Printer Friendly

OPTIMIZATION AND PHYSICOCHEMICAL CHARACTERIZATION OF CHITOSAN AND CHITOSAN NANOPARTICLES EXTRACTED FROM THE CRAYFISH PROCAMBARUS CLARKII WASTES.

INTRODUCTION

Chitosan is considered as one of the smartest natural cationic biopolymers. It is regarded as the second most abundant polymer after cellulose (Aranaz et al. 2009). Chitosan is obtained from the deacetylation of chitin, which is considered as the second most abundant biopolymer after cellulose, which is naturally found in the exoskeletons of many insects, the cell wall of many fungi, and crustacean shells. Moreover, it is found in wastes from processing of marine food products, such as shrimp, crab, and krill shells (Simpson et al. 1999). Chitosan has remarkable properties, including nontoxicity, biodegradability, biocompatibility, and improved solubility, in addition to immunorestorative properties (Shard et al. 2014) and ease of modification that enable it to be applied in various fields, such as aquaculture, agriculture, cancer therapy, other medical fields, cosmetics, textiles, drug carriers, water treatment, and biochemical engineering industries (Zhao et al. 2018).

The field of nanotechnology has experienced a significant growth over the last years. The application of nanotechnology may have the potential to solve many problems related to human and animal health, production, reproduction, and treatment of diseases (Jimenez-Fernandez et al. 2014). They are favored because of their small size, nano-scale, which increases the available surface area to interact with biological support, increases the bioavailability of essential compounds, and enables efficient uptake by body cells and deep penetration into the target sites (Alishahi et al. 2011). Chitosan nanoparticles have been proven to be applied in drug delivery either by ocular administration routes or by mucosal or traditional routes

(Kumar et aI. 2016). Moreover, they were used in the delivery of vaccines and designing of nonviral vectors for gene delivery (Bravo-Anaya et al. 2016). Moreover, Chitosan nanoparticles have been proven to act as feed additives, improving the growth and meat quality status of Oreochromis niloticus (Wang & Li 2011). Different approaches have been used in the synthesis of chitosan nanoparticles, such as ionotropic gelation (Calvo et al. 1997), spray drying (Ngan et al. 2014), and water-in-oil emulsion cross-linking (Riegger et al. 2018). Ionotropic gelation method was considered as the best procedure, as it is regarded as a simple and mild preparation method that does not require any harsh organic solvents or sonication or homogenization (Calvo et al. 1997). It has been found that the variation in the concentration of chitosan and the polyanion tripolyphosphate (TPP), during the nanoparticles' synthesis, affects greatly some physicochemical characteristics of the resultant nanoparticles and accordingly the application that will be performed (Shard et al. 2014).

Wastes of the crayfish Procambarus clarkii are considered as excellent sources of chitin because of their higher chitin and lower protein contents. Their wastes contain 23.5% chitin by weight, whereas shrimp wastes and crab wastes contain 14%-27% and 13%-15% chitin by weight, respectively (No et al. 1989). Moreover, they are considered as feasible sources of chitin as the crayfish industry in Egypt has increased during the last 5 y, and the drastic problem facing crayfish plants is how they can get rid of their solid wastes that reach hundreds of tons, as the inedible part of the animal reaches about 60%-70% of the total body weight (Ibrahim & Khalil 2009). There is no previous data on chitosan nanoparticles extracted from P. clarkii.

The objective of the present study was to evaluate the most optimum condition for preparing chitosan nanoparticles that will, subsequently, give the best physicochemical characterization regarding the nanoparticles' size and surface morphology.

MATERIALS AND METHODS

Materials

Exoskeleton wastes of Procambarus clarkii were obtained from a commercial crayfish processor company (Jiang's Fish Processor Co., Ltd., Cairo, Egypt). Commercial chitosan, with degree of deacetylation 93% and molecular weight (Mw) 161.16 kDa, was purchased from Oxford Laboratory Chemicals, India. Sodium hydroxide 50%, hydrochloric acid 33%, sodium hypochlorite 12.5%, sodium TPP, and glacial acetic acid 99.7% were supplied by Abou-Zaabal Company for Chemicals.

