Lipid Production from Dilute Alkali Corn Stover Lignin by

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Lipids Production from Dilute Alkali Corn Stover Lignin by Rhodococcus Strains Yucai He, Xiaolu Li, Haoxi Ben, Xiaoyun Xue, and Bin Yang ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/ acssuschemeng.6b02627 • Publication Date (Web): 09 Jan 2017 Downloaded from http://pubs.acs.org on January 13, 2017

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ACS Sustainable Chemistry & Engineering

Lipids Production from Dilute Alkali Corn Stover Lignin by Rhodococcus Strains Yucai Hea, Xiaolu Lia, Haoxi Bena, Xiaoyun Xuea, and Bin Yanga* a

Bioproducts, Sciences and Engineering Laboratory, Department of Biological Systems

8

Engineering, Washington State University, 2710 Crimson Way, Richland, WA 99354. Tel: 509-

9

372-7640, Fax: 509-372-7690, E-mail: [email protected]

10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25

*The corresponding author is Dr. Bin Yang.

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ACS Paragon Plus Environment


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ABSTRACT: Biotransformation of lignin to lipids is challenging due to lignin’s recalcitrant

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nature as a phenolic heteropolymer with a non-uniform structure that imparts rigidity and

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recalcitrance of plant cell walls. In this study, wild and engineered Rhodococcus strains (R.

30

opacus PD630 and R. jostii RHA1 VanA-) with lignin degradation and/or lipid biosynthesis

31

capacities were selected to establish fundamental understanding of the pathways and functional

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modules necessary to enable a platform for biological conversion of biomass-derived lignin to

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lipids. Degradation of lignin (39.6%, dry weight) was achieved by performing co-fermentation

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with wild type R. opacus PD630 and engineered R. jostii RHA1 VanA-. Co-fermentation of these

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two strains produced higher lipids yield than single strain fermentation. Profiles of metabolites

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produced by the Rhodococcus strains while growing on alkali technical lignin suggested that

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lignin was depolymerized to reactive intermediates, such as vanillin, 2,3-dihydro-benzofuran, 2-

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methoxy-4-vinylphenol, and 3-hydroxy-4-methoxy-benzaldehyde, for lipid biosynthesis.

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Additionally, fatty acids (C13-C24), especially palmitic acid (C16:0; 35.8%) and oleic acid

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(C18:1; 47.9%), were accumulated in cells of R. opacus PD630 and R. jostii RHA1 VanA- with

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lignin as sole carbon source. Results suggest that the co-fermentation strategy can depolymerize

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lignin into aromatics and promote the lipid production. The lipids produced during co-

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fermentation of lignin by R. opacus PD630 and R. jostii RHA1 VanA- showed promising

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potential in biofuel production.

45 46 47

KEYWORDS: Lignin, Lipid, Rhodococcus opacus PD630, Rhodococcus jostii RHA1 VanA-,

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Co-fermentation

49 2


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■ INTRODUCTION

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Despite the recent advances in bioprocessing carbohydrates in lignocellulosics, the utilization

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of lignin for fungible fuels or chemicals has yet to be achieved 1. Lignin utilization is a major

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factor in reducing cost, minimizing carbon emissions, and maximizing sustainability of

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lignocellulosic biofuels2-3. Although the current cellulosic ethanol platform replaces the fossil

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fuel gasoline and biodiesel from fatty acid methyl esters (FAMEs), a large-scale and robust

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platform for biomass-derived biofuel is largely lacking 1, 4-6.

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Lignin, the second most abundant organic polymer on earth after cellulose, is an energy-dense,

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heterogeneous polymer that is comprised of phenylpropanoid monomers and used by plants for

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structure, water transport, and defense 1. The utilization of lignin in biorefinery waste streams as

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feedstock for value-adding chemicals, materials and biofuels represents a unique opportunity to

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improve cost competitiveness as well as carbon and energy efficiency of biorefineries. The major

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challenge of lignin utilization is that carbon is trapped in the recalcitrant structure. A number of

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acid-catalyzed, base-catalyzed, noble metal-catalyzed, ionic liquids-assisted, and supercritical

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fluids-assisted processes have been used for lignin depolymerization at high temperature and/or

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pressure with various levels of success 6. Recently, biodegradation of lignin has attracted

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attention due to its potential for production of second-generation biofuels and other valuable

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aromatics under mild conditions 5.

