Cellulose Nanofibers from Softwood, Hardwood, and Tunicate

Subscriber access provided by UB + Fachbibliothek Chemie | (FU-Bibliothekssystem)

Article

Cellulose nanofibers (CNF) from softwood, hardwood and tunicate: preparation-structure-film performance interrelation Yadong Zhao, Carl Moser, Mikael E. Lindstrom, Gunnar Henriksson, and Jiebing Li ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b01738 • Publication Date (Web): 28 Mar 2017 Downloaded from http://pubs.acs.org on March 30, 2017

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

ACS Applied Materials & Interfaces is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Applied Materials & Interfaces

Cellulose Nanofibers (CNF) from Softwood, Hardwood and Tunicate: Preparation-Structure-Film Performance Interrelation

Yadong Zhao,† Carl Moser,†,‡ Mikael E. Lindström,† Gunnar Henriksson† and Jiebing Li*†,§

†

Department of Fibre and Polymer Technology, School of Chemical Science and Engineering, Royal Institute of

Technology, KTH, Teknikringen 56-58, 10044 Stockholm, Sweden. ‡

§

Valmet AB, 85194 Sundsvall, Sweden. Research Institute of Sweden, RISE, Bioeconomy/ Biorefinery and energy, Drottning Kristinas väg 61 11486, Stockholm, Sweden.

* Corresponding author: [email protected]

ABSTRACT: This work reveals the structural variations of cellulose nanofibers (CNF) prepared from different cellulose sources, softwood (Picea abies), hardwood (E. grandis x E. urophylla) and tunicate (Ciona intestinalis), using different preparation processes, and their correlations to the formation and performance of the films prepared from the CNF. Here the CNF are prepared from wood chemical pulps and tunicate isolated cellulose by an identical homogenization treatment subsequent to either an enzymatic hydrolysis or a 2,2,6,6-tetramethylpiperidinyl-1-oxyl (TEMPO)-mediated oxidation. They show a large structural diversity in terms of chemical, morphological, and crystalline structure. Among others, the tunicate CNF consist of purer cellulose and have higher degree of polymerization than wood CNF. Introduction of surface

1 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

charges via the TEMPO-mediated oxidation is found to have significant impacts on the structure, morphology, optical, mechanical, thermal, and hydrophobic properties of the prepared films. For example, the film density is closely related to the charge density of the used CNF and the tensile stress of the films is correlated to the crystallinity index of the CNF. In turn the CNF structure is determined by the cellulose sources and the preparation processes. This study provides useful information and knowledge for understanding the importance of the raw material for the quality of CNF for various types of application.

KEY WORDS: cellulose nanofibers (CNF); softwood; hardwood; tunicate; film; comparison; correlation


Page 2 of 40

Page 3 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


1. INTRODUCTION Cellulose is the most abundant biopolymer on earth.1 For different purposes, it has been isolated most commonly from wood, followed by other plants, algae, and bacteria.2 In fact, tunicate, which is widely distributed worldwide, is to our knowledge the only animal resource for cellulose.3 It has become an abundant and competitive resource after large-scale cultivation of certain tunicate species (e.g. Ciona intestinalis) in 3D sea farms successfully, and from which high quality cellulose is isolated.4-5 The cellulose isolation from different sources is achieved following different principles. In wood, cellulose is present in a mixture together with hemicelluloses, lignin, pectin and other substances, so the isolation is conducted by chemical pulping processes, such as kraft pulping, in order to remove mainly lignin. In tunicates cellulose is present as a composite of cellulose-protein fibrils cemented by sulfated mucopolysaccharides or sulfated glycans and lipids.5 This composite structure exists as a cover layer over the entire epidermis of individual animal body, termed “tunic”.6-7 The cellulose can be isolated from the tunic after removal of mainly protein and mucopolysaccharides using e.g. an acid hydrolysiskraft cooking-bleaching procedure.5 The isolated celluloses have different structures depending on the sources and preparation processes. Both softwood and hardwood celluloses have a glucose chain length between 300 and 1700 units. However, no matter what processes and reaction parameters are followed in the isolation, there are certain amounts of lignin and hemicelluloses remained in the isolated celluloses. The native hemicellulose and lignin structures vary between softwood and hardwood species. In the former, mannose is the most common hemicellulosic monomer presenting as Oacetyl-galactoglucomannan.8 In the latter, xylose is most abundant occurring predominately as Oacetyl-4-O-methyglucuronoxylan.9 Moreover, the hardwood lignin is composed of both guaiacyl



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

and syringyl units, while softwood lignin is mainly of guaiacyl units.10 Meanwhile, the isolated cellulose from tunicate has a chain length ranging from 800 to 10,000 units with a purity of > 99%.5 Morphologically, the wood celluloses contain fibrils with lateral dimensions of 3-5 nm, agglomerating into fibrillar aggregates and subsequent bundles with a lateral dimensions of 10 to 35 nm.11 The tunicate cellulose, on the other hand, arranges in microfibrils with a rectangular cross-sectional shape of 10–20 nm in width.5 These differences in fibrillar morphology depends probably on differences in size and structure of the cellulose synthesizing enzyme complexes in different phylum.12 It shall be underlined that cellulose most likely is “invented” independently of tunicates and plants, i.e., convergent evolution. However, irrespective of the sources, all isolated celluloses have many common or inherent characteristics, such as large specific surface area, high crystallinity, good thermal stability and excellent mechanical properties.2 On the other side, it is possible that fibrils of different dimensions might have diverse properties that can be utilized in specific applications. Due to the resource abundance and inherent renewability, sustainability and biocompatibility as well as excellent chemical and physical properties, the isolated celluloses have today become highlighted as a potential candidate for advanced bio-based material development, especially for films or composites.13-15 However, their poor dissolution and suspension properties in water limit their wider applications. To overcome this drawback, individualization of the fibrils into size-reduced cellulosic materials having at least one dimension in the nanometer range, termed nanocellulose, has been intensively investigated.2, 16 With or without an introduction of surface charge during the preparation, the nanocellulose can be suspended in aqueous solution, improving its processability. Films and composites from


Page 4 of 40

Page 5 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


nanocellulose have been shown to have improved mechanical, thermal, barrier and other physiochemical properties compared to macroscopic cellulose fibers.17 Generally, more structural differences among different prepared nanocelluloses will be resulted from the processing of the celluloses.2, 16 Chemically, the nanocellulose preparation is generally a hydrolysis process where chemical and mechanical methods are most commonly used.18 In the former, nanocellulose is generated by liberation of crystalline regions from the cellulose fibrils by hydrolysis of amorphous regions commonly using hydrochloric acid or sulfuric acid, and the obtained nanocellulose is termed cellulose nanocrystals (CNC).19-20 In the latter, the preparation is a process of mechanical defibrillation, including refining, homogenization and grinding, and the obtained nanocellulose is termed cellulose nanofibers (CNF).21 However, high energy consumption is the main drawback of CNF preparation.22 In order to reduce energy consumption, several cellulose pretreatments have been extensively investigated prior to the CNF preparations, especially for the wood celluloses.16 One is enzymatic hydrolysis using e.g. a monocomponent endoglucanase to facilitate the defibrillation via specific hydrolysis of the glucosidic bonds along the cellulose chain, primarily in less crystalline regions.23 Another is 2,2,6,6-tetramethylpiperidine-1-oxyl (TEMPO) mediated oxidation, which introduce anionically charged carboxyl groups not only causing electrostatic repulsion between each other and thus easing the defibrillation but also influencing the applications of the obtained CNF.24-25 Wood CNC prepared by acid hydrolysis has dimensions in the range of 3–5 nm in width and 50–500 nm in length with a crystallinity of 54–88% with 68-94% being the Iβ crystal structure.2, 16 However, wood CNF produced via mechanical refining contains nanofibers with dimensions in the range of 4–20 nm in width and 500–2000 nm in length containing both amorphous and crystalline regions. Although CNF from tunicates prepared by mechanical



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

method is scarcely reported, it is known that acid hydrolysis of tunicate cellulose resulted in ribbon-like shaped CNC with a height of ~8 nm, a width of ~20 nm, and a length of 100–4000 nm (typical aspect ratios of 70–100), being highly crystalline (85–100%) and containing nearly pure Iβ crystal structure. Understanding the structures and properties of different CNF and their dependence on the cellulose sources in the preparation process and their further impacts on film formation and performance is critical for the preparation and application of CNF for more advanced bio-based material developments. For example, a novel type of bio-based materials termed as microcapsules has been fabricated by using CNF.26 Based on these microcapsules, it has demonstrated that the dimensions of CNF are critical to the fabrication of microcapsules with tunable transport properties due to pore dimensions and film layer properties.27-29 In addition, another study generally suggested that the surface area of nanoparticles plays one among key roles for their applications.30 Although it was proposed that the characteristics of CNF have great effects on its performance in different applications, so far, however, no direct and systematic comparisons have been conducted for both wood and tunicate CNF in terms of both preparation and film formation. Numerous studies exist in the literature mainly about either wood or tunicate CNF. On the preparation subject, one comparative study has recently been conducted by Sacui et al. in 2014, in which the measurement and direct comparison of some properties of CNF isolated from bacteria, tunicate, and wood using typical hydrolysis conditions (acid, enzymatic, mechanical, and TEMPO-mediated oxidation) were performed.31 Unfortunately, they had not investigated the effects of variations in chemical compositions on CNF preparation and properties, which was however suggested to be critical for CNF performance.32 On the film formation subject, focus has been paid almost exclusively on wood CNF.32-33 In our previous study, we


