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Biodegradable Nanoparticles Based on Linoleic Acid and Poly(β-malic acid) Double Grafted Chitosan Derivatives as Carriers of Anticancer Drugs Ziming Zhao,†,‡ Miao He,†,‡ Lichen Yin,† Jiamin Bao,‡ Lili Shi,†,‡ Bingqing Wang,†,‡ Cui Tang,† and Chunhua Yin*,†,‡ State Key Laboratory of Genetic Engineering, Department of Pharmaceutical Sciences, School of Life Sciences, Fudan University, Shanghai 200433, China, and Department of Biochemistry, School of Life Sciences, Fudan University, Shanghai 200433, China Received October 27, 2008; Revised Manuscript Received December 18, 2008
Novel chitosan derivatives carrying linoleic acid (LA) as hydrophobic moieties and poly(β-malic acid) (PMLA) as hydrophilic moieties (LA/PMLA double grafted chitosan, LMC) were synthesized. It self-assembled into nanoparticles of 190-350 nm in water, which carried negative surface charges in physiological pH. The critical aggregation concentration of the LMC deceased with an increase in the LA content. Paclitaxel (PTX) was loaded into the LMC nanoparticles with a high loading efficiency and the maximum loading capacity of 9.9 ( 0.4%. PTX-LMC nanoparticles exhibited a sustained release within 24 h in pH 7.4 phosphate-buffered saline (PBS), and the release rate was affected by the LA content and PMLA length. Hemolysis and acute toxicity assessment indicated that the LMC nanoparticles were safe drug carriers for i.v. administration. Additionally, PTX-LMC showed significantly potent tumor inhibition efficacy relative to that of TAXOL in S-180 bearing mice. Therefore, the LMC nanoparticles could be an effective and safe vehicle for systemic administration of hydrophobic drugs, especially PTX.
Introduction Over the past few years, great effort has been made to develop novel polymeric nanoparticles as desirable drug delivery systems (DDSs) for their attractive characteristics, such as longer circulation time, targeted drug delivery, protection from enzymatic degradation, and reduced drug toxicity or side effects.1 Research on polymeric nanoparticles has been mainly focused on amphiphilic block copolymers2,3 and hydrophobically modified water-soluble polymers,4 which can self-assemble to form hydrophobic core and hydrophilic shell structure when dissolved in water due to the intra- or intermolecular hydrophobic interactions. The hydrophobic domain at the same time can serve as a preservatory for hydrophobic drugs and protect them from the aqueous environment. As drug carriers, the safety and potential accumulation of these polymeric nanoparticles in the body must be taken into account. Therefore, a number of nontoxic and biodegradable polymers, both synthetic and natural, have been utilized in formulating polymeric nanoparticles including polyesters (such as poly(lactic acid) (PLA),5 poly(lactic-co-glycolic acid) (PLGA),6 poly(L-caprolactone) (PCL)7), poly(amino acids) (such as poly(γ-glutamic acid),8 poly(Laspartic acid),9 poly(L-lysine),10 and poly(L-ornithine)11), poly(anhydrides),12 poly(orthoesters), and polyphosphazene derivatives.13 Natural polymers such as albumin,14 collagen, gliadin,15 gelatin, chitosan, pullulan,16 and Curdlan4 have also been used. In particular, chitosan has been widely applied due to its structural and physicochemical properties. It shows satisfactory biocompatibility, biodegradability, and low immunogenicity,17 which are beneficial attributes for an ideal drug delivery * Corresponding author. Tel: +86-21-65643797. Fax: +86-21-55522771. E-mail address:
[email protected]. † Department of Pharmaceutical Sciences. ‡ Department of Biochemistry.