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However, biotransformation of lignin to biofuel is challenging due to its recalcitrant nature as

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a phenolic heteropolymer with a non-uniform structure that imparts rigidity and recalcitrance of

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plant cell walls 4. Oxidative enzymes secreted by some fungi and bacteria primarily carry out

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bio-depolymerization of lignin. White-rot fungi are major lignin degraders in nature. White-rot

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fungi (Phanerochaete chrysosporium, Phlebia radiata, Trametes versicolor, etc.) can degrade

3



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lignin either selectively or non-selectively through oxidases

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manganese peroxidase (MnP), and laccase are extracellular lignin degradation enzymes of white-

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rot fungi. LiP executes the H2O2-dependent Ca-Cß cleavage of lignin model compounds,

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including nonphenolic syringyl and biphenyl model compounds, and subsequently catalyzes the

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partial depolymerization of methylated lignin in vitro 9. Although MnP is not effective in

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preventing nonphenolic lignin oxidation 8-9, it can oxidize Mn2+ to chelated Mn3+ using H2O2 as

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oxidant on phenolic or non-phenolic lignin units. Laccase belongs to the family of blue multi-

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copper oxidases that catalyze the one-electron oxidation of aromaticamines, phenolics, and other

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electron-rich substrates via the reduction of O2 to H2O. MnP and laccase can be produced by

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almost all white-rot fungi, but only some fungi produce LiP. MnP plays an important role in

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depolymerizing lignin and chlorolignin as well as in lignin demethylation and pulp bleaching.

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Although white-rot fungi were extensively studied because of their powerful lignin-degrading

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enzymatic systems

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environmental and substrate conditions such as at high pH value, O2 limitation, high extractive

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and lignin concentration 10. Additionally, lignin biodegradation with white-rot fungi has a limited

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application as recalcitrant fungal cell material is concomitantly produced 4.

9

. Lignin peroxidase (LiP),

, they are not very stable in practical biological treatment under

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Compared with lignin degrading fungi, bacteria could have more promising application

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potential for the degradation of lignin and its derivatives because of their immense biochemical

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versatility and environmental adaptability

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studied to understand the fundamentals of lignin metabolism and depolymerization process 11. It

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is well-known that some Rhodococcus strains have the pathways for oxidizing ring opening of

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central aromatic intermediates via the β-ketoadipate pathway

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aromatic-derived carbon into central carbon metabolism via the tricarboxylic acid (TCA) cycle.

11

. Bacterial lignin depolymerization enzymes were

4, 12

and enabling shuttling of

4


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These pathways enable significant versatility for microbes to convert lignin-derived aromatic

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molecules as well as xenobiotic aromatic species to carbon and energy sources. It was reported

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that C. echinulata FR3 could accumulate a high level of lipids by biodegradation of all

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components of the plant cell wall, including cellulose, hemicellulose and lignin 13.

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R. opacus DSM 1069 and PD630 can degrade aromatic compounds, including phenol,

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benzoate, vanillin, 4-hydroxybenzoic acid, vanillic acid and syringic acid, via the β-ketoadipate

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pathway

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aromatics, and long-chain-length alkanes to produce lipids

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opacus DSM 1069 and PD630 were able to degrade lignin through β-ketoadipate pathway 4.

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Ethanol-organosolv-lignin and its ultrasonicated product were good carbon sources for R. opacus

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DSM 1069, and lipids were accumulated up to 4% (based on cell dry weight) 4. Laccase from

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Trametes versicolor (catalog# 51639) and cells of R. opacus PD630 could synergize to degrade

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lignin, and the cell–laccase fermentation led to a 17-fold increase of lipids production

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Furthermore, R. jostii RHA1 with a high proportion of oxidative genes in its genome and its

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mutant VanA- were also reported as efficient degraders of lignin and lignin-like compounds 17-20.

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R. jostii RHA1 was reported to transform lignin into a number of monocyclic phenolic

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compounds

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which encoded a vanillin-degrading enzyme

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accumulated vanillic acid could be utilized as a carbon source by R. opacus PD630

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Moreover, R. opacus PD630 could degrade vanillic acid to lipids via the β-ketoadipate pathway 7,

116

12

7, 12, 14

. R. opacus PD630 reportedly utilizes acetate, phenylacetic acid, gluconate,

18

15

. Several studies showed that R.

16

.

. Its ferredoxin oxygenase (VanA) deletion mutant strain R. jostii RHA1 VanA-, 19

, was able to metabolize lignin, and the 12, 14

.