Page 6 of 40

Page 7 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


evaluated and compared the film formation of tunicate CNF prepared by different methods including mechanical approach, enzymatic treatment and TEMPO-mediated oxidation.34 In this paper, in order to increase the understanding of the role of the raw material for CNF application, CNF were prepared from three different cellulose sources, softwood, hardwood and tunicate, using an identical homogenization conditions after a pretreatment of either enzymatic hydrolysis or TEMPO-mediated oxidation. The prepared CNF were subjected to comprehensive characterizations regarding chemical structure including chemical composition, degree of polymerization, and charge density; morphology of derived nanofibers by measuring fiber size and aspect ratio; and crystal properties by determining the crystalline allomorph, crystallinity index, crystal size, and crystalline Iβ ratio. Cast-films prepared from the various CNF were subjected to characterizations regarding density, morphology, and optical, mechanical, thermal and hydrophobicity properties.

2. EXPERIMENTAL SECTION 2.1 Starting celluloses

Mono-cultured animal bodies of one tunicate species, Ciona intestinalis, were collected at a farm located in Rong, Norway, on November, 2015 and cellulose was then prepared in our lab by using an alkali process similar to kraft pulping, as described earlier.5 Never-dried tunicate cellulose with a consistency of 2% was treated by a disintegrator (Frank-PTI GmbH, Germany) for 10 minutes (at 30,000 revolutions) before further pretreatment. Unbleached hardwood (E. grandis x E. urophylla) and softwood (Picea abies) kraft pulps with 1.6% and 7% acid insoluble lignin were chlorite delignified at a consistency of 3%, using 0.6 g sodium chlorite and acetic



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

acid per gram fibers for 2 hours at 70 °C, after which the pulp was thoroughly washed. A monocomponent endoglucanase enzyme was purchased from FiberCare®, Novozymes, DE and was used without further treatment. The cellulolytic activity of this enzyme supplied by the manufacturer is 4500 ECU g-1 (Endo Cellulase Units per unit mass of material).

2.2 CNF preparation

2.2.1 Pretreatment 2.2.1.1 Enzymatic hydrolysis

Cellulose suspension of 10g/L was prepared in 5 mM phosphate buffer with a pH of ~7. After the addition of a monocomponent endoglucanase enzyme (FiberCare®, Novozymes, DE) at an amount of 24.8 ECU per gram cellulose (in dry mass), the mixture was incubated at 55°C for 1 h.22-23 The enzyme in the reaction system was then denatured by heating to 80 °C for 30 min. Next, the treated cellulose was recovered by vacuum filtration with Whatman® qualitative filter paper, Grade 4, Pore size 20-25 µm, and then washed several times with deionised water.23 2.2.1.2 TEMPO-mediated oxidation

Cellulose suspension (10 g/L, 100 mL) was prepared containing TEMPO (0.016 g, 0.1 mmol) and sodium bromide (0.1 g, 1 mmol). The cellulose sample was stirred continuously at 400-500 rpm. Then, 3.1 mL aqueous NaClO solution (12%, 1.61 M) was added drop-wise.35 After that, the pH of the reaction system was controlled at 10.0±0.1 via the addition of 0.5 M NaOH. When the charge density reached the required number (~ 400 µmol/g), the reaction was fully quenched by the addition of 100 ml of ethanol.36 The oxidised cellulose was recovered by vacuum filtration with Whatman® qualitative filter paper, Grade 4, Pore size 20-25 µm, and then washed several times with ultrapure water to neutral pH. 8 ACS Paragon Plus Environment

Page 8 of 40

Page 9 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


2.2.2 Homogenizer treatment All pretreated celluloses (10 g/L) after either the enzymatic hydrolysis or TEMPOmediated oxidation were mechanically disintegrated in a homogenizer (M-110EH, Microfluidics Corp, USA) using two large chambers in series (400 and 200 µm, respectively) at 925 bar for the first pass and smaller chambers (200 and 100 µm, respectively) at 1600 bar for four passes.22 The yield (%) and concentration (g/L) of CNF were determined gravimetrically by drying a 10 ml CNF suspension.37 Based on the obtained concentrations for different CNF, they were diluted to 5g/L for further use. 2.3 Fabrication of CNF films

The 5g/L CNF suspension prepared as mentioned above was directly casted on Petri dishes and dried at 50 °C overnight to make cast films.32 In order to ensure a uniform thickness of dry films, the Petri dishes were placed on a levelled surface prior to drying.33 2.4 Characterizations

2.4.1 Chemical analysis 2.4.1.1 Chemical composition

Chemical composition of celluloses was determined in terms of carbohydrate and Klason lignin contents following the Tappi test method T222om-06. 200 mg celluloses were fluffed and added to 3 mL 72% H2SO4, before being put under vacuum for 80 min, after which the mixture was diluted with 84 mL mili-Q water and put into an autoclave for 120 min at 125 °C. The solutions were filtered when still warm and the filtered liquids were kept for the polysaccharide analysis. The filtered residue was repeatedly washed with hot distilled water and then dried at 105 °C during 24 h; the dry residue was finally weighed as the Klason lignin. The lignin content



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 10 of 40

was obtained by Lignin (%) = A*100/W, where A (g) and W (g) were the weight of Klason lignin and the oven-dry weight of pulps, respectively. The released carbohydrates were examined by high performance anion exchange chromatography with a pulsed amperometric detection (HPAEC-PAD) on a ICS3000 system (Dionex, Sunnyvale, CA, USA) using a Carbopac PA1 column (Dionex, Sunnyvale, CA, USA) at 30 °C at a flow rate of 1 mL/min. Quantification was performed by external calibration using neutral sugar standards (glucose, mannose, xylose, arabinose, and galactose). The anhydrous content of each monosaccharide was calculated based on different factors (0.88 for xylose and arabinose, 0.90 for glucose, mannose and galactose) according to the SCAN-CM 71:09. 2.4.1.2 Degree of polymerization

Dried CNF samples (30 mg) were dissolved in 8% LiCl/DMAc at a concentration of 8 g/L, by stirring at 4 °C for 5 days, after which the solutions were diluted with DMAc to reach a concentration of 5 g/L. The Size Exclusive Chromatography (SEC) system was equipped with a Rheodyne injector, a DGU-20A3 degasser, a LC-20AD liquid chromatography system and a RID-10A refractive index detector. The separations were achieved using four 20 µm Mixed-A columns with a guard column. The injection volume was 100 µL, and the separations were performed at 80 °C with the flow rate of 0.5 mL/min of 0.5% LiCl/DMAc. Calibration was conducted with pullulan standards of nominal masses ranging from 320 to 800 kDa (Fluka/Riedel-de Haën, Seelze, Germany). Data acquisition and analysis was performed using LC Solution software (Shimadzu, Kyoto, Japan) equipped to the SEC system. 2.4.1.3 Charge density determination

The charge density of CNF was analyzed by conductiometric titration according to a modified SCAN-CM 65:02 standard based on the operations used by other researchers.38-41 10 ACS Paragon Plus Environment

Page 11 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Briefly, each CNF (100 mg) was suspended into 50 ml of 0.01 M hydrochloric acid solution containing 0.01 mM NaCl. After stirring for 10 min, the suspension was titrated with 0.01 M NaOH. The amount of NaOH consumed in the plateau region was used to calculate the total charge density according to the method reported in the literature.40, 42 2.4.1.4 Fourier transform infrared spectroscopy (FTIR) analysis

