Exploring the Nature of Cellulose Microfibrils - Biomacromolecules

Mar 20, 2015 - Ultrathin cellulose microfibril fractions were extracted from spruce wood powder using combined delignification, TEMPO-catalyzed oxidat...
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Exploring the Nature of Cellulose Microfibrils Ying Su, Christian Burger, Hongyang Ma, Benjamin Chu,* and Benjamin S. Hsiao* Department of Chemistry, Stony Brook University, Stony Brook, New York 11794-3400, United States S Supporting Information *

ABSTRACT: Ultrathin cellulose microfibril fractions were extracted from spruce wood powder using combined delignification, TEMPO-catalyzed oxidation, and sonication processes. Small-angle X-ray scattering of these microfibril fractions in a “dilute” aqueous suspension (concentration 0.077 wt %) revealed that their shape was in the form of nanostrip with 4 nm width and only about 0.5 nm thicknesses. These dimensions were further confirmed by TEM and AFM measurements. The 0.5 nm thickness implied that the nanostrip could contain only a single layer of cellulose chains. At a higher concentration (0.15 wt %), SAXS analysis indicated that these nanostrips aggregated into a layered structure. The X-ray diffraction of samples collected at different preparation stages suggested that microfibrils were delaminated along the (11̅0) planes from the Iβ cellulose crystals. The degree of oxidation and solid-state 13C NMR characterizations indicated that, in addition to the surface molecules, some inner molecules of microfibrils were also oxidized, facilitating the delamination into cellulose nanostrips.

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INTRODUCTION Crystalline cellulose microfibrils, embedded in the matrix of polysaccharides (i.e., hemicelluloses and pectins),1 function as the structural support in cell walls of all green plants as well as in some types of algae. It has been shown that the lengths of microfibrils can reach a few microns, and the diameters can vary from 2 to 20 nm, depending on their origins.2,3 Each cellulose microfibril is composed of multiple linear (1,4)-linked β-Dglucan chains in an ordered arrangement, where the glucan chains are synthesized and extruded by the synthase complexes in the plasma membrane.1,4 However, it is still not clear as to where and how the glucan chains assemble into microfibrils, which has been one of the major challenges in understanding cellulose biosynthesis. Among the several notable hypotheses regarding the biosynthesis of cellulose microfibrils, the multistep assembly model based on molecular mechanical calculations is the most popular one.4−7 In this model, the nascent glucan chains first align with each other and form a monolayer by hydrophobic interactions in the terminal complex (TC). Subsequently, the monolayers pass through the rosette aperture and assemble into crystalline cellulose microfibrils via hydrogen bonding.4,6 Experimentally, the fine structure of cellulose microfibrils can be investigated by a wide range of characterization techniques. For example, in highly crystalline cellulose, such as algae cellulose8,9 and tunicate cellulose,10 the microfibril cross-section dimensions and the crystalline structure in the microfibril can be determined by the analysis of negative-staining cross-section images using transmission electron microscopy (TEM), as well as of lattice images using electron diffraction technique, respectively. However, in typical plants, since the cellulose microfibrils have much smaller cross-section sizes, and many © XXXX American Chemical Society

microfibrils tend to aggregate together, it is much more difficult to investigate the morphology of an individual microfibril without the need to extract them from the surrounding polysaccharide matrix first.3 Recently, it has been reported that individual cellulose microfibrils can be extracted from neverdried softwood pulp through a combined 2,2,6,6-tetramethylpiperidinyl-1-oxyl (TEMPO)-mediated oxidation and mechanical treatment.3,11 The results indicated that each microfibril exhibited a regular width of 3−4 nm,12 corresponding to about 5−7 chains parallel arranged along the width. With the assistance of atomic force microscopy (AFM), the fractions of cellulose microfibrils with thicknesses below 1 nm were observed by Renneckar et al., who suggested that the thin microfibril fractions were single layers of cellulose molecular chains resulted from the delamination of thicker cellulose microfibrils.13 Compared to the highly localized observations using microscopic techniques, small-angle X-ray/small-angle neutron scattering (SAXS/SANS) are powerful techniques to investigate the average dimensions of nanostructured cellulose. For example, the cellulose molecules extracted from switchglass were characterized by SANS, and the radius of gyration of these molecules was estimated from the Guinier plot.14 Tunicate cellulose whiskers were studied by both SAXS and SANS, and the lateral dimensions (8.8 × 18.2 nm) were determined from a parallelepiped model.15 Bacterial cellulose pellicles were characterized by SAXS, and it was found that the cross sections of bacterial cellulose microfibrils were of rectangular shape (1 × Received: December 29, 2014 Revised: March 16, 2015

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DOI: 10.1021/bm501897z Biomacromolecules XXXX, XXX, XXX−XXX

