Spherical cellulose nanoparticles preparation from

Powder Technology 261 (2014) 232–240

Contents lists available at ScienceDirect

Powder Technology journal homepage: www.elsevier.com/locate/powtec

Spherical cellulose nanoparticles preparation from waste cotton using a green method Tayebeh Fattahi Meyabadi a,⁎, Fatemeh Dadashian a, Gity Mir Mohamad Sadeghi b, Hamid Ebrahimi Zanjani Asl a a b

Department of Textile Engineering, Amirkabir University of Technology, Tehran, Iran Department of Polymer Engineering & Color Technology, Amirkabir University of Technology, Tehran, Iran

a r t i c l e

i n f o

Article history: Received 10 September 2013 Received in revised form 21 January 2014 Accepted 8 April 2014 Available online 18 April 2014 Keywords: Waste cotton fibers Spherical nanoparticles Enzymatic hydrolysis Sonication

a b s t r a c t Cellulose nanoparticles with spherical morphology were prepared through enzymatic hydrolysis of waste cotton fibers and sonication treatment. The nanoparticles were characterized with viscometer, field emission scanning electron microscope, particle size analyzer, Fourier Transform Infrared Spectroscopy, X-ray diffraction, and thermogravimetric analysis. The results indicated that the average particle size of cotton fibers after hydrolysis and sonication process was 0.526 μm and less than 100 nm, respectively. The crystalline structure of cellulose was preserved following hydrolysis and ultrasonic processes, with the decline in its polymerization degree, though. Additionally, there was no significant change in the crystallinity. Moreover, thermal degradation showed that the hydrolysis and sonication of cotton would have no effect on the chemical fingerprints of the cellulose. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Cellulose, the most abundant biopolymer in the world, has attracted much research interest because of its biosynthesis, chemistry, and ultra structure [1–4]. It is noteworthy that between 1010 and 1011 tons of the cellulose are reportedly synthesized and destroyed each year worldwide [5–8]. Thus, the researchers have developed efficient systems for recycling and utilizing cellulose waste in chemical, food, cosmetic, textile, and polymer industries to deal with the respective environmental and economic concerns [9–13]. There are various methods to further utilize, recycle, and degrade the cellulose waste, including enzymatic and ultrasonic degradation [14]. Of course, enzyme application has gained considerable attention in producing useful products and developing new processes [3,9,15]. Cellulases are a class of enzymes acting as a catalyst for the hydrolysis of the cellulose [16]. Cellobiohydrolase is a sub-group of the cellulase that attacks the crystalline structure of cellulose. Also, another subgroup of the cellulase is endogluconase that catalyzes the hydrolysis of amorphous cellulose. Consequently, enzymatic degradation of the cellulose ends up with a decrease in the chain length of the cellulose [3,9,17, 18]. Ultrasound irradiation (or sonication) is defined as a sound with a higher frequency than the human hearing range, i.e. N 20 kHz. The ultrasound role in degrading polymeric solutions has been well documented, especially in synthetic polymers [19]. It is, also, highly efficient with the ⁎ Corresponding author. Tel.: +98 2164542691; fax: +98 2166400245. E-mail address: [email protected] (T. Fattahi Meyabadi).

http://dx.doi.org/10.1016/j.powtec.2014.04.039 0032-5910/© 2014 Elsevier B.V. All rights reserved.

cellulose degradation through disruption of weak β-glycosidic bonds [19,20]. Today, waste cotton fiber is one of the cellulosic wastes considered for reusing and recycling. Obviously, cotton contains the highest percentage of the cellulose (N 95%) among various cellulosic sources, and occupies almost 1/3 of the global market of textile fibers with annual output of over 23 million tons per year [13,17,18]. Therefore, cotton and its waste may be considered proper for the production of ethanol, biogas, organic acids [12,13,18], microcrystalline cellulose [21], cellulose powders [22] and cellulose nanoparticles [18,23]. Many researchers have reported on the preparation and characterization of the cellulose nanoparticles (with different terms including whiskers, nanowhiskers, nanospheres, nanopowders, nanocrystals, microfibrils, nanofibrils, nanofibers …) prepared from various cellulosic sources [2,6,8,14,21,23–31]. The cellulose nanoparticles have major advantages, such as non-toxicity, low cost, low density, higher availability and higher specific strength as well as modulus [1,8,17,27, 32]. As a result, they can be applied in many areas such as in nanocomposites, tissue engineering scaffolds, and filtration media, and in the pharmaceutical, health & cosmetic, food and automotive industries [1,8,27,32,33]. Many studies suggest that the prepared cellulose nanoparticles are reportedly rod shaped called whiskers [8,18,29,31,32]. Nevertheless, there are a few studies in terms of preparation of spherical cellulose nanoparticles [2,14,30]. To the best of our knowledge, spherical cellulose nanoparticles have been previously produced only with acid hydrolysis which is not an environment friendly process. However, it requires two additional processes (recycling the acid and neutralizing its carbon)

