Mild Alkaline Pretreatment for Isolation of Native-Like Lignin and

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Mild alkaline pretreatment for isolation of native-like lignin and lignin-containing cellulose nanofibers (LCNF) from crop waste Xuran Liu, Yanding Li, Chinomso M. Ewulonu, John Ralph, Feng Xu, Qilin Zhang, Min Wu, and Yong Huang ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/acssuschemeng.9b02800 • Publication Date (Web): 23 Jul 2019 Downloaded from pubs.acs.org on July 25, 2019

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ACS Sustainable Chemistry & Engineering

Mild alkaline pretreatment for isolation of native-like lignin and lignin-containing cellulose nanofibers (LCNF) from crop waste Xuran Liu,a,b Yanding Li,†c,d Chinomso M. Ewulonu,a,e John Ralph,c,d Feng Xu,f Qilin Zhang,f Min Wu,§a,b and Yong Huang*a aTechnical

Institute of Physics and Chemistry, Chinese Academy of Sciences, Beijing 100190,

China. bCenter

of Materials Science and Optoelectronics Engineering, University of Chinese Academy of

Sciences, Beijing 100039, China. cDepartment

of Biological Systems Engineering, University of Wisconsin-Madison, Madison, WI

53706, USA. dDOE

Great Lakes Bioenergy Research Center, University of Wisconsin–Madison, Madison, WI

53726, USA. eDepartment

of Polymer and Textile Engineering, Nnamdi Azikiwe University, P. M. B. 5025,

Awka, Nigeria. fBeijing

Key Laboratory of Lignocellulosic Chemistry, College of Materials Science and

Technology, Beijing Forestry University, Beijing 100083, China. Keywords: biomass, ball-milling, nanocellulose, NMR, cellulosic film Author Information Corresponding Authors †Email: [email protected] §Email: [email protected] *Email: [email protected] “The authors declare no competing financial interest.”

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Abstract The efficient utilization of crop waste is limited because of the plant cell wall recalcitrance. Solving this problem through sustainable processing, meanwhile producing value-added products, is crucial. Here we report a rapid lignocellulose fractionation method, applied to common reed, using milling under mild alkaline conditions without involving any organic solvents or other reagents. Lignincontaining cellulose nanofibers (LCNF) and native-like lignin were obtained in high yield. LCNF films prepared from the LCNF suspension could have their properties, such as transparency and hydrophobicity, adjusted by changing the pretreatment conditions. Overall, this biomass fractionation provides value-added products and is a successful alternative to simply burning the crop waste. Introduction Biomass, including various woods, grasses, bast fibers, and agriculture straw, is a promising feedstock for the development of sustainable materials by reducing the dependence on fossil energy and petrochemical-based polymers.1 Common reed (Phragmites australis), a commelinid monocot, is widespread with an annual production reaching millions of tons in Chinese wetlands.2 In addition to its applications for woven fabric, reed straws were used to block river canals or burned to generate low-value heat. As with most monocot stem tissues, the chemical composition of the reed straw cell wall consists of 35-50% cellulose, 25-30% hemicelluloses, and 10-20% lignin.3 Recently, there have been many studies focusing on the complete valorization of biomass. The main approach to biomass utilization is to separate the three plant cell wall components and use them separately.4 Nanocellulose, including cellulose nanofibers,5 cellulose nanocrystals, 6and

cellulose nanosheet,7 is a high-value product from the cellulose stream. It can be

prepared by various pretreatment methods such as acid hydrolysis,8 high-pressure homogenizer,9 ball milling,10-11 and ultrasonication treatment12 from cellulose. Among those methods, ball milling has been considered as an environmentally friendly and inexpensive method of producing nanocellulose. Conventionally, lignin removal from lignocellulose has been considered to be a necessary step in the preparation of cellulose nanofiber (CNF).13-14 However, the required pulping and bleaching of lignocellulose