Fourier transform infrared (FTIR) spectra were obtained using a Perkin-Elmer Spectrum 2000 Fourier transform infrared spectroscopy spectrometer (Waltham, MA, USA) equipped with an ATR system, Spectac MKII Golden Gate (Creecstone Ridge, GA, USA). The samples were analyzed at wavenumbers ranging from 600-4000 cm-1. All spectra were obtained from dry samples subjected to 16 scans at a resolution of 4 cm-1 and an interval of 1 cm-1 at room temperature. Before data collection, background scanning was performed for background correction. 2.4.2 Morphological analysis Before scanning electron microscopy (SEM) analysis, all samples were coated with gold using a Cressington 208HR high-resolution sputter coater. A Cressington thickness monitor controller was used to control the thickness (3-5 nm). Then, sample morphology was analyzed using a Hitachi S-4800 Field Emission Scanning Electron Microscope. A tapping-mode AFM (Multimode IIIa, Veeco, Santa Barbara, CA, USA) was also used to determine the morphology of CNF. AFM images were recorded under ambient and air conditions (23 °C and 50% relative humidity) using RTESP silica cantilevers (Veeco) having a tip with a radius of 8 nm and a spring constant of 40 N/m oscillated at their fundamental resonance frequencies between 200 and 400 kHz. The width of the CNF was measured using the NanoScope Analysis software, in which the height was measured for 124 - 256 nanofibers.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 12 of 40

Length measurements were conducted based on AFM images following the method reported by Usov et al.43 with mean lengths reported as an average of 92 - 151 nanofibers. The RMS (rootmean-square) roughness was measured based on the 5×1 µm2 height images using the Nanoscope Analysis software and the average value of at least five measurements was reported. 2.4.3 Crystal structure A PANalytical X’Pert PRO Materials Research Diffractometer equipped with an X´Celerator detector was used to determine the diffraction patterns and crystallinity index (CI) of the samples. The analysis was performed using monochromatic CuKα radiation at 30 mA and 40 kV. CI, defined to evaluate the crystallinity of different samples, was calculated using equation (1): =

− × 100 (1)

Where I200 is the intensity of the 200 lattice plane at around 2θ=22.8°, and Iam is the intensity from the amorphous phase at approximately 2θ=18°.44 The Scherrer equation (2) was used to calculate the crystal size, t (nm), which was determined perpendicular to the (200) planes for both cellulose I and cellulose II structures, =

(2)

Where k is the correction factor and usually taken as 0.9; λ is the radiation wavelength; θ is the diffraction angle; and β is the corrected angular width at half maximum intensity in radians.45 By following the method reported by Rondeau-Mouro et al.,46 the relative proportion of cellulose Iβ to Iα allomorph was calculated by integrating the FTIR absorption bands (section


Page 13 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


2.4.1.4) near 710 and 750 cm-1, which were considered to be characteristic for Iα and Iβ allomorphs, respectively. The percentage of Iβ was obtained by equation (3): (%) =

× 100 (3) +

Where A710 and A750 are integrated intensities of the bands around 710 and 750 cm-1. 2.4.4 Transmittance Light transmission (T%) of films was measured using a Shimadzu UV-240 (Japan) spectrophotometer. To compare different samples, readings were taken at a wavelength of 750 nm. Film specimens were cut into rectangles and placed in a spectrophotometer test cell directly, and air was used as the reference. Transmittance (T%= I/I0, where I and I0 were the intensities of emergent and incident radiation, respectively) was used to define the transparency of a measured film. Further, for a visual comparison, the films were photographed. 2.4.5 Mechanical strength measurement The tensile stress, tensile strain and Young’s modulus of the films were determined using an Instron 4411 tensile tester with a 500-N load cell (Instron Ltd., Norwood, MA, USA). The initial grip distance was 25 mm, and the rate of grip separation was 5 mm/min. Two films of each type and 3 specimens from each film were tested. The specimens were 5 mm wide and approximately 60 mm long. The thickness of the specimens was measured at three points using a micrometer (NSK, Japan). 2.4.6 Thermal stability measurement Thermo gravimetrical analysis (TGA) was employed using a Mettler Toledo TGA/SDTA 851e equipped with STARe software for data analysis. The samples were subjected to a heating scan between 30 and 800 °C, with a rate of 10 °C/min under an inert atmosphere of nitrogen at a gas flow of 50 mL/min. 13 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 14 of 40

The differential scanning calorimetry (DSC) analysis was performed through a METTLER TOLEDO DSC 822e instrument previously calibrated with indium standard. Samples of about 5–10 mg were sealed in standard aluminium pans previously pierced and tested under nitrogen atmosphere at a heating rate of 3 °C/min. The samples were first heated to 120 °C to eliminate interferences due to moisture and thermal history. Then the samples were cooled to 20 °C and reheated to 300 °C to determine their glass transition. Glass transition temperature (Tg) was recorded at the midpoint temperature of the heat capacity transition of the second heating run. 2.4.7 Hydrophobicity The contact angle (CA) was determined by the pendant drop method using a water drop and an optical contact angle meter SL 100B from Solon Information Technology Co., Ltd. (Shanghai, China) at a relative humidity (RH) of 50% and 23 °C. To compare different samples, each contact angle was taken at 45 s, and the average value of at least three measurements is presented. 2.4.8 Brunauer-Emmett-Teller (BET) analysis A BET-analysis was conducted on a Micrometritics ASAP 2020 Surface Area and Porosity Analyzer (Micromeritics, USA). First, in order to remove all foreign adsorbed molecules, ~100 mg film sample was heated to 115 °C and degassed under a vacuum for >300 minutes. After that, an inert nitrogen gas was introduced with a controlled amount and adsorbed. The adsorption isotherms were generated by exposing the samples to varying pressures at the temperature of liquid nitrogen (-196 °C). Then the Micromeritics ASAP 2020 V3.00 software was used to calculate the surface area, pore volume and pore size by using the BET equation automatically.

3. RESULTS AND DISCUSSION 14 ACS Paragon Plus Environment

Page 15 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


The used starting celluloses have been prepared from wood, both softwood (Picea abies) and hardwood (E. grandis x E. urophylla), and tunicate (Ciona intestinalis) using different processes to remove most of the other non-cellulose components. From wood cell walls, the fibers were liberated from lignin using kraft pulping and bleaching. The tunicate cellulose was isolated from its composite with protein, lipids and other non-cellulose polysaccharides in the tunic using an acid hydrolysis-kraft cooking-bleaching procedure.5 These three celluloses have inherently a great diversity in terms of chemical composition as can be seen in Table 1. Chlorite bleached softwood and hardwood celluloses had a glucose content of 77.6% and 79.3% respectively. They contain also residual lignin and hemicelluloses, as indicated by 0.2% Klason lignin and 6.6% mannose (predominately present as glucomannan) in the former and 1.5% Klason lignin and 14.8% xylose (predominately present as xylan) in the latter. On the contrary, tunicate cellulose is highly pure as indicated by a glucose content of 99.1% with only traces of mannose (Table 1). Table 1. Chemical composition of starting celluloses (%) Lignin

Glucose

Other sugars Xylose

Mannose

Galactose

Arabinose

Tunicate cellulose

n.d.*

99.1

n.d.*

0.3

n.d.*

n.d.*

Softwood cellulose

0.2

77.6

8.0

6.6

0.3

0.9

Hardwood cellulose

1.5

79.3

14.8

0.3

0.1

n.d.*

* not detected

It should be noted that in this study all these celluloses were processed as never-dried samples. It is known that drying not only is a costly process but also causes hornification resulting in the loss of elasticity and flexibility of the fibers, as well as closing internal pores thus lowering the chemical accessibility of the cellulose.47 For example, hornification has been shown to decrease the rate of a subsequent enzymatic hydrolysis.48 Therefore, never-dried samples 15 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 16 of 40

should be the best possible starting materials since enzymatic or chemical pretreatment was here conducted prior to mechanical defibrillation for the CNF preparation. The yields were relatively high for all CNF, 94-99%, irrespective to either pretreatment method or cellulose source. For ease of discussion, the obtained CNF were labeled with the abbreviations shown in Table 2 indicating their sources and pretreatments applied.

