Processing of Cellulose Nanofiber-reinforced Composites A. BHATNAGAR* AND M. SAIN Faculty of Forestry University of Toronto 33 Willcocks Street, Toronto Ontario, Canada, M5S 3B3 ABSTRACT: Cellulose nanofibers are obtained from various sources such as flax bast fibers, hemp fibers, kraft pulp, and rutabaga, by chemical treatments followed by innovative mechanical techniques. The nanofibers thus obtained have diameters between 5 and 60 nm. The ultrastructure of cellulose nanofibers is investigated by atomic force microscopy and transmission electron microscopy. The cellulose nanofibers are also characterized in terms of crystallinity. Reinforced composite films comprising 90% polyvinyl alcohol and 10% nanofibers are also prepared. The comparison of the mechanical properties of these composites with those of pure PVA confirmed the superiority of the former. KEY WORDS: natural fibers, root crops, cellulose microfibrils, defibrillation.

INTRODUCTION polymeric matrix and a synthetic filler (e.g., glass fiber, carbon, or aramid) as a reinforcement have been widely used in many applications (automotive, packaging, construction, etc.) because of their high performance and great versatility. However, current environmental problems caused by these products at end-oflife disposal, their partial combustibility, and the increasing demand for techniques for the recycling of these materials have resulted in the replacement of synthetic fillers by natural organic ones such as natural fibers, wood fibers, starch, etc. These kinds of fibers, compared to inorganic fillers, have many advantages, including low cost, lower density, no abrasion of the processing equipment, similar moduli, good thermal properties, and biodegradability. The natural fibers used in most of the applications like paper making, textile industry, etc. are bundles of individual fibers held together by thin layers of polysaccharides, pectins, and lignin. Like synthetic fibers, natural fibers in the form of woven or nonwoven mats present outstanding opportunities to develop a new class of advanced lightweight composites. These mats either in the form of nonwoven or even as woven mats are now being used in manufacturing materials including reinforced composites. Generally, these mats are made from long and thick fiber bundles instead

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*Author to whom correspondence should be addressed. E-mail: [email protected]

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0731-6844/05/12 1259–10 $10.00/0 DOI: 10.1177/0731684405049864 ß 2005 Sage Publications

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of lean microfibrils. Until now, focus has been on the long fibers from plants such as hemp, flax jute etc. [1–4]. These long natural fibers are bundles of individual strands held together by thin layers of polysaccharides, pectin, and lignin. Typical primary plant cell wall is composed of cellulose microfibrils (9–25%) and an interpenetrating matrix of hemicelluloses (25–50%) and pectins (10–35%). Primary cell wall composition is of cellulose fibers, bound together by molecules made of sugar units. Approximately 90% of the cell wall consists of carbohydrates (mostly pentose and hexose units). Cellulose forms the framework of the cell wall while hemicelluloses cross-link noncellulosic and cellulosic polymers. Pectins provide cross-links and structural support to the cell wall. The arrangement of the cellulose microfibrils in the primary wall is random. Secondary walls are derived from the primary walls by the thickening and inclusion of lignin into the cell wall matrix and they occur inside the primary wall. Secondary cell walls of plants contain cellulose (40–80%), hemicellulose (10–40%), and lignin (5–25%), where cellulose microfibrils are embedded in lignin. The cellulose molecules are always biosynthesized in the form of nanosized fibrils (referred to as nanofibers in the present study), which are in turn assembled into fibers, films, walls, etc. The cellulose nanofibers can be considered to be an important structural element of natural cellulose. It consists of an assembly of cellulose chains whose degree of perfection in their parallel organization is expressed in its crystallinity percentage. The molecular arrangements of these fibrillar bundles are so small that the average diameter of the bundle is about 10 nm [5–8]. These cellulose nanofibers are with diameters of 5–50 nm and lengths of thousands of nanometers [9]. Many studies were done to prepare composites using cellulose microfibrils as reinforcement from sugar beet, tunicin, etc. [10]. But cellulose microfibrils obtained from these sources had some disadvantages. Bacterial cellulose microfibrils are very expensive and can cause a contamination problem in alimentary applications. Cellulose microfibrils from primary cell walls, as described in almost all the literature cited, can be obtained only from the sources which are principally constituted of parenchyma cells; therefore, the raw material choice is very limited. Surface-modified cellulose microfibrils as described in previous works are actually cellulose derivatives, and not pure cellulose. The objective of this study is to explore new raw materials for extracting cellulose nanofibers, from the plant fibers which contain not only primary cell walls, but also secondary ones. The cellulose nanofibers were extracted from cell walls by chemomechanical treatments, followed by the preparation of cellulose nanocomposites. MATERIALS AND METHODS The raw materials used in the study were industrial grade hemp fiber of Ontario (Source: Hempline, London, Ontario), spring flax fibers (Source: Saskatchewan), Kraft pulp (Source: Bleached northern black spruce-Kimberly Clark), rutabaga (local sources), and polyvinyl alcohol (Source: Dupont; commercial name Elvanol, grade 71–30 in powder). Experimental PREPARATION OF NANOFIBERS Chemical Treatment: Removal of Pectins, Hemicellulose, and Lignin from Long Fibers Hemp and flax bast long fibers were cut into 2–3 mm length and were then soaked in sodium hydroxide solution of 17.5% w/w concentration for 2 h. An important aim of this

