Development of conjugate-by-conjugate structured nanoparticles for oral delivery of docetaxel

Materials Science & Engineering C 107 (2020) 110346

Contents lists available at ScienceDirect

Materials Science & Engineering C journal homepage: www.elsevier.com/locate/msec

Development of conjugate-by-conjugate structured nanoparticles for oral delivery of docetaxel

T

Phuong Ha-Lien Trana,∗, Tao Wangb,c, Chunguang Yangd, Thao T.D. Trane,f, Wei Duana a

Deakin University, Geelong, School of Medicine and Centre for Molecular and Medical Research, Victoria, 3216, Australia School of Nursing, Zhengzhou University, Zhengzhou, 450001, China c Centre for Comparative Genomics, Murdoch University, Perth, WA, 6150, Australia d Medical College, Dalian University, 116622, China e Department for Management of Science and Technology Development, Ton Duc Thang University, Ho Chi Minh City, Vietnam f Faculty of Pharmacy, Ton Duc Thang University, Ho Chi Minh City, Vietnam b

A R T I C LE I N FO

A B S T R A C T

Keywords: Conjugate-by-conjugate nanoparticles Oral administration Docetaxel Permeability Cytotoxic effects against breast cancer cells Caco-2 cells uptake

In the current study, we developed interpolymer-complex structures composed of hydroxypropyl methylcellulose (HPMC) and chitosan knitted with D-α-tocopherol polyethylene glycol succinate (TPGS) to establish oral nanoparticle delivery systems that could keep the drug dose from releasing into the gastrointestinal tract for at least 6 h. Two kinds of nanoparticle formations based on the so-called conjugate-by-conjugate strategy were introduced in the study. In the first conjugate-by-conjugate structured nanoparticle formation, TPGS was conjugated with an HPMC-chitosan conjugate, followed by the drug loading process. In the second approach, the drug was loaded with TPGS directly and subsequently conjugated with the HPMC-chitosan conjugate. Beneficially, polyvinyl alcohol could act not only as a stabilizing agent but also as a crosslinking agent for the nanoparticles. This study created newly modified structures of HPMC and chitosan, altering their physicochemical properties that could then retard drug release. The nanoparticles were cytotoxic towards MDA-MB-231 breast cancer cells when docetaxel was loaded in the nanoparticles, particularly the nanoparticles produced in the second approach, demonstrating their ability to kill cancerous cells and their potential for further applications in cancer therapy. Additionally, when Caco-2 cells were used as an absorption model in a transport study, the nanoparticles in the second approach showed their capacity to increase drug permeability across the monolayers of Caco-2 cells compared to the free-drug solution. This study also illustrated the enhanced uptake of the nanoparticles by the Caco-2 cells, implying enhanced absorption through the intestine. Therefore, these oral nanoparticles can be considered for delivery systems of agents that are sensitive to the gastrointestinal tract so that they can be transported across the epithelial cells to the bloodstream to deliver the loading cargo at an optimal concentration.

1. Introduction Oral administration is a convenient approach for drug delivery for patients who undergo long-term treatment. According to a report collecting data from cancer patients, oral is preferred over intravenous (IV) administration for their treatment [1]. Patient quality of life and treatment experience are important considerations associated with increased survivorship and treatment decisions for continued treatments. However, oral drug delivery encounters major physiological barriers, such as crossing the epithelial membrane in the small intestines to reach the bloodstream [2], the effects of digestive enzymes, and drug

insolubility in the gastrointestinal (GI) tract, leading to variable absorption rates and reduced bioavailability [3]. Polymethacrylates, shellac, cellulose acetate phthalate, and polyvinyl acetate phthalate are common enteric coating polymers in the pharmaceutical industry used to prevent drug disintegration and dissolution in the gastric environment. Although the use of enteric coating polymers is one of the strategies used to overcome drug issues, including instability or irritation in the stomach, these polymers may cause allergic reactions [4] or toxicity [5,6] and may still have limited protection from drug dissolution at low pH (i.e., approximately 10% of the drug may release at gastric pH [7,8]) or fail to disintegrate in the intestinal fluid over the intended time

∗

Corresponding author. School of Medicine, Deakin University, Australia. E-mail addresses: [email protected] (P. Ha-Lien Tran), [email protected] (T. Wang), [email protected] (C. Yang), [email protected] (T.T.D. Tran), [email protected] (W. Duan). https://doi.org/10.1016/j.msec.2019.110346 Received 22 November 2018; Received in revised form 26 September 2019; Accepted 20 October 2019 Available online 22 October 2019 0928-4931/ © 2019 Elsevier B.V. All rights reserved.

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

Fig. 1. Schematic diagrams of the two different fabrication approaches to creating conjugate-by-conjugate structured nanoparticles, where drug loading occurred (A) at the middle (HTCD-P nanoparticles) and final stages (HTCP-D nanoparticles), and (B) at the initial stage (Tact-D-HC-P nanoparticles). The Tact-D-HC-P nanoparticles have smallest size, and more favourable capability of drug encapsulation efficiency and loading capacity, as compared with the other nanoparticles.

taxane class, has clinically significant antitumour activity against breast cancer cells, as it can disrupt microtubule function and cause cell apoptosis and death [18]. However, their clinical use is limited due to their extreme hydrophobicity, low water solubility and low bioavailability [19]. These shortcomings cannot be solved by IV solutions of DTX, as IV administration poses systematic toxicity due to both the high initial DTX blood concentration after IV injection and the side effects caused by polysorbate 80 and ethanol, the excipients used to dissolve DTX [20]. Our study proposed a conjugate-by-conjugate strategy for synthesizing oral hydrogel nanoparticles of DTX, whereby hydroxypropyl methylcellulose (HPMC) was activated to conjugate with D-αtocopherol polyethylene glycol succinate (TPGS) to form the conjugate HT, followed by a second conjugation of the HT conjugate with chitosan (HTC) and a third self-conjugation of HTC with polyvinyl alcohol (PVA) crosslinking and stabilization to form the final HTCP nanoparticles. The drug was loaded either at the middle stage of the conjugation process (right after HTC formation) or at the final stage of HTCP formation. A second approach involved TPGS activation, followed by loading with the drug (Tact-D); meanwhile, HPMC was conjugated with chitosan (HC) and then formed a conjugated outer layer for Tact-D to finally form Tact-D-HC nanoparticles, which were also stabilized and crosslinked by PVA (Tact-D-HC-P nanoparticles). Chitosan is a mucoadhesive polymer known for its outstanding efficacy in cellular permeability and bioavailability of orally administered nanoparticles [21]. Chitosan is protonated at low pH, but it is insoluble at high pH [22]. HPMC is one of the most common hydrophilic polymers used for oral controlled drug delivery systems due to its unique swelling and erosion ability in biological fluid. Drug dissolution and diffusion through the HPMC hydrogel delivery system are associated with

course [9]. Despite these disadvantages, enteric polymers have been used as nanocarriers in oral nanodelivery systems [10,11]; nanotechnology is the latest trend that has been developed over the past few decades for applications of drug delivery systems in biomedical and pharmaceutical areas due to the ability of nanoparticles to deliver drugs to a target site and their capacity to modulate drug solubility and dissolution. Several types of nanodelivery systems have been studied for oral medication, such as drug nanocrystals, lipid-based nanoparticles, polymeric micelles, polymeric nanoparticles, and pH-sensitive nanogels. However, there are still challenges remaining for each kind of nano system. For instance, drug nanocrystals may induce a higher toxicity potential in the GI tract [12]; lipid-based nanoparticles have poor stability under acidic conditions and large aggregation in gastric environments [13]; polymeric micelles’ capability of protecting drug release from stomach acidic pH is still limited [14]; polymeric nanoparticles face the risk of premature drug release through the enteric coating in the stomach or burst drug release upon entering the intestine [15,16]; and pH-sensitive nanogels that use enteric coating polymers may cause some side effects as mentioned above or burst drug release [17]. Furthermore, the fabrication approach of hydrophobic drug delivery has been a hot and challenging research topic, as an increasing number of studies have been investigated newly discovered chemical entities, most of which are poorly water-soluble. In our study, we developed oral hydrogel nanoparticles that overcome the abovementioned limitations seen in traditional pH-sensitive delivery systems and oral nanodelivery systems and that prevent drug release for at least 6 h so that the systems can be transported across the epithelial cells to the blood stream at the optimal concentration of loading cargo. The model drug docetaxel (DTX), a member of the 2