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final concentration of the suspension of cellulose nanostrips was 0.077 wt %, determined by the total organic carbon (TOC) analyzer. A higher concentration (0.15 wt %) suspension was prepared from the 0.077 wt % suspension by water evaporation on a hot plate with magnetic stirrer stirring at ∼250 rpm. For WAXD and NMR measurements, freeze-dried samples were prepared after each stage of sample treatment. To be specific, samples after the delignification step (WPDL), after the TEMPO-mediated oxidation step (WPTO), and after the sonication step (WPSN) were collected, respectively. During the preparation, the suspension was first cooled in liquid nitrogen for 30 min, and then immediately transferred to the freeze-drier (BT53, Millrock Technology Inc.) and kept at −40 °C for 5 days. Degree of Oxidation (DO). The ratio between the amount of oxidized and that of total hydroxymethyl groups in TEMPO-mediated cellulose nanostrips was determined through the conductometric titration method.20 Briefly, 0.1 mol/L of hydrochloric acid aqueous solution were added to 24 mL of cellulose nanostrips suspension with a concentration of 0.05 wt % until the pH reached 2.8. The suspension was titrated with 0.005 mol/L NaOH solution under stirring (450 rpm). The conductivity of the suspension was monitored using a conductivity meter throughout the titration process. The titration performance was terminated when the pH reached 10.7. The conductivity versus the volume of NaOH aqueous solution was plotted (Figure S1 in Supporting Information), where the NaOH volume used to neutralize the carboxyl groups, V, was determined from this curve. The amount of oxidized hydroxymethyl groups was calculated by multiplying V with the concentration, C, of the NaOH solution, while the amount of cellulose was directly measured from the TOC analyzer. Synchrotron SAXS Measurement of Cellulose Nanostrips Suspensions. The solution SAXS measurements (covering both SAXS and WAXD ranges) of cellulose nanostrips suspensions were carried out at the Beamline X9 of the National Synchrotron Light Source (NSLS), Brookhaven National Laboratory (BNL), U.S.A. Two suspension samples having the concentrations of 0.077 and 0.15 wt % were used for the study. In this measurement, 20 μL of suspension was pumped into a glass capillary (diameter of 1 mm) sealed across the vacuum path. The sample was allowed to flow continuously through the capillary during data collection in order to minimize the radiation damage.21 The chosen X-ray wavelength was 0.0918 nm. A PILATUS 300 K detector, located 3.2 m away from the sample, was used to collect the data covering the conventional SAXS angular range. A custom-designed Photonic Science CCD detector, 463 mm from the sample, was used to collect 2D data at the conventional WAXD range. For each sample, three 30-s scans were taken. The average of these three scans was used as the scattering pattern of the sample. A silver behenate standard was used to calibrate the parameters of the scattering geometry (i.e., beam center and sample-to-detector distance). Preliminary data processing was done with a Pythonbased package developed at the X9 Beamline to convert the 2D image into a linear scattering profile, block off dead pixels and pixels behind the beam-stop, merge SAXS and WAXD scans, and subtract the buffer and capillary background from the scattering profile. In this study, only the profile in small-angle region (s < 1 nm−1) was simulated, and the analysis of the wide-angle region was not involved here. The model fitting of the SAXS data was performed using the software Mathematica.22 Synchrotron WAXD Study of Dry Cellulose Samples. Three dried samples, including freeze-dried WPDL, freeze-dried WPTO, and freeze-dried WPSN samples, were measured using the WAXD technique at Beamline X27C of NSLS, BNL. The fluffy freeze-dried sample was carefully and uniformly spread onto a piece of Kapton tape to prevent any preferred orientation effect. The cellulose sample sandwiched between the Kapton tapes was loaded in a metal frame for the X-ray detection. The chosen X-ray wavelength was 0.1371 nm, the exposure time for each sample was 60 s, where 2D WAXD patterns were acquired using a MAR-CCD detector. The Al2O3 standard was used to calibrate the sample-to-detector distance. The diffraction data analysis, including background subtraction and conversion from 2D

16 nm).16 The sulfuric acid hydrolyzed cellulose from cotton, Avicel, and tunicate was also investigated by SAXS in suspension, and the dimensions were determined from the parallelepiped model simulation.17 In addition, bulk wood, pulp, and cellulose nanocrystals in nanocomposites could also be characterized by SAXS, but the interfibril interactions were necessary to be taken into consideration for the data interpretation.18,19 The goal of the present study was to explore the structure and morphology of exfoliated ultrathin cellulose microfibril fractions in aqueous suspension using the combined delignification, TEMPO-mediated oxidation, and successive sonication treatment to delaminate spruce wood cellulose microfibrils. This is because hydroxyl groups at the C6-position of the D-glucose unit can be region-selectively converted into carboxylate groups, where the electrostatic repulsion on the oxidized cellulose crystal surface would facilitate the delamination process and result in the formation of uniform dispersion of individual ultrathin microfibrillar sheets in “dilute” aqueous suspension. The shape and dimensions of these sheets (we termed “nanostrips” hereafter) in aqueous suspension at different concentrations were characterized by solution smallangle X-ray scattering (SAXS). In addition, wide-angle X-ray diffraction (WAXD), TEM, AFM, and solid 13C nuclear magnetic resonance (NMR) techniques were carried out to confirm and supplement the structural information obtained from the solution SAXS analysis. The experimental results are supportive of a possible delamination mechanism of cellulose microfibrils, based on the comparison of intermolecular interactions and an existing cellulose biosynthesis hypothesis.

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EXPERIMENTAL SECTION

In this study, the structure of products is closely correlated with the specific experimental preparation procedures chosen as follows. Materials and Delignification. The Jezo spruce (Picea jezoensis) wood powder (i.e., raw softwood sawdust) with particle size larger than 80-mesh (gift from Prof. Akira Isogai’s group, The University of Tokyo, Tokyo, Japan) was first soaked in 90% (v/v) acetone aqueous solution for 1 day under continuing stirring to remove extractives (e.g., triglycerides, resin acids, etc.). Subsequently, the treated wood powder was washed with acetone and separated by vacuum filtration. Delignification was carried out following the Wise method.11 Specifically, wood powder was repeatedly treated in 1% (w/v) NaClO2 buffer solution (with acetic acid added to keep pH around 5) and heated at temperatures between 60 and 70 °C for 1 h under stirring. The procedure was repeated for 8−12 times until the color of the wood powder became white. After soaking in HCl solution at pH 2 for 1 h to remove any metal ions, the wood powder was washed with deionized (DI) water 5 times and stored in a never-dried state for further use. TEMPO-Mediated Oxidation and Subsequent Mechanical Treatments. The never-dried delignified wood powder (1.0 g) was first suspended in water (90 mL), where sodium bromide (0.10 g) and the TEMPO agent (0.02 g) were subsequently added. After that, 1.5 mol/L of sodium hypochlorite solution (8 mmol per gram of cellulose) was added to initiate the oxidation process and the reaction was stirred mechanically in a sealed bottle for 24 h. The pH of the suspension was maintained between 9.8 and 10.3 during the reaction (monitored with a pH meter) by addition of sodium hydroxide aqueous solution (1 mol/L). The oxidized cellulose suspension was dialyzed in DI water until the pH reached 6.5. After the treatment, 10 mL of the oxidized cellulose suspension was diluted to 130 mL with DI water and disintegrated with a homogenizer (Cole Parmer, VCX400) at the output power of 80% (60 Hz, 155 W) in an ice bath for 10 min. After sonication, the suspension was centrifuged at 4700 g for 20 min and the supernatant was collected. With the above procedures, the B

