A new family of materials comprised of cellulose, cellulose nanomaterials (CNMs), having properties and functionalities distinct from molecular cellulose and wood pulp, is being developed for applications that were once thought impossible for cellulosic materials. Commercialization, paralleled by research in this field, is fueled by the unique combination of characteristics, such as high on-axis stiffness, sustainability, scalability, and mechanical reinforcement of a wide variety of materials, leading to their utility across a broad spectrum of high-performance material applications. However, with this exponential growth in interest/activity, the development of measurement protocols necessary for consistent, reliable and accurate materials characterization has been outpaced. These protocols, developed in the broader research community, are critical for the advancement in understanding, process optimization, and utilization of CNMs in materials development. This review establishes detailed best practices, methods and techniques for characterizing CNM particle morphology, surface chemistry, surface charge, purity, crystallinity, rheological properties, mechanical properties, and toxicity for two distinct forms of CNMs: cellulose nanocrystals and cellulose nanofibrils.

Deconstruction of lignocellulosic plant cell walls to fermentable sugars by thermochemical and/or biological means is impeded by several poorly understood ultrastructural and chemical barriers. A promising thermochemical pretreatment called ammonia fiber expansion (AFEX) overcomes the native recalcitrance of cell walls through subtle morphological and physicochemical changes that enhance cellulase accessibility without extracting lignin and hemicelluloses into separate liquid streams. Multiscale visualization and characterization of Zea mays (i.e., corn stover) cell walls were carried out by laser scanning confocal fluorescence microscopy (LSCM), Raman spectroscopy, atomic force microscopy (AFM), electron microscopy (SEM, TEM), nuclear magnetic resonance (NMR), and electron spectroscopy for chemical analysis (ESCA) to elucidate the mechanism of AFEX pretreatment. AFEX first dissolves, then extracts and, as the ammonia evaporates, redeposits cell wall decomposition products (e.g., amides, arabinoxylan oligomers, lignin-based phenolics) on outer cell wall surfaces. As a result, nanoporous tunnel-like networks, as visualized by 3D-electron tomography, are formed within the cell walls. We propose that this highly porous structure greatly enhances enzyme accessibility to embedded cellulosic microfibrils. The shape, size (10 to 1000 nm), and spatial distribution of the pores depended on their location within the cell wall and the pretreatment conditions used. Exposed pore surface area per unit AFEX pretreated cell wall volume, estimated via TEMtomogram image analysis, ranged between 0.005 and 0.05 nm 2 per nm 3 . AFEX results in ultrastructural and physicochemical modifications within the cell wall that enhance enzymatic hydrolysis yield by 4-5 fold over that of untreated cell walls.

Conversion of lignocellulose to biofuels is partly inefficient due to the deleterious impact of cellulose crystallinity on enzymatic saccharification. We demonstrate how the synergistic activity of cellulases was enhanced by altering the hydrogen bond network within crystalline cellulose fibrils. We provide a molecular-scale explanation of these phenomena through molecular dynamics (MD) simulations and enzymatic assays. Ammonia transformed the naturally occurring crystalline allomorph I(β) to III(I), which led to a decrease in the number of cellulose intrasheet hydrogen bonds and an increase in the number of intersheet hydrogen bonds. This rearrangement of the hydrogen bond network within cellulose III(I), which increased the number of solvent-exposed glucan chain hydrogen bonds with water by ~50%, was accompanied by enhanced saccharification rates by up to 5-fold (closest to amorphous cellulose) and 60-70% lower maximum surface-bound cellulase capacity. The enhancement in apparent cellulase activity was attributed to the "amorphous-like" nature of the cellulose III(I) fibril surface that facilitated easier glucan chain extraction. Unrestricted substrate accessibility to active-site clefts of certain endocellulase families further accelerated deconstruction of cellulose III(I). Structural and dynamical features of cellulose III(I), revealed by MD simulations, gave additional insights into the role of cellulose crystal structure on fibril surface hydration that influences interfacial enzyme binding. Subtle alterations within the cellulose hydrogen bond network provide an attractive way to enhance its deconstruction and offer unique insight into the nature of cellulose recalcitrance. This approach can lead to unconventional pathways for development of novel pretreatments and engineered cellulases for cost-effective biofuels production.

