Lipid Production from Dilute Alkali Corn Stover Lignin by

Research Article pubs.acs.org/journal/ascecg

Lipid Production from Dilute Alkali Corn Stover Lignin by Rhodococcus Strains Yucai He, Xiaolu Li, Haoxi Ben,# Xiaoyun Xue, and Bin Yang*

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Bioproducts, Sciences and Engineering Laboratory, Department of Biological Systems Engineering, Washington State University, 2710 Crimson Way, Richland, Washington 99354, United States

ABSTRACT: Biotransformation of lignin to lipids is challenging due to lignin’s recalcitrant nature as a phenolic heteropolymer with a nonuniform structure that imparts rigidity and recalcitrance of plant cell walls. In this study, wild and engineered Rhodococcus strains (R. opacus PD630 and R. jostii RHA1 VanA−) with lignin degradation and/or lipid biosynthesis capacities were selected to establish fundamental understanding of the pathways and functional modules necessary to enable a platform for biological conversion of biomass-derived lignin to lipids. Degradation of lignin (39.6%, dry weight) was achieved by performing cofermentation with wild type R. opacus PD630 and engineered R. jostii RHA1 VanA−. Co-fermentation of these two strains produced higher lipids yield than single strain fermentation. Profiles of metabolites produced by the Rhodococcus strains while growing on alkali technical lignin suggested that lignin was depolymerized to reactive intermediates, such as vanillin, 2,3-dihydrobenzofuran, 2-methoxy-4-vinylphenol, and 3-hydroxy-4-methoxy-benzaldehyde, for lipid biosynthesis. Additionally, fatty acids (C13−C24), especially palmitic acid (C16:0; 35.8%) and oleic acid (C18:1; 47.9%), were accumulated in cells of R. opacus PD630 and R. jostii RHA1 VanA− with lignin as the sole carbon source. Results suggest that the cofermentation strategy can depolymerize lignin into aromatics and promote the lipid production. The lipids produced during cofermentation of lignin by R. opacus PD630 and R. jostii RHA1 VanA− showed promising potential in biofuel production. KEYWORDS: Lignin, Lipid, Rhodococcus opacus PD630, Rhodococcus jostii RHA1 VanA−, Cofermentation

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INTRODUCTION

utilization is that carbon is trapped in the recalcitrant structure. A number of acid-catalyzed, base-catalyzed, noble metal-catalyzed, ionic liquids-assisted, and supercritical fluids-assisted processes have been used for lignin depolymerization at high temperature and/or pressure with various levels of success.6 Recently, biodegradation of lignin has attracted attention due to its potential for production of second-generation biofuels and other valuable aromatics under mild conditions.5 However, biotransformation of lignin to biofuel is challenging due to its recalcitrant nature as a phenolic heteropolymer with a nonuniform structure that imparts rigidity and recalcitrance of plant cell walls.4 Oxidative enzymes secreted by some fungi and bacteria primarily carry out biodepolymerization of lignin. White-rot fungi are major lignin degraders in nature.

Despite the recent advances in bioprocessing carbohydrates in lignocellulosics, the utilization of lignin for fungible fuels or chemicals has yet to be achieved.1 Lignin utilization is a major factor in reducing cost, minimizing carbon emissions, and maximizing sustainability of lignocellulosic biofuels.2,3 Although the current cellulosic ethanol platform replaces the fossil fuel gasoline and biodiesel from fatty acid methyl esters (FAMEs), a large-scale and robust platform for biomass-derived biofuel is largely lacking.1,4−6 Lignin, the second most abundant organic polymer on earth after cellulose, is an energy-dense, heterogeneous polymer that is comprised of phenylpropanoid monomers and used by plants for structure, water transport, and defense.1 The utilization of lignin in biorefinery waste streams as feedstock for value-adding chemicals, materials, and biofuels represents a unique opportunity to improve cost competitiveness as well as carbon and energy efficiency of biorefineries. The major challenge of lignin © 2017 American Chemical Society

Received: October 31, 2016 Revised: December 20, 2016 Published: January 9, 2017 2302

DOI: 10.1021/acssuschemeng.6b02627 ACS Sustainable Chem. Eng. 2017, 5, 2302−2311

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ACS Sustainable Chemistry & Engineering