.

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During the biodegradation of lignin, loss of β-O-4 bonds and decrease in the molecular

118

weight of lignin to monomers by using extracellular radical molecules could cause

5



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repolymerization due to low biodegradation ability21. Unlike cellulose with β-1,4-glucosidic link

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as the chemical bond and glucose as the monomer, lignin contains various aromatic monomers

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and diverse types of chemical bonds or interunit linkages. Thus, it needs an appropriate redox

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reaction strategy to cleave these bonds thus further complicates the depolymerization process21-23.

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Since co-fermentation with different strains has been employed for synergetic bioconversion of

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various substrates to enhance degradation rate, shorten degradation time, and improve product

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yield, co-fermentation of R. opacus PD630 and R. jostii RHA1 VanA- is a potentially effective

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approach to degrade lignin and accumulate lipids24. In this study, the wild type and engineered

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Rhodococcus strains (R. opacus PD630 and R. jostii RHA1 VanA-) with lignin degradation

128

and/or lipid biosynthesis capacities were employed to establish the functional modules that

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enabled fundamental insights in synergetic pathways of biological conversion of lignin to lipids.

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■ MATERIALS AND METHODS

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Strains. R. opacus PD630 and R. jostii RHA1 VanA- were kindly provided by Dr. Joshua

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Yuan (Texas A&M University, US) and Dr. Lindsay Eltis (University of British Columbia,

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Canada), respectively.

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Alkali-extracted lignin preparation. Corn stover provided by National Renewable Energy

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Laboratory was extracted by 0.1 M NaOH at 80 oC for 2 h to obtain lignin material, which

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consists of 20% glucose, 11% xylose, 3% arabinose, 2% galactose, 53% lignin, and 11% ash. To

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further separate the carbohydrates from this lignin-rich material, it was soaked in 0.1 M NaOH

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solution until it reached pH 12.5. Solubilized lignin was then filtered through 11 µm pore size

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Whatman filters. Then the filtrate was solidified again by decreasing pH to 3.0 with 2 M H2SO4.

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The precipitated lignin was filtrated and washed twice with 70oC deionized water. Finally, the 6


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resulting lignin solids were freeze-dried for 3 days in a freeze drier (VirTis, Warminster, PA).

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The lignin solids obtained contain no glucose, xylose, arabinose, or galactose according to the

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results of high performance liquid chromatography (HPLC) analysis 25-27. This final product, the

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alkali-extracted lignin, was used as the substrate in all experiments in this work.

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Ball milled lignin: Ball milled lignin isolated from corn stover biomass was obtained based on

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the previously reported procedure 27.

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Fermentation conditions. The seed culture was prepared by inoculating a single colony of

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Rhodococcus strain (R. opacus PD630 or R. jostii RHA1 VanA-) into 20 mL Tryptic Soy Broth

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(TSB) medium, and cultivated at 30°C to OD600 1.5. The cultured cells were harvested by

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centrifuging and washing twice with 20 mL 0.85% (w/v) NaCl and then re-suspended in 20 mL

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0.85% (w/v) NaCl. 5 mL of R. opacus PD630, R. jostii RHA1 and/or R. jostii RHA1 VanA- cells

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were added to 100 mL of lignin fermentation medium (RM minimum medium). The

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Rhodococcus Minimal (RM) medium contains (per L): 1.4 g (NH4)2SO4, 1.0 g MgSO4·7H2O,

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0.015 g CaCl2·2H2O, 1.0 mL sterile trace element solution, 1.0 mL of sterile stock A solution,

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and 35.2 mL of sterile 1.0 M phosphate buffer at pH 7.0. The trace element solution contains

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(per liter): 0.050 g CoCl2·6 H2O, 0.0050 g CuCl2·2H2O, 0.25 g EDTA, 0.50 g FeSO4·7H2O,

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0.015 g H3BO3, 0.020 g MnSO4·H2O, 0.010 g NiC12·6 H2O, and 0.40 g ZnSO4·7H2O. Stock A

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solution contains 5.0 g/L FeNa-EDTA and 2.0 g/L NaMoO4·H2O 16. The alkali- extracted lignin

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was added in the RM medium as sole carbon source. The fermentation was carried out in a 250

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mL flask at 180 rpm in a shaker at 30°C for 7 days. 0.5 mL of the fermentation broth was taken

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in each time intervals for further analysis.