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involves harsh treatment with hazardous chemicals, which causes environmental issues.15 It has recently become apparent that a certain amount of added lignin provides useful features to the CNF, such as reducing polarity, lowering hydrophilicity, and delivering additional thermal stability.16-17 CNF produced directly from the lignocellulose rather than purified fiber could therefore advance the material properties of the nanofibers.18-19 Lignin, the most abundant aromatic component on the planet, is quite sensitive to chemical processing.20-21 Conventional biomass pretreatment methods using harsh acid/base conditions and/or high-temperature organic solvents not only have sustainability problems but can also completely destroy lignin’s native structure.22 The condensed lignin that results is almost impossible to characterize and contains C–C bonds that are difficult to cleave. Valuable platform monomers can be obtained from various lignin ether-cleaving depolymerization methods from native or native-like lignins in which the β-ethers are predominant.23 However, once lignin is condensed by a pretreatment process, monomers can only be obtained in low yields. Generally, the only option left for such condensed lignins is to burn them to generate low-value heat. Here, we present a sustainable ball-milling process in aqueous mild-alkaline solution to produce high-quality lignin-containing cellulose nanofibers (LCNF). The composition, morphology and dispersion stability of LCNF, as well as the optical and contact angle properties of the LCNF films, were characterized in this study. LCNF films showed unique properties and extra mechanical strength compared to CNF. Concomitantly, a native-like high-yield lignin stream was obtained using this process. The lignin fraction, characterized by 2D HSQC NMR, showed no significant structural changes to the polymer backbone, nor condensation, after the mild-alkaline treatment. This native-like lignin is therefore suitable for downstream lignin valorization processes. Results and discussion Chemical compositional analysis and yield of the LCNF The compositional analysis of the reed straw feedstock and LCNF was performed for each processing step, using a conventional Klason lignin method followed by monosaccharides analysis.24-25 In raw straw feedstock, the cellulose, hemicellulose, and lignin contents were 31%, 34%, and 16%, respectively. More lignin can be fractionated as the alkali



concentration increases, with 34% removed under 1% alkaline treatment, and up to 67% under 7% alkaline treatment. Milling time was shown to have a significant effect on the fractionation of lignin. When the alkali concentration was 1%, 24% and 45% of the lignin was removed as ball-milling time increased from 0.5 h to 6 h. With the aid of alkali, hemicelluloses can be dissolved and removed during milling.26 Lignin-hemicellulose ester bonds27 can also be hydrolyzed by the alkaline treatment, which benefits the hemicellulose and lignin dissolution. An ultrasonication step was performed to prepare the LCNF after ball-milling. A small amount of lignin and hemicellulose was further removed after the ultrasonication likely because of the retained alkaline solution. The chemical composition of the samples was also investigated by FTIR (Fig. 1). The chemical structure of cellulose remained unchanged during the treatment, as anticipated. The absorbance bands at 1595, 1505, and 1240 cm−1 were assigned to C = O and C = C aromatic skeletal vibrations.28 The bands at 1738 cm−1 represented the carboxyl groups and are attributed to the hemicellulosic acetate and uronate ester groups, lignin γ-acetate, or the ferulate and p-coumarate esters that are on lignin and/or arabinoxylan hemicelluloses.29 The spectra indicate that the presence of retained lignin and hemicellulose in the LCNF decreased as the alkaline concentration increased. The yields of LCNF, compositional analysis, and isolated lignin levels are summarized (Fig. 2 and Table S1). For example, we obtained LCNF with a 74% yield, along with isolation of 38% of the lignin (on a Klason lignin basis), at 1% NaOH, 2 h ball-milling time, 2 h ultrasonication time (1%-M2-U2). Lignin characterization To elucidate the structural transformation of lignocellulose and the structure of isolated lignin, the lignin was precipitated from the suspension following neutralization with HCl to pH 3. The precipitate was washed by dialysis for 72 h to remove all of the small molecules and salts, then freeze-dried, and characterized by 2D-HSQC NMR (Fig. 3). 2DHSQC NMR has been widely used for charactering lignin condensation.30-33 The resolved 1H–13C

correlation signals in the 2D-HSQC spectra were assigned according to the

literature.29,34 The alkali-pretreated lignins were compared to the enzyme lignin (EL, derived by removing polysaccharides via crude cellulase treatment), which closely resembles lignin’s native structure. No obvious condensation peaks appeared, nor were


