Structural modification of hemicelluloses and lignin based on the biorefinery process with white-rot fungal

Carbohydrate Polymers 153 (2016) 7–13

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Carbohydrate Polymers journal homepage: www.elsevier.com/locate/carbpol

Structural modification of hemicelluloses and lignin based on the biorefinery process with white-rot fungal Jian-feng Ma a , Hai-yan Yang b , Wang Kun c,∗ , Xing-e Liu a,∗ a b c

International Center for Bamboo and Rattan, Beijing 100102, China College of Materials Engineering, Southwest Forestry University, Kunming 650224, China Beijing Key Laboratory of Lignocellulosic Chemistry, Beijing Forestry University, Beijing 100083, China

a r t i c l e

i n f o

Article history: Received 23 June 2016 Received in revised form 19 July 2016 Accepted 19 July 2016 Available online 20 July 2016 Keywords: Biodegradation Topochemistry Hemicelluloses Lignin Structural modification

a b s t r a c t On the concept of biorefinery, hemicellulosic and lignin fractions were isolated from white-rot fungal Trametes velutina D10149 biodegraded poplar, and the structural modification was elucidated in detail according to the different incubation duration. Transversal-section Raman images showed that the fiber secondary walls were preferentially degraded, whereas the compound middle lamellae, including the cell corner regions, were mainly intact after 16 weeks incubation. More importantly, lignin and carbohydrates were simultaneously removed within the fiber secondary wall. From wet chemistry analysis, the yields and structural properties for both hemicellulosic and lignin fractions were not significantly altered. The synergistic effect of ligninolytic system finally obviously appeared after 16 weeks incubation, evidenced by the remarkable decrement of hemicellulose and lignin molecular weights. Additionally, the preferential degradation of S units in lignin biomacromolecule was further confirmed by composition analysis of cell wall phenolics and the integration of 2D NMR correlations in the aromatic region. © 2016 Elsevier Ltd. All rights reserved.

1. Introduction Second-generation bioethanol from lignocellulosic biomass hasattracted much interests over last few decades, and enzymatic saccharification has been identified as one of the most costly steps in cellulosic ethanol production. However, in spite of the ubiquity of 2nd bioethanol feedstock, plant cell walls have protective and structural functions and are therefore resistant to degradation: the intrinsic characteristics of cellulose (i.e. crystallinity, degree of polymerization, etc.), the lignin and hemicelluloses network surrounding cellulose limits the accessibility of the enzymes. Many industrial pretreatments (steam-explosion, acid and alkaline pretreatment etc.) have been developed to physically remove lignin and hemicelluloses from cell walls and to expose cellulose to hydrolytic enzymes. These pretreatments are effective but have negative effects, such as high consumption of energy and chemicals, and generation of toxic byproducts. Thereby, fungal pretreatment using white rot fungi has attracted extensive attention for biorefinery, as the remarkable abilities of delignification and certain advantages of low-cost, environmentally friendly and no emission

∗ Corresponding authors. E-mail addresses: [email protected], [email protected] (W. Kun), [email protected] (X.-e. Liu). http://dx.doi.org/10.1016/j.carbpol.2016.07.085 0144-8617/© 2016 Elsevier Ltd. All rights reserved.