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Microbial growth analysis. 0.5 mL samples were taken from the fermentation flask and 7



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mixed with 0.5 mL of Trypan Blue Solution (50% trypan blue, 50% 0.1 M phosphate buffer

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saline (PBS) solution at pH 7.0). Then, 10 µL of the mixture was loaded to a hemocytometer and

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viable cells were counted (cell/mL) under a fluorescence light microscope (NIKON

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LABOPHOT 2, Nikon, Japan). OD600 of cells was analyzed with an UV/vis spectrophotometer

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(UV-2550PC, Shimadzu, Japan).

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Analysis of soluble compounds in fermentation broth. The degradation metabolites of

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lignin in the fermentation broth were determined by gas chromatography-mass spectroscopy

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(GC-MS). 30 mL of ethyl acetate was added to 15 mL of fermentation broth in a 50 mL

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centrifuge tube and was vortexed for 5 min at room temperature. The ethyl acetate layer was

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collected and vortexed with 20 mL of ethyl acetate for 5 min in a centrifuge tube. The collected

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ethyl acetate layer was placed in a rotary evaporator (Heidolph, Elk Grove Village, IL) for 15

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min in a water bath at 30oC. These samples were re-suspended by dissolving in 1.5 mL of ethyl

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acetate and then transferred to GC-MS auto-injection vials. GC-MS analysis was performed on

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Ultra GC-DSQ (Thermo Electron, Waltham, MA) using electron impact ionization. Rxi-5 ms

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was used as the gas chromatographic column (60 m length, 0.25 mm i.d. and 0.25 µm film

178

thickness, Restek, Bellefonte, PA). Helium was used as the carrier gas at a constant flow of 1.5

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mL/min. The injection volume was 1 µL and in the splitless mode. The oven temperature was

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maintained at 50oC for 5 min and raised to 320 oC at 20 oC/min. Mass spectrometer was operated

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in full scan mode.

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Analysis of lignin. Alkali- extracted lignin was characterized by NMR spectroscopy using the

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2-D 1H-13C HSQC and GPC analysis

28

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(FelixNMR, Inc) or MestreNova 6.0.4 (Mestrelab Research) with matched cosine-bell

. NMR spectra were processed with Felix 2007

8


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apodization and sine-bell squared apodization in the indirect dimension, 2X zero filling in both

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dimensions, and forward linear prediction of 30% more points in the indirect dimension. One-

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dimensional 1H spectra were conducted with no apodization or linear prediction and 2X zero

188

filling. For LC, the lignin was dissolved in THF and analyzed using an Agilent 1200 LC with UV

189

detector. A 20 µL sample was injected after filtration through a 0.45 µm membrane filter. The

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lignin concentration was analyzed by mixing the dissolved lignin with Prussian blue reagents and

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the absorbance at 700 nm was detected with an UV/vis spectrophotometer (UV-2550PC,

192

Shimadzu, Japan) 16. All the experiments were performed in triplicates. Total lignin degradation

193

was calculated by the equation below:

Lignin degradation % = 1 −

Lignin concentration after biodegradation × 100 Initial lignin concentration before biodegradation

194

Extraction of total lipids and analysis of fatty acid methyl esters (FAMEs). To quantify the

195

total lipids produced by microbes, 50 mL fermentation broth was centrifuged at 8000 × g for 5

196

min. The supernatant was removed and the pellet was re-suspended with 50 mL 0.75% NaCl and

197

centrifuged again at 8000 × g for 5 min. The supernatant was obtained and combined with the

198

previous supernatant. The collected supernatant (150 mL) was centrifuged at 10,000 × g for 30

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min to collect cells. The pelleted cells were lyophilized in a VirTis lyophilizer (The VirTis Co.,

200

Inc., Gardiner, NY). 3 mL chloroform:methanol (2:1, v/v) mixture was added to cells to

201

homogenize the cells with a sealed lid in a shaker at 30oC and 180 rpm for 3 h, followed by

202

centrifugation at 3000 × g. The supernatant was transferred to a weighted tube, and 500 µL

203

distilled water was added. After phase separation, the upper phase was discarded. The organic

204

phase containing total lipids was extracted again with a solution containing a mixture of

205

chloroform:methanol:water (3:48:47, v/v/v). The upper aqueous phase was removed and the 9



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bottom organic phase was dried under N2 stream and the resulting lipid was weighted

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yield in the cells was calculated as follows:

208

Y L =W L / DC W

209

where WL = Total lipid weight (g), DCW= Dry cell weight (g).

