Precise Targeting of Liver Tumor Using Glycol Chitosan Nanoparticles

Article pubs.acs.org/molecularpharmaceutics

Precise Targeting of Liver Tumor Using Glycol Chitosan Nanoparticles: Mechanisms, Key Factors, and Their Implications Jin Hee Na,†,‡,§,△ Heebeom Koo,∥,△ Sangmin Lee,‡,§ Seung Jin Han,† Kyung Eun Lee,⊥ Sunjin Kim,# Haeshin Lee,# Seulki Lee,‡,§ Kuiwon Choi,¶ Ick Chan Kwon,†,$ and Kwangmeyung Kim*,† †

Center for Theragnosis, Biomedical Research Institute, Korea Institute of Science and Technology (KIST), Hwarangno 14-gil 5, Seongbuk-gu, Seoul 136-791, Republic of Korea ‡ The Russell H. Morgan Department of Radiology and Radiological Science, Johns Hopkins University School of Medicine, 601 North Caroline Street, Baltimore, Maryland 21287, United States § The Center for Nanomedicine, The Wilmer Eye Institute, Johns Hopkins University School of Medicine, 400 North Broadway, Baltimore, Maryland 21231, United States ∥ Department of Medical Lifescience, College of Medicine, The Catholic University of Korea, 222 Banpo-daero, Seocho-gu, Seoul 06591, Republic of Korea ⊥ Advanced Analysis Center, Korea Institute of Science and Technology (KIST), Hwarangno 14-gil 5, Seongbuk-gu, Seoul 136-791, Republic of Korea # Department of Chemistry and Institute for NanoCentury and BioCentury, Korea Advanced Institute of Science and Technology (KAIST), 291 Daehak-ro, Yuseong-gu, Daejeon, 305-701, Republic of Korea ¶ Korea Institute of Science and Technology Europe (KIST-Europe) Forschungsgesellschaft mbH, Campus E7.1, 66123 Saarbrücken, Germany $ KU-KIST School, Korea University, 1 Anam-dong, Seongbuk-gu, Seoul 136-701, Republic of Korea S Supporting Information *

ABSTRACT: Herein, we elucidated the mechanisms and key factors for the tumor-targeting ability of nanoparticles that presented high targeting efficiency for liver tumor. We used several different nanoparticles with sizes of 200−300 nm, including liposome nanoparticles (LNPs), polystyrene nanoparticles (PNPs) and glycol chitosan-5β-cholanic acid nanoparticles (CNPs). Their sizes are suitable for the enhanced permeation and retention (EPR) effect in literature. Different in vitro characteristics, such as the particle structure, stability, and bioinertness, were carefully analyzed with and without serum proteins. Also, pH-dependent tumor cell uptakes of nanoparticles were studied using fluorescence microscopy. Importantly, CNPs had sufficient stability and bioinertness to maintain their nanoparticle structure in the bloodstream, and they also presented prolonged circulation time in the body (blood circulation halflife T1/2 = about 12.2 h), compared to the control nanoparticles. Finally, employing liver tumor bearing mice, we also observed that CNPs had excellent liver tumor targeting ability in vivo, while LNPs and PNPs demonstrated lower tumor-targeting efficiency due to the nonspecific accumulation in normal liver tissue. Liver tumor models were produced by laparotomy and direct injection of HT29 tumor cells into the left lobe of the liver of athymic nude mice. This study provides valuable information concerning the key factors for the tumor-targeting ability of nanoparticles such as stability, bioinertness, and rapid cellular uptake at targeted tumor tissues. KEYWORDS: tumor targeting, nanoparticles, stability, liver tumor, biodistribution

Received: June 7, 2016 Revised: August 28, 2016 Accepted: September 21, 2016

1. INTRODUCTION Recent advances in nanotechnology and biotechnology have led to a flurry of nanoparticle research for biomedical applications.1−3 © XXXX American Chemical Society

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DOI: 10.1021/acs.molpharmaceut.6b00507 Mol. Pharmaceutics XXXX, XXX, XXX−XXX

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Molecular Pharmaceutics

efficiency in vivo in many studies.6,19−21 Different in vitro characteristics, such as the particle structure, stability, and bioinertness, were carefully analyzed in the presence of serum protein. Also, pH-dependent tumor cell uptake in the tumoral acid microenvironment was studied using fluorescence microscopy. Finally, the in vivo biodistribution and tumor targeting efficiency of various nanoparticles was studied using noninvasive fluorescence imaging technology that could evaluate the relationship between their prolonged circulation properties and the ERP effect in liver tumor bearing mice.

