Abstract
Determining the structural changes of lignin during bio-treatment will facilitate the understanding of biomass recalcitrance during the sustainable production of chemicals and fuels. However, the analysis of milled wood lignin (MWL) cannot completely elucidate the complex and irregular structural changes therein. In this study, MWL and lignin degradation compounds were extracted from white-rot fungi-treated poplar in order to unveil the degradation process. Results from MWL revealed that the cleavage of β-O-4′ linkages (from 76.4/100Ar to 31.5/100Ar) and the degradation of β–β′and β-5′ linkages clearly occurred, resulting in a decrease in molecular weight. In addition, G-type lignin was more degraded than S-type lignin, with a slightly elevated S/G ratio from 1.13 to 1.29. Further analyses of lignin degradation compounds confirmed these results by showing a high amount of conjugated and unconjugated C=O functionalities. Furthermore, the degradation product of G-type lignin (vanillin) was detected by 2D HSQC NMR and GC-MS. This study of lignin alterations during white-rot fungi treatment could be beneficial for the sustainable production of chemicals, materials, and fuels from renewable plant resources.
Download PDF
Full Article
Determining Lignin Degradation in White-Rot Fungi-Treated Sacrau Poplar: Lignin Structural Changes and Degradation Compound Analysis
Liming Zhang, Tingting You, Tian Zhou, Lu Zhang, and Feng Xu *
Determining the structural changes of lignin during bio-treatment will facilitate the understanding of biomass recalcitrance during the sustainable production of chemicals and fuels. However, the analysis of milled wood lignin (MWL) cannot completely elucidate the complex and irregular structural changes therein. In this study, MWL and lignin degradation compounds were extracted from white-rot fungi-treated poplar in order to unveil the degradation process. Results from MWL revealed that the cleavage of β-O-4′ linkages (from 76.4/100Ar to 31.5/100Ar) and the degradation of β–β′ and β-5′ linkages clearly occurred, resulting in a decrease in molecular weight. In addition, G-type lignin was more degraded than S-type lignin, with a slightly elevated S/G ratio from 1.13 to 1.29. Further analyses of lignin degradation compounds confirmed these results by showing a high amount of conjugated and unconjugated C=O functionalities. Furthermore, the degradation product of G-type lignin (vanillin) was detected by 2D HSQC NMR and GC-MS. This study of lignin alterations during white-rot fungi treatment could be beneficial for the sustainable production of chemicals, materials, and fuels from renewable plant resources.
Keywords: White-rot fungi; Lignin; Enzyme activity; 2D HSQC NMR; Degradation compounds
Contact information: Beijing Key Laboratory of Lignocellulosic Chemistry, Beijing Forestry University, Beijing, 100083, China; *Corresponding author: xfx315@bjfu.edu.cn
INTRODUCTION
Most terrestrial biomass (including tree trunks and grass stems) is sequestered in lignocellulosic macrostructures that are composed of cellulose, hemicelluloses, and lignin (Yelle et al. 2011). Lignin is recalcitrant to hydrolytic depolymerization because it is a combinatorial and racemic polymer of phenylpropanoid subunits connected via ether and carbon-carbon bonds (Zhao and Dixon 2011), which protects plant structural polysaccharides against microbial attack. White-rot fungi are the only organisms capable of degrading (mineralizing) lignin in wood, thus opening up the plant cell wall (Martínez et al. 2011). With the development of lignocellulose biorefineries, the question of how white-rot fungi circumvent wood and remove lignin has drawn attention because of the demand for the sustainable production of chemicals, materials, and fuels from renewable plant resources (Bozell et al. 2011; Wörmeyer et al. 2011).
Lignin can be oxidized and degraded by a ligninolytic system generated by white-rot fungi, which is composed of lignin peroxidase (LiP), Mn-oxidizing peroxidases, manganese peroxidase, and laccase (Umezawa and Higuchi 1987; Heinfling et al. 1998; Elegir et al. 2005; Mate et al. 2013). Early research on the utilization of white-rot fungi for improving degradation mainly focused on the utilization of a specific number of fungal species and strains. Zadrazil (1985) examined approximately 200 strains of white-rot fungi and found that Phanerochaete chrysosporium,Cyathus stercoreus, and Pleurotus spp. were most commonly used. However, the mechanism of cleaved lignin structures in biodegraded wood attacked by white-rot fungi is poorly understood. In recent years, the mechanism by which lignin model compounds are degraded by the ligninolytic system has been studied (Elegir et al. 2005). Research on the reactivity of the ligninolytic system on lignin model compounds revealed that LiP attacks the β-O-4′ linkage of the lignin model (Umezawa et al. 1987) and that laccase oxidizes β-5′ and 5-5′ lignin substructures in the presence of phenolic groups (Elegir et al. 2005). Manganese peroxidase (MnP) oxidizes and depolymerizes natural and synthetic lignins as well as entire lignocelluloses (Hofrichter 2002). However, whether these mechanisms occur in the biological degradation process is unclear because of the complex macromolecular structure of lignin (a branched polymer comprised of different units and inter-unit bonds) and its intimate association with cell-wall polysaccharides.