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pretreatment was to increase the surface area of the lignocellulosic material to make polysaccharides more susceptible to hydrolysis. The fibers were then washed with abundant distilled water. Rutabaga was peeled and made into pulp by using warring blender (10 : 1 water to pulp ratio). Pretreatment of the fibers was followed by acid hydrolysis to solubilize hemicellulose in dilute acid at a lower temperature. The fibers were then treated with 1 M hydrochloric acid solution at 60–80
C. Pulp thus obtained was neutralized with distilled water. The fibers were then treated with 2% w/w sodium hydroxide solution for 2 h at 60–80
C [11]. Dilute sodium hydroxide treatment helps to separate structural linkages between lignin and carbohydrate, and disrupts the lignin structure. This alkali treatment resulted in the solubilization of lignin and remaining pectins and hemicellulose. The fibers were then washed with abundant distilled water until it became neutral; then the sample was vacuum filtered. Kraft pulp was not chemically treated and was used as it is for mechanical treatments. Isolation of Cellulose Nanosized Fibrils After the chemical treatments, the nanofibers were still associated within the cell walls [12]. A mechanical shear force was applied to defibrillate them. Cryocrushing was done with liquid nitrogen for this purpose, where samples were frozen and ice crystals were formed. A high mechanical impact was applied to slash the cell wall fragments and liberate the nanosized fibrils from the cell wall. The samples were further broken down into leaner fragments by a mechanical treatment using high shear, high energy transfer, and high impact. The process provided high turbulence and shear that created the efficient mechanism of reduction in size (to submicron level). The detailed method for mechanical treatment to rupture fiber cell wall and to enhance production of nanofibers is described in recent patent applications [13,14]. COMPOSITE PREPARATION The objective of composite preparation was to study the effect of chemical purification and mechanical treatments (isolation of cellulose nanofibers) on the mechanical performance of these nanofibers as reinforcement in a polymer matrix. Solid films were prepared using the nanofibers suspension previously obtained. The control sample was prepared with pure PVA. A blend containing 10% nanofibers and 90% polyvinyl alcohol was used for making the nanofiber-reinforced composite. A calculated amount of PVA was slowly dissolved in water at a temperature of 80
C with constant stirring; subsequently 10% nanofibers were added to the solution. The mix was poured into Pyrex glass Petri dishes. Each solution was dried for 24 h at 45–50
C to form the desired films. The films were finally removed from the trays. Chemical Analysis Untreated fibers and acid/alkali treated fibers were chemically analyzed for hemicellulose, lignin, and cellulose contents by using TAPPI standard T222 om-83 and TAPPI standard 250 um-85 represented hemicellulose and lignin contents, respectively. Scanning Electron Microscopy A scanning electron microscope (SEM) from Hitachi (S-2500 model) was used to study the morphology of the fibers after various stages of chemical and mechanical treatments.