In particular, many researchers have developed various nanoparticles for tumor-targeted imaging and drug delivery. A large amount of promising data has been published, and some of these nanoparticles have been commercialized in clinics.4,5 Also, numerous studies have reported that nanoparticles could easily penetrate angiogenic tumor vessels during circulation, which can result in high accumulation in tumor tissue by the enhanced permeation and retention (EPR) effect.6−9 To explain the EPR effect, researchers have primarily focused on the size of particles as the most important key factor for tumor-targeting ability of nanoparticles. The nanoparticles with the size of 10−500 nm are well-known to pass through fenestrated vessels, leading to efficient accumulation in tumor tissues.8 In the animal model, highly permeable tumor vessels could allow nanoparticles to localize at the tumor interstitial region, but larger particles are prevented from entering into the smaller angiogenic tumor vessels. However, the size of nanoparticles could not fully explain the full mechanism of the EPR effect since nanoparticles with similar particle size often showed different tumor-targeting efficiency in various tumor models. Indeed, many nanoparticles intended for tumor-targeting delivery system have proven unsatisfactory during in vivo tests, due to the nonspecific targeting of nanoparticles to normal tissues.10 Furthermore, despite the EPR effect, the final number of nanoparticles eventually delivered to tumor tissue does not meet the criteria for clinical applications.11 Besides the size of nanoparticles, surface chemistry, stability, and bioinertness are also known to be additional key factors that might determine the long-circulating half-life of nanoparticles and the EPR effect of nanoparticles.12−14 The key factors that determine the tumor-targeting ability of different nanoparticles have not been fully elucidated under in vivo conditions, due to the difficulty of directly analyzing the characteristics of different nanoparticles after intravenous injection. Typically, nanoparticles are simply assayed in water or PBS without serum proteins, but these aqueous conditions do not fully explain the physicochemical properties of nanoparticles, such as size and surface chemistry, or recapitulate the complex in vivo environment in which they are used. It has been reported that nanoparticle properties can be dramatically changed by nonspecific protein adsorption in the bloodstream, which can lead to dissociation or aggregation of nanoparticles.15,16 For example, as size increases beyond 150 nm, more and more nanoparticles rapidly bind to serum proteins in blood, in a process known as opsonization.17 These opsonized nanoparticles are then readily removed by the reticuloendothelial system (RES) of liver and spleen. It has been reported that macrophages captured the opsonized nanoparticles in the bloodstream within several seconds after intravenous injection, inhibiting the long circulation needed for targeted drug delivery.18 Ideal nanoparticles should minimize nonspecific protein adsorption and escape immune system uptake in vivo. As a result, there is strong demand for research that evaluates in vivo characteristics of nanoparticles, such as stability in the blood, particle−protein interactions, and in vivo biodistribution, all of which can determine the in vivo tumor-targeting efficacy of different nanoparticles for drug delivery systems and molecular imaging. Herein, various key factors that determine the tumor-targeting efficiency of nanoparticles were analyzed both in vitro and in vivo. Control nanoparticles such as 1,2-dipalmitoyl-sn-glycero-3phosphocholine liposomes (LNPs) and polystyrene nanoparticle (PNPs) were compared to glycol chitosan-5β-cholanic acid nanoparticles (CNPs), which showed excellent tumor-targeting

2. EXPERIMENTAL SECTION 2.1. Materials. Glycol chitosan (Mw = 2.5 × 105 kDa; degree of deacetylation = 82.7%), 5β-cholanic acid (99% purity), 1-ethyl-3-(3-(dimethylamino)propyl)carbodiimide hydrochloride (EDC, 99% purity), N-hydroxysuccinimide (NHS, 98% purity), 4-(dimethylamino)pyridine (DMAP, 99% purity), and 4-methylmorpholine (NMM, 99% purity) were purchased from Sigma (St. Louis, MO). The hydroxysuccinimide ester form of Cy5.5 was purchased from Amersham Biosciences (Piscataway, NJ). Methanol (99.9% purity), chloroform (99.8% purity), anhydrous dimethyl sulfoxide (DMSO, 99% purity), and hexane (99% purity) were purchased from Merck (Darmstadt, Germany). Polystyrene nanoparticles (Polybead Amino Microspheres 0.2 μm) was purchased from Polysciences, Inc. (Warrington, PA). 1,2-Dipalmitoyl-sn-glycero-3-phosphocholine (PC, 99.9% purity) and 1,2-dipalmitoyl-sn-glycero-3-phosphoethanolamine (PE, 99.9% purity) were purchased from Avanti Polar Lipids (Alabaster, AL). D-Luciferin firefly potassium salt (98% purity) was from Caliper Life Sciences (MA, USA). All other chemicals were purchased and used as analytical grade. 2.2. Preparation of Glycol Chitosan-5β-Cholanic Acid Nanoparticles (CNPs). The molecular weight of glycol chitosan was determined by GPC (gel permeation chromatography) on PSS Hema-Bio 40 columns (300 × 8 mm, particle size 10 mm, exclusion limit for dextran 3,000,000) and PSS Hema-Bio 300 (300 × 8 mm, particle size 10 μm, exclusion limit for dextran 500,000).22 Glycol chitosan-5β-cholanic acid nanoparticles (CNPs) were prepared by the self-assembly of amphiphilic glycol chitosan conjugates as in the previous papers.19 First, the water-soluble glycol chitosan polymer (250 kDa) was chemically modified with 5β-cholanic acid in the presence of NHS and EDC. Briefly, glycol chitosan (0.5 g) was dissolved in 60 mL of deionized water, and it was added to 60 mL of methanol. Then, this solution was mixed with 120 mL of methanol containing 150 mg of 5β-cholanic acid in the presence of NHS and EDC (1.5 equiv of 5β-cholanic acid). After stirring at room temperature for 1 day, it was purified via extensive dialysis (against 20 volumes of reaction buffer) for 3 days in distilled water/methanol mixture (1:3, (v/v)) using a cellulose membrane (molecular weight cutoff = 12−14 kDa, Spectrum Laboratories, Laguna Hills, CA, USA). The solution was further dialyzed for 2 days in deionized distilled water and lyophilized to obtain glycol chitosan−5β-cholanic acid conjugates. The degree of substitution (DS), defined as the number of 5β-cholanic acid groups per one glycol chitosan polymer, was determined using a colloidal titration method.23 Briefly, 5 mg of glycol chitosan−5β-cholanic acid conjugate was dissolved in 10 mL of 2% acetic acid/water solution. After adding 0.1 w/v % toluidine blue (20 μL) as indicator, the solution was titrated using N/400 potassium polyvinyl sulfate solution. Next, glycol chitosan−5β-cholanic acid conjugates were labeled with the NIRF dye, Cy5.5, for efficient tracking of nanoparticles. Hydroxysuccinimide ester of Cy5.5 (2 mg) was B


