International Journal of Biological Macromolecules 69 (2014) 489–498

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Chitin extraction from shrimp shell using enzymatic treatment. Antitumor, antioxidant and antimicrobial activities of chitosan Islem Younes a,∗ , Sawssen Hajji a , Véronique Frachet b , Marguerite Rinaudo c , Kemel Jellouli a , Moncef Nasri a a

Laboratory of Enzyme Engineering and Microbiology, University of Sfax, National School of Engineering, PO Box 1173-3038 Sfax, Tunisia AGing Imaging Modeling, CNRS FRE 3405, Université Joseph Fourier, EPHE, Grenoble, France c Biomaterials Applications, 6, rue Lesdiguières 38000 Grenoble, France b

a r t i c l e

i n f o

Article history: Received 1 April 2014 Received in revised form 15 May 2014 Accepted 5 June 2014 Available online 17 June 2014 Keywords: Chitin Chitosan Antimicrobial Antioxidant Antitumor

a b s t r a c t Chitin was recovered through enzymatic deproteinization of the shrimp processing by-products. Different microbial and fish viscera proteases were tested for their deproteinization efficiency. High levels of protein removal of about 77 ± 3% and 78 ± 2% were recorded using Bacillus mojavensis A21 and Balistes capriscus proteases, respectively, after 3 h of hydrolysis at 45 ◦ C using an enzyme/substrate ratio of 20 U/mg. Therefore, these two crude proteases were used separately for chitin extraction and then chitosan preparation by N-deacetylation. Chitin and chitosan samples were then characterized by 13 Cross polarization magic angle spinning nuclear magnetic resonance (CP/MAS)-NMR spectroscopy and compared to samples prepared through chemical deproteinization. All chitins and chitosans showed identical spectra. Chitosans prepared through enzymatic deproteinization have practically the same acetylation degree but higher molecular weights compared to that obtained through chemical process. Antimicobial, antioxidant and antitumoral activitities of chitosan-M obtained by treatment with A21 proteases and chitosan-C obtained by alkaline treatment were investigated. Results showed that both chitosans inhibited the growth of most Gram-negative, Gram-positive bacteria and fungi tested. Furthermore, both chitosans exhibited antioxidant and antitumor activities which was dependent on the molecular weight. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Chitin, a homopolymer of N-acetyl-d-glucosamine residues linked by osidic ␤-1,4 bonds, is the most abundant renewable natural resource after cellulose. Chitin and its derivatives are biomolecules of a great potential, possessing versatile biological activities, demonstrating biocompatibility and biodegradability. Therefore, they found extensive applications in pharmacy, medicine, agriculture, food and textile industries, cosmetics, and wastewater treatment [1]. The main sources of raw material for the production of chitin are cuticles of various crustaceans, principally crabs and shrimps. Chitin existing in the animal world is closely associated with proteins, minerals, lipids and pigments. They all have to be quantitatively removed to achieve the highest purity of chitin necessary for biological applications [2]. To isolate chitin from crustacean

∗ Corresponding author. Tel.: +216 74 27 40 88. E-mail address: [email protected] (I. Younes). http://dx.doi.org/10.1016/j.ijbiomac.2014.06.013 0141-8130/© 2014 Elsevier B.V. All rights reserved.

shells, chemical processing for demineralization and deproteinization have been applied using strong acids and bases to remove calcium carbonate and proteins, respectively [2]. However, the use of these chemicals may cause a partial deacetylation of the chitin and hydrolysis of the polymer resulting in final inconsistent physiological properties [3]. The hydrolysed protein components become also useless during this chemical protein removal. Moreover, chemical treatments bring about hazardous environmental problems like disposal of wastewater making this process ecologically aggressive and a source of pollution. The cost of the chemicals is another drawback of this approach. An alternative approach to these harsh chemical treatments is the use of proteolytic microorganisms [4] or proteolytic enzymes [5]. Bustos and Healy [6] have demonstrated that chitins obtained after deproteinization of shrimp shell waste with various proteolytic microorganisms have higher molecular weights compared to chemically prepared shellfish chitin. In addition, the protein hydrolysates contained bio-active peptides, which may be valuable as pharmacological tools or as a growth stimulating agent in animal feed [7].

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For that purpose, the present research was developed in order to extract chitin from shrimp shells through enzymatic deproteinization. Previously, enzymatic deproteinization was realized using purified proteases: Alcalase 2.4 L, Pancreatin, Delvolase, Cytolase PCL5, Econase CEPi and MP 100, Maxazime NNP and Cellupulin MG [8]. The drawback of this approach is the high cost of purified enzymes. On the opposite, some microbial enzyme preparations could be applicable for deproteinization; this approach is not only a low-cost process but could be also more efficient due to the presence of coexisting proteases. Digestive system of some marine invertebrates contains a variety of highly active proteases that can be used as a source of enzyme preparations. In this study, different non-commercial crude microbial and fish protease extracts were compared for their efficiency against shrimp waste deproteinization. Two different enzyme preparations (one microbial and one fish proteases) allowing the higher deproteinization rate were selected, and used separately for chitin recovery. Chitins were then characterized and converted to chitosans. The antimicrobial, antioxidant and antitumor activities of chitosan obtained by treatment with A21 proteases and that prepared by alkaline treatment were investigated. 2. Material and methods 2.1. Shrimp waste homogenate preparation Fresh shrimp (Metapenaeus monoceros) shells were obtained from the shrimp processing plant located in Sfax, Tunisia. Shrimp waste were washed thoroughly with tap water and mixed with distilled water at a ratio of 1:2 (w/v). Then, they were heated during 20 min at 90 ◦ C to inactivate endogenous enzymes. At end, the samples were drained and homogenized in a Moulinex® blender for about 2 min and then kept at −20 ◦ C without any drying step until further use. 2.2. Chemical analysis

