Functionalizing Biodegradable Dextran Scaffolds ... - ACS Publications

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Functionalizing Biodegradable Dextran Scaffolds Using Living Radical Polymerization: New Versatile Nanoparticles for the Delivery of Therapeutic Molecules Hien T. T. Duong,† Felicity Hughes,† Sharon Sagnella,†,‡ Maria Kavallaris,†,‡ Alexander Macmillan,§ Renee Whan,§ James Hook,∥ Thomas P. Davis,*,† and Cyrille Boyer*,† †

Australian Centre for NanoMedicine ‡Children’s Cancer Institute Australia, Lowy Cancer Research Centre, §Biomedical Imaging Facility, Mark Wainwright Analytical Centre, ∥Nuclear Magnetic Resonance Facility, Mark Wainwright Analytical Centre, The University of New South Wales, Sydney, NSW 2052, Australia S Supporting Information *

ABSTRACT: Conferring biodegradability to nanoparticles is vitally important when nanomedicine applications are being targeted, as this prevents potential problems with bioaccumulation of byproducts after delivery. In this work, dextran has been modified (and rendered hydrophobic) by partial acetalation. A solid state NMR method was first developed to fully characterize the acetalated polymers. In a subsequent synthetic step, RAFT functionality was attached via residual unmodified hydroxyl groups. The RAFT groups were then used in a living free radical polymerization reaction to control the growth of hydrophilic PEG-methacrylate chains, thereby generating amphiphilic comblike polymers. The amphiphilic polymers were then self-assembled in water to form various morphologies, including small vesicles, wormlike rods, and micellar structures, with PEG at the periphery acting as a nonfouling biocompatible polymer layer. The acetalated dextran nanoparticles were designed for potential doxorubicin (DOX) delivery application based on the premise that in the cell compartments (endosome, lysozome) the acetalated dextran would hydrolyze, destroying the nanoparticle structure, releasing the encapsulated DOX. In-vitro studies confirmed minimal cytotoxicity of the (unloaded) nanoparticles, even after 3 days, proving that the hydrolysis products from the acetal groups (methanol and acetone) had no observable cytotoxic effect. An intriguing initial result is reported that in vitro studies of DOX-loaded dextrannanoparticles (compared to free DOX) revealed an increased differential toxicity toward a cancer cell line when compared to a normal cell line. Efficient accumulation of DOX in a human neuroblastoma cell line (SY-5Y) was confirmed by both confocal microscopy and flow cytometry measurements. Furthermore, the time dependent release of DOX was monitored using fluorescence lifetime imaging microscopy (FLIM) in SY-5Y live cells. FLIM revealed bimodal lifetime distributions, showing the accumulation of both DOX-loaded dextran-nanoparticles and subsequent release of DOX in the living cells. From FLIM data analysis, the amount of DOX released in SY-5Y cells was found to increase from 35% to 55% when the incubation time increased from 3 h to 24 h. KEYWORDS: controlled radical polymerization, biodegradable nanoparticles, drug delivery, fluorescence lifetime imaging microscopy (FLIM), self-assembled

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nanostructures, such as vesicles, micelles, and so on.3 Diverse well-defined polymeric structures now can be obtained easily using different controlled radical polymerizations (CRP),4 such as reversible addition−fragmentation chain transfer polymerization, copper mediated polymerization, or nitroxide mediated polymerization. In addition, click-chemistry can be used for efficient functionalization of polymer chains with biomolecules.5,6 Combining CRP with click is now highly prevalent in

INTRODUCTION Polymeric nanoparticles have been widely reported for drug delivery applications, as they prolong circulation times in the bloodstream and allow effective accumulation in vascularized solid tumors due to the enhanced permeability and retention (EPR) effect.1 The attraction of polymeric core−shell nanostructures as drug delivery platforms is further enhanced by the versatile modification and functionalization of the underlying polymer chemistry to incorporate different functionalities and control nanoparticle morphology.2 Polymeric nanoparticles can be constructed from a range of polymer architectures, including dendrimers, nanoparticles, and star polymers, enabling the production of a large range of © 2012 American Chemical Society

Received: Revised: Accepted: Published: 3046

March 14, 2012 September 9, 2012 September 26, 2012 October 18, 2012 dx.doi.org/10.1021/mp300144y | Mol. Pharmaceutics 2012, 9, 3046−3061

Molecular Pharmaceutics

Article

Scheme 1. Synthesis and Self-assembly of Comblike Amphiphilic Copolymer Ac-Dex-g-POEGMA

hydrophobic dextran as a polymer component together with a PEG based polymer (as a stealth polymer12) to generate chains suitable for self-assembly into nanoparticles.13 To our knowledge, this is first report on the use of acetalated dextran together with CRP to generate structures suitable for drug delivery (Scheme 1). Following assembly in water, we anticipate that the nanostructures will break down under mildly acidic conditions (e.g., in lysosomal compartments or in tumor tissue) following hydrolysis of the acetal groups, disassembling the nanoparticles, creating hydrophilic unimers, easily eliminated from the body. We demonstrate that the biodegradable, pH sensitive nanoparticles, with hydrophobic cores can encapsulate doxorubicin (DOX), while maintaining a PEG-based hydrophilic shell, ensuring solubility in biological media and good protein resistance properties. In addition, the nanoparticles were evaluated in vitro as nanocarriers. Finally, the efficient accumulation and intracellular dynamics of DOX-loaded dextran-nanoparticles and released DOX in the living human neuroblastoma cell line (SY-5Y) was monitored by combining confocal microscopy, flow cytometry, and FLIM exploiting the intrinsically fluorescent properties of doxorubicin.

the scientific literature, and some recent examples applied to nanoparticle design for imaging agents or oligonucleotide/drug delivery are given.3f,k,7 Biodegradable drug carriers are highly sought after, as they produce biocompatible degradation products that are not accumulated in the body. Both natural and synthetic biodegradable polymers have been extensively investigated as polymeric biomaterials.8 Biodegradability induced by pH change is particularly attractive for drug delivery, as acidic environments such as cell compartments and tumor tissue can be used to trigger drug release. It is also a useful design feature if polymer composition can be manipulated to tune selfassembly, thereby controlling nanoparticle size to take advantage of passive targeting via the EPR effect. As a biodegradable polymer, dextran (a polysaccharide consisting predominantly of 1,6-glucosidic linkages) is very attractive because of its colloidal biocompatibility and because it is already widely used in commercial drug formulations. Many studies have shown that dextran can be easily functionalized with biomolecules or drugs via its hydroxyl groups either by direct esterification or by the pre-introduction of spacer arms.9 In contrast, limited studies have focused on using dextran as a component in nanoparticles.10 In previous relevant work,10 dextran was hydrophobically modified using deoxycholic acid (DA) or functionalized with lipid-, thiol-, or poly(ethylene glycol) (PEG) using click chemistry to generate self-assembled nanoparticles for the delivery of anticancer drugs.10 In another relevant study, an amphiphilic diblock copolymer, dextranblock-poly(ε-caprolactone) (DEX-b-PCL) was prepared. However, the synthesis of hydrophobically modified dextran was rather complex in the previously reported work. In the present work, we have been inspired by a simple but effective powerful approach, using acetalated dextran as described in recent publications by Fréchet’s group.11 Acetalated dextran is both hydrophobic and pH sensitive, following the introduction of acid-labile acetal groups. In our work, we utilized acetalated

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EXPERIMENT SECTION Materials. All chemicals and solvents were purchased from suppliers as reagent grade and used as supplied, except where specified below. 2,2′-Azobisisobutyronitrile (AIBN) was crystallized twice in methanol before use. Oligo(ethylene glycol) methyl ether methacrylate (OEGMA; Mn = 300 g·mol−1) was deinhibited using a column of basic alumina prior to use. Deionized distilled water (dd-H2O) was purified (to a resistivity of 18.2 MΩ·cm and a total organic carbon content of less than 1 ppb) using a Sartorius Arium 611VF system. Deionized H2O was adjusted to slightly basic (pH 8) by the addition of 0.01% (v/v) triethylamine (TEA) when added to acetal containing materials, in order to prevent the degradation of acetal groups. 3047

dx.doi.org/10.1021/mp300144y | Mol. Pharmaceutics 2012, 9, 3046−3061


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Figure 1. 1H NMR spectrum of acetalated dextran in CDCl3, recorded at 300 MHz at 293 K.