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Biomacromolecules images to 1D profiles was performed using the software XPolar custom-developed at Stony Brook. TEM and AFM Measurements. For TEM measurements to determine the cellulose nanostrip width, samples were prepared by depositing cellulose nanostrips suspension (∼10 μL, 0.077 wt %) onto a carbon-film-coated copper grid (300 mesh). The suspension was allowed to stay on the grid for 3 min and then the excess liquid was drawn off the grid by using a filter paper. Before drying, 5 μL of 1.5 wt % phosphotungstic acid (PTA) aqueous solution was subsequently deposited on the grid and left for 30 s to stain the sample, where the excess liquid was absorbed by filter paper. This staining step was repeated once more and the stained sample was kept in air until completely dry and then stored in a desiccator before use. The TEM measurements were performed at the Center for Functional Nanomaterials (CFN) at BNL using a JEOL JEM-1400 microscope (JEOL, Tokyo, Japan) equipped with a CCD camera (ORIUS SC200, Gatan Inc., U.S.A.) operated at 100 kV. From the TEM images, the widths of the nanofibers were measured using the image processing software Leika. The statistical width information was obtained based on the measurements of 124 nanofibers on each TEM image. For AFM measurements, 0.0015 wt % of cellulose nanostrip suspension was spin-coated onto a freshly cleaved mica substrate. The measurement, in the tapping mode, was performed using a Bruker Dimension ICON scanning probe microscope (Bruker Corporation, U.S.A.) equipped with a Bruker OTESPA tip (radius 7 nm) at the Advanced Energy Research and Technology Center (AERTC), Stony Brook University. The thickness measurement was based on the line scans of a square image area with dimensions ranging from 1 to 3 μm. Solid-State CP-MASS 13C NMR Measurement. Solid-state 13C NMR spectra of four samples, i.e. original wood powder without any treatment (original), freeze-dried WPDL, freeze-dried WPTO, and freeze-dried WPSN samples, were obtained from Bruker 600 MHz wide-bore solid-state NMR spectrometer using cross-polarization magic angle sample spinning. The spinning speed was 12 kHz, pulse delay was 5 s and contact time was 1 ms. The peak assignments were made based on previous studies23−25 and the spectral fitting in the region between 40 and 200 ppm was performed based on the peak deconvolution method developed for cellulose samples.26

Figure 1. Experimental patterns of cellulose nanostrip suspensions at concentrations of 0.077 and 0.15 wt %, respectively, together with the fitting results by using the polydisperse ribbon model having an average thickness a = 0.48 nm with a standard deviation σa = 0.19 nm, and an average width b = 3.94 nm with a standard deviation σb = 0.5 nm. The inset sketch shows the cross-section shape of the ribbon model.

scattering interference peak implied that the increase in concentration led to an aggregation of nanostrips in suspension. We believe that the results extracted from the 0.077 wt % scattering curve: size-weighted average width b = 3.94 nm with a standard distribution σb = 0.5 nm, and size-weighted average thickness a = 0.48 nm with a standard deviation σa = 0.19 nm, should represent a good estimate of the dimensions for the building blocks of cellulose microfibril−nanostrips, which were delaminated from microfibrils and finely dispersed in the “dilute” suspension. Nevertheless, it is also possible that there existed small amounts of microfibrils that were not completely delaminated or nanostrip aggregates due to the attractive hydrogen bonding, but the content of such “thicker” fibrils should be below 15% based on the thickness distribution of the polydisperse ribbon model. To verify these two dimensions (width and thickness), TEM and AFM techniques were used to examine the solid samples evaporated from the suspension. Figure 2a,b shows the typical TEM images of cellulose nanostrips prepared from the 0.077 wt % suspension at different magnifications; Figure 2c illustrates the projected microfibrillar widths measured from the images (ranging from 3.25 to 5.25 nm). This size range was consistent with the width range of 3.4 to 4.4 nm, derived from the SAXS analysis of the 0.077 wt % suspension, as well as the reported widths of spruce cellulose microfibrils in previous literatures.12,28 We believe that the SAXS analysis has yielded the more precise information because the TEM samples were prepared with evaporation of the nanostrip suspension, whereas cellulose nanostrips could aggregate into thicker stacks. In addition, the edges of cellulose nanostrips on TEM images were quite blurry due to the resolution limit and the negative dying might further cause artifacts affecting the measurement of the nanostrip width. Generally, the TEM measurement might lead to a larger width estimate. The size-weighted average thickness a = 0.48 nm having a standard distribution σa = 0.19 nm, determined from the SAXS characterization, indicated that the cellulose nanostrips consist of a single layer of cellulose chains. These results were confirmed by the AFM measurements of solid samples