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All images were taken at 15 kV. Fibers were mounted and gold coated on the top surface of the specially prepared metal studs. Fourier Transform Infrared Spectroscopy [FTIR] Perkin Elmer Spectrum 1000 (using Spectrum for Windows) was used to obtain spectra for the fibers after every chemical treatment. Fibers were ground and mixed with KBr (sample/KBr ratio, 1/99) to prepare pellets. Transmission Electron Microscopy Transmission electron microscopy (TEM) observations were achieved with Philips CM201 operated at 80 kV. A drop of a dilute cellulose nanofiber suspension was deposited on carbon-coated grids and allowed to dry. Atomic Force Microscopy AFM images were obtained using a Digital Instruments Dimension 3100 AFM with a nanoscope IIIa controller. The system was operated in tapping mode with DI tapping mode tips of a resonant frequency of 280 kHz. X-ray Powder Diffraction and Crystallinity Measurement A D8 Advance Bruker AXS diffractometer, a Cu point focus source, Graphite monochromator, and a 2D-area detector GADDS system were used for this purpose. All composite samples were analyzed on transmission mode. The objective of X-ray diffraction procedures is the recording and evaluation of the scattering direction and intensity or radiation diffracted by atom planes a fixed distance apart, according to the well-known Bragg’s law, n ¼ 2d sin , where  is the wavelength of the radiation, d is the distance between parallel planes,  is the angle of incidence and reflection of X-rays with respect to the planes, and n is an integer. Crystallinity of flax, rutabaga, and kraft pulp nanofibers estimated and compared with the crystallinity pattern of microcrystalline cellulose. Testing of Composites The mechanical behavior of PVA-cellulose nanofiber composite was analyzed with a Sintech-1 machine model 3397-36 in tensile mode. A load cell of 50 lb was used as per ASTM D 638 standard. The specimens were cut in dumbbell shapes with a die ASTM D 638 type V. Tensile tests were performed at a crosshead speed of 5 mm/min. The values, reported in this work, result from the average of at least five measurements. RESULTS AND DISCUSSION Plant fibers, wood fibers, and root crop fibers consisted of different cell walls structured together. The isolation of the individual cells required chemical treatments to hydrolyze and solubilize pectins, lignin, and hemicellulose. Cellulose can be partially degraded due to these chemical treatments. To avoid this degradation, alkali extraction was very carefully

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controlled. The alkali extraction hydrolyzes only the surface materials. If the concentration of the soda is high, some undesirable reaction can occur, like ‘peeling’ phenomenon, which occurs on the polysaccharides’ reducing end and continues through recurrent b-elimination. Therefore, a very low concentration of alkali was used to avoid this situation. Effect of Chemical Treatment: Removal of Lignin and Hemicellulose Chemically treated flax and hemp fibers were analyzed for hemicellulose, lignin, and cellulose contents and it was observed that cellulose content in the chemically treated fibers was 95% as compared to the original 75%. Tables 1 and 2 show the decline in hemicellulose and lignin content and increase of cellulose. Higher content of cellulose leads to a better stiffness and strength of the fibers. After each chemical treatment, scanning electron microscopy was done to verify the removal of pectin, hemicellulose, and lignin, and to visualize the separation of fibers from fiber bundles. Figure 1 shows the change in morphology of flax fibers after chemical treatments due to the removal of hemicellulose, lignin, and pectin. Infrared measurements were performed on chemically treated fibers to follow the removal of pectins. Samples were analyzed using FTIR after each chemical treatment. The alkali treatment allowed the ionization of pectin carboxylic groups and formation of sodium carboxylates, which decreased the formation of hydrogen-type intermolecular bond. Heating at elevated temperature led to degradation of pectic substances. Infrared measurements in Figure 2 show the removal of pectins due to vanishing of characteristic bands at approximately 1740 cm1 (carboxylate groups) and at approximately 1590 and 1240 cm1 (acetyl and methyl ester groups, respectively). Cellulose Nanofibers Size Distribution Atomic force micrographs and transmission electron micrographs suggested that no or only few lateral associations occur between adjacent nanofibers. Nanofibers are much more clearly defined probably because of the removal of pectic polysaccharides. The nanofibers obtained after the efficient mechanism of reduction in size (to submicron level) of chemically treated flax, hemp, and rutabaga fibers seem to be more interwoven and Table 1. Chemical analysis of flax bast fibers after selective chemical treatments.