Scheme 2. Functionalization of Acetalated Dextran with RAFT Agent

× 10−2 mol) were both added with stirring, followed by the addition of molecular sieves, almost to the liquid line. Following reaction, the molecular sieve was removed by filtration. The liquid mixture was filtered through a 0.45 μm Millipore filter and then added dropwise to distilled H2O (pH 8), yielding a precipitate that was isolated by centrifugation and repeated washing three times with additional dd-H2O (pH 8), prior to drying in vacuo for 24 h. This procedure yielded Acdextran (1.58 g) as a white powder. The polymer was analyzed by 1H NMR spectroscopy (Figure 1) and DMAc SEC (Mn = 10 500 g/mol, PDI = 1.43). 1H NMR (400 MHz, CDCl3): 1.39 (s, br, −C−CH3, 6 × (a+b) × H), 3.25 (br, −OCH3, 3b × H), 3.45 (br, −CH2−O, 2H), 3.60−4.15 (br, C−H, glucose ring, 4H), 4.92 and 5.13 (br, CH−O, 1H). Note: a and b correspond to cyclic and acyclic acetal rings per glucose unit, respectively.

RAFT agent 4-cyanopentanoic acid dithiobenzoate (CPADB) was prepared according to published procedures.14 Doxorubicin hydrochloride (DOX·HCl) was obtained from Sigma-Aldrich. Dialysis membranes (with a molecular weight cutoff (MWCO) of 3500 Da) were provided by Fisher Biotec (Cellu SepT4, regenerated cellulose-Tubular membrane). Methods. Acetalation of Dextran. The method used for the acetalation of dextran was slightly modified from that published by Fréchet et al.11a Dextran (Mw = 10000 g mol−1, 1.7 g, 1.7 × 10−4 mol) was mixed with dry dimethyl sulfoxide (DMSO) (40 mL) and stirred ensuring dissolution. Approximately 10 cm3 of 5 Å molecular sieves was added, and the mixture was stirred for a further 5 min to complete the dehydration of DMSO. p-Toluene sulfonic acid (p-TsOH) (170 mg, 8.94 × 10−4 mol) and 2-methoxypropene (5 mL, 5.22 3048



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The hydrolysis of Ac-dextran was studied using 1H NMR measurements on samples dissolved in DCl; see Figure S1 in the Supporting Information. Synthesis of Dextran-Based macroRAFT (Scheme 2). Acdextran (0.5 g, 4.1 × 10−5 mol) was dissolved in dichloromethane (DCM) (8 mL), and then 4-cyanopentanoic acid dithiobenzoate (CPADB) (50 mg, 1.79 × 10−4 mol) was added and stirred until dissolved. N,N′-dicyclohexylcarbodiimide (DCC) (100 mg, 4.85 × 10−4 mol) dissolved in additional DCM (1 mL) was added, and the mixture was placed in an ice/ water bath and stirred for 1 h. 4-Dimethylaminopyridine (DMAP) (2 mg, 1.64 × 10−5 mol) was dissolved in DCM (1 mL) and added to the reaction, which was then stirred at room temperature overnight. The reaction mixture was dialyzed against acetone for 2 days, with the solvent replaced twice daily. After dialysis, the acetone was removed by rotary evaporation and the resultant pink powder was dried to yield the Ac-dextran macroRAFT agent (0.54 g). The Ac-dextran macroRAFT agent was analyzed by 1H NMR spectroscopy (Figure S2 in the Supporting Information) and DMAc SEC (Mn = 12000 g/mol, PDI = 1.43). 1H NMR (400 MHz, CDCl3): 1.39 (s, br, −C− CH3 (cyclic acetal), 6(a+b) × H), 1.95 (s, br, C−CH3 (RAFT), 3nH), 2.4−2.8 (m, br, CH2 (RAFT), 4nH), 3.25 (br, −OCH3, 3b × H), 3.45 (br, −CH2−O, 2H), 3.60−4.15 (br, C−H, glucose ring, 4H), 4.92 (br, CH−O, 1H), 5.13 (br, CH−O, 1H), 7.38 (t, br, CH benzyl ring, 2nH), 7.55 (t, br, CH benzyl ring, nH), 7.90 (d, br, CH benzyl ring, 2nH). Note n, a, and b stand for the number of RAFT agents and cyclic and acyclic groups per glucose unit, respectively. RAFT Polymerization of OEGMA Using Dextran-Based macroRAFT. [OEGMA]/[RAFT agent]/[AIBN] = 33:1.0:0.19. The Ac-dextran macroRAFT (100 mg, 4.20 × 10−5 mol of RAFT agent), OEGMA (420 mg, 1.4 × 10−3 mol), and 2,2′azobisisobutyronitrile (AIBN) (1.3 mg, 8.0 × 10−6 mol) were dissolved in dimethylacetamide (DMAc) (4 mL). The reaction mixture was divided equally into four vials to study the kinetics of the polymerization. Each vial was sealed and purged with nitrogen for 30 min. The reaction mixtures were then immersed in an oil bath at 70 °C, and samples were taken over a period of 14.5 h. The polymerization was terminated by placing the samples in an ice bath for 5 min. The polymer was purified three times by precipitation in a mixture of petroleum spirits and diethyl ether (1:1, v/v), and then isolated by centrifugation and dried under vacuum, yielding the Ac-dextran-g-POEGMA polymer as a pink solid. Table S2 in the Supporting Information reports the molecular weights and PDI of the different copolymers. Self-assembly of Ac-Dextran-g-POEGMA. Ac-dextran-gPOEGMA polymer (10 mg) was dissolved in DMSO (200 μL), and a phosphate buffer solution (pH 7.4, 2 mL) was added at a rate of one drop every 10 s. The solution was stirred very gently for 2 h and then dialyzed against buffer solution for 2 days, changing the solvent twice daily to remove DMSO. The targeted final nanoparticle concentration was 4 mg mL−1. Doxorubicin Loading and Release. The micellar structure formed from Ac-dextran-g-POEGMA (Mn = 21000 g mol−1 and PDI = 1.47), obtained after 14.5 h of OEGMA polymerization was used for DOX loading experiments. In the presence of triethylamine (50 μL), 20 mg of Ac-dextran-g-POEGMA and 2 mg of DOX.HCl were dissolved in DMSO (2 mL). Phosphate buffer solution (pH 7.4, 6 mL) was then added and the mixture stirred at room temperature for 1 h. The mixture was first dialyzed (MWCO 3500 Da) against a DMF and buffer solution

