A sustainable approach for efficient conversion of lignin into biodiesel accompanied by biological pretreatment of corn straw

Energy Conversion and Management 199 (2019) 111928

Contents lists available at ScienceDirect

Energy Conversion and Management journal homepage: www.elsevier.com/locate/enconman

A sustainable approach for efficient conversion of lignin into biodiesel accompanied by biological pretreatment of corn straw

T

Kai Zhanga, Rong Xua, Abd El-Fatah Abomohrab, Shangxian Xiec, Zhengsheng Yua, Qian Guoa, ⁎ Pu Liua, Liang Penga, Xiangkai Lia, a

Ministry of Education Key Laboratory of Cell Activities and Stress Adaptations, School of Life Science, Lanzhou University, Lanzhou, Gansu 730000, People’s Republic of China b Botany Department, Faculty of Science, Tanta University, 31527 Tanta, Egypt c School of Life Science & Technology, Huazhong University of Science and Technology, Wuhan, Hubei 430074, People’s Republic of China

A R T I C LE I N FO

A B S T R A C T

Keywords: Lignin conversion Microbial lipids Biofuel Pretreatment Lignocelluloses

Lignin is the second-most abundant biopolymer on the Earth and it is hard to valorize without pretreatment due to its inherent heterogeneity and recalcitrance. Nowadays, it is necessary to develop effective innovative methods for lignin degradation and efficient utilization. In the present study, the lignolytic bacterium Mycobacterium smegmatis LZ-K2 was isolated from rotten wood. The isolate showed high lipid production and high efficiency of lignin degradation. The lipid production of LZ-K2 grown in corn straw medium with alkali pretreatment, acid pretreatment, and without chemical pretreatment were 0.083 g/L, 0.069 g/L, and 0.072 g/L, respectively. Fatty acids (C14-C24), especially palmitic acid (C16:0; 38.9%), were also accumulated in the untreated corn straw cultures. Results confirmed that the enzyme system and Fenton reaction are the major pathways for lignin depolymerization. In addition, the presence of a critical lignin-degrading enzymes, other than cellulase and hemicellulase, was revealed by the genome analysis. Moreover, the proteome of LZ-K2 showed enzymes, mainly glucose-methanol-choline (GMC) oxidoreductases, which are involved in the Fenton reaction and β-ketoadipate pathway. Unique enzymes of oleaginous microorganisms, such as acetyl CoA carboxylase, were also identified in LZ-K2. In conclusion, the present work provides a sustainable approach for efficient conversion of lignin into biodiesel with simultaneous biological pretreatment of lignocelluloses.

1. Introduction The shortage of non-renewable fossil fuels has created the need of new energy sources and the sharp increase in biomass conversion to value-added products [1]. Amongst, lignocellulases receive increasing attention as the main agricultural waste [2–4]. They are composed mainly of cellulose, hemicellulose and lignin [5]. The latter is the second-most abundant biopolymer on the Earth after cellulose, with an estimated global production of 5–36 × 108 tons annually [6,7]. Improper disposal of lignin causes significant risk to the environment and the global economy [8]. Enzymatic hydrolysis of lignocelluloses was proved as efficient method to transform cellulose into glucose for biofuel production. Because of its inherent heterogeneity and recalcitrance, lignin impedes the action of lignocellulolytic enzyme during biomass conversion into biofuel [9,10]. Therefore, lignocellulosic biomass conversion typically involves a pretreatment step before biofuel production [11]. As ligninolytic enzymes are usually too large to

⁎

penetrate plant cell wall, the pretreatment of biomass is necessary to remove lignin shield in order to increase the accessibility of cellulose and hemicellulose to the conversion enzymes [12]. Many pretreatment methods including physical (e.g. liquid hot water, and irradiation), chemical (e.g. alkaline, acidic, and inorganic salts), and biological (bacterial, fungal, and enzymatic) have been investigated, individually or combined [3,13–16]. However, high energy consumption and pollutant byproducts are associated with physicochemical pretreatments, which are the advantages of biological pretreatment [17]. Nevertheless, the relative inefficiency of biological methods poses challenges in their industrial applications. In addition, lignin can not be converted into biofuel through fermentation or anaerobic digestion. Therefore, dual application of biological pretreatment with conversion of lignin into biodiesel may be a promising option for biofuel production. The steps of lignin bioconversion include the degradation of the biopolymer and synthesis of bioproducts [18]. Studies on the microbial degradation of lignin have mainly focused on white-rot and brown-rot

Corresponding author. E-mail address: [email protected] (X. Li).

https://doi.org/10.1016/j.enconman.2019.111928 Received 18 June 2019; Received in revised form 7 August 2019; Accepted 8 August 2019 0196-8904/ © 2019 Elsevier Ltd. All rights reserved.

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

fungi, which produce a series of ligninolytic enzymes [19,20]. In these fungi, lignin is modified by Fenton-based oxidation model making it accessible for enzymolysis [21]. Nonetheless, commercial bioconversion methods have not been popularized since the first report in the mid-1980s owing to the difficulty in handling fungi-mediated processes. Along with lignin degradation, recalcitrant fungal biomass is produced, which limits its further utilization [22]. Cellulolytic and sulfate-reducing bacteria have been applied to improve the biofuel and pollutant treatment yields [23,24]. In the recent years, several enzymes and catabolic pathways for lignin degradation have been elucidated in bacteria, such as Sphingobium sp. SYK-6 and Rhodococcus jostii RHA1 [25–29]. As compared with fungi, easier genetic manipulation and higher environmental tolerance allow bacteria to be utilized as a potential alternative for higher lignin degradation [30]. As the annual global lipid demand has reached nearly 22.5 billion liters and due to the global interest in biodiesel production, exploring new feedstocks or conversion pathways is of great interest [31–33]. The high cost of animal fat and the dependence of plant oils on season, freshwater, climate, and land make it necessary to explore new microbial sources [34–36]. Some bacterial cells naturally store triacyl glycerides (TAG), which are the biodiesel precursors.[37,38] As compared with other microbes, bacteria exhibit key advantages for lipid production from lignocelluloses, such as easier scale-up, cost-effectiveness and waste recycling [11,39]. Recently, Rhodococcus opacus strains DSM 1069 and PD630 have been reported to produce lipids by utilizing lignin-model compounds as a sole carbon source [37,40]. The one-step bioconversions of lignin to biodiesel would be more appropriate because of the simplicity of operation and economic feasibility [41]. However, most studies focused on the lignin-model compounds or pretreated lignocelluloses for enhanced lipid production [42]. For an efficient one-step for dual use of bacteria in biological pretreatment and biodiesel production, it is necessary to isolate an appropriate bacterial species with high ability to grow on raw lignocellulosic biomass and high conversion rate of lignin into lipids. In the present study, possibility of a simple biological platform that could convert lignin into lipids without prior pretreatment was examined. The Actinobacterium Mycobacterium smegmatis LZ-K2 was isolated from rotten wood and its lignin-selective degradation capability was examined. After pretreatment, the reduction in biomass components, Fourier transform infrared spectroscopy (FTIR) and scanning electron microscope (SEM) were performed. In addition, the activities of some critical enzymes involved in lignin depolymerization, aromatic compound degradation, and lipid accumulation were measured. Subsequently, lipid accumulation and fatty acid profile of M. smegmatis grown on corn straw, with and without chemical pretreatment, was evaluated. The findings from this study will provide a practical route for biofuel production through biological pretreatment of lignocelluloses with efficient lignin conversion.