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voltage mode (120 V). CCD camera was used to obtain NIRF images of the gel. Coomassie Brilliant Blue (0.1% w/v in 10% acetic acid solution) was used to stain rat serum. Cy5.5-GC, Cy5.5-CNPs, Cy5.5-PNPs, and Cy5.5-LNPs (1 mg/mL) in PBS (pH 7.4) containing rat serum (15% v/v) were incubated for 24 h at 37 °C. Then, NIRF intensities of each sample were adjusted to the same level using the values measured by the 12-bit CCD camera equipped with a special C-mount lens and a Cy5.5 bandpass emission filter set (680 to 720 nm; Omega Optical). Each solution was filtered by syringe filter membranes (cellulose acetate, Millipore) with decreasing pore sizes of 0.8, 0.45, and 0.2 μm, and the NIRF intensity of the filtered solution was determined by CCD camera. 2.4. Macrophage or Tumor Cell Uptake of Nanoparticles in Vitro. RAW264.7 (murine macrophage) cells and HT29 (human colorectal adenocarcinoma) cells were purchased from ATCC (Manassas, VA, USA). All cultures were mycoplasma free and maintained in RPMI1640 (Welgene, Daegu, Korea), 100 μg/mL streptomycin, and 100 U/mL penicillin (Welgene, Daegu, Korea) in humidified 5% CO2, atmosphere at 37 °C. RAW264.7 cells (passage number 26−30) or HT29 (passage number 55−60) (5 × 104 cells/dish) were grown on 35 mm cover-glass bottom dishes. After 1 day, the medium was replaced by 1 mL of serum-free medium at pH 5.5, pH 6.5, pH 7.4, and pH 8.0 and Cy5.5-CNPs or Cy5.5-LNPs (50 μg/mL) were added. After 2 h, the culture dishes were washed three times with PBS (pH 7.4) and fixed with 4% (w/v) formaldehyde in PBS (pH 7.4) at 4 °C for 30 min. The cellular uptake of Cy5.5-CNPs at pH 5.5, pH 6.5, pH 7.4, and pH 8.0 was visualized with a FluoView FV1000 confocal laser scanning microscope (Olympus, Tokyo, Japan) equipped with Cy5.5 (675 nm) and DAPI (358 nm) lasers. Also, nanoparticle-treated cells were detached with DPBS containing 2% FBS (FACS buffer) and washed twice. We analyzed 20,000 cells per each sample using flow cytometry (BD FACSVerse, BD Biosciences, San Jose, CA, USA) and analyzed the subsequent data using FlowJo software. 2.5. Analyzing Blood Circulation Time of Nanoparticles. All experiments with live animals were performed in compliance with the relevant laws and institutional guidelines of Korea Institute of Science and Technology (KIST), and institutional committees have approved the experiments. To analyze the amount of nanoparticles in the blood, we drew 1 mL of blood samples from five-week-old male athymic nude mice (n = 5) at different time points. After centrifugation at 3000 rpm for 15 min, NIRF intensity graph and images of the supernatant fractions were obtained using a 12-bit CCD camera. 2.6. Liver Tumor Model and in Vivo/ex Vivo Imaging. Liver tumor models were prepared using athymic nude mice (Crl:NU(NCr)-Foxn1nu, BALB/c, Orient Bio, Inc.) via direct injecting 3 × 105 HT29 cells into the left lobe of the liver. After further growing for 7 days, Cy5.5-GC, Cy5.5-CNPs, Cy5.5PNPs, and Cy5.5-LNPs (5 mg/kg, injection volume: 100 μL) were injected by tail vein (n = 5 per each group), and in vivo and ex vivo NIRF images were obtained by eXplore Optix System (Advanced Research Technologies Inc., Montreal, Canada) and KODAK imaging station, respectively. The NIRF intensities of all solutions were adjusted to the same values based on the data obtained by the spectrophotometer, wherein there was no quenching in every sample. NIRF signals were normalized with Cy5.5 standard solution, and signal changes were calculated by subtracting background signal (preinjection). The intensities were recorded as total photons per second per centimeter squared per steradian (p/s/cm2/sr). Optical tomographic images