(pH 7.4) mixture (1:9, v/v) for 24 h and then against buffer solution for 24 h to remove solvent, triethylamine, and free DOX, before lyophilization, yielding a red powder. After dissolving Ac-dextran-g-POEGMA nanoparticles (1 mg) loaded with DOX in DMSO (1 mL), the solution absorbance at 485 nm was measured on a CARY 300 spectrophotometer (Bruker). The amount of encapsulated DOX in Ac-dextran-gPOEGMA nanoparticles was quantified to be 5.1 wt % using a calibration curve of DOX·HCl in DMSO. The unencapsulated DOX was determined by adding 0.5 mL of Ac-dextran-gPOEGMA DOX nanoparticles in a Vivaspin 10 mL centrifugal concentrator (MWCO 3500 Da). The DOX content in the collected tube was determined using fluorescence spectroscopy at the excitation wavelength of 480 nm, emission wavelength of 560 nm, and slit width of 10 nm based on a standard curve of DOX·HCl. The results showed less than 0.25% loss of DOX, revealing the low amount of free drug in the nanoparticles solution. The release rate of DOX from DOX-loaded dextran nanoparticles was performed in vitro in acetate buffer (pH 5.0 to mimic the pH conditions of endosomes and lysosomes) and phosphate buffer (pH 7.4). The DOX-incorporated nanoparticle solution after dialysis (2 mL, 1 mg mL−1, 5.1% of DOX) was placed in a dialysis membrane tube (MWCO 3500 Da), prior to immersion in a 200 mL bottle with 100 mL of acetate buffer (pH 5.0) or phosphate buffer (pH 7.4). The bottles were covered with aluminum foil and placed in a shaking incubator with a stirring speed of 100 rpm at 37 °C. At specific time intervals (6 h, 1, 2, 3, 4, 5, 6, 7 days), 3 mL of the media was taken for DOX concentration analysis. An equal volume of fresh media was replaced, and the sink condition, where the doxorubicin concentrations in the release medium were below 10% of its aqueous solubility (2.1 mg/mL), were maintained during the course of the release study.15 DOX was determined using fluorescence spectroscopy at the emission wavelength of 560 nm, the excitation wavelength of 480 nm, and the slit width of 10 nm. The release experiments were conducted in triplicate, and the result is presented as an average. Cell Culture. The human neuroblastoma cell line SY-5Y and normal fetal lung fibroblasts MRC5 were grown in Dulbecco’s Modified Eagle’s Medium: Nutrient Mix F-12 (DMEM) supplemented with 10% (v/v) Fetal Calf Serum (FCS) in a ventilated tissue culture flask T-75. The cells were incubated at 37 °C in a 5% CO2 humidified atmosphere and passaged every 2−3 days when monolayers at around 80% confluence were formed. Cell density was determined by counting the number of viable cells using a trypan blue dye (Sigma-Aldrich) exclusion test. For passaging and plating, cells were detached using 0.05% trypsin-EDTA (Invitrogen), stained using trypan blue dye, and loaded on the hemocytometer. All the experiments were done in triplicate. Cell Viability. The cytotoxicity of free DOX, dextran nanoparticles, and DOX-loaded dextran nanoparticles was tested in vitro by a standard Alamar Blue Assay. The assay is based on the ability of living cells to convert blue redox dye (resazurin) into bright red resorufin, which can be read in a spectrophometric reader. Nonviable cells rapidly lose metabolic capacity and, thus, do not generate a color signal. The intensity of the color is proportional to the cell viability. The cells were seeded at 10000 cells/well for SY-5Y and 5000 cells/well for MRC5 in 96 well tissue culture plates and incubated for 24 h. The medium was then replaced with fresh medium containing 3049



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trace of acid. Further detailed characterization of acetalated dextran was carried out in both solution and the solid state as described below: Solution state NMR spectroscopy was also performed on a Bruker Avance III 600 NMR spectrometer operating at 14.1 T, with a frequency of 600.13 MHz for 1H NMR and one of 150.9 MHz for 13C NMR. Solid state NMR was performed with a Bruker Avance III NMR spectrometer operating at 300 MHz (7.05 T). Samples (approximately 80 mg) were packed into 4 mm o.d. zirconia rotors, capped with Kel-F rotor caps and spun at the magic angle at 12 kHz under control of a Bruker MAS II unit (to ±2 Hz) to eliminate potential interference from spinning side bands. Spectra were acquired at the 13C frequency of 75.47 MHz, using magic angle spinning (MAS) with crosspolarization (CP). CP was initiated by a 2.5 μs 1H pulse, followed by a 1−10 ms contact time (ramped for 1H; highpowered 1H decoupling using spinal64 and a 5−20 s recycle time, of which the optimum parameters were 3 ms and 3 s, respectively), or with a 13C excitation pulse of 2.0 μs (45° pulse) (high-powered 1H decoupling using spinal64 and a recycle delay of 20 s for direct polarization). The degree of substitution was determined from the relative ratio of the integrated peaks of the CP/MAS spectra. All spectra were processed using BrukerTopSpin 3.0 software. Monomer conversion was calculated by solution 1H NMR using the following equation: α O E G ‑ M A = 100 − [(∫ I CHCH 5.5−6.5 ppm )/(∫ I OCH 2 4.2 ppm ) × 100], where ∫ ICHCH 5.5−6.5 ppm and ∫ IOCH2 4.2 ppm correspond to the intensities from the methacrylate bonds of the OEG-MA and CH2 ester, respectively. Mass Analysis. Electrospray-ionization mass spectrometry (ESI-MS) experiments were carried out using a Thermo Finnigan LCQ Deca ion trap mass spectrometer (Thermo Finnigan, San Jose, CA). The instrument was calibrated with caffeine, MRFA, and Ultramark 1621 (all from Aldrich) in the mass range 195−1822 Da. All spectra were acquired in positive ion mode over the mass to charge range, m/z, 100−2000 with a spray voltage of 5 kV, a capillary voltage of 44 V, and a capillary temperature of 275 °C. Nitrogen was used as sheath gas while helium was used as auxiliary gas. The sample (0.1 mg/1 mL) was prepared by dissolution in water. 56 spectra were recorded in positive ion mode with an instrumental resolution of 0.1 Da. All reported molecular weights were calculated via the program package CS ChemDraw 6.0 and are monoisotopic. The theoretical molecular weight over charge ratios (m/z, assuming z + 0.1) are calculated using the exact molecular mass of the predominant isotope within the structure. Gel Permeation Chromatography (GPC) Measurements. DMAc GPC analyses of the polymers were performed in N,Ndimethylacetamide [DMAc; 0.03% w/v LiBr, 0.05% 2,6-dibutyl4-methylphenol (BHT)] at 50 °C (flow rate = 1 mL·min−1) using a Shimadzu modular system comprised of an SIL-10AD autoinjector, a PL 5.0-mm bead-size guard column (50 mm × 7.8 mm) followed by four linear PL (Styragel) columns (105, 104, 103, and 500 Å), and a RID-10A differential refractiveindex detector. The SEC calibration was performed with narrow-polydispersity polystyrene standards ranging from 168 to 106 g·mol−1. A total of 50 μL of polymer solution (2 mg·mL−1 in DMAc) was injected each time, for analysis. Aqueous SEC was performed using a Shimadzu modular system comprising a DGU-12A solvent degasser, on a LC10AT pump, a CTO-10A column oven, and a RID-10A refractive index detector and a SPD-10A Shimadzu UV Vis