The isolation of lignin from fungi-degraded lignocellulosic material is more intuitive than investigating lignin model compounds. However, elucidating the mechanisms of lignin macromolecule degradation or modification during bio-treatment is challenging because it is difficult to obtain a representative lignin sample. Milled wood lignin (MWL) is a representative source of native lignin and has been extensively used to determine the native lignin structure (Wen et al. 2013). However, the analysis of MWL cannot completely elucidate the complex and irregular structural changes of bio-treated samples, although the primary structure is well understood (Mao et al. 2013). The main reasons for this are: (1) isolated MWL is mainly extracted from the middle lamella, whereas biodegradation occurs in multiple locations (Wen et al. 2013), and (2) some “low-molecular” fractions are lost during separation and extraction. Thus, it is equally important to analyze the degradation products generated during white-rot fungi treatment.
In this study, Sacrau poplar was treated with the white-rot fungi Trametes pubescens C7571 and Trametes versicolor C6915 under solid-state conditions. The manganese peroxidase (MnP), lignin peroxidase (LiP), laccase, and hemicellulase activities were determined using a UV spectrophotometer; lignin structural changes (MWL and lignin degradation products) were characterized by Fourier translation infrared spectrum (FT-IR), two-dimensional 1H-13C NMR (2D HSQC NMR), and gas chromatography-mass spectrometer (GC-MS). This analysis enhances the understanding of structural changes that occur in bio-treated lignin.
EXPERIMENTAL
Fungal Incubation
Sacrau poplar was harvested from the experimental farm at Beijing Forestry University, Beijing, China. Before treatment, the wood sample was ground to pass through a 0.9-mm-sized screen. Two fungal strains, T. pubescens C7571 and T. versicolor C6915, were collected from Guangdong and Hebei provinces in China, respectively. Biological treatment was carried out in a 250-mL Erlenmeyer flask with 5 g of air-dried poplar wood and 12.5 mL of distilled water. The samples were sterilized in the autoclave for 20 min at 121 °C and inoculated with 5 mL of inoculum. The cultures were incubated statically at 28 °C for 4, 8, 12, and 16 weeks. The non-inoculated sample served as the control. All experiments were performed in triplicate.
Enzyme Assays
The enzymes were extracted with 50 mM sodium acetate buffer (pH 5.5) supplemented with Tween 60 (0.1 g/L). The entire contents of each bioreactor was transferred to an Erlenmeyer flask and extracted with 500 mL of extracting solution. Five successive extractions were performed at 120 rpm for 4 h at 10 ± 1 °C. After the second extraction, the wood chips were stored overnight while soaking in the extraction solution at 4 °C. The crude extracts were recovered by filtration through a fine filter paper (Souza-Cruz et al. 2004). Enzyme activities were determined at 25 °C using a Helios gamma UV-Vis spectrophotometer (Thermo Fischer Scientific, California, USA). Manganese peroxidase activity was determined according to the modified method of Heinfling et al. (1998) by the formation of Mn3+− tartrate (ε238 = 6500 M/cm) from 0.10 mM MnSO4 using 100 mM tartrate buffer (pH 5) and 0.10 mM H2O2. Lignin peroxidase activity was monitored at pH 3.0 according to Tien and Kirk (1988), and the formation of veratraldehyde was monitored at 310 nm (ε310 = 9.3 mM−1 cm−1). Laccase was measured following the oxidation of 2.0 mM 2, 2′-azino-bis (3-ethylbenzthiazoline-6-sulphonic acid, ABTS) at 420 nm (Dias et al. 2004). Hemicellulase activity was measured according to Ghose and Bisaria (1987). The liberated reducing sugars were quantified using dinitrosalicylic acid (DNS) reagent (Miller 1959).