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

dissolved in dimethylformamide (DMF) (10 mL), which was then poured in the aqueous phase containing PVA (10%) under constant stirring, followed by ultrasonication (40% amplitude, in 10 min using a Sonics Vibra-Cell VCX130 ultrasonicator). The emulsion was then stirred and evaporated overnight (18 h) to remove the organic solvent prior to autoclaving for crosslinking. The nanoparticles (HTCD-P) were then collected by centrifugation for 30 min at 10,000×g and then dialyzed (MWCO 10,000) against acetone for purification prior to resuspending in distilled water (10 mL) and freeze-drying for 48 h. The drug was also loaded by the incubation method, where drug loading occurred after the formation of HTCP nanoparticles (HTCP-D).

swelling kinetics, polymer chain relaxation and volume expansion [23]. The HPMC matrix has been commonly used to extend the release of hydrophilic drugs [23] or improve the dissolution of poorly water-soluble drugs [24]. Neither HPMC alone nor chitosan alone is able to restrain drug release in the gastric environment. A conjugated chitosan may decrease its solubility at low pH and its positive charges to avoid general cytotoxicity or unselectively when taken up by cells. Therefore, our hypothesis is that the conjugation of HPMC and chitosan would eliminate the disadvantages and inherit advantages of each polymer. TPGS was selected in the study, as it is an ideal pharmaceutical excipient for nanoparticle-based oral drug delivery systems for cancer chemotherapy due to its capacity to enhance cellular uptake, prolong the circulation time of the nanoparticles, and improve drug permeability through cell membranes by inhibiting P-gp and reducing P-gpmediated multidrug resistance in cancer cells [25]. Moreover, TPGS has amphiphilic properties that can facilitate drug loading efficiency. HPMC must be activated in the two proposed approaches, but the difference between the two approaches is that, in the second approach, TPGS was activated and directly loaded with the drug for more efficient drug loading before the drug-TPGS compound was conjugated with the HC conjugate (Fig. 1 depicts the schematic diagrams of the two different approaches).

2.2.2. Second approach: conjugate-by-conjugate structured nanoparticles with drug loading at the initial stage TPGS was activated to have a carboxylic group by dissolving TPGS, SA and DMAP in a stoichiometric molar ratio of 1:3:1 in 50 mL of anhydrous dichloromethane (DCM) for reaction for 24 h at room temperature. Then, the functionalized (–COOH)TPGS was precipitated in cold acetone (300 mL), followed by dialysis (MWCO 1,000) against a 50% ethanol solution for 72 h to remove unreacted SA. The drug and activated TPGS were subsequently dissolved in DCM; meanwhile, HPMC was activated as described in the first approach and then conjugated with chitosan to form the HC conjugate by EDC/NHS reaction in a stoichiometric molar ratio of 1:2:2 in anhydrous DCM for 24 h at room temperature, and then reacted with chitosan as described in the synthesis of the HTC conjugate. The two DCM solutions of Tact-D and HC conjugate were mixed thoroughly under constant stirring. Then, the reaction mixture was added in the aqueous phase containing PVA (10%), followed by further steps as described in the first approach to collect the nanoparticles (Tact-D-HC-P).

2. Materials and methods 2.1. Materials Hydroxypropyl methylcellulose (HPMC, Cat. 09963), chitosan (Cat. 448869), D-α-tocopheryl polyethylene glycol succinate (TPGS, Cat. 57668), polyvinyl alcohol (PVA, Cat. 341584), succinic anhydride (SA, Cat. 239690), 4-dimethylaminopyridine (DMAP, Cat. 107700), 1-ethyl3-(3-dimethylaminopropyl)-carbodiimide (EDC, Cat. 03449), and Nhydroxysuccinimide (NHS, Cat. 130672) were purchased from Sigma Aldrich (USA). The solvents used were high-performance liquid chromatography (HPLC) grade. All other chemicals were of analytical grade and were used without further purification.

2.3. Cell culture Caco-2 cells (human intestinal cells) and MDA-MB-231 (metastatic breast cancer cells) were purchased from the American Type Culture Collection (ATCC, Manassas, VA, USA). The cell lines were cultured in Dulbecco's Modified Eagle's Medium (DMEM, Invitrogen, 12800–017) and supplemented with 10% foetal bovine serum (FBS, HyClone, A50111, USA) and 1% Glutamax (Life Technologies, 35050–061) at 5% CO2 and 37 °C.

2.2. Preparation of docetaxel-loaded nanoparticles 2.2.1. First approach: conjugate-by-conjugate structured nanoparticles with drug loading at the final stage First, HPMC was activated to carboxylic HPMC. HPMC, SA and DMAP in a stoichiometric molar ratio of 1:10:5 were dissolved in 50 mL of anhydrous dichloromethane (DCM) for reaction for 24 h at room temperature. Then, precipitation was conducted in cold acetone (300 mL), followed by dialysis (MWCO 10,000) against a 50% ethanol solution for 72 h to remove unreacted SA. Then, the activated HPMC was conjugated with TPGS. HPMC and CDI in a stoichiometric molar ratio of 1:2 were dissolved in 20 mL of anhydrous DCM for 24 h at room temperature. The homogenous reaction mixture was then precipitated in cold acetone (300 mL) and dried overnight to obtain the HPMC-CDI intermediate. Subsequently, the intermediate and TPGS in a stoichiometric molar ratio of 1:5 were dissolved in DMSO and stirred for 24 h. The reaction was terminated by dialyzing (MWCO 10,000) against DMSO for 48 h and against distilled water for another 48 h. The next step was the formation of an amide bond between the primary amine group of chitosan and the carboxylic group functions of the HT conjugate using EDC/NHS reactions in a stoichiometric molar ratio of 1:2:2 in anhydrous DCM for 24 h at room temperature. After solvent removal, the product was added to an acetic acid solution of chitosan (1% v/v, pH = 4) for reaction for 24 h, which was subsequently terminated by dialyzing (MWCO 10,000) against distilled water for 48 h. The HTC conjugate was then ready to load the drug and form nanoparticles by the solvent evaporation method. Drug and HTC were

2.4. In vitro dissolution and swelling The cumulative release of DTX from the nanoparticles was performed by transferring the nanoparticles into dialysis tubes (molecular weight cutoff: 14 kDa) filled with simulated gastric fluid (pH 1.2) and simulated intestinal fluid (pH 6.8), each with the presence of enzymes of the GI tract, respectively pepsin (for pH 1.2) and pancreatin (for pH 6.8), as well as phosphate buffered saline (PBS, pH 7.4) under sink conditions at 37 °C with stirring. One hundred microliters of the samples were withdrawn at predetermined intervals for UV–vis spectrophotometer analysis, and an equivalent amount of fresh medium was added to maintain a constant dissolution volume. The amount of drug released was expressed as a percentage of total drug and plotted as a function of time. Meanwhile, a swelling study was performed according to a previous report [26], with 100 mg of each kind of nanoparticle for the evaluation of swelling capacity in buffers having the same components as the ones used in the in vitro dissolution study (pH 1.2, 6.8 and 7.4, respectively) for 24 h. Nanoparticles in both studies were placed at pH 1.2 for 2 h, then at pH 6.8 for 6 h, and finally at pH 7.4 for the remaining time. 2.5. Thermal analysis A differential scanning calorimeter (DSC)-Model Q200 (TA Instruments, USA) was used to investigate the thermal behaviours of 3

