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Different Conformations of Surface Cellulose Molecules in Native Cellulose Microfibrils Revealed by Layer-by-Layer Peeling Ryunosuke Funahashi, Yusuke Okita, Hiromasa Hondo, Mengchen Zhao, Tsuguyuki Saito, and Akira Isogai Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.7b01173 • Publication Date (Web): 27 Sep 2017 Downloaded from http://pubs.acs.org on September 28, 2017
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Biomacromolecules
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Different Conformations of Surface Cellulose
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Molecules in Native Cellulose Microfibrils
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Revealed by Layer-by-Layer Peeling
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Ryunosuke Funahashi, Yusuke Okita, Hiromasa Hondo, Mengchen Zhao, Tsuguyuki Saito, and
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Akira Isogai*
7 8
Department of Biomaterials Science, Graduate School of Agricultural and Life Sciences, The
9
University of Tokyo, Tokyo 113-8657, Japan
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ABSTRACT: Layer-by-layer peeling of surface molecules of native cellulose microfibrils was
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performed using a repeated sequential process of 2,2,6,6-tetramethylpiperidine-1-oxyl
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radical-mediated oxidation followed by hot alkali extraction. Both highly crystalline algal and
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tunicate celluloses and low-crystalline cotton and wood celluloses were investigated. Initially,
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the C6-hydroxy groups of the outermost surface molecules of each algal cellulose microfibril
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facing the exterior had the gauche–gauche (gg) conformation, whereas those facing the interior
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had the gauche–trans (gt) conformation. All the other C6-hydroxy groups of the cellulose
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molecules inside the microfibrils contributing to crystalline cellulose I had the trans–gauche
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(tg) conformation. After surface peeling, the originally 2nd-layer molecules from the
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microfibril surface became the outermost surface molecules, and the original tg conformation
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changed to gg and gt conformations. The plant cellulose microfibrils likely had disordered
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structures for both the outermost surface and 2nd-layer molecules, as demonstrated using the
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same layer-by-layer peeling technique.
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KEYWORDS: cellulose microfibril, layer-by-layer peeling, conformation, nuclear magnetic
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resonance, TEMPO-mediated oxidation
27 28
INTRODUCTION
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Cellulose, the most abundant extracellular polysaccharide on Earth, is a linear
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homopolysaccharide consisting of D-glucopyranosyl units linked by β-1,4-glucoside bonds.1
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Plant cellulose molecules are polymerized by cellulose-synthesizing enzyme complexes in the
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plasma membrane, directed to the exit channel of the complexes, and crystallized into cellulose
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microfibrils.2‒9 The cellulose microfibrils are self-assembled and then deposited on primary
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cell walls as a scaffold to form secondary cell walls, which contribute to the tough physical
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properties of skeletal tissues.3 Cellulose microfibrils are predominately generated by vascular
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plants as well as a large number of algae,6 certain bacteria,2,3 and tunicates.5 It has been
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proposed that plant cellulose microfibrils have disordered surface regions; however, the
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cross-sectional structures of cellulose microfibrils including the number of cellulose chains in
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each microfibril remain under debate.10‒15
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Native cellulose microfibrils have received considerable attention for nanotechnology
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applications because they are the most abundant reproducible bio-based nanofibers on Earth.
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Moreover, unlike other inorganic and petroleum-based organic nanomaterials, cellulose
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microfibrils have unique characteristics such as small widths of 3‒20 nm (depending on the
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origin of the cellulose), high aspect ratios, high mechanical strength,16 high moduli,17,18 low
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coefficients of thermal expansion,19 and large surfaces areas. Therefore, many researches have
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explored methods for efficiently preparing nanosized celluloses (or nanocelluloses) from
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micrometer-sized plant cellulose fibers and the use of nanocelluloses as fillers and scaffolds for
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composites and bulk materials such as films, porous materials, and hydrogels.20–25 Some of the
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resultant materials have exhibited unique mechanical, thermal, optical, catalytic, electric,
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oxygen-barrier, absorbent, and biological properties, which demonstrate their potential
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application in high-tech material fields. The nanosized morphologies, characteristics, and
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functions of the prepared nanocelluloses intrinsically originate from the biosynthesized
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cellulose microfibrils. The structures of cellulose microfibrils and their molecular-level surface
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characteristics thus significantly affect the efficient conversion of cellulose fibers into
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nanocelluloses as well as on their functions and applications.