Fig. 1. Lignin isolation and enzyme activity analysis of bio-treated poplar
a FT-IR, Fourier translation infrared spectrum; b GC-MS, gas chromatography-mass spectrometer;c GPC, gel permeation chromatography; d NMR, nuclear magnetic resonance
Determination of Lignin
MWL was isolated and purified according to the method of Björkman (1954). MWL0 was lignin extracted from the untreated sample, and MWL1, MWL2, MWL3, and MWL4 were extracted from poplar treated with T. pubescens C7571 for 4 weeks, 8 weeks, 12 weeks, and 16 weeks, respectively. The lignin degradation products in these four bio-treated poplar samples were extracted with ethyl acetate and labeled L1, L2, L3, and L4 (Fig. 1). The weight average (Mw) and number-average (Mn) molecular weights of the lignin fractions were determined using gel permeation chromatography (GPC; Agilent 1200, California, USA). The sugar concentration in lignin was analyzed using a Dionex high-performance anion exchange chromatography (HPAEC) system (model ICS-3000, California, USA). FT-IR spectra of the lignin fractions were collected using a Thermo Scientific Nicolet iN10 FT-IR Microscope (Thermo Nicolet Corporation, Madison, WI, USA) equipped with a liquid nitrogen-cooled MCT detector. The dried fractions were ground and pelletized using KBr, and the spectra were recorded in the range of 4000 to 700 cm−1 at a 4-cm−1 resolution, with 128 scans per sample. Before data collection, background scanning was performed for correction. NMR spectra were recorded on a Bruker AVIII 400 MHz spectrometer (Saarbrucken, Germany) at 25 °C in DMSO-d6. Analytical Py-GC/MS of the lignin was performed using a CDS Pyroprobe 5200HP pyrolyzer (Chemical Data Systems, California, USA) connected to a Perkin Elmer GC/MS apparatus (Clarus 560, California, USA) equipped with an Elite-35MS capillary column (30 × 0.25 mm inner diameter, 0.25-μm film thickness). Compounds were identified by comparing their mass spectra with the NIST library (Wen et al. 2014).
RESULTS AND DISCUSSION
Enzyme Activities
The enzyme activities of the fungi with different treatment times are presented in Fig. 2. The MnP activity of T. pubescens C7571 developed gradually over the treatment period, showing a maximum value of 0.60 U/mL during week 16. T. versicolor C6915 showed maximum MnP activity (0.43 U/mL) during week 4 and then declined, and it showed lower activity overall than T. pubescens C7571 (Fig. 2(a)). In the case of laccase activity (Fig. 2(b)), the more active producer was T. versicolor C6915, with a maximum value (2.23 U/mL) during week 12, while for T. pubescens C7571, the peak was observed at week 8 (0.69 U/mL). In contrast, LiP activity was higher for T. pubescens C7571 for all treatment times (Fig. 2(c)). Furthermore, ligninase activity was much higher than that of hemicellulase, and the hemicellulase of T. pubescens C7571 showed maximum activity during week 8 (0.03 U/mL), without obvious differences from the other treatment periods (Fig. 2(d)).
The production of ligninolytic enzymes during degradation by white-rot fungi has already been reported by several authors. Using a culture medium containing wheat straw incubated with fungi, Kapich et al. (2005) reported that F. fomentarius had an MnP activity value of 0.17 U/mL and B. adusta had a laccase activity value of 0.22 U/mL. More recently, Arora and Sharma (2010) presented values for laccase and MnP in T. versicolor of approximately 2.40 and 2.00 U/mL, respectively. The differing values reported in the literature were mainly because of the diversity of growth media and fungi strains.
Fig. 2. Enzyme activities of white-rot fungi during the incubation period
a MnP, Manganese peroxidase; b LiP, lignin peroxidase
Lignin Component Changes
The decrease in lignin content differed between the two fungi-treated samples during different periods (Fig. 3). A lignin decrease of 6.6% (from 23.5% to 16.9%) and 5.1% (from 23.5% to 18.4%) was measured in the initial degradation period of T. pubescens C7571 and T. versicolorC6915, respectively. During this treatment period (week 4), LiP was the only enzyme whose activity was higher in T. pubescens C7571 than in T. versicolor C6915 (Fig. 2). This result was explained by the fact that LiP oxidizes non-phenolic lignin (Umezawa et al. 1987). Between week 4 and week 8, the decrease was approximately 2.7% for both types of white-rot fungi. Thus, there was a first phase (0 to 4 weeks) of growth, during which a rapid decrease in lignin was observed, and a second phase in which the values remained fairly stable. Additionally, during this treatment period, nearly all enzyme activity for T. pubescens C7571 was higher than that of T. versicolorC6915, except for that of hemicellulase. Hemicellulases are extremely important in the degradation of lignocellulosic biomass because of their lignin-carbohydrate complexes (LCCs) (Dinis et al. 2009). During week 16, T. pubescens C7571 showed higher activity for all three enzymes during the last treatment period (Fig. 2), thus yielding a low lignin content of 10.4%. A recent study also demonstrated the low enzyme producing capabilities of T. versicolor that laccase and MnP activity was only detectable and no LiP activity was detected (Kuhar et al. 2015). The results of enzyme activities as well as lignin component changes confirmed that the ability of T. versicolor to remove lignin was weaker than T. pubescens C7571. Chemical analyses in other studies have also indicated that this species is a non-selective type and that it removes all types of cell-wall constituents not only lignin (Bari et al. 2015). In summary, lignin degradation varied during different treatment periods; because T. pubescens C7571 showed a higher capacity for lignin degradation, it was considered for more specific studies.
