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Preparation of Chitin Nanofibers with a Uniform Width as r-Chitin from Crab Shells Shinsuke Ifuku,*,† Masaya Nogi,‡ Kentaro Abe,‡ Masafumi Yoshioka,† Minoru Morimoto,§ Hiroyuki Saimoto,† and Hiroyuki Yano‡ Department of Chemistry and Biotechnology, Graduate School of Engineering, Tottori University 4-101 Koyamac-cho Minami, Tottori, Japan, Research Center for Bioscience and Technology, Tottori University 4-101 Koyamac-cho Minami, Tottori, Japan, and Research Institute for Sustainable Humanosphere, Kyoto University, Uji, Japan Received February 5, 2009; Revised Manuscript Received April 3, 2009
Chitin nanofibers were prepared from dried crab shells by a simple grinding treatment in a never-dried state under an acidic condition after the removal of proteins and minerals. The obtained nanofibers were observed by FE-SEM and found to have a uniform width of approximately 10-20 nm and high aspect ratio; both these findings were similar to those for nanofibers from prawns. Furthermore, it was confirmed that the nanofibers were extracted from the natural chitin/protein/mineral composites of crab shell in their original state. That is, the N-acetyl group was not removed and the R-chitin crystal structure was maintained, as confirmed by elemental analysis data, FT-IR spectra, and X-ray diffraction profiles.
Introduction Chitins are highly abundant biomacromolecules occurring mainly in the exoskeletons of shellfish and insects and are synthesized at a rate of 1010 to 1011 tons per year.1 Although native chitin is a semicrystalline biopolymer with microfibrillar morphology and excellent material properties, most of the biomass is thrown away as industrial waste (shrimp and crab shells) without effective utilization. Thus, it is important to make efficient use of this biomass resource as a natural and environmentally friendly material. Because of their linear (1,4)-β-N-acetyl glycosaminoglycan structure with two hydroxyl groups and an acetamide group, native chitins in these crustacean shells are highly crystalline with strong hydrogen bonding, and are arranged as R-chitin microfibrils in an antiparallel fashion. These microfibrils consist of nanofibers about 2-5 nm diameter and about 300 nm in length embedded in a protein matrix.2,3 Because chitin nanofibers are considered to have great potential, various methods have been employed for their preparation, including acid hydrolysis,1,4 TEMPO-mediated oxidation,5 an ultrasonic technique,6 and an electrospinning method.7 However, the nanofibers obtained by these methods were substantially different from the native chitin nanofibers in terms of width, aspect ratio, crystallinity, chemical structure, and/or homogeneity. Although Fan et al. reported a procedure for preparing chitin nanofibers 3-4 nm in width from squid pen β-chitin by ultrasonication treatment under acidic conditions,8 crystallinity of the nanofibers from the squid pen is relatively low. In addition, the biomass quantity of the pen is considerably lower than those of crab and shrimp shells. Recently, Abe et al. succeeded in obtaining cellulose nanofibers with a uniform width of approximately 15 nm from wood.9 Because cellulose nanofibers in cell walls of wood are embedded * To whom correspondence should be addressed. Phone and Fax: +81857-31-5592. E-mail:
[email protected]. † Department of Chemistry and Biotechnology, Tottori University. ‡ Research Institute for Sustainable Humanosphere, Kyoto University. § Research Center for Bioscience and Technology, Tottori University.
in matrix substances such as lignin and hemicellulose, the cellulose can be isolated by removal of the matrix substances as cellulose nanofibers after a very simple mechanical treatment. Similarly, the exoskeletons of crustacea have a strictly hierarchical organization consisting of crystalline R-chitin nanofibers and various types of proteins and minerals, as shown in Figure 1.2,3,10 Thus, chitin nanofibers, like cellulose nanofibers, are encased in embedding matrix components. Although the authors described that the preparation method can be used to isolate cellulose nanofibers from any natural plant containing lignin and hemicellulose, we consider that this isolation method would be universally applicable to any natural nanofiber source consisting of nanofibers and other embedding matrixes. Accordingly, we here studied the extraction of natural R-chitin nanofibers with a uniform width of 10-20 nm from crab shells.