T. Fattahi Meyabadi et al. / Powder Technology 261 (2014) 232–240

to be an environment friendly one. The present paper reports a simple and versatile approach for preparing the spherical nanoparticles from waste cotton fibers using enzymatic treatment coupled with ultrasonication. Needless to say, both the enzymatic hydrolysis and the ultrasound treatments are deemed to be green processes. Therefore, they could replace conventional chemical treatment (acid hydrolysis) to produce the cellulose nanoparticles leaving zero environmental pollution. The applied method in this study is a top-down method for preparing the cellulose nanoparticles. It is also applicable for many other natural fibrous materials. 2. Experiment

233

The excess water of suspension was excluded with centrifuging at 6000 rpm for 10 min as far as possible. Then, the concentrated suspension was frozen and put into a freeze-dryer (Christ Alpha 1-2 LD Freeze dryer (Germany)) for 24 h to remove the remained water. The dried powder was then stored in desiccator for characterizations. 3. Characterization 3.1. Fluidity analysis The fluidity (α viscosity−1) of 0.5 and 2% solutions of the samples in cuprammonium was measured using a standard Shirely X-type CUAM viscometer in accordance with cotton standard (Cotton, Silk and Manmade Fibers Research Association).

2.1. Materials 3.2. Microscopic analysis The waste cotton fibers attained from comber noil of a Textile Mill were used to prepare the nanoparticles. The cellulase enzyme used in this work was a Cellusoft L from Novozymes (Biotechnology incorporated) with activity of 750 EGU/g. It was a full-value cellulase system encompassing cellobiohydrolases, endoglucanases, and glucosidases and would completely hydrolyze both the amorphous and crystalline cellulose. Cellusoft L was chosen due to its lower level of glucosidase enzyme, since our aim was to avoid complete hydrolysis to monomer (glucose). In addition, technical grade and high purity nonionic surfactant (Irgasol NA) supplied from Ciba Co. (Tehran, Iran). It was an ethoxylated fatty alcohol with a density of 1.0 g/mL. Also, other chemicals such as acetic acid (99.8% purity, analytical grade) and anhydrous sodium acetate (99.99% suprapur, analytical grade) were purchased from Merck Co. (Germany). 2.2. Enzymatic hydrolysis The waste cotton fibers were chopped into pieces of less than 2 mm in length, washed with nonionic surfactant (1 g/L Irgasol) for an hour, rinsed with distilled water, and then oven-dried at 105 °C for 3 h. Using the result of response surface methodology, the hydrolysis was performed in 0.05 M acetate buffer (pH = 4.8) at 48 °C under optimum conditions including: 2.3% cellulase enzyme and 5 g/L substrate concentration for 175 h [34]. Having been heated up to 80 °C for 15 min, the enzymatic hydrolysis was terminated. Besides, using a Hettich centrifuge EBA 20 (Tuttlingen, Germany), the hydrolysate was washed with distilled water through repeated centrifugations at 5000 rpm. 2.3. Sonication The suspension of the hydrolyzed cotton was sonicated with Heilscher Ultrasonics UP200S (200 W, 24 kHz) and sonotrode S14 (tip diameter of 14 mm) at 50% amplitude. In each batch, 50 mL of the 5 g/L suspension was sonicated for 15 min. Then, it was allowed to remain for 20 min after the sonication. The top turbid part was then decanted. Subsequently, water was added to the residue to reach the final volume of 50 mL. After stirring, the sonication was done for another 15 min cycle and the top turbid part was decanted. In the third cycle, there was no sign of turbidity in the suspension; since turbidity is a sign of the presence of the nanoparticles, it was found that as many as two cycles of sonication (30 min) were sufficient to attain almost the entire nanoparticles.

Optical micrographs were obtained using a ZEISS optical microscope (200×) by placing a drop of suspension on a glass slide and air-drying. The surface morphology of the samples was also examined with a Field emission scanning electron microscope (FESEM) (Hitachi, model S4160, voltage 15 kV, Japan), after gold coating. 3.3. Particle size analysis The particle size distribution of hydrolyzed cotton was measured using Mastersizer 2000 particle size analyzer (Malvern Instruments Limited, Malvern, UK), with a detection range of 0.02–2000 μm. A 0.5% w/v suspension of the hydrolyzed sample was used and d (0.5) was applied to report the median particle size. d (0.5) means that 50% of all particles were finer than this size. Additionally, Zetasizer analyzer (Malvern Instruments Limited, Malvern, UK) with a detection range of 0.3 nm–10 μm was used to measure the particle size distribution of the cellulose nanoparticles. 100 μL of the nanoparticle suspension was dispersed in 4 mL distilled water and the particle size distribution of nanoparticles was determined. 3.4. Fourier Transform Infrared Spectroscopy (FTIR) The FTIR spectra of nanoparticles were measured with a Thermo Nicolet Nexus 870 spectrophotometer in absorbance mode, ranging 4000–400 cm−1 at a resolution of 4 cm−1. 40 scans were recorded per sample. The untreated cotton and hydrolyzed cotton were also analyzed for comparison purpose. Obtained spectra were normalized to the absorbance of the band at 2902 cm−1 due to any obtained changes in this band among all examined samples [35]. In doing so, the Thermo Nicolet OMNIC software was used to analyze the data of FTIR spectroscopy. 3.5. X-ray diffraction analysis (XRD) The X-ray diffraction analysis was used to investigate the crystalline structural changes of the cellulose created with the enzymatic and ultrasound treatments. Hence, the crystalline phases present in the samples were determined using an X-ray diffractometer (Equinox 3000, France) at 40 kV and 30 mA. Then, the diffracted intensity of Cu Kα radiation (λ = 1.5418 Å) was recorded in a 2θ range between 5° and 40° with a step scan of 0.03° and a step time of 2 s. Plus, the crystallinity index (CrI) of samples was calculated from diffraction intensity data using the empirical method proposed by Segal et al. [36] (Eq. (1)) for native cellulose:

2.4. Freeze drying CrI ¼ ððIc−IaÞ=IcÞ 100 The required volume of the hydrolyzed cotton and nanoparticles suspensions was separated for the optical microscopic and particle size analyses, and the rest was freeze dried according to the following method.

ð1Þ

where Ic is the peak intensity of crystal plane (002) (maximum intensity at 2θ = 22.5°) and Ia is the minimum in intensity corresponding to amorphous content at 2θ = 18° [36].

234

T. Fattahi Meyabadi et al. / Powder Technology 261 (2014) 232–240 Table 1 The fluidity of different samples. Sample

Fluidity (Poise−1) (cotton std ± 0.1)

Untreated cotton Hydrolyzed cotton Cotton nanoparticles

2.21 45.18 53.34

The interplanar distance (d-spacing) and average crystallite size (Lc) of samples were also calculated using the Bragg low (Eq. (2)) and the Scherrer equation (Eq. (3)), respectively: d‐spacing ¼ λ=ð2 sinθÞ

ð2Þ

Lc ¼ Kλ=β cosθ

ð3Þ

where λ is the wavelength of the X-ray used (1.5418 Å); θ, the diffraction angle corresponding to (002) crystal plane; K, the shape factor (0.94); and β, the full width at half maximum of the peak angle of (002) crystal plane (FWHM) [17,18,21]. The measurements were performed at least three times. 3.6. Thermogravimetric analysis (TGA) The thermal stability of the samples was studied with thermogravimetric analyzer (TGA-50, Shimadzu, Japan) in air atmosphere at a heating rate of 10 °C/min from ambient temperature to 600 °C.

ultrasonic process was used to produce nanoparticles. When the cellulose is subjected to ultrasonic treatment, the structure of cellulose gets further loosened indicating disruption of the weaker β-D-(1 → 4) glycosidic linkages. The disruption results from the hydrodynamic force and shear stress generated from the collapse of bubbles through the ultrasonication process [19,37]. The bubble collapse is strong enough to affect the chemical structures, and break some of the weaker bonds [20]. As a result, the ultrasonic could effectively disintegrate the micron-sized particles into nanoparticles applying no specific chemicals. To ensure the noticeable impact of enzymatic process on nanoparticle production, untreated cotton fibers were sonicated with the same time and power, but excluded enzymatic treatment (control experiment). Consequently, no change was noticed in the appearance of fibers and a colloidal suspension was not formed; however, enzymatic hydrolysis conjoined with sonication resulted in a stable colloidal suspension. As a result, it was found that the applied low power and short sonication could disrupt the weakened linkages of the hydrolyzed cotton and change the micron-sized particles into nanoparticles. Nevertheless, there have been several studies considering a lengthened sonication and nanofiber production from natural materials such as cellulose and silk [37,38], but our objective, here, was to use short and low power ultrasonic. It is noteworthy that the prepared nanoparticle yield (calculated as the percentage of the ratio of the dry mass of nanoparticles to the initial dry mass of cotton) was low (less than 20%). The low yield could be attributed to the fact that the enzyme converts a significant amount of cellulose into glucose, cellobiose, cellotriose, and cellotetraose. It seems that the low yield of nanoparticles could be, in part, improved through short periodic hydrolysis and ultrasonic, surfactant addition, and a shaker during the hydrolysis.

4. Results and discussion 4.1. Fluidity analysis The respective method in the present study was to produce spherical nanoparticles from the waste cotton fibers. The enzymatic hydrolysis of cellulose involves dissociation of glucosidic bonds which is a very complex process. Because the cellulose is insoluble, the cellulase enzyme could not diffuse into the cellulose structure. Accordingly, the enzymatic hydrolysis is a slow process proceeding from surface to core of the cellulose. On the other hand, the cotton cellulose is highly recalcitrant owing to the dominance of cellulose Iβ structure. Therefore, a long-term hydrolysis process (175 h) was applied to produce small particles ranging several hundred nanometers. Subsequently, a short and low power

Fluidity of a given solution is inversely proportionate to viscosity considered as an indicator of the degree of polymerization. This parameter is often measured to assess the length of cellulose chains [16]. Fluidity measurement is a very sensitive method for measuring the amount of cellulose degradation [39]. As shown in Table 1, enzymatic hydrolysis and sonication cause a significant increase in the fluidity. This indicates that cellulose chains are shortened due to the applied treatments. This would, therefore, pertain to the extreme reduction of the degree of polymerization in the treated samples.