Table 1 Characteristics of corn straw used in the present study. Parameters

Value

TS (%) VS (% TS) TC (% VS) TN (% VS) Lignin (dw%) Cellulose (dw%) Hemicellulose (dw%)

97.1 ± 92.1 ± 47.34 0.76 15.2 ± 48.6 ± 26.2 ±

0.1 0.05

3.5 2.1 1.6

0.18 g/L; CaCl2, 0.09 g/L; K2HPO4 3H2O, 0.234 g/L; and KH2PO4, 0.18 g/L supplemented with 2% (w/v) of glucose, alkaline lignin, or corn straw as the carbon source. The main characteristics of the used corn straw are shown in Table 1. The alkaline lignin used in these experiments was purchased from Tokyo Chemical Industry Co., Ltd (Tokyo, Japan). The experiments on lipid accumulation included two stages, namely the adaptation of bacteria to high nitrogen concentration (using 0.07 g/L of (NH4)2SO4 with C/N ratio of 30), followed by lipid accumulation stage under low nitrogen levels (without additional nitrogen source at C/N ratio of 60). 2.2. Compositional analysis Lignin, cellulose and hemicellulose contents of the untreated and pretreated corn straw were measured according to the National Renewable Energy Laboratory’s (NREL) analytical procedures [43]. Total solids (TS) and volatile solids (VS) were measured using the APHA standard method [44]. Total carbon (TC) and total nitrogen (TN) were analyzed with an elemental analyzer (vario EL cube; Elementar Analysensysteme GmbH, Germany) [44]. 2.3. SEM and FTIR The raw corn straw and that pretreated with LZ-K2 were assessed by SEM (S-3400 N, Hitachi, Japan) after sequential freeze-drying for 16 h and metal spraying [45]. The functional groups present in the corn straw from the untreated and treated samples (9 days) were investigated by the FTIR (Nexus 670; Nicolet, USA) with a resolution of 4 cm−1. The spectra were obtained with 32 scans from 4000 to 500 cm−1 [44]. 2.4. Enzymes, Fe3+-reducing activities and hydrogen peroxide concentration assay LZ-K2 cells were grown in the corn straw, alkaline lignin, and glucose media as previously described in section 2.1. At 24-h intervals, 2 mL was withdrawn from the cultures for further analysis. Laccase, lignin peroxidase-like, and manganese-like peroxidase activities were measured by monitoring the transformation of 2,2,-azino-bis (3ethylben-zothiazoline-6-sulfonic acid) (ABTS) at 420 nm, veratryl alcohol at 310 nm, and 2,6-dimethyl phenol (2,6-DMP) at 469 nm, respectively [46,47]. The Fe3+-reducing activity was determined by monitoring the changes in absorbance at 562 nm [48]. The samples were centrifuged (8000 rpm, 4 °C, 2 min) and the supernatants were collected for H2O2 analysis. Hydrogen peroxide assay kits (Beyotime Institute of Biotechnology, Haimen, China) were used to determine the H2O2 concentration.

2. Materials and methods 2.1. Bacterial isolation and identification The strain used in the present study was isolated from rotten wood and deposited in the China Center for Type Culture Collection (CCTCC, AB2018066). The bacterium was identified by Beijing Genomics Institute based on 16S rRNA gene sequencing as is Mycobacterium smegmatis LZ-K2. The draft genome of LZ-K2 was sequenced at Shanghai Majorbio Bio-pharm Technology Co., Ltd (Shanghai, China) using the Illumina MiSeq platform. The data were analyzed on the free Majorbio I-Sanger Cloud Platform (www.i-sanger.com). The Whole Genome Shotgun project has been deposited at DDBJ/ENA/GenBank under the accession SOMM00000000. The strain LZ-K2 was grown at 37 °C under aerobic conditions in mineral salt (MS) medium containing Na2HPO4·12H2O, 0.125 g/L; NaH2PO4, 0.45 g/L; NaCl, 0.9 g/L; (NH4)2SO4, 0.9 g/L; MgSO4·7H2O,

2.5. Chemical pretreatment All experiments were conducted in 500-mL bioreactors with 300 mL/each working volume. For alkali pretreatment, corn straw was reacted with 70 mg of NaOH/g of dry straw at a solid loading of 2% (w/ v) [38]. The dilute-acid pretreatment of corn straw was performed by 2

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

mixing with diluted sulfuric acid to a final concentration of 1.5% (w/w) at a solid loading of 2% (w/v) [49]. The bioreactors were subsequently autoclaved at 121 °C for 1 h, then the pH of the cultures was adjusted to 6.7.