(VanA) deletion mutant strain R. jostii RHA1 VanA−, which encoded a vanillin-degrading enzyme,19 was able to metabolize lignin, and the accumulated vanillic acid could be utilized as a carbon source by R. opacus PD630.12,14 Moreover, R. opacus PD630 could degrade vanillic acid to lipids via the β-ketoadipate pathway.7,12 During the biodegradation of lignin, loss of β-O-4 bonds and decrease in the molecular weight of lignin to monomers by using extracellular radical molecules could cause repolymerization due to low biodegradation ability.21 Unlike cellulose with β-1,4-glucosidic link as the chemical bond and glucose as the monomer, lignin contains various aromatic monomers and diverse types of chemical bonds or interunit linkages. Thus, it needs an appropriate redox reaction strategy to cleave these bonds thus further complicates the depolymerization process.21−23 Since cofermentation with different strains has been employed for synergetic bioconversion of various substrates to enhance degradation rate, shorten degradation time, and improve product yield, cofermentation of R. opacus PD630 and R. jostii RHA1 VanA− is a potentially effective approach to degrade lignin and accumulate lipids.24 In this study, the wild type and engineered Rhodococcus strains (R. opacus PD630 and R. jostii RHA1 VanA−) with lignin degradation and/or lipid biosynthesis capacities were employed to establish the functional modules that enabled fundamental insights in synergetic pathways of biological conversion of lignin to lipids.

White-rot fungi (Phanerochaete chrysosporium, Phlebia radiata, Trametes versicolor, etc.) can degrade lignin either selectively or nonselectively through oxidases.7,8 Lignin peroxidase (LiP), manganese peroxidase (MnP), and laccase are extracellular lignin degradation enzymes of white-rot fungi. LiP executes the H2O2-dependent Ca−Cß cleavage of lignin model compounds, including nonphenolic syringyl and biphenyl model compounds, and subsequently catalyzes the partial depolymerization of methylated lignin in vitro.9 Although MnP is not effective in preventing nonphenolic lignin oxidation,8,9 it can oxidize Mn2+ to chelated Mn3+ using H2O2 as oxidant on phenolic or nonphenolic lignin units. Laccase belongs to the family of blue multicopper oxidases that catalyze the oneelectron oxidation of aromaticamines, phenolics, and other electron-rich substrates via the reduction of O2 to H2O. MnP and laccase can be produced by almost all white-rot fungi, but only some fungi produce LiP. MnP plays an important role in depolymerizing lignin and chlorolignin as well as in lignin demethylation and pulp bleaching. Although white-rot fungi were extensively studied because of their powerful lignindegrading enzymatic systems,9 they are not very stable in practical biological treatment under environmental and substrate conditions such as at high pH value, O2 limitation, and high extractive and lignin concentration.10 Additionally, lignin biodegradation with white-rot fungi has a limited application as recalcitrant fungal cell material is concomitantly produced.4 Compared with lignin degrading fungi, bacteria could have more promising application potential for the degradation of lignin and its derivatives because of their immense biochemical versatility and environmental adaptability.11 Bacterial lignin depolymerization enzymes were studied to understand the fundamentals of lignin metabolism and depolymerization process.11 It is well-known that some Rhodococcus strains have the pathways for oxidizing ring opening of central aromatic intermediates via the β-ketoadipate pathway4,12 and enabling shuttling of aromatic-derived carbon into central carbon metabolism via the tricarboxylic acid (TCA) cycle. These pathways enable significant versatility for microbes to convert lignin-derived aromatic molecules as well as xenobiotic aromatic species to carbon and energy sources. It was reported that C. echinulata FR3 could accumulate a high level of lipids by biodegradation of all components of the plant cell wall, including cellulose, hemicellulose and lignin.13 R. opacus DSM 1069 and PD630 can degrade aromatic compounds, including phenol, benzoate, vanillin, 4-hydroxybenzoic acid, vanillic acid and syringic acid, via the β-ketoadipate pathway.7,12,14 R. opacus PD630 reportedly utilizes acetate, phenylacetic acid, gluconate, aromatics, and long-chain-length alkanes to produce lipids.15 Several studies showed that R. opacus DSM 1069 and PD630 were able to degrade lignin through β-ketoadipate pathway.4 Ethanol-organosolv-lignin and its ultrasonicated product were good carbon sources for R. opacus DSM 1069, and lipids were accumulated up to 4% (based on cell dry weight).4 Laccase from Trametes versicolor (catalog no. 51639) and cells of R. opacus PD630 could synergize to degrade lignin, and the cell−laccase fermentation led to a 17-fold increase of lipids production.16 Furthermore, R. jostii RHA1 with a high proportion of oxidative genes in its genome and its mutant VanA− were also reported as efficient degraders of lignin and lignin-like compounds.17−20 R. jostii RHA1 was reported to transform lignin into a number of monocyclic phenolic compounds.18 Its ferredoxin oxygenase

