhodamine B (RhB) at the amino end of PMAA block via aminolysis of the succinimidyl ester of RhB (12.5 mg) in the presence of 4dimethylaminopyridine (2.5 mg) as the catalyst in DMSO at 25 oC for 24 h. After dialysis (Cellu Sep MWCO 3500) against deionized water for 3 days, the purified copolymer was then collected by lyophilization. To obtain mPEG-b-PHMA, the PMAA blocks of RhB-labeled mPEG-bPMAA were subjected to thorough amidation with histamine (in 3-fold molar excess with respective to the MAA residues) using the DCC/NHS coupling system in DMSO at 60 °C for 72 h. The crude product was filtered (0.45 µm PTFE filter) twice to remove dicyclohexylurea,


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followed by dialysis (Cellu Sep MWCO 3500) against deionized water for 7 days to remove unreacted histamine. The product was then collected by lyophilization. The detailed synthetic route is illustrated in Figure S2a. The compositions of mPEG-b-PMAA and mPEG-b-PHMA were determined by 1H-NMR in DMSO-d6 (Figures S2b and S2c). The conjugation efficiency of histamine with mPEG-b-PMAA, leading to the production of mPEG-b-PHMA, was evaluated to be 92.3%. The weight-average molecular weight (Mw 7700 g/mol) and polydispersity (1.13) of mPEG-b-PMAA were determined by gel permeation chromatography (GPC) (PL-Aquagel-OH columns: GF083; 100-30K, GF084; 10K-200K, and GF086; 200K-10M, calibrated with PEG standards) using tris buffer (0.01 M) as the eluent at a flow rate of 1.0 mL/min under RI detection. The average molecular weight of mPEG-b-PHMA was estimated to be 1.32 ×104 g/mol from the molecular weight of mPEG-b-PMAA and the content (mol%) of histamine residues within the copolymer. The purity was confirmed by GPC using citrate buffer (pH 5.0) as the eluent at a flow rate of 1.0 mL/min under RI detection. 2.4 pH-Induced Phase Separation of mPEG-b-PHMA in Aqueous Phase The pH-evolved phase separation of mPEG-b-PHMA in aqueous phase was characterized by changes in fluorescence emission of pyrene serving as a nonpolar probe. Aliquots (20.0 µL) of the pyrene solution (6.0×10-5 M) in acetone were placed in vials and evaporated under vacuum. The aqueous polymer suspension (0.1 mg/mL, 1.0 mL) at prescribed pH was added into the vial, while the final pyrene concentration reached 6.0×10-7 M. The solution was kept at 4 oC overnight. Fluorescence characterization was conducted by measuring the intensity ratio (I3/I1) of the third vibronic band at 383 nm to the first at 373 nm of the pyrene fluorescence emission spectra in aqueous copolymer solutions with different pH. The excitation was performed at 337


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nm and the emission recorded in the range 350~500 nm on a Hitachi F-7500 fluorescence spectrometer. The light scattering intensity of aqueous mPEG-b-PHMA suspensions (0.10 mg/mL) at preset pH was monitored by a Malvern Zetasizer Nano-ZS instrument at a scattering angle of 173° equipped with a 4 mW He-Ne laser operating at λ = 632.8 nm. Prior to measurement, the sample was equilibrated at 25 °C for 6 h. The data reported herein represent an average of at least triplicate measurements. The relative light scattering intensity of each sample at preset pH was normalized against the mean intensity of the polymer suspension at pH 7.4. 2.5 Nanoparticle Preparation HA-g-PLGA (6.0 mg) and SN38 (1.2 mg) were dissolved in DMSO (600 µL). The polymer/drug solution was added dropwise into phosphate buffer (pH 7.4, 4.0 mL, ionic strength 0.01 M) under pulse sonication (on/off 5/5 s, Sonics Vibracell VCX 750) for 20 min in an ice/water bath. After dialysis against phosphate buffer (pH 7.4) to remove organic solvent and filtration with 0.45 µm PVDF filter to eliminate unloaded SN38 aggregates, the SN38-loaded nanoparticles (referred to hereinafter as NP-I) used in this work were obtained. Modification of nanoparticle surfaces with mPEG-b-PHMA was carried out by slow addition of the aqueous diblock copolymer solution (200 µL, pH 3.0, 10 mg/mL) into the NP-I suspension (pH 7.4, 1.0 mL). With the pH being kept at 7.4, the mixture was stirred at 4 oC for 12 h to obtain the mPEGb-PHMA decorated NPs (denoted as NP-II). Instead of mPEG-b-PHMA, the pH-insensitive diblock copolymer, mPEG-b-PLGA (MW of PEG 5000 g/mol, MW of PLGA 10000 g/mol; LA/GA 75/25) was also used to prepare the other PEGylated NPs in this work as an important negative control (NP-III). Owing to the solubility limitation of mPEG-b-PLGA in aqueous phase,