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In this study, we investigated the surface structures of highly crystalline cellulose
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microfibrils from Cladophora sp. and Halocynthia roretzi and low-crystalline microfibrils
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from cotton lint, cotton linters, and wood cellulose using layer-by-layer peeling of surface
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molecules. Solid-state
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conductivity titration, and X-ray diffraction (XRD) analyses of the resultant structures were
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performed focusing on the conformations of the C6-hydroxy groups of the cellulose
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microfibrils and their changes during repetitions of the surface peeling process. Furthermore,
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we determined the local mobility of cellulose molecules and their C6-OH groups in wet and
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dry states using 13C NMR spectroscopy.
13
C nuclear magnetic resonance (NMR) spectroscopy, electrical
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MATERIALS AND METHODS Materials. Marine green alga, Cladophora sp., was collected at the sea of Chikura (Chiba,
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Japan). A tunicin, Halocynthia roretzi, was obtained at a local fish market. These algal and
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tunicate celluloses were washed thoroughly with water, soaked in aqueous 0.1 M HCl at RT
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overnight, and then repeatedly bleached with fresh aqueous 0.3% (w/v) NaClO2 at pH 4–5 and
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70 °C until the products turned white. These algal and tunicate celluloses were cut into short
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filaments and small pieces, respectively, using scissors and further purified with 0.1 M HCl
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and then 4% (w/w) NaOH at room temperature (RT) overnight. Next, 0.5% slurries (200 mL
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each) of the purified algal and tunicate cellulose in water were mechanically disintegrated at
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7500 rpm for 10 min using a double-cylinder-type homogenizer (Physcotron NS-56, Microtec
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Nition, Chiba, Japan). To obtain the cotton lint cellulose, an American cotton ball was
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purchased from Japan Cotton Promotion Institute (Tokyo, Japan). The cotton lint cellulose was
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soaked in aqueous 90% (v/v) acetone at RT for 1 day to remove the wax components, followed
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by rinsing with 90% (v/v) acetone and then thoroughly with water using filtration. The cotton
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cellulose was cut into short filaments with lengths of ~3 mm, and the wet cellulose (~2 g) was
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added to a mixture containing NaClO2 (2.28 g) and 0.1 M acetate buffer (200 mL, pH 4.8).
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Bleaching was performed by stirring the mixture at RT for 3 days, followed by washing
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thoroughly with water using filtration. A commercial filter pulp (Advantec, Tokyo, Japan) was
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used as the cotton linters cellulose. A never-dried bleached softwood kraft pulp containing
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~90% α-cellulose and ~10% mostly hemicelluloses was obtained from Nippon Paper Industry
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(Tokyo, Japan). All the reagents and solvents were laboratory grade (Wako Pure Chemicals,
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Japan) and used as received.
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TEMPO-Mediated Oxidation. The wet cellulose sample with a weight corresponding to a
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dry weight of 2 g was suspended in water (200 mL) containing TEMPO (0.032 g) and NaBr
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(0.2 g).26,27 The oxidation was started by adding 1.8 M NaClO solution to the cellulose
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suspensions: 10 mmol/g NaClO for the algal, tunicate, and cotton celluloses and 3.8 mmol/g 4 ACS Paragon Plus Environment
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NaClO for the wood cellulose. The suspension was maintained at pH 10 by continuous
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addition of 0.5 M NaOH using a pH stat (AUT-701, DKK-TOA, Tokyo, Japan) for 4 h for the
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algal, tunicate, and cotton celluloses and for 40 min for the wood cellulose. The oxidized
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celluloses were thoroughly washed with water using filtration and stored at 4 °C without
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drying.
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TEMPO-Oxidized Cellulose Nanofibrils. The 0.1% (w/v) TEMPO-oxidized algal, tunicate,
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cotton, and wood cellulose/water slurries were mechanically disintegrated at 7500 rpm for 2 min
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using the double-cylinder-type homogenizer. The obtained TEMPO-oxidized cellulose
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nanofibril/water dispersions were then sonicated for 6‒12 min using an ultrasonic homogenizer
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with a 7-mm probe tip diameter at 19.5 kHz and 300-W output power (US-300T, Nihon Seiki,
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Tokyo, Japan). The unfibrillated fraction, if present in the dispersion, was removed by
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centrifugation at 12 000×g for 10 min.