of inhibitors to fermentation (Alvira, Tomas-Pejo, Ballesteros, & Negro, 2010; Kumar, Revathi, & Khanna, 2014). Furthermore, effective fractionation of each component in high yield and purity could provide multiple high-valued chemicals, breaking through the fuelonly production model by the traditional fractionation process. Lignin, as one of the most recalcitrant to biodegradation of all natural polymers, comprises roughly 15% of all terrestrial biomass. Structurally, lignin is a polymer of heterogeneous phenylpropanoid units in vascular plants that is built randomly by oxidative coupling between hydroxyphenyl (H), guaiacyl (G), and syringyl (S), which are derived from three corresponding monolignols: p-coumaryl alcohol (pCoumA), coniferyl alcohol (ConA), and sinapyl alcohol (SinA). The extremely complex structure of lignin is finally formed by the variety of a series of characteristic linkages (e.g. ˇ-O-4, ˛-O-4, and ˇ-␤,) and distribution in cell wall. Lignin plays an important role in providing strength, protection against pathogens, improving water conduction and preventing degradation of structural polysaccharides by hydrolytic enzymes. To improve the biological removal of lignin, many white-rot fungi have been tested in microbial pretreatments for many kinds of lignocellulosic materials (Dong, Yang, Zhu, Wang, & Yuan, 2013). Overall, lignin biodegradation by white rot fungi is an oxidative process and phenol oxidases are the key enzymes, acting synergistically with xylanases to disrupt the hemicelluloses-lignin association (Rabinovich, Bolobova, & Vasil’chenko, 2004). A new

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fungus, Trametes velutina D10149, was recently isolated and identified in Institute of Microbiology, Beijing Forestry University, and reported to be a promising microorganism for biopretreatment. 1.3 Times increment of bioconversion and 4 times enhancement of ethanol production were achieved after being pretreated for 8 weeks with this fungus (Wang, Yuan, Cui, & Dai, 2012). As the principal barrier to enzymatic hydrolysis and direct objective of pretreatment process, lignin and hemicelluloses were selected to study as the independent variables in present work. As the continuation of our previous work (Wang, Yang, Wang, & Sun, 2013), the fractionation efficiency and physicochemical characterization of hemicelluloses and lignin after biodegradation treatment is the emphasis of this investigation. In order to get the full view of the biopretreatment, the investigation on cell wall topochemistry was also integrated into the present work. Information on the structural modification of these recalcitrant barriers at multiscale is important not only for providing additional insight into the mechanism of fungal biodegradation, but also for the industrial process of the bioethanol production, as the efficiently utilization of hemicelluloses and lignin affects the benefit for an integrated lignocellulosic biorefinery. 2. Methods 2.1. Substrate and fungal strain Details of the substrate and strain used in this study are given in the previous paper (Yang, Wang, Wang, & Sun, 2013). The main components of poplar were determined as: glucose 44.4 ± 0.6%, xylose 23.0 ± 0.5%, Klason lignin 19.4 ± 0.3%, and acid soluble lignin 4.9 ± 0.1% (weight% of starting material).The white rot fungal strain, Trametes velutina D10149, was isolated from Jilin province in China and preserved in Institute of Microbiology, Beijing Forestry University. After being activated and cultured, this suspension would act as inocula. 2.2. Biodegradation and alkaline fractionation The biorefinery process based on the synergistic treatment with biodegradation and alkaline fractionation was detailedly described in the previous paper (Yang et al., 2013). After removing the solid residues with filtration under vacuum, the liquid fractions were collected and neutralized. The hemicellulosic components were fractionated with alcohol precipitation, and the lignin macromolecules were obtained by acidification with HCl (Wang, Yang, Yao, Xu, & Sun, 2012). All the samples were freeze-dried and kept in a desiccator at room temperature for further analysis. To reduce errors and confirm the results, each experiment was repeated in double under the same condition. The notations of the hemicellulosic and lignin fractions were HX and LX , respectively, while the subscript X (1–4) represents the increasing incubation period (4, 8, 12 and 16 weeks). H0 and L0 were obtained from poplar with the sole alkaline fractionation process as controls. 2.3. Analysis procedures 2.3.1. Structural carbohydrates The compositions of the structural carbohydrates were determined using National Renewable Energy Laboratory (NREL) protocol (Sluiter, Hames, Ruiz, Scarlata, & Sluiter, 2007), and analyzed by high-performance anion exchange chromatography (HPAEC) (Dionex, ICS 3000, Sunnyvale, CA, USA) on a CarboPac PA 20 analytical column (4 × 250 mm) with pulsed-amperometric detection (PAD).