Untreated fibers Fibers after acid hydrolysis Fibers after acid and alkali treatment

a Cellulose

Hemicellulose

Total lignin

Others

73 (3) 84 (6) 95 (1)

13 (2) 10 (5) 1 (1)

5 (1) 3 (1) 3 (1)

9 3 1

Table 2. Chemical analysis of hemp fibers after selective chemical treatments.

Untreated fibers Fibers after acid hydrolysis Fibers after acid and alkali treatment

a Cellulose

Hemicellulose

Total lignin

Others

76 (4) 85 (5) 94 (1)

11 (1) 6 (4) 2 (1)

7 (2) 6 (1) 3 (1)

6 3 1

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Figure 1. Scanning electron micrographs showing the change in morphology of fiber after chemical treatments for flax bast and hemp fibers, respectively. (A) Pretreated swollen fibers; (B) acid hydrolyzed fibers; (C) acid hydrolysis followed by alkali treatment, resulted in opening of the fiber bundles. Infrared spectra of flax bast fibres after each chemical treatment 100 90 80

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Figure 2. Removal of pectins after acid and alkali treatments in long fibers, vanishing of carboxylate group at approximately 1740 cm1, acetyl ester group at approximately 1590 cm1 and methyl ester group at 1240 cm1, respectively.

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their diameters were estimated between 5 and 80 nm as shown in Figure 3. Figure 4 shows the diameter distribution of the nanofibers obtained through chemomechanical treatments. It was observed that most of the nanofibers had a diameter range of 10–60 nm.

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Figure 3. (A) Transmission electron micrograph of rutabaga nanofibers negatively stained; (B) atomic force micrograph of flax bast nanofibers; (C) atomic force micrograph of hemp nanofibers; and (D) atomic force micrograph of bleached kraft pulp nanofibers.

Size distribution of flax bast nanofibres 0.3

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0.25 0.2 0.15 0.1 0.05 0 120

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Figure 4. Size distribution graph of flax bast nanofibers showing the nanofibers diameter range between 5 and 60 nm.

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Effect of Chemomechanical Treatment on Crystallinity X-ray diffractograms of nanofibers of flax, hemp, rutabaga, and kraft pulp are shown in Figure 5. The main diffraction intensity was at about 2 ¼30
for each sample and this corresponds to a unit cell length of 4.47 A˚. The crystalline nature of the cellulose nanofibers is not only influenced by the chain conformation, but also by the packing of adjacent chains. Chemical treatments and processing of cellulose fibers gives different X-ray patterns (Figure 5). A dominant feature of most of the celluloses is the molecular orientation in cellulose fibrils and along the cell wall axis of fibers. Strong orientation along these axes is associated with high tensile strength and has an intense effect on the mechanical properties of the bulk material. The nanofibers demonstrated high crystallinity as evidenced by the sharper main peak. Crystallinity of flax and rutabaga nanofibers was estimated as 59 and 64%, respectively, while nanofibers from wood kraft pulp showed estimated crystallinity of 54%. These results show proximity with microcrystalline cellulose (MCC) crystallinity of 64%. Therefore, these nanofibers may find application in the biomedical field as a replacement of MCC (Figure 6).

Figure 5. X-ray patterns of cellulose flax bast fibers before treatment and after chemomechanical treatments showing the cellulose nanofibers exhibit sharp peaks and diffraction pattern is similar to that of native cellulose.

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Figure 6. X-ray diffractions patterns for cellulose nanofibers: (a) flax nanofibers; (b) rutabaga nanofibers; (c) kraft pulp nanofibers; and (d) microcrystalline cellulose.

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Figure 7. Mechanical performance study of nanocomposites prepared by using 10% cellulose nanofibers and 90% polyvinyl alcohol from kraft pulp, hemp, rutabaga, and flax fibers. (A) Tensile strength and (B) Young’s modulus.