conjugated to glycol chitosan−5β-cholanic acid conjugate (220 mg) in DMSO (80 mL) at 25 °C in the dark for 1 day. Unreacted Cy5.5 molecules were removed by dialysis (against 50 volumes of reaction buffer) for 1 day in DMSO, and for a further 2 days in deionized distilled water, and lyophilized. We also checked that there was no dye in the outer solution after 3 days using a UV/vis spectrophotometer showing that all unreacted dye was removed. The amount of Cy5.5 was calculated based on the extinction coefficient of Cy5.5 at 675 nm (2.5 × 105 M−1 cm−1) using a fluorescence spectrophotometer (Hitachi F-7000, Tokyo, Japan). Finally, Cy5.5-labeled glycol chitosan− 5β-cholanic acid conjugate in PBS (pH 7.4) was sonicated three times with a probe-type sonicator (Ultrasonic Processor, GEX600) for 2 min at 90 W. Liposomes (LNPs) were prepared by 5 wt % of Cy5.5-PE and 95 wt % of PC. In brief, the lipid mixtures were dissolved in chloroform/methanol (2:1) cosolvent and dried under nitrogen.24 PBS (pH 7.4) was added to the obtained film, and then sonicated three times using a probe-type sonifier (ultrasonic processor, Cole-Parmer Instrument Co., USA) at 90 W for 2 min. Next, the liposomes were passed through a 0.8 μm syringe filter membrane (cellulose acetate, Millipore). For labeling of polystyrene beads (PNPs), hydroxysuccinimide ester of Cy5.5 (2 mg) was conjugated to polystyrene nanoparticles (220 mg) in DMSO (80 mL) at 25 °C in the dark for 1 day. They were purified by washing with distilled water using a 10 kDa molecular weight cutoff Amicon Ultra centrifugal filter device (cellulose, Millipore) and concentrated. To label LNPs with Cy5.5, Cy5.5-labeled PE was synthesized by conjugating Cy5-NHS to PE similarly. Briefly, PE (3.5 mg, 5 μmol) in 3.5 mL of chloroform/methanol mixture (2:1, v/v) was incubated with Cy5-NHS (6 mg, 10 μmol) overnight at room temperature. The obtained Cy5-PE was precipitated in diethyl ether three times and lyophilized. Additionally, for both PNPs and LNPs, unreacted Cy5.5 molecules were similarly removed by dialysis (against 50 volumes of reaction buffer) for 2 days in deionized distilled water using a cellulose membrane (molecular weight cutoff = 12−14 kDa), and the product was lyophilized. After dialysis, outer solution was precisely checked using UV/vis spectrophotometer Lambda 7 (PerkinElmer, USA). There was no dye in the outer solution, showing that all dye molecules were stably conjugated to nanoparticles. The mean size and surface charge of CNPs, PNPs, and LNPs were analyzed in PBS (pH 7.4) by dynamic light scattering (DLS) at 25 °C and 633 nm. Zeta-potential values of nanoparticles were determined in PBS (pH 7.4) with a zeta-potential analyzer (Malvern Zetasizer Nano ZS, Malvern, U.K.). We evaluated the polydispersity factor, presented as μ2/Γ2, using the cumulant method.25 The morphology of nanoparticles (1 mg/mL in distilled water) was observed using a transmission electron microscope (TEM, CM 30 electron microscope, Philips). 2.3. In Vitro Stability Test and Filtration Test of Nanoparticles. The in vitro stability of CNPs in serum was evaluated by an SDS−polyacrylamide gel electrophoresis (PAGE) assay. Cy5.5-labeled GC (Cy5.5-GC), Cy5.5-CNPs, Cy5.5-PNPs, and Cy5.5-LNPs (0.25 mg/mL) were dispersed in PBS (pH 7.5), and incubated with or without rat serum (15% v/v); 15% rat serum was used for in vitro stability test of nanoparticles in physiological conditions in other papers.26,27 The NIRF intensities of all samples were adjusted to the same level using the values measured by fluorescence spectrophotometer (F-7000, Hitachi, Tokyo, Japan). After 1 h incubation, each solution was transferred onto a vertical slab gel in a Tris/glycine/SDS buffer, and it was run at a constant C


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Scheme 1. Schematic Illustration of in Vivo Tumor-Targeting Processes for Nanoparticlesa

were obtained using luciferase-expressing HT29 cells during liver tumor model fabrication. CNPs in the liver tumor sites were specifically visualized by the fluorescence emission of Cy5.5. We also observed that luminescence signals were specifically visualized in liver tumor sites. The fluorescence and luminescence of mapping was recorded using a CCD camera, implemented in a high-sensitivity imaging system (IVIS-spectrum, PerkinElmer, USA). For the ex vivo imaging, liver tissues were obtained from liver tumor mouse models. They were fixed in 4% (v/v) buffered formaldehyde solution at room temperature for 10 min and embedded in OCT tissue compound (Sakura, Tokyo). After freezing by a dry ice bath, they were cut on a cryostat (10 μm in thickness). The tissue sections were dried at 45 °C in dark conditions and fluorescence images were obtained using IX81-ZDC focus drift compensating microscope (Olympus, Tokyo, Japan). 2.7. CryoTEM and Correlative Light−Electronic Microscopy (CLEM) Analysis. For the CryoTEM assay, the dissected livers were fixed with 4% formaldehyde and 1% glutaraldehyde dissolved in 0.1 M cacodylate buffer overnight at 4 °C and with 2% OsO4 for 3 h. Then en bloc staining was performed with 2% uranyl acetate aqueous solution. The specimens were dehydrated with ethanol series, infiltrated with Spurr’s resin series, and polymerized at 60 °C overnight. The embedded tissues were cut with a diamond knife on an ultramicrotome (MTX-L, RMC). The sections were mounted on Formvar coated slot grids, and then they were stained with 2% uranyl acetate in 50% methanol for 20 min and Reynold’s lead citrate for 10 min. The grids were examined with TEM (Tecnai F20, FEI, Netherlands) at 200 kV. For CLEM, the dissected liver was fixed similarly and sectioned using a vibratome (Vibratome 3000 Plus, USA). Then, it was dehydrated and embedded in resin. The sections were observed by both confocal microscopy (Zeiss, Germany) and TEM (Tecnai F20, FEI, Netherlands).

a

(A) Stably maintaining their nanostructure without dissociation or aggregation during blood circulation after intravenous injection to body (stability). (B) Preventing the nonspecific uptake by the reticuloendothelial system (RES) of spleen and liver (bioinertness). (C) Passing through fenestrated vessels in tumor tissue via enhanced permeation and retention (EPR) effect and providing fast uptake into tumor cells (size, surface property).

3. RESULTS AND DISCUSSION 3.1. Preparation and Characterization of Nanoparticles in Aqueous Conditions. To maximize tumor targeting efficiency of nanoparticles at targeted tumor tissues: (1) Nanoparticles should stably maintain their nanostructure without dissociation or aggregation during blood circulation after intravenous injection (Scheme 1A). (2) They should also prevent nonspecific uptake by the reticuloendothelial system (RES) to minimize in vivo particle rejection in liver and spleen (Scheme 1B). (3) Finally, they should pass through the fenestrated vessels in tumor tissue via the EPR effect, and they are finally rapidly taken up by tumor cells (Scheme 1C). Therefore, we hypothesized that the tumor targeting nanoparticles should have excellent stability, minimal nonspecific uptake in normal tissue, enhanced EPR effect at targeted tumor tissues, fast uptake into tumor cells, etc. Furthermore, the key factors that determined the tumor targeting ability of different nanoparticles should be fully analyzed to explain how they can overcome all biological barriers, such as the stability in bloodstream, bioinertness against nonspecific uptake by in vivo RES, and adequate accumulation through the EPR effect at targeted tumor tissues, wherein the nanoparticles are being rapidly taken up by targeted tumor cells. As a potential tumor targeting particle, glycol chitosan-5βcholanic acid nanoparticles (CNPs) were synthesized and used in our experiments, due to the their higher tumor targeting efficiency in different tumor bearing mouse models.6,19−21,28 Briefly, hydrophilic glycol chitosan polymer (GC) (Mw = 250 kDa) was chemically modified with 159 ± 3.9 molecules of hydrophobic