Table 2. CNF from softwood, hardwood and tunicate and their charge density and crystalline structures Chemical structure Cellulose

Pretreatment

Abbreviation

DP

source

Crystalline structure

Charge density

Crystallinity

Crystal

Iβ ratio

(µmol/g)

index (%)

size (nm)

(%)

Softwood

Enzymatic

eSW

1900

99

75.47

3.26

80.14

Softwood

TEMPO

tSW

1300

443

76.71

2.42

81.74

Hardwood

Enzymatic

eHW

1300

76

77.13

3.43

70.87

Tunicate

Enzymatic

eTC

4200

43

89.94

7.70

93.58

Tunicate

TEMPO

tTC

3900

428

86.17

7.06

94.29

3.1 Characteristics of CNF

3.1.1 Chemical structure As verified by FTIR measurements of CNF (Figure 1), the tunicate CNF was also nearly pure cellulose. For both eTC and tTC, the peaks at 3333 and 2900 cm-1 originated from the –OH stretching and C-H symmetrical stretching, respectively (Figure 1a). The peak at 1161 cm-1 arose from the C–O anti-symmetric bridge stretching (Figure 1b). The shoulder band at 1112 cm-1 was 16 ACS Paragon Plus Environment

Page 17 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


attributed to the C–OH skeletal vibration. The C–O–C pyranose ring skeletal vibration gave prominent bands at 1057 and 1030 cm-1. The small peak at 895 cm-1 corresponded to the glycosidic –CH deformation with a ring vibration and –OH bending in β-glycosidic linkages between glucoses in the cellulose.49 A band at 664 cm-1 originated from the out of plane deformation of C-H functional group.50 These peaks are characteristics of pure cellulose.5, 51 Wood CNF showed all these characteristic peaks, although with broader peaks than their tunicate counterpart at 3327-3331 cm-1 for the –OH stretching, 2893-2897 cm-1 for C-H symmetrical stretching, 1159-1161 cm-1 for the C–O anti-symmetric bridge stretching, 1023-1028 cm-1 for the C–O–C pyranose ring skeletal vibration and 661 cm-1 for the out of plane deformation of C-H functional group. This difference might arise from the presences of remaining lignin and hemicelluloses. The celluloses are still cross-linked by the residual lignin and hemicelluloses, showing wavenumber shifts from the pure cellulose from the tunicate. In addition, the peaks from the carbohydrates in the hemicelluloses would interfere the characteristic peaks from cellulose. The residual lignin would be observed by a characteristic peak at 1505 cm-1 arising from the aromatic skeletal vibrations. However, this peak was barely seen in either fully bleached wood CNF due to the low content of lignin in those samples (0.2-1.6%) (Table 1), although it was clearly visible in unbleached wood CNF (data not shown).



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 18 of 40

Figure 1. Two regions of FTIR spectra of softwood, hardwood and tunicate CNF. (a) 4000-2500 cm-1 and (b) 2000-600 cm-1 As seen from the SEC analysis (Table 2, DP), all CNF exhibit high molecular mass structures with a long chain of repeating glucose units. The degree of polymerization (DP) for wood CNF is between 1300 and 1900 with softwood CNF being longer than the hardwood CNF after the same enzymatic hydrolysis followed by homogenization. Comparatively, tunicate CNF has a significantly higher DP, 4200, than wood CNF. The TEMPO-mediated oxidation pretreatment tends to hydrolyze cellulose structures, as indicated by the lower DP of 3900 for tTC compared to 4200 for eTC and 1300 for tSW compared to 1900 for eSW, respectively. As determined by conductometric titration, after the enzymatic hydrolysis, eTC has the lowest charge density (43 µmol/g), implying that it is nearly perfectly pure cellulose (Table 2). A slightly higher charge density (76-99 µmol/g) is observed for wood CNF, eSW and eHW, due to the presence of hemicelluloses and lignin. In contrast, the TEMPO-mediated oxidation significantly increased the charge density irrespective of the celluloses source, reaching 428 µmol/g for tTC and 443 µmol/g for tSW respectively due to the introduction of carboxylic groups


Page 19 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(Table 2). The introduction of charged carboxyl groups in both tTC and tSW is further confirmed by their FTIR spectra as shown in Figure 1b. The peaks at 1740 and 1716 cm-1 arose from the C=O vibration stretching and the pronounced peak at 1609 cm-1 generated from the asymmetrical carboxylate COO- vibration overlapping with the O-H bending vibration of the absorbed water.52 3.1.2 Crystalline structure As illustrated in Figure 2, the XRD curves of eTC and tTC displayed a similar cellulose I structure, with strong crystalline peaks at 14.7°, 16.8° and 22.8°, corresponding to the (1ī0), (110), and (200) crystal planes, respectively.34 As discussed above, eTC has the least modified cellulose crystalline structure after the preparation process, and the observed similarity of tTC to eTC verified that the carboxyl groups introduced by TEMPO-mediated oxidation are selectively present on cellulose fibril surfaces without any interfering of the internal cellulose crystallites.42 On the other hand, both softwood and hardwood CNF showed similar diffraction patterns to tunicate CNF, although the peaks for their crystal planes were located at a shifted 2θ angles of around 15.0-15.6° (1ī0), 16.3-16.5° (110), and 22.3-22.6° (200), respectively, relating also to the presence of residual hemicelluloses and lignin. Based on the crystallinity indexes (CI) values calculated from the XRD analysis, the wood CNF have a CI between 75-77%. No significant CI difference was found between softwood and hardwood CNF. On the contrary, a higher CI is observed for tunicate CNF, 86-90% (Table 2), which might result from their natively highly ordered crystalline structure. The higher cellulose purity of the tunicates compared to wood might be another reason, since the hemicellulose and lignin present in wood are generally considered to be amorphous.53 After calculations, the crystal size of eTC is 7.70 nm, the highest among all CNF. Also after the identical enzymatic pretreatment, the crystal size for wood CNF (eSW and eHW) is smaller (3.26-3.43 nm) (Table 2). Irrespective of the celluloses used, the TEMPO-mediated 19 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 20 of 40

oxidation results in smaller crystal sizes in the obtained CNF than those after the enzymatic pretreatment, namely 7.06 nm for tTC vs. 7.70 nm for eTC and 2.42 nm for tSW vs. 3.26 nm for eSW, respectively, which is attributed to the higher degradation of the cellulose’s amorphous regions during the oxidation.54

Figure 2. XRD spectra of softwood, hardwood and tunicate CNF In nature, cellulose is commonly present in the crystalline form of cellulose I, although this form is not the thermodynamically minimum, but rather a local minimum. Commonly several cellulose chains are synthesized simultaneously in the cellulose synthetase complexes. There are two forms of cellulose I, cellulose Iα and cellulose Iβ, 55 that differs in how cellulose layers are organized towards each other in the fibrils.12 As calculated from FTIR data, all CNF are mainly in cellulose Iβ structures. However, the Iβ ratios of softwood CNF (around 80%) are higher than hardwood CNF (around 70%). This agreed well with the literature values; in woody cellulose, both cellulose Iα and cellulose Iβ are present, but the Iβ phase prevails with a proportion of above 64%.55 On the contrary, tunicate CNF is an almost pure cellulose Iβ structure (~94%) (Table 2). The reason for this might be that the shape and size of the tunicate fibrils is different from the 20 ACS Paragon Plus Environment

Page 21 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


wood fibrils, and in line with this it has been suggested that cellulose Iα in wood fibrils mainly occur on the surface. Neither pretreatment showed any indication of significant alternation of the Iβ ratios, and the only difference came from the cellulose sources. 3.1.3 Morphological structure As shown in Figure 3, all CNF after enzymatic pretreatment show clear nanofibrillar structures irrespective of the cellulose source and the differences in chemical structures. However, eTC consists of mainly individual fibrils while fibrillar aggregates are abundant in the wood CNF. This could be attributed to the differences in the chemical compositions (Table 1), but also due to the differences in crystallinity and crystal structure between wood and tunicate cellulose; a higher content of unordered cellulose and cellulose Iα on the fibril surface of plant cellulose fibrils might make them “stickier” and more prone to aggregate than the tunicate fibrils. During the homogenization, the wood celluloses are thus more difficult to fully disintegrated, and this might also be due to the presence of residual hemicelluloses and lignin that are cross-linked with the cellulose.56 In addition, eTC is morphologically flexible and its curved structures could be clearly observed in Figure 3. The nanofibers found in eTC have a width of 8.55±3.37 nm and a length of 2040±840 nm to several µm (Figure 4), and thus an aspect ratio (length/diameter) of at least 239. In the literature, tunicate nanocellulose was reported to have a width of 9.2±2.1 nm and a length of 1073±719 nm,57 to which the width value observed in this study agrees well although the observed length is higher. In addition, the aspect ratio for eTC in this study is significantly higher than the reported 70-148 for tunicate nanocellulose in the literature16, 31, and this lower value is a result from the harsh conditions used during the acid hydrolysis compared to the mechanical methods utilized in this work. Wood CNF exhibited similar lateral dimension with eSW being slightly wider (5.63±3.42 nm) than eHW (4.93±2.61 nm) (Figure 4).These 21 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

dimensions also correspond well with the 4-20 nm reported in the literature for similarly treated CNF.2, 16 Unfortunately, the length could not be accurately measured from AFM analysis for the enzymatically hydrolyzed wood CNF due to their aggregated fibrillary structures and curved morphologies.