Table 1 Optimal conditions for the different crude proteases used in the deproteinization of shrimp wastes. Crude enzyme

B. mojavensis A21 B. subtilis A26 B. licheniformis NH1 B. licheniformis MP1 V. metschnikovii J1 A. clavatus ES1 Sardinelle (S. aurita) Goby (Z. ophiocephalus) Grey triggerfish (B. capriscus)

Deproteinization conditions Temperature (◦ C)

pH

50 40 50 50 40 40 45 45 45

10 8 10 10 11 8.5 8 9 9

Reference

[11] [12] [13] [14] [15] [16] [17] [18] [19]

alkaline proteases. Shrimp waste homogenate (15 g) was mixed with 45 ml distilled water. The pH and temperature of this suspension were adjusted to optimal conditions for each enzyme (Table 1). Then, the shrimp waste proteins were digested during 3 h with enzymatic preparations containing 20 U/mg. Reactions were stopped by heating the solution at 90 ◦ C during 20 min to inactivate enzymes. The mixtures were pressed manually through four layers of gauze in order to separate solid and liquid phases. Deproteinized products in the solid phase were washed thoroughly until a neutral pH, and then dried overnight at 50 ◦ C. Degree of deproteinization (DDP) was expressed as a percentage and computed by the following equation [21]: DDP(%) =

[(PO × O) − (PR × R)] × 100 PO × O

where PO and PR are the protein concentrations (%) before and after hydrolysis; while, O and R represent the mass (g) of the original sample and the hydrolyzed residue on dry weight basis, respectively.

The moisture and ash content were determined at 105 and 550 ◦ C, respectively, according to the AOAC [9] standard methods 930.15 and 942.05. Total nitrogen content of shrimp waste was determined using the Kjeldahl method. Separately, pure chitin was prepared to determine its nitrogen contribution allowing to estimate the crude protein content by multiplying nitrogen content attributed to proteins by the factor of 6.25 [10]. Lipids were determined gravimetrically after soxhlet extraction on dried raw samples with hexane.

2.5. Deproteinization of shrimp waste by alkali

2.3. Enzymatic preparations

Demineralization was carried out in HCl medium according to a method developed by Madhavan and Nair [22]. Solid fractions obtained after hydrolysis by A21 or B. capriscus proteases were treated with 1.5 M HCl at a ratio of 1:10 (w/v) for 6 h at 25 ◦ C under constant stirring (150 rpm). The chitin residues were filtered through four layers of gauze with the aid of a vacuum pump, washed to neutrality with deionized water and then dried overnight at 50 ◦ C. Demineralization efficiency was determined from the ash content percentage in the chitin samples.

Proteolytic preparations from Bacillus mojavensis A21 [11], Bacillus subtilis A26 [12], Bacillus licheniformis NH1 [13], B. licheniformis MP1 [14], Vibrio metschnikovii J1 [15], Aspergillus clavatus ES1 [16] and crude alkaline protease extracts from Sardinelle (Sardinella aurita) [17], Goby (Zosterisessor ophiocephalus) [18] and Grey triggerfish (Balistes capriscus) [19] were prepared and characterized in our laboratory. Optimal conditions for deproteinization (pH and temperature) are given in Table 1. Protease activity was measured by the method described by Kembhavi et al. [20] using casein as a substrate. In these conditions, one unit of protease activity was defined as the amount of enzyme required to liberate 1 ␮g of tyrosine per minute. 2.4. Deproteinization of shrimp waste by proteases Deproteinization tests were carried out in a thermostated stirred Pyrex reactor (300 ml) using several microbial and fish

Shrimp waste was treated with 2.5 M NaOH under standard autoclaving conditions (15 psi/121 ◦ C) for 20 min. After filtration, the solid residue was washed thoroughly until a neutral pH, and then dried overnight at 50 ◦ C.

2.6. Chemical demineralization

2.7. Deacetylation of chitins The purified chitins were treated with 12.5 M NaOH at a ratio of 1:10 (w/v) for 4 h at 140 ◦ C to obtain perfectly water soluble chitosans in acidic conditions. After filtration, the residues were washed with deionized water, and the crude chitosans were obtained by drying in a dry heat incubator at 50 ◦ C for 12 h.

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2.8.