2.3.2. Molecular weight and distribution Measurement of the molecular weights of the hemicellulosic and lignin fractions by gel permeation chromatography (GPC) were described in the previous paper (Wang, Yang et al., 2012), using sodium phosphate buffer and tetrahydrofuran (THF) as eluents, respectively. By comparing with the reference standards of known molecular weights, the curves and values of samples were calibrated and calculated on a HPLC system equipped with refractive index detector (RID). 2.3.3. Monolignol components and linkages The wet-chemical degradation processes were carried out to determine the monolignol components and ester/ether bound linkages. Alkaline nitrobenzene oxidation was employed to illustrate the structural differences in non-condensed phenolics of the lignin samples (Wang, Jiang, Xu, & Sun, 2009). 1 N and 4 N alkaline (NaOH) solution at room temperature and 170 ◦ C could efficiently break the ester- and ether-bound, respectively, as described in previous paper (Lozovaya, Ulanov, Lygin, Duncan, & Widholm, 2006), which were introduced to examine the main linkages between lignin subunits in this study. Qualitative and quantitative analysis of the released phenolic acids/aldehydes was performed on a HPLC system (1200 Series, ZORBAX Eclipse XDB-C18 column4.6 × 250 mm, Agilent Technologies, USA) by comparison of retention times and UV spectra (DAD, diode array detector) of the eluting peaks and the authentic standard compounds (p-hydroxybenzoicacid, vanillic acid, syringic acid, ferulic acid, pcoumaricacid, p-hydroxybenzaldehyde, vanillin, syringaldehyde, acetovanillone, and acetosyringone) (Sigma–Aldrich Corp.; St. Louis, MO, USA). 2.3.4. Spectral analysis Fourier transform infrared (FT–IR) spectra were recorded on an FT–IR spectrophotometer (Nicolet iN10, Thermo Scientific, USA) in the attenuated total reflection (ATR) mode, ranging from 4000 to 800 cm−1 . The measured spectra were further analyzed by resolution enhancement approaches, as second derivatives, to resolve the overlapped peaks and to follow their variations under investigation. Second derivative spectra were obtained by applying program DERIV to the original data, and no smoothing function was applied. Solution 1D–and 2D–NMR spectra were recorded at room temperature on a Bruker AVANCE 400 MHz spectrometer using a z-gradient triple resonance probe. The hemicellulosic and lignin samples were dissolved in D2 O and DMSO-d6 , respectively, and chemical shifts were calibrated relative to the HOD signal (ı=4.70 ppm) and the central DMSO peak (ıC = 39.5 ppm; ıH = 2.49 ppm). HSQC cross-signals of lignin were assigned by comparing them with previously reported literature (Capanema, Balakshin, & Kadla, 2005; Ralph, Bunzel et al., 2004; Ralph, Ralph, & Landucci, 2004). 2.3.5. Topochemical analysis Small sample blocks of approximately 15 mm (longitudinal) × 5 mm (tangential) × 10 mm (radial) were cut out from the stem of Populustomentosa (3 years old). The 10-␮m-thick crosssections for chemical imaging were cut on a sliding microtome (Leica 2010R). For chemical imaging, samples were placed on a glass slide with a drop of D2 O, covered by a coverslip (0.17 mm thickness) and sealed with nail-polish to prevent evaporation during measurement. Raman spectra were acquired with a LabRam HR800 confocal Raman microscope (Horiba JobinYvon). Measurements were conducted with an Olympus 100 × Oil objective (NA = 1.25) and a linear-polarized 633-nm laser. The laser power on the sample was approximately 8 mW. The Raman light was detected by an air-cooled, front-illuminated spectroscopic charge-coupled device

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Fig. 1. Raman images of poplar wood fiber decayed by Trametes velutina D10149. (A) 2789–3000 cm−1 (Reference fiber, Overall morphology); (B) 2789–3000 cm−1 (Decayed fiber, Overall morphology); (C) 2789–2938 cm−1 (Reference fiber, Carbohydrates); (D) 2789–2938 cm−1 (Decayed fiber, Carbohydrates); (E) 1540–1720 cm−1 (Reference fiber, Lignin); (F) 1540–1720 cm−1 (Decayed fiber, Lignin); (G) Ratio image of reference fiber; (H) Ratio image of decayed fiber.