Experimental Section Materials. Purified R-chitin was purchased from Nacalai Tesque, and the other chemicals were purchased from Aldrich or Kanto chemical and used as received. Removal of Matrix Components. Dried crab shell powder of Paralithodes camtschaticus (Red king crab) was used for this study (Kawai Hiryo Co., Japan). The crab shells were purified to prepare the chitin nanofibers according to the general methods.1,11 First, crab shell powder was refluxed in 5 wt % of potassium hydroxide for 6 h under vigorous stirring to remove most of the proteins. This suspension was cooled to room temperature, then filtered and washed thoroughly with distilled water. Next, the chitin samples were treated with 7% hydrochloric acid solution for 2 days at room temperature to remove the mineral salts, which consisted mainly of calcium carbonate. After filtration and rinsing with an abundance of distilled water, the treated sample was dispersed and boiled in a 5% KOH solution for 2 days to remove residual proteins completely. The pigment composition in the sample was then removed using 1.7 wt % of sodium chlorite in 0.3 M sodium acetate buffer for 6 h at 80 °C followed by filtration and washing with distilled water. The yield of chitin from the crab shells was estimated to be 12.1 wt %. Fibrillation. The purified wet chitin from dry crab shells was dispersed in water at 1 wt %, and acetic acid was added to adjust the
10.1021/bm900163d CCC: $40.75 2009 American Chemical Society Published on Web 04/27/2009
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Figure 1. Schematic presentation of the exoskeleton structure of crustacean shell.
pH value to 3 to facilitate fibrillation. The suspension was treated with a domestic blender. Finally, the slurry of 1 wt % purified chitin was passed through a grinder (MKCA6-3; Masuko Sangyo Co., Ltd.) at 1500 rpm.9,12 Grinder treatment was performed with a clearance gauge of -1.5 (corresponding to a 0.15 mm shift) from the zero position, which was determined as the point of slight contact between the grinding stones. In principle, there is no direct contact between grinding stones due to the presence of chitin suspension. Scanning Electron Microscopy (SEM). The suspension of chitin fibrils was subjected to oven-drying at 105 °C after replacement from water to ethanol, and the obtained sheets were coated with an approximately 2 nm layer of platinum by an ion sputter coater and observed with a field emission scanning electron microscope (JSM6700F; JEOL, Ltd.). Elemental Analysis. The DS values of amino group of the chitin nanofibers were calculated from the C and N contents in the elemental analysis data by using elemental analyzer (Elementar Vario EL III, Elementar). X-ray Diffraction. Equatorial diffraction profiles were obtained with Ni-filtered Cu KR from an X-ray generator (Shimadzu XRD-6000) operating at 40 kV and 30 mA. Fourier Transform Infrared (FT-IR) Spectroscopy. Infrared spectra of the chitin samples were recorded using potassium bromide pellets with an FT-IR spectrometer (FTIR 8300, Shimadzu). All the spectra were obtained by accumulation of 20 scans, with resolution of 2 cm-1, at 400-4000 cm-1.
Results and Discussion Preparation of Chitin Nanofibers. In general, the exoskeletons of crustacea have a strictly hierarchical organization which reveals various structural levels, as shown in Figure 1.2,3,13,14 The molecular level is the chitin itself. These chitin molecules are aligned in an antiparallel manner that gives rise to R-chitin crystals in the form of thinner nanofibers of about 2-5 nm diameter. These nanofibers are wrapped in protein layers, which can be regarded as the next level. The next level in the scale consists of the clustering of some of these nanofibers into chitin/ protein fibers of about 50-300 nm thicker diameter. The next step is the formation of a planar woven and branched network
of such chitin-protein fibers with a variety of thickness. These strands are embedded in a variety of proteins and minerals. The minerals mainly consist of crystalline calcium carbonate. Thickness of strands vary widely among crustaceans. Furthermore, these woven and network planes form twisted plywood pattern. This structure is formed by the helicoidal stacking sequences of the fibrous chitin-protein layers. The thickness of the twisted plywood layer corresponds to a certain stacking density of planes, which are gradually rotated about their normal axis. Dried crab shell powder was used as a starting material; this powder is commercially available as a fertilizer at low cost. First, the matrix substances were removed from the crab shell powder. To extract chitin nanofibers from such a natural composite, proteins and minerals were removed according to the conventional method using KOH and HCl solutions, respectively.1,11 It is well-known that almost these matrix substances can be removed by the alkali and acid treatment.15 Because the process of drying chitin nanofibers generates strong hydrogen bonding between the bundles, which makes it difficult to obtain thin and uniform nanofibers, the material was kept wet after the removal of the matrix, as described in Abe et al.9 Figure 2 shows SEM images of the crab shell surface after the removal of the matrix. Although these images do not represent the whole chitin fiber structure from crab shell because the crab shell has hierarchical complex structure, these images should be from endoskeleton, which is the main part of the crab shell. The samples were prepared by oven drying after solvent replacement from water to ethanol to prevent coagulation between chitin fibers. Although the crab shell structure was still maintained, we could observe chitin nanofibers approximately 10 to 20 nm wide as shown in Figure 1, which were bundles of crystalline chitin nanofibers having a width of 2-5 nm with strong interaction between each thinner nanofibers. Furthermore, as shown in Figure 2b, thicker chitin-protein fibers of around 100 nm diameter were confirmed to be bundles of nanofibers of 10-20 nm in width. The obtained chitin slurry in distilled water with a concentration of 1 wt % was passed through a grinder under a neutral
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Figure 2. FE-SEM micrographs of crab shell surface after the removal of matrix (without grinder treatment). The length of the scale bar is (a) 1000 and (b) 100 nm, respectively.