material.18,19 However, the extended applications of chitosan are limited because of its insolubility in physiological solution (pH 7.4) and lack of amphiphilicity, which prohibits it from forming micelles in water. Therefore, a lot of chitosan derivatives have been developed to be more appropriate for drug delivery carriers. Calvo et al.20 reported the formation of hydrophilic chitosan-polyethylene oxide nanoparticles and studied their potential application as protein carriers. Miwa et al.21 synthesized a novel chitosan derivative with lauryl groups attached to amino groups as the hydrophobic moieties and carboxymethyl groups to hydroxyl groups as the hydrophilic moieties (N-lauryl-carboxymethyl-chitosan, LCC). Kwon et al.22 prepared hydrophobically modified glycol chitosans (HGCs) by covalent attachment of 5β-cholanic acid to glycol chitosan. Zhang et al.23 synthesized a series of novel chitosan derivatives carrying long chain alkyl groups (n ) 8, 10, 12) as hydrophobic moieties and sulfated groups as hydrophilic moieties. Park et al.24 reported N-acetyl histidine-conjugated glycol chitosan (NAcHis-GC) self-assembled nanoparticles as a promising system for intracytoplasmic delivery of drugs. Wang et al.25 synthesized a cholesterol-modified chitosan conjugate (CHCS) with succinyl linkage. Poly(β-malic acid) (PMLA) is well-known as a water-soluble and biodegradable polymer26 that can be degraded into malic acid, an intermediate of tricarboxylic acid cycle. It also bears lateral carboxyl groups, which allow further conjugation of biologically active molecules. Cammas et al.27 synthesized degradable macromolecular micelles based on amphiphilic block copolymers of PMLA as hydrophilic units and poly(β-malic acid alkyl esters) as hydrophobic blocks, and their properties and stability were proven to be pH-dependent. Lee et al.28 designed a new nanoconjugate, Polycefin, where PMLA was used as the scaffold, and the drug or functional modules (such as targeting
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agent, degradation protecting module, and fluorescent agent) were conjugated to the pendant carboxyl groups of PMLA. Although chemotherapy is one of the major cancer therapeutic methods in clinical practice, there lies a serious problem in that most anticancer drugs have no specific efficacies to tumors, and normal cells (especially bone marrow cells and endothelial cells) can also be killed. At the same time, water insolubility, biological barriers in body, and toxicity induced by high dosage also limit the application of many anticancer drugs. Nanoparticle DDSs offer major improvements in therapeutics through their solubilization capacity, the ability to bypass biological barriers, and sustained release. More importantly, targeted drug delivery can be achieved by surface modification or introduction of targeting ligands. Therefore, nanoparticle DDSs have wide application prospects in cancer therapy. The aim of this study was to develop a novel polymeric nanoparticle system that had good biodegradability and could provide modification sites on the surface for the introduction of targeting ligands. Linoleic acid (LA) and PMLA double grafted chitosan derivatives (LMCs) were synthesized, where chitosan acted as the backbone, LA as the hydrophobic moiety, and PMLA as the hydrophilic moiety. It was assumed that LMCs would exhibit desirable biodegradability in that the ester or amide linkage between chitosan and LA/PMLA could be easily hydrolyzed by lipase and amidase while the glycoside linkage could be degraded by glycosidase. Moreover, LA is an essential fatty acid involved in human fatty acid metabolism, and PMLA is biodegradable as mentioned above. The lateral carboxyls and residual aminos provided abundant sites that could be modified. The physicochemical properties of LMCs were characterized by Fourier transform infrared (FTIR) spectroscopy, 1H NMR and X-ray diffraction (XRD). LMC nanoparticles were prepared by a sonication method and were characterized by dynamic light scattering (DLS), transmission electron microscopy (TEM), scanning electron microscopy (SEM), and fluorescence spectroscopy. As a model hydrophobic drug, paclitaxel (PTX) was loaded into the LMC nanoparticles; the loading capacity as well as the loading efficiency were determined, and in vitro release profiles were studied. Hemolysis and acute toxicity assessment were performed to evaluate the safety of LMCs. Finally, the comparable antitumor efficacies of PTX-LMC and TAXOL were monitored in Sarcoma180 (S-180) bearing mice.