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Layer-By-Layer Surface Peeling. A solution of 2 M aqueous NaOH (200 mL) was slowly
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poured into the 0.2% TEMPO-oxidized cellulose suspension (200 mL) containing NaBH4
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(0.01 g). The mixture was kept at 105 °C for 12 h after bubbling N2 gas into the mixture for 10
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min. After being washed thoroughly with water using centrifugation, the water-insoluble
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residue was oxidized again using the same TEMPO/NaBr/NaClO system in water at pH 10
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described above, and the obtained TEMPO-oxidized cellulose was then extracted with 1 M
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NaOH under the same conditions described above.
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Analyses. The carboxylate contents of the oxidized celluloses were determined using the
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electrical conductivity titration method.28 AFM images of the TEMPO-oxidized cellulose
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nanofibrils were captured using an atomic force microscope (NanoScope IIIa, Veeco
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Instruments, Inc., USA) with a Bruker MPP-11100-10 tip operating in tapping mode. The largest
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height value measured along each isolated nanofibril was taken as the true height value, as the 5 ACS Paragon Plus Environment
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tapping step (512×512 pixel images for 5-µm-wide squares) was large compared with the narrow
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(~10 nm wide) nanofibrils. More than 60 nanofibrils were measured for each sample. The
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freeze-dried samples (~0.1 g each) were pressed at approximately 750 MPa for 1 min to make
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pellets. XRD patterns were recorded for the pellets for 2θ diffraction angles between 10° and 30°
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in reflection mode using a diffractometer (RINT 2000, Rigaku, Tokyo, Japan) with Ni-filtered
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Cu Kα radiation (λ = 0.1542 nm) at 40 kV and 40 mA. The crystal widths of cellulose I were
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calculated from the full widths at half maximums of the (2 0 0) diffraction peaks of the tunicate,
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cotton and wood celluloses using Scherrer’s equation.29 Two peaks centered at approximately
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14.8° and 16.8° in the XRD patterns of the algal celluloses, which correspond to d spacings of
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0.60‒0.61 and 0.53‒0.54 nm, respectively, were deconvoluted via curve-fitting using a
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pseudo-Voigt function.30 The water contents of the never-dried and re-wetted samples inside an NMR rotor were
127 128
approximately
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Cross-polarization/magic angle sample spinning (CP/MAS)
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performed using an NMR spectrometer (JNM-ECAII 500, JEOL, Japan) operating at 125.77
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MHz for
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performed at 298 K under the following conditions: sample spinning rate of 6 kHz, proton 90°
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pulse time of 2.5 µs, and relaxation delay of 5 s. The CP transfer was achieved using a ramped
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amplitude sequence (RAMP/CP) for a CP contact time of 2 ms and an amplitude graduation of
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7%. Adamantane was used as an external standard for the ppm calculation. Each CP/MAS
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spectrum was deconvoluted into Gaussian and Lorentzian components (i.e., Voigt components)
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at 60‒62, 62.5‒64.5, and 65.6‒66.6 ppm assigned to C6-OH conformations of gg, gt, and tg,
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respectively, using FeakFit version 4.12 (Seasolve Software Inc., San Jose, CA, USA).31,32 The
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13
13
50‒60%;
the
dried
samples
were
prepared 13
after
freeze-drying.
C NMR measurements were
C with a 3.2-mm HXMAS probe and a ZrO2 rotor. All the measurements were
C T1-measurement was performed using the pulse sequence developed by Torchia,33 with a 6 ACS Paragon Plus Environment
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relaxation delay of 10 s and eight τ values between 0.1 and 60 s for the cotton and wood
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celluloses. The
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between 0.1 and 400 s. The 13C T1 values were calculated based on the time-dependent signal
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intensities. The evolution of the signal intensity with τ was modeled using weighted linear
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least-squares fitting.