Extraction of Chitin from the Exoskeleton of Procambarus clarkii and Its Conversion (Deacetylation) into Chitosan

Raw material. Exoskeleton wastes of Procambarus clarkii were brought to the laboratory at Faculty of Science, Ain Shams University. The exoskeletons were crushed into small pieces and then washed with tap water to remove soluble organics, adherent proteins, and other impurities. Finally, the crushed exoskeletons were left to dry at room temperature.

Chitin extraction. Chitin was extracted from the exoskeletons of Procambarus clarkii following the standard procedure, according to No et al. (1989), with slight modifications. It involves three main steps, namely, demineralization, deproteinization, and decoloration.

Demineralization. The demineralization step was carried out by treatment of the crushed exoskeletons with 1N hydrochloric acid (HO) at room temperature for 30 min, until the effervescence completely disappeared, with a solid-to-solvent ratio of 1:15 (w/v). The demineralized crushed exoskeletons were washed with distilled water until the pH became neutral.

Deproteinization. Deproteinization was performed using alkaline treatment of the samples with 3.5% sodium hydroxide (NaOH) solution at 65[degrees]C for 2 h with a solid-to-solvent ratio of 1:10 (w/v). This step was followed by washing with distilled water until neutrality.

Decoloration. Decoloration was carried out by immersing the samples in acetone for the removal of the carotenoid astaxanthin (the pigment that is responsible for the dark red color of the exoskeleton of Procambarus clarkii). For the removal of other pigment traces, the crushed exoskeletons were placed in 10% sodium hypochlorite (NaOC1) solution, with a solid-to-solvent ratio of 1:15 (w/v), and left overnight. Moreover, final bleaching was performed by a night-stand immersion of the crushed exoskeletons in 50% hydrogen peroxide (H202) and 33% hydrochloric acid (HC1), at a ratio of 9:1 (v/v) and with a solid-to-solvent ratio of 1:15 (w/v) (Arbia et al. 2013). The samples were washed with distilled water until neutral pH was reached and left to dry at room temperature. Finally, the resultant white chitin was ground several times using a kitchen grinder and stored in polyethylene bags at room temperature for further processing.

Deacetylation of the extracted chitin. The extracted chitin from Procambarus clarkii was treated with 50% sodium hydroxide (NaOH) solution at 100[degrees]C using a hot plate magnetic stirrer for 3, 6, 9, and 12 h, with a solid-to-solvent ratio of 1:10 (w/v) (No & Meyers 1989). The mixture was cooled for 30 min at room temperature, then washed with distilled water several times until neutrality, and filtered to retain the solid extract. This obtained chitosan was left to dry at room temperature and then stored in polyethylene bags for further use.

Preparation of chitosan nanoparticles. Chitosan nanoparticles were synthesized by the ionotropic gelation method according to Calvo et al. (1997), with slight modifications.

Preliminary experiments were performed so as to determine the optimum condition for the formation of nanoparticles.

Three different ratios of chitosan and the polyanion sodium TPP were experimented: 1:1, 3:1, and 1:3.

The procedure is detailed as follows:

Chitosan was dissolved in 1% glacial acetic acid (CH3COOH) solution and sodium TPP was dissolved in 1% deionized water.

The chitosan solution was then placed on the magnetic stirrer for 2 h at room temperature until a clear solution was obtained.

Sodium TPP solution was added dropwise to the chitosan solution, at three different ratios, 1:1, 1:3, and 3:1 (w/w), under magnetic stirring at room temperature for 1 h until a milky white suspension, chitosan--TPP nanoparticles, was obtained.

The resultant suspension was kept in the refrigerator at 4[degrees]C overnight to allow sufficient reaction. The nanoparticles were collected by cooling ultracentrifugation at 4[degrees]C for 30 min at 16,000 rpm and washed extensively to remove the unreacted reagents. The precipitate was re-dispersed in minimal amount of distilled water, freeze-dried, and finally stored at 4[degrees]C until further use.

Physicochemical Characterization of Chitosan and

Chitosan Nanoparticles

Fourier-transform IR (FTIR) spectroscopy. Fourier-transform IR spectra of the chitosan nanoparticles were recorded with an FTIR 4100 (Jasco, Japan) spectrophotometer at the Central Laboratory of Faculty of Science, Ain Shams University. The spectral region between 4,000 and 400 [cm.sup.-1] was scanned.