Fig. 3. Changes in lignin content during the incubation period
Chemical Component Analysis and Molecular Weight of Extracted Lignin
To reveal the effect of white-rot fungi treatment on the structural changes of lignin, MWL1, MWL2, MWL3, and MWL4 from T. pubescens C7571-treated poplar were further investigated. The sugar analysis of the extracted lignin fractions is listed in Table 1. The lignin fraction from the untreated sample had a relatively higher amount of sugar (6.49%) than the treated samples. Comparatively, MWL4 obtained from treated samples during the last 16 weeks had the lowest amount of bound sugars (0.97%). This result might be caused by the destruction of linkages between lignin and carbohydrates by the hemicellulase in white-rot fungi (Dinis et al. 2009). In all cases, xylose was the predominant sugar (0.28 to 3.06%), followed by mannose, glucose, and galactose, with a small amount of arabinose. An increase in treatment time from 4 to 16 weeks resulted in a noticeable decrease in xylose content (from 0.92% to 0.28%).
Table 1. Percentage of Neutral Sugars in the Isolated Lignin Fractions
a ND, not detected
The molecular weight distribution of the extracted lignin samples was analyzed using gel permeation chromatography (GPC). Changes in the molecular weights of lignin provide insights to lignin fragmentation and re-condensation reactions during bio-treatment. As shown in Table 2, all lignin fractions possessed narrow molecular weight distributions (Mw/Mn, lower than 1.5). The molecular weight of the MWL0 sample was Mw 3585.6 g/mol and Mn 2451.5 g/mol. When treated for 4 to 12 weeks, the extracted lignin had low molecular weights (2555.3 to 3226.4 g/mol and 1773.0 to 2250.0 g/mol). A significant reduction in molecular weight was observed in the lignin fraction MWL4 (Mw 2555.3 g/mol and Mn 1773.0 g/mol). The decrease in molecular weight indicated that lignin was fragmented during bio-treatment (discussed below).
Table 2. Weight-average (Mw), Number-average (Mn) Molecular Weights, and Polydispersity (Mw/Mn) of the Isolated Lignin Fractions
Fig. 4. FT-IR spectra of the extracted lignin fractions
FT-IR Spectra of the Extracted Lignin
To better understand lignin structural changes, the extracted lignin fractions were characterized by FT-IR spectroscopy (Fig. 4). The spectra of samples from weeks 4, 8, 12, and 16 were rather similar. The band at 1024 cm−1 was indicative of the aromatic C-H in-plane deformation. The strong band at 1219 cm−1 was caused by C-C, C-O, and C=O stretching. Syringyl (S) and condensed guaiacyl (G) absorptions were clearly observed at 1329 cm−1, whereas guaiacyl ring breathing with C=O stretching appeared at 1373 cm−1. The bands at 1420, 1503, and 1591 cm−1corresponding to the aromatic skeletal vibrations, and the C-H deformation combined with the aromatic ring vibration at 1462 cm−1 were present in these five spectra. The absorption at 1662 cm−1 was attributed to the carbonyl stretching in conjugated p-substituted aryl ketones. A wide absorption band at 3452 cm−1 originated from the OH stretching vibration in aromatic and aliphatic OH groups, whereas the bands at 2840, 2887, and 2937 cm−1 arose from the C-H asymmetric and symmetrical vibrations in the methyl and methylene groups, respectively. The lignin degraded by white-rot fungi revealed an intensity at the 1662 cm−1 band assigned to C=O in conjugated carbonyl groups, and untreated lignin showed this absorbance to a lesser extent. Moreover, a strong increasing intensity at the 1688 to 1708 cm−1 band was recorded in degraded lignin, which indicated unconjugated carbonyl groups. The results showed that during the bio-treatment of lignin, white-rot fungi generated conjugated and unconjugated C=O, which suggests that lignin biodegradation involved oxidation.