13
C T1 measurements of algal cellulose were performed with eight τ values
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RESULTS AND DISCUSSION
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TEMPO-Mediated Oxidation and Alkali Treatment of Algal Cellulose. Almost all the
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C6-hydroxy groups exposed on crystalline cellulose microfibril surfaces in native celluloses
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are position-selectively oxidized to Na C6-carboxylate groups by TEMPO-mediated oxidation
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under suitable conditions.22,27 As a result, one of every two glucosyl units on the cellulose
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microfibril surfaces are mostly converted into sodium glucuronosyl units by the
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TEMPO-mediated
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C6-carboxylate groups on the TEMPO-oxidized cellulose microfibril surfaces could thus be
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removed by alkali extraction. Native celluloses are stable without swelling and most glycoside
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bonds in cellulose molecules are not significantly cleaved with treatment using 1 M NaOH at
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105°C.38 Thus, the treatment with 1 M NaOH at 105°C for 12 h was used to remove as many
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of the C6-carboxylate-group-containing cellulose molecules as possible. This treatment was
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much harsher than that previously used to obtain glucose/glucuronic acid alternating
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co-polysaccharides from TEMPO-oxidized native celluloses by surface peeling with 10%
160
(w/w) NaOH at RT for 30 min.37 As a result, new cellulose microfibrils with one surface layer
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less than the original microfibrils were expected to be formed by the surface peeling (Figure 1).
oxidation.22,27,34‒37
The
surface
cellulose
molecules
containing
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Figure 1 Schematic model of layer-by-layer peeling of cellulose microfibril surface by
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TEMPO-mediated oxidation and subsequent alkali extraction.
170 0.6
171 172 173 174 175 176
Carboxylate content (mmol/g)
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0.5 0.4 0.3 0.2 0.1 0.0
Or i gi
na l
1s to
xi d i
1s ta ze d
2n 2n 3rd 3rd do da ox alk lka lka xi d i di ali li -e li -e ize ze -ex xtra d d xtra tra cte cte cte d d d
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Figure 2 Carboxylate contents of algal cellulose samples prepared by repetitions of
178
TEMPO-mediated oxidation and subsequent hot alkali extraction.
179 180
The carboxylate content of the 1st TEMPO-oxidized algal cellulose was 0.51 mmol/g, which
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is consistent with previously reported results.36 This carboxylate content decreased to 0.06
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mmol/g after the 1st hot alkali treatment. When this 1st TEMPO-oxidized and alkali-treated
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sample was re-oxidized using the TEMPO/NaBr/NaClO system, the resulting 2nd
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TEMPO-oxidized sample had a carboxylate content of 0.53 mmol/g. This value decreased to
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0.06 mmol/g after the 2nd hot alkali treatment. The carboxylate contents of the original,
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TEMPO-oxidized, and TEMPO-oxidized/alkali-treated algal cellulose samples up to three
187
repetitions are shown in Figure 2. Although complete removal of carboxylate groups in 8 ACS Paragon Plus Environment
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TEMPO-oxidized algal celluloses could not be achieved in each hot alkali treatment,
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approximately 90% of the carboxylate groups present in each TEMPO-oxidized algal cellulose
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sample were removed by the hot alkali treatment.
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XRD and Atomic Force Microscopy Analyses of TEMPO-Oxidized/Alkali-Treated
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Algal Celluloses. The cellulose microfibril widths of the original, TEMPO-oxidized, and
193
alkali-treated algal cellulose samples after the 1st, 2nd, and 3rd treatments shown in Figure 2
194
were examined using XRD and AFM. The XRD patterns of the algal cellulose samples are
195
presented in Figure S1 (see Supporting Information). The changes in the crystal sizes
196
corresponding to the diffraction peaks at 2θ = 14.8° and 16.8° for the algal cellulose samples
197
are shown in Figure 3. The cross sections of the algal cellulose microfibrils were assumed to be
198
rectangular or almost square,39–44 and the crystal sizes were calculated from the XRD patterns
199
based on the microfibril model shown in Figure S2 using Scherrer’s equation.29
200 20
201 202 203 204 205 206
Cellulose I crystal width (nm)
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10
5 Crystal width at 14.8 ° (1 1 0) Crystal width at 16.8 ° (1 -1 0)
0
Or i gi
na l
1s
to
xi d i
1s ta ze d
2n 2n 3rd 3rd da do alk ox lka xi d lka id i alili -e li -e i ze z e ex xtr d d xtra tra ac cte cte ted d d
207
Figure 3 Crystal widths of (1 1 0) and (1 ‒1 0) planes of cellulose I of algal cellulose samples
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prepared by repetitions of TEMPO-mediated oxidation and subsequent hot alkali extraction.