Figure 3. AFM and SEM observations of softwood, hardwood and tunicate CNF


Page 22 of 40

Page 23 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 4. Width and length distributions of softwood, hardwood and tunicate CNF After the TEMPO-mediated oxidation pretreatment, the obtained tTC and tSW are dominated by individual fibrils. Apparently, the negatively charged carboxyl groups formed after the oxidation of the primary hydroxyl groups at the cellulose surface cause electrostatic repulsion between each other improving the fibril disintegration.24 In contrast to the flexible eTC, tTC shows more stretched rod-like structures, possibly due to the anionic charges.34 However, tSW shows fibrillar structures with more kinks than tTC (Figure 3). These kinks had been proposed to be generated from the processing conditions used in the preparation of the CNF43 or the alternating amorphous and crystalline domains of cellulose fibrils.54 tTC has a width of 7.75±2.36 nm and a length of 2.04±1.27 µm to several µm (Figure 4), thereby an aspect ratio of >263. 23 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Though eTC and tTC had similar measured mean lengths, longer fibrils with lengths of >5 µm exist in tTC than eTC (Figure 4). In contrast to tTC, tSW has thinner and shorter fibrils with a width of 2.33±0.83 nm and a mean length of 0.48±0.23 µm, thus an aspect ratio of ~209. In the literature, wood elementary fibril was reported to have a diameter of 3.5 nm and a length of 0.050.5 µm,16 with which our data for tSW agree well. Comparatively, the aspect ratio of tSW (209) is lower than for tTC (263). 3.2 Characteristics of CNF films

It was demonstrated that the physicochemical properties of CNF are closely related to the cellulose source and preparation method, and these differences play a critical role in their various applications. In order to investigate that, cast films, as one of the major utilization forms of CNF, have been prepared from different CNF and the comparison in terms of film performance has been conducted. Firstly, a suspension-casting-drying procedure was applied to prepare films from both wood and tunicate CNF. After casting from a dilute concentration (5g/L) the evaporation process was in fact a self-assembly process during which the CNF formed an intermingling network structure. Under the experimental setup applied, the thickness of the prepared films ranged from 16 to 22 µm (Table 3). 3.2.1 Film density All the prepared films have a density of 1.14-1.48 g/cm3 (Table 3), which is consistent to the density value (0.96-1.52 g/cm3) for the CNF films reported in the literature.33 Compared to the reported density of 1.68 g/cm3 for crystalline cellulose with cellulose Iβ crystal structure,58 the lower densities of ~1.5 g/cm3 for all the films observed in this study indicated the presence of empty space or pores among the fibrils in the films, which is confirmed by the pore volume of


Page 24 of 40

Page 25 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


0.001-0.068 cm3/g determined by BET analysis (Table 3) and the visible pore structures in the SEM images (Figure 5). Tunicate eTC and wood eSW or eHW films have a similar density of ~1.2 g/cm3 while tunicate tTC and wood tSW show higher densities of 1.37 and 1.48 g/cm3 respectively. Correspondingly, tTC and tSW films exhibited a lower surface area and a lower pore volume than their enzymatically pretreated counterparts (Table 4). As discussed above, the TEMPO-mediated oxidation resulted in shorter and thinner fibrils than the enzymatic hydrolysis. These shorter and thinner fibrils forms a denser network during the film preparation, thus resulting in the smaller surface area and lower pore volume.33 For all films, the density is found to increase with increasing charge density as shown in Figure 6a (R2=0.9194). With higher charge density the fibrils become better dispersed, which results in better packing of the fibrils during the film formation and eventually a film with smaller surface area and higher density.59

Table 3. Profiles of softwood, hardwood and tunicate CNF films (single value or mean±SD) CNF

Thickness (µm)

Density

RMS roughness

Surface area

Pore volume

Pore size

(g/cm3)

(nm)

(m2/g)

(cm3/g)

(nm)

eSW

20.0±1.0

1.17

21.1±4.3

14.6

0.013

3.6

tSW

16.0±1.0

1.48

2.9±0.5

7.9

0.005

2.6

eHW

21.6±0.5

1.14

19.5±7.6

2.8

0.001

1.4

eTC

17.7±0.9

1.18

38.0±7.6

31.4

0.068

8.7

tTC

19.2±1.5

1.37

3.8±0.3

19.9

0.051

10.3



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 26 of 40

Table 4. Properties of softwood, hardwood and tunicate CNF films (single value or mean±SD) Optical

Mechanical properties

Thermal property*

Hydrophobicity

property CNF

Transmit-

Tensile stress

Young’s

Tensile strain

DSC

tance (%) at

(MPa)

modulus

(%)

Tg (°C)

To (°C)

Tp (°C)

(°)

750 nm

TGA

Contact angle

(GPa)

eSW

14.5±1.4

79.0±3.9

5.8±0.7

3.1±0.5

202.7

273.3

313.8

17.4±0.8

tSW

90.2±0.6

81.8±3.8

10.6±1.2

0.9±0.1

232.1

252.1

305.2

10.8±1.5

eHW

14.0±0.7

69.0±0.6

4.6±0.1

5.3±1.1

203.7

284.7

320.3

17.7±1.7

eTC

2.2±0.1

177.5±20.3

12.6±0.1

4.1±0.3

228.4

325.1

359.6

60.5±1.4

tTC

22.0±1.1

144.9±12.3

12.7±1.4

1.4±0.1

201.8

268.2

314.9

35.8±1.5

*(Tg) glass transition temperature, (To) degradation onset temperature and (Tp) degradation peak temperature.

Figure 5. SEM images of softwood, hardwood and tunicate CNF films


Page 27 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 6. Correlations between film performance and CNF structures, (a) film density against charge density and (b) film tensile stress against crystallinity index

3.2.2 Film morphology Morphologically, CNF were randomly orientated in a mesh-like network, and numerous aggregates were observed in all the films from enzymatically pretreated CNF (Figure 5). The presence of aggregates resulted in the high RMS surface roughness of the films, 19-38 nm (Table 3). Especially, eTC film showed an intermingled network of long and thick microfibrils resulting in a particularly higher pore size of 8.7 nm compared to the shorter and more flexible wood CNF films with pore size of 1.4-3.6 nm (Table 3). The increased charge density of tTC and tSW caused the individual fibrils to stretch, and the stretched structure was preserved even after complete water evaporation (Figure 5). Both tTC and tSW films exhibited a lower RMS roughness (2.9-3.8 nm) than their enzymatically pretreated counterparts (Table 3), which is due to the overall smaller lateral size as shown in Figure 4. This effect was most pronounced in the tSW film which is denser than the tTC film when analyzing the SEM images, which is consistent to the higher density of 1.48 g/cm3 for tSW than 1.37 g/cm3 for tTC (Table 3). 27 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 28 of 40

3.2.3 Optical property All films fabricated from CNF after enzymatic pretreatment are visually transparent although their transmittances are only within the range of 2-14% at 750 nm (Figure 7 and Table 4). Irrespective of wood species, all films made from wood CNF have a transmittance of around 14%. eTC forms the opaquest film with a transmittance of 2.2% significantly lower than 14.5% and 14.0% for eSW and eHW, respectively. This difference might originate from that larger fibrils are present in eTC than the wood CNF, which would cause more light scattering and thus reduce the film transmittance.60 Additionally, the high porosity and presence of air bubbles within the films is another reason for the low transmittance. On the contrary, after TEMPO-mediated oxidation tTC (22.0%) and tSW (90.2%) films showed much higher transmittance than eTC (2.2%) and eSW (14.5%) (Table 4) due to the surface charge introduction, which improves the cellulose disintegration resulting in more elementary fibrils making the films more uniform and of higher density.59

Figure 7. Digital photos of softwood, hardwood and tunicate CNF films placed partly above a logo of the Royal Institute of Technology, KTH. The logotype is used under a formal permission form the Royal Institute of Technology, KTH. 28 ACS Paragon Plus Environment

Page 29 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


In wood, lignin has been bio-synthesized by polymerization of monolignols during which the electronic conjugation of the vinyl side chain group to the aromatic ring was lost. However, the polymerization resulted also in the generation of UV chromophores by which the lignin possesses absorption capability in the UV region.61 Therefore the presence of residual lignin in the wood CNF films exhibits a certain UV barrier phenomenon (Figure 8). However, both eTC and tTC also showed UV barrier properties, although lignin was absent in these samples (Table 1). This UV barrier effect should be caused by the nearly complete light reflection or scattering resulting from the thick fibrils of tunicate CNF, and this type of light scattering effect had been previously reported for wood nanofibers with width of larger than 50 nm.62

Figure 8. UV-vis spectra of softwood, hardwood and tunicate CNF films in the range of 200-400 nm 3.2.4 Mechanical property Among all the films, eTC film exhibited the highest tensile stress of 177.5 MPa, followed by 144.9 MPa for tTC (Table 4). However, both tunicate films have similar Young’s modulus values, 12.6 and 12.7 GPa, respectively. For the softwood samples a reversed trend was noted as