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the diblock copolymer along with HA-g-PLGA and SN38 was dissolved in DMSO with the weight ratio equivalent to mPEG-b-PHMA in NP-II. The solution was then added dropwise into phosphate buffer (pH 7.4, 4.0 mL, ionic strength 0.01 M) under pulse sonication (on/off 5/5 s) for 20 min in an ice/water bath, followed by the same procedure pertinent to the nanoparticle purification described previously. To determine the level of drug loading, an aliquot of nanoparticle suspension was withdrawn and diluted with DMSO to a volume ratio of DMSO/water = 9/1, followed by the UV/Vis spectroscopic measurements. The destruction of nanoparticle structure by DMSO was confirmed by dynamic light scattering. The absorbance measurement of the solution of drug-free nanoparticles in the DMSO/water co-solvent system was also carried out to ensure the exclusive absorbance of SN38 at 378 nm. Drug loading efficiency and loading content were calculated according to the following formulae, respectively: Drug loading efficiency (%) =

Drug loading content (%) =

weight of loaded SN38 × 100 % weight of SN38 in feed

weight of loaded SN38 × 100 % weight of the nanoparticles

2.6 Structural Characterization The measurements of the mean hydrodynamic diameter (Dh), size distribution (polydispersity index, PDI) and ζ-potential of nanoparticles in aqueous solution were conducted on a Malvern Zetasizer Nano-ZS Instrument. The results reported herein represent an average of at least triplicate measurements. The sample for transmission electron microscopy (TEM) examination was prepared by placing a few drops of aqueous nanoparticle suspensions on a 300mesh copper grid covered with carbon and allowed to stand at 20 °C for 20 s. Excess solution on the grid was gently removed with absorbent paper. The sample was then dried under vacuum for


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2 days. The TEM image was obtained on a JEOL JEM-1200 CXII microscope operating at an accelerating voltage of 120 kV. 2.7 Förster Resonance Energy Transfer (FRET) Assay A hydrophobic fluorescence dye, 3,3'-dilinoleyloxacarbocyanine perchlorate (DiO) (1.0 µg), capable to form the FRET donor-acceptor pairing with RhB, was loaded during nanoparticle preparation. The emission spectra of the aqueous suspensions of DiO-loaded NP-II (1.0 mg/mL) at different pH were determined by the fluorescence spectrometer. For the FRET measurements, the excitation was performed at 488 nm and the emission spectra were recorded in the wavelength range 500~700 nm. 2.8 Cellular Uptake Cells (Raw 264.7, Tramp-C1 or NIH T3T cells) were seeded to each well of a 6-well plate (2.0×105 cells/well). Nanoparticles loaded with DiO by hydrophobic association were used for fluorescence detection. Cells were co-incubated with NP-I, NP-II, acid-pretreated NP-II and NP-III, respectively, at an SN38 concentration of 10.0 µM at 37 oC for 15 min. With being washed with phosphate buffered saline (PBS) twice, cells were detached by trypsin and dispersed in PBS. Cellular uptake was analyzed on a FACSCalibur flow cytometer (BD Biosciences). At least 10,000 events (cells) were analyzed for each sample. Note that the acidic pretreatment to induce the detachment of mPEG-b-PHMA from NP-II surfaces was carried out by suspending NP-II in succinate buffer of pH 6.5 for 24 h prior to the co-incubation with cells. The cellular uptake of NP-II by Raw 264.7, Tramp-C1 and NIH T3T cells was performed, respectively, in comparison with acid-pretreated NP-II in order to elucidate the effect of HA on mediating


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cellular internalization of NPs by CD44-expressed tumor cells; i.e., Raw 264.7 and Tramp-C1 cells adopted in this work. 2.9 Cytotoxicity Analysis Raw 264.7 and Tramp-C1 cells were seeded in a 96-well culture plate at a density of 1×104 cells/well in RPMI 1640 and DMEM, respectively, containing 10% FBS and 1% (100 units/mL) penicillin and incubated at 37 °C for 24 h. The medium was then replaced with 200 µL of culture medium containing either free SN38, pristine nanoparticles or NP-I with varying SN38 concentrations. With the cells being washed twice with PBS to remove nanoparticles or free drug that was not internalized, the MTT solution (10 µL, 5.0 mg/mL) was added into each well at each preset time point (24, 48 and 72 h post incubation). The plates were further incubated for 4 h at 37 °C. The culture medium was then replaced with DMSO (100 µL), and the absorbance of the resulting solution in each well at 570 nm determined using a SpectraMax M5 microplate reader. 2.10 In Vitro Intercellular Drug Delivery Study Raw 264.7 cells were added to each well of a 6-well plate (2.5×105 cells/well). NP-I was labeled