Fig. 1. Optical microscopy images of cotton before hydrolysis (left) after hydrolysis (right).


235

4.2. Microscopic analysis Optical microscopic images of the cotton fibers, before and after the enzymatic hydrolysis, are shown in Fig. 1. As it is evident, the width and length of fibers were reduced and the fine fiber amount increased through the enzymatic treatment. Fig. 2 demonstrated the FESEM images of samples in different magnification levels. The FESEM images of cotton fibers clearly revealed a relatively smooth and uniform surface before enzymatic hydrolysis process (Fig. 2a and b). Following hydrolysis process, the fibers were swollen and separated into cellulose products with much shorter chains. Of course, many distinct disruptions were observed on the fiber surface. In addition, short fragments were released from the fibers (Fig. 2c and d). Fig. 2e and f show that the morphology structure of cellulose fibers changed following ultrasound treatment. The nanoparticles would apparently form a membrane in the freeze drying process due to high concentration of the nanoparticles suspension. In order to achieve a more obvious image, a drop of very dilute suspension of nanoparticles was placed on a glass slide and was observed after freeze drying (Fig. 2 g). It is clear that nanoparticles have spherical shape with sizes less than 100 nm in diameter. Some researchers have previously studied the production of spherical cellulose nanoparticles [2,14,30]. For example, Li et al. [14] prepared nanospherical cellulose structures with diameters of several hundred nanometers from short-staple cotton through pre-swelling the fibers with dimethyl sulfoxide (DMSO) and sodium hydroxide (NaOH) as well as acid hydrolysis in a sonicator. Additionally, Zhang et al. [2]

Fig. 3. Particle size distribution of hydrolyzed cotton as analyzed by Mastersizer particle size analyzer.

used a pre-treatment with NaOH and DMSO and 8-hour acid hydrolysis in a sonicator to produce spherical cellulose nanoparticles with sizes ranging 60–570 nm from cellulose fibers. Further, Lu and Hsieh [30] have recently reported that a great deal of the products from acid hydrolysis and freeze drying of the cotton cellulose were 10–100 nm spherical cellulose nanocrystals. Acid hydrolysis is the most commonly used method for the preparation of cellulose nanoparticles, whereas mineral acids aren't environment friendly and produce by-products. Also, mineral acids are to some extent associated with difficult process controlling, handling and transportation as well as maintenance problems. Therefore, the used

Fig. 2. Field emission scanning electron microscopy images of (a and b) untreated cotton, (c and d) hydrolyzed cotton, and (e, f and g) cotton nanoparticles.

236


This smaller size of the nanoparticles as compared to the hydrolyzed cotton ends up with a stable suspension. Furthermore, the results of dynamic light scattering studies confirm the FESEM results. 4.4. Fourier Transform Infrared Spectroscopy

Fig. 4. Particle size distribution of cotton nanoparticles as analyzed by Zetasizer particle size analyzer.

method in this study (enzymatic hydrolysis) is the preferred method as compared to the acid hydrolysis.

4.3. Particle size analysis The results of light scattering studies on the particle size and distribution of the hydrolyzed cotton and the respective nanoparticles are presented in Figs. 3 and 4, respectively. The figures indicated a notable shift in the particle size distribution towards lower values for the nanoparticles as compared to the hydrolyzed cotton. The particle size analysis of hydrolyzed cotton suspension showed a bimodal distribution with average 0.417 and 3.802 μm for each peak (Fig. 3). The average particle size of hydrolyzed cotton suspension was d (0.5) = 0.526 μm indicating that 50% of the particles were smaller than 0.526 μm. The particle size distribution of the nanoparticle suspension indicated that the product mixture consisted of two different particle sizes with approximately 70 nm and 200 nm (Fig. 4). But, most of the particles were 40–90 nm, while the number of particles ranging 100–300 nm was negligible.