13 C

CP/MAS-NMR spectroscopic chitin and chitosan analyses

Chitin and chitosan structural analyses were carried out by 13 C NMR (nuclear magnetic resonance) with CP/MAS technique (crosspolarization, magic-angle-spinning) using a BRUKER-ASX300 instrument. NMR spectra were recorded at a 13 C frequency of 75.5 MHz (field of 7.04 T). CP/MAS sequence was used with the following parameters: the 13 C spin lattice relaxation time was 5 s; powdered samples were placed in an alumina rotor used for the double air bearing type MAS system and spun as fast as 8 kHz; contact time was 8 ms. Acetylation degrees (DA) of chitosan samples were determined by dividing the integral of the resonance of the methyl group carbon by the average integral of the resonances of the glycosyl ring carbon atoms. Acetylation degrees were calculated using the following relationship [23]: %DA =

I [CH3] × 100 (I [C1] + I [C2] + I [C3] + I [C4] + I [C5] + I [C6])/6

I represent the integral of the particular resonance signal. Deacetylation degrees (DD) of the chitin samples were determined as follow: %DD = 100 − %DA

2.9. Viscosity-average molecular weight The viscosity measurements were performed using an Ubbelohde viscometer and recording the efflux time of the solution and solvent in a constant temperature bath at 25 ± 0.1 ◦ C. Chitosan samples were dissolved in 0.3 M acetic acid/0.1 M sodium acetate buffer. Intrinsic viscosity ([]) was obtained from linear plots of reduced viscosity (sp/C) against polymer concentration (C, g/ml), extrapolating to zero concentration. The viscosity-average molecular weight (Mv ) of chitosans was determined by Mark–Houwink equation [24]: [] = K MVa with K = 7.95 × 10−2 and a = 0.79 [25]. The average on four replicates was taken for each viscosity data. 2.10. Antimicrobial activity Antibacterial activities of chitosans were tested against eight strains of bacteria; four Gram-negative: Escherichia coli (ATCC 25922), Pseudomonas aeruginosa (ATCC 27853), Klebsiella pneumoniae (ATCC 13883) and Salmonella typhi; and four Gram-positive: Staphylococcus aureus (ATCC 25923), Micrococcus luteus (ATCC 4698), Bacillus cereus (ATCC 11778) and Enterococcus faecalis (ATCC 29212). Antifungal activities were tested using Fusarium oxysporum, Fusarium solani and Fusarium sp., provided by the Center of Biotechnology, Sfax-Tunisia. Antimicrobial activity assays were performed according to the method described by Berghe and Vlietinck [26]. Chitosan samples (50 mg/ml) were prepared under stirring in 0.1% acetic acid. Culture suspension (200 ␮l) of the tested microorganisms (106 colonyforming units/ml of bacteria cells estimated by absorbance at 600 nm and 108 spores/ml of fungal strains measured by Malassez blade) were spread on Muller–Hinton agar and Potato Dextrose Agar medium, respectively. Then, bores (3 mm depth, 5 mm diameter) were made using a sterile borer and loaded with 50 ␮l of chitosan samples at 50 mg/ml. A well with 50 ␮l of 0.1% acetic acid was used as a negative control. Gentamycine and amphotericin B were used as positive references for bacteria and fungi activities, respectively. The Petri dishes were kept, first for 1 h at 4 ◦ C, and

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then incubated for 24 h at 37 ◦ C for bacteria and 72 h at 30 ◦ C for fungal strains. Antimicrobial activity was evaluated by measuring the diameter of growth inhibition zones in millimeters (including well diameter of 5 mm). 2.11. Antioxidant activity Antioxidant activity was tested using three different methods involving different mechanisms: free radical scavenging assay, reducing activity against iron(III) and protective activity on ␤Carotene in presence of free radicals. 2.11.1. DPPH radical scavenging assay The DPPH (1,1-diphenyl-2-picrylhydrazyl) radical-scavenging activity of prepared chitosans was determined as described by Bersuder et al. [27]. A volume of 500 ␮l of each sample, at different concentrations (1 to 5 mg/ml) in 0.1% acetic acid, was mixed with 375 ␮l of 99.5% ethanol and 125 ␮l of 0.02% DPPH in 99.5% ethanol. The mixtures were then incubated for 60 min in the dark at room temperature and the reduction of DPPH radical was measured at 517 nm. A control was conducted in the same manner, except that distilled water was used instead of chitosan sample. In its radical form, DPPH has an absorption band at 517 nm which disappears upon reduction by antiradical compounds. Lower absorbance of the reaction mixture indicated higher DPPH free radical-scavenging activity. DPPH radical-scavenging activity was calculated as follows: Radical-scavenging activity(%) =