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significant structural changes found even from the harshest 7% alkaline milling condition. The NMR spectra (Fig. S1) and molecular weight distribution (Fig. 4) of the pretreated lignins were almost identical to the EL regardless of the alkali concentration used. The only difference was that more lignin and hemicellulose were isolated when higher alkali concentrations were used, as discussed above. Ferulic acid and p-coumaric acid (Fig. 3) are present in most of the monocot species as ester conjugates to lignin or arabinoxylans; a fraction of the ferulate on arabinoxylans, and the ferulate dimers that result from them, integrate combinatorially into lignins, but such components are not readily seen in NMR spectra;35 the same is the case with the usually low levels of ferulate acylating the γ-position of lignin sidechains.36-37 Our alkaline pretreatment extensively cleaved these ester bonds, as readily observed in the NMR and IR analysis. In the aromatic region, the original correlation peak from the C8/H8 position (δC/δH: 103.5/6.3 ppm, FA8 and pCA8 in Fig. 3a) and C7/H7 (δC/δH: 144.4/7.5 ppm, FA7 and pCA7 in Fig. 3a) of the ferulate and p-coumarate moieties disappeared after alkaline treatment. New peaks for the C8/H8 position (δC/δH: 103.5/6.3 ppm, FA8 and pCA8 in Fig. 3b) and C7/H7 (δC/δH: 143.7/7.6 ppm, FA7 and pCA7 in Fig. 3b) of ferulic acid and p-coumaric acid appeared correspondingly. In addition, two new peaks for C2/H2 position (δC/δH: 110.9/7.3 ppm, FA2 in Fig. 3b) and C6/H6 (δC/δH: 122.0/7.3 ppm, FA6 in Fig. 3b) of the ferulic acid appeared as resolved peaks. The chemical shift changes of these correlation signals unambiguously indicated the cleavage of ester conjugates and the formation of the corresponding carboxylic acids. Such cleavage obviously benefits the separation of lignin and hemicellulose from the lignin-hemicellulose matrix. An interesting observation is that tricin, a valuable flavonoid compound found in the grass cell wall,38 was not soluble and precipitated out with the lignin. We have noted that the solubility of tricin is low especially in aqueous solution. It is highly likely that tricin remained in the cell wall during our pretreatment process. The purity of the isolated lignin is reported in the supporting information (Table S3). In general, there was only a trace amount of cellulose residue in the isolated lignin (less than 8% from the highest 7% NaOH condition). With an increase in alkali concentration, more xylans were observed to be coisolated with the lignin (increasing from 4% to 39%). Overall, the isolated lignin preserved its native structure which had a high β-ether content and minimal condensation/degradation.



The isolated high-quality lignin is suitable for further upgrading by various lignin depolymerization processes. Morphology of the LCNF The morphology of the reed straw and ball-milled samples was determined by scanning electron microscopic imagine (SEM) (Fig. S2). The raw reed straw showed to be macro-fabric materials (with diameter >100um) with smooth surface (Fig. S2a). After the ball-milled treatment without any alkali, the reed straw was broken into small pieces which subsequently aggerated into a bulk material (Fig. S2b). After ball-milled with alkali, the reed straw turned into a porous bulk material without visibly fibric structure (Fig. S2c-e). This was attributed to extraction of the dissolved lignin and hemicellulose by alkali solution from the small pieces of reed straw broken down by ballmilling. After ball-milling, the samples were defibrated by ultrasonication and the as-prepared LCNF examined by atomic force microscopic (AFM) imaging (Fig. 5a-c). The LCNF prepared under 1%M2-U2 condition was more uniform with a diameter of less than 5 nm. The 4%-M2-U2 mainly consisted of fibers with ~5 nm diameter, and a few fibers of larger diameter. LCNF prepared under the 7%-M2-U2 condition consisted of fibers with over 10 nm diameter and the fibers were shorter. As a control, the defibrillation of the LCNF was incomplete when there was no alkali present during the ball-milling (Fig. 5d). This result can be attributed to that the retaining lignin facilitate cellulose fibrillation. As an antioxidant, lignin stabilizes free radicals which potentially counteract recombination reactions between reactive cellulose radicals generated in the alkali ball-milling process. Similarly, hemicellulose residues also affect the charge densities and structure of the of the refined cellulose and further promote cellulose fibrillation.39 A similar result could be confirmed by Transmission electron microscopy (TEM) images (Fig. 5 e-h). Crystallinity and dispersion stability of the obtained LCNF The crystal morphology and crystallinity of LCNF were examined by X-ray diffraction (XRD) analysis (Fig. 6). Mild alkaline ball-milling and ultrasonication did not change the crystal form of the LCNF. However, the crystallinity decreased from 51% to 23% as the alkali concentration increased from 1% to 7% at a constant 2 h of ball-milling time. It can be rationalized that the higher the percentage of retained lignin in the LCNF, the harder it