followed by corn straw then alkaline lignin. The degradation rate of lignin, cellulose and hemicellulose at day 8 of growth on corn straw reached 51.17%, 7.43% and 22.4%, respectively (Fig. 2). Fig. 3A shows the FTIR spectra of corn straw before and after pretreatment with Mycobacterium smegmatis LZ-K2. Comparing with the untreated corn straw, the increase in absorption peaks at 1106, 1162, 1253, and 1321 cm−1 were due to the cleavage of the acyl and O–H phenolic groups present in the treated sample [55]. The decrease in the intensities of bands near 1730 cm−1 and 1654 cm−1 indicates that lignin side chains were cleaved. These results can be further verified by SEM results shown in Fig. 3B, where the treated corn straw was fully destroyed by the strain LZ-K2, while the surface of the untreated corn straw remained smooth. The genome of LZ-K2 was sequenced and a total of 6,995,354 bp were detected encompassing 6805 protein coding sequences (CDS) and a GC content of 67.41% (Table S1). Based on the genome analysis, the strain was inferred to possess several key enzymes for lignin degradation, such as glucose-methanol-choline (GMC) oxidoreductases and catalase-peroxidase. Interestingly, genes which have been identified as the crucial ones for lipid accumulation in the most of oleaginous microorganisms were also recorded in the genome of LZ-K2, such as those coding for malic enzyme and acetyl-CoA carboxylase (Table S2). However, several genes that are critical for cellulolytic proteins were not detected, such as exo-1,4-β-glucanase, which can be confirmed in the present study by the low degradation levels (7.43%) of cellulose. In addition, few genes involved in hemicellulose degradation, such as xylose isomerase and glucosidase, were detected (Table S2). Overall, the present results confirmed that LZ-K2 could effectively degrade lignin in the corn straw. As reported in the previous studies, the genus Mycobacterium has been proven to degrade polycyclic aromatic hydrocarbons (PAH) [56]. However, lignin degradation ability of M. smegmatis has not yet been reported. The strain LZ-K2 showed comparable lignin-degradation abilities to the other reported bacteria [46]. The cleavage of chemical bonds, such as β-O-4, indicates that the strain LZ-K2 degrades lignin by mechanisms similar to those reported in bacteria [57]. Based on the FTIR spectral observation, the variations in the band intensities confirm the breakdown of lignin. In addition, the slight change in the intensity of peaks corresponding to carbohydrates implies that hemicellulose and cellulose were partially utilized by the strain [44]. The results of the growth suggested that LZ-K2 tends to utilize lignin as a carbon source. Similarly, it is reported that Ceriporiopsis subvermispora depolymerizes lignin prior to cellulose [58]. In the present study, the reduction pattern of lignin and cellulose contents confirms that lignin was preferentially utilized by LZ-K2, while cellulose was almost undegraded. Due to the mature industrial applications of cellulose, the leftover cellulose might be more suitable for further processing, such as bioethanol production [59]. Therefore, the treatment by the strain LZ-K2 is a promising approach for pretreatment of lignocellulose by lignin disruption to make cellulose more accessible. The absence of glycoside hydrolase family 7(GH7) and abundance of oxidoreductase genes in LZ-K2 are similar to those in C. subvermispora, which was reported to exhibit pronounced lignin degradation enzymes with very little cellulose degrading enzymes [60].

2.6. Measurement of lipid production and composition Total lipids were extracted using sulfuric acid–methanol method [50]. The extracted lipids were directly converted to fatty acid methyl esters (FAMEs) by transesterification using sodium methoxide [51]. The extracted FAMEs were dried in pre-weighed tubes for 3 days. The tubes were then weighed to measure the FAMEs. A Thermo system GC–MS with HP-5MS was employed for the analysis of FAMEs [52]. Samples (2 µL) were injected at split ratio of 20:1 using He as a carrier gas. A 37compound FAME mix diluted in CH2Cl2 at 0.1, 0.25, 0.5, and 1.0 mg/ mL concentrations was used as external standard. 2.7. Proteomics analysis Cultures of M. smegmatis LZ-K2 grown in MS medium with corn straw (1%, w/v) or glucose (1%, w/v) as a carbon source were harvested at 24 h intervals [18]. After filtration through 10 µm membrane to remove corn straw residues, cells were separated by centrifugation (8000 rpm, 20 min, 4 °C). After centrifugation, the supernatant was filtered using 0.2 µm membranes. The cells were sonicated for 15 min and digested overnight with 0.01 mg/mL trypsin at 37 °C. Thereafter, the peptides were desalted, dried by vacuum centrifugation, and resuspended in 0.1% formic acid (FA) for protein analysis [53]. The protein-containing supernatant obtained from cultures of each medium was concentrated at 4 °C and loaded on sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) for electrophoresis. After destaining, each sample lane was excised into 1 mm2 pieces. Gel fragments were washed and destained twice, incubated with acetonitrile (ACN) for dehydration and then with tris-(2-carboxyethyl) phosphine (TCEP) at 56 °C for 1 h, and subsequently cooled to the room temperature. The TCEP was discarded, and the gel was incubated with 40 mmol/L iodoacetamide (IAA) in the dark for 45 min, washed thrice, dried, then incubated with ACN at the room temperature for 15 min. The gel was air dried for 5–10 min and incubated with 0.01 mg/mL trypsin solution at 37 °C for 8–24 h. The digesting solution was transferred to a new tube. The gel was extracted thrice with 50% ACN/5% trifluoroacetic acid (TFA) at 37 °C for 5–15 min. Peptides were collected from the extracted solution, dried by vacuum centrifugation, and resuspended in 0.1% formic acid solution for mass spectrometric analysis [54]. 2.8. Statistical analysis All experiments were performed in triplicates and results were presented as the mean ± SD. The different treatment groups were compared using T-test or ANOVA. Two-tailed P < 0.05 were considered statistically significant. R software version 3.4.4 was used to perform the differential expression of proteins.

3.2. Lignocellulolytic enzymes activities and Fenton reaction 3. Results and discussion Since the genes involved in lignin depolymerization (such as those encoding peroxidase and oxidoreductase) were recorded in the genome (Table S2), the activities of several lignocellulolytic enzymes and the existence of Fenton reaction in the strain were investigated to further confirm its degradation ability (Fig. 4). The maximum activities of laccase-like (Fig. 4A) and lignin peroxidase-like enzymes (Fig. 4B) were observed in the alkaline lignin culture on day 4, with the values of 0.44 ± 0.05 U/mL and 0.35 ± 0.03 U/mL, respectively. However, these enzymes reached their maximum activity levels on day 3 in the corn straw medium, with the values of 0.18 ± 0.07 U/mL and 0.12 ± 0.03 U/mL, respectively. The highest activities of manganese