Fig. 5 shows the Fourier Transform Infrared spectra of untreated cotton, hydrolyzed cotton and the respective nanoparticles. All samples showed analogous spectra with the crystalline Iβ characteristic peaks: OH stretching at 3270 cm−1 and OH out of plane bending at 710 cm−1 [15,30]. It was clear that the enzymatic hydrolysis and the ultrasound treatment retained the cellulose Iβ allomorph in the cotton. Some characteristic peaks in cotton are at 3800–3000, 2902, 1635, 1430, 1372, 1162, and 899 cm−1 [18,40]. Accordingly, the band 3800–3000 cm−1 corresponds to OH stretching and flexural vibration of intra- and intermolecular hydrogen bonds of cellulose [41,42]. Of course, any possible change in the number and strength of hydrogen bonds brings about change in intensity (height) and width of the related band [40,43]. Fig. 5 shows that the hydrogen bonds partially change after the enzymatic hydrolysis and ultrasonication. The vibrations located at 2902 and 2850 cm−1 are attributed to CH2 asymmetric vibrations [43]. The peak 1635 cm−1 associated with the OH bending of adsorbed moisture (H2O) was stronger for the nanoparticles than that of other samples because of the larger surface area of nanoparticles as compared to the untreated and hydrolyzed cotton. The band at 1430 cm−1 is associated with the HCH and OCH in-plane bending vibrations [18]. This band is designated as a crystalline absorption band [40]. Additionally, the vibration located at 1372 cm−1 is attributed to the CH deformation vibration [18,40]. Also, the band at 1162 cm−1 is assigned to an asymmetric bridge oxygen stretching vibration (i.e. of C1\O\C4 groups). Furthermore, the band at 899 cm−1 designated as an amorphous absorption band is related to the COC, CCO, and CCH deformation modes and stretching vibrations of the C-5 and C-6 atoms [18,40]. Overall, all the bands retained their position after the hydrolysis and ultrasonication. 4.5. X-ray diffraction analysis The X-ray diffraction patterns of untreated cotton, hydrolyzed cotton and nanoparticles are depicted in Fig. 6.

Fig. 5. Fourier Transform Infrared spectra of (a) untreated cotton, (b) hydrolyzed cotton, and (c) cotton nanoparticles.


237

Fig. 6. X-ray diffraction of (a) untreated cotton, (b) hydrolyzed cotton, and (c) cotton nanoparticles.

All the samples showed three cellulose Iβ characteristic peaks at 2θ = 14.7° (101), 16.3° (10 1 ), and 22.5° (002) [17,19,30,42]. The peak intensity of the (002) plane (2θ = 22.5°) in the diffraction patterns of treated samples (hydrolyzed cotton and the nanoparticles) increased slightly indicating higher perfection of the crystal lattice in the (002) plane than untreated cotton. The peak of the (101) plane (2θ = 14.7°) also became partially more intense for treated samples. The crystallinity index (CrI) values of cellulose revealed no significant changes after the hydrolysis and ultrasonication processes (Table 2). Of course, the difference in the CrI of the samples was marginal. Moreover, the crystallinity indices for the untreated and enzyme treated cotton were 80.46% and 82.80%, respectively. To date, certain research has confirmed the increase in crystallinity during the enzymatic hydrolysis [15,42]. For example, Wang et al. [15] reported an increase in the cotton crystallinity from 84.8% to 86.8, 90.6, and 90.7% after 6, 12, and 18-day enzymatic hydrolysis, respectively. Also, Cao and Tan [42] reported the crystallinity of softwood pulp increased from 75.83% to 79.59, 81.22, and 82.12% after 2, 7, and 15-hour hydrolysis, respectively. The partial increase in CrI (Table 2) resulted from the difference in reactivity between the crystalline and amorphous cellulose leading to a rapid removal of amorphous cellulose near the surface. Also, the loss of the surface amorphous cellulose did not considerably change the total crystallinity, because its amount was negligible as compared to the bulk amorphous cellulose [44]. After the sonication, the crystallinity index decreased to 78.98%. Of course, the marginal reduction in CrI of the nanoparticles pertained to

Table 2 The parameters of X-ray diffraction for different samples. Sample

2θ (°)

CrI (%)

FWHM (°)

Lc (nm)

d-spacing (nm)


22.53 22.38 22.35

80.46 82.80 78.98

1.90 1.91 1.91

4.452 4.431 4.428

0.394 0.396 0.397

more disruption during the ultrasonic treatment as reported by Yang et al. [45]. As can be seen in Table 2, no significant change in the crystallite size and d-spacing of hydrolyzed cotton and nanoparticles was observed as compared to untreated cotton. The slight decrease in the crystallite size of hydrolyzed cotton may be due to this fact that the enzyme cannot easily penetrate the crystals; therefore, crystal surface erosion occurs. Also, a negligible reduction in the nanoparticles crystallite size may be because of cavitations bubbles' collapsing energy. It is worth noting that the method in the present study retained cellulose I crystalline structure in spherical nanoparticles. But, the other methods have led the crystalline structure of nanoparticles to change into cellulose II [2,14]. To our knowledge, there is only one report about the spherical nanoparticles in which cellulose I crystalline structure of the cotton fibers was retained following the acid hydrolysis [30]. 4.6. Thermogravimetric analysis To evaluate the thermal stability of untreated cotton, hydrolyzed cotton and nanoparticles, thermogravimetric analysis was performed. Fig. 7 exhibits steps of weight loss indicative of degradation processes in the samples. As can be seen, hydrolysis and sonication of cotton fibers did not result in significant changes in the characteristic degradation temperatures. The degradation patterns in terms of shape appear to be quite the same for all samples. The dominant peak between 300 and 350 °C was a result of concurrent degradation processes such as depolymerization, dehydration, and decomposition of the glycosyl rings and subsequent formation of a charred residue [19,46]. The peak above 350 °C was also due to the oxidation and breakdown of the charred residue into gaseous products with low molecular weight [46,47]. The untreated cotton showed the typical decomposition with onset temperature above 300 °C. Also, the degradation behavior of the hydrolyzed cotton showed partial differences from that of the untreated cotton implying that the hydrolyzed cotton degradation started at a