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is for hydroxide ions to access the inner glycosidic bonds of amorphous cellulose and then liberate shorter cellulose crystals. It also illustrated that the crystallinity of 1%-M2 is slightly reduced from 39% to 34% for 1%-M2-U2, i.e., after ultrasonication, in which the hydrodynamic forces are known to play an important role in decreasing the cellulose crystallinity.40 The surface charge of the LCNF colloidal suspension was obtained by measuring the zeta potential (ZP), which reflects its colloidal stability.41 The higher the absolute value of the ZP, the more stable the suspension. The ZP of the produced LCNF ranged between – 32 to –25 mV. The 1%-M2-U2 LCNF suspension exhibited the highest ZP (–32 mV) which also had the highest stability among all of the conditions. The transmittance appearance of each of the LCNF suspensions was consistent with the ZP analysis (Fig. 7). The suspension at 1%-M2-U2 condition was more transparent compared to other suspensions. The light transmittance of a 0.5 wt% LCNF suspension was 69% at 800 nm wavelength light. We contend that the LCNF of 1%-M2-U2 are longer, allowing more surface negative charges along the length, which results in a higher electrostatic repulsion of the nanofibers. Structure and properties of the LCNF films LCNF films were prepared from 1%-M2-U2, 4%-M2-U2 and 7%-M2-U2 LCNF suspension respectively. The LCNF films had a light brownish color and good transparency (Fig. 8a). The highest light transmittance was 82% at 650 nm, in the sample prepared from 1%-M2-U2 LCNF. Because the diameter of 1%-M2-U2 LCNF was less than 5 nm, the fibers and the voids between them were much smaller than the wavelength of visible light which cause the transparency of the resultant film. The surface of the LCNF films was viewed under AFM (Fig. 8b). The compact fiber network, in which the nanofibers were interconnected with each other, illustrates that the film surface is uniform and smooth. The distribution of the lignin on the surface of the LCNF films was visualized by monitoring lignin’s aromatic ring signatures via Confocal Laser-Raman Spectrometry (Fig.8c-d); the intensity differences reflected the different concentrations of aromatic rings. The distribution of lignin was clearly uneven, likely due to the deconstruction of the plant cell wall followed by lignin redistribution.



The static water contact angles of pure CNF and LCNF films were measured to show the change of hydrophilicity due to the presence of lignin (Fig. 9a). Compared to the ligninfree cellulosic CNF film (ball-milled in water without alkali as described in the Experimental Section), the water contact angle of the LCNF film was significantly higher because of the retained lignin and hemicellulose. It increased from 45° (CNF film) to 74° (1%-M2-U2 LCNF film). The higher water contact angle gives the LCNF film a better hydrophobicity that can be beneficial for specific applications. The mechanical properties of the LCNF films were measured by Young’s modulus (Fig. 9b). The 1%-M2-U2 LCNF films had representative values of tensile strength of 104.4 MPa and Young’s modulus of 4612.4 MPa; these results were 79% and 28% higher than those of the cellulosic CNF films. The retained lignin and hemicellulose dramatically promoted the strength and modulus of the films because of the lignin’s bonding effect just like in the natural straws. Conclusions When designing a new biomass pretreatment process, most of the components in the biomass should be considered while attempting to preserve their ideal properties for further utilization. More importantly, new processes should not only be more efficient but also less harmful to the environment compared to conventional methods. LCNF can be obtained from reed straw in high yield by a fast aqueous mild-alkaline ball-milling process. The isolated native-like lignin appears to be ideal for further lignin upgrading. The LCNF films prepared from our LCNF showed better properties compared with traditional CNF films. We contend that this study is a good example of an improved utilization strategy for grassy biomass, especially crop wastes. Experimental Materials Reed (Phragmites australis) straw was collected from Baiyang Lake in Hebei Province, China. Reed straws were cut into small pieces using scissors, washed with distilled water and dried overnight in an oven at 50 °C. The dried straw pieces were pulverized and then sieved through a 60-mesh screen. Sodium hydroxide (NaOH) was analytical grade and


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provided by Beijing Chemical Reagent Co. The cellulose material used in this study as a control was purchased from Sigma-Aldrich (α-cellulose, CAS:9004-34-6).

Biomass fractionation and LCNF preparation Reed flour (1 g) and 20 mL of NaOH solution (1, 4, 7 wt %) were added to a 45 mL zirconia pot containing seven zirconia balls (d = 10 mm). Control experiments were also performed by adding 20 mL distilled water. Ball-milling was carried out under programmed punctuated operation (working time 20 min and interval of 2 min) with a rotation speed of 300 rpm at room temperature for 0.5, 2, and 6 h. After ball-milling, the mixture was diluted with excess distilled water and washed several times until the supernatant pH was 7-8, followed by centrifugation at 13,000 rpm for 5 min. Pure CNF was prepared as a control. α-Cellulose (1 g) was ball-milled with 20 mL distilled water for 2 h. Enzyme lignin from the ball-milled biomass was prepared as previously reported.42 The washed fibers were then ultrasonicated (JY99-IIDN ultrasonic homogenizer, Ningbo Scientz Biotechnology Co., Ltd, China) at an output power of 450 W for 2 h (working time 2 s and interval of 2 s) in an ice-bath. The resulting LCNF were labelled X-MY-UZ; where X is the percentage of alkali used, Y is the ball-milling time, and Z is the ultrasonication time. Preparation of LCNF film An CNF/LCNF suspension (5 mL, 0.5 wt%) was vacuum-filtered through a 0.45 micron Nylon membrane. The wet film was peeled from the membrane and dried in oven at 60 °C overnight. The thickness of the film was controlled by the amount of suspension used.