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

was used as the hydrophobic drug DTX in formulations of drug solutions and drug-loaded nanoparticles. Caco-2 cells were seeded in an 8chamber slide (Lab-Tek I; Nunc) for 24 h. The culture medium was replaced with Hank's balanced salt solution (HBSS) when cells reached 80% confluence, prior to incubation at 37 °C for 30 min. Cells were then washed thrice with HBSS and subsequently incubated with 200 μL of the samples for 1 h prior to the washing and imaging steps. Nuclei were stained blue using Hoechst 33342 in the last 10 min of the incubation. Cells were imaged using a FluoView FV10i laser scanning confocal microscope (Olympus).

the raw materials of DTX, HPMC, chitosan, TPGS, conjugates and nanoparticles. The amounts of the samples ranged from 5 to 7 mg. The samples were weighed in a standard open aluminium pan, while an empty pan of the same type was used as a reference. The temperature used to run each sample was set from 30 to 200 °C at a rate 10 °C/min, using nitrogen as a purge gas. Calibration of temperature and heat flow was performed with indium. 2.6. Powder X-ray diffraction (PXRD) PXRD patterns were obtained with an XPert Powder diffractometer (PANalytical) using Cu–K radiation at a voltage of 40 kV and a current of 30 mA. The samples, which included the raw material of DTX, HPMC, chitosan, TPGS, conjugates and nanoparticles, were scanned in increments of 0.0130° from 6.6946° to 79.9756° (diffraction angle 2θ) at a rate of 1 s per step using a zero-background sample holder.

2.11. Transport study Caco-2 cells cultured in Corning® Transwell® inserts (0.4 μm pore diameter, 1.12 cm2 area) were used for this study after 21 days post seeding. Notably, the inserts were washed twice and equilibrated with the transport medium, 25 mM HEPES in HBSS (pH 7.4) before the analysis. DMSO (1%) in transport buffer was used as a co-solvent. Lucifer yellow was used to check the integrity of the monolayers. Drug solutions and nanoparticles (drug concentration 40 μg/mL) in the transport buffer were added on the apical side (0.5 mL). The basolateral side contained 1.5 mL of transport buffer. The transport study was conducted at 37 °C. Determination of the apical-to-basolateral permeability coefficient (Papp in centimetres per second) followed the equation: Papp = (dQ/dt)/(A × C0 × 60), where dQ/dt is the drug amount in the basolateral compartment as a function of time (mg/min), A is the monolayer area (cm2), and C0 is the initial drug concentration in the apical compartment (mg/mL).

2.7. Fourier transform infrared spectroscopy (FTIR) The spectra of the samples (again including the raw materials of DTX, HPMC, chitosan, TPGS, conjugates and nanoparticles) were recorded using an FTIR spectrophotometer (Bruker Vertex 70 FTIR) in the attenuated total reflectance (ATR) mode. The wavelength ranged from 4000 to 600 cm−1 with a resolution of 4 cm−1 and 32 scans. 2.8. Particle size and morphology of nanoparticles The particle size, size distribution and zeta potential of the nanoparticles were measured at 25 °C by dynamic light scattering (DLS) (Malvern Zetasizer-Nano ZS, Malvern Instruments Limited, Worcestershire, UK). The nanoparticle suspension (5 μL) was placed on a carbon-coated grid (300 mesh) to observe the morphology using transmission electron microscopy (TEM) (JEOL 2100 LaB6, Japan) at 100 kV. The samples were stained with a 50 μL drop of 2% methyl cellulose/4% uranyl acetate mixture (6:1) in 10 min, and rinsed three times with distilled water. The grids were dried at room temperature. The excess fluid was removed using Whatman filter paper.

3. Results and discussion 3.1. Investigation of the conjugate-by-conjugate formations in establishing nanoparticles resistant to the GI tract FTIR spectra of pure HPMC, reacted and activated HPMC (Hact), the HPMC-TPGS (HT) conjugate, and the HPMC-TPGS-chitosan (HTC) conjugate are shown in Fig. 2A. Noticeable bands of pure HPMC are observed, such as the bands associated with the hydroxyl groups in the region of ∼3500 cm-1 ʋ(OH), the band at 2800-3000 cm-1 for ʋ(CH), and the strong characteristic vibration band at 1070.5 cm-1 for ʋ(CO). The presence of the carboxyl group is implied in the spectrum of Hact with a broader band of ʋ(OH) at ∼3400 cm-1, most obviously with the peak associated with the characteristic carbonyl group (C=O) stretching frequency at 1734.8 cm-1, a strong absorption band for ʋ(CO) at 1057.2 cm-1 and the peak associated with the OH deformation vibration of the carboxyl group at 957.8 cm-1. This result is evidence of the activated COOH group of HPMC for the subsequent conjugation with TPGS and chitosan. In the first approach, activated HPMC was conjugated with TPGS through a carbonyldiimidazole (CDI) reaction. Of note, Fig. 2A shows an obvious right shifting OH band of the HT conjugate to ∼3290 cm-1 (compared to the OH band positions of the activated HPMC and TPGS). Additionally, the ʋ(CH) stretching region is narrowed to ∼2900-2940 cm-1. Overlapping of the carbonyl stretching vibration in the ester group of TPGS and in the carboxylic group of the activated HPMC are observed in the HT spectrum with a slight shift to 1745.3 cm−1 and 1727.1 cm−1, respectively. The absorption bands from 1100 to 1148 cm−1 are attributed to the characteristic C–O–C stretching vibrations of the repeated –OCH2CH2 units of TPGS. Subsequently, chitosan was able to conjugate with HPMC by conjugating the amino-terminal groups of chitosan with the remaining carboxyl group of activated HPMC through the EDC/NHS coupling reaction. As shown in the HTC spectrum, the O–H stretching and amide A (NH stretch coupled with hydrogen bonding) overlap at 3354.1 cm-1, and the ʋ(CH) stretching region is observed at ∼2870-2977 cm-1. The amide formation between chitosan and activated HPMC is observed through the appearance of bands at approximately 1650 cm-1, 1570 cm-1 and

2.9. Cytotoxicity study Cells were cultured in 96-well plates (100 μL/well) with 1500 MDAMB-231 cells per well for incubation at 37 °C with 5% CO2 for 24 h. The culture medium was Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% foetal bovine serum (FBS) and 5% streptomycin sulfate and penicillin. After 24 h incubation, the medium in each well was discarded, and then 100 μL of fresh medium containing pure drug or drug formulations was added at different concentrations, and cells were incubated at 37 °C with 5% CO2 for 72 h. In addition, medium control (only cell culture medium without cells and drugs) was used as the background, and medium containing cells without drug was used as positive control for the study. At the determined time, the cells were washed twice with PBS to eliminate the remaining drug, and 20 μL of the MTT (Sigma, Cat. M5655) solution (5 mg/mL in PBS) was added to each well. Cells were incubated for 4 h at 37 °C, and the medium was carefully removed. Then, 150 μL of DMSO was added into each well, and the plate was shaken for 10 min at 100 rpm to solubilize the MTT crystal. The absorbance of each well was measured at 570 nm using a VICTORTM X5 Multilabel Plate Reader (PerkinElmer Life and Analytical Sciences). The half maximal inhibitory concentration (IC50), which is defined as the required dose that inhibits 50% of cell growth, was calculated by plotting the cell survival rate of each drug concentration against the natural logarithm of the drug concentration. 2.10. Confocal microscopy study for cellular uptake In this study, 6-coumarin, a lipophilic fluorescent marker (green), 4