238


Fig. 7. Thermogravimetric analysis of (a) untreated cotton, (b) hydrolyzed cotton, and (c) cotton nanoparticles.

partially higher temperature (305 °C). Since, according to Wang et al. [48], more ordered cellulose fibers require more energy for polymer degradation, this partial change is the outcome of the higher crystallinity of hydrolyzed cotton as compared to the untreated cotton. The decomposition of cotton nanoparticles started at a lower temperature about 250 °C because of the lower crystallinity of nanoparticles as compared to the other samples. The high surface area of nanoparticles had a significant effect on reducing their thermal stability owing to the increased exposure surface area to heat. Furthermore, the decomposition at lower temperatures would also show faster transfer of heat in the cellulose nanoparticles [30]. According to Table 3, the amount of charred residue was also larger in samples with higher crystallinity. The char yield from 11.09% for the untreated cotton reached 25.21 and 7.34% for hydrolyzed cotton and nanoparticles, respectively. These nanoparticles showed higher thermal stability as compared to spherical nanoparticles prepared with the acid hydrolysis. Lu and Hsieh [30] reported that the decomposition of spherical nanocrystals prepared with the sulfuric acid hydrolysis started at around 150 °C. Thus, lower thermal stability of nanoparticles prepared with the sulfuric acid hydrolysis suggests a different decomposition mechanism. It has been reported that the activation energies of the degradation of cellulose nanoparticles were significantly minimized through introducing sulfate groups with the sulfuric acid hydrolysis [49]. According to the results of experiments, the proposed schematic image for the formation of nanoparticles is presented in Fig. 8.

Table 3 The amount of charred residue (Char yield) for different samples. Sample

Char yield (%)


11.09 25.21 7.34

As Fig. 8a shows, the cellulase enzyme attaches to the cellulose chains. Cellobiohydrolase attacks the crystalline structure of cellulose and endogluconase hydrolyzes the amorphous regions. Based on the results of fluidity analysis, the enzymatic hydrolysis decreases the chain length of the cellulose through splitting the β-D-(1,4) glycosidic linkages (Fig. 8b). Additionally, change in the intensity and width of the FTIR band in 3800–3000 cm−1 of hydrolyzed cotton showed that the cellulase enzyme could weaken and split the hydrogen bonds in the cellulose structure as reported by others [50,51] (Fig. 8c). The degree of polymerization significantly decreases after 175 h enzymatic hydrolysis. Consequently, according to the results of particle size analysis (Fig. 8d), the enzymatic hydrolysis leads to formation of particles with low aspect ratio (d 0.5 = 0.526 μm). In conclusion, based on the nanoparticle size analysis results, sonication may convert hydrolyzed fibers into particles with an average of 70 nm (Fig. 8e).

5. Conclusions Spherical cellulose nanoparticles were produced through the enzymatic hydrolysis followed by ultrasonication of waste cotton fibers. The FESEM images and the particle size analysis showed that the average size of the spherical nanoparticles was less than 100 nm. Also, the XRD results along with the FTIR data confirmed that cotton fibers retained the cellulose Iβ crystalline structure following the enzymatic hydrolysis and ultrasound treatment. In addition, the crystallinity did not change so much while the degree of polymerization significantly decreased, during these treatments. Also, the hydrolysis and sonication of cotton fibers did not lead to significant changes in the degradation patterns of the cellulose. To sum up, this study showed a green process for reusing of waste cotton fibers and converting them into useful cellulose powder without significant change in their beneficial properties and structure applicable for other waste cellulosic fibers.


Fig. 8. Proposed schematic image of nanoparticles formation.

References [1] P. Chen, S.Y. Cho, H.-J. Jin, Modification and applications of bacterial celluloses in polymer science, Macromol. Res. 18 (2010) 309–320. [2] J. Zhang, T.J. Elder, Y. Pu, A.J. Ragauskas, Facile synthesis of spherical cellulose nanoparticles, Carbohydr. Polym. 69 (2007) 607–611. [3] M. Aliyu, M.J. Hepher, Effects of ultrasound energy on degradation of cellulose material, Ultrason. Sonochem. 7 (2000) 265–268. [4] Y. Habibi, L.A. Lucia, O.J. Rojas, Cellulose nanocrystals: chemistry, self-assembly, and applications, Chem. Rev. 110 (2010) 3479–3500. [5] G. Agoda-Tandjawa, S. Durand, S. Berot, C. Blassel, C. Gaillard, C. Garnier, J.L. Doublier, Rheological characterization of microfibrillated cellulose suspensions after freezing, Carbohydr. Polym. 80 (2010) 677–686.