Samples were prepared as KBr pellets. Dried chitosan powder was mixed thoroughly with KBr and then pressed to form an ultimate thin homogenous disc with a thickness of 0.5 mm. The chitosan concentration in the samples was 2%, calculated with respect to KBr.

Elemental analysis. An Elementar Vario EL III apparatus (Elementar, Germany) was used in the present study for the analysis of the amount of C and N in chitosan. Briefly, approximately 2 mg of chitosan was placed inside a thin capsule followed by heating in a combustion tube. The weight percentages of carbon and nitrogen were measured using thermal conductivity detection.

The degree of deacetylation of chitosan was determined using the following equation according to Abdou et al. (2008):

DDA(%) _ [(6.857 - C/N)/1.7143] x 100, where C/N is the ratio of carbon/nitrogen that was determined from the elemental analysis of the resultant chitosan.

Nuclear magnetic resonance spectroscopy ('H-NMR). The 1H-NMR spectra were measured on a Varian Gemini 300-MHz spectrometer, with chemical shift ([delta]) expressed in ppm downfield, with tetramethylsilane as initial standard, in DMSO-d6, and coupling constants J in Hz at the Central Laboratory of Cairo University.

Calculation of the Mw of chitosan using viscometry. Viscometry is the best method for the measurement of the Mw of chitosan. The viscosity ratio (relative viscosity) [??].sub.rei], which is given by the ratio of the outflow time for the solution (t) to the outflow time for the pure solvent (to), was calculated by the following equation:

Arel = t/to.

The (to) and (t) were measured by using an Ostwald viscometer; specific viscosity 010, which is the relative increment in viscosity of the solution over the viscosity of the solvent, was calculated by the following equation:

[mathematical equation not reproducible]

The relative viscosity (Tirai) and specific viscosity (Thp) values for different concentrations of chitosan (1.3, 1, 0.7, and 0.5 g %) were calculated. The intrinsic viscosity [i] is calculated from the plot of rlsp/c versus concentrations.

The Mw of the prepared chitosan was calculated in kDa according to the Mark-Houwink equation:

[11] = KM[degrees], where k = 3.5 x 10-4 (cm3 g-1) and a = 0.76.

Surface Morphology of Chitosan and Chitosan Nanoparticles Using Scanning Electron Microscopy (SEM) and Transmission Electron Microscopy (TEM)

Scanning electron microscopy of chitosan. The chitosan powder was further grinded until it was smooth and then coated with gold by spraying gold powder. The chitosan powder was examined in the solid state using an SEM Model Quanta 250 Field Emission Gun attached with an EDX unit (energy-dispersive X-ray analyses), with an accelerating voltage of 30 kV, magnification 14X up to 1,000,000, and resolution for Gun. In (FEI Company, The Netherlands), at the SEM Unit, Egyptian Mineral Resources Authority "Egyptian Geological Survey", Ministry of Petroleum.

Transmission electron microscopy of chitosan and chitosan nanoparticles. The morphological examination of chitosan and chitosan nanoparticles, and the particle size measurement of nanoparticles were determined by TEM. For TEM, 0.5 g of chitosan nanoparticles was suspended in 2% glacial acetic acid and sonicated in an ultrasonic bath for 30 min. Then, a drop of the suspension was dripped onto carbon-coated copper grids and left to dry at room temperature overnight. Finally, the grids were examined with a JEOL JEM-1400 transmission electron microscope at the Faculty of Agriculture Research Park, Cairo University.

Determination of the zeta potential ofthe chitosan nanoparticles. Chitosan nanoparticles (0.05% w/v) were dissolved in 1% acetic acid and the solution was sonicated using a probe sonicator (Osonic 450 kHz, amplitude 30%) for 11 min. Sodium hexametaphosphate solution (0.1% w/v) in distilled water was prepared and added to the chitosan solution at a rate of 1.5 mL/min until the transmittance of the suspension reaches up to 70% at an operating wavelength of 632 nm. The whole solution was sonicated for 7 min. To stabilize the particle suspension, 1% PVA was added. The zeta potential measurements of chitosan nanoparticles were performed using the dynamic light scattering instrument NICOMP ZLS-380 (PSS, Santa Barbara, CA), using the 632-nm line of an He Ne laser as the incident light with angle 90[degrees] and zeta potential with external angle 18.9[degrees] at Nanomaterial Investigation Laboratory, Central Laboratory, National Research Center.