Absorbance of control − Absorbance of sample × 100 Absorbance of control

2.11.2. Reducing power assay The ability of chitosan samples to reduce iron(III) was determined according to the method of Yildirim et al. [28]. Sample solutions (1 ml) containing chitosan at different concentrations (1 to 5 mg/ml) were mixed with 2.5 ml of 0.2 M phosphate buffer (pH 6.6) and 2.5 ml of 1% (w/v) potassium ferricyanide. The mixtures were incubated for 30 min at 50 ◦ C, followed by the addition of 2.5 ml of 10% (w/v) trichloroacetic acid. The reaction mixtures were then centrifuged for 10 min at 10,000 rpm. Finally, 2.5 ml aliquot of the supernatant solution, from each sample mixture, was mixed with 2.5 ml of distilled water and 0.5 ml of 0.1% (w/v) ferric chloride. After 10 min reaction time, the absorbance of the resulting solution was measured at 700 nm. Higher absorbance of the reaction mixture indicated higher reducing power. 2.11.3. ˇ-Carotene bleaching method The ability of chitosan samples to prevent the bleaching of ␤carotene was determined as described by Koleva et al. [29]. A stock solution of ␤-carotene/linoleic acid was prepared by dissolving 0.5 mg of ␤-carotene, 25 ␮l of linoleic acid, and 200 mg of Tween-40 in 1 ml chloroform. The chloroform was completely evaporated under vacuum in a rotatory evaporator at 45 ◦ C, then 100 ml distilled water was added, and the resulting mixture was vigorously stirred. The emulsion obtained was freshly prepared before each experiment. Aliquots (2.5 ml) of the ␤-carotene/linoleic acid emulsion were transferred to test tubes containing 0.5 ml of each chitosan sample at different concentrations in 0.1% acetic acid. The tubes were immediately placed in a water bath and incubated at 50 ◦ C for 2 h. Thereafter, the absorbance of each sample was measured at 470 nm. 0.5 ml of distilled water instead of the chitosan solution is used as negative control. Butylated hydroxyanisole

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(a) 100

(a)

100 80

60

D D P (% )

DDP (%)

80

40 20

A21

J1

MP1

A26

NH1

0

ES1

100

3

5

20

30

20

30

(b) 100 80 D D P (% )

60 40 20 0

1

E/S ratio (U/mg)

80 DDP (%)

40 20

0

(b)

60

S. aurita

Z. ophiocephalus

B. capriscus

Fig. 1. Deproteinization of shrimp waste (%) using an enzyme/substrate ratio of 20 U/mg. (a) Microbial proteases: B. mojavensis A21; V. metschnikovii J1; B. licheniformis MP1; B. subtilis A26; B. licheniformis NH1 and A. clavatus ES1. (b) Fish digestive proteases: grey triggerfish (B. capriscus); goby (Z. ophiocephalus) and sardinelle (S. aurita).

(BHA), a synthetic antioxidant, was used as positive standard in the same range as chitosan weight concentration. 2.12. Antitumor activity 2.12.1. Cell culture Bladder cancer cell lines were cultured in RPMI 1640 medium containing 2 mM l-glutamine (Invitrogen Life Technologies, Cergy Pontoise, France) and supplemented with 10% (v/v) fetal calf serum, 2.5 U/ml penicillin, and 2.5 mg/ml streptomycin (Invitrogen, Life Technologies). Cells were cultured at 37 ◦ C in a 5% CO2 -humidified atmosphere and tested to ensure absence of mycoplasma contamination. 2.12.2. MTT tetrazolium salt colorimetric assay RT112 bladder cancer cells (3 × 103 cells/well) were seeded in 96-well plates and incubated for 24 h at 37 ◦ C, before treatment with chitosan. Each chitosan sample was prepared under stirring in aqueous solution containing the stoichiometric amount of HCl (1 mg/ml or ∼6 mM). Successive dilutions of initial chitosan solution were then made up with RPMI medium at pH = 6.5 (using 1 M HCl and HEPES as buffer) to get 50, 100, 500 and 1000 ␮M chitosan concentrations. RT112 cells were treated with these chitosan solutions during 2 h allowing chitosan penetration into cells and preventing cell death at pH = 6.5 as demonstrated separately. Then, the initial mixture was replaced by pure RPMI medium at pH = 7.5 for a total of 24, 48 and 72 h. Ten microliters of tetrazolium dye (MTT) (at 5 mg/ml in phosphate-buffer; PBS) was then added to each well and incubated for 2 h; thereafter, the plate was centrifuged at 1800 × g for 5 min at 4 ◦ C. After careful removal of the medium, 100 ␮l of DMSO was added to each well and plates were shaken. Absorbance was recorded on a microplate reader (Sunrise; Tecan, Lyon, France) at the wavelength of 570 nm. The effect of each chitosan sample on growth inhibition was assessed as percent cell viability, where vehicle-treated cells were taken as 100% viable.

60 40 20 0

1

3

5

E/S ratio (U/mg)

Fig. 2. Effect of the E/S ratio on the deproteinization of shrimp waste using A21 (a) and B. capriscus (b) proteases.