Mechanical Performance of Nanofiber-reinforced Composites When 10% nanofibers were used as a reinforcement, tensile strength of the film increased to 118 MPa when compared to 69 MPa for a nonreinforced pure PVA film. This implies that in practical applications, the materials made from these nanocomposites can withstand much greater stresses without undergoing irreversible deformation. It is predicted that cellulose nanofibers appreciably reinforce the PVA matrix. Since PVA is a hydrophilic polymer, there is a strong interface bonding between hydroxyl groups of nanofibers with polymer, resulting in an increase in tensile strength of the composite film. Four to fivefold increases in Young’s modulus was observed in nanofiber-reinforced composite films compared to nonreinforced polymer (Figure 7). The mechanical defibrillation at submicron level led to isolation of nanofibers and that induced an increase in Young’s modulus. This is probably due to the fact that mechanical treatments form a network of nanofibers as displayed in AFM. CONCLUSIONS Various chemomechanical treatments were done to obtain cellulose nanofibers from flax bast fibers, hemp, wood kraft pulp fibers, and rutabaga. Removal of hemicellulose, pectin, and lignin was quantified by chemical analysis; FTIR and SEM were also done to visualize the removal of the same. High shear mechanical defibrillation was the most important step to isolate cellulose nanofibers. The characterization of these nanofibers was made using AFM and TEM. Estimated crystallinity of the nanofibers was in the range of

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55–64% for flax and rutabaga nanofibers. The resultant suspension was used as a cheap and environmentally friendly reinforcement to process composite materials using polyvinyl alcohol as a polymer matrix. A polymer/nanofibers matrix was prepared where the nanofibers reinforcement content was 10% in 90% PVA. When the film was examined, apparently, the nanofiber-reinforced film looked the same as the pure polymer film under optical microscope; but when the films were tested for mechanical performance, it was observed that even 10% nanofibers provide a remarkable reinforcing potential. Nanofibers have great potential in high-end applications. These nanocomposites can be used in medical devices like biocompatible drug delivery system. Due to their lightweight and high strength, they can be utilized as high strength components in aerospace and automotive sector. Since these nanocomposites can be made biodegradable with tremendous stiffness and strength, they find application in the medical field such as blood bags, cardiac devices, and valves as reinforcing biomaterials. REFERENCES 1. Saheb, D. N. et al. (1999). Review: Advances in Polymer Technology, Natural Fiber Polymer Composites: A, 18(4): 351–363. 2. Bledzki, A. K. et al. (1996). Properties and Modification Methods for Vegetable Fibers for Natural Fiber Composites, Appl. Poly. Sci., 9(8): 1329–1336. 3. Hornsby, P. R. et al. (1997). Preparation and Properties of Polypropylene Composites Reinforced with Wheat and Flax Straw Fibers, J. Material Science, 32: 1009–1015. 4. Simonsen, J. (1996). Utilizing Straw as Reinforcement in Thermoplastic Building Materials, Constr. and Build. Mat., 10(6): 435–440. 5. Bacic, A. et al. (1988). Structure and Function of Plant Cell Walls, The Biochemistry of Plants: A Comprehensive Treatise, 14: 297–371. 6. Reiter, W.-D. (1998). The Molecular Analysis of Cell Wall Components, Trends Plant Sci., 3: 27–32. 7. Carpita, N. C. et al. (1993). Structural Models of Primary Cell Walls in Flowering Plants, Plant J., 3: 1–30. 8. Stamboulis, A. et al. (2001). Effects of Environmental Conditions on Mechanical and Physical Properties of Flax Fibers, Comp. Part A: Appl. Sci. Manuf., 32, 1105–1115. 9. Hepworth, D. G. et al. (2000). The Mechanical Properties of a Composite Manufactured from Non Fibrous Vegetable Tissue and PVA, Comp. Part A: Appl. Sci. Manuf., 31: 283–285. 10. Cavaille, Jean-Yves et al. (1997). Surface-modified Cellulose Microfibrils, Method for Making Same, and Use Thereof as Reinforcement in Composite Materials, Canadian Pat. No. WO1997/012917. 11. Helbert et al. (1997). Characterization of Native Crystalline Cellulose in the Cell Walls of Oomycota, J. Biotech., 57(1–3): 29–37. 12. Dufresne, A. et al. (1997). Mechanical Behavior of Films Prepared from Sugar Beet Cellulose Microfibrils, J. Appl. Poly. Sci., 64(6): 1185–1194. 13. Sain, M. and Bhatnagar, A. (2003). Canadian Patent Pending 2003, Application No. 2,437,616. 14. Sain, M. and Bhatnagar, A.. US Patent Pending, Application No. 60/512,912.