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

unchanged positions, except for the broad –OH stretching band that overlaps with the same positions of the –OH and N–H vibrations in HTC (broadened peak, which manifests the enhanced hydrogen bonding in the formation of the nanoparticles) and the disappearance of C–O stretching at 1144 cm-1. The peaks involved in the strong intermolecular interactions between chitosan and PVA are the symmetric deformation of –NH3+ groups at approximately 1555 cm-1, the –COOH peak at 1428.2 cm-1, the peak at 1657.8 cm-1 characteristic of carboxylic acid dimers overlapped with the amide I of HTC, and a small new peak at 1242.7 cm-1 attributed to the –C-O stretching of carboxylic acid. These results demonstrate the hydrogen bonding interaction between PVA and the chitosan section of HTC, implying PVA's role as a crosslinking agent in addition to its stabilization effect for nanoparticles [16,28,29]. This determination also implies avoiding the use of any common toxic crosslinking agent, such as glutaraldehyde. Additionally, the FTIR spectra (Fig. 2A) imply successful loading of DTX. Specifically, the FTIR spectrum of DTX displays the following characteristic absorption bands: N–H stretching and O–H stretching at 3490.68 cm-1 and 3343.12 cm-1, respectively, C–H stretching vibrations at 2988.84 cm-1, –C=O anhydrous ketone stretch at 1747.16 cm-1 and 1715.88 cm-1, the aromatic C=C stretching vibration at 1507.8 cm-1, and the out-of-plane vibration of N–H at 710.84 cm−1 (amide Ⅴ band). For the drug-loaded nanoparticles, the peak C=O stretching at 1715.88 cm-1 was significantly lowered, and the N–H absorption bands disappeared. For HTCD-P, the absence of the absorption bands at 3490.68 cm-1 and 1747.16 cm-1 indicated the formation of hydrogen bonds between DTX (N–H group) and HTC (carbonyl groups, C=O, hydrogen receptor and hydroxyl groups, O–H, hydrogen donor). In the case of HTCP-D, the peak at 1747.16 cm-1 did not disappear, but it was lowered. On the other hand, the absence of absorption bands at 2988.84 cm-1, 1507.8 cm-1 and 710.84 cm−1 indicate that a hydrophobic interaction could be another intermolecular force between DTX and HTC or HTCP. The method where drug loading occurred at the step before PVA was added hence induced more chemical interactions from DTX, which may lead to a stronger binding of the drug to the nanoparticles and an accordingly slower drug release. In the second approach, Hact was conjugated with chitosan and added to the drug-loaded TPGS by the EDC/NHS coupling reaction (Fig. 2B). For specifying the successful H–C conjugate, the following absorption bands were observed: a broadened N–H and O–H stretching region shifted right to 3310.5 cm-1, the narrower ʋ(CH) stretching region between 2919.2 and 2945.8 cm-1, the disappearance of C=O stretching at 1739.7 cm-1 and OH deformation vibration of the carboxyl group at 957.8 cm-1, and the left shift of the strong absorption of ʋ(CO) to 1096.4 cm-1. Meanwhile, TPGS was activated successfully, as evidenced by the appearance of the broad –OH band at ∼3340 cm-1, the right shift of the C=O carbonyl group to 1725.4 cm-1, the two new peaks at 1445.92 and 1414.64 cm-1, the right shift of the C–O–C stretching vibrations of the repeated –OCH2CH2 units from 1025 to 1092 cm-1 and the disappearance of the ʋ(CH) stretching region at ∼2900 cm-1. For DTX-loaded TPGS, the ʋ(CH) stretching region was observed at ∼2870-2940 cm-1, whereas the positions of the absorption bands of the N–H and O–H stretching region, the C=O stretch at 1747.16 cm-1, C–H stretching vibrations at 2988.84 cm-1, the aromatic C=C stretching vibration at 1507.8 cm-1 and the out-of-plane vibration of N–H at 710.84 cm−1 were absent. These results indicated chemical interactions between DTX and TPGS. Hence, the drug was persistently entrapped in the TPGS structure before being further engineered by the H–C conjugate, followed by crosslinking and stabilization of PVA for the protection of the nanoparticles in the GI tract. This engineering was confirmed by the presence of the broad N–H and O–H stretching region at ∼3330 cm-1, which was broadened further in the presence of PVA, the significantly lowered band of the ʋ(CH) stretch at 2929 cm-1, and the disappearance of the C=O stretch at 1715.88 cm-1. Therefore, the drug release from this type of nanoparticle would be slower than the nanoparticles loaded with the drug at the final stage (HTCP-D), as the

Fig. 2. FTIR spectra of (A) the first approach of the drug loading process with the HPMC-TPGS (HT) conjugate, the HPMC-TPGS-chitosan (HTC) conjugate, HTCD-P nanoparticles and HTCP-D nanoparticles (drug-loaded HTC before and after PVA stabilization, respectively); (B) the second approach of the drug loading process with the drug loaded into activated TPGS (Tact-D), protected by the HPMC-TPGS conjugate (Tact-D-HC) and finally stabilized by PVA (Tact-DHC-P).

1267.12 cm-1, which are associated with the presence of the C=O stretching of the amide I band, bending vibrations of the N–H coupled with C–N stretching (amide II band), and C–N stretching and N–H bending (amide III), respectively, whereas the C=O stretching of amide I, the N–H bending of the primary amine of chitosan and amide III were at 1657.92 cm-1, 1589.17 cm-1 and 1589.17 cm-1, respectively. Meanwhile, the peaks at 1423.62 cm-1, 1151.33 cm−1, 950.5 cm-1 and 878.13 cm−1 correspond to C–H bending, OH bending, the skeletal vibration involving C–O stretching and the CH bending out of the plane of the ring of monosaccharides of chitosan, respectively. The C–O–C asymmetric stretching and the C–O stretching of chitosan are overlapped by the C–O–C stretching vibrations of TPGS from 1070.82 cm-1 to 1155.85 cm-1. Additionally, the peak for –C–C– stretching in the aromatic ring of TPGS appears at 1456.19 cm-1. These peaks confirm the formation of the HTC conjugate by the EDC/NHS coupling reaction. PVA was used in the study, as the polymer blend of chitosan and PVA has been widely reported to possess good mechanical and chemical properties through hydrophobic side-chain aggregation and intermolecular and intra-molecular hydrogen bonds between the two components [27]. The absorption peaks of pure PVA are found at 3297.2 cm−1 (–OH stretching), 2918.5 cm-1 and 2947.2 cm-1 (alkyl stretching), 1435.9 cm-1 (C−H bending), 1333.7 cm−1 (O–H bending), 1095.7 cm−1 for the –C–O group, and 1144 cm-1 and 843 cm-1 for the C–O stretching. These peaks are observed in the HTCP spectrum with 5

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

Table 1 Average particle size and zeta potential of various nanoparticles. Nanoparticles

Average particle size (nm)

PDI

Zeta potential (mV)

HTCP-D HTCD-P Tact-D-HC-P (10% PVA) Tact-D-HC-P (5% PVA)