Table 1 Yields (w/w) and composition of sugars in the hemicelluloses (relative%, w/w) and lignin fractions (weight%, w/w). Hemicellulosesa

Yieldb Galc Glc Xyl Mw Mn Mw /Mn

Lignina

H0

H1

H2

H3

H4

L0

L1

L2

L3

L4

3.2 12.9 6.7 76.4 1.96 × 105 2.15 × 104 9.1

2.2 7.6 25.6 66.9 4.82 × 104 2.64 × 104 1.8

1.6 5.3 42.7 52.0 4.94 × 104 2.63 × 104 1.9

1.3 4.7 43.7 51.6 4.43 × 104 2.58 × 104 1.7

6.4 5.7 49.0 45.4 2.15 × 104 1.33 × 104 1.6

7.1 NDd 0.4 0.6 3687 1390 2.7

5.5 ND 0.4 0.5 2737 1340 2.0

3.4 ND ND 0.3 2833 1407 2.0

3.4 ND ND 0.3 2493 1394 1.8

3.2 ND ND 0.2 1362 794 1.7

a HX and LX represent the hemicellulosic and lignin fractions, respectively, isolated from the biodegraded poplar, while the subscript X (1–4) represent the different cultivation cycles (4, 8, 12 and 16 weeks). H0 and L0 were obtained from poplar with the sole alkaline fractionation process as controls. b Weight% of the starting material as the average of the replicates, and the standard deviation is less than 5%. c Gal, galactose; Glc, glucose; Xyl, xylose. d Not detectable.

(CCD) behind a grating (600 grooves mm−1 ) spectrometer with a spectral resolution of 2 cm−1 . For mapping 0.5 ␮m steps were chosen and every pixel corresponds to one scan. The spectrum from each location was obtained by averaging 2-s cycles. TheLabspect6 software was used for spectral and image processing and analysis. Before a detailed analysis, the calculated average spectra were baseline corrected using the Savitsky-Golay algorithm (linesmethod, 7 points). 3. Results and discussion 3.1. Topochemical changes of poplar wood fiber after biodegradation Raman microspectroscopy can be used to provide a spatial distribution of intact plant cell wall components in their native form. Besides, both carbohydrates and lignin can be mapped simultaneously by selecting Raman bands that are specific to these cell wall components. The overall morphology of the fiber were highlighted

by integrating over the spectral range from 2789 to 3000 cm−1 involving C H and C H2 stretching vibrations, in which all cell wall polymers (cellulose, hemicelluloses, pectin, lignin) contribute to the Raman signal (Gierlinger & Schwanninger, 2007). The morphologically distinct cell wall regions (Cell corner middle lamella, Ccml; Compound middle lamella, Cml and Secondary wall, S) of fiber were clearly differentiated (Fig. 1A). After 16 weeks decay, the fiber S layer displayed the shrinkage and cubical pattern of cracks (Fig. 1B). When integrating over the characteristic band range from 2789 to 2938 cm−1 (C H and C H2 stretching in carbohydrates) and from 1540 to 1720 cm−1 (aromatic ring stretching), the expected high levels of carbohydrates in fiber S layer as well as high lignin concentration in Ccml and Cml was visualized for the reference material (Fig. 1C and E). However, after 16 weeks fungal degradation strong depletion of carbohydrates in the morphologically distinct fiber walls occurred (Fig. 1D), which indicated that fungus was able to utilize the carbohydrates fraction of the wall effectively. Although lignin located in the fiber S layer was also removed, the Cml, including the cell corner regions were mainly intact at this