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RESULTS AND DISCUSSION Dimensions of Cellulose Nanostrips. From our previous study,27 we demonstrated that a simplified polydisperse ribbon model could efficiently describe the SAXS intensity profiles of cellulose nanofibers in suspension, extracted from dried wood pulp using the TEMPO-oxidation method. Based on the model fitting, the structural information, including cross-section shape (ribbon like), size, and size distribution of cellulose nanofibers, could be obtained, while the fiber length was assumed to be infinitely long. This is because the fiber length was in microns, exceeding the detectable size range by SAXS. For cellulose nanostrip suspensions (extracted from neverdried softwood pulp) having concentrations of 0.077 and 0.15 wt %, the corresponding SAXS intensity profiles were also fitted with the polydisperse ribbon model, where the results are illustrated in Figure 1. It was found that this model could fit the 0.077 wt % curve with high confidence (the coefficient of determination R2 = 0.9987) yielding the size-weighted average width b = 3.94 nm with a standard distribution of width σb = 0.5 nm, size-weighted average thickness a = 0.48 nm with a standard deviation of thickness σa = 0.19 nm. However, this model could not fit the 0.15 wt % curve with any satisfaction. For example, the discrepancy between the model fit using the same set of parameters and the experimental curve was notable, especially near the region s = 0.3 nm−1, where the presence of a scattering shoulder on the experimental curve was seen. This C

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Figure 2. (a, b) TEM images of cellulose nanostrips at different magnifications. (c) Width distribution of nanostrips measured from TEM images based on a count of 124 widths at different positions from five images.

distributions of a single layer of cellulose molecules in reality (considering that the hydrogen atoms distribute to the external layer of a molecular chain compared to the carbon and oxygen atoms). The expression of the monodisperse nanostrip model is shown in eq 1, and the derivation of this model from the monodisperse parallelepiped model is illustrated in the Supporting Information.

prepared by using a very dilute (0.0015 wt %) suspension of cellulose nanostrips. Figure 3 illustrates two representative AFM images of cellulose nanostrips spin-coated onto a freshly cleaved mica substrate. It was seen that the smallest thickness value was around 0.5 nm, indicating a single nanostrip, where thickness values larger than 1 nm were also seen, indicating the presence of thicker microfibrils or aggregation of nanostrips. In addition, the bending and twisting of nanostrips on the mica substrate could also result in the apparent value of a larger thickness. For the 0.15 wt % sample, due to the appearance of the small scattering shoulder at s = 0.3 nm−1 in the SAXS intensity profile, it was hypothesized that nanostrips exhibited the tendency to aggregate, due to attractive hydrogen bonding between the nanostrips, at a higher concentration, thereby forming a loose stacking structure. Based on this hypothesis, a “1D paracrystalline” model was developed to fit the 0.15 wt % intensity curve. To achieve this, the “polydisperse ribbon” model for extracting the nanostrip dimensions (i.e., the sizeweighted average width b = 3.94 nm with a standard distribution σb = 0.5 nm, and size-weighted average thickness a = 0.48 nm with a standard deviation σa = 0.19 nm) was first simplified into a “monodisperse nanostrip” model with rectangular cross sections. To be specific, in the monodisperse nanostrip model, the width of the cross-section was assumed to be uniform with no size distribution. Meanwhile, the thickness was also assumed to be monodisperse, but it was composed of a Gaussian distribution of electron density across the thickness. Although this assumption was made mainly for the efficient calculation purpose, it also simulated the electron density

I (s ) = f

⎛1 3 ⎞ 1 exp( − 2πs 2d 2)1F2⎜ ; , 2; −π 2s 2b2⎟ + IB ⎝2 2 ⎠ s (1)

In eq 1, I(s) is the scattered intensity, s is the modulus of scattering vector (s = 2 sin θ/λ), b is the width of the crosssection of the nanostrip, d is the integral width of the Gaussian distribution of the electron density across the thickness, IB is the constant background intensity, f is the scaling factor of the intensity, and 1F2(1/2; 3/2, 2; −π2s2b2) is a hypergeometric function. To demonstrate the validity of this nanostrip model, eq 1 was used to fit the 0.077 wt % scattering curve using nonlinear leastsquares fitting, and the results are shown in Figure 4. It was seen that by using the width b = 3.93 nm and the integral width of the Gaussian distribution of the electron density across the thickness d = 0.40 nm, the nanostrip model could fit the experimental scattering curve with excellent confidence (coefficients of determination R2 = 0.9996). In fact, this fitting result indicated that compared to the polydisperse ribbon model yielding the coefficient of determination R2 = 0.9987, the simpler monodisperse nanostrip model could even be better or at least equally fit the scattered intensity of nanostrip D

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Figure 3. AFM height images of cellulose nanostrips (top two diagrams). The height profiles of the fibers marked on the images are shown below the images.

width at half-maximum (fwhm; i.e., 0.38 nm) or the 3-sigma width (i.e., 0.48 nm) of the Gaussian distribution to represent the thickness of the nanostrips. Therefore, the difference between d = 0.4 nm (the integral width of the Gaussian distribution of the electron density across the thickness in the nanostrip model) and a = 0.48 nm (the average thickness in the polydisperse ribbon model) represents the difference between the two width definitions. As discussed earlier, the appearance of the small scattering shoulder at s = 0.3 nm−1 in the 0.15 wt % intensity curve was probably due to the aggregation of nanostrips from hydrogen bonding. Using the nanostrip as the building block, a “1D paracrystalline” model was developed to simulate the aggregation of cellulose nanostrips in aqueous suspension at higher concentrations. Specifically, we assumed that the cellulose nanostrips with thickness (d′) of 0.5 nm (this is close to d = 0.4 nm, the integral width of the Gaussian distribution of the electron density across the thickness in the nanostrip model) stacked up with their large surfaces facing each other. The distances between two adjacent nanostrips followed a Gaussian distribution, and the numbers of nanostrips in each stack followed an exponential distribution. The expression of the “1D paracrystalline” model can be shown as follows:

Figure 4. Experimental pattern for cellulose nanostrip suspensions at concentration 0.077 wt % and the fitting result of the nanostrip model with width b = 3.93 nm and integral width of the thickness d = 0.40 nm. The inset sketch shows the cross-section dimensions of the nanostrip model.

suspension at low concentrations. As a result, the dimensions of the nanostrip extracted by the nanostrip model were used as the building units to describe the stacking structure at higher concentrations. It is worth pointing out that we used the integral width of the Gaussian distribution to represent the thickness in the nanostrip model, but one could also use the full E

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Biomacromolecules exp(− 2πd′2 s 2) I(s) = f s

∫0

π /2

⎛ ⎜ 3b′2 ⎜−1 3 + π 2b′2 s 2 sin 2 ψ ⎜ ⎝

⎡ ⎢ 1 + 2Re⎢ ⎢ 1 − exp − 2πia0s cos ψ − 2π 2σa2s 2 cos2 ψ − ⎣

(

2 n0

)

model suggested we included some more diffused interfaces between nanostrips and their surrounding matrix in the stacking structure. Molecular Structure in Cellulose Microfibrils. Based on the above finding that cellulose microfibrils could be delaminated into monomolecular sheets or nanostrips, now the question is when and how the delamination can happen. To explore the possible mechanisms for the delamination process, we investigated the structure and functionality of the samples collected at different stages of sample preparation, including delignification, TEMPO-mediated oxidation, and sonication. The WAXD technique was used to track the changes of the crystalline structure in these samples, where the results are illustrated in Figure 6. From the comparison of different

⎤⎞ ⎥⎟ ⎥⎟dψ ⎥⎟ ⎦⎠

(2)

where f is the scaling factor, b′ is the width of each nanostrips, d′ is the corresponding thickness, a0 is the average interlayer distance, σa is the standard deviation of the interlayer distance, n0 is the average number of nanostrips in each stack following an exponential distribution, IB is the background constant, and ψ is the azimuthal angle. The detailed derivation of the “1D paracrystalline” model is illustrated in the Supporting Information. Figure 5 illustrates the fitting results of the 0.15 wt % scattering curve using the “1D paracrystalline” model, where

Figure 6. Comparison of WAXD patterns from samples collected after the delignification step (WPDL), after the TEMPO-mediated oxidation step (WPTO), and after the sonication step (WPSN).

diffraction patterns, it was seen that the (11̅0) reflection peak intensity decreased significantly after the TEMPO-oxidation step, and it almost disappeared after the sonication step. This observation implies that the cellulose crystallites were delaminated or exfoliated along the (11̅0) planes, where delamination started during the prolonged oxidation process, and the sonication step further intensified and completed the delamination process. For the WPSN sample (after the sonication step), both (110) and (200) reflections shifted toward the amorphous peak position (s ∼ 2.1 nm−1), suggesting the degradation of the crystalline structure. This could be explained by the hypothesis that, as the microfibril delaminated into nanostrips, a large amount of the inner crystalline chains became surface chains with higher disordering, leading to the shift of all major diffraction peaks toward the amorphous peak position. For native cellulose materials obtained from certain biological sources (e.g., tunicate or Valonia), the cellulose microfibrils usually show planar orientation before fibrillation, which would influence the relative reflection intensities of the diffraction patterns.29,30 However, for spruce cellulose, it has been known that the cellulose microfibrils only has uniaxial orientation in the cell wall before fibrillation, where no planar orientation can be obtained.30 In this case, the intensity decrease in the (11̅0) reflection after the oxidation and mechanical fibrillation treatments should be attributed to the

Figure 5. Experimental pattern for cellulose nanostrip suspensions at concentration 0.15 wt % and the fitting result of the “1D paracrystalline” model using width b′ = 3.93 nm, integral width of the electron density distribution across the thickness of each nanostrip d′ = 0.5 nm, average distance between adjacent nanostrips a0 = 1.4 nm, standard deviation of the interlayer distance σa = 0.5 nm, and average number of nanostrips in each stack n0 = 1.6. The sketch in the upper right-hand corner shows the layered structure of the “1D paracrystalline” model; the inset in the bottom left-hand corner shows the exponential distribution curve of the number of layers in each stack.

the following parameters were obtained: b′ = 3.93 nm, d′ = 0.5 nm, a0 = 1.4 nm, σa = 0.5 nm, and n0 = 1.6. These results suggested that in average, there were 1.6 nanostrips in each stack, and each nanostrip was of 3.93 nm wide and 0.5 nm thick. The average distance between two adjacent nanostrips was 1.4 nm with standard deviation 0.5 nm. It should be noted that, n0 is statistically a decimal because it is the average number of the continuous exponential distribution [f(n) = (1/n0) exp(−n/n0)] of the number of nanostrips n in each stack. The probability density curve (inset of Figure 5) indicated that, with n0 = 1.6, the aggregates could be composed of more than two nanostrips, but single nanostrips still dominated the suspension system. Also, compared to the d = 0.4 nm in the nanostrip model, the selection of d′ = 0.5 nm in the “1D paracrystalline” F