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MATERIALS AND METHODS

Strains. R. opacus PD630 and R. jostii RHA1 VanA− were kindly provided by Dr. Joshua Yuan (Texas A&M University, US) and Dr. Lindsay Eltis (University of British Columbia, Canada), respectively. Alkali-Extracted Lignin Preparation. Corn stover provided by National Renewable Energy Laboratory was extracted by 0.1 M NaOH at 80 °C for 2 h to obtain lignin material, which consists of 20% glucose, 11% xylose, 3% arabinose, 2% galactose, 53% lignin, and 11% ash. To further separate the carbohydrates from this lignin-rich material, it was soaked in 0.1 M NaOH solution until it reached pH 12.5. Solubilized lignin was then filtered through 11 μm pore size Whatman filters. Then the filtrate was solidified again by decreasing pH to 3.0 with 2 M H2SO4. The precipitated lignin was filtrated and washed twice with 70 °C deionized water. Finally, the resulting lignin solids were freeze-dried for 3 days in a freeze drier (VirTis, Warminster, PA). The lignin solids obtained contain no glucose, xylose, arabinose, or galactose according to the results of high performance liquid chromatography (HPLC) analysis.25−27 This final product, the alkali-extracted lignin, was used as the substrate in all experiments in this work. Ball Milled Lignin. Ball milled lignin isolated from corn stover biomass was obtained based on the previously reported procedure.27 Fermentation Conditions. The seed culture was prepared by inoculating a single colony of Rhodococcus strain (R. opacus PD630 or R. jostii RHA1 VanA−) into 20 mL Tryptic Soy Broth (TSB) medium and cultivated at 30 °C to OD600∼1.5. The cultured cells were harvested by centrifuging and washing twice with 20 mL 0.85% (w/v) NaCl and then resuspended in 20 mL 0.85% (w/v) NaCl. 5 mL of R. opacus PD630, and/or R. jostii RHA1 VanA− cells were added to 100 mL of lignin fermentation medium (RM minimum medium). The Rhodococcus minimal (RM) medium contains (per liter): 1.4 g (NH4)2SO4, 1.0 g MgSO4·7H2O, 0.015 g CaCl2·2H2O, 1.0 mL sterile trace element solution, 1.0 mL of sterile stock A solution, and 35.2 mL of sterile 1.0 M phosphate buffer at pH 7.0. The trace element solution contains (per liter): 0.050 g CoCl2·6 H2O, 0.0050 g CuCl2·2H2O, 0.25 g EDTA, 0.50 g FeSO4·7H2O, 0.015 g H3BO3, 0.020 g MnSO4·H2O, 0.010 g NiC12·6 H2O, and 0.40 g ZnSO4·7H2O. Stock A solution contains 5.0 g/L FeNa-EDTA and 2.0 g/L NaMoO4·H2O.16 The alkali-extracted lignin was added in the RM medium as sole carbon source. The fermentation was carried out in a 250 mL flask at 180 rpm 2303


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ACS Sustainable Chemistry & Engineering in a shaker at 30 °C for 7 days. Here, 0.5 mL of the fermentation broth was taken in each time interval for further analysis. Microbial Growth Analysis. For this, 0.5 mL samples were taken from the fermentation flask and mixed with 0.5 mL of Trypan Blue Solution (50% trypan blue, 50% 0.1 M phosphate buffer saline (PBS) solution at pH 7.0). Then, 10 μL of the mixture was loaded to a hemocytometer and viable cells were counted (cell/mL) under a fluorescence light microscope (NIKON LABOPHOT 2, Nikon, Japan). OD600 of cells was analyzed with an UV/vis spectrophotometer (UV-2550PC, Shimadzu, Japan). Analysis of Soluble Compounds in Fermentation Broth. The degradation metabolites of lignin in the fermentation broth were determined by gas chromatography−mass spectroscopy (GC-MS). A 30 mL portion of ethyl acetate was added to 15 mL of fermentation broth in a 50 mL centrifuge tube and was vortexed for 5 min at room temperature. The ethyl acetate layer was collected and vortexed with 20 mL of ethyl acetate for 5 min in a centrifuge tube. The collected ethyl acetate layer was placed in a rotary evaporator (Heidolph, Elk Grove Village, IL) for 15 min in a water bath at 30 °C. These samples were resuspended by dissolving in 1.5 mL of ethyl acetate and then transferred to GC-MS autoinjection vials. GC-MS analysis was performed on Ultra GC-DSQ (Thermo Electron, Waltham, MA) using electron impact ionization. Rxi-5 ms was used as the gas chromatographic column (60 m length, 0.25 mm i.d. and 0.25 μm film thickness, Restek, Bellefonte, PA). Helium was used as the carrier gas at a constant flow of 1.5 mL/min. The injection volume was 1 μL and in the splitless mode. The oven temperature was maintained at 50 °C for 5 min and raised to 320 °C at 20 °C/min. Mass spectrometer was operated in full scan mode. Analysis of lignin. Alkali-extracted lignin was characterized by NMR spectroscopy using the 2D 1H−13C HSQC and GPC analysis.28 NMR data were processed with Felix 2007 (FelixNMR, Inc.) or MestreNova 6.0.4 (Mestrelab Research) with matched cosine-bell apodization and sine-bell squared apodization in the indirect dimension, 2X zero filling in both dimensions, and forward linear prediction of 30% more points in the indirect dimension. One-dimensional 1 H spectra were conducted with no apodization or linear prediction and 2X zero filling. For LC, the lignin was dissolved in THF and analyzed using an Agilent 1200 LC with UV detector. A 20 μL sample