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 30 of 40

the tensile stress of tSW and eSW was similar (81.8 and 79.0 MPa) while the modulus was significantly higher for tSW (10.6 GPa) than eSW (5.8 GPa). This implies that the introduction of anionically charged carboxyl groups could induce formation of additional hydrogen bonds to those from the original hydroxyl groups, contributing to more inter-fiber bonding and thus a stronger network.63-64 Another possible explanation for this might be that the TEMPO-oxidized CNF are more evenly distributed prior to the film formation, and thus fewer “weak points” are formed. However, this effect was not observed for the eTC film perhaps due to the larger cellulose fibrils in the tunicate CNF making these effects less important. Films prepared from softwood (eSW) are slightly stronger than those made from hardwood (eHW), as seen by the tensile stress of 79.0 vs. 69.0 MPa and the Young’s modulus of 5.8 vs. 4.6 GPa. The strength of a film is closely correlated to the DP, crystallinity and aspect ratio of the CNF used. For example, the length of tunicate CNF is significantly longer than wood CNF, correspondingly the tensile stress of tunicate CNF films (144-177 MPa) is higher than those of wood CNF films (68-82 MPa). Furthermore, among these factors, it was demonstrated for all films that a higher crystallinity index yield higher tensile stress values with a correlation coefficient of R2=0.9704 (Figure 6b). After enzymatic hydrolysis, a tensile strain value of 4.1% was observed for the eTC films (Table 4), which is related to eTC’s flexible and long fibrillar structure. For wood CNF, the eSW films are more brittle than the eHW films, with a tensile strain value of 3.1 vs. 5.3% (Table 4). On the other hand, after TEMPO-mediated oxidation, the tensile strains are 1.4 and 0.9% for tTC and tSW films, respectively, lower than 4.1 and 3.1% for eTC and eSW counterparts, respectively. Here, both tTC and tSW have high charge densities, 428 µmol/g and 443 µmol/g (Table 2),


Page 31 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


respectively. The highly charged groups at the surface cause CNF to stretch in the formed film networks, thus reducing the films’ tensile strains.

3.2.5 Thermal property By the thermogravimetric analysis of all the films, the highest degradation onset temperature (To) (325°C) and the highest degradation peak temperature (Tp) (360°C) were observed for eTC film (Table 4 and Figure 9). In contrast to eTC film, all wood CNF films after the same enzymatic pretreatment showed lower To (273-285°C) and Tp (314-320°C) (Table 4). This might be attributable to their lower DP, thinner fibrils and lower crystallinities or the fact that the tunicate CNF has a lower content of contaminations, i.e. non-cellulose materials (Table 1). A detail comparison between eSW and eHW films shows that eHW film is slightly more thermally stable than eSW film, To of 285 vs. 273°C and Tp of 320 vs. 314°C. This is also in line with the idea that non-cellulosic compounds lower the terminal stability, since the cellulose content appears to be somewhat higher in the hardwood material (Table 1), but the higher thermal stabilities might also originate from the slightly higher CI of eHW than eSW, 77.13 vs. 75.47% (Table 2). It was previously reported that the thermal decomposition of cellulose shifted to higher temperatures with higher CI.65 After TEMPO-mediated oxidation, To (268 and 252°C) and Tp (315 and 305°C) for tTC and tSW are lower than To (325 and 273°C) and Tp (360 and 314°C) for their counterparts after enzymatic hydrolysis (eTC and eSW, respectively) (Table 4). This is due to the fact that during heating the additional carboxyl groups bring about additional decarboxylation reactions from the anhydroglucuronate units present on the CNF surfaces which cause thermal instability, as it was reported that carboxymethyl cellulose sodium salt (CMC-Na) had a lower To of ~ 230 °C and a lower Tp of ~280 °C.66 31 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 9. TGA (a) and DSC (b) curves of softwood, hardwood and tunicate CNF films. As learned from differential scanning calorimetry (DSC) analysis, the glass transition temperature (Tg) for all films is generally between 200-230°C. For the films of CNF after enzymatic hydrolysis, eTC film has the highest Tg (228 °C) and the films from wood CNF showed lower but similar Tg (203-204 °C). For the CNF films after TEMPO-mediated oxidation, tTC film has a Tg of 202°C, lower than the counterpart after enzymatic hydrolysis (eTC film of 228 °C). However, tSW film shows a higher Tg (232 °C) than the corresponding eSW film (203 °C). 3.2.6 Hydrophobicity Contact angle measurements showed that all CNF films are hydrophilic, as seen by the low contact angles (11-60°) (Table 4). There are several factors closely related to the contact angle including chemical composition, surface texture (roughness and particle shape) and surface chemistry (heterogeneity) etc. of the tested materials.67 The abundance of hydroxyl groups on the cellulose is the primary reason for the hydrophilicity seen in all samples.68 In particular, the contact angles for the tTC and tSW films are only 36° and 11°, respectively, significantly lower 32 ACS Paragon Plus Environment

Page 32 of 40

Page 33 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


than their counterparts after enzymatic hydrolysis, namely 60° and 17° for eTC and eSW, respectively (Table 4). This is because of the introduced COO- groups that have higher water affinity than hydroxyl groups.69 However, it is surprising to see the highest contact angle for eTC film (60°) since the inherent cellulose structure is identical to other samples. The presence of pores as shown in Figure 5 might be the reason for the high contact angle. According to the Cassie and Baxter’s model, when the size of the pores is small enough the probe water drop cannot penetrate into the pores but will instead trap air in between the sample and water droplet. This causes an increased contact angle reading.70

4. CONCLUSIONS Irrespective of cellulose sources, homogenization after a pretreatment could process wood or tunicate celluloses into cellulose nanofibers (CNF). There is a structural diversity present among the CNF prepared in terms of chemical composition, morphology and crystalline structure. These differences may stem from the raw materials or be related to the procedures used for the CNF preparation. Tunicate CNF chemically consist of purer cellulose and have higher degrees of polymerization (DP) and crystallinity than the wood CNF that contain residual hemicelluloses and lignin. The crystalline structure is nearly pure Iβ in the tunicate CNF whereas the wood CNF contains both Iα and Iβ. Physically, more fibrillar aggregates are present in the wood CNF while tunicate CNF are dominated by individual fibrils; however, the tunicate elementary fibrils are significantly larger than their wood counterparts. The pretreatment by enzymatic hydrolysis alters slightly cellulose’s chemical structure while the pretreatment by TEMPO-mediated oxidation noticeably results in CNFs with lower DP, smaller nanofiber and crystal size. The Iβ ratio of CNF is independent to pretreatment, but determined by cellulose source. 33 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

The source of CNF has a significant impact on the structure, morphology, optical, mechanical, thermal and hydrophobic properties of the prepared films. Tunicate CNF films exhibit better mechanical properties and higher thermal stability, while the wood CNF films are more transparent due to their thinner and shorter fibrillar structures. Irrespective of the cellulose sources, all the films obtained from the CNF prepared after TEMPO-mediated oxidation pretreatment are denser and smoother than the enzymatic hydrolysis pretreated counterparts. This results in the formation of more uniform and transparent films. However, the oxidation deteriorates the thermal stability and hydrophobicity, although the mechanical properties are not affected significantly. Distinctively, this study has been conducted on direct and comprehensive comparisons among different CNF isolated from softwood, hardwood and tunicate, and their films. Conclusively, the wood and tunicate CNF should be considered as two distinctly different types. They should further be grouped into two categories, one obtained after enzymatic hydrolysis and another after TEMPO-mediated oxidation pretreatment. In fact, the structures and physiochemical properties of CNF and their films could be tuned by varying cellulose source and preparation processes as well as blending different CNFs together to meet different application demands. The results from this study can provide useful information and knowledge for the practical preparations and applications of wood and tunicate cellulose or CNF.


Page 34 of 40

Page 35 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


AUTHOR INFORMATION Corresponding Author *E-mail: [email protected] Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ORCID Yadong Zhao: 0000-0001-8208-4938 Carl Moser: 0000-0002-8125-7734 Jiebing Li: 0000-0002-4521-1122 Notes The authors declare no competing financial interest.