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point (Fig. 1F). In order to glean more detailed information on the correlation between lignin and carbohydrates concentration at cellular level, images constructed according to lignin/carbohydrates ratio were obtained. A relative increase of the ratio in Ccml and Cml was due to the partial loss of carbohydrates in these areas. However, throughout the fiber S layer the ratio was uniform before and after fungal decay, indicating the degradation of lignin followed that of carbohydrates in the fiber S layer (Fig. 1G and H). Average spectra were calculated for the different cell wall layers (Ccml, Cml and S) by marking the distinct areas on the chemical images. The well-characterized Raman spectral features of lignin have been assigned in Supporting Materials Table S1. (Agarwal & Ralph, 1997). Comparing the average spectra extracted from different fiber cell wall regions (Ccml, Cml and S) before and after 16 weeks exposure to fungus revealed big differences in carbohydrates and lignin concentration (Supporting Materials Fig. S1). Distinct band increments assigned to lignin (1660 cm−1 , 1605 cm−1 , 1333 cm−1 and 1273 cm−1 ) and band decrements assigned to carbohydrates (2897 cm−1 , 1380 cm−1 , 1121 cm−1 , and 1099 cm−1 ) were seen in Raman spectra of decayed fiber Ccml and Cml compared to the reference. However, for the fiber S layer an obvious decrease in band intensity at lignin and carbohydrates related bands was found after 16 weeks bioprotreatment. This is the results of the simultaneous degradation of carbohydrates and lignin in the fiber S layer by fungus. 3.2. Physicochemical characterization of hemicellulosic fractions 3.2.1. Yields and sugar components Probably due to the mild fractionation process (1% NaOH (w/v), 75 ◦ C), the lower yields (1.3-6.4%) of hemicellulosic fractions were finally obtained (Table 1). On the other hand, the alkaline treatment with low severity minimized the structural variation of hemicelluloses in the starting materials, truthfully reflecting the effect of biodegradation process on the physicochemical characterization of hemicelluloses. Clearly, the sugar components were obviously changed with the incubation periods, although the yields were relatively maintained. The gradual decreased content of xylose, corresponding to the increased glucose, indicated that cellulose was partial degraded and could be dissolved during the alkaline fractionation. This result further confirmed that the bioconversion efficiency of cellulose could be improved regardless of the non-selective degradation of lignin with this fungus (Wang, Yang, Wang & Sun, 2013). FT-IR analysis, as an efficient qualitative method, provided non-significant difference by comparing the original spectra (Supporting Materials Fig. S2). Closer analysis by 1st order derivation revealed that the hemicellulosic polymer obtained after biopretreatment process contained less associated lignin (1500 cm−1 ) and partial degraded cellulose (1170 cm−1 ). Similarly, the basic glycosidic linkages and subunits were maintained after biodegradation since the relevant patterns were shown in the 1 H NMR spectra (Supporting Materials Fig. S3). The chemical shifts (ı = 3.1–4.3) are originated from the equatorial proton and other protons of the anhydroxylose residues. Signal at ∼4.4 ppm is due to the anomeric protons of ␤-d-xylose substituted at C-3 (monosubstituted) residues, and the signal at ∼5.2 ppm is ascribed to the ␣-configuration. 3.2.2. Molecular weight distribution Analysis by GPC of H0 from the unbiotreated material revealed the presence of minor amount of higher Mw portion (a shoulder maximum at ∼5E5 Da) together with high proportion of low Mw components (peak maximum at ∼5E4 Da) (Fig. 2 A), and polydispersity (9.1) was consequently auto-calculated, indicating its inherent characteristics of molecular weight distribution (Table 1). After being incubated with white-rot fungus Trametes velutina D10149,

Fig. 2. Molecular weight distribution curves of the hemicellulosic (A) and lignin (B) fractions.