was injected after filtration through a 0.45 μm membrane filter. The lignin concentration was analyzed by mixing the dissolved lignin with Prussian blue reagents and the absorbance at 700 nm was detected with an UV/vis spectrophotometer (UV-2550PC, Shimadzu, Japan).16 All the experiments were performed in triplicates. Total lignin degradation was calculated by the equation below:

Lignin degradation (%) ⎛ ⎞ lignin concentration after biodegradation = ⎜1 − ⎟ initial lignin concentration before biodegradation ⎠ ⎝ × 100 Extraction of Total Lipids and Analysis of Fatty Acid Methyl Esters (FAMEs). To quantify the total lipids produced by microbes, 50 mL fermentation broth was centrifuged at 8000g for 5 min. The supernatant was removed and the pellet was resuspended with 50 mL 0.75% NaCl and centrifuged again at 8000g for 5 min. The supernatant was obtained and combined with the previous supernatant. The collected supernatant (150 mL) was centrifuged at 10 000g for 30 min to collect cells. The pelleted cells were lyophilized in a VirTis lyophilizer (The VirTis Co., Inc., Gardiner, NY). A 3 mL chloroform:methanol (2:1, v/v) mixture was added to cells to homogenize the cells with a sealed lid in a shaker at 30 °C and 180 rpm for 3 h, followed by centrifugation at 3000g. The supernatant was transferred to a weighted tube, and 500 μL distilled water was added. After phase separation, the upper phase was discarded. The organic phase containing total lipids was extracted again with a solution containing a mixture of chloroform:methanol:water (3:48:47, v/v/v). The upper aqueous phase was removed and the bottom organic phase was dried under N2 stream, and the resulting lipid was weighted.16 Lipid yield in the cells was calculated as follows:

YL = WL /CDW where WL = total lipid weight (g), CDW= cell dry weight (g). The FAMEs were obtained by the sulfuric acid−methanol method.13 The lipid composition of FAMEs was determined by GC/MS using an Agilent 7890 GC (Agilent Technologies, Santa

Figure 1. 2D HSQC analysis of lignin samples (left two spectra are ball milled lignin and right two spectra are alkali-extracted lignin). 2304


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ACS Sustainable Chemistry & Engineering Clara, CA) coupled with an Agilent 5975 mass spectrometer according to the reported method.13

PD630, respectively. Wild strain R. opacus PD630 and mutant strain R. jostii RHA1 VanA− have different capability to degrade the alkali-extracted lignin.19 Thus, the growths of these two strains were dramatically different. Higher degradation (39.6%) of lignin was obtained by cofermentation with these two strains (Figure 3a). It was reported that R. jostii RHA1 and its mutant

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RESULTS AND DISCUSSION Characterization of Lignin by HSQC NMR Analysis. The HSQC NMR analysis of alkali-extracted lignin provided chemical shift evidence for relevant substructures of lignin, indicating the presence of aromatic p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) units as well as the major lignin linkages (β-O-4, β-β, and β-5). The detailed assignments for the major S, G, and H aromatic C−H bonds and aliphatic C−H bonds in the major lignin linkages are shown in Figure 1. The 2D NMR analysis indicated that compared to the ball milled lignin, the alkali-extracted lignin contains relatively less β-O-4 linkages and less aromatic C−H bonds in all the S, G, and H units. Therefore, the alkali-extracted lignin presented a relatively condensed structure, resulting in some barriers to biochemical degradation. Successful biochemical conversion for this type of lignin indicates a potential universal method to convert various types of lignins, including the waste lignin from biorefinery. Microbial Growth and Lignin Degradation Using Lignin As the Sole Carbon Source. In this study, lignin was used as the sole carbon source for Rhodococcus strains. Microbial growth curves of R. opacus PD630 and R. jostii RHA1 VanA− on lignin as the sole carbon source are shown in Figure 2.