with

1,1'-dioctadecyl-3,3,3'3'-tetramethylindocarbocyanine

perchlorate

(DiI)

by

hydrophobic association for fluorescence detection prior to their cellular uptake. Instead of DiO, DiI was used as a probe in order to prevent the overlap in fluorescence signal with green fluorescence protein (GFP) tagged on Tramp-C1 as described below. After 24 h pre-incubation, cells were co-incubated with NP-I at an SN38 concentration of 10.0 µM at 37 oC for 4 h. After being washed with PBS, the cargo-loaded cells were seeded onto 22 mm round glass coverslips containing 2.5×105 GFP-tagged Tramp-C1 cells in RPMI medium and further co-incubated for 48 h. With being washed twice with PBS, cells were fixed with 4% formaldehyde and directly visualized with a fluorescence microscope (Olympus IX70).


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For the quantitative evaluation of cytotoxicity, the NP-I engulfed Raw 264.7 cells were placed into the insert (PET membrane with 1.0 µm pore size, BD Falcon) of the transwell system. The insert was then accommodated within the well of 6-well plate containing GFPtagged Tramp-C1 cells (2.5×105 cells/well) in RPMI medium and further co-incubated for 24, 48 and 72 h, respectively. Cancer cells were detached at the preset time by trypsin and then dispersed in PBS. The in vitro cell viability was evaluated by flow cytometry. 2.11 Biodistribution and Tumor Growth Inhibition Tumors were produced by subcutaneous inoculation of 3 × 106 viable Tramp-C1 cells into the right hind legs of C57BL/6JNarl mice. Tumors with the size of ca 4 mm in diameter were selected and randomly allocated into 5 groups (5 mice per group) for the in vivo study. Tumor-bearing mice were injected intravenously with different NP formulations, respectively, at a daily SN38 dosage of 5.0 mg/kg for a total of three doses. Due to the limited solubility of free SN38 in aqueous phase, a water-soluble SN38 prodrug, irinotecan (hydrochloride) was used to serve as the free drug at an equivalent molecular dosage. Tumor volume was measured with calipers and calculated using the formula (a×b2)/2 where a is the longest and b the shortest orthogonal tumor diameter. The mice were scarified and the tumors harvested 72 h post therapeutic treatment. Differences in the postmortem tumor weights from various therapeutic groups were compared using an unpaired two-sample Student’s t-test. Nanoparticles were stained with NIR fluorophore, Cy5.5, for IVIS fluorescence detection. Tumor-bearing mice were intravenously injected with different formulations at day 1 with an equivalent SN38 dose of 5.0 mg/kg. Mice were imaged by IVIS (Cy5.5 ﬁlter: λex = 675 nm; λem = 720 nm) 72 h post injection under anesthesia via inhalation of oxygen-isoﬂurane. Heart, liver, spleen, kidney and tumors were then collected for ex vivo optical imaging. Gray-


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scale photographic and fluorescence images of each sample were analyzed and overlaid using Living Image® 3.1 software (Xenogen, Alameda, CA). The regions of interest (ROI) were drawn over the signals and the average radiant efficiency was quantified in p/s/cm2/sr. 2.12 H&E examination and IHC Analysis For H&E histologic examination, the tumor-bearing mice were treated with NP-I, NP-II, NP-III and drug-free NP-II, respectively, via tail vein injection with the same doses as described above. The major organs, including heart, liver, spleen, lung and kidney, and tumors were harvested at 2 day post the third NP treatment. The organs and tumors were embedded in OCT compound (Sakura Finetek, Torrance, CA, USA) and stored at -80 °C for further H&E staining. The tissue sections (10 µm) were mounted onto slides and the tissue slides were then fixed with pre-cooled methanol for 5 min and washed twice with PBS. Tissue sections were then stained with H&E and examined under optical microscopy. As for IHC staining of tumor hypoxia, mice received PIMO (pimonidazole hydrochloride, 60 mg/kg) suspended in PBS (10 mg/mL; Hypoxyprobet-1 Kit, Chemicon, Billerica, MA, USA) via intraperitoneal injection 1 h prior to tumor harvest. To prevent non-specific binding of antibodies, tissue slides were blocked with the blocking buffer containing 4% FBS, 1% normal goat serum and 0.01% Tween-20 in PBS for 1 h at room temperature. The slides were subsequently incubated with anti-PIMO, rat anti-mouse CD11b, CD31, and rabbit anti-mouse caspase-3 antibodies (BD Pharmingen) at 4 °C. After 24 h reaction, slides were washed and incubated with secondary antibodies conjugated with Alexa Fluor 488 (Invitrogen) for 1 h at ambient temperature. After being washed thoroughly with PBS, slides were stained with DAPI (5.0 µg/mL; Invitrogen) to identify the loci of cell nuclei. Images were captured with the Olympus IX70 and Pursuit USB Slider microscopes. 3. RESULTS AND DISCUSSION