Characterization of LCNF and lignin Compositional analysis The Klason lignin and compositional analysis of the raw reed flour and the obtained samples were determined according to the standard of Determination of Structural Carbohydrates and Lignin in Biomass by the National Renewable Energy Laboratory (NREL), USA. The LCNF yield is defined as:



𝐘=

𝑴𝑳 𝑴𝑹

× 𝟏𝟎𝟎

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(1)

In this equation, Y is the yield of LCNF, WL is the dry weight of LCNF (after dialysis and filtration), and WR is the dry weight of reed straws. The experiments were repeated three times to take the average mass fraction of the original straw weight to obtain the final yield of LCNF. Nuclear magnetic resonance spectroscopy (NMR) NMR spectra were acquired on a Bruker Biospin AVANCE-III 700 MHz spectrometer fitted with a cryogenically cooled 5-mm QCI 1H/31P/13C/15N gradient probe with inverse geometry (proton coils closest to the sample) and spectral processing used Bruker’s Topspin 4.0.6 (Mac) software. For NMR experiments, enzyme lignin and isolated lignins were dissolved in 4:1 v/v DMSOd6/pyridine-d5. The central solvent peaks were used as the internal references (δC/δH: DMSO, 39.5/2.49 ppm). Standard Bruker implementations of the traditional suite of one-dimensional (1D) and two-dimensional (2D) 1H–13C heteronuclear single-quantum coherence (HSQC) NMR experiments were used for structural elucidation. Adiabatic 2D-HSQC (“hsqcetgpsisp2.2”) experiments were carried out using the parameters described previously.29 Processing used typical matched Gaussian apodization in F2 (LB = −0.5 GB = 0.001) and squared cosine-bell apodization in F1. Gel-permeation chromatography (GPC) Enzyme lignin and isolated lignins (5 mg) were dissolved in DMF (5 mL) and then filtered through a PTFE membrane (0.2 μm). Molecular weight distributions of lignin were determined by GPC using a Shimadzu LC20-AD LC pump equipped with a Shimadzu SPD-M20A UV-vis detector set at 280 nm and a Polymer Standard Services GPC column and guard column [PSS PolarSil analytical Linear S, 8-mm inner diameter (ID) × 5 cm and 5-mm particle size → PSS PolarSil analytical Linear S, 8-mm ID × 30 cm and 5-mm particle size]. The column oven was held at 50 °C during analysis. The mobile phase was dimethylformamide (DMF) with 0.1 M lithium bromide, and the flow rate was 0.3 mL/min. Molecular weight distributions were determined using


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Shimadzu GPC postrun software via a conventional calibration curve using a ReadyCal polystyrene Kit from Sigma-Aldrich [Aldrich # 76552, M(p) 250-70000]. Calibration curve (3rd order fitting, R2=0.996): y = ―4.449𝑒 ―5𝑥3 + 0.015𝑥2 ― 1.772𝑥 + 72.981

(2)

Fourier-transform infrared spectroscopy (FTIR) FTIR spectra were collected using Varian 3100 Excalibur Series Fourier Transform Infrared Spectrometer (FTIR) with Attenuated Total Reflectance (ATR). The spectra collected were within the data range of 600 cm-1 to 4000 cm-1 at a resolution of 4 cm-1. Thin sheets were formed for the FTIR analysis using the freeze-dried samples by pressing them in a hydraulic press. Scanning Electron Microscopy (SEM) SEM images (Hitachi S-4800) were used for analyzing the surface morphology of the ballmilled samples. A thin layer of gold was coated onto the dried samples using a vacuum-ion sputter-coater (Hitachi MC1000) before subjecting them to the SEM. X-ray diffraction (XRD) and degree of crystallinity The XRD patterns of the pellet were recorded by an X’Pert PRO X-ray diffractometer (Bruker AXS GmbH) with Curadiation (λ = 0.154184 nm) from 5° to 60°. The increment step was 0.02°, and the scan speed was 0.05 s/step. The crystallinity degree was calculated by equation 2, where I002 is the intensity of the crystalline region of cellulose (2θ = 22.2°) and Iam is the intensity of the amorphous phase (2θ = 18.6°). 𝐂𝐈（%） =

𝑰𝟎𝟎𝟐 ― 𝑰𝒂𝒎 𝑰𝟎𝟎𝟐

× 𝟏𝟎𝟎

(3)

UV-vis spectroscopy and light transmittance The UV-vis spectroscopy of aqueous LCNF suspensions was measured in the wavelength range of 300–800 nm by using a TU-1900 spectrophotometer (Beijing Purkinje General Instrument Co. Ltd, China). Light transmittance of LCNF films was measured from 200 to