free DOX, dextran nanoparticles, and DOX-loaded dextran nanoparticles over an equivalent DOX concentration range of 0.001−50 μM. At 72 h post drug/nanoparticle incubation, treatments were removed and fresh media was added (100 μL) followed by the addition of Alamar Blue dye (20 μL) to each well, and the cells were incubated for 6 h. Cell viability was determined as a percentage of untreated control cells, and IC50 values were calculated via regression analysis using Microsoft Excel. In-Vitro Accumulation and Intracellular Dynamics of DOX-Loaded Dextran-Nanoparticles and Released DOX by Flow Cytometry, Confocal Laser Scanning Microscopy, and FLIM Analysis. To examine the intracellular accumulation of DOX by flow cytometry, SY-5Y cells were seeded at a density of 2 × 105 cells/well in six-well tissue culture plates. The cells were left to grow for 24 h in DMEM media containing 10% FBS at 37 °C in 5% CO2 atmosphere. After 24 h, doxorubicin loaded nanoparticles were added to the wells (concentration of 0.125 μM based on DOX) and the cells were incubated for 30 min, 1 h, 6 h, or 24 h. Following particle incubation, cells were rinsed twice with PBS to remove any non-uptaken nanoparticles. Cells were harvested by trypsinization and resuspended in 500 μL of PBS for flow cytometry analysis using the FACScanto flow cytometer (BD Biosciences). Data shown are the mean fluorescent signal for 10,000 cells. SY-5Y cells that were not treated with nanoparticle solution were used as a control. Confocal laser scanning microscopy (CLSM) was also used to examine the accumulation and biodistribution of free DOX and DOX-loaded dextran nanoparticles by the SY-5Y cells. CLSM images of cells were obtained using a confocal microscope (Leica TCS SP5 II) with a 60× oil immersion objective (1.4 numerical aperture). The 488-nm line from an argon ion laser was used for excitation, and emission was collected between 565 and 630 nm. For this purpose, SY-5Y (5000 cells/well) were plated into four-well Lab Tek chamber slides precoated with poly-D-lysine hydrobromide (SigmaAldrich) for 10 min. After 48 h of seeding, the medium was replaced with fresh medium containing either free DOX or DOX-loaded dextran nanoparticles (0.5 μM equivalent DOX concentration), and cells were incubated at 37 °C for 6 h. Cells were then rinsed with PBS and fixed in 4% paraformaldehyde for 10 min at room temperature. Following fixation, slides were left to air-dry briefly, and then a drop of ProLong Gold mounting media was placed onto the slide and a clean coverslip was carefully placed on top to avoid trapping any air bubbles. The slide was stored in the dark overnight and then sealed with nail polish. Slides were stored at 4 °C until required for imaging. For FLIM analysis, SY-5Y cells were seeded into sterilized glass bottom dishes (Mat-Tek) (100000 cells/well). After 48 h of seeding, the medium was replaced with fresh medium containing either free DOX or DOX-loaded dextran nanoparticles (0.5 μM equivalent DOX concentration) and cells were incubated at 37 °C for 3 and 24 h. Lifetime images were recorded using a 60× water-immersion objective (1.2 numerical aperture, Olympus). Analysis. Nuclear Magnetic Resonance (NMR). Basic structural analyses were performed using solution 1H NMR spectroscopy in CDCl3 on a Bruker Avance III 300 spectrometer (300.13 MHz). To prevent acid-catalyzed hydrolysis of the acetal groups, CDCl3 was passed through a plug of basic alumina prior to preparing samples to remove any 3050



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Scheme 3. Reaction Scheme for the Acetalated of Dextran

performed using SimFCS software developed at the Laboratory for Fluorescence Dynamics, University of California at Irvine.

detector (flow rate: 1 mL/min). The column system was equipped with a Polymer Laboratories 5.0 mm bead-size guard column (50 mm × 7.8 mm) followed by two PL aquagel MIXED−OH columns (8 μm). Calibration was performed with PEO standards ranging from 106 to 909,500 g/mol. UV−Visible Spectroscopy. UV−visible spectra were recorded using a CARY 300 spectrophotometer (Bruker). Dynamic Light Scattering (DLS). DLS measurements were performed using a Malvern Zetasizer Nano Series running DTS software and using a 4 mW He−Ne laser operating at a wavelength of 633 nm and an avalanche photodiode (APD) detector. The scattered light was detected at an angle of 173°. The temperature was stabilized to ±0.1 °C of the set temperature. To reduce the influence of larger aggregates, the number-average hydrodynamic particle size is reported. The dispersity index is used to describe the width of the particle size distribution, as calculated from the DTS software using a cumulant analysis of the measured intensity autocorrelation function; it is related to the standard deviation of the hypothetical Gaussian distribution (i.e., dispersity = s2/ZD2, where s is the standard deviation and ZD is the Z average mean size). Transmission Electron Microscopy (TEM). The sizes and morphologies of nanoparticles were observed using a transmission electron microscopy JEOL1400 TEM at an accelerating voltage of 100 kV. The particles were dispersed in water (1 mg/ mL) and deposited onto a 200 mesh, holey film, copper grid (ProSciTech). Osmium vapor (OsO4) staining was applied. Atomic Force Microscopy (AFM). Samples for AFM imaging were prepared by simply depositing Dox-loaded dextran nanoparticles in aqueous solution (0.5 mg mL−1) onto cleaved mica prior to drying under an ambient atmosphere. The morphology of the resulting deposits was analyzed by Multimode 8 (Bruker) operating in tapping mode in air using standard Si probes. Fluorescence Lifetime Imaging Microscopy (FLIM). Fluorescence lifetime imaging (FLIM) was performed on a Picoquant Microtime200 inverted confocal microscope with a 60×, 1.2NA water-immersion objective. Excitation was via a fiber-coupled, pulsed laser diode operating at 470 nm with a pulse width below 200 ps. The emission was collected using a 550 nm long-pass filter and a single-photon avalanche diode (SPAD) (PDM, MicroPhoton Devices) connected to timecorrelated single-photon counting (TCSPC) electronics (Picoharp300, Picoquant). The data was acquired and analyzed using SymphoTime software (Picoquant). Phasor analysis of FLIM data was

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RESULTS AND DISCUSSION Acetalation of Dextran. Dextran is an appealing starting material to make nanoparticles for bioapplications, as it is biodegradable and approved for in vivo use as a plasma expander.16 Dextran was modified in the presence of 2methoxypropene and p-toluene sulfonic acid in DMSO (Scheme 3), as inspired by Fréchet’s work, converting the hydroxyl groups into pH-responsive acetal groups.17 The acetalation has two crucial purposes in modifying dextran properties: first, to switch the solubility of the dextran from hydrophilic to hydrophobic and, second, to make the chains pH sensitive. The switch in solubility is important for encapsulating hydrophobic payloads, such as many anticancer drugs. The pH sensitivity is imbued as the acid catalyzed hydrolysis of the acetal groups, and it is reversible at low pH values, thus regenerating hydrophilic dextran, releasing small amounts of acetone and methanol. This pH sensitivity provides a switch mechanism for drug delivery applications, as areas targeted for therapy are often mildly acidic (such as the lysosome compartment or tumor tissue). An important aspect of this study centered on characterizing the acetalation reaction. We used both solution NMR and solid state NMR techniques to gain comprehensive information on the acetalation process, allowing us to tune the reaction conditions, with some control over the degradation time and grafting density of the POEGMA side chains. The acetalation reaction results in two different acetal structures attached to the dextran: cyclic and acyclic (the structures are shown in Scheme 4). The cyclic and acyclic structures form at very different rates and also subsequently degrade at different rates. It is known that acyclic acetals form more quickly; however, cyclic acetals are more stable and degrade more slowly.17 The total number of acetals and the distribution between cyclic and acyclic forms therefore affects the degradation time Scheme 4. Types of Acetals Attached to Dextran Rings during Acetalated

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of any subsequent material made from the modified dextran. For nanoparticles carrying a drug, the degradation profile of dextran plays an important role for the drug release and is, therefore, crucial. Careful acetal analysis also yields information about the number of free (unreacted) hydroxyl groups remaining available for RAFT group attachment. The relative distribution of cyclic and acyclic groups and RAFT functionality (subsequently used for attaching hydrophilic chains) alters the self-assembly of the final polymers, thus dictating the nanoparticle morphology (vide infra). The acetalation method we used was a modified version of that published by Fréchet et al.11a,c 1H NMR analysis of Ac-dextran in deuterated chloroform confirmed the successfully modification of dextran, as indicated by the signal of CH3− at 1.2 ppm. The signals at 5.13, 4.92, and 4.15−3.60 ppm can be attributed to the anomeric and ring protons of dextran, confirming a quantitative modification (Figure 1) in good agreement with Fréchet’s work.11a The exact proportion of acyclic and cyclic acetals was determined using two different methods: a hydrolysis method and a solid state NMR method. The Fréchet group reported on the identification of acetalation degree via hydrolysis of the acetal groups,11a as the hydrolysis of acyclic acetal releases both acetone and methanol, while cyclic acetal releases only acetone (Scheme S1 in the Supporting Information). This method was attempted in the present study to determine the amount and composition of acetal groups after acetalation (Figure S1 in the Supporting Information). While this method gives a good estimate of the degree and type of acetal coverage, some problems with reproducibility were evident (e.g., a sample measured on two different instruments within a few hours gave conflicting results). The hydrolysis method assumes that all acetals are hydrolyzed and that no acetone or methanol is lost by evaporation after hydrolysis; with care, this problem can be avoided. However, two potential sources of error remain: methanol and dextran signals are proximate and difficult to resolve, thus affecting their relative integration. Second, a potential complication is proton exchange of acetone in acidic solutions, potentially affecting the integration of the acetone signal, skewing the results. A new method for characterizing the degree and type of acetalation was developed using solid-state cross-polarization magic angle spinning (CP/MAS) 13C NMR spectroscopy. The premise behind the characterization approach was that the quaternary carbons from the cyclic and acyclic acetals would appear at different chemical shifts because of the ring strain present in the cyclic acetals, and this shift could be used for quantification. To facilitate discussion, we refer the reader to Figure 2, where we specify our nomenclature for the relevant carbon atoms. On the basis of previous published work on CP/MAS NMR analysis of dextrans,18 it was predicted that the anomeric carbon signal would appear at around 95 ppm with the remainder of the dextran carbons producing signals between 60 and 80 ppm. A study on the chemical shifts of 1,3-diol acetonides suggested that the methyl resonances from the cyclic acetals would appear at either 24−25 ppm or 30 and 19 ppm, depending on the conformation of the acetal ring (anti and syn, respectively).19 The same study indicated that the quaternary carbon of a 6membered acetonide would appear between 98 and 100 ppm. We theorized that this would be similar to an acyclic quaternary carbon, as a 6-membered ring is relatively unstrained; therefore, the cyclic quaternary carbon in a 5-membered ring would