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The original crystal sizes of the (1 1 0) and (1 ‒1 0) planes were unchanged after the 1st
211
TEMPO-mediated oxidation; however, these sizes decreased by ~1 nm after the 1st hot alkali 9 ACS Paragon Plus Environment
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treatment. This value was unchanged after the 2nd TEMPO-mediated oxidation but decreased
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by ~1 nm after the 2nd hot alkali treatment. Similar results were observed for the 3rd
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TEMPO-oxidized and alkali-extracted algal cellulose samples (Figure 3). These XRD results
215
indicate that the crystal size of the algal cellulose decreased by ~1 nm after each round of
216
TEMPO-mediated oxidation and subsequent hot alkali extraction.
217 218 219 220 221 222
Cellulose nanofibril width measured from AFM height image (nm)
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15
10
5
0
Or igin al
1s 1s 2n 2n 3rd 3rd to ta do da ox alk xid lka lka xid idiz aliize li-e li-e zie ed ex d xtr d x tra tra ac cte c ted ted d
223
Figure 4 TEMPO-oxidized cellulose nanofibril widths measured from AFM height images of
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algal cellulose samples prepared by repetitions of TEMPO-mediated oxidation and subsequent
225
hot alkali extraction.
226 227
The TEMPO-oxidized algal celluloses prepared by the 1st, 2nd, and 3rd treatments shown in
228
Figures 2 and 3 were converted into individual nanofibrils dispersed in water using mechanical
229
disintegration.22,26,27 AFM examination was performed to measure the average heights of
230
individual TEMPO-oxidized cellulose nanofibrils (Figure S3). The average microfibril size of
231
the 1st TEMPO-oxidized algal cellulose nanofibrils was 14.2 ± 2.2 nm (Figure 4). This size
232
decreased to 13.3 ± 2.2 nm after the 1st hot alkali treatment and the 2nd TEMPO-mediated
233
oxidation. This size further decreased to 12.2 ± 3.0 nm with the 2nd hot alkali treatment and
234
subsequent TEMPO-mediated oxidation. These AFM observations of the TEMPO-oxidized
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algal cellulose nanofibrils indicate that the microfibril width decreased by ~1 nm with one
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sequential treatment of TEMPO-mediated oxidation/hot-alkali extraction, as also observed in
237
the XRD results presented in Figure 3.
238
The results in Figures 2‒4 show that the layer-by-layer peeling of the surface molecules of
239
algal cellulose microfibrils was successful with repetitions of the TEMPO-mediated oxidation
240
followed by hot alkaline extraction. This peeling was possible because the thicknesses of each
241
layer of a cellulose I crystallite corresponding to the (1 1 0) and (1 ‒1 0) planes are 0.53 and
242
0.61 nm, respectively, or ~0.5 nm (Figure S2).1,43 The alkali-soluble fractions obtained from
243
TEMPO-oxidized algal celluloses at each stage were highly degraded and depolymerized under
244
the alkaline conditions used.45
245
Similar results were obtained for highly crystalline tunicate cellulose, which has pure
246
cellulose Iβ and a parallelogram cross-section of microfibrils,46‒50 in layer-by-layer peeling
247
experiments (Figures S4–S6).