800 nm using a UV-vis-NIR spectrometer (Varian Cary 5000) containing an integrating sphere (air was used as the reference). The obtained LCNF suspensions were frozen in liquid nitrogen and dried under high vacuum at -85 °C using a freeze-drier (Eyela FDU-2110, Japan). Atomic force microscopy (AFM) AFM was carried out to evaluate the extent of defibrillation of the straws using the atomic force microscope (Bruker Multimode-8, Germany). The LCNF images obtained were analyzed by NanoScope Analysis software version 1.40 (Bruker Company, Germany). A drop of a suspension from the prepared LCNF was diluted with about 10 mL distilled water and sonicated for 5 mins in an ultrasonic cleaner (Kunshan Shu Mei KQ-250DE, China) to obtain good dispersion of the nanofibrils. An aliquot (50 µL) of the dispersion was deposited onto a mica plate and left to air-dry at room temperature. Transmission electron microscopy (TEM) TEM images obtained were analyzed to confirm the surface diameter of the LCNF using Nano Measurer version 1.2.5 software (Department of Chemistry, Fudan University, China). Several drops of diluted LCNF suspension were deposited on glow-discharged carbon-coated electron microscope grids after sonication to achieve good dispersion. After the samples were completely dry, the specimens were observed using JEOL JEM 2100 Transmission Electron Microscope, Japan. The zeta potential was measured using Malvern 3000HSA Zetasizer (Malvern Instruments, UK). An average reading was calculated from 5 measurements at 25 °C. Raman spectroscopy Raman spectra were recorded using a Microscopic Confocal Laser-Raman spectrometer (Renishaw). Data was acquired by using a laser power 100 mW at 532 nm. The surface of the film (20 μm × 20 μm) was randomly selected, and the aromatic ring fingerprint peak (1605 cm-1) of lignin was measured. Static water contact angles (WCA)


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WCA of the samples were measured using OCA20 contact angle meter (DataPhysics Instruments GmbH, Germany) at room temperature. Water (4 µL) was dropped onto the surface of the LCNF films for the measurement. The final WCA were calculated by averaging readings from 5 different positions on each sample. Mechanical properties measurement Tensile mechanical properties of CNF films were measured by an MTS Sintech-1 (MTS Systems) at a strain rate of 4 mm/min. Specimen strips were 5 mm wide and 30 mm long, with 20 mm gauge length. The thickness of the CNF films was measured separately before the measurements.

Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: The compositional analysis, NMR, SEM characterization of the lignin and LCNF samples, Tables S1-S3, and Figures S1-S2.

Acknowledgements This study was supported by the National Natural Science Foundation of China (No. 51733009), Chinese Academy of Sciences-President’s International Fellowship Initiative (CAS-PIFI) Postdoctoral Research in China (No. 2017PS0019) and Funding to YL and JR from the DOE Great Lakes Bioenergy Research Center (DOE BER Office of Science DESC0018409).



Figure and Table captions

7%-M2-U2

4%-M2-U2 1%-M2-U2

3500

3000

1500

Wavenumber/cm-1 Fig. 1 FTIR spectra of reed straw, pretreated reed straw, and LCNF.


897

1505 1465 1423 1376 1240

1595

1738

2900

0%-M2-U2 reed flour

3407

Transmittance/a.u.

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

Page 14 of 35

1000

Page 15 of 35

100

lignin

cellulose

87.2

80

76.5

hemicellulose

80.0

79.5

74.0

71.3

70.3 69.0

others

64.4

60

60.1

40

2 -U

2

2 7%

-M

2 -U

2 4%

-M

2 -U

2 1%

-M

2 -U

-M

-M 6

0%

1%

-M 0.5

-M 2

1%

7%

-M 2

4%

-M 2

1%

0%

ree

ds

0

-M 2

20 tra w

Percentage / %

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


Fig. 2 Compositional analysis (Klason lignin and sugar analysis) of the reed straw, pretreated reed straw, and LCNF. X%-MY-UZ represents the concentration of NaOH concentration (%, wt/v), ball-milling time, and ultrasonication time, respectively.



Fig. 3 Partial 2D HSQC NMR of reed straw enzyme lignins (EL) and alkali-pretreated reed lignin (1% NaOH). The retained polysaccharides are mainly xylan in the alkali-soluble lignin. The assignment of xylan peaks is shown in Figure S1.


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EL 1% AL 4% AL 7% AL

a. u.

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


80

90

100

110

120

min

Fig. 4 Molecular weight distribution of EL and alkali-pretreated lignin (AL) by gel-permeation chromatography (GPC). The x axis is the retention time. The y axis shows the response of a UV-light detector (at 280 nm) normalized to the most abundant signal in each chromatogram.