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In order to enhance selective cellular uptake and attain deep penetration of nanotherapeutics in tumors, the surface detachment of mPEG-b-PHMA from HA/PLGA-based particles in response to tumor acidic microenvironment and the exposure of HA shells that allows the active targeting of NPs to CD44 presenting tumor cells, including both cancer cells and TAMs, is the core design of this work. The intratumoral deep penetration of nanotherapeutics is thus achieved by means of the hitchhiking transport while the loaded TAMs being further chemotactically recruited towards hypoxia regions. The capability of mPEG-b-PHMA as a major component deposited on the NP outer surfaces to undergo the hydrophobic/hydrophilic transition in aqueous phase in response to pH change was first characterized by the fluorescence technique using pyrene as a non-polar probe.31,32 A dramatic change in the fluorescence intensity ratio (I3/I1) of pyrene at the vibronic bands of 383 and 373 nm in the aqueous copolymer solutions occurred mainly from pH 6.0 to 7.0 (Figure 1a), being the range as reported in solid tumor microenvironment.20,33 Since the I3/I1 ratio represents a quantitative measure of the nonpolarity nature of the microenvironment where most hydrophobic pyrene molecules reside, the abrupt decrease illustrated in Figure 1a is primarily a result of the phase transition of the mPEG-bPHMA from insoluble to soluble state due to the extensive protonation of the imidazole groups of the PHMA block with the external pH being reduced. Similar observations from the polyhistidine-based derivatives containing imidazole rings with a pKa of ca 6.0 were reported elsewhere.34,35 In agreement with the pyrene fluorescence data, the relative light scattering intensity determined by dynamic light scattering (DLS) was abruptly reduced as a consequence of the protonation of the PHMA block in virtually the same pH range (Figure 1b). These results strongly support the transition of the PHMA block from hydrophobic to hydrophilic state when exposed to the tumor acidic environment. This will then enable the detachment of mPEG-b-


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PHMA from NP surfaces and exposure of HA shells, allowing the selective cellular uptake and transport subsequently occurring via CD44-expressed tumor cells.

mPEG-b-PHMA

0.72

O

O

0.70

O 113

NC O

S

30

Blank

RhB

NH

0.68

I383/I373

mPEG-b-PHMA

HN

0.66

N

0.64

pH reduction pH reduction

0.62 O O 113

O

0.60

30

NC O

0.58 0.56

S

RhB

NH

H2N

(a)

0.54

N

3

4

5

6

pH

Relative laser light scattering intensity (I pH/ I pH 7.4)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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7

8

Tumor pH range

1.2 1.0 0.8 pH pHdecrease reduction

0.6 0.4 0.2

(b) 3

4

5

6

7

8

pH

Figure 1. (a) Fluorescence intensity ratio (I383/I373) of pyrene at the vibronic bands of 383 and 373 nm with the excitation being performed at 337 nm in aqueous solution of mPEG-b-PHMA as a function of pH. (b) pH-dependent relative light scattering intensity of aqueous mPEG-b-


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PHMA solution. The relative light scattering intensity at preset pH was attained by normalization to the mean light scattering intensity at pH 7.4. The drug-loaded NPs with a mean hydrodynamic diameter (Dh) of ca 175 nm in the absence of mPEG-b-PHMA decoration (NP-I) were prepared from assembly of the HA-g-PLGA copolymer with SN38 in the continuous aqueous phase of pH 7.4 by the nanoprecipitation technique.36 SN38 was adopted in this work for chemotherapy of tumor hypoxia because of its capability to overcome the chemoresistance of cancer cells under hypoxic conditions through the expression inhibition of a cell survival factor, hypoxia-inducible factor 1α.37 The NP-I was further elaborated with mPEG-b-PHMA to obtain NP-II through deposition of hydrophobic deprotonated PHMA chain segments on the particle surfaces in aqueous solution at pH 7.4. The data in Table 1 show that the NP-II have a Dh of ca 220 nm and a PDI of 0.16. The DLS size distribution profiles of the nanoparticles at pH 7.4 are also illustrated in Figure S3a. The further enlarged particle size, yet with a similar size distribution as compared to NP-I strongly suggests the successful decoration of the mPEG-b-PHMA outlayer on the NP-I surface without additional NP formation from the diblock copolymer alone. The TEM image shown in Figure S3b confirms the spherical shape of NP-II with a size of ca 200 nm. The slight size reduction of NPII observed by TEM as compared to the DLS measurement is ascribed to the dehydration of NPs under vacuum during TEM imaging. NP-II also shows an excellent colloidal stability in PBS upon aging at 37 oC over a period of at least 6 days by virtue of the PEG protection (Figure S3c). Owing to the nonpolar nature of SN38, a large quantity of drug species was loaded in the hydrophobic domain of NPs via hydrophobic association. With the loading efficiencies being the same at ca 87% for NP-I and NP-II, the drug loading contents are ca 16.3 and 14.5 wt%, respectively. The particle surface modification with mPEG-b-PHMA accounts for the slight