5β-cholanic acid (CA) in the presence of chemical catalysts, resulting in amphiphilic GC−CA conjugates (Figure 1A), and the amphiphilic GC−CA conjugates could be self-assembled into very stable nanoparticle structure in aqueous conditions.19 For in vitro and in vivo fluorescent imaging analysis, we labeled the GC−CA conjugates with 1 wt % of near-infrared fluorescent (NIRF) dye, Cy5.5, and the Cy5.5-labeled GC-CA conjugates were finally self-assembled into glycol chitosan-5β-cholanic acid nanoparticles (CNPs) in aqueous conditions, due to their amphiphilic structure.29 The critical aggregation of concentration (CAC), the threshold concentration of self-aggregate formation, value of CNPs was 0.047 mg/mL.23,30 It means that CNPs could maintain their nanoparticle structure at the diluted low concentration in the body. As control nanoparticles, polystyrene nanoparticles (PNPs; polybead amino microspheres 200 nm, Polysciences, Inc.) and 1,2-dipalmitoyl-sn-glycero-3phosphocholine liposomes (LNPs) prepared by traditional film casting method were selected for in vitro and in vivo experiments.31−33 Both nanoparticles were also labeled with 1 wt % of Cy5.5 using a stable chemical conjugation method. The average size of the Cy5.5-labeled CNPs in PBS at 25 °C was determined to be 284.5 ± 35.5 nm using dynamic light scattering (DLS), and the spherical shape of CNPs in distilled water was also confirmed using transmission electron microscopy (TEM) images (Figure 1B,C). Also, the control nanoparticles were about 200−300 nm in size and spherical in shape, D


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Figure 1. Preparation and characterization of nanoparticles. (A) Chemical structures of glycol chitosan (GC), glycol chitosan-5β-cholanic acid nanoparticles (CNPs), polystyrene nanoparticles (PNPs), and 1,2-dipalmitoyl-sn-glycero-3-phosphocholine liposome (LNPs). (B) Dynamic light scattering (DLS) data of nanoparticles. (C) Transmission emission microscopy (TEM) images of nanoparticles.

Table 1. Size Distribution and Zeta-Potential Values of Nanoparticles

similar to CNPs, as shown in DLS data and TEM images. Furthermore, the zeta-potential values showed that CNPs, PNPs, and LNPs showed slightly cationic, anionic, and neutral surface charges, respectively, confirmed by zeta potentiometer in PBS at 25 °C. The detailed properties of each nanoparticle are summarized in Table 1. 3.2. The Stability of CNPs in Serum Conditions. The average sizes of different particles are usually determined in water or PBS without serum proteins, but this size information is not sufficient to predict their particle size and stability after injection into the body. This is because nanoparticles may readily dissociate or aggregate after exposure to serum proteins. Thus, the stability of nanoparticles in serum is very important to determine the real size of nanoparticles in the blood. To test the stability of nanoparticles in serum proteins, first, all particles were incubated in PBS without or with 15% v/v of rat serum solution at 37 °C

a

samples

μ2/Γ2

sizea (nm)

zeta-potential (mV)

CNPs PNPs LNPs

0.009 0.013 0.011

284.5 ± 36.57 226.59 ± 17.31 195.63 ± 18.74

13.25 ± 0.21 −35.1 ± 0.61 3.37 ± 0.28

Mean diameter measured by dynamic light scattering.

and their stability was observed using the SDS−PAGE test. After incubating for 1 day, Cy5.5-labeled CNPs and PNPs dropped at the upper site of SDS−PAGE still remained at the loading site, indicating their stability in PBS and serum conditions (Figure 2A). However, most Cy5.5-labeled LNPs rapidly moved down in PBS at 37 °C, indicating that they could be dissociated in the presence of SDS. In the presence of serum, some of LNP also aggregated at the upper site of SDS−PAGE, suggesting nonspecific binding of E