248
Solid-State 13C NMR Spectroscopy Analysis of TEMPO-Oxidized/Alkali-Treated Algal
249
Celluloses. The hydroxymethyl or C6-OH groups can have three conformations in cellulose
250
microfibrils: trans–gauche (tg), gauche–trans (gt), and gauche–gauche (gg) appearing at 60‒62,
251
62.5‒64.5, and 65.6‒66.6 ppm, respectively, in solid-state 13C NMR spectra.31,32,48. The C6-OH
252
signals of the original and 1st TEMPO-oxidized algal celluloses are presented in Figures 5A
253
and 5B, respectively. The signal for the C6-OH groups with the gg conformation appeared at
254
61.6‒61.7 ppm for the original algal cellulose, and its intensity decreased after the
255
TEMPO-mediated oxidation. The C6-OH groups present on the cellulose microfibril surfaces
256
were position-selectively converted into Na C6-carboxylate groups by the TEMPO-mediated
257
oxidation.22,26,27,34,35 Thus, the C6-OH groups present on the algal cellulose microfibril surfaces
258
likely had the gg conformation, which is consistent with a previously reported hypothesis.51
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Figure 5 Solid-state 13C NMR spectra of C6 signals of the original algal cellulose (A) and the
273
1st TEMPO-oxidized algal cellulose (B), with their deconvoluted patterns to tg, gt, and gg
274
conformations. The molar ratios of the tg, gt, and gg conformations of C6-OH groups of algal
275
cellulose samples prepared by repetitions of TEMPO-mediated oxidation and subsequent hot
276
alkali extraction (C) from solid-state 13C NMR spectra.
277 278
The carboxylate content of the 1st TEMPO-oxidized algal cellulose was 0.51 mmol/g
279
(Figure 2), which corresponds to 0.084 mol/mol of repeating units (i.e., glucosyl and
280
glucuronosyl units) according to a previously reported calculation (see Eq. S1 in Supporting
281
Information).36 The 8.4% glucosyl units in the original algal cellulose were converted into
282
C6-carboxylate units by the 1st TEMPO-mediated oxidation. As shown in Figure 5C, the
283
fraction of gg conformation in the original algal cellulose measured by solid-state
284
spectroscopy analysis was 6.9%, which is close to 8.4%. These results indicate that one of
13
C NMR
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every two C6-OH groups in the molecules present on each algal cellulose microfibril surface
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facing the exterior was likely to have the gg conformation.52‒54
287
Note that the gg:gt conformation ratios in the original algal celluloses as well as the 1st, 2nd,
288
and 3rd alkali-extracted algal cellulose samples were almost 1:1 (Figure 5C). This result
289
indicates that one of every two C6-OH groups in the molecules on each algal cellulose
290
microfibril surface facing toward the interior of the microfibril had the gt conformation (Figure
291
6). The glucosyl units with gg and gt conformations of C6-OH groups were, therefore,
292
alternatingly linked to each other in each cellulose molecule present on the algal microfibril
293
surface.37 Based on the cross-sectional model of the algal cellulose microfibril,1,36,39–44 16% of
294
the glucosyl units were present on the microfibril surfaces, and 84% were present inside the
295
microfibrils. Figure 5C shows that 86% of the glucosyl units in the 1st TEMPO-oxidized algal
296
cellulose had the tg conformation, which is consistent with the XRD data.
297 298 299 300 301 302 303 304
Figure 6 Schematic model of C6-OH conformations in the 1st, 2nd, and 3rd layers from the
305
algal cellulose microfibril surface..
306 307
The same interpretations were made for the 2nd and 3rd TEMPO-oxidized algal celluloses
308
and 2nd and 3rd alkali-extracted algal celluloses based on the layer-by-layer peeling model of 13 ACS Paragon Plus Environment
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309
the algal cellulose microfibrils by the TEMPO-oxidation/alkali-extraction process (Figures 5C,
310
S7, and S8). Thus, the cellulose molecules inside the algal cellulose microfibril surfaces always
311
had the tg conformation as cellulose I crystallites.1 A schematic model of the layer-by-layer
312
surface peeling of the algal cellulose microfibrils is presented in Figure 7.
313 314 315 316 317 318 319
Figure 7 Schematic model of the layer-by-layer peeling of algal cellulose microfibril surface
320
by repetitions of TEMPO-mediated oxidation and subsequent hot alkali extraction.