3.1. Bacterial growth and lignocellulose degradation Comparing of 16S rRNA gene sequences showed that the strains LZK2 was found to be closely related to Mycobacterium smegmatis ATCC 19420 (Fig. 1A). To investigate the potential of lignin-degrading capability of the strain LZ-K2, cells were incubated in media containing glucose, alkaline lignin, corn straw, xylan, or sodium carboxymethylcellulose (CMC) as a carbon source. The growth results revealed that the strain LZ-K2 grows slowly in the CMC medium (Fig. 1B). However, the growth of the strain in glucose showed the highest value 3

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

Fig. 1. Phylogenetic tree of the strain LZ-K2 (A), and growth curve of bacterial strain on glucose (Glu), corn straw (CS), alkaline lignin (ALK), xylan (Xyl) and sodium carboxymethylcellulose (CMC) as a sole carbon source (B).

peroxidase-like enzyme were 0.98 ± 0.05 U/mL on day 5 and 0.52 ± 0.05 U/mL on day 4, respectively, on alkaline lignin and corn straw (Fig. 4C). However, all of these enzyme activities were almost undetectable in the glucose medium. Although the Fe3+-reducing activities in the corn straw and glucose substrates maintained the same pattern of variation, the capacities in the presence of corn straw (0.026 ± 0.0008 A/min) were significantly higher than those in glucose (0.0026 ± 0.0005 A/min) on day 3 (Fig. 4D). Moreover, the maximum H2O2 concentration in corn straw (117.79 ± 6.91 µmol/L) at day 4 was significantly greater than that in the glucose culture (3.16 ± 0.22 µmol/L) (Fig. 4E). All of the detected enzyme activities in alkaline lignin medium were higher than those in corn straw and glucose. It is suggested that the lignin components induced the expression of these enzymes [18]. By comparing the maximum laccase-like activity of LZ-K2 (0.44 ± 0.05 U/mL) with other reported bacteria (~1 U/mL), it is inferred that this enzyme might play a crucial role in lignin oxidation. Moreover, the estimated activities of lignin peroxidase-like

Fig. 2. Cellulose, hemicellulose and lignin degradation rates of Mycobacterium smegmatis LZ-K2 grown on corn straw for 8 days.

4

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

Fig. 3. Changes in FTIR spectra (A) and scanning electron microscope (B) of corn straw (CK) after pretreatment with Mycobacterium smegmatis LZ-K2.

pentadecanoic acid (C15:0), 7-hexadecenoic acid (C16:1), palmitic acid (C16:0), heptadecanoic acid (C17:0), oleic acid (C18:1), octadecanoic acid (C18:0), eicosanoic acid (C20:0), behenic acid (C22:0), and tetracosanoic acid (C24:0), were detected in the FAMEs of LZ-K2 grown in corn straw medium with and without pretreatment (Table 2). In all groups, palmitic acid (C16:0) was the main fatty acid, with its content in the control, dilute acid pretreated, and alkali pretreated groups being 38.93%, 31.55%, and 28.07%, respectively. Thus, alkali pretreatment enhanced the unsaturation of fatty acids which resulted in increase of MUFAs by 81.4% and 116.1% over the untreated and acid-pretreated corn straw, respectively. The comparative proteomic analysis was used to elucidate the potential of lignin degradation and lipid accumulation. A total of 1536 proteins were detected in the comparative proteomic analyses, among which 868 were upregulated and 279 were downregulated. Various proteins involved in lignin degradation were upregulated in the strain, especially catalase-peroxidases, GMC family oxidoreductase and MnSODs (Fig. 6). As expected, the strain LZ-K2 did not express the proteins related to cellulose degradation. These proteomic data were in consistent with the genomic observations. The overexpression of enzymes involved in fatty acids and TAG synthesis in LZK2 is similar to that reported in R. jostii RHA1 [63]. The overexpression enzymes involved in fatty acid and TAG biosynthesis, such as acetyl CoA carboxylase (ACC) and fatty acid synthase (FAS), confirmed that LZ-K2 has the capability of lipid accumulation. Owing to the high degradation rate of lignin accompanied by the

(0.35 ± 0.03 U/mL) and manganese peroxidase-like (0.98 ± 0.05 U/ mL) enzymes were less than that reported by others (~1 U/mL and ~3 U/mL, respectively) [61]. However, the activities of traditional lignin degradation enzymes were all detected in strain LZ-K2. It is indicated that strain LZ-K2 has good lignin degradation ability. Moreover, the Fe3+-reducing activity and H2O2 generation capability were higher than those of P. ananatis Sd-1, which was the first bacterium in which Fenton reaction was observed [46]. During lignin degradation, several antioxidant enzymes and low-molecular weight proteins, such as catalase and thioredoxin, overexpressed by 2–30 times. The overexpression of theses enzymes could protect the cells from oxidative stress due to elevated concentrations of the produced H2O2 [62]. Therefore, it can be confirmed that strain LZ-K2 exhibits a relatively complete lignin degradation system similar to that of fungi. 3.3. Comparison of lipid production and composition The presence of lignin degradation and lipid biosynthesis enzymes suggests that LZ-K2 has the potential to produce lipids. For enhanced practical application, raw corn straw and that pretreated with alkali or acids were investigated as a feedstock for lipid accumulation (Fig. 5). The maximum lipid production of the alkali-pretreated straw (0.083 g/ L at day 8) was higher than those of the control (0.072 g/L) and acidpretreated (0.069 g/L) groups. However, lipid production was not significantly different among these groups. The fatty acids, including 5

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

Fig. 4. Time course of laccase-like activity (A), lignin peroxidase-like activity (B), manganese peroxidase-like activity (C), Fe3+-reducing activity (E) and H2O2 concentration (D) of LZ-K2 in the presence of different substrates including glucose (Glu), alkaline lignin (Alk), and corn straw (CS).