536.73 ± 65.34 314.37 ± 40.63 180 ± 46.19

0.451 ± 0.14 0.381 ± 0.05 0.312 ± 0.02

-23.7 ± 3.1 -30.4 ± 6.3 -35.9 ± 4.7

252.21 ± 12.32

0.358 ± 0.08

-34.8 ± 2.5

release that should have been faster—where drug was loaded after HTCP formation (HTCP-D), as the drug would be theoretically located on the outer membrane—ended up having a slower release than those from the nanoparticles where the drug was loaded to the HTC in the first approach (HTCD-P) or where the drug was first loaded with TPGS in the second approach (Tact-D-HC-P). The reason for this discrepancy was that the particle size of the HTCP-D nanoparticles was larger than that of the HTCD-P and Tact-D-HC-P nanoparticles (Table 1); indeed, the Tact-D-HC-P nanoparticles, where the drug was loaded with TPGS prior to shielding with the HC conjugate, had the smallest size, approximately 200 nm. It is in accordance with the literature that the smaller the particle size, the faster the drug release since a smaller particle size has a larger surface area per unit mass or volume [31]. Furthermore, the polydispersity index (PDI) of the nanoparticles is below 0.5, indicating a stable dispersion [32]. The lower the PDI is, the narrower the size distribution or the more uniform the nanoparticles. The Tact-D-HC-P nanoparticles hence demonstrated a more homogenous dispersion. The smaller particle size indicated a more rigid formation of the particles by the crosslinking effect. TEM morphology images of these nanoparticles confirmed their size and their round shape (Fig. 1). In addition, a zeta potential below −30 mV indicates stable nanoparticle samples [33]. The dissolution profiles of these nanoparticles was further elucidated through the swelling study (Fig. 4), which, at the same conditions of the in vitro dissolution study, was performed over 24 h. Typically, the swelling degree reflects the release behaviour of the drug loaded in the nanoparticles, probably due to the change of pH and/or temperature [34]. The Tact-D-HC-P nanoparticles expressed an explicitly higher swelling degree than the other two nanoparticles, which is in accordance with the fact that the increased drug release occurred probably due to the increased swelling level of the polymers that facilitates a porous structure and diffusion path for drug release [35]. The swelling profiles hence correspond to the dissolution profiles over 24 h. Moreover, the swelling degree of the nanoparticles began experiencing a significant increase after 6 h for Tact-D-HC-P and 8 h for the other nanoparticles. The swelling study also indicated that a higher swelling percentage could be achieved at a higher pH. In contrast, the swelling

Fig. 3. Dissolution profiles of HTCP-D, HTCD-P and Tact-D-HC-P nanoparticles in simulated gastric fluid (pH 1.2) (the first 2 h), simulated intestinal fluid (pH 6.8) (the next 6 h) and PBS (pH 7.4) (the remaining time) at 37 °C. Data shown are means ± SD, n = 3.

drug in this latter case may be located partially on the outer layer of the nanoparticles in the competitive reaction with PVA. This GI resistance effect of the final nanoparticles is expected to further protect substances such as proteins and nucleic acids from damage by the GI tract. Accordingly, the drug dissolution profiles of these nanoparticles were interesting to determine and are presented in the section below.

3.2. Investigation of in vitro dissolution profiles of various nanoparticles in their capacity to resist a mimetic GI pH environment The in vitro dissolution test was performed in simulated gastric fluid (pH 1.2) and simulated intestinal fluid (pH 6.8) with the presence of enzymes of the GI tract, respectively, pepsin (for pH 1.2) and pancreatin (for pH 6.8), as described in a previous report [30], and in phosphate buffered saline (PBS, pH 7.4). As shown in Fig. 3, while HTCP-D and HTCD-P nanoparticles could protect drug release for 8 h, Tact-D-HC-P nanoparticles could prevent drug release for 6 h. After 24 h, the amounts of drug released from HTCP-D, HTCD-P and Tact-D-HC-P nanoparticles were approximately 8%, 12.5%, and 26%, respectively. Subsequently, the drug released from these nanoparticles was approximately 56%, 75.6% and 86%, respectively, after 30 days of the in vitro test. Meanwhile, the free-drug solution almost completely released the entire drug content within the first 24 h. Hence, the nanoparticles could display a GI resistance effect for between 6 h and 8 h depending on the fabrication approach of the nanoparticles, ensuring the greatest concentration of drug released when the nanoparticles enter the bloodstream. The chemical properties of the pure polymers, such as HPMC and chitosan, were transformed, further conjugated to each other and/or to TPGS and hence improved their capacity to prevent drug release for a determined amount of time. Notably, the Tact-D-HC-P nanoparticles achieved the best percentage of drug release, approximately 90% after 30 days. Interestingly, the drug

Fig. 4. Swelling degree of HTCP-D, HTCD-P and Tact-D-HC-P nanoparticles in simulated gastric fluid (pH 1.2) (the first 2 h), simulated intestinal fluid (pH 6.8) (the next 6 h) and PBS (pH 7.4) (the remaining time) at 37 °C. Data shown are means ± SD, n = 3. *P < 0.05; **P < 0.01 compared with Tact-D-HC-P (10% PVA). 6

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

particles due to the crosslinking effect [36], the lower swelling capacity also specifies an immense crosslinking degree [37]. Therefore, the nanoparticles have a low swelling degree in the low pH condition of the stomach, and the drug hence can be protected against digestion by its enzymes. The swelling degree increases as the nanoparticles pass down the GI tract. Furthermore, DSC and PXRD studies were conducted to investigate whether thermal or crystallinity changes may contribute to the differences in drug dissolution of the nanoparticles. The DSC results (Fig. 5A) show that only a broad endotherm of the polymers or TPGS appeared on the thermograms of the drug-loaded nanoparticles or drug-loaded TPGS (Tact-D), while these drug-loaded forms did not show the endothermic peak of DTX at 189 °C, reflecting the absence of a crystalline structure [38]. On the other hand, PXRD diffractograms of these drugloaded delivery systems revealed more evidence of drug crystallinity changes (Fig. 5B and C). The more crystalline peaks of the drug are observed, the lower the drug dissolution [39]. The HTCP-D nanoparticles showed crystalline drug peaks that remained in their structures at 9.45°, 11.02° and 16.77°; whereas the HTCD-P and Tact-D-HC-P nanoparticles exposed their amorphous structure, as the numerous crystalline peaks of DTX disappeared in these diffractograms. The second fabrication approach could not only generate smaller particles but also induce changes from the crystalline to amorphous state, leading to higher drug dissolution. This was probably due to the way DTX was introduced to the system before applying PVA at the final step, facilitating a greater embedment capacity of the drug in the HTC conjugate for amorphous transformation. This was confirmed by the result in which the HTCD-P and Tact-D-HC-P nanoparticles have a higher drug encapsulation efficiency and higher loading capacity than the HTCP-D nanoparticles (Fig. 1). The amorphous transformation tendency of the HTCD-P nanoparticles also implies a major role in the transformation of the HTC structure rather than that of PVA. This role was also evidenced by examining the diffractogram of drug-loaded TPGS, where some crystalline peaks of drug were observed at 9.45°, 11.02°, 16.77°, 22.91°, 24.95°, 25.91°, 26.87°, 37.58° and 42.62° in contrast to the Tact-D-HC and Tact-D-HC-P diffractograms, where no peaks were found. In addition, the stability of the HTCP-D, HTCD-P and Tact-D-HC-P nanoparticles against various pH values and temperature conditions were investigated (Fig. 6). The particle sizes of the Tact-D-HC-P nanoparticles at different pH values were almost unchanged, whereas the particle sizes of the HTCP-D and HTCD-P nanoparticles were slightly decreased at pH 7.4 (Fig. 6A). Fig. 6A also shows that HTCP-D nanoparticles have higher PDI values than Tact-D-HC-P and HTCD-P nanoparticles. In other words, HTCP-D nanoparticles may have a higher risk of aggregation than the other two kinds of nanoparticles. In addition, the PDI values of HTCD-P nanoparticles at pH 6.8 and pH 7.4 were slightly increased compared to those at pH 1.2. However, the PDI values of Tact-D-HC-P nanoparticles at different pH values were not significantly different. With regard to the effect of temperature on storage conditions, all three kinds of nanoparticles were stable at 4 °C, 25 °C and 37 °C after 15 days. However, after 21 days, while Tact-D-HC-P nanoparticles showed their consistent stability at the three different temperatures, the particle sizes of HTCP-D and HTCD-P nanoparticles were only unchanged at 4 °C and 25 °C and were likely to increase at 37 °C. Therefore, in terms of pH and temperature, Tact-D-HC-P nanoparticles demonstrated the best stability, as they were pH-independent and unaffected by different temperatures, even at 37 °C, for at least 21 days.