321 322
When the cellulose molecules originally present in the 2nd or 3rd layer from the microfibril
323
surface were exposed to the outermost surface by the layer-by-layer peeling, the gg and gt
324
conformations again appeared at the same 1:1 ratio (Figure 7). The signal patterns of C1
325
carbon at ~105 ppm in the solid-state
326
indicate that the ratio of cellulose Iα/Iβ was unchanged during the repetitions of the
327
TEMPO-oxidation/alkali extraction treatment (Figures S 7 and S8).32,55,56 Moreover, the gg, gt,
328
and tg conformation ratios for the original and alkali-extracted samples shown in Figure 5C
329
were almost unchanged between the wet and dry states (Figure S9).
330
13
C NMR spectra for these algal cellulose samples
Spin-Lattice Relaxation Times of Algal Cellulose Carbons. The
13
C T1 relaxation times
331
were measured using a pulse-sequence developed by Torchia.33 13C atoms with more freedom
332
of movement have shorter 13C T1 values, whereas those in rigidly ordered domains have longer 14 ACS Paragon Plus Environment
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333
13
C T1 values.57
13
334
including disordered regions. For the highly crystalline algal cellulose microfibril, the mobile
335
components should originate from the outermost microfibril surfaces, whereas the rigid
336
components should originate from inside the crystalline regions. As described in the previous
337
section, the C6-OH groups with the gg and gt conformations were located on the outermost
338
microfibril surfaces, whereas those with the tg conformation were located inside the crystalline
339
microfibrils (Figures 6 and 7).
C atoms with short T1 values are found on cellulose microfibril surfaces
340 341
Table 1. T1 Relaxation Times (ms) of C4 and C6 Carbons of Algal Cellulose.
C4
Never-dried
Freeze-dried
35
8.0
Freeze-dried and then re-wetted 25
1700
860
1600
gg conformation
0.4
2.0
0.4
tg conformation
940
430
1100
Surface Inside
C6 342 343
The T1 values of the C4 and C6 carbons in the original algal cellulose in the never-dried,
344
freeze-dried, and re-wetted states are listed in Table 1. These 13C T1 values clearly demonstrate
345
that the C4 and C6 carbons at the outermost surfaces of the microfibrils had much shorter T1
346
values than those inside the microfibril or crystalline regions for all three dry and wet states.
347
This result is reasonable because of the different molecular environments between the
348
microfibril surfaces and inside the crystallites. Moreover, the local mobility of C4 and C6
349
carbons (except for the C6 carbons with the gg conformation) was much more highly restricted
350
in the wet state than in the dry state. This difference in mobility can possibly be explained
351
using the following hypothesis: the hydration of algal cellulose microfibril surfaces or
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352
hydrogen bond formation between water molecules and C6-OH groups with the gg
353
conformation on the microfibril surfaces improve the rigidity of C4 carbons of both microfibril
354
surfaces and interiors and of C6 carbons inside the microfibrils. Because the C6-OH groups
355
with the gg conformation on the microfibril surface could form hydrogen bonds with water
356
molecules in the wet states, their
357
states. In contrast, there may not be opportunities for the C4 carbons on the microfibril surfaces
358
to form hydrogen bonds with water molecules even in the wet states, resulting in the surface
359
C4 carbons having higher T1 values in the wet states than in the dry state.
13
C T1 values were higher in the dry state than in the wet
360
Layer-by-Layer Peeling of Cotton and Wood Celluloses. It has been proposed that plant
361
cellulose microfibrils have more disordered structures on their surfaces than highly crystalline
362
algal cellulose.7‒12 The layer-by-layer peeling technique described above was thus applied to
363
cotton and wood celluloses. The carboxylate contents of the 1st and 2nd TEMPO-oxidized
364
cotton and wood celluloses and those of the 1st and 2nd hot alkali extractions are shown in
365
Figure 8A. Significant amounts of carboxylate groups were formed in the TEMPO-oxidized
366
celluloses, and most of them were removed by the hot alkali extraction, as for the algal and
367
tunicate celluloses. These results indicate that the layer-by-layer peeling of the
368
TEMPO-oxidized cellulose molecules in the plant cellulose microfibrils is likely to be
369
achieved by the repetition of the TEMPO-mediated oxidation and subsequent hot alkali
370
extraction process.
371
The crystal sizes of the original cotton and wood celluloses did not, however, decrease by ~1
372
nm after each hot alkali extraction, in contrast to the results for the algal and tunicate celluloses
373
(Figure 8B). These findings indicate that the plant cellulose microfibrils have disordered
374
structures not only in the 1st layer but also in part of the 2nd layer from the microfibril surface.