239

[6] T. Zimmermann, N. Bordeanu, E. Strub, Properties of nanofibrillated cellulose from different raw materials and its reinforcement potential, Carbohydr. Polym. 79 (2010) 1086–1093. [7] D.S. Hon, Cellulose: a random walk along its historical path, Cellulose 1 (1994) 1–25. [8] M.A.S. Azizi Samir, F. Alloin, A. Dufresne, Review of recent research into cellulosic whiskers, their properties and their application in nanocomposite field, Biomacromolecules 6 (2005) 612–626. [9] D. Dienes, A. Egyhazi, K. Reczey, Treatment of recycled fiber with Trichoderma cellulases, Ind. Crop. Prod. 20 (2004) 11–21. [10] M.T. Halimi, M. Ben Hassen, F. Sakli, Cotton waste recycling: quantitative and qualitative assessment, Resour. Conserv. Recycl. 52 (2008) 785–791. [11] Krystyna Wrześniewska-Tosik, Ewa Wesołowska, Dariusz Wawro, H. Struszczyk, Improvement of the enzymatic utilisation of textile waste from cellulose/polyester blends, Fibers text. East. Eur. 11 (2003) 63–66. [12] A. Isci, G.N. Demirer, Biogas production potential from cotton wastes, Renew. Energy 32 (2007) 750–757. [13] A. Jeihanipour, M.J. Taherzadeh, Ethanol production from cotton-based waste textiles, Bioresour. Technol. 100 (2009) 1007–1010. [14] X.F. Li, E.Y. Ding, G.K. Li, A method of preparing spherical nano-crystal cellulose with mixed crystalline forms of cellulose I and II, Chin. J. Polym. Sci. 19 (2001) 291–296. [15] L. Wang, Y. Zhang, P. Gao, D. Shi, H. Liu, H. Gao, Changes in the structural properties and rate of hydrolysis of cotton fibers during extended enzymatic hydrolysis, Biotechnol. Bioeng. 93 (2006) 443–456. [16] M.A. Stewart, Biopolishing Cellulosic Nonwovens, North Carolina State University, Raleigh, 2005. 159. [17] P.B. Filson, B.E. Dawson-Andoh, D. Schwegler-Berry, Enzymatic-mediated production of cellulose nanocrystals from recycled pulp, Green Chem. 11 (2009) 1808–1814. [18] P. Satyamurthy, P. Jain, R.H. Balasubramanya, N. Vigneshwaran, Preparation and characterization of cellulose nanowhiskers from cotton fibres by controlled microbial hydrolysis, Carbohydr. Polym. 83 (2011) 122–129. [19] S.S. Wong, S. Kasapis, Y.M. Tan, Bacterial and plant cellulose modification using ultrasound irradiation, Carbohydr. Polym. 77 (2009) 280–287. [20] G. Markevičius, Pressure Variation Assisted Fiber Extraction and Development of High Performance Natural Fiber Composites and Nanocomposites, Department of Physics, Southern Illinois University, Carbondale, 2010. 166. [21] K. Das, D. Ray, N.R. Bandyopadhyay, T. Ghosh, A.K. Mohanty, M. Misra, A study of the mechanical, thermal and morphological properties of microcrystalline cellulose particles prepared from cotton slivers using different acid concentrations, Cellulose 16 (2009) 783–793. [22] J. Rojas, A. Lopez, S. Guisao, C. Ortiz, Evaluation of several microcrystalline celluloses obtained from agricultural by-products, J. Adv. Pharm. Technol. Res. 2 (2011) 144–150. [23] E.D. Teixeira, A.C. Correa, A. Manzoli, F.D. Leite, C.R. Oliveira, L.H.C. Mattoso, Cellulose nanofibers from white and naturally colored cotton fibers, Cellulose 17 (2010) 595–606. [24] S. Elazzouzi-Hafraoui, Y. Nishiyama, J.L. Putaux, L. Heux, F. Dubreuil, C. Rochas, The shape and size distribution of crystalline nanoparticles prepared by acid hydrolysis of native cellulose, Biomacromolecules 9 (2008) 57–65. [25] W. Bai, J. Holbery, K. Li, A technique for production of nanocrystalline cellulose with a narrow size distribution, Cellulose 16 (2009) 455–465. [26] D. Liu, X. Chen, Y. Yue, M. Chen, Q. Wu, Structure and rheology of nanocrystalline cellulose, Carbohydr. Polym. 84 (2011) 316–322. [27] K. Abe, S. Iwamoto, H. Yano, Obtaining cellulose nanofibers with a uniform width of 15 nm from wood, Biomacromolecules 8 (2007) 3276–3278. [28] H.M.C. Azeredo, Nanocomposites for food packaging applications, Food Res. Int. 42 (2009) 1240–1253. [29] J.K. Pandey, J.-W. Lee, W.-S. Chu, C.-S. Kim, S.-H. Ahn, C.S. Lee, Cellulose nano whiskers from grass of Korea, Macromol. Res. 16 (2008) 396–398. [30] P. Lu, Y.L. Hsieh, Preparation and properties of cellulose nanocrystals: rods, spheres, and network, Carbohydr. Polym. 82 (2010) 329–336. [31] S.J. Eichhorn, Cellulose nanowhiskers: promising materials for advanced applications, Soft Matter 7 (2011) 303–315. [32] S.J. Eichhorn, A. Dufresne, M. Aranguren, N.E. Marcovich, J.R. Capadona, S.J. Rowan, C. Weder, W. Thielemans, M. Roman, S. Renneckar, W. Gindl, S. Veigel, J. Keckes, H. Yano, K. Abe, M. Nogi, A.N. Nakagaito, A. Mangalam, J. Simonsen, A.S. Benight, A. Bismarck, L.A. Berglund, T. Peijs, Review: current international research into cellulose nanofibres and nanocomposites, J. Mater. Sci. 45 (2009) 1–33. [33] T. Fattahi Meyabadi, G. Mir Mohamad Sadeghi, F. Dadashian, H. Ebrahimi Zanjani Asl, From cellulosic waste to nanocomposites. Part 2: synthesis and characterization of polyurethane/cellulose nanocomposites, J. Mater. Sci. 48 (2013) 7283–7293. [34] T. Fattahi Meyabadi, F. Dadashian, Optimization of enzymatic hydrolysis of waste cotton fibers for nanoparticles production using response surface methodology, Fiber Polym. 13 (2012) 313–321. [35] E. Gumuskaya, M. Usta, H. Kirci, The effects of various pulping conditions on crystalline structure of cellulose in cotton linters, Polym. Degrad. Stab. 81 (2003) 559–564. [36] L. Segal, J.J. Creely, A.E. Martin Jr., C.M. Conrad, An empirical method for estimating the degree of crystallinity of native cellulose using the X-ray diffractometer, Text. Res. J. 29 (1959) 786–794. [37] H.-P. Zhao, X.-Q. Feng, H. Gao, Ultrasonic technique for extracting nanofibers from nature materials, Appl. Phys. Lett. 90 (2007) 073112. [38] W. Chen, H. Yu, Y. Liu, P. Chen, M. Zhang, Y. Hai, Individualization of cellulose nanofibers from wood using high-intensity ultrasonication combined with chemical pretreatments, Carbohydr. Polym. 83 (2011) 1804–1811. [39] V.C. Jelineki, Flow characteristics of dispersions of cotton and regenerated cellulose rayon fabrics in cuprammonium their significance in fluidity calculations, Ind. Eng. Chem. 16 (1944) 172–178.