could accumulate lipids up to 0.06–0.12 g/L (Table 3). A high concentration of fatty acids, such as hexadecanoic acid, signifies the potential of biodiesel production. Proteomic analysis indicated that catalase-peroxidases and MnSODs were the most likely critical enzymes based on their relatively high abundance. The overexpression of GMC family oxidoreductase, involved in hydrogen peroxide generation for Fenton chemistry in most of the reported fungi, was observed in the corn straw medium [64]. Moreover, the upregulation of catalase and thioredoxin protects the strain from the oxidative stress and produced toxic compounds [65]. The overexpression of proteins, such as ACC and FAS, asserts the possibility of lipid accumulation in the strain LZ-K2. The similar fatty acid and TAG synthesis pattern in LZ-K2 with R. jostii RHA1 suggested that LZ-K2 could convert aromatic lignocelluloses into bio-oils [63,66]. In the future, avenues for deriving a higher lipid content to ensure the economic feasibility of the process should be explored. However, the present study confirmed the effective conversion of corn straw to lipids by LZ-K2. Compared to the traditional pretreatment, the treatment using LZK2 not only treat lignin as a waste, but also preferentially convert lignin

poor breakdown of cellulose, the lipid production by the strain LZ-K2 could be chiefly attributed to lignin degradation. Although the chemical treatment was more efficient in depolymerization, there were no significant differences between lipid production of corn straw without and with pretreatment. Therefore, it can be hypothesized that the strain LZK2 could convert lignin to lipid, even without chemical pretreatment. The lipid production of the strain (0.072 g/L) from corn straw without pretreatment was comparable to that of Rhodococcus opacus DSM 1069 (0.069 g/L) from kraft lignin with oxygen-pretreatment [37]. The lipid production of Rhodococcus opacus PD630, cultured in kraft lignin medium with laccase- Rhodococcus opacus PD630 co-fermentation reached 0.145 g/L. Comparing lipid content of strain LZ-K2 (0.877 g/g CDW) with the R. opacus PD630 and R. jostii RHA1 VanA− cofermentation (0.39 g/g CDW) showed higher lipid content of LZ-K2, which might be attributed to the higher lignin degradation rate (51.17%). In ethanol organosolv lignin (EOL) and ultrasonicated EOL medium, the lipid content of R. opacus DSM 1069 reached 5.6 × 10−4 g/g EOL and 4.0 × 10−3 g/g EOL, respectively. In pyrolysis light oil fraction from switch grass, R. opacus PD630 and DSM 1069 6

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

Fig. 5. The cellular dry weight, lipid content, and lipid production of Mycobacterium smegmatis LZ-K2 grown on corn straw for 8 days with or without chemical pretreatment.

4. Conclusions

Table 2 Fatty acid composition of the strain LZ-K2 in raw corn straw (CK) and after chemical treatment for 8 days. Fatty acids

CK

Acidic treatment

Alkali treatment

Dodecanoic acid (C12:0) Tetradecanoic acid (C14:0) Pentadecanoic acid (C15:0) 7-Hexadecenoic acid (C16:1) Palmitic acid (C16:0) Heptadecanoic acid (C17:0) Oleic acid (C18:1) Octadecanoic acid (C18:0) Eicosanoic acid (C20:0) Behenic acid (C22:0) Tetracosanoic acid (C24:0) SFAs MUFAs PUFAs

0 5 6.83 8.20 38.93 11.58 5.24 10.67 1.97 3.18 8.40 86.56 13.44 0

0 4.06 4.75 4.96 31.55 7.01 6.32 9.5 4.24 7.14 20.49 88.72 11.28 0

0.32 3.60 0.91 9.71 28.07 2.57 14.67 9.85 3.65 4.38 22.28 75.62 24.38 0

The present study investigated the possibility of a simple biological platform using Actinobacterium Mycobacterium smegmatis LZ-K2 to convert lignin into lipids without prior pretreatment of lignocellulosic biomass. It can be concluded from the present study that;

• The peroxidase system and Fenton reaction are the major pathways

•

to lipid for biodiesel production. The leftover cellulose could be further used for other industrial applications because of the absence of genes of cellulose degradation in the strain LZ-K2. The recorded activities of laccase-like and peroxidase-like enzymes may be due to the expression of catalase-peroxidase and MnSOD. In Sphingibacterium sp. T2, catalaseperoxidase was found to be active in the oxidation of ABTS [67]. The more powerful Fenton reaction and enzymatic activities indicated that LZ-K2 possesses substantial lignin-degradation ability. The overexpression of ACC and the ability to generate acetyl-CoA, which forms the basis for fatty acid synthesis, may share a good correlation with lipid accumulation in the strain [68]. Similar lipid contents were obtained with or without chemical pretreatment in the strain LZ-K2. In general, fungal lipid production was found to be higher than that in LZK2. However, wider range of industrial application, simpler genetic manipulation, and greater adaptability give the bacteria the privilege of practical usage than fungi. The conversion of lignin into lipid by the strain LZ-K2 thus offers a significant opportunity for biodiesel production.

•

responsible for lignin depolymerization. Simultaneously, catalaseperoxidase involvement in lignin degradation is expected, which needs further validation. In addition, several highly expressed genes were annotated as uncharacterized proteins and, therefore, further studies are of great importance to understand their involvement in the lignin degradation and detoxification-stress-responsive mechanisms of strain LZ-K2. The present study identified the strain’s capability for lignin degradation and lipid accumulation for the first time. Moreover, substantial lipid production was obtained without chemical pretreatment of the corn straw. This capacity to convert lignin and lipid accumulation can be applied in the future for efficient biodiesel production from agricultural wastes. Although the technological advance and yield obtained are still far below the requirements for an industrial process, this study provided the theoretical possibility for a novel approach. Further studies are needed to completely understand the metabolic pathways and to identify the enzymes involved in lignin degradation. In addition, microbial lipid production from cheap, readily available, and renewable plant materials need to be explored. The present study will comprehensively help to explore novel biological strategies for the effective conversion of biopolymers without relying on chemical or physical pretreatment.