2D HSQC NMR Analysis of the Extracted Lignin
Two-dimensional 1H-13C NMR (2D NMR) provided important structural information and allowed for the resolution of otherwise overlapping resonances observed in either the 1H or 13C NMR spectra. To understand the detailed structural changes in the lignin fractions, they were characterized by 2D HSQC NMR. The HSQC NMR spectra of lignin showed two regions corresponding to the side chain (Fig. 5(a) and (b)) and aromatic 13C-1H correlations (Fig. 5(c) and (d)). The HSQC spectra of untreated wood lignin and treated lignin were similar, which meant that the main structure of lignin remained the same. In the side-chain regions of the HSQC spectra of these two lignin fractions, cross-signals of methoxyls (δC/δH 56.0/3.75) and side chains in β-O-4′ aryl ether linkages were the most prominent. The Cγ-Hγ correlations in β-O-4′ substructures were observed at δC/δH 72.1/4.85 (structures A, A′, and A′′ (Fig. 6). The Cβ-Hβcorrelations corresponding to the erythro and threo forms of the S-type β-O-4′ substructures were distinguished at δC/δH 86.0/4.10 and 86.0/3.98, respectively. These correlations shifted to δC/δH 83.9/4.29 in structure A linked to the G/H lignin units and γ-acylated β-O-4′ aryl ether substructures (A′ and A′′) linked to the S lignin unit. The Cγ-Hγ correlations in structure A were observed at δC/δH 59.5 to 59.7/3.41 to 3.64. In addition to the β-O-4′ ether substructures, β-β′ (resinol, B) and β-5′ (phenylcoumaran, C) linkages were observed. Strong signals for resinol substructures B were observed with Cγ-Hγ, Cβ-Hβ, and the double Cγ-Hγ correlations at δC/δH 85.0/4.70, 53.8/3.09, and 71.1/4.19/3.84, respectively. Phenyl coumarin substructures C were found in lower amounts. The signals for their Cγ-Hγ and Cβ-Hβ correlations were discovered at δC/δH 87.0/5.49 and 53.1/3.47, respectively, whereas the Cγ-Hγ correlations overlapped with other signals at approximately δC/δH 62.5/3.73. In addition, Cγ-Hγ correlations (at δC/ δH 61.4/4.10) in the p-hydroxycinnamyl alcohol end group (I) and various signals from the associated carbohydrates and polysaccharide from fungi (δC/δH 65.0 to 77.1/2.9 to 4.2) were found in the side-chain regions of the lignin HSQC spectra.
In the aromatic regions of the HSQC spectra, cross-signals from the S- and G-lignin units were observed. The S-lignin units showed a prominent signal for the C2,6-H2,6 correlation at δC/δH 104.0/6.70, whereas the G units showed different correlations for C2-H2, C5-H5, and C6-H6 at δC/δH 111.1/6.99, 115.1/6.78, and 119.1/6.81, respectively. Signals corresponding to the C2,6-H2,6correlations in Cɑ-oxidized S units (S′) (δC/δH 105.3/7.31) were present in all of the HSQC spectra of these two lignin fractions. The correlation for the C2-H2 in oxidized ɑ-ketone structures G′ was observed in the spectra of MWL0, and became stronger in the spectra of MWL4. The C2,6-H2,6 correlations of PB were observed as a strong signal at δC/δH 131.3/7.69. The signal at δC/δH 123.0/7.6 might be assigned to vanillic acid or its analogue (Huang et al. 1993; Ali et al.2011), which was one of the lignin-degraded products (Fig. 7).
Fig. 5. 2D HSQC NMR spectra of the lignin fractions: a) side-chain regions of MWL0; b) aromatic regions of MWL0; c) side-chain regions of MWL4; d) aromatic regions of MWL4; e) changes in lignin side-chain linkage abundances, as percentage of total side-chains (calculated from the 2D-NMR spectra); f) percentage of lignin structural unit changes.
The different structural features among the treated and untreated lignin fractions were quantitatively investigated (Fig. 5(e) and (f)). The percentages of lignin side chains involved in the primary substructures A-C and the lignin units were calculated from the corresponding HSQC spectra (Wen et al. 2013). As expected, the main substructures present in all lignin fractions were the β-O-4′ linked ones (A, A′, and A′′), which ranged from 31.5/100 Ar to 76.4/100 Ar. The β-β′ resinol substructure (B) appeared to be the secondary major substructure, comprising 7.5/100 Ar to 18.0/100 Ar. The phenylcoumaran substructures (C) were calculated to be minor amounts, ranging from 1.9/100 Ar to 5.6/100 Ar. The content of β-O-4′ aryl ether in MWL0 was 76.4/100Ar, and it decreased to 31.5/100 Ar in MWL4. These data suggested that the cleavage of the β-O-4′ aryl ether (depolymerization) is the predominant reaction during bio-treatment. This might be because of the high activity of LiP in T. pubescens C7571, which attacks the β-O-4′ linkage of lignin (Umezawa et al. 1987). In addition to β-O-4′ aryl ether linkage cleavage and carbon-carbon linkage degradation, the S/G ratio in lignin was another prominent structural alteration observed after bio-treatment. The S, G, and PB ratios decreased from 53.1/100 Ar, 46.9/100 Ar, and 4.5/100 Ar to 23.0/100 Ar, 17.9/100 Ar, and 0.6/100 Ar for MWL0 and MWL4, respectively. Interestingly, the data also suggested that the G-type lignin was further degraded than the S-type, as revealed by a slightly elevated S/G ratio in MWL0 (1.13) and MWL0 (1.29). Thus, G-type lignin was more easily degraded. Another possible reason is that the isolated MWL was mainly extracted from the middle lamella of the cell wall (Whiting and Goring 1982), which is more easily penetrated by the hyphae of white-rot fungi. Because G-type lignin has less methoxyl than S-type lignin, the cross-signals of methoxyls (δC/δH 56.0/3.75) of MWL4 were stronger.