Table 2 Yields of phenolic acids and aldehydes (w/w, ␮g/mg) from alkaline nitrobenzene oxidation of the lignin fractions.

p-hydroxybenzoic acid p-hydroxybenzaldehyde vanillic acid syringic acid vanillin syringaldehyde acetovanillone acetosyringone Total S/Vb

L0

L1

L2

L3

L4

8.3 0.8 3.0 5.1 32.3 49.6 1.2 1.5 101.8 1.54

8.9 0.6 2.4 4.5 30.1 43.2 NDa ND 89.7 1.47

8.1 0.6 1.8 4.1 29.9 38.7 ND ND 83.2 1.35

7.1 0.4 3.9 4.2 31.1 39.6 0.4 0.5 87.2 1.25

5.0 0.7 2.2 3.1 33.6 36.8 ND ND 81.4 1.11

a

Not detectable. S represents the total mass of syringaldehyde, syringic acid and acetosyringone, and V represents the total mass of vanillin, vanillic acid and acetovanillone. b

the values of the Mw were immediately dropped from 1.96 × 105 Da (H0 ) to the region of 4.94 × 104 –4.43 × 104 Da (H1–3 ), as well as the polydispersity (1.7–1.9). This phenomenon was visually illustrated in Fig. 2. The high Mw peak was substantially reduced even after 4 weeks incubation, and the whole Mw distribution curves was gradually shifted to the lower molecular region with prolonging the incubation cycles. Further degradation was observed after 16 weeks biodegradation, clearly evidenced by the appearance of a shoulder representing low-molecular-size components (Mwaround 5E3 Da), together with the markedly declined value of Mw (2.15 × 104 Da).

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Fig. 3. Unit ratio of the obtained lignin fractions. (A) 1N NaOH (ester-bound phenolics); (B) 4N NaOH (ether-bound phenolics); (C) alkaline nitrobenzene oxidation (noncondensed phenolics). S represents the total mass of syringaldehyde, syringic acid and acetosyringone; V represents the total mass of vanillin, vanillic acid and acetovanillone; H represents the total mass of p-hydroxybenzoic acid and p-hydroxybenzaldehyde.

Fig. 4. HSQC spectra (aliphatic region: left; aromatic region: right) of the lignin fractions L0 , L1 and L3 .

3.3. Physicochemical characterization of lignin fractions 3.3.1. Yields and purity Lignification is considered to be a primary factor limiting the biodegradation of the cell wall by rumen microbes, and the knowledge of lignin content in the plant is consequently of primary importance to access the mechanisms involved in the inhibition of structural carbohydrate digestion. However, on the basis of biorefinery concept, efficient fractionation of each component is the primary objective. It is clear that the yields of lignin were indeed gradually decreased with prolonging the biodegradation time, especially after 8 weeks incubation. As reported by Dinis et al., the

activity of ligninolytic system (manganese-dependent peroxidase (MnP), lignin peroxidase (LiP), and laccase) varied greatly at different cultivation times, and the synergistic role among these enzymes normally occurred after 4 weeks incubation (Dinis et al., 2009). The non-selective degradation of lignin with Trametes velutina D10149 could also be observed from the fact that the yields of lignin almost leveled off after 12 week incubation, which is benefit for the lignin utilization. Xylose, as one of the major monosaccharides in lignincarbohydrate complex, was slightly detected in all lignin fractions, indicating that neither the biodegradation process nor the alkaline treatment could completely cleave this prevalent ester bond (Popescu, Popescu, & Vasile, 2010). In addition, hardly any differ-

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ence on the subunits and linkages of lignin biomacromolecular could be observed from FT-IR analysis, ever after the derivation process (Supporting Materials Fig. S4).