Fig. 5. (A) DSC thermograms of the HPMC-TPGS (HT) conjugate, the HPMCTPGS-chitosan (HTC) conjugate, HTCD-P and HTCD-P nanoparticles, drug loaded into activated TPGS (Tact-D), and Tact-D-HC and Tact-D-HC-P nanoparticles, and PXRD diffractograms of (B) HT conjugate, HTC conjugate, and HTCD-P and HTCP-D nanoparticles, and (C) Tact-D, Tact-D-HC and Tact-D-HCP nanoparticles.

release profile of chitosan nanoparticles is generally high at low pH and decreases at higher pH [26]. This result implies a successful conjugation of chitosan in association with HPMC in the formation of a new oral controlled-release nanocarrier. The role of PVA as a crosslinking agent was also evidenced by NTA analysis and the swelling study. Specifically, the average particle size of Tact-D-HC-P nanoparticles when PVA was added at 5% was 250 nm, larger than the Tact-D-HC-P nanoparticles with 10% PVA (the nanoparticles investigated throughout the whole study), and the higher PVA concentration also induced a slower swelling degree for the nanoparticles than the lower PVA concentration. While the smaller particle size indicates a more rigid structure of

3.3. Investigation of biological activities of the nanoparticles 3.3.1. Cytotoxicity of the nanoparticles against MDA-MB-231 breast cancer cells The MTT assay revealed that the capacity of the studied nanoparticles to inhibit the proliferation of cancer cells was best in the TactD-HC-P nanoparticles, followed by the HTCD-P and then the HTCP-D nanoparticles (Fig. 7). The free DTX formulation was dissolved in 7

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

Fig. 6. (A) Particle sizes of HTCP-D, HTCD-P and Tact-D-HC-P nanoparticles in simulated gastric fluid (pH 1.2), simulated intestinal fluid (pH 6.8) and PBS (pH 7.4) after 2 h of incubation at 37 °C. Particle sizes of (B) Tact-D-HC-P, (C) HTCD-P, and (D) HTCP-D nanoparticles after 1, 7, 14, and 21 days of storage at different temperatures (4 °C, 25 °C, and 37 °C). Data shown are means ± SD, n = 3.

explaining why they were less toxic than the other two nanoparticle treatments [43]. The viability of the MDA-MB-231 cells after treatment with the various studied nanoparticles after 24 h and 48 h is shown in Fig. 8. No cytotoxic activity was observed for the empty nanoparticles (no drug), assuring that the inhibition of the cell growth or the cytotoxicity resulted only from the DTX packaged in the nanoparticles and reflecting the efficient DTX delivery of the nanoparticles to the cells to perform the pharmacological activity of the drug [44]. Cytotoxicity was influenced by drug concentration and incubation time, in which higher drug concentrations and longer incubation times inhibited a greater number of cells. At 24 h (Fig. 8A), the percentage of cell viability at a drug concentration of 7.5 μM was 55.33%, 69.56%, 79.20% and 92.22% for the HTCD-P, HTCP-D, and Tact-D-HC-P nanoparticles and the free-drug solution, respectively. The highest drug concentration, 75 μM, resulted in significant decreases in cell viability at 24 h: 5.89%, 14.57%, 42.79% and 60.23% for the Tact-D-HC-P, HTCD-P, HTCP-D nanoparticle treatments and the free-drug solution, respectively. After 48 h, a drug concentration of 75 μM (Fig. 8B) resulted in cell viabilities of 1.18%, 6.33%, 22.79% and 41.70% for the Tact-D-HC-P, HTCD-P, HTCP-D nanoparticles and the free-drug solution, respectively. The order of the nanoparticles in terms of increasing cytotoxicity was the same as that in terms of increasing in vitro drug dissolution and was consistent at 24 h to 48 h at drug concentrations from 15 μM to 75 μM. Therefore, the drug-loaded nanoparticles were more effective in suppressing cell viability of the MDA-MB-231 cancer cell line than free drug and, most importantly, avoided the toxicity associated with cosolvents (ethanol and Tween 80) used in the commercial product. For further studies on cellular uptake and transport studies of the drug-loaded nanoparticles in Caco-2 cells, cytotoxicity of HTCD-P, HTCD-P and Tact-D-HC-P nanoparticles as well as free-drug solution against Caco-2 cells was also determined (Fig. 9). The results show that IC50 of drug solutions and the drug-loaded nanoparticles are not

Fig. 7. IC50 of DTX determined after 24 h and 48 h of incubation of various drug formulations on MDA-MB-231 cells. Data shown are means ± SD, n = 3. *P < 0.05; **P < 0.01 compared with the free-drug solution.

0.25% DMSO as described in previous reports, and DMSO below 0.5% was not cytotoxic [40,41]. The DTX solution in DMSO was much less toxic compared to the nanoparticles, specifically approximately 26-fold less toxic than the Tact-D-HC-P nanoparticles, 7.5-fold less toxic than the HTCD-P nanoparticles, and 3-fold less toxic than the HTCP-D nanoparticles after 48 h incubation of treatments; the cytotoxicity levels of all treatments increased over time. However, the retarded drug release of the nanoparticles may account for their lower cytotoxicity [42]. Additionally, the larger particle size of the HTCP-D nanoparticles was a factor to take into account for 8

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

3.3.2. Cellular uptake and transport studies of the nanoparticles in Caco2 cells The use of the Caco-2 cellular model as a permeability assay to predict human oral absorption has grown exponentially over the last 20 years [46]. Caco-2, a human intestinal cell line, has the ability to undergo spontaneous differentiation to form a monolayer of cells that has the typical morphology and functions of mature absorptive enterocytes with a brush border layer, as found in the small intestine [47]. Therefore, the use of Caco-2 is a well-characterized intestinal in vitro model to evaluate the ability of substances to cross the intestinal barrier and to study their transport mechanisms [46]. Confocal microscopy was conducted to investigate the cellular uptake of the nanoparticles in Caco2 cells. For this study, coumarin-6 was loaded in the nanoparticles by the same method as the nanoparticles prepared in the previous sections. Fig. 10 shows that the best cellular uptake was that of the Tact-D-HC-P nanoparticles, followed by, in decreasing order, the HTCD-P and HTCPD nanoparticles; particularly, Tact-D-HC-P nanoparticles showed significantly enhanced uptake in comparison to the pure dye solution, which mimicked the formulation of DTX in 0.25% DMSO. The increased cellular uptake implied an enhanced absorption through Caco-2 cells, a model of the intestinal epithelial barrier [47,48]. The presence of TPGS accounts for the cellular uptake of TPGS-based nanoparticles due to the role of TPGS as a permeation enhancer and by the endocytosis mechanism [49]. Meanwhile, the internalization of solutions containing poorly water-soluble molecules has been extensively studied to show that as a result of the lipophilic characteristics, it commonly occurs via passive diffusion, which is less efficient than the endocytosis [50,51]. The cellular uptake level of nanoparticles however, also depends on the size of nanoparticles [52]. The ability to adhere to and interact with the cells of nanoparticles greatly depend on their sizes, as particles of 100–200 nm are best internalized by receptor-mediate endocytosis, while larger particles are taken up by phagocytosis [53]. The smallest size of Tact-D-HC-P nanoparticles falling into 100–200 nm size range facilitated their easier entry into and translocation across the cells [54]. Therefore, that the Tact-D-HC-P nanoparticles exhibited the best cellular uptake could be attributed not only to the ability of TPGS to enhance cellular uptake, but also to the smallest size of these particles. In addition, a transport study was performed to evaluate the permeability of the drug-loaded nanoparticles into Caco-2 cells. To check the integrity of the cell monolayers before performing the transport studies of the nanoparticles, lucifer yellow, a paracellular leakage marker, was monitored for its permeability across the monolayers. The apparent permeability coefficient was less than 0.5 × 10-6, implying that the monolayers were tight enough for the transport experiments [48]. The colloidal integrity of the nanoparticles after permeating the cell monolayers was investigated in terms of particle size and zeta potential of the nanoparticles versus time. DLS analysis of the basolateral media was recorded in 4 h, compared to that of the apical media, which was depicted at the zero time point (Fig. 11). The results demonstrated that the size and surface charge of the nanoparticles before and after the transport through the cell monolayers were resembled. This determination indicates that the nanoparticles were intact after the permeation and affirmed their capability of crossing the cell monolayers without being degraded inside cells. The result also shows that the nanoparticles seemed not likely to agglomerate or encounter other physical changes in culture media, the most challenging aspects commonly occurred with metal nanoparticles [55]. Thus, these results ensure the nanoparticles subjected to the following permeability study possess a colloidal integrity for an investigation of the best nanoparticles for effective drug delivery in further in vivo applications. Drug-loaded Tact-D-HC-P nanoparticles increased the permeability 3.5-fold compared to the free-drug solution. The permeability coefficient of the drug-loaded nanoparticles across the Caco-2 cells was found to be 6.20 × 10-5 for the Tact-D-HC-P nanoparticles, 3.22 × 10-5 for the HTCD-P nanoparticles, 2.11 × 10-5 for the HTCP-D nanoparticles, and 1.73 × 10-5 for the free-drug solution. On the other hand, the difference