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reduction of the drug content in NP-II. The in vitro SN38 release performance of the NPs in aqueous solution was also evaluated. As shown in Figure S3d, the NP-II demonstrated a rather slow drug release profile under normal physiological conditions (37 oC, PBS) by dialysis, merely 18% SN38 unloading over a period of 48 h. By contrast, more than 80% of free drug serving as a control was eliminated from the dialysis tube under the same conditions. The sustained drug release from the NPs is presumably due to the pronounced hydrophobic association of SN38 with NP cores consisting of PLGA, thus promoting the drug retention and reducing the leakage from the NPs. This ensures a low cytotoxicity of therapeutic cargos against host cells while being cellularly transported via TAMs towards hypoxia. Based upon the inherent pH-inducible phase transition property of mPEG-b-PHMA in aqueous phase as shown in Figure 1, the detachment of surface mPEG-b-PHMA from NP-II in response to the decreased pH in external medium was further studied. Owing to the strong susceptibility of resonance energy transfer between spectrally complementary fluorophores (i.e., donor and acceptor) to spacious distance,38 the FRET effect on fluorescence intensity as a function of medium pH was evaluated herein to verify the pH-evolved shedding of the block copolymer from NP surface. By serving as a donor, the hydrophobic fluorescence probe, DiO, was incorporated into hydrophobic PLGA domains of NPs. In contrast, RhB acting as an acceptor was labeled at the end of the PHMA block anchored onto NPs via hydrophobic association at pH 7.4 at which the PHMA block remained mostly uncharged and thus hydrophobic in nature. As shown in Figure 2, a fluorescence signal of RhB appears concomitantly with the fluorescence quenching of DiO for NP-II at pH 7.4. This indicates the occurrence of FRET between donor and acceptor virtually induced by the spatially close contact with each other. By contrast, only the DiO fluorescence was detected when pH was reduced from 7.4 to 6.5. This implies that mPEG-b-PHMA desorbed out of NP-II surfaces as


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primarily a result of the increased protonation of the PHMA blocks (i.e., the enhanced hydrophilicity of the protonated PHMA block). Further evidences to support the pH-mediated detachment of mPEG-b-PHMA from NP-II in aqueous solution were provided by the DLS characterizations. For comparison, NP-I coated by a pH-insensitive mPEG-b-PLGA copolymer (NP-III) was also included in this study. The SN38 loading efficiency and loading content of NPIII are ca 90% and 13.8 wt%, respectively. The data of Dh, PDI and zeta potential (ζ) of various NPs at three different pH values are summarized in Table 1. With NP-II exhibiting similar Dh and ζ to NP-III at pH 7.4, both NP-II and NP-III showed larger particle size and lower ζ arising from the shift of the shear plane within the electric double layer away from the particle surface (termed the PEG shielding effect) as compared to NP-I at pH 7.4. More importantly, while both Dh and ζ of NP-III remained essentially unchanged when pH is decreased from 7.4 to 6.5 or even lower (5.5), the Dh of NP-II was reduced from 219 to ca 175 nm, which is quite comparable to that (~172 nm) of NP-I. It is also interesting to note that the values of ζ for NP-II are quite close to those of NP-I at pH 6.5 and 5.5 (Table 1), which is consistent with the FRET effect shown in Figure 2. All these factors provide supporting evidence for the capability of mPEG-b-PHMA to undergo pH-induced detachment from NP-II surfaces.


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Table 1. Effects of pH on particle size, polydispersity index (PDI) and ζ-potential of varying NPs in aqueous solution. pH 7.4

Sample IDa

NP-I

pH 6.5

pH 5.5

Dh (nm)

PDI

ζ-potential (mV)

Dh (nm)

PDI

ζ-potential (mV)

Dh (nm)

PDI

ζ-potential (mV)

175.2±1.0

0.13

-37±0.9

172.3±4.6

0.15

-33.8±0.9

172.7±0.5

0.16

-28±2.1

219.6±8.0

0.16

-27±1.0

179.5±1.7

0.18

-33.0±1.2

174.6±1.5

0.21

-29±1.2

222.1±4.0

0.16

-27.5±1.0

216.4±3.2

0.17

-27.3±0.9

211.4±1.8

0.18

-26.2±1.4

NP-II

NP-III

a

NP-I: NPs without PEG coating; NP-II: NPs coated with pH-sensitive mPEG-b-PHMA copolymers; NP-III: NPs coated with pH-insensitive mPEG-b-PLGA copolymers.