2.12.3. Clonogenic assay of cells in vitro RT112 bladder cancer cell lines (2 × 103 cells/well) were plated in six-well plate and incubated for 24 h at 37 ◦ C, before treatment with chitosans. Initial solutions of chitosan were diluted with RPMI medium at pH = 6.5 to get 500 ␮M of chitosan concentration. The cells were incubated with these solutions for 2 h and then replaced by the RPMI medium. After incubation for 48 h at 37 ◦ C in a humidified incubator, cultures in presence and absence of chitosan samples were observed using an inverted microscope at 200× magnifications (Nikon, Tokyo, Japan). The harvesting of cells was performed using trypsinization. Then, cell suspension were diluted at 1/1000 and seeded into six-well plates for clonogenic assay according to the method elaborated by Franken et al. [30]. Plates were placed in the incubator until cells in control dish have formed large clones (7 days). Colonies were then fixed and stained using a mixture of 6.0% glutaraldehyde and 0.5% crystal violet. 3. Results and discussion 3.1. Chitin isolation and characterization 3.1.1. Enzymatic deproteinization The application of proteolytic enzymes for deproteinization of marine crustacean waste is a current research trend for the conversion of wastes into useful biomass. Hence, in the present study, nine microbial and fish crude alkaline proteases were tested for their efficiency in the chitin isolation from shrimp waste. As shown in Fig. 1, high deproteinization degrees were recorded using 20 U/mg of microbial or fish proteases. Concerning microbial enzyme preparations, high degrees of deproteinization (DDP) were obtained with proteases from B. mojavensis A21 (77 ± 3%) followed by those of B. subtilis A26, V. metschnikovii J1 and B. licheniformis MP1 (75 ± 3%), while NH1 and ES1 proteases gave lower values (65 ± 3% and

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493

Fig. 3. 13C CP/MAS-NMR solid-state spectra of chitins. (Chitin-M) chitin obtained by deproteinization with microbial A21 proteases; (chitin-F) chitin obtained by deproteinization with B. capriscus fish proteases; (chitin-C) chitin obtained by chemical deproteinization.

59 ± 3%, respectively) (Fig. 1a). The lowest DDP obtained with ES1 enzyme preparation could be attributed to the presence of only one protease [16]. The DDP of microbial crude proteases used in this study were similar to many microbial proteases reported in previous studies [5,31]. Fish crude protease extracts were also efficient in shrimp waste deproteinization (Fig. 1b), and the DDP obtained using B. capriscus crude extract was slightly higher (78 ± 2%) than those obtained with proteases from Z. ophiocephalus (76 ± 2%) and S. aurita (75 ± 2%). Therefore, B. mojavensis A21 enzyme preparation and B. capriscus crude protease extract were selected for the chitin isolation. Previous studies reported that enzymatic deproteinization was found to be dependent on the enzyme/substrate (E/S) ratio [32]. Thus, different E/S ratios from 1 to 30 U/mg were applied to compare the shrimp waste deproteinization efficiencies. As shown in Fig. 2, the percentage of protein removal increased with increasing E/S ratio and reached about 77 ± 3% and 78 ± 2%, respectively, for A21 and B. capriscus proteases, with an E/S ratio of 5 U/mg protein. Further increase in enzyme concentration did not increase the deproteinization rates. Deproteinization degree obtained using chemical alkaline process was 94 ± 2%. Although such deproteinization percentage is higher than that obtained using enzymatic treatment, enzymatic process helps to avoid many drawbacks of chemical treatment as reported previously.

3.1.2. Chemical demineralization The ground shrimp waste contained a relatively high content of ash (35.3 ± 0.6%) on the basis of dried initial raw material. In the recovery of chitin from shrimp waste, the associated minerals should be removed as a second step. As a consequence,

deproteinized products were subjected to acid treatment in order to remove minerals. Residual minerals were then determined and results showed that obtained samples contained at around of 1.3% minerals. These values are as low as those reported in previous studies such as that of Percot et al. (1.8%) who suggested that low mineral content must be considered as one of the principal factors determining the good quality of chitin [33]. 3.1.3. 13 C CP/MAS-NMR spectroscopic analysis of chitin The structure of chitin and its purity was evaluated using 13 C CP/MAS solid-state NMR spectroscopy. 13 C CP/MAS-NMR spectra of the chitin samples prepared by enzymatic (chitin-M and chitinF) and alkaline (chitin-C) treatments are shown in Fig. 3. As can be seen all chitins had practically identical spectra. Each spectrum consisted of eight well-defined resonances. Six signals corresponding to glucosamine ring were observed between 50 and 110 ppm, indicating high structural homogeneity. The signal corresponding to carbonyl group (C O) appeared as a sharp and symmetric profile at 173 ppm indicating a unique conformational state, typical of ␣chitin structure. The signal corresponding to the methyl substituent is located at 23 ppm. In addition, the 13 C signals for C3 (at about 76 ppm) and C5 (at about 74 ppm) were clearly separated into two signals. This situation is also characteristic of ␣-chitin. Solid-state NMR studies support the isomorphic form of prepared chitins. Deacetylation degree (DD) is considered as the most important characteristic of chitin, and its value depends on the raw material and the processes used for the deproteinization and the demineralization. Solid-state 13 C CP/MAS-NMR spectroscopy appears to be suitable for the evaluation of the DD and is known to be very sensitive to changes in the local structure [34]. Deacetylation

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Fig. 4. 13C CP/MAS-NMR solid-state spectra of chitosans. (Chitosan-M) chitosan prepared from chitin obtained by deproteinization with microbial A21 proteases; (chitosan-F) chitosan prepared from chitin obtained by deproteinization with B. capriscus fish proteases; (chitosan-C) chitosan prepared from chitin obtained by chemical deproteinization.