Declaration of Competing Interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

7

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

Fig. 6. An integrated model elucidating the lignocellulose degradation mechanisms by Mycobacterium smegmatis LZ-K2. CP catalase peroxidase; MnSOD Superoxide dismutase [Mn]; QRD quinone reductase; and GMC GMC oxidoreductase. Table 3 Comparison of lignin degradation and biodiesel production of the present study with other reported studies. Pretreatment

Microorganism

Conditions

Lipids

Lignin degradation rate

References

Biological

Mycobacterium smegmatis LZ-K2

Limited nitrogen source

51.17%

This study

Oxygen-pretreated kraft lignin Laccase

Rhodococcus opacus DSM 1069 Rhodococcus opacus PD630

28% NA

[37] [40]

Biological

R. opacus PD630 and R. jostii RHA1 VanA− R. opacus DSM 1069 R. opacus DSM 1069 R. opacus PD630 and DSM 1069

Limited nitrogen source Laccase – Rhodococcus opacus PD630 cofermentation R. opacus PD630 and R. jostii RHA1 VanAcofermentation Limited nitrogen source

0.0715 g/L (0.877 g/g CDW) 0.043 g/L 0.145 g/L 0.39 g/g CDW

39.6%

[64]

0.004 g/g EOL

23%

[22]

23%

[22]

NA

[51]

Ultrasonicated EOL (us-EOL) Ethanol organosolv lignin (EOL) Pyrolysis light oil fraction from switch grass

−4

g/g EOL

Limited nitrogen source

5.6 × 10

Limited nitrogen source

0.06–0.12 g/L

Acknowledgements This work was supported by National Natural Science Foundation Grant (No: 31870082). 8

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

Appendix A. Supplementary data

[26] Bhatia SK, Kim S-H, Yoon J-J, Yang Y-H. Current status and strategies for second generation biofuel production using microbial systems. Energy Convers Manage 2017;148:1142–56. [27] McLeod MP, Warren RL, Hsiao WW, Araki N, Myhre M, Fernandes C, Miyazawa D, Wong W, Lillquist AL, Wang D, Dosanjh M, Hara H, Petrescu A, Morin RD, Yang G, Stott JM, Schein JE, Shin H, Smailus D, Siddiqui AS, Marra MA, Jones SJ, Holt R, Brinkman FS, Miyauchi K, Fukuda M, Davies JE, Mohn WW, Eltis LD. The complete genome of Rhodococcus sp. RHA1 provides insights into a catabolic powerhouse. PNAS 2006;103:15582–7. [28] Kontur WS, Bingman CA, Olmsted CN, Wassarman DR, Ulbrich A, Gall DL, et al. Novosphingobium aromaticivorans uses a Nu-class glutathione S-transferase as a glutathione lyase in breaking the beta-aryl ether bond of lignin. J Biol Chem 2018;293:4955–68. [29] Lin L, Wang X, Cao L, Xu M. Lignin catabolic pathways reveal unique characteristics of dye-decolorizing peroxidases in Pseudomonas putida. Environ Microbiol 2019;21:1847–63. [30] Wu W, Liu F, Singh S. Toward engineering E. coli with an autoregulatory system for lignin valorization. PNAS 2018(115):2970–5. [31] Abomohra AE-F, Jin W, Tu R, Han S-F, Eid M, Eladel H. Microalgal biomass production as a sustainable feedstock for biodiesel: Current status and perspectives. Renew Sustain Energy Rev 2016;64:596–606. [32] Ma X, Gao Z, Gao M, Ma Y, Ma H, Zhang M, et al. Microbial lipid production from food waste saccharified liquid and the effects of compositions. Energy Convers Manage 2018;172:306–15. [33] Xu R, Zhang K, Liu P, Han H, Zhao S, Kakade A, et al. Lignin depolymerization and utilization by bacteria. Bioresour Technol 2018;269:557–66. [34] Abomohra AE, Shang H, El-Sheekh M, Eladel H, Ebaid R, Wang S, et al. Night illumination using monochromatic light-emitting diodes for enhanced microalgal growth and biodiesel production. Bioresour Technol 2019;288:121514. [35] Abomohra AE, Eladel H, El-Esawi M, Wang S, Wang Q, He Z, et al. Effect of lipidfree microalgal biomass and waste glycerol on growth and lipid production of Scenedesmus obliquus: Innovative waste recycling for extraordinary lipid production. Bioresour Technol 2018;249:992–9. [36] Soccol CR, Dalmas Neto CJ, Soccol VT, Sydney EB, da Costa ESF, Medeiros ABP, et al. Pilot scale biodiesel production from microbial oil of Rhodosporidium toruloides DEBB 5533 using sugarcane juice: Performance in diesel engine and preliminary economic study. Bioresour Technol 2017;223:259–68. [37] Wei Z, Zeng G, Huang F, Kosa M, Huang D, Ragauskas AJ. Bioconversion of oxygenpretreated Kraft lignin to microbial lipid with oleaginous Rhodococcus opacus DSM 1069. Green Chem. 2015;17:2784–9. [38] Linger JG, Vardon DR, Guarnieri MT, Karp EM, Hunsinger GB, Franden MA, et al. Lignin valorization through integrated biological funneling and chemical catalysis. PNAS 2014;111:12013–8. [39] Kumar M, Sundaram S, Gnansounou E, Larroche C, Thakur IS. Carbon dioxide capture, storage and production of biofuel and biomaterials by bacteria: A review. Bioresour Technol 2018;247:1059–68. [40] Zhao C, Xie S, Pu Y, Zhang R, Huang F, Ragauskas AJ, et al. Synergistic enzymatic and microbial lignin conversion. Green Chem 2016;18:1306–12. [41] Sun S, Zhang L, Liu F, Fan X, Sun RC. One-step process of hydrothermal and alkaline treatment of wheat straw for improving the enzymatic saccharification. Biotechnol Biofuels 2018;11:137. [42] Liu ZH, Xie S, Lin F, Jin M, Yuan JS. Combinatorial pretreatment and fermentation optimization enabled a record yield on lignin bioconversion. Biotechnol Biofuels 2018;11:21. [43] Sluiter BHA, Ruiz R, Scarlata C, Sluiter DTJ, Crocker D. Determination of structural carbohydrates and lignin in biomass. Natl Renew Energy Lab 2012. [44] Yao Y, He M, Ren Y, Ma L, Luo Y, Sheng H, et al. Anaerobic digestion of poplar processing residues for methane production after alkaline treatment. Bioresour Technol 2013;134:347–52. [45] Wu W, Chen Y, Faisal S, Khan A, Chen Z, Ling Z, et al. Improving methane production in cow dung and corn straw co-fermentation systems via enhanced degradation of cellulose by cabbage addition. Sci Rep 2016;6:33628. [46] Ma J, Zhang K, Huang M, Hector SB, Liu B, Tong C, et al. Involvement of Fenton chemistry in rice straw degradation by the lignocellulolytic bacterium Pantoea ananatis Sd-1. Biotechnol Biofuels 2016;9:211. [47] Nakagawa Y, Sakamoto Y, Kikuchi S, Sato T, Yano A. A chimeric laccase with hybrid properties of the parental Lentinula edodes laccases. Microbiol Res 2010;165:392–401. [48] Du W, Yu H, Song L, Zhang J, Weng C, Ma F, et al. The promoting effect of byproducts from Irpex lacteus on subsequent enzymatic hydrolysis of bio-pretreated cornstalks. Biotechnol Biofuels 2011;4:37. [49] Xie S, Qin X, Cheng Y, Laskar D, Qiao W, Sun S, et al. Simultaneous conversion of all cell wall components by an oleaginous fungus without chemi-physical pretreatment. Green Chem. 2015;17:1657–67. [50] O'Fallon JV, Busboom JR, Nelson ML, Gaskins CT. A direct method for fatty acid methyl ester synthesis: application to wet meat tissues, oils, and feedstuffs. J Anim Sci 2007;85:1511–21. [51] Wells T, Wei Z, Ragauskas A. Bioconversion of lignocellulosic pretreatment effluent via oleaginous Rhodococcus opacus DSM 1069. Biomass Bioenergy 2015;72:200–5. [52] Wei Z, Zeng G, Kosa M, Huang D, Ragauskas AJ. Pyrolysis oil-based lipid production as biodiesel feedstock by Rhodococcus opacus. Appl Biochem Biotechnol 2015;175:1234–46. [53] Li X, He Y, Zhang L, Xu Z, Ben H, Gaffrey MJ, et al. Discovery of potential pathways for biological conversion of poplar wood into lipids by co-fermentation of Rhodococci strains. Biotechnol Biofuels 2019;12. [54] Ma J, Zhang K, Liao H, Hector SB, Shi X, Li J, et al. Genomic and secretomic insight