NP-I (DiO stained) NP-II at pH 7.4 NP-II at pH 6.5 NP-II at pH 7.4 (without DiO stained)

1500

Fluorescence intensity (a.u.)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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1000

Without FRET reaction FRET reaction 500

(a) 0 500

550

600

650

700

Wavelength (nm)

Figure 2. Fluorescence emission spectra for NP-I and NP-II at 25 °C with excitation conducted at 480 nm for DiO. Occurrence of FRET reaction depends on the distance between DiO (λem 510 nm) and RhB (λem 565 nm).


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Figure 3. Representative flow cytometric histogram profiles of fluorescence intensity for Raw 264.7 cells (a) and Tramp-C1 cells (b), respectively, incubated with different DiO-stained NPs for 15 min at 37 oC. Schematic of selective uptake of NP-II deprived of mPEG-b-PHMA in tumor acidic microenvironment (similar to NP-I) by CD44-presenting tumor cells (cancer cells and TAMs) is also included.

To gain a better insight into the cellular uptake of the NPs, the flow cytometry measurements were conducted, using the murine macrophage-like Raw 264.7 cell line as a TAM


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model for evaluating phagocytic internalization in vitro. The NPs were loaded with a green fluorophore, DiO (1.0 wt%), prior to the reaction in order to facilitate the fluorescence detection by flow cytometry. As shown in Figure 3a, the greatly enhanced cellular uptake of NP-I and NPII after mPEG-b-PHMA shedding under acidic conditions was evidenced by a dramatic increase in the signal of intracellular DiO fluorescence. By contrast, the cellular pickup of PEGylated NPII and NP-III was substantially reduced. The enhanced NP uptake is primarily attributed to the HA/CD44-mediated endocytosis (phagocytosis) in the absence of steric repulsion provided by mPEG-b-PHMA physically attached onto NP surfaces.39,40 Similar enhanced uptake of NPs via HA/CD44-mediated endocytosis was also observed on Tramp-C1 cells, a murine prostate cancer cell line characterized to be highly metastatic.36 The NP-II with the mild acidic treatment to detach the surface PEG adducts were internalized significantly higher than NP-II and III by the malignant prostate cancer cells (Figure 3b). The Tramp C1 cancer cells were known to express CD44 molecules on cell membranes, thereby being capable of undergoing CD44-mediated endocytosis of HA outlayered NPs, in particular for those without PEG intervention. On the other hand, when NIH 3T3 fibroblast cells were adopted to serve as a negative control due to the absence of CD44 expression,41 the internalization of NP-I and acid-pretreated NP-II by NIH 3T3 was reduced as compared to Tramp-C1 cells (Figure S4). The PEG intervention effect on cellular uptake of NP-II and NP-III by the fibroblast cells was also observed. The difference in NP uptake efficiency between macrophages and cancer cells was also noted. With the much enhanced NP uptake by macrophage compared to Tramp-C1, this demonstrates the great potential of the TAM-involved hitchhiking strategy by taking advantage of those cells belonging to phagocytes while overexpressing CD44 molecules as the HA receptor. The cytotoxicity of free SN38 and the NP-I (mimicking the scenario that mPEG-b-


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PHMA detaches from NP-II surfaces in tumor microenvironment) against either Raw 264.7 or Tramp-C1 cells was evaluated using MTT assay. As presented in Figure S5, the viability of Raw 264.7 or Tramp-C1 cells when exposed to NP-I was reduced proportionally with the dose of SN38, while the survival rates of cells incubated with drug-free NPs still remained high. This result implies that the pristine NP-I is virtually non-toxic. The cytotoxic effect on both cells was essentially induced by chemotherapy of SN38 that was released slowly from NPs upon PLGA degradation. From Figure S5, it seemed as though Raw 264.7 cells were more sensitive to SN38 than Tramp-C1 cells. This is because of the enhanced uptake of the chemotherapy in either free drug or NP form by the macrophages compared to the cancer cells during the co-incubation reaction. Since the in vitro therapeutic efficacy of the NPs against tumor cells including macrophages and cancer cells was demonstrated, it is important to further validate their therapeutic potential on cancer cells via cellular hitchhiking. To evaluate the intercellular drug transport from macrophages to cancer cells and its cytotoxic activity against cancer cells, two different approaches, i.e. the in vitro mixed co-culture and transwell insert systems illustrated in Figure 4 were adopted. For the mixed co-incubation system, the NP-I engulfed Raw 264.7 cells were incubated together with Tramp-C1 cells genetically engineered to express GFP. The fluorescence micrographs shown in Figure 4a confirm considerable overlaps in fluorescence signals from DiI-stained NPs and GFP-tagged Tramp-C1 at 48 h post co-incubation in contrast to the appearance of the DiI signals exclusively within Raw 264.7 cells at the onset of the reaction. Along with intercellular transport of nanomedicine, apoptosis of Tramp-C1 cells was also observed, as shown by typical apoptotic morphological features including shrinking and blebbing.42 In agreement with the fluorescence microscopic observation, the cytotoxic effect of therapeutics from intercellular transport against cancer cells using the transwell insert system by