According to the automatic integration of the processing software, the S/G ratio was also calculated to be gradually decreased with incubation time from 1.39 (L0 ) to 1.32 (L1 ), and to 1.19 (L3 ) 1

3.3.2. Molecular weight distribution A clear decrement of obtained lignin Mw was observed with biopretreatment process, straightly from 3687 Da (L0 ) to 1362 Da (L4 ) (Table 1). Correspondingly, the main peak of molecular weight distribution was remarkably shifted to the lower molecular weight region (Fig. 2 B). Except for the initial stage of incubation, a sudden degradation of lignin was clearly detected after 16 weeks biodegradation with Trametes velutina D10149, although the Mw peak was still higher than 1000 Da. Combining with the Mw analysis of obtained hemicellulosic fraction (a sudden degradation after 16 weeks incubation), it is reasonable to believe that the extensive cleavage of lignin-carbohydrate complexes occurred at this moment. As known, the covalent bonds between polysaccharides or hemicelluloses play a key role in preventing degradation. Dong et al. confirmed that the esterase participated the degradation procedures of hemicellulose–lignin matrix by cooperation with the other ligninolytic enzymes, although the esterase along could not mineralize the lignin (Dong et al., 2013). The increased effectiveness of the culture filtrates where the possible presence of lignin-degrading enzymes perhaps increased the accessibility to the carbohydrate fraction. This phenomenon also partially suggested that biopretreatment is a time-consuming process, not satisfying the economic feasibility of industrial application. 3.3.3. Composition of cell wall phenolics The ratios of phenolic composition in the isolated lignin fractions are presented in Fig. 3. Clearly, the H units predominated in eater bound cell wall phenolics in all lignin fractions, and its proportion was gradually decreased from 81.5% to 80.5% with white-rot fungi degradation (Fig. 3A), as well as the yields of the detectable monolignol derivatives (Supporting Materials Table S2). In terms of the ether bonds, S-derived lignin units were determined to be the major contributor (Fig. 3B), and the yields of the detectable phenolics were also decreased as incubation processed (Supporting Materials Table S3). Although the relative proportion of H unit almost maintained the same level (5.6%-6.6%), the S/V ratio was found to be gradually decreased from 1.6 to 1.4. The similar trend was also exhibited in the alkaline nitrobenzene oxidation test, which is usually used to analyze the structural features of side chain and extent of carbon-carbon linkages in lignin subunits. The data in this study further confirmed the conclusion that white-rot fungi preferred the degradation of syringyl units, resulting in the decreased S/V ratio from 1.54 (L0 ) to 1.47(L1 ), to 1.35 (L2 ), to 1.25 (L3 ), and to 1.11 (L4 ) (Table 2). Despite the degradation mechanism was not clearly identified, three white-rot fungi, selected by Dong’s group, were all proved to preferably degrade S units of lignin in bagasse (Dong et al., 2013). Two dimensional 1 H-13 C NMR was also employed to further investigate the structural alteration of lignin macrobiomolecular (Fig. 4). In the aliphatic region, the ␤-O-4 aryl ether (I) (identified by 72.5/4.9 ppm (I␣), 84.1/4.3 and 86.0-87.4/4.0-4.2 ppm (I␤), and 59.5-60.5/3.2-3.6 and 62.8-63.5/3.4-3.6 ppm (I␥ )) and resinol substructure (II) (identified by85.4/4.7 ppm (II␣), 54.2/3.1 ppm (II␤), and 71.6/3.8 and 71.6/4.2 ppm (II␥ )) were clearly presented in all samples. It is indicated that the main and basic subunit-linkages were still maintained in the biodegraded lignin fractions. In the aromatic region, the identifications of H, G and S units were also be easily achieved by their specific correlations at 104.7/6.7, 111.7/7.0, 115.0–115.6/6.6–7.0, 119.5/6.8, and 131.9/7.8 ppm, respectively (As shown in Fig. 4). A small quantity of syringic acid was also detected (Popescu, Popescu, & Vasile, 2011).