Fig. 6. Main substructures of lignin involving different side-chain linkages and aromatic units identified by HSQC: (A) β-O-4′ aryl ether linkages with a free –OH at the γ carbon; (A′) β-O-4′ linkages with a carbonyl group at Cγ; (A′′) β-O-4′ linkages with p-hydroxybenzoated –OH at Cγ; (B) resinol structures formed by β-β′/α-O-γ′/γ-O-α′linkages; (C) phenylcoumaran structures formed by β-5′/α-O-4′ linkages; (G) guaiacyl unit; (G′) oxidized guaiacyl unit with an α-ketone; (S) syringyl unit; (S′) oxidized syringyl unit with a carbonyl group at Cα (phenolic); (I) p-hydroxy-cinnamyl alcohol end group; (PB) p-hydroxybenzoate substructures; (VA) vanillic acid or its analogue
Fig. 7. Possible lignin degradation processes
Fig. 8. FT-IR spectra of the lignin degradation products
FT-IR Analysis of the Extracted Lignin
To identify the chemical groups of the degradation compounds formed during white-rot fungi treatment, the degradation products were analyzed using FT-IR spectroscopy. The wide absorption band at 3452 cm−1 originated from the OH stretching vibration in aromatic and aliphatic OH groups, whereas the bands at 2937, 2887, and 2840 cm−1 were attributed to the C-H asymmetric and symmetrical vibrations in the methyl and methylene groups, respectively. The absorption band at 1708 cm−1 was attributed to the C=O in the conjugated carbonyl groups, and the absorption band at 1658 cm−1 was attributed to the α-C=O. The bands at 1460 cm−1, corresponding to aromatic skeletal vibrations and the C-H deformation, combined with aromatic ring vibration at 1459 cm−1, were present in these four spectra of lignin degradation products. The strong band at 1258 cm−1 was caused by the C-C, C-O, and C=O stretching of the syringyl units. The bands at 1085 (1011) cm−1 and 797 cm−1 were attributed to the polysaccharides of white-rot fungi and hexose. As shown in Fig. 8, high amounts of conjugated and unconjugated C=O functionalities were shown in the spectra of lignin degradation products. Similar results were observed in T. versicolor treated beech wood that unconjugated C=O functionalities increased after fungi treatment (Bari et al. 2015). The FT-IR spectra of lignin degradation products revealed that side chain oxidation was one of the main reactions that occurred during bio-treatment, which was also observed by the aforementioned 2D-HSQC analysis.
GC-MS Analysis of the Lignin Degradation Compounds
Gas chromatography/mass spectrometry (GC-MS) is a convenient tool for the rapid analysis of lignin, and it has been used to characterize different patterns of wood decay by fungi. To obtain more detailed insight into the degradation of lignin during fungal decay, lignin degradation products were subjected to Py-GC/MS analysis. The complete series of Py-GC/MS compounds identified in the degradation products are included in Table 3 and Fig. 9.
According to the chemical structure, the degradation products were divided into five categories: aldehydes, phenols, organic acids, alkanes, and benzenes. In both degraded samples (L2 and L4), debris from S- and G-type units were observed. The two peaks at 11.2 (3,5-dimethyl-benzaldehyde) and 15.6 (2,4-bis(1,1-dimethylethyl) phenol) most likely originated in the S units, whereas the peaks at 12.4 (4-methoxy-benzaldehyde) and 14.4 (vanillin) were attributed to the degradation of the G-type units in lignin. Specifically, vanillin (peak 2, Fig. 9), an important degradation product, was also found in the 2D NMR. Simultaneously, a small increase in the abundance of 1,2-benzenedicarboxylic acid (peak 14, Fig. 9) and other organic acids (peak 4, 13, 15, 16, and 18) were observed, confirming that some oxidative alteration of the lignin side-chains (β-O-4′, β-β′, and β-5′) occurred during white-rot decay, as shown by FT-IR and 2D HSQC NMR. Furthermore, the high abundance of alkanes released upon Py-GC/MS analysis indicates that a ring-opening reaction occurred in the white-rotted wood, and the alkyl chain length was between 12 and 27.
The other benzene-type compounds, including benzene propanol (peak 7), 2-(methylthio)-benzothiazole (peak 11), 1,6-dimethyl-4-(1-methylethyl)-naphthalene (peak 12), 1,2-benzenedicarboxylic acid (peak 14), 3-methyl-n,n-bis(3-methylphenyl)-benzenamine (peak 19), dioctyl phthalate (peak 21), and 1-nitro-2-(1,1,2,2-tetrafluoroethoxy)-benzene (peak 22) may be the derivatives of degradation products. These results confirmed the oxidative alteration of lignin side-chains by the detection of debris from units.