degrees of the two chitins isolated by enzymatic deproteinization (at about 9%) were lower than that of chitin obtained by alkaline treatment (16%). Observed differences in the DD values could be attributed to the partial deacetylation of polysaccharide occurring in the alkaline solution during the deproteinization process. The aim of substituting chemical method by the enzymatic one for chitin extraction process is mainly providing pure chitin retaining a structure as close as possible to the native form (less deacetylated and less depolymerized). Indeed, Roberts [35] concluded that the severity of deproteinization step conditions affects, as in the case for the demineralization step, the properties of the final product.

chitosan-M and chitosan-F showed the highest intrinsic viscosity, 197 ml/g and 180 ml/g, respectively, while chitosan-C had the lowest (75 ml/g), which suggests a lower molecular weight. The estimated molecular weight for chitosan-C was 5.820 g/mol, while those of chitosan-F and chitosan-M, were 17,650 and 19,780 g/mol, respectively. These results are in agreement with Oh et al. [36] studies which reported that the molecular weight of chitosan is very sensitive to chitin extraction conditions including, alkali solution, concentration of alkali, high temperature, reaction time, etc. Chitosan-M and chitosan-C which have the same acetylation degree but different average molecular weights were selected for testing their antimicrobial, antioxidant and antitumor activities.

3.2. Chitosan preparation and characterization

3.3. Antimicrobial activities of chitosans

Three chitosans were then prepared in the same conditions from chitins obtained by chemical (chitosan-C) and enzymatic deproteinizations (chitosan-M and chitosan-F). Solid-state 13 C CP/MAS-NMR was used to evaluate the homogeneity and the acetylation degrees (DA) of chitosan samples (Fig. 4). NMR analysis of enzymatic chitosans gave similar peak patterns to that of alkaline chitosane. The residual acetyl group is determined from the carbonyl group signal at around 173 ppm, while that of the residual methyl group appears at around 23 ppm. The lower the signals of carbonyl and methyl groups are, the more efficient the deacetylation reaction is. From these NMR spectra, the DA value was the same for chitosan-M and chitosan-F (DA = 20%). DA of chitosan-C was not significantly different (DA = 19%). Conventionally, a deacetylated chitin with a rate of 70–90% and low protein content is considered as a good final product for application. Viscosity-average molecular weights of prepared chitosans were also determined. Among the three samples used in this study,

Antimicrobial activity of chitosans against several microbial species has been recognized and is considered as one of the most important properties linked directly to their possible biological applications. In this work, the antimicrobial activity of chitosanC and chitosan-M was investigated against four Gram-positive and four Gram-negative bacteria, and three fungi. The potency of chitosan against bacteria and fungi was determined through agar diffusion method. As shown in Table 2, both chitosans prepared by enzymatic and alkaline treatments inhibited the growth of all fungi and bacteria tested except P. aeruginosa and E. faecalis. The inhibitory effect differed with regard to the microbial specie. Both chitosans showed stronger inhibitory effects for fungi than bacteria. Further, they were more active against Gram-negative than Gram-positive bacteria. The antifungal activity is more potent against F. oxysporum, while the antibacterial activity is more potent against E. coli followed by K. pneumoniae. Otherwise, the inhibitory effect was

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slightly influenced by the average molecular weight. Indeed, for all Gram-negative bacteria, the antibacterial effect was significantly higher with chitosan with lower MW ; whereas, opposite effect was observed with Gram-positive bacteria as well as fungi since chitosan with higher MW was more effective. Concerning the antibacterial activity of chitosan, our results are in concordance with previous studies [37,38]. Zheng and Zhu [38] demonstrated for Gram-positive S. aureus, the antimicrobial activity increases with increasing of the chitosan MW . On the opposite, for Gram-negative E. coli, they indicated that the antibacterial activity increases with decreasing MW . These authors suggested the two following mechanisms for the antimicrobial activity: (1) in the case of S. aureus, the chitosan on the surface of the cell forms a polymeric membrane, which inhibits nutrients from entering the cell and, (2) for E. coli, chitosan with a lower MW enters the cell through pervasion. Concerning the antifungal activity, it has been reported that chitosan reduces the fungal infection, mycelial growth, sporangial production, release of zoospores and germination of fungi [39,40]. Chitosan activity on Fusarium fungus was previously studied by many authors. For example, Guo et al. [41] found that 0.5 ␮g/ml of chitosan (DA = 3%, MW = 200 kDa) inhibit 15% of F. oxysporum growth. The effect of chitosan on the growth and morphology of F. oxysporum and its ability to elicit a defence reaction against this fungus in date palm roots were also been demonstrated by El Hassni et al. [42]. Up to the present, many researchers have targeted chitosan characteristics effect on antibacterial activity, whereas, few investigations have been performed on the effect on antifungal activity. Effect on F. oxysporum f. sp. vasinfectum has been investigated by Guo et al. [43] which found that the inhibitory effect on mycelial growth occurred when this fungi grew on media with high MW chitosan (MW = 200 kDa) compared to lower MW . The same effect was observed with Alternaria solani and Valsa mali [46]. However, this trend was not observed with other fungi [44]. Our findings suggest that chitosan behaviour towards fungi is dependent on its molecular weight. However, it is necessary to carry out more basic studies on the mode of action and the effect of the molecular weight of chitosan on the development of fungi. 3.4. Antioxidant activity Synthetic antioxidants, such as butylated hydroxyanisole (BHA), butylated hydroxytoluene (BHT), t-butylhydroquinone (TBHQ) and propyl gallate, have been widely used in food products to delay the deterioration caused by lipid oxidation. However, these antioxidants create potential health hazards, and their use has been restricted in some countries. Thus, it is essential to develop safe and natural antioxidants as alternatives to synthetic ones. For that purpose, antioxidant properties of chitosan were assessed using