RESULTS AND DISCUSSION

Chitosan Production and Its Preliminary Identification

These findings prompted us to extract chitosan from the exoskeleton wastes of Procambarus clarkii; the color of the obtained chitosan powder was buff compared with the white color of commercial chitosan that was extracted from the exoskeleton wastes of shrimps (Fig. 1). This is in accordance with Ghannam et al. (2016). Upon adding 1% glacial acetic acid to the resultant chitosan with mild stirring at ambient temperature, solubility occurred and the solution turned transparent, which confirmed the presence of chitosan due to the presence of the amino group. Chitosan is also soluble in 1% hydrochloric acid; however, it is insoluble in sulfuric acid and phosphoric acid. The solubility of chitosan is controlled by the degree of deacetylation, which must be at least 85% to achieve the desired solubility (Toan & Hanh 2013).

With regard to the chemical structure of chitin and chitosan

(Fig. 2), they have a similar structure. Chitin is formed of a linear chain of acetylglucosamine groups, whereas chitosan is obtained by removing enough acetyl groups (CH3-CO). This process is called deacetylation.

To obtain the highly deacetylated chitosan ratio, four trials were performed, at 3, 6, 9, and 12 h. The FTIR spectra of the obtained products indicated that in the case of 3, 6, and 9 h, as the time increased the degree of deacetylation increased, whereas at 12 h the degradation of chitosan occurred.

Degree of Deacetylation

The degree of deacetylation of the obtained chitin/chitosan is considered as the most important parameter that affects their properties, including physicochemical, biological, and mechanical properties. Moreover, the degree of deacetylation depends mainly on the method of extraction; several reaction conditions must be taken into consideration to obtain the optimum condition for the resultant chitosan before using it in any application.

The degree of deacetylation of the extracted chitosan from the exoskeleton wastes of Procambarus clarkii was 87% compared with that of the commercial chitosan (93%). The degree of deacetylation was detected using elemental analysis, in which the C:% H:% N was 39.42:7.09:7.34, respectively.

Fourier- Transform IR Spectroscopy

Chitin is known to have three crystal forms: [alpha], [beta], and y

(Jiang & Xu 2006). Fourier-transform IR spectroscopy is one of the most popular methods to identify the [alpha], [beta], or y form of chitin (Jang et al. 2004). Existing studies have revealed that chitin with [alpha]-crystallinity displays bands around 1,650, 1,620, and 1,550 cm ' (Lavall et al. 2007, Rinaudo 2006, Brugnerotto et al. 2001). In this study, these bands were at 1,653 [cm.sup.-1], 1,621 [cm.sup.-1] (C = O secondary amide stretch), and 1,550 [[cm].sup.-1

(N--H band, C--N stretch). The existence of these bands suggested that the extracted chitin was in the a-form. In addition, there were no bands at 1,540 [[cm].sup.-1 in the spectral evaluation, which suggests the absence of protein residues in the chitin

(Morin & Dufresne 2002). This shows that the deproteinization process during chitin isolation was sufficient.

The four trials of the degree of deacetylation were studied by using the FTIR technique. The main difference between chitin and chitosan was the formation of the amino group ([NH.sub.2]) in chitosan and the disappearance of the amidic carbonyl group of chitin. The FTIR of the four trials are illustrated in Fig. 3A--D.

The FTIR showed that as the time increased, the forked peak of the amino group at 3,442 [cm.sup.-1] appeared and the abundance of the carbonyl of the acetamido group of the chitin at approximately 1,700 [cm.sup.-1] decreased (Fig. 3A--C). The aforementioned data indicated the increase in the degree of deacetylation. Unfortunately, in the case of 12 h, partial degradation occurred (Fig. 3D).