Table 3. Py-GC/MS Analysis of Lignin Degradation Compounds
a NO. represents peaks labeled in Fig. 9
b RT, reaction time
Fig. 9. Py-GC/MS analysis of lignin degradation products
CONCLUSIONS
- T. pubescens C7571 showed higher enzyme activity (particularly LiP) than T. versicolor C6915, thus giving a low lignin content of 10.4%. The cleavage of β-O-4′ aryl ether was the predominant reaction, which may be caused by the high activity of LiP.
- The decrease in lignin molecular weights suggested fragmentation reactions, which was further confirmed by FT-IR spectroscopy. G-type lignin was more easily degraded than S-type lignin, and the degradation products of the G units (e.g., vanillin) were found by 2D HSQC NMR and GC-MS analyses.
- The study of lignin alterations during white-rot fungi treatment could be beneficial for the sustainable production of chemicals, materials, and fuels from renewable plant resources.
ACKNOWLEDGMENTS
This work was supported by Fundamental Research Funds for the Central Universities (No. BLYJ201512), the National Science Fund for Distinguished Young Scholars (31225005), and the Chinese Ministry of Education (113014A).
REFERENCES CITED
Ali, K., Maltese, F., Fortes, A. M., Pais, M. S., Choi, Y. H., and Verpoorte, R. (2011). “Monitoring biochemical changes during grape berry development in Portuguese cultivars by NMR spectroscopy,” Food Chem. 124(4), 1760-1769. DOI: 10.1016/j.foodchem.2010.08.015
Arora, D. S., and Sharma, R. K. (2010). “Ligninolytic fungal laccases and their biotechnological applications,” Appl. Biochem. Biotech. 160(6), 1760-1788. DOI: 10.1007/s12010-009-8676-y
Bari, E., Nazarnezhad, N., Kazemi, S. M., Ghanbary, M.A.T., Mohebby, B., Schmidt, O., and Clausen, C.A. (2015). “Comparison between degradation capabilities of the white rot fungi Pleurotus ostreatus and Trametes versicolor in beech wood,” Int. Biodeter. Biodegr. 104, 231-237. DOI: 10.1016/j.ibiod.2015.03.033
Björkman, A. (1954). “Isolation of lignin from finely divided wood with neutral solvents,” Nature 174, 1057-1058. DOI: 10.1038/1741057a0
Bozell, J. J., O’Lenick, C. J., and Stacy, W. (2011). “Biomass fractionation for the biorefinery: Heteronuclear multiple quantum coherence–nuclear magnetic resonance investigation of lignin isolated from solvent fractionation of switchgrass,” J. Agr. Food Chem. 59(17), 9232-9242. DOI: 10.1021/jf201850b
Dias, A. A., Bezerra, R. M., and Pereira, A. N. (2004). “Activity and elution profile of laccase during biological decolorization and dephenolization of olive mill wastewater,” Bioresour. Technol. 92(1), 7-13. DOI: 10.1016/j.biortech.2003.08.006
Dinis, M. J., Bezerra, R. M. F., Nunes, F., Dias, A. A., Guedes, C. V., Ferreira, L. M. M., Cone, J. W., Marques, G. S. M., Barros, A., and Rodrigues, M. A. M. (2009). “Modification of wheat straw lignin by solid state fermentation with white-rot fungi,” Bioresour. Technol. 100, 4829-4835. DOI: 10.1016/j.biortech.2009.04.036
Elegir, G., Daina, S., Zoia, L., Bestetti, G., and Orlandi, M. (2005). “Laccase mediator system: Oxidation of recalcitrant lignin model structures present in residual kraft lignin,” Enzyme Microb. Tech. 37(3), 340-346. DOI: 10.1016/j.enzmictec.2005.02.017
Ghose, T. K., and Bisaria, V. S. (1987). “Measurement of hemicellulase activities. Part 1: xylanases,” Pure Appl. Chem. 59(12), 1739-1752. DOI: 10.1351/pac198759121739
Heinfling, A., Ruiz-Dueñas, F. J., Martı́nez, M. J., Bergbauer, M., Szewzyk, U., and Martı́nez, A. T. (1998). “A study on reducing substrates of manganese-oxidizing peroxidases from Pleurotus eryngii and Bjerkandera adusta,” Febs Lett. 428(3), 141-146. DOI: 10.1016/S0014-5793(98)00512-2
Hofrichter, M. (2002). “Review: Lignin conversion by manganese peroxidase (MnP),” Enzyme Microb. Tech. 30(4), 454-466. DOI: 10.1016/S0141-0229(01)00528-2
Huang, Z., Dostal, L., and Rosazza, J. P. (1993). “Mechanisms of ferulic acid conversions to vanillic acid and guaiacol by Rhodotorula rubra,” J. Biol. Chem. 268(32), 23954-23958.