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(a)

(b)

Fig. 5. Antioxidant activities of chitosan samples at different concentrations. (a) DPPH free radical-scavenging activities, (b) inhibition of ␤-carotene bleaching.

three tests: DPPH radical-scavenging ability, chelating activity on Fe2+ and the ␤-carotene bleaching method. 3.4.1. DPPH radical-scavenging assay Free radical-scavenging is a primary mechanism by which antioxidants inhibit oxidative processes. The DPPH radical scavenging assay is a widely used method for evaluating the ability to scavenge free radicals generated from DPPH reagent. DPPH is a stable free radical, which can be reduced by a proton-donating substrate such as an antioxidant, causing the decolorization of DPPH and the reduction of the absorbance at 517 nm. The decrease in absorbance is taken as a measure for radical scavenging activity. DPPH radical scavenging capacity of chitosans prepared by enzymatic and alkaline treatments, and BHA (used as positive control) was investigated at different concentrations (0; 1; 2; 3; 4 and 5 mg/ml). As shown in Fig. 5a, both chitosans exhibited high antioxidant activity against DPPH and the scavenging activity was concentration-dependant. However, both chitosans showed lower

Table 2 Diameters of inhibition zones (expressed in mm) against bacteria and fungi in the presence of chitosans after incubation 24 h at 37 ◦ C for bacteria and 72 h at 30 ◦ C for fungi.

Gram−

Gram+

Fungi

E. coli P. aeruginosa K. pneumoniae S. typhi S. aureus B. cereus E. faecalis M. luteus F. oxysporum F. solani Fusarium sp.

Positive control

Chitosan M (DA = 20%; MW ≈ 19,800 g/mol)

Chitosan C (DA = 19%; MW ≈ 5800 g/mol)

20.0 ± 0.5 18.0 ± 0.8 12.0 ± 0.5 18.0 ± 0.5 25.0 ± 0.0 20.0 ± 0.5 20.0 ± 1.0 19.0 ± 0.5 14.0 ± 0.5 16.0 ± 0.5 13.0 ± 0.5

11.0 ± 0.5b R 11.5 ± 0.0b 8.0 ± 0.5b 9.5 ± 0.5a 9.5 ± 0.3a R 8.0 ± 0.3a 16.0 ± 0.3a 18.5 ± 0.5a 16.0 ± 0.8a

12.5 ± 0.5a R 12.5 ± 0.5a 9.0 ± 0.0a 8.0 ± 0.3b 9.0 ± 0.0b R 7.0 ± 0.5b 13.0 ± 0.5b 17.0 ± 0.5b 14.0 ± 0.3b

Diameter well: 5 mm; R resistant; chitosan samples at pH 5.5; Gentamycin and Amphotericin B were used as positive control at a concentration of 15 ␮g/well and 20 ␮g/well, respectively; acetic acid 0.1% as negative control. a-b Means with different superscripts within a column indicate significant difference (P < 0.05).

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Fig. 6. Cytotoxic effects of chitosan samples on human bladder cancer (RT112) cells. (a) Morphological observation of the chitosan tested after 48 h of incubation with RT112 cells. Cell morphology was observed using an inverted microscope at 200× magnification (Nikon, Tokyo, Japan). (b) Clonogenic assay performed in six-well plates and comparative effects of chitosan-M and chitosan-C on the growth of RT112 cells. The first well represent the negative control in which RT112 cells were incubated only with the medium. (c) MTT assay and determination of the IC50 of each chitosan sample. The values are represented as the percentage of cell inhibition, where vehicle-treated cells were regarded as 100%.

radical-scavenging activity than BHA at the same concentrations. For example, at a concentration of 5 mg/ml the scavenging activity of BHA was 90%, while those of the chitosan-M and chitosan-C were 75 and 80% for chitosan-M, respectively. Therefore, the obtained results suggest that chitosan could be applied as proton donors and could react with free radicals to convert them to more stable products and terminate the radical chain reaction. From these results, half maximal inhibitory concentration (IC50 ) values were determined. The lower IC50 indicates the higher free radical-scavenging ability. Chitosan-C which have the lowest MW was found to be the most active radical-scavenger with an IC50 of 1.62 mg/ml compared to chitosan-M which have the highest MW (IC50 = 2.20 mg/ml). 3.4.2. Reducing power The reducing power assay could be used to evaluate the ability of antioxidant to transfer electron or hydrogen [28]. The presence of reducing agents (antioxidants) in tested samples results in the reduction of Fe3+/ ferric cyanide complex to ferrous form. Some studies have reported that there is a direct correlation between antioxidant activities and reducing power of certain bioactive