Figure 3. Effects of single fermentation and cofermentation of R. opacus PD630 and R. jostii RHA1 VanA− on lignin degradation rate (a), pH of biodegradation media (b), and cell concentrations (OD600) (c) (initial lignin concentration was 10 g/L; 120 h).

R. jostii RHA1 VanA− with high oxygenase activity could degrade lignin and some aromatic compounds.17 Moreover, R. opacus PD630 could degrade lignin and aromatic compounds to lipids via the β-ketoadipate pathway.7,12 Thus, the improvement of lignin degradation by cofermentation might be explained by the combined action of different ligninolytic enzymes and metabolic pathways of R. jostii RHA1 VanA− and R. opacus PD630 in lignin biodegradation. The pH of the fermentation broth had no significant changes throughout fermentation, remaining at ∼7.0 (Figure 3b). It was also found that cell concentration (OD600) increased with the increase of lignin degradation (Figure 3c). It was reported that the lignin degradation metabolites could be further used as carbon source by Rhodococcus strains.7 Therefore, cofermentation of these two strains (R. jostii RHA1 VanA− and R. opacus PD630) might reduce product inhibition thus promote the growth of bacteria and lignin biodegradation. Another possibility was that these metabolites could be used as carbon source and fed into TCA for better cellular growth. Variation of Major Metabolites with Single Strain Fermentation and Cofermentation on Lignin by GC/MS Assays. To better understand lignin biodegradation, GC/MS was used to determine fermentation metabolites of lignin by single strain fermentation of R. opacus PD630 and R. jostii RHA1 VanA− as well as their cofermentation (Figure 4). Different aromatic compounds were detected in the single fermentation

Figure 2. Microbial growth of R. opacus PD630 and R. jostii RHA1 VanA− on lignin (10 g/L) as the sole carbon source.

Results showed that both R. opacus PD630 and R. jostii RHA1 VanA− successfully grew on the alkali-extracted lignin as sole carbon source. Differences were observed between these two species in terms of their growth rates. R. opacus PD630 grew relatively slower compared with the growth of R. jostii RHA1 VanA−. Additionally, higher maximum cell concentration (7.6 × 105 cell/mL, 96 h) was achieved by R. jostii RHA1 VanA− compared to R. opacus PD630 (5.5 × 105 cell/mL, 120 h). It was also observed that microbial cell concentration of R. jostii RHA1 VanA− remained relatively constant up to 144 h after reaching the maximum value at 96 h. However, cell concentration of R. opacus PD630 significantly decreased after reaching the maximum value at 96 h. Cell concentrations of R. jostii RHA1 VanA− and R. opacus PD630 decreased 11% and 24%, respectively, within the first 24 h of death phase. Furthermore, the depolymerization of lignin was investigated after 5 days of fermentation. Here, 33.6% and 21.2% of lignin were degraded by R. jostii RHA1 VanA− and R. opacus 2305


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Figure 4. Major aromatic compounds variation in R. opacus PD630 single fermentation on lignin at 24, 72, 120, and 168 h, respectively (a). The major aromatic compounds variation in R. jostii RHA1 VanA− single fermentation on lignin 24, 72, 120, and 168 h, respectively (b). The major aromatic compounds variation in cofermentation with R. opacus PD630 and R. jostii RHA1 VanA− on lignin 24, 72, 120, and 168 h, respectively (c).

in reaction intermediates at different time points (Table 3) between the two strains, which indicated that the lignin degradation by R. jostii RHA1 VanA− was a complex process. Only one compound (2-methoxy-4-vinylphenol) appeared in both R. opacus PD630 and R. jostii RHA1 VanA− single fermentation within 72−168 h. This showed the growth of the two bacteria on lignin was very different. GC-MS analysis showed that the cofermentation of R. opacus PD630 and R. jostii RHA1 VanA− on lignin involved 4 kinds of aromatic compounds, including 2,3-dihydro-benzofuran, 2-methoxy-4-vinylphenol, 3-hydroxy-4-methoxy-benzaldehyde, and vanillin (Figure 4c). Three of these compounds, including 2,3-dihydro-benzofuran, 2-methoxy-4-vinylphenol, and vanillin, were detected in all singleand cofermentation. 2,3-Dihydro-benzofuran and 2-methoxy-4vinylphenol could be detected from 24 to 168 h. It was worth