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. Lipid

13

210

The fatty acid methyl esters (FAMEs) were obtained by the sulfuric acid–methanol method

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The lipid composition of FAMEs was determined by GC/MS using an Agilent 7890 GC (Agilent

212

Technologies, Santa Clara, CA) coupled with an Agilent 5975 mass spectrometer according to

213

the reported method 13.

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■ RESULTS AND DISCUSSION

.

215

Characterization of lignin by HSQC NMR analysis. The HSQC NMR analysis of alkali-

216

extracted lignin provided chemical shift evidence for relevant sub-structures of lignin, indicating

217

the presence of aromatic p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) units as well as the

218

major lignin linkages (β-O-4, β-β, and β-5). The detailed assignments for the major S, G, and H

219

aromatic C-H bonds and aliphatic C-H bonds in the major lignin linkages are shown in Fig. 1.

220

The 2D NMR analysis indicated that compared to the ball milled lignin, the alkali-extracted

221

lignin contains relatively less β-O-4 linkages and less aromatic C-H bonds in all the S, G and H

222

units. Therefore, the alkali-extracted lignin presented a relatively condensed structure, resulting

223

in some barriers to biochemical degradation. Successful biochemical conversion for this type of

224

lignin indicates a potential universal method to convert various types of lignins, including the

225

waste lignin from biorefinery.

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Microbial growth and lignin degradation using lignin as sole carbon source. In this study, 10


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lignin was used as the sole carbon source for Rhodococcus strains. Microbial growth curves of R.

228

opacus PD630 and R. jostii RHA1 VanA- on lignin as the sole carbon source are shown in Fig. 2.

229

Results showed that both R. opacus PD630 and R. jostii RHA1 VanA- successfully grew on the

230

alkali-extracted lignin as sole carbon source. Differences were observed between these two

231

species in terms of their growth rates. R. opacus PD630 grew relatively slower compared with

232

the growth of R. jostii RHA1 VanA-. Additionally, higher maximum cell concentration (7.6 × 105

233

cell/mL, 96 h) was achieved by R. opacus PD630 compared to R. jostii RHA1 VanA- (5.5 × 105

234

cell/mL, 120 h). It was also observed that microbial cell concentration of R. opacus PD630

235

remained relatively constant up to 144 h after reaching the maximum value at 96 h. Nevertheless,

236

cell concentration of R. jostii RHA1 VanA- significantly decreased after reaching the maximum

237

value at 96 h. Cell concentrations of R. opacus PD630 and R. jostii RHA1 VanA- decreased 11%

238

and 24%, respectively, within the first 24 h of death phase.

239

Furthermore, the depolymerization of lignin was investigated after 5 days of fermentation.

240

31.6% and 21.2% of lignin were degraded by R. jostii RHA1 VanA- and R. opacus PD630,

241

respectively. Wild strain R. opacus PD630 and mutant strain R. jostii RHA1 VanA- have different

242

capability to degrade the alkali-extracted lignin19. Thus, the growths of these two strains were

243

dramatically different. Higher degradation (39.6%) of lignin was obtained by co-fermentation

244

with these two strains (Fig. 3a). It was reported that R. jostii RHA1 and its mutant R. jostii RHA1

245

VanA- with high oxygenase activity could degrade lignin and some aromatic compounds.17

246

Moreover, R. opacus PD630 could degrade lignin and aromatic compounds to lipids via the β-

247

ketoadipate pathway 7, 12. Thus, the improvement of lignin degradation by co-fermentation might

248

be explained by the combined action of different ligninolytic enzymes and metabolic pathways

249

of R. jostii RHA1 VanA- and R. opacus PD630 in lignin biodegradation. The pH of the 11



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250

fermentation broth had no significant changes throughout fermentation, remaining at ~ 7.0 (Fig.

251

3b). It was also found that cell concentration (OD600) increased with the increase of lignin

252

degradation (Fig. 3c). It was reported that the lignin degradation metabolites could be further

253

used as carbon source by Rhodococcus strains 7. Therefore, co-fermentation of these two strains

254

(R. jostii RHA1 VanA- and R. opacus PD630) might reduce product inhibition thus promote the

255

growth of bacteria and lignin biodegradation. Another possibility was that these metabolites

256

could be used as carbon source and fed into TCA for better cellular growth.