Figure 2. Nomenclature for carbons in acetalated dextran.

appear downfield. This hypothetical signal assignment was subsequently reinforced by data from a published study on the acetonation of glycols, indicating that the cyclic quaternary carbon would appear at around 111 ppm.20 The hypothetical signal predictions proved to be accurate, as shown in the spectrum shown in Figure 3. Although the anomeric carbon

Figure 3. Solid state 13C CP/MAS NMR spectrum of acetalated dextran with labeled signals.

and the acyclic quaternary carbons were not resolved, this NMR method still gave a spectrum with unique and easily identifiable signals for both the cyclic and acyclic acetals, in the cyclic quaternary and methoxy signals, respectively. The nomenclature for expressing the degree of acetal substitution (DS) was defined by Fréchet and co-workers as the number of each type of acetal substituent in 100 repeat units of dextran.11c This nomenclature is particularly facile, as it is simple to calculate DSby calibrating the dextran carbon signal to 5; it simply requires multiplying the individual integrals by 100 to determine the acetal number per 100 rings. The DS of cyclic acetals was taken as 100 times the integral for the cyclic quaternary carbon, while the DS of the acyclic was taken as the average of the DS determined individually for both the acyclic quaternary and the methoxy peaks (as a crosscheck). Integer DS values are reported. The spectrum shown in Figure 3 yielded DS values of 93 for cyclic acetal and 40 for 3052



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comparing the intensity of vinyl proton peaks (at 6.1 and 5.6 ppm) to that of the −CH2 ester signal at 4.2 ppm (data not shown). As shown in Figure 4A, the monomer conversion

acyclic acetal contents. The ratio of the different substituents can then be defined easily by dividing the DS cyclic by the DS acyclic, giving 1.0:2.3 acyclic to cyclic acetals in this example case. The acetal characterization results proved reproducible over three independent measurements on the same material. It is also relevant to define the remaining functional hydroxyl groups, simply accessible from the DS values. Taking 100 repeating units, there are 300 hydroxyls that are available for substitution (three on each ring). The number of substituted hydroxyls is the DS acyclic plus twice the DS cyclic, as a cyclic acetal covers two hydroxyls, allowing easy calculation of the percentage of total substituted hydroxyls. SEC analysis showed a slight increase in molecular weight of dextran after acetalation and coupling with RAFT functionality from 10000 g/mol to 12000 g/mol, while the polydispersity remains relatively constant (1.34−1.43) (Table S1 in the Supporting Information). Synthesis of Dextran Based Chain Transfer Agent. RAFT agents were attached to the acetalated dextran to form macroRAFT agents capable of exerting control over the polymerization of OEG-MA via a “grafting from” process. It is very important to choose the appropriate RAFT agent for the specific monomer of interest in order to maintain living polymerization conditions. 4-Cyanopentanoic acid dithiobenzoate (CPADB), an example of the dithiobenzoate class of RAFT agents, was used to control the polymerization of OEGMA, as it is known to be effective for methacrylates, acrylates, and styrene.21,22 CPADB was attached to the acetalated dextran via a coupling reaction with the free hydroxyls present on the dextran molecules, yielding a dextran-based macroRAFT agent as a pink powder. The presence of the RAFT functionality on the dextran chains was confirmed using UV/vis spectroscopy via the characteristic absorption of the thiocarbonyl group at 305 cm−1 (Figure S3 in the Supporting Information).23 In addition, 1H NMR analysis confirms the attachment of CPADB on the acetalated dextran by the appearance of a new signal at 7.1−7.8 ppm attributed to the benzyl ring of the RAFT agent (Figure S2 in the Supporting Information). Using the signal of the benzyl ring at 7.1−7.8 ppm and the CH of dextran at 3.5− 4.5 ppm, we are able to estimate the number of RAFT per chain (Comment of Figure S2 in the Supporting Information). On average, there are 5 RAFT agents per dextran chain. The attachment of RAFT agents does not alter the structure of the acetalated dextran polymer, as indicated by 1H NMR, or the physical-chemistry properties (solubility). In this particular example, the number of RAFT groups per chain was also calculated by UV−visible spectroscopy (Figure S3 in the Supporting Information) to be 5.0 (± 0.2) per chain using the following equation: nRAFT = [RAFT]0/[polymer]0, where [RAFT]0 is calculated by Beer−Lambert’s law, i.e. Abs305nm/(εRAFT × l), where Abs305nm, εRAFT, and l correspond to the absorbance of RAFT agent at 305 nm, the molar extinction coefficient of the RAFT group (15000 L mol−1 cm−1),24 and the path length; [polymer]0 stands for polymer concentration. This value is in good agreement with the value obtained by 1H NMR spectroscopy. It is important to underline that the number of RAFT agents can be tuned according to the experimental conditions. In this current example, we used a dextran with around 5.0 RAFT agents per chain. Polymerization of OEGMA. The Ac-dextran macroRAFT agent was subsequently employed to control the polymerization of POEGMA in DMAc at 70 °C. The conversion of the monomer was determined using 1H NMR spectroscopy,

Figure 4. RAFT polymerization of OEGMA using Ac-dextran macroRAFT in DMAc at 70 °C: (A) Monomer conversion at different time intervals and a pseudo-first-order kinetic plot for the copolymerization; (B) molecular weight and PDI evolution versus monomer conversion; (C) W(log M) versus log M of Ac-dextran-gPOEGMA at 23, 58, 76, and 89% OEGMA monomer conversions.

increased with reaction time and the radical concentration was constant during the course of polymerization, as evidenced by a pseudo-first-order kinetic plot. The experimental molecular weight increased linearly with monomer conversion during the course of polymerization, indicating a well-controlled RAFT process (Figure 4B). However, the experimental molecular weights appear lower than the theoretical values, calculated by the following equation: Mn,theoretical = MnAc‑dextran + MWOEGMA × [OEGMA]0/[Ac-dextran]0, with MnAc‑dextran, [OEGMA]0, and [Ac-dextran]0 corresponding to the molecular weight of 3053