(S/G = 2 ⁄). This result once again confirmed that the relatively preferential degradation of S units by white-rot fungi. Besides, the unsaturated C C bond in lignin was gradually decreased, and oxidation reactions occurred during the biodegradation process. The related degradation mechanism is in progress. 4. Conclusions Hemicellulose and lignin fractions in biopretreated poplar were successfully recovered by using alkaline fractionation process. Not only the non-selectivity degradation of Trametes velutina D10149 was exhibited from Raman imaging, also the structural modifications for hemicellulose and lignin were not significantly detected from wet chemistry analysis. The gradually decreased S/G ratio, as the strongest characteristics in obtained lignin samples, indicated the preferential degradation of S units by white-rot fungi. Besides, the ovbious decrement of Mw after 16 weeks incubation suggested that the secretion of ligninolytic enzymes is a time-consuming process, and mainly worked byparticipating the degradationof hemicellulose-lignin matrix. Acknowledgement The authors are grateful for grants from the Fundamental Research Funds of ICBR (Grant No 1632015002). Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.carbpol.2016.07. 085. References Agarwal, U. P., & Ralph, S. A. (1997). FT-Raman spectroscopy of wood: Identifying contributions of lignin and carbohydrate polymers in the spectrum of black spruce. Applied Spectroscopy, 51, 1648–1650. Alvira, P., Tomas-Pejo, E., Ballesteros, M., & Negro, M. J. (2010). Pretreatment technologies for an efficient bioethanol production process based on enzymatic hydrolysis: A review. Bioresource Technology, 101, 4851–4861. Capanema, E. A., Balakshin, M. Y., & Kadla, J. F. (2005). Quantitative characterization of a hardwood milled wood lignin by nuclear magnetic resonance spectroscopy. Journal of Agriculture and Food Chemistry, 53, 9639–9649. Dinis, M. J., Bezerra, R. M. F., Nunes, F., Dias, A. A., Guedes, C. V., Ferreira, L. M. M., et al. (2009). Modification of wheat straw lignin by solid state fermentation with a white-rot fungi. Bioresource Technology, 100, 4829–4835. Dong, X. Q., Yang, J. S., Zhu, N., Wang, E. T., & Yuan, H. L. (2013). Sugarcane bagasse degradation and characterization of three white-rot fungi. Bioresource Technology, 131, 443–451. Gierlinger, N., & Schwanninger, M. (2007). The potential of Raman microscopy and Raman imaging in plant research. Spctroscopy, 21, 69–89. Kumar, M., Revathi, K., & Khanna, S. (2014). Biodegradation of cellulosic and lignocellulosic waste by Pseudoxanthomonas sp. R-28. Carbohydrate Polymers, 134, 761–766. Lozovaya, V., Ulanov, A., Lygin, A., Duncan, D., & Widholm, J. (2006). Biochemical features of maize tissues with different capacities to regenerate plants. Planta, 224, 1385–1399. Popescu, C.-M., Popescu, M.-C., & Vasile, C. (2010). Structural changes in biodegraded lime wood. Carbohydrate Polymers, 79, 362–372. Popescu, C.-M., Popescu, M.-C., & Vasile, C. (2011). Carbon-13 CP/MAS solid state NMR and X-ray diffraction spectroscopy studies on lime wood decayed by Chaetomium globosum. Carbohydrate Polymers, 83, 808–812. Rabinovich, M. L., Bolobova, A. V., & Vasil’chenko, L. G. (2004). Fungal decomposition of natural aromatic structures and xenobiotics: A review. Applied Biochemistry and Microbiology, 40, 1–17. Ralph, J., Bunzel, M., Marita, J. M., Hatfield, R. D., Lu, F., Kim, H., et al. (2004). Peroxidase-dependent cross-linking reactions of p-hydroxycinnamatesin plant cell walls. Phytochemistry Reviews, 3, 79–96. Ralph, S.A., Ralph, J., & Landucci, L. (2004b). NMR database of lignin and cell wall model compounds, US Forest Prod. Lab., One Gifford Pinchot Dr. Madison, WI

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