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number of PNPs did not even pass the 0.45 μm syringe filter (white circles in Figure 2B), unlike their easy passage through the 0.2 μm syringe filter after incubation in PBS. This result can be explained by the fact that both PNPs and LNPs rapidly aggregate in the presence of serum proteins, resulting in the formation of large aggregates that cannot pass through even the largest filter pore sizes.34 Furthermore, we also obtained TEM images of CNPs in serum solution to show the stability of CNPs. As shown in Figure 2C, there were no significant changes in TEM images, and their sizes were 289.9 ± 29.6 and 295.6 ± 6.7 nm after incubation in 15% v/v of rat serum solution for 1 and 24 h, respectively. 3.3. Macrophage Uptake and Blood Circulation Time of Nanoparticles. To analyze nonspecific uptake of nanoparticles in normal tissues, we tested the uptake of different nanoparticles into RAW264.7 murine macrophage cells. In spite of their similar sizes, RAW264.7 cells captured PNPs most efficiently among the nanoparticles (Figure 3A). Also, a large number of LNPs was rapidly taken up by the macrophage cells. However, the amounts of GC polymers and CNPs in RAW264.7 cells were even less than those of PNPs and LNPs. It has been reported that surface hydrophobicity and positive charged surface of nanoparticles causing the nonspecific protein have been also important factors on phagocytosis of nanoparticles by macrophage.35−37 Interestingly, CNPs with slightly cationic charges showed less phagocytosis than anionic PNPs or neutral LNPs. These different macrophage uptakes of nanoparticles might be closely related to the bioinertness of CNPs. The glycol groups of CNPs could be one of the potential reasons for low uptake in macrophage, but it is still not clear, and many further studies are ongoing to figure out this. In addition, MTT assay data of CNPs on RAW264.7 cells showed that the cell viability was not much changed after CNP treatment with high concentration (1 mg/mL) (Figure S1). As macrophage uptake is the main step in RES and liver clearance of nanoparticles, these data suggest that CNPs are bioinert and do not strongly stimulate RES in vivo. Importantly, blood analysis data of intravenously injected nanoparticles may reflect the bioinertness and prolonged circulation time of different nanoparticles in vivo. After intravenous injection of Cy5.5-labeled nanoparticles (5 mg/kg) into mice, each blood sample (100 μL) obtained at a predetermined time point was analyzed by measuring the fluorescent intensity of nanoparticles in the blood (Figure 3B).27,38,39 Most Cy5.5labeled PNPs completely disappeared from the blood 1 h postinjection, due to their poor bioinertness in vivo. The fluorescence signal from Cy5.5-labeled LNPs also decreased quickly and almost completely disappeared from blood after 3 h. The decreasing rates of signals from Cy5.5-labeled GC and Cy5.5-labeled CNPs were slower than those of PNPs and LNPs. Furthermore, Cy5.5-labeled CNPs showed even longer blood circulation and extended half-life (12.2 h) than Cy5.5-labeled GCs likely because their larger particle size could minimize renal clearance, compared to the water-soluble GC polymers. Around 26.9% of injected CNPs still remained in blood even after 1 day (Figure 3C). This data demonstrates that bioinert nanoparticle structure is essential for prolonged blood circulation. 3.4. Tumor Cell Uptake of Nanoparticles in Cell Culture System. After long circulation of nanoparticles in the blood, the particles should pass through the fenestrated tumor vessels by the EPR effect, and they should be selectively taken up by tumor cells for tumor-targeted drug delivery. When we tested the uptake of nanoparticles into HT29 tumor cells in pH 7.4 RPMI1640

Figure 2. In vitro stability and filtration test. (A) SDS gel running test after incubation in 15% v/v serum-containing PBS. (B) Filtration test of nanoparticles in PBS (left) and serum solution (right) measuring nearinfrared fluorescence (NIRF) of samples after filtration through cellulose acetate syringe filters at specified pore sizes. (C) Transmission emission microscopy (TEM) images of CNPs after incubation in 15% v/v of rat serum solution for 1 h (left) and 24 h (right).

serum proteins on Cy5.5-LNPs. However, both CNPs and PNPs maintained their particle structure in the presence of 15% v/v of serum solution at 37 °C, indicating that both particles were very stable in PBS or serum solution. We also observed the aggregation of nanoparticles in serum solution via the syringe filtration test, wherein each nanoparticle incubated in PBS or serum solution at 37 °C was filtered with the syringe filters with different pore sizes from 0.2 to 0.8 μm (Figure 2B). As expected, the soluble Cy5.5-labeled GC polymers in PBS and serum solution easily passed through all the syringe filter tests. This is because GC polymers are very soluble in aqueous conditions and the NIRF intensity of each polymer solution was not changed before and after syringe filtration test. Also, large amounts of both Cy5.5-labeled CNPs and LNPs in PBS passed through all syringe filter tests, indicating that the stable smaller particles without their particle aggregation can pass through the syringe filter. However, large amounts of PNPs were filtered out by syringe filter in 0.2 μm size after 1 day postincubation in PBS. It suggested that the rigid structure PNPs cannot pass through the smaller pore size of the syringe filter. Interestingly, most Cy5.5-labeled GC polymers and CNPs in serum solution successfully passed through the smaller size of syringe filters due to the very stable particle structure against the serum proteins. However, after 1 day incubation of PNPs and LNPs in serum solution, most particles were trapped by the 0.2 μm syringe filter. Interestingly, most LNPs and a large F


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Figure 3. Analysis of nonspecific uptake by macrophages and blood circulation. (A) Microscopic images showing the cellular uptake of nanoparticles to RAW264.7 murine macrophage cells after incubation for 2 h (blue, DAPI (nucleus staining); red, Cy5.5-labeled nanoparticles). (B) Time-dependent NIRF images of blood from mice after injection of nanoparticles. (C) Fluorescence intensity graph of B.

pH (Figure 4D). These results suggest that protonation of amine groups of CNPs in tumoral pH can significantly enhance the cellular uptake of CNPs, which may be instructive in the future design of tumor-targeting nanoparticles. 3.5. In Vivo Biodistribution of Nanoparticles in Liver Tumor Models. To analyze the tumor-targeting ability of these nanoparticles in vivo, liver tumor model was chosen because it is very difficult to target nanoparticles at the liver tumor, due to vigorous nonspecific entrapment of nanoparticles in normal liver tissue. Consequently, targeted imaging or drug delivery for liver tumors is in high demand.44 In our study, the liver tumor model was produced using athymic nude mice. Briefly, mice were anesthetized with ketamine, and xylazine was injected intraperitoneally (ip). After anesthesia, laparotomy was performed with direct injection of 3 × 105 HT29 tumor cells in 20 μL of saline into the left lobe in the liver.20,45−47 In order to target the liver tumor, the tumor-targeting ability of nanoparticles should be effective enough to overcome normal accumulation of nanoparticles in normal liver tissue, due to the immune response against foreign materials.11 After intravenous injection of Cy5.5labeled GCs, CNPs, PNPs, and LNPs (5 mg/kg of all samples), their biodistribution was analyzed by an in vivo NIRF imaging technique.48 For CNPs, a bright spot was observed in part of the liver, from 3 h postinjection onward, with the NIRF signal intensity increasing with time (Figure 5A). For PNPs, there was immediate accumulation in the entire liver tissue with the NIRF intensity being maintained for 1 day. For GCs and LNPs, there