pH condition, with a careful adjustment of the clearance between grinding stones. The oven-dried chitin slurry was observed using SEM after platinum coating. Although we have already succeeded in preparing chitin nanofibers with a uniform width of 10-20 nm from prawn shells of Penaeus monodon (black tiger prawns) by using the same method as shown in Figure 3 (the findings will be reported elsewhere), the widths of the fibers derived from crab shell after grinder treatment were widely distributed over a range from 10 to 100 nm (Figure 4a). Twisted plywood structure (Figure 1) looks broken completely by grinder treatment after removal of matrix substances, in which the chitin-protein fiber was embedded. As a result, the thicker fibers with the width of about 100 nm derived from chitin-protein fibers could be isolated. However, the thicker fibers, corresponding to chitin-protein fibers, which were bundles of nanofibers of 10-20 nm in width (Figure 2), were not fibrillated successfully by grinder treatment even though the protein layers were removed under a never-dried condition. A plausible explanation for this difference in fiber thickness is as follows. The cuticle is made up of mainly two parts, the exocuticle (outer) and the endocuticle (inner), as shown in Figure 1. Although the exocuticle has a chitin-protein matrix with a very fine twisted plywood-type structure, the endocuticle is characterized by a much coarser matrix structure with fibers of thicker diameter.14 The endocuticle makes up about 90 vol % of the crab exoskeleton.3 On the other hand, the exoskeleton of Natantia, which has a semitransparent soft shell, such as black tiger prawn, is mainly made up of fine exocuticle part.16,17 As
Figure 3. FE-SEM micrographs of chitin nanofibers from black tiger prawn after one pass through the grinder at neutral pH condition. The length of the scale bar is (a) 1000 and (b) 100 nm, respectively.
a result, due to the differences of cuticle structure and fiber thickness, it is more difficult to fibrillate the crab shell than the black tiger prawn shell by grinder treatment, because the crab shell has a coarser and more complex hierarchical structure than the prawn shell with greater fiber thickness in the endocuticle region. Fan et al. reported a preparation method for chitin nanofibers from squid pen β-chitin in water at pH 3-4.8 Cationization of the C2 amino groups in the β-chitin at pH 3-4 is important to maintain the stable dispersion state by electrostatic repulsions. Therefore, we considered that the purified R-chitin from crab shell would also be well dispersed under an acidic condition by cationization of the amino groups on the fiber surface, which in turn would facilitate fibrillation into chitin nanofibers. Thus, the pH value of the purified chitin suspension was adjusted to about 3 by the addition of acetic acid, and then the protein-free chitin was subjected to grinder treatment. Interestingly, the chitin slurry thus obtained formed a gel after a single grinder treatment as with cellulose nanofibers, suggesting fibrillation was accomplished, because of its high dispersion property in water and high surface-to-volume ratio of nanofiber.9 Figure 4b and c shows SEM images of the oven-dried chitin gel. The isolated chitin is observed as highly uniform nanofibers with a width of 10-20 nm, which were similar to nanofibers from black tiger prawn, suggesting the fibrillation process was facilitated in acidic
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Figure 4. FE-SEM micrographs of chitin nanofibers from crab shell after one pass through the grinder (a) without acetic acid (pH 7) and (b,c) with acetic acid (pH 3). The length of the scale bar is (a) 200, (b) 200, and (c) 100 nm, respectively.