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ppm33 in the spectrum of the original wood powder disappeared completely after the delignification treatment. Meanwhile, some other polysaccharide components (e.g., hemicellulose and pectin) were also dissolved during the delignification process, which was evidenced by the signal decrease at peak 62 ppm (pectin)34 and the region of 81−82 ppm (hemicellulose)35 in the spectrum of WPDL. Comparing the spectra between WPDL and WPTO, the peak assigned to the carboxylate group (COONa) at 175 ppm appeared after the oxidation reaction and the intensity of the peak at 60−69 ppm corresponding to the C6 position decreased gradually. Furthermore, the intensity of both the noncrystalline C6 (∼63 ppm) and crystalline C6 (∼66 ppm) signals decreased after oxidation, with the crystalline C6 peak area changing from 12.7% to 7.3% and the noncrystalline C6 peak area changing from 4.6% to 3.4%, respectively. This data indicated that not only the surface cellulose chains, but also some of the inner chains, were oxidized during the chosen oxidation process. The total peak area at 81−92 ppm, assigned to the C4 position remained unchanged (∼12%), however, about 10% of the crystalline C4 signal (at 87−92 ppm) turned into the noncrystalline C4 signal (at 81−87 ppm) based on the difference of the peak areas before and after oxidation. Moreover, the decrease in the C1 peak area from 16.5% to 14.3%, and the appearance of a shoulder at 94 ppm could be attributed to the formation of C1 reducing ends after the break of the (1,4) glycosidic bonds,36 indicating that besides oxidation, certain cellulose chains were hydrolyzed during the oxidation process. For the WPSN sample, the broadening of all the peaks in the region of 50−110 ppm was seen, indicating that the system became more disordered after the sonication step. Specifically, the area of the crystalline C4 peak (87−92 ppm) decreased from 4.4% to 0.7% after sonication, and the area of the crystalline C6 peak (∼66 ppm) decreased from 3.4% to 1.0%. The extended decrease of the crystalline C4 and C6 signals suggested that delamination of cellulose microfibrils occurred thoroughly during the sonication processing. The degree of oxidation (DO) of cellulose nanostrips in a “dilute” suspension (0.05 wt %), determined from the conductometric titration was 0.50 (Figure S1), indicating that about 50% of hydroxymethyl groups on cellulose nanostrips were oxidized into carboxylate groups during TEMPOmediated oxidation. It is well-known that the interplane distances of (11̅0) and (110) for Iβ cellulose are 0.61 and 0.54 nm, respectively.8,37 Assuming the (11̅0) and (110) crystal planes are the exposed surfaces of the cellulose microfibrils9 and assuming the cross-section of softwood cellulose microfibril is of near-square shape with the width of 4.1 nm and thickness of 3.5 nm,28 there would be 4.1/0.54 ≈ 7 chains in the (11̅0) plane and 3.5/0.61 ≈ 6 chains in the (110) plane, respectively, where 22 out of the total 42 chains are on the surface of the microfibril. Assuming only the hydroxymethyl groups exposed on the surface could be oxidized, the maximum DO would be (20 × 0.5 + 2)/42 = 0.29. However, the actual DO = 0.50 was much higher than this maximum value, suggesting that besides the exposed hydroxymethyl groups, some “inner” hydroxymethyl groups also participated in the oxidation. Discussion of the Delamination Mechanism. It is wellknown that the (200) plane of cellulose Iβ crystal structure is held together by intrasheet hydrogen bonds, whereas the hydrophobic interactions and weak intersheet hydrogen bonds are the main binding forces in the (11̅0) plane.37 For some

change of the crystalline structure in microfibril rather than due to the change in orientation. This conclusion was further verified by the freeze-dry treatment of the cellulose suspension sample, resulting in a three-dimensional structure with a random distribution of cellulose microfibrils that prevented the effect of orientation on the diffraction intensity change. The WAXD results suggested that delamination occurred along the (11̅0) plane, which is in contrast to the proposed delamination along the (200) plane of the Iβ crystalline structure in the study by Renneckar et al.13 Such a discrepancy may be due to the following reasons. (1) Cellulose from different biological sources may have different microfibrillar dimensions and crystalline properties.3 Certainly, this can lead to different delamination process. But this hypothesis needs to be verified. (2) The different delignification process may have different effect on the resulting cellulose microfibrillar structure. In other words, as lignin and hemicellulose molecules play the role of binding and tethering cellulose microfibrils together in the plant cell wall, it is likely that some portions of hemicellulose molecules are entrapped in the cellulose crystalline domains.1,31 The different dissolution pathway to remove lignin and hemicellulose may result in a different structure of cellulose microfibrils.32 (3) As cellulose microfibrils are highly anisotropic, it is conceivable that a preferred orientation can be created during sample preparation, especially by compressing the anisotropic material into a pallet form. In this study, we have carefully minimized the preferred orientation of the sample. Otherwise, the sample anisotropy can notably affect the corresponding reflection intensities. The functionality of the four solid samples (i.e., original spruce wood powder, WPDL, WPTO, and WPSN) was determined by using the solid-state 13C NMR technique, where the corresponding spectra are shown in Figure 7. It was seen that the lignin signals at 56 ppm and between 110 and 155

Figure 7. Solid-state CP-MASS 13C NMR spectra of original spruce wood powder, WPDL, WPTO, and WPSN. For the peak assignment, “I” denotes inner/crystalline signal, “S” denotes surface/noncrystalline signal. C1′ represents the reducing end of cellulose chain. G

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of these sheets could take place at higher concentrations, as observed by the SAXS results of a more concentrated suspension.

time, it was believed that the intrasheet hydrogen bonds were the strongest attracting force in the cellulose crystalline entity13 and were often attributed to the insolubility of cellulose crystals.38,39 However, recent molecular dynamics simulation suggested that the hydrophobic pairing energy between cellulose oligomer chains were much higher than the intermolecular hydrogen bonds,40,41 which would favor the delamination of cellulose microfibrils along the (11̅0) planes especially under the influence of external forces. Since the cellulose chains on the surface of microfibrils are very disordered, their C6 hydroxyl groups, when exposed to the oxidant, can be readily oxidized into carboxylate groups with negative charges (Figure 8). As a result, electrostatic repulsive interactions between two opposing (11̅0) planes could be considered as external forces that would facilitate the delamination process.