and cofermentation. Five aromatic compounds (vanillin, 2-methoxy-4-vinylphenol, 2-ethoxy-4-anisaldehyde, 2,3-dihydro-benzofuran, 2,3-dimethoxybenzoic acid) were found throughout the single fermentation process of R. opacus PD630 (Figure 4a). Most compounds, including 2,3-dihydrobenzofuran, vanillin, 2-ethoxy-4-anisaldehyde, and 2,3-dimethoxybenzoic acid, appeared in the first 24 h of R. opacus PD630 fermentation. 2-Methoxy-4-vinylphenol could be detected from 24 to 168 h (Table 2). Six aromatic compounds, including vanillin, 6-methoxycoumaran-7-ol-3-one, 2-methoxy4-vinylphenol, 3-hydroxy-4-methoxy-benzaldehyde, 2,3-dihydrobenzofuran, and 3-(3-hydroxyphenyl)-2-propenoic acid, significantly more than those during fermentation with R. opacus PD630, appeared throughout the single fermentation process by R. jostii RHA1 VanA− (Figure 4b). There were big differences 2306


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ACS Sustainable Chemistry & Engineering Table 1. Changes of Vanillin and Its Analogues in the Fermentation Broths of Lignina

a

“+” represents detected; “−” represents undetected.

Table 2. Major Aromatic Compounds Variation in R. opacus PD630 Single Fermentation on Lignina

a


Table 3. Major Aromatic Compound Variation in R. jostii RHA1 VanA− Single Fermentation on Lignina

a


During both single fermentation and cofermentation, vanillin (4-hydroxy-3-methoxybenzaldehyde) and its analogues (e.g., 3-hydroxy-4-methoxy-benzaldehyde, 2-methoxy-4-vinylphenol, and 2,3-dihydro-benzofuran) were detected by GC/MS (Table 1). Vanillin is one of the most important flavor additives in the food industry. It is also used for the production of fragrance,

noting that vanillin and 3-hydroxy-4-methoxy-benzaldehyde only appeared in the cofermentation after 120 and 168 h, respectively (Table 4) although the GC/MS showed their contents were not high. Clearly, the bacterial growth on the lignin under cofermentation conditions was different from those of two single fermentations. 2307


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Table 4. Major Aromatic Compound Variation in Cofermentation with R. opacus PD630 and R. jostii RHA1 VanA− on Lignina

a


pharmaceuticals and other fine chemicals.29 However, the alkali-extracted lignin was not effectively biotransformed to vanillin with R. opacus PD630 and R. jostii RHA1 VanA− singlestrain or cofermentation (Table 1). During R. opacus PD630 fermentation, vanillin only appeared in small amounts at 24 h and disappeared after 72 h. R. jostii RHA1 VanA− produced vanillin between 24h and 120 h. However, this substance only appeared at 120 h in cofermentation. It was reported that R. jostii RHA1 could break down lignin and lignin model compounds by DypB peroxidase and yield vanillin in a considerable amount.18,19 R. jostii RHA1 VanA−, which encoded vanillindegrading enzyme, could further degrade vanillin to vanillic acid to some extent, and it was able to grow on 1 mM vanillic acid but not on 1 mM vanillin.19 Iṅ this study, vanllic acid was produced in R. jostii RHA1 VanA− single fermentation (data not shown). R. opacus PD630 could convert vanillic acid via β-ketoadipate pathway.4,7,12,14 Thus, no vanillic acid was detected in the R. opacus PD630 single fermentation and cofermentation while vanillin and vanillic acid existed in the R. jostii RHA1 VanA− single fermentation. 3-Hydroxy-4-methoxy-benzaldehyde (isovanillin), an isomer of vanillin, is a phenolic aldehyde and a selective inhibitor of aldehyde oxidase. It can be metabolized by aldehyde dehydrogenase into isovanillic acid. In this study, isovanillin appeared at 72 and 168 h in R. jostii RHA1 VanA− single fermentation. However, vanillin was not detected at 72 and 168 h in its single fermentation. Probably, R. jostii RHA1 VanA− could regulate vanillin and isovanillin metabolism. In this study, it presented a very significant peak in GC/MS results. It appeared in all the single-strain fermentation and cofermentation. 2-Methoxy-4-vinylphenol was detected after 72 h of R. jostii RHA1 VanA− single-strain fermentation. Notably, its structure is similar to vanillin. It was reported that a high concentration of vanillin might be toxic to the R. jostii RHA1 and its mutant VanA−.15,18 It was plausible that vanillin could be transformed into two intermediates, 3-hydroxy-4methoxy-benzaldehyde and 2-methoxy-4-vinylphenol. 2,3-Dihydro-benzofuran existed in all single strain and cofermentations but at different time periods and concentrations. In R. opacus PD630 single fermentation, 2,3-dihydro-benzofuran was only detected at 24 h, and none was detected after 72 h. Probably, 2,3-dihydro-benzofuran was bio-oxidized by deoxygenation.30 Clearly, 2,3-dihydro-benzofuran could be formed during lignin utilization by R. jostii RHA1 VanA− single fermentation and cofermentation. As shown in Figure 4 and Table 1, it was found that the single strain fermentation and cofermentation with R. opacus PD630 and R. jostii RHA1 VanA− could effectively degrade lignin and metabolize its degradation metabolites (e.g., vanillin,