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flow cytometric analyses was found dramatically enhanced when the incubation time was increased from 24 to 72 h (Figure 4b). These data clearly demonstrate the chemotherapeutic potential of the NPs against cancer cells in vitro by means of the drug release from host cells through PLGA degradation and subsequent diffusion into the cancer cells nearby.

Figure 4. (a) Fluorescence microscopic images showing the locations of NP-I (similar to NP-II deprived of mPEG-b-PHMA in tumor acidic microenvironment) in cell mixtures of Raw 264.7 and GFP-tagged Tramp-C1 cells at the beginning (0 h) and 48 h post incubation, respectively. (b) Schematic illustration of experimental design confirming the intercellular drug transport and the viability of Tramp-C1 cells co-incubated with NP-I engulfed Raw 264.7 cells for 24, 48 and 72 h, respectively. Prior to cell co-incubation, NP-I was reacted with Raw 264.7 cells for 4 h at an SN38 concentration of 10.0 µM. The viability of Tramp-C1 cells was determined by flow cytometry.


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The in vivo performance of NPs in terms of tumor accumulation, intratumoral distribution and antitumor efficacy was further investigated. The in vivo and ex vivo tumor accumulation of NPs stained with NIR fluorophore, Cy5.5, for fluorescence imaging prior to intravenous injection in Tramp-C1 murine prostate tumor-bearing mice (C57BL/6J) is illustrated in Figure S6. The PEGylated NP-II and NP-III show significant EPR effect, as reflected by the much stronger tumor accumulation in virtue of the greatly retarded uptake in liver and spleen as compared to NP-I in the absence of surface PEG chains.2,43 Furthermore, the pH-induced detachment of mPEG-b-PHMA from NP-II surfaces, which results in the enhanced uptake of naked NP-II by tumor cells, accounts for its higher tumor targeting than that of NP-III.44,45 This is further confirmed by the intratumoral NP distribution study exploiting IHC staining of tumor sections along with the intense red fluorescence of DiI stained on NPs. The TAMs in tumor tissue sections were identified with the CD11b marker (in green), while DAPI (in blue) was used for nucleus identification. Figure 5a shows only few NP-I reaching the tumor site due to the considerable liver uptake as aforementioned. The pH-insensitive NP-III was barely found at those loci where tumor macrophages were largely identified, implying the reduced TAM pickup by effective PEGylated protection.46 This is consistent with the flow cytometric measurements of NP uptake by macrophages as shown in Figure 3a. Distinct from the intratumoral cellular uptake of NP-III, the loci showing signals of NPs (in red) were appreciably overlapped with the signals of macrophage (in green) for the NP-II group. This indicates the effective hitchhiking of naked NP-II by TAMs for further transport toward deep tumor tissue regions (Figure 5b). With tumor sections being further stained with an angiogenic CD31 marker, the merged panel validates the successful deep NP transport by showing NP-II signals at the loci 100 µm distant from the nearest blood vessels, which is beyond the characteristic distance of ca 70 µm for hypoxic


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regions in human solid tumors.47,48 In conventional PEG-elaborated NP delivery designs (e.g., NP-III in this work), NPs were usually observed in the region within solid tumor rather close to blood vessels.49 Although NP-I exhibited the HA-rich shell capable of facilitating their uptake by TAMs, such an effect was severely limited by the poor accumulation of NPs in tumor. A significant amount of NP-I were captured by RES as evidenced by the ex vivo images showing significant NP accumulation in live and spleen (Figure S6b). With the hypoxic and apoptotic cells being specified by the pimonidazole (PIMO) and caspase 3 markers, respectively, in the contiguous tumor sections, Figure 5c shows the therapeutic efficacies of various formulations in tumor hypoxia. The best efficacy was observed for the NP-II group, as evidenced by the significant co-localization of the NP signal (in red) with the caspase-3 marker signal (in green) in PIMO+ regions. By contrast, both NP-I and NP-III were rarely found in these avascular (PIMO+) areas. As a consequence, very few tumor cells underwent the chemotherapy-induced apoptosis in these regions. To evaluate the in vivo therapeutic efficacy of this approach, three doses with SN38 5 mg/kg each dose on day 1, 2 and 3 were given intravenously into tumorbearing mice. A water-soluble SN38 prodrug, irinotecan, was employed as a control at an equivalent molecular dosage. Among different formulations, NP-II demonstrates the most remarkable inhibition in tumor growth in terms of either tumor volume or mass (Figure S7). The antitumor efficacy from the free irinotecan group was appreciably reduced compared to other NP groups, most likely because of the limited accumulation of free drug at tumor site due to the nonspecific cellular uptake and the rapid clearance of drug species.50 In addition to the reduced liver/spleen uptake for both NP-II and NP-III (Figure S6), the more pronounced antitumor efficacy of NP-II is ascribed to the appreciably enhanced cellular uptake and transport within tumor by virtue of the detachment of mPEG-b-PHMA in tumor acidic microenvironment and