Fig. 8. Cell viability following HTCD-P, HTCD-P and Tact-D-HC-P nanoparticle treatments to MDA-MB-231 cells, compared to that of the free-drug solution after (A) 24 h and (B) 48 h of incubation. Data shown are means ± SD, n = 3. *P < 0.05; **P < 0.01 compared with the free-drug solution.

Fig. 9. Determination of cytotoxicity of HTCD-P, HTCD-P and Tact-D-HC-P nanoparticles, compared to that of the free-drug solution after (A) 24 h and (B) 48 h of incubation, on Caco-2 cells. Data shown are means ± SD, n = 3.

statistically different between the formulations within 24 h or 48 h, and comparable to each other between 24 h- and 48 h-time points. In addition, this study also suggested the drug concentration could be used up to 40 μg/mL without cytotoxicity, which is also in accordance with other previous reports [25,45]. Therefore, these drug preparations were confirmed to be safe at the certain concentration used for the following studies.

9

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

Fig. 10. Confocal microscopy of the cellular uptake of coumarin 6-loaded nanoparticles. Nuclei were stained blue using Hoechst 33342. Images were taken after excessive washing and 1 h of incubation. (For interpretation of the references to colour in this figure legend, the reader is referred to the Web version of this article.)

plays an integral part in their adhesion to and interaction with the biological cells, as it is necessary for cell absorption of the nanoparticles at a sufficient level [56]. Particles in the size range of 100-200 nm are assumed to be internalized by receptor-mediated endocytosis, whereas the cellular uptake of larger particles is by phagocytosis [57]. The

between the permeability profiles of the drug-loaded nanoparticles other than the Tact-D-HC-P nanoparticles were not statistically significant from each other nor from that of the free-drug solution (Fig. 12). The increased permeability may be attributed to the higher uptake of nanoparticles by endocytosis in Caco-2 cells. Particle size

Fig. 11. Colloidal integrity of nanoparticles after permeating Caco-2 cell monolayers in terms of (A) particle size and (B) zeta potential. Data shown are means ± SD, n = 3. 10