3-hydroxy-4-methoxy-benzaldehyde, 2-methoxy-4-vinylphenol, and 2,3-dihydro-benzofuran). Notably, cofermentation could degrade vanillin and its analogues during fermentation of lignin. Variation of Major Aromatics with Cofermentation on Lignin and Glucose by NMR Assays. A further investigation for the cofermentation metabolites of lignin and glucose by NMR was accomplished (Figure 5). Results indicated that, compared to the fermentation metabolites of lignin, the fermentation metabolites of glucose after 24 and 72 h did not have any aromatic protons while glucose was found being consumed. For the fermentation metabolites of lignin after 24 and 72 h, the total aromatic protons (∼6−8 ppm) slightly decreased, which may indicate the consumption of soluble lignin fractions. 2D HSQC NMR was also employed to identify fermentation metabolites of lignin samples, and the methoxylaromatic structure (i.e., guaiacol, vanillin or similar structures), assigned to the peak at ∼3.7 ppm, was found being consumed during the fermentation. There is a sharp peak ∼3.5 ppm in both fermentation metabolites of glucose and lignin after 72 h. It can be assigned to small molecules containing −OCH3 structure such as methanol, methyl formate, or similar structures, which can be the byproducts during the fermentation. The NMR results supported the GC-MS analysis for the fermentation metabolites in Table 1, which indicated that lignin was decomposed by biological conversion. Lipids Accumulated in Cells under Cofermentation with R. opacus PD630 and R. jostii RHA1 VanA−. Recently, oleaginous fungi and bacteria were used for the production of lipids from lignocellulosic materials.4,12,31−39 However, few reported effective biotransformation of lignin to lipids by cofermentation. It was reported that R. opacus PD630 was able to accumulate TAGs up to 76% of the cell dry weight (CDW) when it was incubated in gluconate medium,40 and reached 78 g/L (CDW) composed of 38% TAGs by high cell density batch fermentation using a high concentration of glucose as carbon source.41 Clearly, R. opacus PD630 showed high potential in the biosynthesis of TAGs. To test the lipids accumulated in cells under the cofermentation with R. opacus PD630 and R. jostii RHA1 VanA−, effects of lignin concentration on the lipids accumulation in cells were investigated. As presented in Figure 6a, it was found that high lipids yield was obtained at lignin loading from 0.1 to 10 g/L. The incremental lipid yield of 0.33 g lipid/g CDW was achieved at 0.5 g/L of lignin loading after 5 days. When the lignin loading was 10 g/L, the incremental lipid yield reached 0.29 g lipid/g CDW after 5 days. At over 15 g/L of lignin loading, the incremental lipid yield significantly decreased to 0.15 g lipid/g CDW. In order to load more lignin for accumulating lipids, the appropriate lignin loading in fermentation was 10 g/L. The mutant strain R. jostii 2308


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Figure 5. 1H NMR for assaying the 24 and 72 h fermentation samples from the cofermentation of R. opacus PD630 and R. jostii RHA1 VanA− using lignin (a) and glucose (b) as the carbon source. All the spectra have been normalized by using the same intensity for the H2O peak (∼4.8 ppm).

RHA1 VanA−, which encoded a vanillin-degrading enzyme,19 was able to metabolize lignin, and the accumulated vanillic acid could be utilized as a carbon source by R. opacus PD630.12,14 Moreover, R. opacus PD630 could degrade vanillic acid to lipids via the β-ketoadipate pathway.7,12 Cofermentation with R. jostii RHA1 VanA− and R. opacus PD630 is a potential strategy for degrading lignin into some metabolites that could be used as carbon source and fed into β-ketoadipate pathway and TCA for better cellular growth. Furthermore, the distribution of fatty acids accumulated in cells of R. opacus PD630 and R. jostii RHA1 VanA− grown on lignin (10 g/L) as the sole carbon source was tested. As presented in Figure 6b, the monounsaturated and saturated fatty acids, including tridecanoic acid (C13:0), myristic acid (C14:0), palmitic acid (C16:0), palmitoleic acid (C16:1), oleic acid (C18:1), stearic acid (C18:0), heneicosanoic acid (C21:0), and methyl lignocerate (C24:0), were found in cells of R. opacus PD630 and R. jostii RHA1 VanA− grown on lignin as sole carbon source. Especially, palmitic acid (C16:0; 35.8%) and oleic acid (C18:1; 47.9%) were the major compositions in cell lipids. A high concentration of palmitic acid (C16:0) 35.8% and oleic acid (C18:1) 47.9% in the FAMEs content showed promising potential for biodiesel production. Clearly, the alkaliextracted lignin could be used as a good carbon source for the