compounds. Reducing power capacity of both chitosans was investigated at different concentrations (0; 1; 2; 3; 4 and 5 mg/ml). Both chitosans showed relatively low reducing power, indicating that chitosans were not able to reduce Fe3+ to Fe2+ by donating electron (data not shown). 3.4.3. ˇ-Carotene bleaching inhibition assay The ␤-carotene–linoleic bleaching inhibition assay simulates membrane lipid oxidation and can be considered as a good model for membrane based lipid peroxidation. In this oil–water emulsionbased system, linoleic acid acts as a free radical generator producing peroxyl radicals under thermally induced oxidation. These free radicals attack the ␤-carotene chromophore resulting in a bleaching effect, which can be inhibited by a free-radical scavenger [29]. Antioxidant activity of enzymatic and alkaline chitosans was analyzed using the ␤-carotene–linoleate bleaching inhibition assay in comparison with BHA at different concentrations (0; 1; 2; 3; 4 and 5 mg/ml). Results are represented in Fig. 5b. Antioxidant activity of both chitosans increased with increasing chitosan concentration. For instance, as the concentration of the chitosan-C raised from 1 to 5 mg/ml, ␤-carotene–linoleic bleaching inhibition increased

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from 33% to 70%. Chitosan-C with low molecular weight showed significantly higher ability to prevent the bleaching of ␤-carotene than chitosan-M with high molecular weight. This is in line with the work reported by Chien et al., [45] who showed that low molecular weight chitosan (12 kDa) exhibited stronger scavenging activity towards DPPH radicals, superoxide anion radicals and hydrogen peroxide, compared to higher MW chitosans (95 and 318 kDa). However, both chitosans showed lower antioxidant activity than BHA at the same concentrations. These results demonstrated that shrimp waste chitosans have strong effects against the discoloration of ␤-carotene.

497

MW (19,780 g/mol). The significance of this effect must be investigated further across a wide variety of chitosan molecular weights. Nevertheless, these important properties make chitosan an exciting and promising excipient for agriculture, packaging and pharmaceutical industry.

Acknowledgements This work was funded by the Ministry of Higher Education and Scientific Research, Tunisia. The authors would like to thank M. Abdelmajid Dammak from the National School of Engineering of Sfax for his help with English.

3.5. Antitumor activity The use of naturally occurring compounds with antitumor properties has attracted much interest in chemotherapy and treatment of cancers. Currently, bladder cancer is estimated to be the seventh most common malignant neoplasm and the eighth leading cause of cancer-related death worldwide [46]. In light of this, the potential cytotoxic effect of chitosan samples was investigated against human bladder cell carcinoma (RT112). Microscopic observation (Fig. 6a) and clonogenic assay (Fig. 6b) of RT112 cells after 48 h of treatment with chitosan samples showed an important reduction in cell number, as compared to the control. Chitosan-C with lower MW was much more active on bladder human cancer cells than chitosan-M. MTT assay showed that both chitosans were able to induce cytotoxicity in RT112 cells in a dose- and time-dependent manner (Fig. 6c). Chitosan-C with lower MW was most effective with an estimated IC50 of 140 ␮M after 48 h and 41 ␮M after 72 h of incubation. On the other hand, the IC50 values of chitosan with higher MW were 321 and 52 ␮M after 48 and 72 h of incubation, respectively. Antitumor effect of chitosan on bladder cancer cells was previously investigated by Kuppusamy and Karuppaiah [47] but there are no studies on its dependence on the chitosan molecular weight. They studied effect of chitosan on T24 human bladder cancer cells and reported a cytotoxic effect of chitosan. Their results are in accordance with our findings showing an important antitumor activity of chitosan against human bladder cell carcinoma. Our results showed that this effect seemed to be dependent on chitosan molecular weight. However, in-depth studies of this antitumor effect must be investigated further. Antitumor effect of chitosan has been also studied towards other carcinoma cells. Shen et al. [48] demonstrated the protective effect of chitosan on the proliferation of HepG2 cells and suppress tumor growth in HepG2-bearing SCID mice. Chakraborty et al. [49] reported that modified chitosan had in vitro cytotoxicity against HeLa cell lines in Swiss mice. Hosseinzadeh et al. [50] proved that chitosan nanoparticls inhibit cell viability of HT-29 colon carcinoma cell line. 4. Conclusion In this study, the ability of nine different protease preparations to deproteinize shrimp waste for chitin extraction was investigated. B. mojavensis A21 and B. capriscus proteases, which were able to eliminate up to 77 ± 3% of the shell proteins using with an E/S ratio of 5, were selected for chitin extraction. Chitin samples were then characterized and compared to sample prepared through chemical deproteinization. The biological activities of chitosan-M and chitosan-C obtained by enzymatic and alkaline treatment, respectively, were studied. Chitosan-C with lower MW (5.820 g/mol) exhibited higher antioxidant activity. Further, chitosan-C exhibited higher antitumor activity on bladder carcinoma cells (RT112) and showed slightly higher antimicrobial activity than chitosan-M with higher

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