Page 18 of 35

a

b

c

d

e

f

g

h

200 nm

200 nm

200 nm

200 nm

Fig. 5 The morphology of prepared LCNF. AFM images of (a) 1%-M2-U2, (b) 4%-M2-U2, (c) 7%-M2-U2, (d) 0%-M2-U2. TEM images of (e) 1%-M2-U2, (f) 4%-M2-U2, (g) 7%-M2-U2, (h) 0%-M2-U2.


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Fig. 6 X-ray diffraction spectrum (a), (b) and calculated crystallinity (c) of the pretreated samples and LCNF.



Fig. 7 (a) Zeta potential plot of the LCNF suspensions. (b) UV-vis transmittance spectra of 0.5 wt% LCNF suspensions.


Page 20 of 35

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Fig. 8 (a) UV-vis transmittance spectra of the LCNF films. (b) Photograph and AFM image of the surface of the 1%-M2-U2 LCNF films. (c) Raman image of lignin distribution in the 1%-M2U2 LCNF film.



Fig. 9 (a) Water contact angles (WCA) of pure CNF and LCNF films. (b) Tensile strength and Young’s modulus of the LCNF films. The green bars are Tensile strength, the red bars are Young’s modulus.


Page 22 of 35

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17. Wang, X.; Sun, H.; Bai, H.; Zhang, L.-p., Thermal, mechanical, and degradation properties of nanocomposites prepared using lignin-cellulose nanofibers and poly (lactic acid). BioResources 2014, 9 (2), 3211-3224. 18. Dou, J.; Kim, H.; Li, Y.; Padmakshan, D.; Yue, F.; Ralph, J.; Vuorinen, T., Structural characterization of lignins from willow bark and wood. Journal of Agricultural & Food Chemistry 2018, 66 (28), 7294-7300. 19. Shi, Z.; Yang, Q.; Ono, Y.; Funahashi, R.; Saito, T.; Isogai, A., Creation of a new material stream from Japanese cedar resources to cellulose nanofibrils. Reactive and Functional Polymers 2015, 95, 19-24. 20. Tobimatsu, Y.; Takano, T.; Umezawa, T.; Ralph, J., Solution-state multidimensional NMR of lignins: Approaches and applications. In Lignin: Biosynthesis, Functions, and Economic Significance, Lu, F.; Yue, F., Eds. Nova Science Publisher, Inc.: Hauppauge, NY, USA, 2019; pp 79-110. 21. Ralph, J.; Lapierre, C.; Boerjan, W., Lignin structure and its engineering. Current Opinion in Biotechnology 2019, 56, 240-249. 22. Schutyser, W.; Renders, T.; Van den Bosch, S.; Koelewijn, S.-F.; Beckham, G. T.; Sels, B. F., Chemicals from lignin: an interplay of lignocellulose fractionation, depolymerisation, and upgrading. Chemical Society Reviews 2018, 47 (3), 852-908. 23. Shuai, L.; Amiri, M. T.; Questell-Santiago, Y. M.; Héroguel, F.; Li, Y.; Kim, H.; Meilan, R.; Chapple, C.; Ralph, J.; Luterbacher, J. S., Formaldehyde stabilization facilitates lignin monomer production during biomass depolymerization. Science 2016, 354 (6310), 329-333. 24. Dence, C. W., The determination of lignin. In Methods in Lignin Chemistry, Lin, S. Y.; Dence, C. W., Eds. Springer-Verlag: Heidelberg, 1992; pp 33-61. 25. Sluiter, A.; Hames, B.; Ruiz, R.; Scarlata, C.; Sluiter, J.; Templeton, D.; Crocker, D., Determination of structural carbohydrates and lignin in biomass. Laboratory Analytical Procedure (LAP)-Technical Report 2012, NREL/TP-510-42618. 26. Shi, Z.; Yang, Q.; Cai, J.; Kuga, S.; Matsumoto, Y., Effects of lignin and hemicellulose contents on dissolution of wood pulp in aqueous NaOH/urea solution. Cellulose 2014, 21 (3), 1205-1215. 27. Lapierre, C.; Voxeur, A.; Karlen, S. D.; Helm, R. F.; Ralph, J., Evaluation of feruloylated and p-coumaroylated arabinosyl units in grass arabinoxylans by acidolysis in dioxane/methanol. Journal of Agricultural and Food Chemistry 2018, 66 (21), 54185424. 28. McClelland, D. J.; Motagamwata, A. H.; Li, Y.; Rover, M. R.; Wittrig, A. M.; Wu, C.; Buchanan, J. S.; Brown, R. C.; Ralph, J.; Dumesic, J. A.; Huber, G., Functionality and molecular weight distribution of red oak lignin before and after pyrolysis and hydrogenation. Green Chemistry 2017, 19 (5), 1378-1389. 29. Kim, H.; Ralph, J., Solution-state 2D NMR of ball-milled plant cell wall gels in DMSOd6/pyridine-d5. Organic & Biomolecular Chemistry 2010, 8 (3), 576-591. 30. Gioia, C.; Lo Re, G.; Lawoko, M.; Berglund, L., Tunable thermosetting epoxies based on fractionated and well-characterized lignins. Journal of the American Chemical Society 2018, 140 (11), 4054-4061. 31. Lancefield, C. S.; Wienk, H. L. J.; Boelens, R.; Weckhuysen, B. M.; Bruijnincx, P. C. A., Identification of a diagnostic structural motif reveals a new reaction intermediate