257

Variation of major metabolites with single strain fermentation and co-fermentation on

258

lignin by GC/MS assays. To better understand lignin biodegradation, GC/MS was used to

259

determine fermentation metabolites of lignin by single strain fermentation of R. opacus PD630

260

and R. jostii RHA1 VanA- as well as their co-fermentation (Fig. 4). Different aromatic

261

compounds were detected in the single fermentation and co-fermentation. Five aromatic

262

compounds

263

benzofuran, 2,3-dimethoxybenzoic acid) were found throughout the single fermentation process

264

of R. opacus PD630 (Fig. 4a). Most compounds, including 2,3-dihydro-benzofuran, vanillin, 2-

265

ethoxy-4-anisaldehyde, and 2,3-dimethoxybenzoic acid, appeared in the first 24 h of R. opacus

266

PD630 fermentation. 2-Methoxy-4-vinylphenol could be detected from 24 to 168 h (Table 2). 6

267

aromatic compounds,

268

vinylphenol,

269

hydroxyphenyl)-2-propenoic acid, significantly more than those during fermentation with R.

270

opacus PD630, appeared throughout the single fermentation process by R. jostii RHA1 VanA-

271

(Fig. 4b). There were big differences in reaction intermediates at different time points (Table 3)

272

between the two strains, which indicated that the lignin degradation by R. jostii RHA1 VanA-

(vanillin,

2-methoxy-4-vinylphenol,

2-ethoxy-4-anisaldehyde,

including vanillin, 6-methoxycoumaran-7-ol-3-one,

3-hydroxy-4-methoxy-benzaldehyde,

2,3-dihydro-benzofuran,

2,3-dihydro-

2-methoxy-4and

3-(3-

12


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273

was a complex process. Only one compound (2-methoxy-4-vinylphenol) appeared in both R.

274

opacus PD630 and R. jostii RHA1 VanA- single fermentation within 72-168 h. This showed the

275

growth of the two bacteria on lignin was very different. GC-MS analysis showed that the co-

276

fermentation of R. opacus PD630 and R. jostii RHA1 VanA- on lignin involved 4 kinds of

277

aromatic compounds, including 2,3-dihydro-benzofuran, 2-methoxy-4-vinylphenol, 3-hydroxy-

278

4-methoxy-benzaldehyde, and vanillin (Fig. 4c). Three of these compounds, including 2,3-

279

dihydro-benzofuran, 2-methoxy-4-vinylphenol, and vanillin, were detected in all single- and co-

280

fermentation. 2,3-Dihydro-benzofuran and 2-methoxy-4-vinylphenol could be detected from 24

281

to 168 h. It was worth noting that vanillin and 3-hydroxy-4-methoxy-benzaldehyde only

282

appeared in the co-fermentation after 120 and 168 h, respectively (Table 4) although the GC/MS

283

showed their contents were not high. Clearly, the bacterial growth on the lignin under co-

284

fermentation conditions was different from those of two single fermentations.

285

During

both

single

fermentation

and

co-fermentation,

vanillin

(4-hydroxy-3-

286

methoxybenzaldehyde) and its analogues (e.g., 3-hydroxy-4-methoxy-benzaldehyde, 2-methoxy-

287

4-vinylphenol, and 2,3-dihydro-benzofuran) were detected by GC/MS (Table 1). Vanillin is one

288

of the most important flavor additives in the food industry. It is also used for the production of

289

fragrance, pharmaceuticals and other fine chemicals 29. However, the alkali-extracted lignin was

290

not effectively biotransformed to vanillin with R. opacus PD630 and R. jostii RHA1 VanA-

291

single-strain or co-fermentation (Table 1). During R. opacus PD630 fermentation, vanillin only

292

appeared in small amounts at 24 h and disappeared after 72 h. R. jostii RHA1 VanA- produced

293

vanillin before 24 h. However, this substance only appeared at 120 h in co-fermentation. It was

294

reported that R. jostii RHA1 could break down lignin and lignin model compounds by DypB

295

peroxidase and yield vanillin in a considerable amount

18-19

. R. jostii RHA1 VanA-, which 13



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

Page 14 of 43

296

encoded vanillin-degrading enzyme, could further degrade vanillin to vanillic acid to some

297

extent, and it was able to grow on 1 mM vanillic acid but not on 1 mM vanillin 19. Đn this study,

298

vanllic acid was produced in R. jostii RHA1 VanA- single fermentation (data not shown). R.