375
The solid-state
13
C NMR spectra of C6 carbons of these plant celluloses after 16 ACS Paragon Plus Environment
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376
TEMPO-mediated oxidation and hot alkali extraction were too broad in both the dry and wet
377
states for accurate signal deconvolution to separate the tg, gt, and gg conformations. Further
378
studies are thus needed to investigate the layer-by-layer peeling of these plant celluloses.
379 380 381 382 383
Carboxylate content (mmol/g)
2.0
384
A
1st oxidized 1st alkali-extracted 2nd oxidized 2nd alkali-extracted
1.5
1.0
0.5
0.0 Wood cellulose
385
B
386 387 388 389 390
Cotton lint cellulose
Cotton linters cellulose
20
Cellulose I crystal width (nm)
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
Biomacromolecules
Cotton linters Cotton lint Wood cellulose
15
10
5
0
Or
igi
na
l
1s to
391
xid iz
1s ed
ta
lka
2n li- e
do
xtr ac t ed
2n xid i
ze d
da
lka
li- e
xtr a
cte
d
392
Figure 8 Carboxylate contents of wood, cotton lint, and cotton linters cellulose samples
393
prepared by repetitions of TEMPO-mediated oxidation and subsequent hot-alkali extraction
394
(A) and corresponding changes in crystal width of the (2 0 0) plane of cellulose I (B).
395 396
CONCLUSION
397
The layer-by-layer peeling and solid-state 13C NMR analysis of algal cellulose revealed that the
398
C6-OH groups of surface cellulose molecules facing both outside and inside the microfibrils
399
have alternating gg and gt conformations, respectively, even after layer-by-layer peeling. In 17 ACS Paragon Plus Environment
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400
contrast, the C6-OH groups inside the microfibrils forming cellulose I crystallites always had
401
the tg conformation. The weight ratios of the three C6-OH conformations in the original algal
402
cellulose and those of the TEMPO-oxidized/hot alkali extracted samples remained almost
403
unchanged between the wet and dry states. The local mobility of entire cellulose molecules
404
except for C6 carbons with the gg conformation, evaluated by the
405
restricted in the wet state. Thus, the conformations of the C6-OH groups and their T1 values in
406
cellulose molecules were different for the microfibril surface and interior. When the
407
layer-by-layer peeling technique was applied to low-crystalline plant celluloses, no clear
408
decrease in the cellulose I crystal width was observed, indicating that the molecules in the 1st
409
and 2nd layers from the surface of these plant cellulose microfibrils have disordered structures.
410
The surface-peeling technique developed in this study can thus be used to perform structural
411
analyses of native cellulose microfibrils and to prepare new nanocelluloses with specific
412
widths.
13
C T1 values, was highly
413 414
ASSOCIATED CONTENT
415
Supporting Information
416
Calculation model for cellulose crystal sizes; XRD patterns, AFM images, and solid-state 13C
417
NMR spectra of layer-by-layer surface peeled algal celluloses; changes in carboxylate content
418
and microfibril width of layer-by-layer-surface peeled tunicate cellulose; molar ratios of tg, gt,
419
and gg conformations of C6-OH groups of surface-peeled algal celluloses in dry and wet states.
420 421
AUTHOR INFORMATION
422
Corresponding Author
423
*Tel: +81 3 5841 5538. E-mail:
[email protected] 18 ACS Paragon Plus Environment
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424 425
Notes
426
The authors declare no competing financial interest.
427 428
ACKNOWLEDGMENTS
429
This research was supported by Core Research for Evolutional Science and Technology
430
(CREST, Grant number JPMJCR13B2) of the Japan Science and Technology Agency (JST).
431 432
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573 574
Different Conformations of Surface Cellulose Molecules in Native
575
Cellulose Microfibrils Revealed by Layer-by-Layer Peeling
576 577
Ryunosuke Funahashi, Yusuke Okita, Hiromasa Hondo, Mengchen Zhao, Tsuguyuki Saito, and
578
Akira Isogai*
579 580 581 582 583 584 585 586
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