Figure 5. FT-IR spectra of (a) commercially available pure R-chitin chemical derived from crab shell, (b) newly prepared chitin nanofibers from crab shell by removal matrix components, and (c) dried crab shell powder derived from red king crab without purification.
solution as expected. It should be emphasized that the width of the nanofibers extracted in this study corresponds to the width of the chitin nanofibers observed from crab shell surface after removal of matrix as in Figure 2b. Furthermore, because broken fibers are not observed even over a wide observation area, the aspect ratios of the nanofibers are very high. These results indicate that chitin nanofibers were successfully isolated from crab shells without altering their natural shape. Thus, the neverdried process that has previously been used to extract the cellulose nanofibers from wood cell walls was here shown to be applicable for the preparation of chitin nanofibers from crustacean shells by mechanical grinding under an acidic condition. Characterization of Chitin Nanofibers. The degree of N-acetylation of the obtained nanofibers was calculated by comparing the C and N content in the elemental analysis data (found: C, 44.26; H, 6.86; N, 6.54%) and was found to be 0.95. Thus, the degree of substitution of amino group was just 0.05, indicating deacetylation did not occur after the treatment sequence, and chitin nanofibers were obtained as natural fibers. Interestingly, even though ratio of the amino group was very small, cationization of amino groups in the chitin from crab shell at pH 3 facilitate fibrillation of chitin fiber and maintain the stable dispersion state in the water by electrostatic repulsions between the chitin nanofibers with cationic surface charges.
Figure 6. X-ray diffraction profiles of (a) commercially available R-chitin chemical derived from crab shell, (b) newly prepared chitin nanofibers from crab shell by removal of matrix components, and (c) dried crab shell powder derived from red king crab without purification.
Figure 5 shows the normalized FT-IR spectra of (a) commercially available pure R-chitin chemical derived from crab shell, (b) newly prepared chitin nanofibers from crab shell by removal of matrix components, and (c) dried crab shell powder derived from red king crab without purification. The spectrum of the dried crab shell powder is very different from the other two spectra because of the matrix components included in the shell. On the other hand, spectrum of newly prepare chitin nanofibers is in excellent agreement with the spectrum of commercial pure R-chitin, indicating that the matrix, protein, and minerals are well-removed by conventional processing. In particular, the absorption band at 1420 cm-1 derived from protein has completely disappeared, suggesting that these treatments were sufficient to eliminate all the proteins and to obtain pure chitin. Furthermore, the OH stretching band at 3482 cm-1, NH stretching band at 3270 cm-1, amide band I at 1661 and 1622 cm-1, and amide II band at 1559 cm-1 of the chitin nanofibers are observed. These strong absorption peaks in the carbonyl region are especially characteristic of anhydrous R-chitin.1 Figure 6 shows normalized X-ray diffraction profiles of (a) commercially available pure R-chitin chemical derived from crab shell, (b) newly prepared chitin nanofibers from crab shell by removal of matrix components, and (c) dried crab shell powder derived from red king crab without purification. The diffraction peak at 29.6°, which is typical for calcium carbonate, was completely absent from the profile of chitin
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nanofibers, indicating the mineral component was well removed from newly prepared chitin nanofibers. The crystallinity of prepared chitin nanofibers appeared to be lower than that of commercial chitin because the amorphous part of commercial pure chitin is considerably removed by acid hydrolysis through the purification process. However, the four diffraction peaks of chitin nanofibers observed at 9.5, 19.5, 20.9, and 23.4°, which corresponded to 020, 110, 120, and 130 planes, respectively, are typical crystal patterns of R-chitin18 and are closely coincident with commercial R-chitin. Thus, the chitin nanofibers were extracted from the crab shell, and the original molecular structure and R-chitin crystalline structure was maintained even after the removal of the matrix and the grinder treatments.
Conclusion Chitin nanofibers were prepared from dried-crab shell, which has a complex hierarchical structure with a uniform width of approximately 10-20 nm by conventional chemical treatment, followed by mechanical treatment. This study demonstrates that the grinding treatment in a never-dried state after the removal of the matrix is applicable to not only wood but also crab shell to isolate nanofibers, and the mechanical treatment under an acidic condition is the key to fibrillate thicker fibers in the endocuticle region. Because this simple but powerful method allows us to obtain homogeneous chitin nanofibers in their original state from waste crab shell in large amounts, we expect that these nanofibers with a uniform width and very high
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surface-to-volume ratio will be developed into novel green nanomaterials.
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