Supplementary data to this article can be found online at https:// doi.org/10.1016/j.enconman.2019.111928. References [1] Lal R. World crop residues production and implications of its use as a biofuel. Environ Int 2005;31:575–84. [2] Elsayed M, Abomohra AE, Ai P, Wang D, El-Mashad HM, Zhang Y. Biorefining of rice straw by sequential fermentation and anaerobic digestion for bioethanol and/or biomethane production: Comparison of structural properties and energy output. Bioresour Technol 2018;268:183–9. [3] Elsayed M, Abomohra AE-F, Ai P, Jin K, Fan Q, Zhang Y. Acetogenesis and methanogenesis liquid digestates for pretreatment of rice straw: A holistic approach for efficient biomethane production and nutrient recycling. Energy Convers Manage 2019;195:447–56. [4] Peng J, Abomohra AE-F, Elsayed M, Zhang X, Fan Q, Ai P. Compositional changes of rice straw fibers after pretreatment with diluted acetic acid: Towards enhanced biomethane production. J Cleaner Prod 2019;230:775–82. [5] Chen D, Gao A, Cen K, Zhang J, Cao X, Ma Z. Investigation of biomass torrefaction based on three major components: Hemicellulose, cellulose, and lignin. Energy Convers Manage 2018;169:228–37. [6] Abdelaziz OY, Brink DP, Prothmann J, Ravi K, Sun M, García-Hidalgo J, et al. Biological valorization of low molecular weight lignin. Biotechnol Adv 2016;34:1318–46. [7] Ragauskas AJ, Beckham GT, Biddy MJ, Chandra R, Chen F, Davis MF, et al. Lignin valorization: improving lignin processing in the biorefinery. Science 2014;344:1246843. [8] Zhang T, Cai G, Liu S. Application of lignin-based by-product stabilized silty soil in highway subgrade: A field investigation. J Cleaner Prod 2017;142:4243–57. [9] Li H-Y, Wang B, Wen J-L, Cao X-F, Sun S-N, Sun R-C. Availability of four energy crops assessing by the enzymatic hydrolysis and structural features of lignin before and after hydrothermal treatment. Energy Convers Manage 2018;155:58–67. [10] Varman AM, He L, Follenfant R, Wu W, Wemmer S, Wrobel SA, et al. Decoding how a soil bacterium extracts building blocks and metabolic energy from ligninolysis provides road map for lignin valorization. PNAS 2016;113:E5802–11. [11] Jin M, Slininger PJ, Dien BS, Waghmode S, Moser BR, Orjuela A, et al. Microbial lipid-based lignocellulosic biorefinery: feasibility and challenges. Trends Biotechnol 2015;33:43–54. [12] Yang B, Wyman CE. Pretreatment: the key to unlocking low-cost cellulosic ethanol. Biofuels Bioprod Biorefin 2008;2:26–40. [13] Li R, Xie Y, Yang T, Li B, Wang W, Kai X. Effects of Chemical-Biological pretreatment of corn stalks on the bio-oils produced by hydrothermal liquefaction. Energy Convers Manage 2015;93:23–30. [14] Wang S, Li F, Zhang P, Jin S, Tao X, Tang X, et al. Ultrasound assisted alkaline pretreatment to enhance enzymatic saccharification of grass clipping. Energy Convers Manage 2017;149:409–15. [15] Tsegaye B, Balomajumder C, Roy P. Optimization of microwave and NaOH pretreatments of wheat straw for enhancing biofuel yield. Energy Convers Manage 2019;186:82–92. [16] Wei W, Zhang H, Jin Y. Comparison of microwave-assisted zinc chloride hydrate and alkali pretreatments for enhancing eucalyptus enzymatic saccharification. Energy Convers Manage 2019;186:42–50. [17] Sindhu R, Binod P, Pandey A. Biological pretreatment of lignocellulosic biomass–An overview. Bioresour Technol 2016;199:76–82. [18] Lin L, Cheng Y, Pu Y, Sun S, Li X, Jin M, et al. Systems biology-guided biodesign of consolidated lignin conversion. Green Chem 2016;18:5536–47. [19] Bilal M, Adeel M, Rasheed T, Zhao Y, Iqbal HMN. Emerging contaminants of high concern and their enzyme-assisted biodegradation – A review. Environ Int 2019;124:336–53. [20] Kamimura N, Sakamoto S, Mitsuda N, Masai E, Kajita S. Advances in microbial lignin degradation and its applications. Curr Opin Biotechnol 2019;56:179–86. [21] Daniel DF, Eastwood C, Binder Manfred, Andrzej Majcherczyk AA, Schneider Patrick, Asiegbu Fred O, Baker Scott E, Kerrie Barry MB, Bendiksby Mika, Coutinho Pedro M, Cullen Dan, de Ronald P, Vries BG Allen, Gathman Bernard Henrissat, Katarina Ihrmark AK, Kauserud Hävard, LaButti Kurt, Lapidus Alla, Lavin José L, Yong-Hwan Lee EL, Lilly Walt, Lucas Susan, Emmanuelle Morin JAO, Murat Claude, Park Jongsun, Pisabarro Antonio G, Robert Riley AS, Rosling Anna, Schmidt Olaf, Schmutz Jeremy, Skrede Inger, Jan Stenlid XXAd, Wiebenga Ursula Kües, Hibbett David S, Dirk Hoffmeister FM, Högberg Nils, Grigoriev Igor V, Watkinson Sarah C. The plant cell wall–decomposing machinery underlies the functional diversity of forest fungi. Science 2011;333. [22] Kosa M, Ragauskas AJ. Lignin to lipid bioconversion by oleaginous Rhodococci. Green Chem 2013;15:2070. [23] Di Donato P, Finore I, Poli A, Nicolaus B, Lama L. The production of second generation bioethanol: The biotechnology potential of thermophilic bacteria. J Cleaner Prod 2019;233:1410–7. [24] Li X, Lan S-M, Zhu Z-P, Zhang C, Zeng G-M, Liu Y-G, et al. The bioenergetics mechanisms and applications of sulfate-reducing bacteria in remediation of pollutants in drainage: A review. Ecotoxicol Environ Saf 2018;158:162–70. [25] Masai E, Katayama Y, Fukuda M. Genetic and Biochemical Investigations on Bacterial Catabolic Pathways for Lignin-Derived Aromatic Compounds. Biosci Biotechnol Biochem 2014;71:1–15.