The structure of chitosan was strongly confirmed by comparing the FTIR of commercial chitosan that was obtained from shrimp (Oxford Laboratory Chemicals, India) and the selected extracted chitosan (9 h deacetylated) from Procambarus clarkii wastes. Moreover, the chemical structures of the chitosan and the synthesized chitosan nanoparticles were confirmed by FTIR analysis (Fig. 3C, E).

There were no major differences in the FTIR spectra of the obtained chitosan and chitosan nanoparticles. In both of them, the band at 3,423-3,727 (br.) [cm.sup.-1] was assigned to [NH.sub.2] and OH stretching vibrations. The band at 1,631-1,633 [cm.sup.-1] was attributed to the carbonyl amide. The only difference was observed at 1,107 cm-1 in chitosan nanoparticles, which is characteristic of the P = O and could be attributed to the linkage between ammonium ions and phosphate groups, as illustrated in Fig. 3F.

Nuclear Magnetic Resonance Spectroscopy

The 1 H-NMR spectrum of the extracted chitosan showed singlet peaks at [delta] 1.8 and at 4.3 ppm, which correspond to [CH.sup.3] of the acetyl group and H-1 (Ac) of the acetylated chitosan, in addition to the characteristic peaks of deacetylated chitosan that were found at 2.8 and 4.7 ppm, which are characteristic to H-1 and H-2 of the deacetylated form, respectively.

Further support for the structure and ratio of acetylated and deacetylated chitosan was gained from 'H-NMR. Figure 4 reveals that the percentage of methyl group of the acetamido group of acetylated chitosan at [delta] = 1.8 ppm and the peak of H-1(A) acetylated chitosan at [delta] = 4.6 ppm decrease, whereas the degree of H-1 (D) deacetylated chitosan at [delta] = 4.7 ppm increases; this indicates that the degree of deacetylation increases as the time of the deacetylation increases (Fig. 4).

Calculation of the Mw of Chitosan

The weight-average Mw of chitosan is an important characteristic that greatly affects its physicochemical and physiological properties and, hence, the application that will be manipulated. The Mw of chitosan is proportional to its viscosity

(Kanauchi et al. 1995).

Viscometry is considered as the best technique for determination of the Mw of chitosan. The relative viscosity ([[eta].sub.rei]) and specific viscosity [[eta].sub.sp] values for different concentrations of chitosan (1.3, 1, 0.7, and 0.5 g %) are shown in Table 1. The intrinsic viscosity [eta] = 1.87 was calculated from the plot of [mathematical expression not reproducible] versus concentrations shown in Fig. 5; the Mw of the prepared chitosan was found to be 80.72 kDa.

The aforementioned results indicated that chitosan obtained from Procambarus clarkii had a low Mw.

Surface Morphology of Chitosan and Chitosan Nanoparticles

Scanning Electron Microscopy of Chitosan

Scanning electron microscopy analyses conducted on chitosan have revealed that it has three main surface morphologies:

(1) a hard and rough surface without pores or nanofibers, as those isolated from Daphnia sp. (Kaya et al. 2013); (2) a surface solely composed of nanofibers, as in certain species of insects and crustaceans (Wang et al. 2013; Kaya et al. 2014); and (3) a surface with both pores and nanofibers, which is the most common morphology (Paulino et al. 2006). The morphology of chitosan extracted from Procambarus clarkii belonged to the first type. Chitosan appeared as flakes that possess a hard and rough surface that exhibits neither pores nor nanofibers

(Fig. 6A). Moreover, magnification of the flakes indicated that they consist numerous fine fibers (Fig. 6B).

Transmission Electron Microscopy of Chitosan and Chitosan Nanoparticles

The chitosan structure was further examined by TEM to confirm the results of the SEM and reveal its detailed structure, and it is noteworthy to mention that the present study was the first attempt to examine the structure of chitosan by TEM. It was found that it consists of an anastomosing network of fibers, which agrees with the previous data of the SEM (Fig. 7A).

The surface morphology of the three trials of the chitosan nanoparticles with different concentrations of the chitosan and the polyanion TPP, 3:1, 1:3, and 1:1, respectively, was examined by TEM.