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POEGMA copolymers were dissolved in DMF, and an aqueous phosphate buffer solution was slowly added followed by the dialysis against water to remove DMF. The morphology of selfassembled aggregates was then investigated using both DLS and TEM. The relative ratio of hydrophobic and hydrophilic lengths had a significant influence on the measured hydrodynamic diameters (see Table S2 in the Supporting Information). As shown in Table 1, the hydrodynamic diameter increased

acetalated dextran and to the initial concentrations of OEGMA and [Ac-dextran], respectively. This molecular weight anomaly can be explained by the qualitative calibration of the SEC system, with linear polystyrene standards failing to account for the different hydrodynamic radii of the grafted copolymers, and by the difference of polymer nature. SEC traces are presented in Figure 4C, revealing the clear shift of polymer peaks toward higher molecular weight with increasing conversion when compared to the SEC trace of the dextran macroRAFT. As expected, the polydispersity index (PDI) of the dextran-gPOEGMA is relatively high (around 1.40), following the broad PDI of the initial macroRAFT dextran agent. FTIR-ATR was employed to analyze the different polymers at different stages of structure development: dextran polymer (Figure 5A), acetalated dextran (Figure 5B), Ac-dextran

Table 1. Structure Obtained of Ac-dextran-g-POEGMA in Aqueous Mediaa reaction time (h)

monomer conversion (%)b

ratio of hydrophobic/ hydrophilic (%)c

0 2 6 9

0 23 58 76

100 68 45 38

14.5

89

34

structure not soluble small vesicle rodlike sperical nanoparticles sperical nanoparticles

diameter (nm)d 30 90 45 50

a Note: f1 and f 2 correspond to the proportion of DOX loaded nanoparticles and free DOX in the SY-5Y cells. bThe monomer conversion was calculated from the 1H NMR of the reaction mixture by comparing the intensity of vinyl proton peaks (at ca. 6.1 and 5.6 ppm) to that of aliphatic proton peaks (at ca.4.2 ppm); cRatio of hydrophobic/hydrophilic calculated by the following equation: % = mass of Ac-dextran/[mass of Ac-dextran + (OEGMA conversion × mass of OEGMA)] × 100. dHydrodynamic diameter (number size) of self-assembled Ac-dextran-g-POEGMA determined by DLS analysis (concentration of copolymer: 1 mg mL−1).

initially with an increase in hydrophilic side chain (POEGMA) length. It is generally accepted that a longer hydrophobic block length compared to the hydrophilic chain length usually leads to the formation of vesicles and worm-like rods as the results of the stretching of the core-forming blocks and a repulsive interaction among corona chains.26 Extending the length of the hydrophilic chain (or increasing the ratio hydrophilic versus hydrophobic) favors a transition of assembled morphology to nanoparticles. As the hydrophilic POEGMA side chain length increased, the hydrodynamic diameter decreased (as expected). Data on hydrodynamic radii alone is insufficient for structural characterization, as information on specific morphology is also needed to fully understand the self-assembly process. A morphology study using TEM is therefore necessary to complement the DLS studies. The self-assembly of amphiphilic copolymers, such as block copolymers27 and graft copolymers28 into vesicles, wormlike structures, and nanoparticles, has been reported many times previously. In this present work, we hoped to achieve a similar library of morphologies from Ac-dextran-gPOEGMA copolymers in aqueous media simply by modifying the length of the hydrophilic side chain block. As can be seen from Figure 6, the change in particle sizes (measured by DLS) is associated with morphology transitions. DLS confirms the synthesis of well-defined nanoparticles with a relative low dispersity for polymeric nanoparticles (∼0.2). The dispersity values (determined by DLS) obtained for these nanoparticles are in the range of dispersity obtained with the polymeric nanoparticles in previous studies.3g,i,m,8b,29 As the hydrophilic chain length increased, the type of self-assembled aggregate changed from small vesicles (Figure 6A) to wormlike (Figure 6B) and to spherical nanoparticles (Figure 6C and D). A TEM

Figure 5. FTIR-ATR spectra of (A) dextran, (B) Ac-dextran, (C) Acdextran macroRAFT, and (D) Ac-dextran-g POEGMA (14.5 h copolymerization).

macroRAFT (Figure 5C), and Ac-dextran-g-POEGMA (Figure 5D). The key feature to note is the significant decrease of the −OH peak at around 3400 cm−1, indicating the extent of reaction of the hydroxyl groups during the acetalation process (Figure 5B). The IR spectrum of Ac-dextran macroRAFT showed the appearance of expected peaks, including a nitrile peak at around 2100 cm−1 and an aromatic peak at around 3000 cm−1 (Figure 5C). Figure 5D displays the FTIR spectra of purified dextran-g-POEGMA copolymers after 14.5 h of polymerization time. The appearance of a high intensity carbonyl absorption at around 1730 cm−1 is indicative of poly(OEG-MA) segments. Self-assembly Ac-dextran-g-POEGMA Copolymers in Aqueous Media. Eisenberg et al. have done extensive research on the factors that influence the morphologies of self-assembled aggregates to give nanoparticles, vesicles, or rodlike structures and so on. As well as the absolute and relative block lengths, the water content of the solvent mixture, and the nature and the presence of additives (ions, homopolymers, and surfactants), it was also found that micellization is, to a significant extent, controlled by the ratio length of the hydrophobic and hydrophilic blocks.25 The Ac-dextran-g-POEGMA copolymers, where the length of the hydrophobic acetalated dextran backbone was fixed and the length of the OEGMA hydrophilic block was varied by stopping the polymerizations after 2, 6, 9, and 14.5 h, were self-assembled in water. Ac-dextran-g3054



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Figure 6. TEM analyses of the self-assembled aggregates (concentration of polymer 1 mg mL−1 in water) of Ac-dextran-g-POEGMA after (A) 2 h, (B) 6 h, (C) 9 h, and (D) 14.5 h of copolymerization of OEGMA, scale bar 200 nm.

Figure 7. AFM micrograph of self-assembled nanoparticles of Ac-dextran-g-POEGMA (14.5 h of copolymerization of OEGMA) (left) (scale bar =100 nm) and TEM image of DOX-loaded Ac-dextran-g-POEGMA nanoparticles (right) (scale bar is 1 μm).

S13 in the Supporting Information), the acetal groups are very stable (less than 5% of degradation in 72 h). In a second step, Ac-dextran-g-POEGMA (Mn = 21 000 g/ mol, PDI = 1.47) obtained after 14.5 h of OEGMA was investigated for the physical encapsulation of DOX. We used this structure because it offers a higher drug loading compared to the other structures. DOX was loaded into Ac-dextran-gPOEGMA nanoparticles using a codialysis method. A grafted copolymer with Mn = 2 000 g mol−1 and PDI = 1.47, obtained after 14.5 h of OEGMA polymerization, was used for DOX loading experiments. TEM analysis indicated that the particle size of Ac-dextran-g-POEGMA spherical nanoparticles was about 50 nm (Figure 6D), consistent with atomic force microscopy (Figure 7A). Copolymer and hydrophobic DOX were dissolved in DMSO, a good solvent for both copolymer and drug. PBS (pH 7.4) was added dropwise, and the mixture was dialyzed against a PBS and DMF mixture (9:1, v/v) for 1 day and then PBS solution only for 2 days. It is noteworthy that acetal groups, at pH 7.4, are relatively stable (less than 5% of acetal groups degraded over 48 h, as confirmed by 1H NMR spectroscopy). A slight change in hydrodynamic volume was observed during dialysis; the nanoparticle size decreased from 110 nm to ∼90 nm. In addition, during the codialysis of the polymer and DOX, the hydrophobic interactions are present among acetal groups in the acetalated dextran backbone and also among DOX and acetal groups influencing DOX loading efficiency. The drug content was 5.1 wt % (±0.2%), as measured by UV−visible analysis (Figures S6 and S7 in the Supporting Information). When DOX was encapsulated in the Ac-dextran-g-POEGMA nanoparticles, the spherical nanoparticle morphology was retained and the particle size of DOX-loaded Ac-dextran-g-POEGMA nanoparticles (about 90 nm, determined by TEM (Figure 7B)) was significantly larger than that of unloaded Ac-dextran-g-POEGMA (with a size 50