Good-quality Raman spectra of most wood species can now be obtained by using near-infrared Fourier transform Raman spectroscopy. To make effective use of such spectroscopic information, one needs to interpret the data in terms of contributions from various wood components and, for each component polymer, in terms of vibrational modes of its substructural units/groups. In the present work, Raman spectral features of black spruce (Picea mariana) wood were associated with lignin and/or carbohydrate polymers. Lignin's spectral contributions were recognized in several ways. In addition to spectra of milled-wood and enzyme lignins, a spectrum of native lignin was obtained by subtracting the spectrum of acid chlorite delignified black spruce from the spectrum of an untreated wood sample. A comparison of lignin spectra indicated that the Raman features of the three lignins are very similar. Raman contributions of carbohydrate polymers, namely, those of cellulose and hemicellulose, were identified by using authentic and/or isolated samples and, in the case of cellulose, by using previously published spectra. Such an analysis showed that the hemicellulose present in black spruce did not give rise to any new, unique features that were not already present due to cellulose. Therefore, it was concluded that the hemicellulose contribution is broad and is hidden under the Raman contribution of cellulose. Also, peak positions of lignin contributions did not overlap with those of cellulose, and there were spectral regions where either lignin or cellulose contributed.

A detailed understanding of the structural organization of the cell wall of vascular plants is important from both the perspectives of plant biology and chemistry and of commercial utilization. A state-of-the-art 633-nm laser-based confocal Raman microscope was used to determine the distribution of cell wall components in the cross section of black spruce wood in situ. Chemical information from morphologically distinct cell wall regions was obtained and Raman images of lignin and cellulose spatial distribution were generated. While cell corner (CC) lignin concentration was the highest on average, lignin concentration in compound middle lamella (CmL) was not significantly different from that in secondary wall (S2 and S2-S3). Images generated using the 1,650 cm(-1) band showed that coniferaldehyde and coniferyl alcohol distribution followed that of lignin and no particular cell wall layer/region was therefore enriched in the ethylenic residue. In contrast, cellulose distribution showed the opposite pattern-low concentration in CC and CmL and high in S2 regions. Nevertheless, cellulose concentration varied significantly in some areas, and concentrations of both lignin and cellulose were high in other areas. Though intensity maps of lignin and cellulose distributions are currently interpreted solely in terms of concentration differences, the effect of orientation needs to be carefully considered to reveal the organization of the wood cell wall.

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Two new methods based on FT-Raman spectroscopy, one simple, based on band intensity ratio, and the other using a partial least squares (PLS) regression model, are proposed to determine cellulose I crystallinity. In the simple method, crystallinity in cellulose I samples was determined based on univariate regression that was first developed using the Raman band intensity ratio of the 380 and 1,096 cm -1 bands. For calibration purposes, 80.5% crystalline and 120-min milled (0% crystalline) Whatman CC31 and six cellulose mixtures produced with crystallinities in the range 10.9-64% were used. When intensity ratios were plotted against crystallinities of the calibration set samples, the plot showed a linear correlation (coefficient of determination R 2 = 0.992). Average standard error calculated from replicate Raman acquisitions indicated that the cellulose Raman crystallinity model was reliable. Crystallinities of the cellulose mixtures samples were also calculated from X-ray diffractograms using the amorphous contribution subtraction (Segal) method and it was found that the Raman model was better. Additionally, using both Raman and X-ray techniques, sample crystallinities were determined from partially crystalline cellulose samples that were generated by grinding Whatman CC31 in a vibratory mill. The two techniques showed significant differences. In the second approach, successful Raman PLS regression models for crystallinity, covering the 0-80.5% range, were generated from the ten calibration set Raman spectra. Both univariateRaman and WAXS determined crystallinities were used as references. The calibration models had strong relationships between determined and predicted crystallinity values (R 2 = 0.998 and 0.984, for univariateRaman and WAXS referenced models, respectively). Compared to WAXS, univariate-Raman referenced model was found to be better (root mean square error of calibration (RMSEC) and root mean square error of prediction (RMSEP) values of 6.1 and 7.9% vs. 1.8 and 3.3%, respectively). It was concluded that either of the two Raman methods could be used for cellulose I crystallinity determination in cellulose samples.

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