299

opacus PD630 could convert vanillic acid via β-ketoadipate pathway 4, 7, 12, 14. Thus, no vanillic

300

acid was detected in the R. opacus PD630 single fermentation and co-fermentation while vanillin

301

and vanillic acid existed in the R. jostii RHA1 VanA- single fermentation.

302

3-Hydroxy-4-methoxy-benzaldehyde (isovanillin), an isomer of vanillin, is a phenolic

303

aldehyde and a selective inhibitor of aldehyde oxidase. It can be metabolized by aldehyde

304

dehydrogenase into isovanillic acid. In this study, isovanillin appeared at 72 h and 168 h in R.

305

jostii RHA1 VanA- single fermentation. However, vanillin was not detected at 72 h and 168 h in

306

its single fermentation. Probably, R. jostii RHA1 VanA- could regulate vanillin and isovanillin

307

metabolism. In this study, it presented a very significant peak in GC/MS results. It appeared in

308

all the single-strain fermentation and co-fermentation. 2-Methoxy-4-vinylphenol was detected

309

after 72 h of R. jostii RHA1 VanA- single-strain fermentation. Notably, its structure is similar to

310

vanillin. It was reported that a high concentration of vanillin might be toxic to the R. jostii RHA1

311

and its mutant VanA-

312

intermediates, 3-hydroxy-4-methoxy-benzaldehyde and 2-methoxy-4-vinylphenol. 2,3-Dihydro-

313

benzofuran existed in all single strain and co-fermentations but at different time periods and

314

concentrations. In R. opacus PD630 single fermentation, 2,3-dihydro-benzofuran was only

315

detected at 24 h, and none was detected after 72 h. Probably, 2,3-dihydro-benzofuran was bio-

316

oxidized by deoxygenation

317

utilization by R. jostii RHA1 VanA- single fermentation and co-fermentation.

15, 18

. It was plausible that vanillin could be transformed into two

30

. Clearly, 2,3-dihydro-benzofuran could be formed during lignin

14


Page 15 of 43

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318

As shown in Fig. 4 and Table 1, it was found that the single strain fermentation and co-

319

fermentation with R. opacus PD630 and R. jostii RHA1 VanA- could effectively degrade lignin

320

and metabolize its degradation metabolites (e.g., vanillin, 3-hydroxy-4-methoxy-benzaldehyde,

321

2-methoxy-4-vinylphenol, and 2,3-dihydro-benzofuran). Notably, co-fermentation could degrade

322

vanillin and its analogues during fermentation of lignin.

323

Variation of major aromatics with co-fermentation on lignin and glucose by NMR assays.

324

A further investigation for the co-fermentation metabolites of lignin and glucose by NMR was

325

accomplished (Fig. 5). Results indicated that, compared to the fermentation metabolites of lignin,

326

the fermentation metabolites of glucose after 24 and 72 h did not have any aromatic protons

327

while glucose was found being consumed. For the fermentation metabolites of lignin after 24 and

328

72 h, the total aromatic protons (~ 6-8 ppm) slightly decreased, which may indicate the

329

consumption of soluble lignin fractions. 2D HSQC NMR was also employed to identify

330

fermentation metabolites of lignin samples, and the methoxyl-aromatic structure (i.e. guaiacol,

331

vanillin or similar structures), assigned to the peak at ~ 3.7 ppm, was found being consumed

332

during the fermentation. There is a sharp peak ~ 3.5 ppm in both fermentation metabolites of

333

glucose and lignin after 72 h. It can be assigned to small molecules containing –OCH3 structure

334

such as methanol, methyl formate or similar structures, which can be the byproducts during the

335

fermentation. The NMR results supported the GC-MS analysis for the fermentation metabolites

336

in Table 1, which indicated that lignin was decomposed by biological conversion.