media, all nanoparticles (CNPs, PNPs, LNPs) showed similar uptake and the fluorescence signals in tumor cells were much brighter than that of GC (Figure 4A). Generally, polymeric nanoparticles could be continuously internalized by cells through multiple routes including clathrin-mediated and caveolae-mediated endocytosis and macropinocytosis. Therefore, the saturation of the cell surface begins slower than if they are dissociated as polymers, and it will be one of the reasons that explain increased cellular uptake of glycol chitosan nanoparticle compared to GC polymer.40,41 It is generally known that tumor tissues are more acidic than normal tissue due to hyperactive glycolysis producing lactates. Therefore, to closely mimic the tumor tissue in vivo, we tested the uptake of CNPs into HT29 tumor cells at various pHs ranging from 5.5 to 8.0. In both microscopic images and flow cytometry data, more Cy5.5-labeled CNPs were bound to and entered HT29 tumor cells at acidic pH (Figure 4B,C). This finding suggests that the tumor cell uptake of CNPs may be enhanced in acidic tumor tissues, as compared to normal tissue. Interestingly, this accelerated cellular uptake of nanoparticles at acidic pH could not be observed in the case of Cy5.5-LNPs (Figure S2). Recently, Crayton et al. reported that the amine group of glycol chitosan (pKa ≈ 6.5) can be protonated at acidic pH and it results in fast binding and cellular uptake to cells.42,43 To analyze this, we measured the zeta-potential value of CNPs at various pHs. The size of CNPs did not vary significantly with pH, but the zeta-potential value increased from 8 mV at pH 8.0 to 53 mV at pH 5.5, supporting the protonation of CNPs at acidic G


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Figure 4. Uptake of nanoparticles to tumor cells. (A) Microscopic images showing the cellular uptake of nanoparticles to HT29 tumor cells at pH 7.4 after incubation for 2 h (blue, DAPI (nucleus staining); red, Cy5.5-labeled nanoparticles). (B) Microscopic images showing the cellular uptake of CNP to HT29 tumor cells at various pHs after incubation for 2 h (blue, DAPI (nucleus staining); red, Cy5.5-labeled CNPs). (C) Flow cytometer data showing the cellular uptake of CNP to HT29 tumor cells at various pHs. (D) Size and zeta-potential values of CNP at various pHs.

was no significant NIRF intensity in the liver tissue. After 1 day postinjection, all major organs, including the liver, lung, spleen, kidney, and heart, were then harvested and imaged ex vivo (Figure 5B). From the ex vivo images, GCs and LNPs had a moderate accumulation in the liver tumor tissue and GCs also had a bright NIRF signal in the kidney. We think that it originates from single GC polymers being smaller than nanoparticles. Furthermore, they can be degraded by proteins, so that they can be excreted from the body by rapid renal clearance.13,49 In comparison with these, CNPs had much higher NIRF signal in liver tumor tissue than any other organs. However, PNPs had significant accumulation in surrounding liver tissue, but very little localized in the tumor tissue. When comparing the harvested livers, this trend is clearly observed. As shown in Figure 5C, the tumor tissue has a white color in bright field images (red dotted circles) and the NIRF images presented substantial accumulation of CNPs in tumor tissue (white dotted circles). The NIRF intensity of tumor tissue was about 2-fold higher than that of surrounding liver tissue in the case of GCs and LNPs, but CNPs showed more than 5-fold difference, showing its superior tumortargeting ability (Figure 5D). However, PNPs showed more than a 4-fold intense NIRF signal in normal liver tissue, due to the nonspecific accumulation of PNPs in normal liver tissue. One day postinjection, mice were sacrificed, frozen, and cut transversely for total cross-section imaging by the Kodak imaging

station. In accordance with the whole body image of CNPinjected mice, the cross-section images of liver clearly presented the higher NIRF intensity of CNPs at the targeted liver tumor tissue, not in normal liver tissue (Figure 6A). After the sacrificing of all mice, we analyzed the sliced liver tissue after DAPI staining (blue color), wherein normal cells and tumor cells could be differentiated by the cell morphology in DIC images (Figure 6B). In the cases of GC- and LNP-treated mice, NIRF signals were very low in whole slides showing low accumulation in both tumor and normal liver tissue. In accordance with ex vivo images, PNPs were mainly localized in normal liver tissue, while high NIRF intensity of CNPs was observed in tumor tissue. The number of red spots from CNPs significantly changes across the interface between the tumor and normal liver tissue (red dotted lines in DIC image). These images demonstrate that the tumor targeting CNPs can differentiate between tumor tissue and normal liver tissue at a level of hundreds of micrometers. Furthermore, normal and tumor tissue of the mouse model were also imaged by CryoTEM after intravenous injection of materials and mouse sacrifice. From CryoTEM images, the particles of CNPs, PNPs, and LNPs in distilled water were shown as spherical shapes (Figure 6C). We also observed the layered structure of LNPs which is composed of a lipid membrane, but CNPs had no layered structure because its structure is a multicore aggregate, not a single core nanoparticle.36 Importantly, the CryoTEM images H


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Figure 5. In vivo and ex vivo NIRF images of liver tumor bearing mouse model after intravenous injection of Cy5.5-labeled nanoparticles. (A) Timedependent noninvasive NIRF images of liver tumor bearing mice after injection of nanoparticles. (B) Ex vivo NIRF images of various organs from 1 day postinjection of nanoparticles. (C) Ex vivo bright field and NIRF images of tumor-containing livers from B. (D) NIRF intensity graph of normal and tumor tissues in livers from B.