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

DE160100900) and the 2015 Alfred Deakin Postdoctoral Research Fellowship. References [1] D. Eek, M. Krohe, I. Mazar, A. Horsfield, F. Pompilus, R. Friebe, A.L. Shields, Patient Prefer. Adherence 10 (2016) 1609–1621. [2] P. Artursson, S.D. Knight, Science 347 (2015) 716. [3] L. Lin, H. Wong, Pharmaceutics 9 (2017) 41. [4] S. Das, S.E. Jacob, Dermatitis 22 (2011) 220–222. [5] P.F. Holmes, M. Bohrer, J. Kohn, Prog. Polym. Sci. 33 (2008) 787–796. [6] D.R. Schoneker, C.C. DeMerlis, J.F. Borzelleca, Food Chem. Toxicol. 41 (2003) 405–413. [7] P.J. Tarcha, Polymers for Controlled Drug Delivery, CRC Press, 1990. [8] K. Chaturvedi, A.R. Kulkarni, T.M. Aminabhavi, Ind. Eng. Chem. Res. 50 (2011) 10414–10423. [9] M. Luangtana-anan, S. Limmatvapirat, J. Nunthanid, C. Wanawongthai, R. Chalongsuk, S. Puttipipatkhachorn, J. Agric. Food Chem. 55 (2007) 687–692. [10] S. Beyer, A. Moosmann, A.S. Kahnt, T. Ulshöfer, M.J. Parnham, N. Ferreirós, S. Wagner, M.G. Wacker, Pharm. Res. 32 (2015) 3986–3998. [11] S. Chen, C. Xu, L. Mao, F. Liu, C. Sun, L. Dai, Y. Gao, Food Bioprod. Process. 107 (2018) 88–96. [12] T.-L. Chang, H. Zhan, D. Liang, J.F. Liang, Front. Chem. Sci. Eng. 9 (2015) 1–14. [13] Y. Luo, Z. Teng, Y. Li, Q. Wang, Carbohydr. Polym. 122 (2015) 221–229. [14] G.S. Karagiannis, J.M. Pastoriza, Y. Wang, A.S. Harney, D. Entenberg, J. Pignatelli, V.P. Sharma, E.A. Xue, E. Cheng, T.M. D'Alfonso, J.G. Jones, J. Anampa, T.E. Rohan, J.A. Sparano, J.S. Condeelis, M.H. Oktay, Sci. Transl. Med. 9 (2017). [15] P. Zou, H. Chen, H.J. Paholak, D. Sun, Mol. Pharm. 10 (2013) 4185–4194. [16] M. Cetin, A. Atila, Y. Kadioglu, AAPS PharmSciTech 11 (2010) 1250–1256. [17] Z. Zeng, Y. She, Z. Peng, J. Wei, X. He, RSC Adv. 6 (2016) 8032–8042. [18] D.L. Morse, H. Gray, C.M. Payne, R.J. Gillies, Mol. Cancer Ther. 4 (2005) 1495. [19] M.F. Sohail, M. Rehman, H.S. Sarwar, S. Naveed, O. Salman, N.I. Bukhari, I. Hussain, T.J. Webster, G. Shahnaz, Int. J. Nanomed. 13 (2018) 3145–3161. [20] F. Esmaeili, R. Dinarvand, M.H. Ghahremani, M. Amini, H. Rouhani, N. Sepehri, S.N. Ostad, F. Atyabi, J. Pharm. Sci. 98 (2009) 2718–2730. [21] K. Bowman, K.W. Leong, Int. J. Nanomed. 1 (2006) 117–128. [22] E. Szymańska, K. Winnicka, Mar. Drugs 13 (2015) 1819–1846. [23] J. Siepmann, N.A. Peppas, Adv. Drug Deliv. Rev. 48 (2001) 139–157. [24] S.A. Mitchell, T.D. Reynolds, T.P. Dasbach, Int. J. Pharm. 250 (2003) 3–11. [25] H. Liu, L. Tu, Y. Zhou, Z. Dang, L. Wang, J. Du, J. Feng, K. Hu, Sci. Rep. 7 (2017) 43372. [26] P.M. Raj, R. Raj, A. Kaul, A.K. Mishra, A. Ram, RSC Adv. 8 (2018) 20809–20821. [27] S.N. Alhosseini, F. Moztarzadeh, M. Mozafari, S. Asgari, M. Dodel, A. Samadikuchaksaraei, S. Kargozar, N. Jalali, Int. J. Nanomed. 7 (2012) 25–34. [28] E.A. Kamoun, X. Chen, M.S. Mohy Eldin, E.-R.S. Kenawy, Arabian Journal of Chemistry 8 (2015) 1–14. [29] R. Kumar, P.F. Siril, Mater. Today: Proc. 3 (2016) 2261–2267. [30] D.M. Mudie, G.L. Amidon, G.E. Amidon, Mol. Pharm. 7 (2010) 1388–1405. [31] K. Yin Win, S.-S. Feng, Biomaterials 26 (2005) 2713–2722. [32] N. Mali, P. Wavikar, P. Vavia, AAPS PharmSciTech 16 (2015) 59–66. [33] C. Jacobs, O. Kayser, R.H. Müller, Int. J. Pharm. 196 (2000) 161–164. [34] M. Amoli-Diva, R. Sadighi-Bonabi, K. Pourghazi, Mater. Sci. Eng. C 76 (2017) 242–248. [35] P.H.-L. Tran, T.T.-D. Tran, J.B. Park, B.-J. Lee, Pharm. Res. 28 (2011) 2353–2378. [36] K.-K. Yun, K.-K. Kim, W. Choi, H.J. Yeon, Polymers (2017) 9. [37] E. Larrañeta, M. Henry, N.J. Irwin, J. Trotter, A.A. Perminova, R.F. Donnelly, Carbohydr. Polym. 181 (2018) 1194–1205. [38] P.H.L. Tran, H.T.T. Tran, B.-J. Lee, J. Control. Release 129 (2008) 59–65. [39] P.H.-L. Tran, T.T.-D. Tran, K.-H. Lee, D.-J. Kim, B.-J. Lee, Expert Opin. Drug Deliv. 7 (2010) 647–661. [40] A.N. Srivastava, R. Ahmad, M.A. Khan, Sci. Pharm. 84 (2016) 41–59. [41] R. Prudent, É. Vassal-Stermann, C.-H. Nguyen, M. Mollaret, J. Viallet, A. DesrochesCastan, A. Martinez, C. Barette, C. Pillet, G. Valdameri, E. Soleilhac, A. Di Pietro, J.J. Feige, M. Billaud, J.-C. Florent, L. Lafanechère, Br. J. Pharmacol. 168 (2013) 673–685. [42] F.M. Veronese, O. Schiavon, G. Pasut, R. Mendichi, L. Andersson, A. Tsirk, J. Ford, G. Wu, S. Kneller, J. Davies, R. Duncan, Bioconjug. Chem. 16 (2005) 775–784. [43] A.R. Gliga, S. Skoglund, I. Odnevall Wallinder, B. Fadeel, H.L. Karlsson, Part. Fibre Toxicol. 11 (2014) 11-11. [44] P.H.-L. Tran, T.T.-D. Tran, T.V. Vo, C.L.-N. Vo, B.-J. Lee, J. Biomed. Nanotechnol. 9 (2013) 1416–1431. [45] K. Hu, S. Cao, F. Hu, J. Feng, Int. J. Nanomed. 7 (2012) 3537–3545. [46] I.D. Angelis, L. Turco, Current Protocols Toxicol. 47 (2011) 20.26.21-20.26.15. [47] T. Lea, Caco-2 cell line, in: K. Verhoeckx, P. Cotter, I. López-Expósito, C. Kleiveland, T. Lea, A. Mackie, T. Requena, D. Swiatecka, H. Wichers (Eds.), The Impact of Food Bioactives on Health: in Vitro and Ex Vivo Models, Springer International Publishing, Cham, 2015, pp. 103–111. [48] G. Joshi, A. Kumar, K. Sawant, Drug Deliv. 23 (2016) 3492–3504. [49] S. Tan, C. Zou, W. Zhang, M. Yin, X. Gao, Q. Tang, Drug Deliv. 24 (2017) 1831–1842. [50] M. Zhou, X. Zhang, Y. Yang, Z. Liu, B. Tian, J. Jie, X. Zhang, Biomaterials 34 (2013) 8960–8967. [51] S. Majumdar, N. Kobayashi, J.P. Krise, T.J. Siahaan, Mol. Pharm. 4 (2007) 749–758. [52] J. Mosquera, I. García, L.M. Liz-Marzán, Acc. Chem. Res. 51 (2018) 2305–2313.

Fig. 12. In vitro permeability profiles of DTX from various nanoparticles compared to that of the free-drug solution in transport medium (HBSS, containing 25 mM HEPES, pH 7.4). Data shown are means ± SD, n = 3. *P < 0.05 compared with the free-drug solution.

mechanisms of particles passing through the GI barriers have been observed to occur through paracellular passage, where the particles with a size < 50 nm can transport across the epithelium by passing through the intercellular spaces between epithelial cells due to their extremely small sizes [58], through endocytotic uptake, where the particles with a size < 500 nm are absorbed by intestinal enterocytes through endocytosis, or through lymphatic uptake, where larger particles (< 5 μm) are adsorbed by the M cells of Peyer's patches [31]. It has been observed that nanoparticles of 100-200 nm rather than smaller particles achieve the best cellular uptake, challenging the conception that the smaller the particle size is, the better the cellular uptake [31]. The Tact-D-HC-P nanoparticles, which expressed the best enhanced uptake and increased permeability in the current study, fall into the particle size range of 100-200 nm. 4. Conclusion The nanoparticulate drug delivery system in the current study, despite the absence of enteric coating polymers, could take advantage of oral delivery in manifesting drug-release resistance against the GI environment and further express their cytotoxicity in MDA-MB-231 breast cancer cells. The conjugate-by-conjugate strategy promises an effective method to prevent drug release at the initial stage of GI tract travel in its dose form. The study also confirmed that the selection of an approach for introducing a drug into the loading process of the nanoparticles is critical in determining drug encapsulation efficiency. The use of PVA not only formed an interpolymer complex with a chitosanHPMC conjugate to protect the drug from diffusing out of the dose form but also itself played the role of a crosslinking agent for the nanoparticles to avoid commonly used toxic crosslinking agents. Since the amino group of chitosan was involved in the conjugation formation with HPMC, the risk of dissolving chitosan at an acidic pH could be avoided. Hence, the nanoparticles in the current study could protect the loaded therapeutic agent against the acidic denaturation and enzymatic degradation of the GI tract and facilitate the transport of drug-loaded nanoparticles from the small intestine into blood circulation for further direction to a tumour site, as evidenced by the enhanced cellular uptake and increased permeability in Caco-2 cells. Declaration of competing interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Aknowlegement Dr. Phuong Ha Lien Tran is the recipient of Australian Research Council's Discovery Early Career Researcher Award (project number 11

Materials Science & Engineering C 107 (2020) 110346

P. Ha-Lien Tran, et al.

[57] P. Couvreur, F. Puisieux, Adv. Drug Deliv. Rev. 10 (1993) 141–162. [58] S. Maiti, Chapter 6 - nanometric biopolymer devices for oral delivery of macromolecules with clinical significance, in: A.M. Grumezescu (Ed.), Multifunctional Systems for Combined Delivery, Biosensing and Diagnostics, Elsevier, 2017, pp. 109–138.

[53] S.A. Kulkarni, S.-S. Feng, Pharm. Res. 30 (2013) 2512–2522. [54] P. Foroozandeh, A.A. Aziz, Nanoscale research letters 13 (2018) 339-339. [55] K.M. Williams, K. Gokulan, C.E. Cerniglia, S. Khare, J. Nanobiotechnol. 14 (2016) 62-62. [56] K.A. Foster, M. Yazdanian, K.L. Audus, J. Pharm. Pharmacol. 53 (2010) 57–66.

12