ACKNOWLEDGEMENTS We would like to thank Viktor Bladholm, Adam Britts, Alexandra Edberg and Anmol Khawaja from the Royal Institute of Technology, KTH, for their help regarding the production of wood CNF. Christofer Troedsson, Eric Thompson and Jean-Marie Bouquet from University of Bergen are acknowledged for collecting tunicate samples in Norway. Carl Moser would like to thank the Stiftelsen för kunskaps- och kompetensutveckling (KK-stiftelsen) for supporting his Ph.D. study at KTH. 35 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 36 of 40

REFERENCES (1) Huber, T.; Müssig, J.; Curnow, O.; Pang, S.; Bickerton, S.; Staiger, M. P. A Critical Review of All-cellulose Composites. J. Mater. Sci. 2012, 47, 1171-1186. (2) Habibi, Y.; Lucia, L. A.; Rojas, O. J. Cellulose Nanocrystals: Chemistry, Self-assembly, and Applications. Chem. Rev. 2010, 110, 3479-3500. (3) Kimura, S.; Itoh, T. Biogenesis and Function of Cellulose in the Tunicates. In Cellulose: Molecular and Structural Biology: Selected Articles on the Synthesis, Structure, and Applications of Cellulose, Brown, R. M.; Saxena, I. M., Eds. Springer Netherlands: Dordrecht, 2007, pp 217236. (4) Zhao, Y.; Li, J. Ascidian Bioresources: Common and Variant Chemical Compositions and Exploitation Strategy - Examples of Halocynthia roretzi, Styela plicata, Ascidia sp. and Ciona intestinalis. Z. Naturforsch., C: J. Biosci. 2016, 71, 165-80. (5) Zhao, Y.; Li, J. Excellent Chemical and Material Cellulose from Tunicates: Diversity in Cellulose Production Yield and Chemical and Morphological Structures from Different Tunicate Species. Cellulose 2014, 21, 3427-3441. (6) Hirose, E. Ascidian Tunic Cells: Morphology and Functional Diversity of Free Cells outside the Epidermis. Invertebrate Biology 2009, 128, 83-96. (7) Hirose, E. Acid Containers and Cellular Networks in the Ascidian Tunic with Special Remarks on Ascidian Phylogeny. Zool. Sci. 2001, 18, 723-731. (8) Sella Kapu, N.; Trajano, H. L. Review of Hemicellulose Hydrolysis in Softwoods and Bamboo. Biofuels, Bioprod. Biorefin. 2014, 8, 857-870. (9) Butterfield, B. The Structure of Wood: Form and Function. In Primary Wood Processing: Principles and Practice, Walker, J. C. F., Ed. Springer Netherlands: Dordrecht, 2006, pp 1-22. (10) Liu, Q.; Wang, S.; Zheng, Y.; Luo, Z.; Cen, K. Mechanism Study of Wood Lignin Pyrolysis by Using TG-FTIR Analysis. J. Anal. Appl. Pyrolysis 2008, 82, 170-177. (11) Meier, H. Chemical and Morphological Aspects of the Fine Structure of Wood. Pure and Applied Chemistry 1962, 5, 37-52. (12) Brown Jr, R. M.; Saxena, I. M. Cellulose Biosynthesis: A Model for Understanding the Assembly of Biopolymers. Plant Physiol. Biochem. 2000, 38, 57-67. (13) Miao, C.; Hamad, W. Y. Cellulose Reinforced Polymer Composites and Nanocomposites: A Critical Review. Cellulose 2013, 20, 2221-2262. (14) Kalia, S.; Dufresne, A.; Cherian, B. M.; Kaith, B. S.; Avérous, L.; Njuguna, J.; Nassiopoulos, E. Cellulose-Based Bio- and Nanocomposites: A Review. Int. J. Polym. Sci. 2011, 2011, 837875. (15) Mariano, M.; El Kissi, N.; Dufresne, A. Cellulose Nanocrystals and Related Nanocomposites: Review of Some Properties and Challenges. J. Polym. Sci., Part B: Polym. Phys. 2014, 52, 791-806. (16) Moon, R. J.; Martini, A.; Nairn, J.; Simonsen, J.; Youngblood, J. Cellulose Nanomaterials Review: Structure, Properties and Nanocomposites. Chem. Soc. Rev. 2011, 40, 3941-3994. (17) Siró, I.; Plackett, D. Microfibrillated Cellulose and New Nanocomposite Materials: A Review. Cellulose 2010, 17, 459-494. (18) Islam, M. T.; Alam, M. M.; Patrucco, A.; Montarsolo, A.; Zoccola, M. Preparation of Nanocellulose: A Review. AATCC J. Res. 2014, 1, 17-23. (19) Agustin, M. B.; Nakatsubo, F.; Yano, H. The Thermal Stability of Nanocellulose and Its Acetates with Different Degree of Polymerization. Cellulose 2016, 23, 451-464. (20) Araki, J.; Wada, M.; Kuga, S.; Okano, T. Flow Properties of Microcrystalline Cellulose Suspension Prepared by Acid Treatment of Native Cellulose. Colloids Surf., A 1998, 142, 75-82. 36 ACS Paragon Plus Environment

Page 37 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(21) Ummartyotin, S.; Manuspiya, H. A Critical Review on Cellulose: From Fundamental to An Approach on Sensor Technology. Renewable Sustainable Energy Rev. 2015, 41, 402-412. (22) Moser, C.; Lindström, M. E.; Henriksson, G. Toward Industrially Feasible Methods for Following the Process of Manufacturing Cellulose Nanofibers. BioResources 2015, 10, 23602375. (23) Henriksson, M.; Henriksson, G.; Berglund, L. A.; Lindström, T. An Environmentally Friendly Method for Enzyme-assisted Preparation of Microfibrillated Cellulose (MFC) Nanofibers. Eur. Polym. J. 2007, 43, 3434-3441. (24) Saito, T.; Kimura, S.; Nishiyama, Y.; Isogai, A. Cellulose Nanofibers Prepared by TEMPOMediated Oxidation of Native Cellulose. Biomacromolecules 2007, 8, 2485-2491. (25) Rodionova, G.; Saito, T.; Lenes, M.; Eriksen, Ø.; Gregersen, Ø.; Fukuzumi, H.; Isogai, A. Mechanical and Oxygen Barrier Properties of Films Prepared from Fibrillated Dispersions of Tempo-Oxidized Norway Spruce and Eucalyptus Pulps. Cellulose 2012, 19, 705-711. (26) Xu, W.; Steinschulte, A. A.; Plamper, F. A.; Korolovych, V. F.; Tsukruk, V. V. Hierarchical Assembly of Star Polymer Polymersomes into Responsive Multicompartmental Microcapsules. Chem. Mater. 2016, 28, 975-985. (27) Ye, C.; Malak, S. T.; Hu, K.; Wu, W.; Tsukruk, V. V. Cellulose Nanocrystal Microcapsules as Tunable Cages for Nano- and Microparticles. ACS Nano 2015, 9, 10887-10895. (28) Svagan, A. J.; Musyanovych, A.; Kappl, M.; Bernhardt, M.; Glasser, G.; Wohnhaas, C.; Berglund, L. A.; Risbo, J.; Landfester, K. Cellulose Nanofiber/Nanocrystal Reinforced Capsules: A Fast and Facile Approach Toward Assembly of Liquid-Core Capsules with High Mechanical Stability. Biomacromolecules 2014, 15, 1852-1859. (29) Liu, J.; Lan, Y.; Yu, Z.; Tan, C. S. Y.; Parker, R. M.; Abell, C.; Scherman, O. A. Cucurbit[n]uril-Based Microcapsules Self-Assembled within Microfluidic Droplets: A Versatile Approach for Supramolecular Architectures and Materials. Acc. Chem. Res. 2017, 50, 208-217. (30) Korolovych, V. F.; Bulavin, L. A.; Prylutskyy, Y. I.; Khrapatiy, S. V.; Tsierkezos, N. G.; Ritter, U. Influence of Single-Walled Carbon Nanotubes on Thermal Expansion of Water. Int. J. Thermophys. 2014, 35, 19-31. (31) Sacui, I. A.; Nieuwendaal, R. C.; Burnett, D. J.; Stranick, S. J.; Jorfi, M.; Weder, C.; Foster, E. J.; Olsson, R. T.; Gilman, J. W. Comparison of the Properties of Cellulose Nanocrystals and Cellulose Nanofibrils Isolated from Bacteria, Tunicate, and Wood Processed Using Acid, Enzymatic, Mechanical, and Oxidative Methods. ACS Appl. Mater. Interfaces 2014, 6, 61276138. (32) Spence, K. L.; Venditti, R. A.; Habibi, Y.; Rojas, O. J.; Pawlak, J. J. The Effect of Chemical Composition on Microfibrillar Cellulose Films from Wood Pulps: Mechanical Processing and Physical Properties. Bioresour. Technol. 2010, 101, 5961-5968. (33) Kumar, V.; Bollström, R.; Yang, A.; Chen, Q.; Chen, G.; Salminen, P.; Bousfield, D.; Toivakka, M. Comparison of Nano- and Microfibrillated Cellulose Films. Cellulose 2014, 21, 3443-3456. (34) Zhao, Y.; Zhang, Y.; Lindström, M. E.; Li, J. Tunicate Cellulose Nanocrystals: Preparation, Neat Films and Nanocomposite Films with Glucomannans. Carbohydr. Polym. 2015, 117, 286296. (35) Okita, Y.; Saito, T.; Isogai, A. Entire Surface Oxidation of Various Cellulose Microfibrils by TEMPO-mediated Oxidation. Biomacromolecules 2010, 11, 1696-700. (36) Okita, Y.; Fujisawa, S.; Saito, T.; Isogai, A. TEMPO-oxidized Cellulose Nanofibrils Dispersed in Organic Solvents. Biomacromolecules 2011, 12, 518-522. 37 ACS Paragon Plus Environment