of tumor tissues showed a similar trend with ex vivo and histological images of CNPs (Figure 6D). Many CNPs (red arrows) were observed in the tumor tissue, but not in normal liver tissue. We also observed PNPs (blue arrows) primarily in normal liver tissue, while LNPs were not observed in either normal or tumor tissue in liver. 3.6. Optical Tomography and Correlative Light− Electronic Microscopy Analysis. To demonstrate the potential of the tumor-targeting ability of CNPs, we also conducted three-dimensional optical tomographic imaging of mice. In this experiment, we used a similar liver tumor model with luciferaseexpressing HT29 cell line. Seven days postinjection of these cells, Cy5.5-labeled CNPs (5 mg/kg) was injected intravenously and the mice were imaged by the IVIS spectrum.50 Luminescence of HT29 cells was used to mark the location of tumor tissue noninvasively in the whole body following injection of luciferin (Figure 7A, left). Interestingly, this location is exactly overlapped with NIRF signals of Cy5.5-labeled CNPs demonstrating the superior tumor-targeting ability of CNPs (Figure 7A, right). This optical tomographic imaging data shows the potential of CNPs as

tumor-targeting imaging agents for clinical cancer diagnosis. Furthermore, we used correlative light−electronic microscopy (CLEM), which enables the visualization of the structures of interest within fixed cells at the level of confocal microscopy.51 It can provide comparison of TEM and microscopic images in the same field of view and allows observation of merged images. CLEM images of tumor tissues demonstrate that NIRF signals of CNPs are colocalized with tumor cells even when observed at high resolutions (Figure 7B). Red spots in NIRF images exactly overlap with the dark spots in CLEM images; thus, it is evident that these spots are clearly CNPs in liver tumor tissues. This colocalization can also be observed at 10-fold magnification as indicated by the dotted square box in Figure 7C. Figure 7D shows even higher magnified images indicated by the boxes in Figure 7C. The electron dense particles inside of the cells in the TEM image confirmed that the CNPs were efficiently taken by tumor cells. Some CNPs were stored in the cells (boxes 1 and 2), and others were degraded to small pieces within cellular vacuoles (box 3). In all, this data demonstrates the superior tumor-targeting ability of CNPs compared to control particles in a liver tumor bearing mouse model. I


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Figure 7. Three-dimensional optical tomographic images and CLEM images of liver tumor bearing mouse model 1 day post intravenous injection of nanoparticles. (A) Optical tomographic image showing luciferase-expressing HT29 tumor in liver (left) and merged image with the signals from the injected Cy5.5-CNPs (right). (B) Ex vivo CLEM images of liver tissue 1 day postinjection of nanoparticles. (C) Higher magnified CLEM image in dotted square box in B. (D) Even higher magnified CLEM images in square boxes in B.

Figure 6. Cross-section images, histological images, and CryoTEM images of liver tumor bearing mouse model from 1 day post intravenous injection of nanoparticles. (A) Cross-section NIRF images of liver tumor bearing mice after injection of Cy5.5-CNPs. (B) DIC and fluorescence images of liver tissue after intravenous injection of nanoparticles. (C) CryoTEM images of nanoparticles. (D) CryoTEM images of liver tissue from 1 day postinjection of nanoparticles.

experiments. We selected a liver metastasis model to analyze CNPs because of the difficulty in targeting the liver with nanoparticles and the need for targeted therapeutics in the treatment of hepatocellular carcinoma. The superior tumor-targeting ability of CNPs in liver tumor bearing mice was demonstrated and analyzed in various in vivo and ex vivo NIRF images and CryoTEM data. We expect this study to not only reveal the tumor-targeting ability of CNPs in a liver tumor model but also to provide valuable information about key factors such as stability, deformability, bioinertness, and surface property for tumor-targeting of nanoparticles.

In this paper, we focused on intravenous injection of nanoparticles, but amphiphilic glycol chitosan nanoparticles also showed valuable results for oral delivery.52,53 We think that different administration route can result in different biodistribution, and glycol chitosan nanoparticles will be useful in oral delivery as well as intravenous injection. In addition, we used nanoparticles without drugs in this paper, but it will be also considered that the targetability of nanoparticles can be changed after drug loading.54

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4. CONCLUSION We analyzed the key factors underlying the tumor-targeting ability of glycol chitosan-5β-cholanic acid nanoparticles (CNPs) in liver tumor model. CNPs have sufficient stability to maintain nanoparticle structure during blood flow and circulate through narrow sinuses in spleen. Furthermore, the amine groups of CNPs were protonated in tumoral acidic pH, which may contribute to fast binding and uptake to tumor cells in tumor tissue in vivo. For precise analysis of these factors, CNPs were compared to control nanoparticles including glycol chitosan polymer (GC), polystyrene nanoparticle (PNP), and 2-dipalmitoyl-sn-glycero-3phosphocholine liposome (LNP) in various in vitro and in vivo

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.molpharmaceut.6b00507. Additional cell images and graphs (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Author Contributions △

J. H. Na and H. Koo contributed equally to this work. J. H. Na and H. Koo designed and performed the experiments, J


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Sangmin Lee and S. J. Han characterized the materials, Sangmin Lee and K. E. Lee performed animal experiments, S. Kim and H. Lee contributed materials and analyzed the data, and J. H. Na, H. Koo, Sangmin Lee, K. Choi, I. C. Kwon, and K. Kim cowrote the paper. All authors discussed the results and commented on the manuscript. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This study was funded by the Global Research Laboratory Project (NRF-2013K1A1A2A02050115), High Medical Technology Project (HI14C2755) of KHIDI, the Nano-Convergence Foundation (www.nanotech2020.org) funded by the Ministry of Science, ICT and Future Planning (MSIP, Korea) & the Ministry of Trade, Industry and Energy (MOTIE, Korea) (R201501510), and the Intramural Research Program (Theragnosis) of KIST.

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Precise Targeting of Liver Tumor Using Glycol Chitosan Nanoparticles

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