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CONCLUSIONS Monolayer cellulose nanostrips could be produced by the combined delignification, oxidation and sonication treatment of never-dried spruce wood powder using specific sample preparation procedures. The cellulose nanostrips were found to have a width of ∼4 nm and a thickness of ∼0.5 nm. These results were obtained by the SAXS analysis of delaminated cellulose microfibrils in aqueous suspension at a relatively “low” concentration (0.077 wt %). At higher concentrations (e.g., 0.15 wt %), these nanostrips exhibited the tendency to stack up and form a multilayered aggregate. The TEM images of samples cast from the “dilute” suspension (0.077 wt %) verified the width range of cellulose nanostrips determined by SAXS, and the AFM measurement of samples cast from a very dilute (0.0015 wt %) suspension confirmed the thickness of nanostrips determined by SAXS. The WAXD analysis of the samples collected at different stages of preparation indicated that the delamination of microfibril occurred along the (11̅0) plane in Iβ cellulose, leading to the formation of cellulose nanostrips. As the negative charges were introduced into the cellulose crystalline entity, the electrostatic repulsion between the (11̅0) planes might be the driving force for delamination. The degree of oxidation measurement indicated that the DO value of cellulose nanostrips was higher than the projected DO value if one assumes that only the surface cellulose chains are oxidized. Finally, the solid-state 13C NMR analysis revealed that some inner cellulose chains within the microfibrils were also oxidized during the oxidation process, while delamination of the microfibril mainly occurred during the intensive sonication step. It should be noted that our structural results were obtained based on the analysis of samples prepared using the procedures described in the Experimental Section. Without question, the sample preparation scheme can play an important role in influencing the structure of the final product.

Figure 8. Schematic representation of the oxidation process in the cellulose microfibril. Red circles represent the oxidized hydroxymethyl groups both on the surface and partially inside the microfibril.

In the perspective of cellulose biosynthesis, there can be another reason to explain the proposed delamination mechanism of cellulose microfibrils. As hypothesized in previous literatures,4−7 during biosynthesis in the plant cell wall, the assembly of cellulose molecular chains requires multiple steps for cellulose crystallization to take place. It is generally thought that the hydrophobic interaction-associated (11̅0) planes form inside the terminal complex TC first. Only after these planes are extruded from the TC, such monolayer 2D sheets can stack into the crystalline microfibril assembly through hydrogen bonding. During the assembly of the cellulose sheets, some hemicellulose chains can be entrapped between the sheets. It is conceivable that during the delignification and TEMPO-mediated oxidation steps, the hemicellulose chains, entrapped between the (11̅0) planes, can be degraded and dissolved by the reaction system,42,43 creating molecular cracks/channels immediately occupied by water. These channels would provide pathways for oxidant molecules to penetrate into the microfibril, resulting in the partial oxidation of inner cellulose molecules, as observed by the NMR spectra. The monomolecular cellulose sheets (i.e., cellulose nanostrips), resulted from the delamination of microfibrils, could disperse homogeneously in water at low concentrations because of the negative charges induced by the TEMPO-mediated oxidation. However, since the oxidation occurred mainly on the surface of cellulose microfibril and only partially inside of the microfibril, the electrostatic repulsion among the monomolecular sheets might be relatively weak, where the aggregation

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ASSOCIATED CONTENT

S Supporting Information *

The mathematical derivations of the nanostrip model and the “1D paracrystalline” model, the model fitting of SAXS pattern of low concentration (0.077 wt %) suspension of cellulose microfibrils using the “1D paracrystalline” model, as well as the conductometric titration curve are included. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Authors

*E-mail: [email protected]. Tel.: (+631) 6327793. Fax: (+631) 632-6518. *E-mail: [email protected]. Tel.: (+631) 6327928. Fax: (+631) 632-6518. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The financial support for this work was provided by a grant from the SusChEM Program of the National Science Foundation (DMR-1409507). The authors thank Prof. Akira Isogai and Prof. Tsuguyuki Saito at The University of Tokyo H

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(31) Hayashi, T. Annu. Rev. Plant Phys. 1989, 40, 139−168. (32) Iwamoto, S.; Abe, K.; Yano, H. Biomacromolecules 2008, 9, 1022−1026. (33) Maciel, G. E.; Odonnell, D. J.; Ackerman, J. J. H.; Hawkins, B. H.; Bartuska, V. J. Makromol. Chem. 1981, 182, 2297−2304. (34) Ha, M. A.; Apperley, D. C.; Jarvis, M. C. Plant Physiol. 1997, 115, 593−598. (35) Hult, E. L.; Larsson, P. T.; Iversen, T. Cellulose 2000, 7, 35−55. (36) Whitney, S. E. C.; Brigham, J. E.; Darke, A. H.; Reid, J. S. G.; Gidley, M. J. Plant J. 1995, 8, 491−504. (37) Nishiyama, Y.; Langan, P.; Chanzy, H. J. Am. Chem. Soc. 2002, 124, 9074−9082. (38) Bodvik, R.; Dedinaite, A.; Karlson, L.; Bergstrom, M.; Baverback, P.; Pedersen, J. S.; Edwards, K.; Karlsson, G.; Varga, I.; Claesson, P. M. Colloids Surf., A 2010, 354, 162−171. (39) Zhang, L. N.; Ruan, D.; Gao, S. J. J. Polym. Sci., Part B: Polym. Phys. 2002, 40, 1521−1529. (40) Bergenstrahle, M.; Wohlert, J.; Himmel, M. E.; Brady, J. W. Carbohydr. Res. 2010, 345, 2060−2066. (41) Medronho, B.; Romano, A.; Miguel, M. G.; Stigsson, L.; Lindman, B. Cellulose 2012, 19, 581−587. (42) Kumar, R.; Hu, F.; Hubbell, C. A.; Ragauskas, A. J.; Wyman, C. E. Bioresour. Technol. 2013, 130, 372−381. (43) Okita, Y.; Saito, T.; Isogai, A. Holzforschung 2009, 63, 529−535.