Kapich, A. N., Prior, B. A., Lundell, T., and Hatakka, A. (2005). “A rapid method to quantify pro-oxidant activity in cultures of wood-decaying white-rot fungi,” J. Microbiol. Meth. 61, 261-271.DOI: 10.1016/j.mimet.2004.12.010
Kuhar, F., Castiglia, V., and Levin, L. (2015). “Enhancement of laccase production and malachite green decolorization by co-culturing Ganoderma lucidum and Trametes versicolor in solid-state fermentation,” Int. Biodeter. Biodegr. 104, 238-243. DOI: 10.1016/j.ibiod.2015.06.017
Mao, J. Z., Zhang, X., Li, M. F., and Xu, F. (2013). “Effect of biological pretreatment with white-rot fungus Trametes hirsuta C7784 on lignin structure in Carex meyeriana Kunth,” BioResources8(3), 3869-3883. DOI: 10.15376/biores.8.3.3869-3883
Martínez Ferrer, Á. T., Rencoret, J., Nieto Garrido, L., Jiménez-Barbero, J., Gutiérrez Suárez, A., and Río Andrade, J. C. (2011). “Selective lignin and polysaccharide removal in natural fungal decay of wood as evidenced by in situ structural analysis,” Environ. Microbiol. 13(1), 96-107. DOI: 10.1111/j.1462-2920.2010.02312.x
Mate, D. M., Garcia-Ruiz, E., Camarero, S., Shubin, V. V., Falk, M., Shleev, S., Ballesteros, A. O., and Alcalde, M. (2013). “Switching from blue to yellow: Altering the spectral properties of a high redox potential laccase by directed evolution,” Biocatal. Biotransfor. 31(1), 8-21. DOI: 10.3109/10242422.2012.749463
Miller, G. L. (1959). “Use of dinitrosalicylic acid reagent for determination of reducing sugar,” Anal. Chem. 31(3), 426-428. DOI: 10.1021/ac60147a030
Souza-Cruz, P. B. D., Freer, J., Siika-Aho, M., and Ferraz, A. (2004). “Extraction and determination of enzymes produced by Ceriporiopsis subvermispora during biopulping of Pinus taeda wood chips,” Enzyme Microb. Tech. 34(3–4), 228-234. DOI: 10.1016/j.enzmictec.2003.10.005
Tien, M., and Kirk, T. K. (1988). “Lignin peroxidase of Phanerochaete chrysosporium,” Method Enzymol. 161, 238-249. DOI: 10.1016/0076-6879(88)61025-1
Umezawa, T., and Higuchi, T. (1987). “Mechanism of aromatic ring cleavage of β-O-4 lignin substructure models by lignin peroxidase,” Febs Lett. 218(2), 255-260. DOI: 10.1016/0014-5793(87)81057-8
Wen, J. L., Yuan, T. Q., Sun, S. L., Xu, F., and Sun, R. C. (2013). “Understanding the chemical transformations of lignin during ionic liquid pretreatment,” Green Chem. 16(1), 181-190. DOI: 10.1039/c3gc41752b
Wen, J. L., Sun, S. L., Yuan, T. Q., and Sun, R. C. (2014). “Structural elucidation of whole lignin from Eucalyptus based on preswelling and enzymatic hydrolysis,” Green Chem. 17(3),1589-1596. DOI: 10.1039/c4gc01889c
Whiting, P., and Goring, D. A. I. (1982). “Chemical characterization of tissue fractions from the middle lamella and secondary wall of black spruce tracheids,” Wood Sci. Technol. 16(4), 261-267. DOI: 10.1007/BF00353149
Wörmeyer, K., Ingram, T., Saake, B., Brunner, G., and Smirnova, I. (2011). “Comparison of different pretreatment methods for lignocellulosic materials. Part II: Influence of pretreatment on the properties of rye straw lignin,” Bioresource Technol. 102(5), 4157-4164. DOI: 10.1016/j.biortech.2010.11.063
Yelle, D. J., Wei, D., Ralph, J., and Hammel, K. E. (2011). “Multidimensional NMR analysis reveals truncated lignin structures in wood decayed by the brown rot basidiomycete Postia placenta,” Environ. Microbiol. 13(4), 1091-1100. DOI: 10.1111/j.1462-2920.2010.02417.x
Zadrazil, F. (1985). “Screening of fungi for lignin decomposition and conversion of straw into feed,” Angew. Bot. 59, 252-433. DOI: 10.1007/BF02826558
Zhao, Q., and Dixon, R. A. (2011). “Transcriptional networks for lignin biosynthesis: More complex than we thought,” Trends Plant Sci. 16(4), 227-233. DOI: 10.1016/j.tplants.2010.12.005
Article submitted: January 8, 2016; Peer review completed: February 12, 2016; Revised version received and accepted: February 28, 2016; Published: March 14, 2016.
DOI: 10.15376/biores.11.2.3972-3986