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exposure of HA shells on NP surfaces which in part further promotes the cellular pickup for deep tumor tissue penetration. The tumor regrowth was observed for the mice treated with NP-II at day 8 primarily because of a relatively low dosage of SN38 (5 mg/kg) used in this study in order for the efficacy comparison with other formulations. With mice receiving various therapeutic treatments, the H&E examination indicated no apparent abnormality in major organs, most likely also due to the relatively low drug dosage at 5 mg/kg (Figure S8a). Nevertheless, the H&E stained tumor sections from the group treated with NP-II showed enhanced cell death while the dead cell areas were appreciably reduced for the NP-III group (Figure S8b). Cell death was barely observed in the tumor sections from the NP-I and drug-free NP-II groups. The reduced tumor accumulation of NP-I as shown in Figure S6 is apparently responsible for the negligible tumor cell death found in H&E histologic examination. Based on the results of the biodistribution, the tumor growth inhibition and the IHC and H&E tissue examination, it strongly suggests that the delivery strategy developed herein demonstrates prominent accumulation of chemotherapy in tumor, particularly in tumor hypoxia and promising antitumor efficacy.


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Figure 5. Multiple staining of tumor sections from Tramp-C1 tumor-bearing mice 72 h post intravenous injections (the daily SN38 dosage of 5.0 mg/kg for three consecutive days) to demonstrate differences among those NP delivery systems. Intratumoral macrophages, angiogenic blood vessels, hypoxia and apoptotic cells were identified by CD11b marker (in green in a), CD31 marker (in green in b), PIMO marker (in green in c, upper) and caspase 3


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marker (in green in c, lower), respectively. NPs were labeled with DiI in red and cell nuclei stained with DAPI in blue. Scale bar: 50 µm.

4. CONCLUSIONS In this study, a tumor microenvironment-responsive NP-based therapeutics system capable of enhancing selective uptake by tumor cells through HA/CD44-mediated endocytic/phagocytic pathway and hitchhiking via TAM into deep tumor tissues for improving chemotherapeutic efficacy was developed. The HA-shelled NP decorated with a surface layer of mPEG-b-PHMA capable of undressing in tumor acidic microenvironment endows its selective uptake by CD44-expressed tumor cells and TAM hitchhiking for further cellular transport of nanotherapeutics into deep tumor regions. The IHC examination of tumor sections proved the successful delivery of chemotherapeutics and, thus, pronounced apoptosis in tumors, particularly in tumor avascular regions.

ASSOCIATED CONTENT Supporting Information. The Supporting Information is available free of charge on the ACS Publications website at DOI:


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Chemical structures, synthetic routes and 1H-NMR spectra of HA-g-PLGA (Fig. S1), mPEG-bPMAA and mPEG-b-PHMA (Fig. S2), TEM images of nanoparticles, colloidal stability of NP-II and NP-III in PBS and drug release profiles for NP-II and NP-III in PBS (Fig. S3), cellular uptake of NPs by NIH 3T3 (Fig. S4) viability of Tramp-C1 and Raw 264.7 cells treated with free drug, drug-free and drug-loaded NPs (Fig. S5), in vivo NIR fluorescence images of tumorbearing mice (C57BL/6JNarl) and ex vivo fluorescence images of major organs and tumors (Fig. S6), tumor growth curves of Tramp-C1 tumor-bearing mice and tumor images (Fig. S7).

AUTHOR INFORMATION Corresponding Author *E-mail: [email protected]. ACKNOWLEDGMENT This work is supported by the Ministry of Science and Technology, Taiwan (102-2221-E-007032-MY3 and 104-2627-M-007-009), the National Tsing Hua University, Taiwan (104N2742E1 and 105N522CE1) and National Taiwan University Hospital-Hsinchu Branch (HCH103-009). REFERENCES (1)

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