for providing the raw materials and helpful discussions. Special thanks are due to Dr. Henri Chanzy for his enlightening suggestions and discussions. The authors also thank Drs. Lin Yang and Vito Graziano for their assistance at Beamline X9, Dr. Maya K. Endoh for her assistance at Beamline X27C, and Mr. Kim Kisslinger at CFN for his assistance with the TEM measurements. In addition, the authors gratefully acknowledge Dr. Tong Wang of the Biology Department at BNL for the TEM sample preparation, Dr. Chung-Chueh Chang at AERTC for his assistance with the AFM measurement, and Dr. Martine Ziliox at the Center for Structural Biology, Stony Brook University, for her assistance with the NMR characterization.

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REFERENCES

(1) Cosgrove, D. J. Nat. Rev. Mol. Cell Biol. 2005, 6, 850−861. (2) Saito, T.; Nishiyama, Y.; Putaux, J. L.; Vignon, M.; Isogai, A. Biomacromolecules 2006, 7, 1687−1691. (3) Nishiyama, Y. J. Wood Sci. 2009, 55, 241−249. (4) Saxena, I. M.; Brown, R. M. Ann. Bot. (London, U.K.) 2005, 96, 9−21. (5) Moon, R. J.; Martini, A.; Nairn, J.; Simonsen, J.; Youngblood, J. Chem. Soc. Rev. 2011, 40, 3941−3994. (6) Cousins, S. K.; Brown, R. M. Polymer 1995, 36, 3885−3888. (7) Delmer, D. P. Annu. Rev. Plant Phys. 1999, 50, 245−276. (8) Sugiyama, J.; Vuong, R.; Chanzy, H. Macromolecules 1991, 24, 4168−4175. (9) Sugiyama, J.; Harada, H.; Fujiyoshi, Y.; Uyeda, N. Planta 1985, 166, 161−168. (10) Helbert, W.; Nishiyama, Y.; Okano, T.; Sugiyama, J. J. Struct. Biol. 1998, 124, 42−50. (11) Saito, T.; Kuramae, R.; Wohlert, J.; Berglund, L. A.; Isogai, A. Biomacromolecules 2013, 14, 248−253. (12) Isogai, A.; Saito, T.; Fukuzumi, H. Nanoscale 2011, 3, 71−85. (13) Li, Q. Q.; Renneckar, S. Biomacromolecules 2011, 12, 650−659. (14) Pingali, S. V.; Urban, V. S.; Heller, W. T.; McGaughey, J.; O’Neill, H. M.; Foston, M.; Myles, D. A.; Ragauskas, A. J.; Evans, B. R. Acta Crystallogr., Sect. D 2010, 66, 1189−1193. (15) Terech, P.; Chazeau, L.; Cavaille, J. Y. Macromolecules 1999, 32, 1872−1875. (16) Astley, O. M.; Chanliaud, E.; Donald, A. M.; Gidley, M. J. Int. J. Biol. Macromol. 2001, 29, 193−202. (17) Elazzouzi-Hafraoui, S.; Nishiyama, Y.; Putaux, J. L.; Heux, L.; Dubreuil, F.; Rochas, C. Biomacromolecules 2008, 9, 57−65. (18) Leppanen, K.; Andersson, S.; Torkkeli, M.; Knaapila, M.; Kotelnikova, N.; Serimaa, R. Cellulose 2009, 16, 999−1015. (19) Oliveira Patricio, P. S.; Miranda, I. P.; Silva, N. C. F.; Ayres, E.; Pereira, F. V.; Orefice, R. L. Eur. Polym. J. 2013, 49, 3761−3769. (20) Habibi, Y.; Chanzy, H.; Vignon, M. R. Cellulose 2006, 13, 679− 687. (21) Allaire, M.; Yang, L. J. Synchrotron. Radiat. 2011, 18, 41−44. (22) Wolfram Research, Inc. Mathematica, Version 8.0; Wolfram Research, Inc.: Champaign, IL, 2010. (23) Saito, T.; Shibata, I.; Isogai, A.; Suguri, N.; Sumikawa, N. Carbohydr. Polym. 2005, 61, 414−419. (24) Wickholm, K.; Larsson, P. T.; Iversen, T. Carbohydr. Res. 1998, 312, 123−129. (25) Vietor, R. J.; Newman, R. H.; Ha, M. A.; Apperley, D. C.; Jarvis, M. C. Plant J. 2002, 30, 721−731. (26) Kato, Y.; Matsuo, R.; Isogai, A. Carbohydr. Polym. 2003, 51, 69− 75. (27) Su, Y.; Burger, C.; Hsiao, B. S.; Chu, B. J. Appl. Crystallogr. 2014, 47, 788−798. (28) Okita, Y.; Saito, T.; Isogai, A. Biomacromolecules 2010, 11, 1696−1700. (29) Sugiyama, J.; Chanzy, H.; Revol, J. F. Planta 1994, 193, 260− 265. (30) Horikawa, Y.; Itoh, T.; Sugiyama, J. Cellulose 2006, 13, 309− 316. I

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