Experimental Section Materials. Chitosan (deacetylation degree of 85% and MW of 105 000 Da) was purchased from Golden-shell Biochemical Co. Ltd. (China). LA (BR) and D,L-aspartic acid (CP) were purchased from Sinopharm Chemical Reagent Co. Ltd. (China). Trifluoroacetic acid (TFA) was supplied by Sigma (USA). All other reagents were of analytical grade. Kunming mice (6 weeks old, body weight 18-22 g) were obtained from the Animal Centre of Fudan University, and raised under normal conditions with free access to food and water. Animal experiments were performed according to the Guiding Principles for the Care and Use of Experiment Animals in Fudan University. Synthesis of LMC. Benzyl malolactonate was synthesized from D,Laspartic acid as described before.29,30 Synthesis of poly(β-benzyl malate) (PMLABz) was as follows. Lactic acid was dried under vacuum for 24 h at room temperature (RT) and 0.18 g of it was added into a flask as the initiator. TFA (0.02 g) was added as the catalyst. Benzyl malolactonate (4.12 g) was kept under N2 stream for 2 h and then transferred under N2 to the previous flask. The polymerization was conducted at 80 °C for 3 days. TFA was eliminated by vacuum distillation after polymerization, and PMLABz1 (lactic acid-to-lactone molar ratio of 1:10) was obtained. PMLABz2 (lactic acid-to-lactone
Zhao et al. molar ratio of 1:20) was obtained when 0.09 g of lactic acid was used for polymerization. Linoleic chloride was synthesized as follows. LA was dissolved in dichloromethane into which 1.5 equiv of oxalyl chloride was smoothly added in the ice bath. The mixture was stirred at 40 °C under N2 for 6 h. Excess oxalyl chloride and dichloromethane were eliminated by vacuum distillation, and linoleic chloride was obtained. PMLABz acyl chloride was obtained using the same method. Synthesis of LMC was as follows. Chitosan (1.67 g) was dissolved in 20 mL of MeSO3H into which 2.99 g of linoleic chloride (0.9 equiv/ sugar unit of chitosan) was added. After stirring at RT for 2 h, PMLABz1 acyl chloride (2.15 g) was added. The mixture was stirred for 4 h at RT before 30 g of crushed ice was added to stop the acylation reaction. The acidic mixture was stirred at RT for 0.5 h and dialyzed (molecular weight cutoff (MWCO) 3600 Da) for 24 h to remove most of the acid, followed by adjusting the pH to 7.4 with NaHCO3. The mixture was then filtered to remove insoluble matter and dialyzed (MWCO 3600 Da) again for more than 3 days. Finally the pH of dialysis solution was adjusted to 6.0 and allowed to precipitate for 10 min at 4 °C. After filtration, the precipitate was washed twice with water and twice with acetone and dried in vacuum, and LMC1 was obtained. LMC2 was obtained when 1.99 g of linoleic chloride (0.6 equiv/ sugar unit of chitosan) was used. LMC3 was obtained when 0.99 g of linoleic chloride (0.3 equiv/sugar unit of chitosan) was used. When PMLABz1 acyl chloride was replaced by PMLABz2 acyl chloride, LMC4, 5, and 6 were obtained using the same method as LMC1, 2, and 3, respectively (Figure 1). Characterization of LMC. 1H NMR measurements were performed on an AVANCE DMX 500 spectrometer (Bruker, Germany). Chitosan was dissolved in the mixed solvent of D2O and F3CCOOD, and chitosan derivatives were dissolved in the mixed solvent of DMSO-d6 and F3CCOOD. The internal reference was tetramethylsilane (TMS). IR spectra were recorded on a Nexus 470 FTIR spectrometer (Nicolet, USA) in KBr discs. XRD spectra were obtained using a D/max-γB multicrystal diffraction meter (Rigaku, Japan) with Cu KR radiation in the range of 5-40 °(2θ) at 40 kV and 60 mA. Preparation of LMC Nanoparticles. Nanoparticles were prepared by a sonication method. LMCs (10 mg) were dissolved in 10 mL of water, which was then sonicated using a probe-type sonifier (JY92-R, Scientz Biotechnology, China) at 100 W for 10 min in an ice bath. The pulse was turned off for 2 s with the interval of 5 s to avoid increase in temperature. The resulting solution was filtered through a 0.8 µm membrane, and a LMC nanoparticle solution was obtained. DLS Measurement. The particle size and zeta potential of the nanoparticles were determined by DLS (Nicomp380/ZLS, Santa Barbara, CA). The particle size measurements were performed at a wavelength of 635 nm and a scattering angle of 90° at 23 °C. The zeta potential measurements were monitored at a wavelength of 635 nm and a scattering angle of 14° at 23 °C. Sample concentration was kept at 1.0 mg/mL. Morphology. The morphological examination of LMC nanoparticles was conducted by TEM (H-600A, Hitachi, Japan). The samples were placed on copper grids and dehydrated for TEM observation. The morphology of LMC nanoparticles was also observed by SEM (XL30, Philips-FEI, Netherlands) at 20 kV after dehydration, desiccation, fixing, and gold spraying. Fluorescence Spectroscopy. The critical aggregation concentrations (CACs) of the LMC nanoparticles were determined by fluorescence spectroscopy (Cary Eclipse, Varian, USA) with pyrene as a hydrophobic probe. A pyrene solution (6.0 × 10-4 M) was prepared in acetone and stored at 4 °C until use. The pyrene solution was added to 5 mL volumetric flasks and evaporated to dryness under nitrogen. Then 5 mL of LMC solutions with various concentrations (1 × 10-4 to 0.1 mg/mL) were added, which were then sonicated for 1 h before measurement of steady-state fluorescence spectra. Fluorescence excita-
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Figure 1. Reaction scheme of LMC derivatives.