The present study showed that using chitosan and TPP in a ratio of 3:1 resulted in the largest particle size with a large range, whose size varied from 170 to 265 mn (Fig. 7B), whereas when a ratio of 1:3 chitosan to TPP was used, both large size and aggregated particles were spotted, with the particle size ranging from 40 to 77 nm (Fig. 7C). However, the optimum size of chitosan nanoparticles and less aggregation were obtained from the lowest chitosan and TPP concentration, ratio 1:1, in which the particle size fluctuated from 3 to 9 nm (Fig. 7D).

Determination of the Zeta Potential of the Chitosan Nanoparticles

The zeta potential is the electrostatic potential at the boundary dividing the compact layer and the diffuse layer of the colloidal particles. It is considered as an important parameter for various applications, including characterization of biomedical polymers, electrokinetic transport of particles or blood cells, membrane efficiency, and microfluidics (Shim et al. 2002).

The importance of the potential to so many applications in science and engineering led to the development of a number of theories (Ostolska & Wisniewska 2014).

The zeta potential of the chitosan nanoparticles synthesized from different ratios of chitosan to TPP was calculated so as to determine the surface charge of the resulted chitosan nanoparticles and, hence, distinguish the optimum condition for the chitosan nanoparticles. Results showed that surface charge of the chitosan nanoparticles obtained from the three different ratios of chitosan to TPP was positive, and this may be due to the presence of unreacted [NH.sub.2] groups available on the surface of chitosan for the attachment of other negatively charged molecules (Shard et al. 2014).

The average zeta potential of chitosan nanoparticles obtained from ratio 3:1 of chitosan to TPP was 43.93 mV (Fig. 8A).

However, the average zeta potential of chitosan nanoparticles that resulted from ratio 1:3 of chitosan to TPP was 40.24 mV

(Fig. 8B), whereas, using concentration 1:1 chitosan to TPP resulted in an average zeta potential of 1.49 mV (Fig. 8C), which was considered as the best value for the zeta potential.

CONCLUSION

In the present study, the method of extraction of chitosan from the exoskeleton wastes of Procambarus clarkii gave rise to chitosan with degree of deacetylation 87%. Viscometry indicated that the obtained chitosan had a low Mw and this can be useful in many applications.

It was clear that as the concentration of the chitosan or the TPP increased, the particle size and the aggregation of the nanoparticles increased. Results revealed that there is a significant effect of the concentration of both chitosan and TPP on the particle size and dispersion of the resultant nanoparticles. The best physicochemical characteristics were obtained by adding chitosan to TPP in a ratio of 1:1. Moreover, the chitosan nanoparticles appear spherical in shape with smooth surfaces. The small particle size and narrow range distribution of the obtained chitosan nanoparticles may increase the chance for easy and efficient manipulation for several applications in various fields. In addition, the obtained positively charged chitosan nanoparticles can be used for the delivery of negatively charged compounds.

ACKNOWLEDGMENTS

This article is part of the results of Project ID: 9450781006-

ASRT JESOR 2015, entitled "Novel extraction and industry from wastes and exoskeleton of the introduced freshwater crawfish," so we thank the Academy of Scientific Research and Technology for funding this work. Mrs. Diana Fakhry, Faculty of Agriculture, Biochemistry Department, Cairo University, is most appreciated for introducing a novel method for dissolving the chitosan nanoparticles for TEM.

LITERATURE CITED

Abdou, E. S., K. S. Nagy & M. Z. Elsabee. 2008. Extraction and characterization of chitosan from local sources. Bioresour. Technol. 99:1359-1367.

Alishahi, A., A. Mirvaghefi, M. Rafiee, H. Farahmand, S. A. Shojaosadati, F.

Dorkoosh & M. Z. Elsabee. 2011. Shelf life and delivery enhancement of vitamin C using chitosan nanoparticles. Food Chem. 126:935-940.

Aranaz, I., M. Mengibar, R. Harris, I. Panos, B. Miralles, G. G.

Acosta & A. Heras. 2009. Functional characterization of chitin and chitosan. Curr. Chem. Biol. 3:203-230.

Arbia, W., L. Arbia, L. Adour & A. Amrane. 2013. Chitin recovery using biological methods. Food Technol. Biotechnol. 51:12-25.

Bravo-Anaya, L. M., J. F. A. Soltero & M. Rinaudo. 2016. DNA/chitosan electrostatic complex. Int. J. Biol. Macromol. 88:345-353.