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Page 26 of 35

For Table of Contents Use Only Ball-milling with the aid of mild alkaline solution was performed on the reed to fractionate nativelike lignin and high-quality lignin-containing cellulose nanofibers for crop waste valorization.

NaOH

reed

Ultrasonic

LCNF

Ball-milling

Native-like lignin


LCNF film

Page 27 of 35

7 % -M 2 -U 2

4 % -M 2 -U 2 1 % -M 2 -U 2

3 5 0 0

3 0 0 0

1 5 0 0

W a v e n u m b e r /c m - 1 ACS Paragon Plus Environment

8 9 7

1 2 4 0

1 5 0 5 1 4 2 3

1 4 6 5 1 3 7 6

1 5 9 5

1 7 3 8

2 9 0 0

0 % -M 2 -U 2 r e e d flo u r

3 4 0 7

T r a n s m itta n c e /a .u .

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


1 0 0 0

1 0 0

lig n in

c e llu lo s e

Page 28 of 35

h e m ic e llu lo s e

o th e rs

8 7 .2

8 0

8 0 .0

7 6 .5 7 0 .3

6 9 .0

7 9 .5 7 1 .3

7 4 .0 6 4 .4

6 0

6 0 .1

4 0 2 0


1 % -M 2 -U 2 4 % -M 2 -U 2 7 % -M 2 -U 2

-M

2 -U 2

0 re e d s tra w 0 % -M 2 1 % -M 2 4 % -M 2 7 % -M 2 1 % -M 0 .5 1 % -M 6 0 %

P e rc e n ta g e / %

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


a) Reed_EL Page 29 of 35

Polysaccharides Methoxy

Bβ

Cβ

b) Reed_alkaline lignin 50& Engineering ACS Sustainable Chemistry HO MeO Bβ Cβ

Methoxy

HO

γ

α

β O 4'

2'

MeO

6' β

OH

O Aγ Aγ OMe 1 60 6 2 γ α 2 6 2 A Bγ 3 Bγ (β–O–4) B 4 70 (β–5) Cγ Cγ γ O Aα 5 Aα 6 β OMe 6 O α 7 6′ 80 O 2 Aβ-G 8 8 9 HO O Aβ-G C OMe 2 2′ (β–β) 9 Bα Bα Aβ-S Aβ-S Cα Cα 6 3 T 10 tricin 90 OH O 11 6 5 4 3 6 5 4 3 90 12 T8 13 T6 6 2 6 2 6 2 100 14 T2′/6′ T3 5 S2/6 S2/6 5 3 OMe MeO OMe 15 O O O G2 G2 110 FA 2 16 FA8 S H G 17 FA8 syringyl p-hydroxyphenyl guaiacyl pCA8 G5/6 G 5/6 pCA8 120 18 O O HO O O O HO O FA6 FA5 FA5 19 9 9 9 9 pCA3/5 pCA2/6 pCA2/6 pCA3/5 8 8 8 8 20 H2/6 130 7 7 7 7 H2/6 21 6 2 6 2 6 2 6 2 22 140 5 3 5 3 5 5 FA7 OMe OMe Pyridine 23FA7 ACS Paragon Plus Environment pCA7 pCA7 Unresolved OH OH O OH 13C 24 pCA pCA FA FA 1H ppm 7 825 7 6 8 p-Coumarate p-Coumaric acid ferulate ferulic acid


EL 1% AL 4% AL 7% AL

a. u.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

Page 30 of 35


80

90

100

110

120

min

Page 31 of 35

a

b

e

f

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 200 nm 20


c

d

g

h


200 nm

200 nm

200 nm



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ACS Sustainable Chemistry & Engineering 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20


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Mild Alkaline Pretreatment for Isolation of Native-Like Lignin and

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