240


[40] S.Y. Oh, D.I. Yoo, Y. Shin, G. Seo, FTIR analysis of cellulose treated with sodium hydroxide and carbon dioxide, Carbohydr. Res. 340 (2005) 417–428. [41] F. Dadashian, Changes in the fine structure of lyocell fibers during enzymatic degradation, Proceeding of the World Textile Conference, 2nd Autex ConferenceBruges, Belgium, 2002, pp. 464–473. [42] Y. Cao, H.M. Tan, Structural characterization of cellulose with enzymatic treatment, J. Mol. Struct. 705 (2004) 189–193. [43] N. Abidi, E. Hequet, L. Cabrales, Applications of fourier transform infrared spectroscopy to study cotton fibers, in: G. Nikolic (Ed.), Fourier Transforms — New Analytical Approaches and FTIR Strategies, 2011. [44] H. Zhao, J. Kwak, Z. Conradzhang, H. Brown, B. Arey, J. Holladay, Studying cellulose fiber structure by SEM, XRD, NMR and acid hydrolysis, Carbohydr. Polym. 68 (2007) 235–241. [45] F. Yang, L. Li, Q. Li, W. Tan, W. Liu, M. Xian, Enhancement of enzymatic in situ saccharification of cellulose in aqueous-ionic liquid media by ultrasonic intensification, Carbohydr. Polym. 81 (2010) 311–316.

[46] M. Grunert, Cellulose Nanocrystals: Preparation, Surface Modification, and Application in Nanocomposites, State University of New York, New York, 2002. [47] N. Abidi, E. Hequet, L. Cabrales, J. Gannaway, T. Wilkins, L.W. Wells, Evaluating cell wall structure and composition of developing cotton fibers using Fourier transform infrared spectroscopy and thermogravimetric analysis, J. Appl. Polym. Sci. 107 (2008) 476–486. [48] N. Wang, E. Ding, R. Cheng, Thermal degradation behaviors of spherical cellulose nanocrystals with sulfate groups, Polymer 48 (2007) 3486–3493. [49] M. Roman, W.T. Winter, Effect of sulfate groups from sulfuric acid hydrolysis on the thermal degradation behavior of bacterial cellulose, Biomacromolecules 5 (2004) 1671–1677. [50] V. Arantes, J.N. Saddler, Access to cellulose limits the efficiency of enzymatic hydrolysis: the role of amorphogenesis, Biotechnol. Biofuels 3 (2010) 1–11. [51] S. Janardhnan, M.M. Sain, Isolation of cellulose microfibrils — an enzymatic approach, Bioresources 1 (2006) 176–188.