Brugnerotto, J., J. Lizardi, F. M. Goycoolea, W. Arguelles-Monal, J. Desbrieres & M. Rinaudo. 2001. An infrared investigation in relation with chitin and chitosan characterization. Polymer. 42:3569-3580.

Calvo, P., C. Remunan-Lopez, J. L. Vila-Jato & M. J. Alonso. 1997.

Novel hydrophilic chitosan-polyethylene oxide nanoparticles as protein carrier. J. Appl. Polym. Sci. 63:125-132.

Ghannam, H., A. S. Talab, N. V. Dolganova, A. M. S. Hussein & N. M.

Abdelmaguid. 2016. Characterization of chitosan extracted from different crustacean shell wastes. J. Appl. Sci. 16:454-461.

Ibrahim, A. M. & M. T. Khalil. 2009. The red swamp crayfish in Egypt

(a fast spreading freshwater invasive crustacean). Publication of the Center of Reseach & Studies of Protectorates, Ain Shams University, Cairo, Egypt. 153 pp.

Jang, M. K., B. G. Kong, Y. I. Jeong, C. H. Lee & J. W. Nah. 2004.

Physicochemical characterization of a-chitin, 13-chitin, and y-chitin separated from natural resources. J. Polym. Sci. A Polym. Chem. 42:3423-3432.

Jiang, J. C. & M. Q. Xu. 2006. Kinetics of heterogeneous deacetylation of 13-chitin. Chem. Eng. Technol. 29:511-516.

Jimenez-Fernandez, E., A. Ruyra, N. Roher, E. Zuasti, C. Infante & C.

Fernandez-Diaz. 2014. Nanoparticles as novel delivery system for vitamin C administration in aquaculture. Aquaculture. 432:426-433.

Kanauchi, O., K. Deuchi, Y. Imasato, M. Shizukuishi & E. Kobayashi. 1995. Mechanism for the inhibition of fat digestion by chitosan and for the synergistic effect of ascorbate. Biosci. Biotechnol. Biochem. 59:786-790.

Kaya, M., T. Baran, A. Mentes, M. Asaroglu, G. Sezen & K. O. Tozak. 2014. Extraction and characterization of a-chitin and chitosan from six different aquatic invertebrates. Food Biophys. 9:145-157.

Kaya, M., I. Sargin, K. O. Tozak, T. Baran, S. Erdogan & G. Sezen. 2013. Chitin extraction and characterization from Daphnia magna resting eggs. Int. J. Biol. Macromol. 61:459-464.

Kumar, A., A. Vimal & A. Kumar. 2016. Why chitosan? From properties to perspective of mucosal drug delivery. Int. J. Biol. Macromol. 91:615-622.

Lavall, R. L., O. B. G. Assis & S. P. Campana-Filho. 2007.13-chitin from the pens of Loligo sp.: extraction and characterization. Bioresour.

Technol. 98:2465-2472.

Morin, A. & A. Dufresne. 2002. Nanocomposites of chitin whiskers from Riftia tubes and poly (caprolactone). Macromolecules. 35:2190-2199.

Ngan, L. T. K., S.-L. Wang, D. M. Hiep, P. M. Luong, N. T. Vui, T. M.

Dinh, T. M. & N. A. Dzung. 2014. Preparation of chitosan nanoparticles by spray drying, and their antibacterial activity. Res. Chem.

Intermed. 40:2165-2175.

No, H. K. & S. P. Meyers. 1989. Crayfish chitosan as a coagulant in recovery of organic compounds from seafood processing streams.

J. Agric. Food Chem. 37:580-583.

No, H. K., S. P. Meyers & K. S. Lee. 1989. Isolation and characterization of chitin from crawfish shell waste. J. Agric. Food Chem. 37:575-579.

Ostolska, I. & M. Wisniewska. 2014. Application of the zeta potential measurements to explanation of colloidal Cr203 stability mechanism in the presence of the ionic polyamino acids. Colloid Polym.

Sci. 292:2453-2464.

Paulino, A. T., J. I. Simionato, A. J. Garcia & J. Nozaki. 2006. Characterization of chitosan and chitin produced from silkworm crystalides.