9

Energy Conversion and Management 199 (2019) 111928

K. Zhang, et al.

[55] [56]

[57]

[58] [59]

[60]

into lignocellulolytic system of an endophytic bacterium Pantoea ananatis Sd-1. Biotechnol Biofuels 2016;9:25. Bugg TD, Rahmanpour R. Enzymatic conversion of lignin into renewable chemicals. Curr Opin Chem Biol 2015;29:10–7. Pagnout C, Frache G, Poupin P, Maunit B, Muller JF, Ferard JF. Isolation and characterization of a gene cluster involved in PAH degradation in Mycobacterium sp. strain SNP11: expression in Mycobacterium smegmatis mc(2)155. Res Microbiol 2007;158:175–86. Salvachúa D, Katahira R, Cleveland NS, Khanna P, Resch MG, Black BA, et al. Lignin depolymerization by fungal secretomes and a microbial sink. Green Chem 2016;18:6046–62. Chen Z, Wan C. Biological valorization strategies for converting lignin into fuels and chemicals. Renew Sustain Energy Rev 2017;73:610–21. Ko JK, Lee SM. Advances in cellulosic conversion to fuels: engineering yeasts for cellulosic bioethanol and biodiesel production. Curr Opin Biotechnol 2018;50:72–80. Elena Fernandez-Fueyo FJR-D, Ferreira Patricia, Floudas Dimitrios, Hibbett David S, Paulo Canessa LFL, James Tim Y, Seelenfreund Daniela, Lobos Sergio, Polanco Rubén, Mario Tello TW, Honda Yoichi, Watanabe Takashi, Ryu Jae San, Kubicek Christian P, Monika Schmoll KEH, Gaskell Jill, Franz J St, John Amber Vanden, Sandra Grzegorz SabatWymelenberg KS, BonDurant Splinter, Yadav Jagjit S, Harshavardhan Doddapaneni JL, Subramanian Venkataramanan, Oguiza José A, Perez Gumer, Antonio G, Pisabarro Lucia Ramirez, Francisco Santoyo EM, Coutinho Pedro M, Henrissat Bernard, Lombard Vincent, Magnuson Jon Karl, Ursula Kües CH, Igarashi Kiyohiko, Samejima Masahiro, Held Benjamin W, Barry Kerrie W, Kurt AL, LaButti M, Lindquist Erika A, Lucas Susan M, Riley Robert, Salamov Asaf A, Dirk Hoffmeister DS, Hadar Yitzhak, Yarden Oded, de Vries Ronald P, Ad Wiebenga DE,

[61]

[62]

[63]

[64]

[65]

[66] [67]

[68]

10

Stenliddd Jan, Grigorievz Igor V, Berkaff Randy M, Blanchettey Robert A, Phil Kerstenm RV, Martinez Angel T, Cullen Dan. Comparative genomics of Ceriporiopsis subvermispora and Phanerochaete chrysosporium provide insight into selective ligninolysis. PNAS 2012;109:8352. Chen Y, Chai L, Tang C, Yang Z, Zheng Y, Shi Y, et al. lignin biodegradation by Novosphingobium sp. B-7 and analysis of the degradation process. Bioresour Technol 2012;123:682–5. Abucayon E, Ke N, Cornut R, Patelunas A, Miller D, Nishiguchi MK, et al. Investigating catalase activity through hydrogen peroxide decomposition by bacteria biofilms in real time using scanning electrochemical microscopy. Anal Chem 2014;86:498–505. Davila Costa JS, Herrero OM, Alvarez HM, Leichert L. Label-free and redox proteomic analyses of the triacylglycerol-accumulating Rhodococcus jostii RHA1. Microbiology 2015;161:593–610. Kersten P, Cullen D. Extracellular oxidative systems of the lignin-degrading Basidiomycete Phanerochaete chrysosporium. Fung Genet Biol FG & B 2007;44:77–87. Olson KR, Gao Y, DeLeon ER, Arif M, Arif F, Arora N, et al. Catalase as a sulfidesulfur oxido-reductase: An ancient (and modern?) regulator of reactive sulfur species (RSS). Redox Biol 2017;12:325–39. Wells Jr. T, Ragauskas AJ. Biotechnological opportunities with the beta-ketoadipate pathway. Trends Biotechnol 2012;30:627–37. Brown ME, Walker MC, Nakashige TG, Iavarone AT, Chang MCY. Discovery and characterization of heme enzymes from unsequenced bacteria: application to microbial lignin degradation. J Am Chem Soc 2011;133:18006–9. Ratledge C. Regulation of lipid accumulation in oleaginous micro-organisms. Biochem Soc Trans 2002;30:1047–50.