micrograph showing small vesicles under different magnifications can be seen in Figure S4 in the Supporting Information. For a full description of the spectrum of structures possible, the reader is referred to the work of Eisenberg et al.25−27,30,31 It should be noted that the vesicle sizes were rather small, possibly attributable to the PDI of the grafted copolymer originating from the relatively broad distribution of dextran macroRAFT. Terreau et al. investigated the effect of the PAA block PDI on the aggregate morphology for a series of PS-b-PAA copolymers and found that the vesicle sizes generally decreased as the PAA PDI increased.30 Luo and Eisenberg suggested that a decrease in vesicle size originates from a segregation of the long chains preferentially to the vesicle exterior whislt the short chains segregate to the vesicle interior.31 The stability of the nanoparticles was investigated using a serum media to mimic biological conditions. The nanoparticles were found to stay perfectly dispersed and stable in serum, as indicated by visual inspection and by DLS measurement (data not shown). DLS did not reveal an increase of the size after 8 h, confirming that the POEGMA confers antifouling characteristics on the nanostructures. DOX Loading. In the following part, we focused only on the use of the micellar structure formed from Ac-dextran-gPOEGMA (Mn = 21000 g/mol, PDI = 1.47) obtained after 14.5 h of OEGMA polymerization for the drug delivery study. First, we monitored the acetal degradation of the polymer at pH 7.4 and 5.0 using 1H NMR spectroscopy. In this experiment, we followed the release of acetone (signal at 2.10 ppm) and methanol (signal at 3.5 ppm) obtained by the degradation of cyclic and acyclic acetals. The kinetics of degradation was in good accord with the data reported by Fréchet and coworkers.11a The half-life time of acetal groups was estimated to be 72 h at pH 5.0 in our experiment, while, at pH 7.4 (Figure 3055



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nanoparticles and the amount of drug retained in the nanoparticles. As can be seen in Figure 8, concurrently with the release of DOX, the nanoparticles dissociate quickly at low pH (within 2 days), while they are relatively stable at pH 7, where the formation of unimers only occurred after nearly one week. DOX released from the nanoparticles was analyzed by fluorescence spectroscopy and mass spectroscopy (Figures S8 and S9 in the Supporting Information) to confirm the structural integrity of doxorubicin. Electrospray mass spectroscopy confirmed that the molar mass of doxorubicin was not changed (theoretical mass (DOX/H+) 544.52, experimental mass (DOX/H+) 544.18, Figure S9 in the Supporting Information). In addition, a fluorescence study confirmed the characteristic peak of doxorubicin after release from the nanoparticles (data not shown), corresponding exactly to the initial spectrum of doxorubicin (before encapsulation). The degradation of doxorubicin can be monitored by fluorescence spectroscopy due to the appearance of new absorption peaks and by mass spectroscopy.33 In-Vitro Uptake and Toxicity of Doxorubicin Loaded Dextran Nanoparticles. DOX accumulation in the cells was examined over time via flow cytometry (Figure S10 in the Supporting Information) and confocal laser scanning microscopy (CLSM) (Figure 9) using the SY-5Y neuroblastoma cell line using the characteristic fluorescent emission of DOX at 595 nm. DOX accumulation in the cell can be seen following 30 min incubation with the DOX-dextran nanoparticles, as indicated by an increase in fluorescence intensity versus the nontreated cells with DOX-dextran nanoparticles by flow cytometry. DOX accumulation continues to increase over a 24 h period, as indicated by an increase in fluorescence intensity over time. CLSM was also exploited to confirm the accumulation of DOX from DOX-loaded dextran nanoparticles. As can be seen from Figure 9, cells treated by DOX and DOX loaded nanoparticles show an accumulation of DOX in the cell nuclei of SY-5Y neuroblastoma cells (Figures 9B and C) when compared to control cells (Figure 9A) after 6 h of incubation. It is important that DOX-loaded dextran nanoparticles can accumulate in the cell nucleus. Indeed, one of the wellknown toxicity mechanisms of DOX is attributed to its intercalation with DNA and inhibition of macromolecular biosynthesis. Both flow cytometry and CLSM analyses confirm that the DOX accumulates in the SY-5Ycells. However, both techniques (flow cytometry and CLSM) do not unequivocally confirm the cell uptake of DOX-dextran nanoparticles. To investigate the cell uptake, we used fluorescence lifetime imaging microscopy (FLIM) and phasor analysis. To achieve this, it is desired to differentiate between encapsulated DOX in the nanoparticles and free DOX inside the live cells. With conventional intensity based microscopic techniques, it is not possible to distinguish these two forms of DOX, as their spectral profiles are not significantly different (Figure S11 in the Supporting Information). Fluorescence lifetime imaging microscopy (FLIM) and phasor analysis are useful tools for differentiating between polymer-mediated and free drugs inside living cells.34 Phasor analysis presents the fluorescence lifetime data in a graphical form, which negates the use of exponential fitting to the fluorescence decay.35 The application of phasor plotting was originally developed to overcome some of the drawbacks of FLIM, such as the low photon counts per pixel which renders differentiating between one and two lifetimes difficult.36 The potential application of

nm) (Figure 6D). This result was consistent with DLS analysis (Figure S5 in the Supporting Information). The fluorescence of doxorubicin (after loading) was determined, confirming that the structure of DOX remained unaltered during the encapsulation process (Figure S8 in the Supporting Information). Finally, mass spectroscopy confirmed the DOX structure after encapsulation (Figure S9 in the Supporting Information). Nanoparticle Degradation Studies. An important feature of this drug delivery system is a built-in sensitivity to pH, originating from the acetal groups on the dextran, that are known to hydrolyze at low pH values, converting back to native hydrophilic dextran, inducing micellar dissociation. This is significant for drug delivery applications, as the targeted disease sites are often moderately acidic, for example tumor cells and lysosomal compartments.32 The degradation of the acetal groups and subsequent dissociation of nanoparticles will trigger release of encapsulated drug and subsequent excretion of the small polymer residues of the carrier. Nanoparticle dissociation and release of DOX were monitored over time under two different conditions, pH 7.4 and pH 5.0, using fluorescence spectroscopy (calibrated using free DOX). As predicted, the release rate of DOX is higher in mildly acidic media (Figure 8). After 3 days, the nanoparticles

Figure 8. Release profile of DOX as evaluated by fluorescence spectroscopy at pH 5.0 (open square) and pH 7.4 (close square) and the time-dependent change in the hydrodynamic diameter of the nanoparticles in water at pH 5.0 (open star) and pH 7.4 (close star) (nanoparticles concentration 4 mg mL−1). Experiments were done in triplicate.

were completely disassembled (size around 10 nm). However, only 60% of DOX was found to be released. This release efficiency was attributed to the presence of a hydrophobic domain in the polymer due to the partial hydrolysis of acetal groups. Indeed, only 45% of acetal groups (i.e., 30% and 90% of cyclic and acyclic acetal groups) were hydrolyzed. After one week, over 80% of the drug was released at pH 5.0 (corresponding to 85% of the degradation of acetals; see Figure S13 in the Supporting Information); however, only about 40% released at pH 7.4 and no initial burst of drug was observed. These results confirm that DOX loaded Ac-dextran-gPOEGMA nanoparticles are more stable at physiological pH than endosomal pH. The amount of DOX released was also determined by the measurement of the residual drug in the nanoparticles at each sampling point by spectrophotometry measurement at λmax = 475 nm of an aliquot taken from inside the dialysis bag, as a cross-check. The amount of drug released was calculated from the amount of drug initially present in the 3056



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Figure 9. Confocal laser scanning microscopy showing cell uptake of doxorubicin using SY-5Y cells after 6 h of incubation: (A) control experimental (nanoparticles without doxorubicin); (B) free doxorubicin; (C) doxorubicin loaded dextran nanoparticles. Note: DOX f luorescent images were acquired at λex = 485 nm and λem = 595 nm. (Scale bar corresponds to 10 μm.)