tion spectra were recorded at the emission wavelength (λem) of 395 nm and excitation and emission bandwidths at 5 nm. Drug Loading and In Vitro Release. LMC (50 mg) was dissolved in 10 mL of 0.15 M phosphate-buffered saline (PBS) (pH 7.4), and 1 mL of PTX solution in ethanol (10 mg/mL) was added. The mixture was then sonicated using a probe-type sonifier at 100 W for 10 min in an ice bath. To remove ethanol, the solution was dialyzed (MWCO 3600 Da) against 0.15 M PBS (pH 7.4) overnight, and the nanoparticle solution was filtered through a 0.8 µm membrane to remove insoluble PTX. The PTX-loaded LMC nanoparticle solution was loaded onto the silica column (50-100 mesh) and eluted with 0.15 M PBS (pH 7.4). The fraction of LMC nanoparticles was collected, and its volume (V1) was measured. Then, the column was eluted in gradient mode with ethanol/water to collect free PTX fraction, and its volume (V2) was measured. The whole elution process was monitored with a UV detector (UV-WXJ 9388 spectrophotometer). Methanol (4.5 mL) was added to 0.5 mL of the nanoparticles fraction, which was sonicated for 5 min and centrifuged at 12 000 rotations/min for 10 min. The concentration (C1) of PTX in the supernatant was determined by high-performance liquid chromatography (HPLC). The mobile phase was acetonitrile/ water (46/54, v/v); the flow rate was 1.0 mL/min, and the detection wavelength was 227 nm. The column was a Hypersil C18 column (5 µm, 150 mm × 4.6 mm, Yilite, China). The fraction of free PTX was treated in the same way, and the concentration (C2) was determined. The drug loading capacity (LC) and loading efficiency (LE) were defined as follows:
LC(%, w ⁄ w) ) [(C1 × V1 × 10) ⁄ (C1 × V1 × 10 + Mass of LMC)] × 100% LE(%, w ⁄ w) ) [(C1 × V1) ⁄ (C1 × V1 + C2 × V2)] × 100% In vitro release of PTX from the LMC nanoparticles was investigated in PBS (0.15 M, pH 7.4) containing 0.1% (w/v) Tween 80. Briefly,
0.5 mL of PTX-loaded LMC nanoparticle solution was added to 99.5 mL of the release medium, which was incubated at 37 °C and 100 rotations/min. At predetermined time intervals, 1 mL of the sample was taken out and centrifuged at 12 000 rotations/min. The amount of PTX in the supernatant was determined by HPLC, and the cumulative released amount was calculated. Hemolysis. Erythrocytes were isolated from the whole blood of a healthy New Zealand rabbit, washed three times with 0.15 M PBS (pH 7.4), and resuspended to obtain a concentration of 2% (v/v). PBS was added into 0.1 mL, 0.2 mL, and 0.3 mL nanoparticle suspensions (10 mg/mL) to obtain a final volume of 2.5 mL, into which 2.5 mL of the erythrocyte suspension was added. Samples were incubated at 37 °C for 1 h and centrifuged at 3000 rotations/min for 10 min. Erythrocytes incubated with 0.15 M PBS and water served as negative and positive controls, respectively. Nanoparticle solution without erythrocytes was used as the blank to compensate the turbidity of free LMC nanoparticles. The hemolysis ratio (HR) was determined spectrophotometrically at 541 nm and calculated from the following equation:
HR(%) ) (ODsample - ODnegative control) × 100 % ⁄ (ODpositive control ODnegative control) Acute Toxicity. Mice were fasted for 12 h and randomly grouped (five males and five females in each group). LMC was dissolved in saline and administered via i.v. at a dose of 625 mg/kg body weight once or twice a day. Dominant signs of toxicity and mortality were monitored for the subsequent two weeks. At the end of the observational period, the animals were sacrificed and a thorough autopsy was carried out. Antitumor Efficacy of PTX-Loaded LMC Nanoparticles. S-180 bearing mice were obtained by subcutaneous inoculation of S-180 cells at the axillary region of male mice. When the tumor volume reached 50-200 mm3, mice were grouped and administered with saline (control), PTX injection at 6 mg/kg, PTX-loaded LMC nanoparticles