Carbohydr. Polym. 64:98-103.

Riegger, B. R., B. Baurer, A. Mirzayeva, G. E. M. Tovar & M. Bach. 2018. Systematic approach for preparation of chitosan nanoparticles via emulsion crosslinking as potential adsorbent in wastewater treatment. Carbohydr. Polym. 180:46-54.

Rinaudo, M. 2006. Chitin and chitosan: properties and applications.

Prog. Polym. Sci. 3:603-632.

Shard, P., A. Bhatia & D. Sharma. 2014. Optimization and physicochemical parameters on synthesis of chitosan nanoparticles by ionic gelation technique. Int. J. Drug Deliv. 6:58-63.

Shim, Y., H. J. Lee, S. Lee, S. H. Moon & J. Cho. 2002. Effects of natural organic matter and ionic species on membrane surface charge. Environ. Sci. Technol. 36:3864-3871.

Simpson, B. K., N. Gagne & M. V. Simpson. 1999. Bioprocessing of chitin and chitosan. In: Martin, A. M., editor. Fisheries processing.

Boston, MA: Springer. pp. 155-173.

Toan, N. V. & T. T. Hanh. 2013. Application of chitosan solutions for rice production in Vietnam. Afr. J. Biotechnol. 12:382-384.

Wang, Y., Y. Chang, L. Yu, C. Zhang, X. Xu, Y. Xue, Z. Li & C. Xue. 2013. Crystalline structure and thermal property characterization of chitin from Antarctic krill (Euphausia superba). Carbohydr. Polym. 92:90-97.

Wang, Y. & J. Li. 2011. Effects of chitosan nanoparticles on survival, growth and meat quality of tilapia, Oreochromis nilotica. Nanotoxicology. 5:425-431.

Zhao, D., S. Yu, B. Sun, S. Gao, S. Guo & K. Zhao. 2018. Biomedical applications of chitosan and its derivative nanoparticles. Polymer. 10:462.

MARWA M. EL-NAGGAR, (1*) WAEL S. I. ABOU-ELMAGD, (2*) ASHRAF SULOMA, (3) HAMZA A. EL-SHABAKA, (1) MAGDY T. KHALIL (1) AND FAWZIA A. ABD EL-RAHMAN (1)

(1) Departments of Zoology and (2) Chemistry, Faculty of Science, Ain Shams University, Abassia, Cairo 11566, Egypt; (3) Department of Animal Production, Fish Nutrition Lab, Faculty of Agriculture, Cairo University, Giza 12613, Egypt

(*) Corresponding authors. E-mails: m.ainggar@hotmail.com or waelmagd97@yahoo.com

DOI: 10.2983/035.038.0220
TABLE 1.
Relative viscosity ([[eta].sub.rel]) and specific viscosity
([[eta].sub.sp]) values for different concentrations of chitosan.

C%   T      t/[t.sub.0]  [[eta].sub.specific]  [[eta].sub.reduced]
                         = t/[t.sub.0] - 1     = [[eta].sub.sp]/c

1.3  41.96  6.34         5.34                  4.11
1    26.66  4.03         3.03                  3.03
0.7  17.25  2.60         1.60                  2.28
0.5  15.79  2.38         1.38                  2.72
COPYRIGHT 2019 National Shellfisheries Association, Inc.
No portion of this article can be reproduced without the express written permission from the copyright holder.
Copyright 2019 Gale, Cengage Learning. All rights reserved.

Article Details
Printer friendly Cite/link Email Feedback
Author:El-Naggar, Marwa M.; Abou-Elmagd, Wael S.I.; Suloma, Ashraf; El-Shabaka, Hamza A.; Khalil, Magdy T.;
Publication:Journal of Shellfish Research
Article Type:Report
Geographic Code:7EGYP
Date:Aug 1, 2019
Words:4970
Previous Article:THE EFFECTS OF SEASONAL TEMPERATURE AND PHOTOPERIOD MANIPULATION ON REPRODUCTION IN THE EASTERN ELLIPTIO ELLIPTIO COMPLANATA.
Topics:

Terms of use | Privacy policy | Copyright © 2020 Farlex, Inc. | Feedback | For webmasters