the FLIM technique for study of cellular uptake behavior has been demonstrated in previous publications.34,37 First, the fluorescence lifetimes of free DOX and DOXloaded nanoparticles in aqueous solution were measured using time-correlated single photon counting (TCSPC). The concentration-independent single exponential decay corresponded to a lifetime of 1.0 ns for free DOX, which is consistent with a published lifetime value of free DOX,34 and a lifetime of 3.5 ns for DOX encapsulated in nanoparticles. The increase in DOX lifetime was also observed when DOX was encapsulated in other polymeric nanoparticles.34,37 The fluorescence lifetimes of free DOX and DOX-loaded nanoparticles are substantially different, allowing the differentiation between these two species in solution. FLIM was then performed using DOX and DOX-loaded nanoparticles at concentrations of 0.5 μM (equivalent DOX concentration) in SY-5Y neuroblastoma live cells. DOX and DOX-loaded nanoparticles present two different lifetimes (around 1 and 4.0 ns) in live cells, allowing us to distinguish these two forms of DOX in the cells. FLIM images were recorded at 3 h (region of interest (ROI) (A)) and 24 h (ROI) (B) after incubation of DOX and DOX-loaded nanoparticles in live cells. As can be seen in Figure 10, the ROI (A) includes longer lifetime DOX-loaded nanoparticles and released DOX, and it is clearly evident that DOX is released from the DOXloaded nanoparticles over time as the change in the phasor plot of ROI (A) and (B) toward free doxorubicin (ROI (C)) was observed. Using FLIM, we are able to estimate, after 3 h, that around 35% of doxorubicin has been released from the nanoparticles in the cells, while, after 24, and over 55% of the doxorubicin has been released from dextran nanoparticles (Table 2). As a control experiment, the lifetime of DOX in the SY-5Y cells was measured. A slight decrease of the DOX lifetime was observed due to the interactions of DOX with the

Figure 10. Region of interest (ROI) examined (up) and the corresponding phasor plot of DOX-dextran nanoparticles (bottom) after 3 h (ROI A) and 24 h (ROI B) in the SY-5Y cells. Note: Free doxorubicin (ROI C) after 3 and 24 h in the SY-5Y cells added as a reference.

cellular environment (from 1.0 to 0.9 ns). The release of DOX from the nanoparticles determined by FLIM is relatively close to the values determined in the model reaction at pH 5.5. FLIM analysis confirms the efficient cell-uptake and subsequent 3057



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Table 2. Lifetime of DOX in the Cells under Different Forms (DOX and DOX-dextran Nanoparticles): τ1 (ns) and τ2 (ns) Correspond to the Lifetimes of DOX-Dextran Nanoparticles and Free DOX and, Respectively DOX-dextran 3 h DOX-dextran 24 h DOX 3 h DOX 24 h

τ1 (ns)

f1

τ2 (ns)

f2

3.4 3.3

0.65 0.45

1.0 0.9 1.0 0.9

0.35 0.55 100 100

Table 3. Comparison of IC50 (μM) Values for Doxorubicin and Doxorubicin Loaded Particles in SY-5Y Neuroblastoma Cells and MRC5 Normal Fibroblast Cells doxorubicin doxorubicin loaded nanoparticles

SY-5Y

MRC5

0.036 (± 0.004) 0.178 (± 0.002)

0.298 (± 0.03) 5.28 (± 0.52)

(demonstrated by FLIM) and can effectively deliver their drug load to SY-5Y neuroblastoma cell nuclei as demonstrated by CLSM.

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doxorubicin release from the DOX loaded dextran nanoparticles in the SY-5Y cells. The toxicities of dextran nanoparticles and dextran nanoparticles containing doxorubicin were assessed using the SY-5Y neuroblastoma cancer cell line and normal MRC5 lung fibroblasts (Figure 11). The nanoparticles alone were nontoxic at all concentrations examined after 72 h, proving that the hydrolysis products from the acetal groups (methanol and acetone) had no observable cytotoxic effect, and any future influence on cell viability from DOX loaded nanoparticles could not be attributed to the polymeric nanoparticles alone (Figure S12 in the Supporting Information). The normal MRC5 cells were less susceptible than the SY-5Y cells to both free and encapsulated doxorubicin. IC50 values for the MRC5 were ∼8fold higher for free doxorubicin and ∼30-fold higher for the encapsulated doxorubicin than those for SY-5Y cells (Table 3). In the case of SY-5Y neuroblastoma cells, the IC50 for free DOX was only ∼5-fold lower (36 nM) than that for the encapsulated DOX (178 nM), while there was a ∼ 17-fold difference for the MRC5. Thus, while both cell types show less sensitivity to the encapsulated doxorubicin, the difference is more pronounced for the normal MRC5 cells. A reduction of drug toxicity encapsulated in polymeric nanoparticles has been previously reported in other studies. It has been found, for example, that a 10−20-fold reduction exists in in vitro toxicity of encapsulated doxorubicin versus the free drug.38 Different explanations have been suggested, such as changes in a drug’s distribution in the cells resulting from a significant decrease of drug activity38b or from the drug encapsulation.39 In the case of DOX-HCl, DOX HCl is converted to its basic form, which is known to be chemically unstable, inducing drug degradation, possibly resulting in an increase in the IC50 value, as seen for the DOX-dextran nanoparticles.39 In summary, despite some requirement for optimization, our studies have demonstrated that doxorubicin loaded dextran nanoparticles are taken up

CONCLUSIONS Dextran was effectively acetalated, inducing a change from hydrophilic to hydrophobic chains. Solid state NMR proved a very useful and reproducible technique to characterize the acetalation of dextran, determining the exact proportion of acyclic and cyclic acetal functionalization. Subsequent modification of acetalated dextran to form a macroRAFTagent enabled control to be exerted over the structure of a comblike graft copolymer of acetalated dextran and POEGMA. The acyclic and cyclic acetal groups on the dextran backbone functioned as a hydrophobic segment that together with the hydrophilic PEG segment provided a driving force for selfassembly to nanoparticles in aqueous media. The nanoparticles displayed a range of morphologies, and they could be used to load and deliver doxorubicin (DOX). DOX loaded Ac-dextrang-POEGMA showed a more pronounced cytotoxicity to a SY5Y neuroblastoma cancer cell line than a MRC5 fibroblast lung normal cell line. This approach, described herein, to nanoparticle formation for drug delivery is extremely powerful and versatile, as the structures are inherently nonfouling and biodegradable and can be triggered at known disease sites. The system is highly tunable and can be adapted to different requirements. We also demonstrate the application of FLIM as a powerful technique in drug delivery research in combination with an intensity-based imaging method, facilitating studies on the uptake and release of DOX from dextran-based nanoparticles in live cells. We are currently exploiting this nanoparticle system in vivo for the treatment of neuroblastoma cancer.

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ASSOCIATED CONTENT

S Supporting Information *

1 H NMR spectrum of the Ac-dextran macroRAFT agent, UV− visible specturm of the Ac-dextran macroRAFT agent,

Figure 11. Toxicity of doxorubicin (black squares) and doxorubicin loaded dextran nanoparticles (blue triangles) in SY-5Y neuroblastoma cells (A) and MRC5 normal fibroblast cells (B), done in triplicate for each sample. 3058



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determination of doxorubicin loading into the nanoparticles, mass spectroscopy, fluorescence spectroscopy of doxorubicin, Scheme S1, Figures S1−S12, and Tables S1 and S2. This material is available free of charge via the Internet at http:// pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*Australian Centre for NanoMedicine, The University of New South Wales, Sydney, NSW 2052, Australia. C.B.: e-mail, [email protected]. T.P.D.: e-mail, [email protected]. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS All the authors are grateful for UNSW’s funding and Australian Research Council (ARC) funding. C.B. is thankful for his APD fellowship from ARC (DP1092640). This work was supported by the Children’s Cancer Institute Australia for Medical Research, which is affiliated with the UNSW and Sydney Children’s Hospital. M.K. is supported by a NHMRC Senior Research Fellowship. We acknowledge the DVCR, Professor Les Field, at UNSW, for significant strategic funding to set up the Australian Centre for Nanomedicine.

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REFERENCES

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