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at 6 mg/kg, and PTX-loaded LMC nanoparticles at 3 mg/kg. Drug administration was performed three times via tail vein injection with 2 days spaced between each administration. Tumor size and body weight were recorded every other day. The long diameter (L(t), mm) and the short diameter (W(t), mm) of the tumor were measured with a vernier caliper, and the change of body weight of each mouse was recorded every other day. The tumor volume (V(t))was calculated using the following equation:
V(t) ) L(t) × W(t)2 ⁄ 2 The mice were sacrificed on the third day after the last administration, and the tumors were excised and weighed. The tumor inhibition ratio (TIR) was calculated according to the following equation:
TIR(%) ) (1 - WT ⁄ WC) × 100% where WT and WC refer to the tumor weight of the test and control groups, respectively.
Results and Discussion Synthesis of LMC. Lactic acid was selected as the initiator because it was a hydroxy acid that could initiate the ring-opening polymerization of lactone (benzyl malolactonate) and thus form PMLABz with lactic acid as its carboxyl terminal. Chitosan was a macromolecule with steric hindrance, so it was difficult for pure PMLABz to be conjugated onto chitosan because of the high steric hindrance of its lateral benzyl. Comparatively, lactic acid without side chains was suitable for conjugation onto chitosan via the carboxyl terminal group. On the other hand, the length of PMLABz could be controlled by adjusting the molar ratio of lactic acid to lactone. Linoleic chloride was added prior to PMLABz acyl chloride because the latter was immiscible with MeSO3H and the reaction would be uneven. The feeding sequence resulted in a homogeneous and complete reaction. MeSO3H acted as a solvent and catalyst in the acylation of chitosan. After ice was added, the benzyl groups could be hydrolyzed directly by the strong acid of MeSO3H. Compared with other amphiphilic chitosan derivatives reported, such as LCC21 or N-octyl-O-sulfate chitosan (OCS),23 LMC had longer alkyl chains and longer hydrophilic chains. It was reported that the solubility of hydrophobic compounds was increased with an increase in the carbon length in hydrophobic moieties, so the long chains of LA might be helpful for the encapsulation of PTX.31 The long PMLA chains might form a hydration layer and reduce the absorption of plasma opsonins and thus might decrease the clearance of the reticuloendothelial system (RES). In addition, the lateral carboxyls of PMLA provided modification sites for conjugation of bioactive molecules, and the jointed substances on the outer surface would be easily recognized. Characterization of LMC. FTIR spectra of chitosan and its derivatives are shown in Figure 2. New peaks at 2924, 2853, 1467 cm-1 were attributed to the long alkyl chain of LA, and the peak at 1733 cm-1 was assigned to the carboxyl group of PMLA. Peaks at 1173 and 1088 cm-1 showed the formation of an ester bond. There was a slight peak at 1650 cm-1 in LMC1 representing the amide linkage, which was not detected in LMC2 or LMC3. Such results suggested that LA and PMLA were linked to chitosan mainly by ester linkage, and when the C-6 position was saturated they could also be linked to NH2 via amide bond. This accorded with previous investigations that C-6 OH was more active than C-3 OH, and the positively charged NH2 was inactive for acylation reaction.32
Figure 2. IR spectra of chitosan (a), LMC3 (b), LMC2 (c), and LMC1 (d).
Figure 3. 1H NMR spectra of chitosan (a) and LMC1 (b).