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 38 of 40

(37) de Morais Teixeira, E.; Corrêa, A. C.; Manzoli, A.; de Lima Leite, F.; de Oliveira, C. R.; Mattoso, L. H. C. Cellulose Nanofibers from White and Naturally Colored Cotton Fibers. Cellulose 2010, 17, 595-606. (38) da Silva Perez, D.; Montanari, S.; Vignon, M. R. TEMPO-mediated Oxidation of Cellulose III. Biomacromolecules 2003, 4, 1417-1425. (39) Habibi, Y.; Chanzy, H.; Vignon, M. TEMPO-mediated Surface Oxidation of Cellulose Whiskers. Cellulose 2006, 13, 679-687. (40) Rusli, R.; Shanmuganathan, K.; Rowan, S. J.; Weder, C.; Eichhorn, S. J. Stress Transfer in Cellulose Nanowhisker Composites—Influence of Whisker Aspect Ratio and Surface Charge. Biomacromolecules 2011, 12, 1363-1369. (41) Araki, J.; Wada, M.; Kuga, S. Steric Stabilization of A Cellulose Microcrystal Suspension by Poly(ethylene glycol) Grafting. Langmuir 2000, 17, 21-27. (42) Isogai, A.; Saito, T.; Fukuzumi, H. TEMPO-oxidized Cellulose Nanofibers. Nanoscale 2011, 3, 71-85. (43) Usov, I.; Nystrom, G.; Adamcik, J.; Handschin, S.; Schutz, C.; Fall, A.; Bergstrom, L.; Mezzenga, R. Understanding Nanocellulose Chirality and Structure-properties Relationship at the Single Fibril Level. Nat. Commun. 2015, 6, 7564. (44) Reddy, N.; Yang, Y. Structure and Properties of High Quality Natural Cellulose Fibers from Cornstalks. Polymer 2005, 46, 5494-5500. (45) French, A. D.; Santiago Cintrón, M. Cellulose Polymorphy, Crystallite Size, and the Segal Crystallinity Index. Cellulose 2013, 20, 583-588. (46) Rondeau-Mouro, C.; Bouchet, B.; Pontoire, B.; Robert, P.; Mazoyer, J.; Buléon, A. Structural Features and Potential Texturising Properties of Lemon and Maize Cellulose Microfibrils. Carbohydr. Polym. 2003, 53, 241-252. (47) Fernandes Diniz, J. M. B.; Gil, M. H.; Castro, J. A. A. M. Hornification—Its Origin and Interpretation in Wood Pulps. Wood Sci. Technol. 2004, 37, 489-494. (48) Luo, X.; Zhu, J. Y. Effects of Drying-induced Fiber Hornification on Enzymatic Saccharification of Lignocelluloses. Enzyme Microb. Technol. 2011, 48, 92-99. (49) Sun, J. X.; Sun, X. F.; Zhao, H.; Sun, R. C. Isolation and Characterization of Cellulose from Sugarcane Bagasse. Polym. Degrad. Stab. 2004, 84, 331-339. (50) Pavithra, R.; Gunasekaran, S.; Sailatha, E.; Kamatchi, S. Investigations on Paper Making Raw Materials and Determination of Paper Quality by FTIR-UATR and UV-Vis DRS Spectroscopy. Int. J. Curr. Res. Aca. Rev. 2015, 3, 42-59. (51) Nakashima, K.; Sugiyama, J.; Satoh, N. A Spectroscopic Assessment of Cellulose and the Molecular Mechanisms of Cellulose Biosynthesis in the Ascidian Ciona intestinalis. Marine Genomics 2008, 1, 9-14. (52) Jiang, F.; Hsieh, Y. L. Chemically and Mechanically Isolated Nanocellulose and Their Selfassembled Structures. Carbohydr. Polym. 2013, 95, 32-40. (53) Célino, A.; Freour, S.; Jacquemin, F.; Casari, P. The Hygroscopic Behavior of Plant Fibers: A Review. Front. Chem. 2014, 1, 43. (54) Montanari, S.; Roumani, M.; Heux, L.; Vignon, M. R. Topochemistry of Carboxylated Cellulose Nanocrystals Resulting from TEMPO-Mediated Oxidation. Macromolecules 2005, 38, 1665-1671. (55) Gray, D. Nanocellulose: From Nature to High Performance Tailored Material. Holzforschung 2013, 67, 353-353.


Page 39 of 40

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


(56) Lawoko, M.; Henriksson, G.; Gellerstedt, G. Characterisation of Lignin-carbohydrate Complexes (LCCs) of Spruce Wood (Picea abies L.) Isolated with Two Methods. Holzforschung 2006, 60, 156-161. (57) Elazzouzi-Hafraoui, S.; Nishiyama, Y.; Putaux, J.-L.; Heux, L.; Dubreuil, F.; Rochas, C. The Shape and Size Distribution of Crystalline Nanoparticles Prepared by Acid Hydrolysis of Native Cellulose. Biomacromolecules 2008, 9, 57-65. (58) Diddens, I.; Murphy, B.; Krisch, M.; Müller, M. Anisotropic Elastic Properties of Cellulose Measured Using Inelastic X-ray Scattering. Macromolecules 2008, 41, 9755-9759. (59) Fukuzumi, H.; Saito, T.; Iwata, T.; Kumamoto, Y.; Isogai, A. Transparent and High Gas Barrier Films of Cellulose Nanofibers Prepared by TEMPO-mediated Oxidation. Biomacromolecules 2009, 10, 162-165. (60) Borch, J.; Marchessault, R. H. Light Scattering by Cellulose. I. Native Cellulose Films. J. Polym. Sci., Part C: Polym. Symp. 1969, 28, 153-167. (61) Dean, J. C.; Navotnaya, P.; Parobek, A. P.; Clayton, R. M.; Zwier, T. S. Ultraviolet Spectroscopy of Fundamental Lignin Subunits: Guaiacol, 4-methylguaiacol, Syringol, and 4methylsyringol. J. Chem. Phys. 2013, 139, 144313. (62) Bhattacharya, M.; Malinen, M. M.; Lauren, P.; Lou, Y.-R.; Kuisma, S. W.; Kanninen, L.; Lille, M.; Corlu, A.; GuGuen-Guillouzo, C.; Ikkala, O.; Laukkanen, A.; Urtti, A.; Yliperttula, M. Nanofibrillar Cellulose Hydrogel Promotes Three-dimensional Liver Cell Culture. J. Controlled Release 2012, 164, 291-298. (63) Zhao, C.; Zhang, H.; Zeng, X.; Li, H.; Sun, D. Enhancing the Inter-fiber Bonding Properties of Cellulosic Fibers by Increasing Different Fiber Charges. Cellulose 2016, 23, 1617-1628. (64) Fujisawa, S.; Okita, Y.; Fukuzumi, H.; Saito, T.; Isogai, A. Preparation and Characterization of TEMPO-oxidized Cellulose Nanofibril Films with Free Carboxyl Groups. Carbohydr. Polym. 2011, 84, 579-583. (65) Kim, U.-J.; Eom, S. H.; Wada, M. Thermal Decomposition of Native Cellulose: Influence on Crystallite Size. Polym. Degrad. Stab. 2010, 95, 778-781. (66) de Britto, D.; Assis, O. B. G. Thermal Degradation of Carboxymethylcellulose in Different Salty Forms. Thermochim. Acta 2009, 494, 115-122. (67) Chau, T. T.; Bruckard, W. J.; Koh, P. T. L.; Nguyen, A. V. A Review of Factors That Affect Contact Angle and Implications for Flotation Practice. Adv. Colloid Interface Sci. 2009, 150, 106-115. (68) Osovskaya, I. I.; Poltoratskii, G. M.; Dmitrieva, E. A.; Kamyshanskaya, Y. V. Hydrophilic Characteristics of Cellulose Treated with Saturated Water Vapor. Russ. J. Appl. Chem. 2005, 78, 1182-1184. (69) Hakalahti, M.; Mautner, A.; Johansson, L.-S.; Hänninen, T.; Setälä, H.; Kontturi, E.; Bismarck, A.; Tammelin, T. Direct Interfacial Modification of Nanocellulose Films for Thermoresponsive Membrane Templates. ACS Appl. Mater. Interfaces 2016, 8, 2923-2927. (70) Michael, N.; Bharat, B. Roughness-induced Superhydrophobicity: A Way to Design Nonadhesive Surfaces. J. Phys.: Condens. Matter 2008, 20, 225009.



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

TOC Figure


Page 40 of 40

Cellulose Nanofibers from Softwood, Hardwood, and Tunicate

Recommend Documents