biosynthesis of biodiesel in cells through cofermentation of R. opacus PD630 and R. jostii RHA1 VanA−. In addition to C16 and C18 fatty acids, a certain amount of long chain fatty acids, e.g. heneicosanoic acid (C21:0) and methyl lignocerate (C24:0), were produced from lignin by cofermentation with R. opacus PD630 and R. jostii RHA1 VanA−. Therefore, lignin could be effectively utilized and further metabolized into lipids by cofermentation with R. opacus PD630 and R. jostii RHA1 VanA− (Table 1; Figure 4). A synergetic metabolic pathway of lignin bioconversion to lipids was proposed in this study. Probably, R. jostii RHA1 VanA− could degrade lignin to vanillic acid. R. opacus PD630 could convert vanillic acid or its derivatives to produce TAGs7 via the β-ketoadipate pathway.4,7,14,19 In this study, cofermentations by R. opacus PD630 and R. jostii RHA1 VanA− to effectively convert lignin to lipids is successfully demonstrated for the first time.

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CONCLUSION

Both natural and engineered Rhodococcus strains with lignin degradation and/or lipid biosynthesis capacities were selected to establish a fundamental understanding of the pathways and functional modules necessary to enable a platform for biological conversion of lignin to lipids. The Rhodococcus strains have the 2309


Research Article


Environment, Southeast University, Nanjing 210096, P.R. China. ORCID

Bin Yang: 0000-0003-1686-8800 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS This work was supported by U.S. Department of Energy (DOE) award no. DE-EE0006112 with the Bioproducts, Science & Engineering Laboratory and Department of Biological Systems Engineering at Washington State University. This work was performed in part at the William R. Wiley Environmental Molecular Science Laboratory (EMSL), a national scientific user facility sponsored by the U.S. Department of Energy’s Office of Biological and Environmental Research and located at the Pacific Northwest National Laboratory, operated for the Department of Energy by Battelle. The authors would like to thank Dr. Melvin Tucker from the National Renewable Energy Laboratory for kindly providing corn stover for use in this work. Y.H. was partially supported by Jiangsu Government Scholarship for Overseas Studies. We thank Mr. Peiyu Leu, Ms. Marie S. Switab, Dr. Daochen Zhu, and Drs. Hasan Bugra Coban for technical support. We also thank Dr. John Cort who helped us to collect part of NMR data for this project and for insightful discussions.

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Figure 6. Effects of different substrate lignin loading on the incremental lipid yield for cofermentation fermentation of R. opacus PD630 and R. jostii RHA1 VanA− (a). Distribution of fatty acids of the accumulated in cells of PD630 and VanA− grown on lignin as carbon source (b).

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REFERENCES

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pathways for degrading lignin to aromatics and oxidizing ring opening of central aromatic intermediates via the β-ketoadipate pathway, shuttling aromatic-derived carbon into central carbon metabolism via TCA cycle, and accumulating lipids via lipid biosynthetic pathways. Our results suggest that R. jostii RHA1 VanA− could grow with higher cell concentrations using lignin as sole carbon source compared to R. opacus PD630. Furthermore, it was first reported that lignin was successfully utilized in cofermentations by these two Rhodococcus strains. GC-MS and NMR results showed that different lignin aromatics were produced during the single-strain fermentation and cofermentation. Finally, lignin to lipids (with a yield of 0.39 g lipid/g CDW) by cofermentations has been achieved, yet the yield is still low at the current stage. However, the results lead to support for our hypothesis: synthetic reconstruction and balanced modification of key regulators and enzymes in lignin depolymerization, aromatic compound catabolism, lipid biosynthesis, and other relevant processes, will enable Rhodococcus strains to efficiently convert lignin to lipid.

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LIST OF ABBREVIATIONS GC-MS = gas chromatography−mass spectroscopy GPC = gel permeation chromatography HPLC = high performance liquid chromatography FAME = fatty acid methyl ester

AUTHOR INFORMATION

Corresponding Author

*Tel.: 509-372-7640. Fax: 509-372-7690. E-mail: binyang@ tricity.wsu.edu (B.Y.). Present address #

(Haoxi Ben) Key Laboratory of Energy Thermal Conversion and Control of Ministry of Education, School of Energy and 2310


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Lipid Production from Dilute Alkali Corn Stover Lignin by

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