In the 1H NMR spectrum of chitosan (Figure 3), the peak at 1.9 ppm was assigned to the NHAc, while peaks at 2.9-3.8 ppm were attributed to the H-2,3,4,5,6 and OH. In comparison, LMC1 showed the peak at 0.8 ppm, which was assigned to the methyl hydrogen. The peaks at 1.1-1.6 ppm were attributed to the methene hydrogen of LA. The peak at 4.5 ppm was assigned to the CH of the terminal lactic acid residue, and the peaks at 4.7 ppm represented the CH of PMLA. Such results evidenced both LA and PMLA in the modified chitosan. The degree of substitution (DS) of LA in LMC was calculated by comparing the ratio of linoleic methyl protons (δ ) 0.8 ppm) to sugar protons (δ ) 2.9-3.8 ppm), which was defined as the
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Table 1. Particle Size, Zeta Potential, and DS of LMC Nanoparticles (n ) 3) sample
mean diameter (nm)a
zeta potential (mV)
DS (LA)b
DS (PMLA)b
LMC1 LMC2 LMC3 LMC4 LMC5 LMC6
190.0 ( 69.7 (0.14) 223.3 ( 66.1 (0.09) 240.7 ( 185.6 (0.59) 207.9 ( 78.4 (0.14) 266.2 ( 116.6 (0.19) 328.8 ( 127.6 (0.15)
-17.1 ( 0.7 -10.3 ( 4.0 -16.9 ( 2.5 -13.6 ( 4.6 -10.0 ( 3.6 -10.7 ( 2.6
72.9 47.0 22.5 67.1 41.1 24.9
2.9 3.6 3.2 1.8 1.5 1.5
a Values in parentheses represent the polydispersity index. b Determined by 1H NMR.
Figure 5. TEM (a) and SEM (b) images of LMC1 nanoparticles.
Figure 4. XRD patterns of chitosan (a), LMC1 (b), LMC2 (c), and LMC3 (d).
number of LA groups per 100 anhydroglucose units of chitosan. The DS of PMLA was calculated by comparing the ratio of methylene protons of PMLA (δ ) 4.5 ppm) to sugar protons (δ ) 2.9-3.8 ppm), which was defined as the number of PMLA groups per 100 anhydroglucose units of chitosan. The results were shown in Table 1. XRD graphs of chitosan and LMC derivatives were shown in Figure 4. Chitosan showed two sharp diffraction peaks at 2θ ) 11° and 20°, and a small shoulder peak at 2θ ) 22°. The reflection fall at 2θ ) 11° was assigned to crystal form I, which was orthorhombic and represented hydrated crystal structure of chitosan. The strongest reflection at 2θ ) 20° corresponded to crystal form II, which was also orthorhombic.33 LMC1, LMC2 and LMC3 showed only one broad peak at around 2θ ) 20°, and the peak intensity decreased, which suggested that the hydrogen bond was decreased after chemical modification, and the crystalline structure of LMC appeared to be amorphous. The molecules of amorphous LMC were in a disorder state, and the long chains of them were easy to twist together, which was beneficial to encapsulate drugs and form nanoparticles. Morphology of LMC Nanoparticles. The morphology of the LMC nanoparticles as examined by TEM and SEM are shown in Figure 5. The nanoparticles appeared spherical in shape. Size and Zeta Potential of LMC Nanoparticles. Particle size of LMC nanoparticles demonstrated a unimodal distribution (figure not shown). The hydrodynamic diameter and the polydispersity index of the nanoparticles are illustrated in Table 1. The size of LMC nanoparticles was affected by the DS of LA and the length of PMLA. At a certain PMLA chain length, the particle size decreased with the increase of DS (LA), possibly
due to the increasing hydrophobic interactions between LA moieties, which made the hydrophobic core more compact. When the DS (LA) was fixed, the particle size increased with the length of PMLA, which was possibly due to an increase of the extending hydrophilic shell. Long-circulating nanoparticles are able to penetrate into tumor tissues, accumulate, and release the therapeutic drug locally at tumor sites because of the “enhanced permeability and retention” (EPR) effect. Particle size is a key factor that affects the blood circulation half-life of nanoparticles. As a rough approximation, nanoparticles with diameters larger than 200 nm, providing that they are rigid structure, are readily scavenged by macrophages and the RES. For very small particles (