337

Lipids accumulated in cells under co-fermentation with R. opacus PD630 and R. jostii

338

RHA1 VanA-. Recently, oleaginous fungi and bacteria were used for the production of lipids

339

from lignocellulosic materials

4, 12, 31-39

. However, few reported effective biotransformation of

15



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

Page 16 of 43

340

lignin to lipids by co-fermentation. It was reported that R. opacus PD630 was able to accumulate

341

TAGs up to 76% of the dry cell weight (DCW) when it was incubated in gluconate medium

342

and reached 78 g/L (DCW) composed of 38% TAGs by high cell density batch fermentation

343

using a high concentration of glucose as carbon source 41. Clearly, R. opacus PD630 showed high

344

potential in the biosynthesis of TAGs. To test the lipids accumulated in cells under the co-

345

fermentation with R. opacus PD630 and R. jostii RHA1 VanA-, effects of lignin concentration on

346

the lipids accumulation in cells were investigated. As presented in Fig. 6a, it was found that high

347

lipids yield was obtained at lignin loading from 0.1 to 10 g/L. The incremental lipid yield of 0.33

348

g lipid/ g DCW was achieved at 0.5 g/L of lignin loading after 5 days. When the lignin loading

349

was 10 g/L, the incremental lipid yield reached 0.29 g lipid/ g DCW after 5 days. At over 15 g/L

350

of lignin loading, the incremental lipid yield significantly decreased to 0.15 g lipid/ g DCW. In

351

order to load more lignin for accumulating lipids, the appropriate lignin loading in fermentation

352

was 10 g/L. The mutant strain R. jostii RHA1 VanA-, which encoded a vanillin-degrading

353

enzyme 19, was able to metabolize lignin, and the accumulated vanillic acid could be utilized as a

354

carbon source by R. opacus PD630 12, 14. Moreover, R. opacus PD630 could degrade vanillic acid

355

to lipids via the β-ketoadipate pathway 7, 12. Co-fermentation with R. jostii RHA1 VanA- and R.

356

opacus PD630 is a potential strategy for degrading lignin into some metabolites that could be

357

used as carbon source and fed into β-ketoadipate pathway and TCA for better cellular growth.

40

,

358

Furthermore, the distribution of fatty acids accumulated in cells of R. opacus PD630 and R.

359

jostii RHA1 VanA- grown on lignin (10 g/L) as the sole carbon source was tested. As presented

360

in Fig. 6b, the monounsaturated and saturated fatty acids, including tridecanoic acid (C13:0),

361

myristic acid (C14:0), palmitic acid (C16:0), palmitoleic acid (C16:1), oleic acid (C18:1), stearic

362

acid (C18:0), heneicosanoic acid (C21:0), and methyl lignocerate (C24:0), were found in cells of 16


Page 17 of 43

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363

R. opacus PD630 and R. jostii RHA1 VanA- grown on lignin as sole carbon source. Especially,

364

palmitic acid (C16:0; 35.8%) and oleic acid (C18:1; 47.9%) were the major compositions in cell

365

lipids. A high concentration of palmitic acid (C16:0) 35.8% and oleic acid (C18:1) 47.9% in the

366

FAMEs content showed promising potential for biodiesel production. Clearly, the alkali-

367

extracted lignin could be used as a good carbon source for the biosynthesis of biodiesel in cells

368

through co-fermentation of R. opacus PD630 and R. jostii RHA1 VanA-. In addition to C16 and

369

C18 fatty acids, a certain amount of long chain fatty acids, e.g. heneicosanoic acid (C21:0) and

370

methyl lignocerate (C24:0), were produced from lignin by co-fermentation with R. opacus

371

PD630 and R. jostii RHA1 VanA-.

372

Therefore, lignin could be effectively utilized and further metabolized into lipids by co-

373

fermentation with R. opacus PD630 and R. jostii RHA1 VanA- (Table 1; Fig. 4). A synergetic

374

metabolic pathway of lignin bioconversion to lipids was proposed in this study. Probably, R.

375

jostii RHA1 VanA- could degrade lignin to vanillic acid. R. opacus PD630 could convert vanillic

376

acid or its derivatives to produce TAGs 7 via the β-ketoadipate pathway 4, 7, 14, 19. In this study, co-

377

fermentations by R. opacus PD630 and R. jostii RHA1 VanA- to effectively convert lignin to

378

lipids is successfully demonstrated for the first time.

379 380

■ CONCLUSION

381

Both natural and engineered Rhodococcus strains with lignin degradation and/or lipid

382

biosynthesis capacities were selected to establish a fundamental understanding of the pathways

383

and functional modules necessary to enable a platform for biological conversion of lignin to

384

lipids. The Rhodococcus strains have the pathways for degrading lignin to aromatics and

17



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Page 18 of 43

385

oxidizing ring opening of central aromatic intermediates via the β-ketoadipate pathway, shuttling

386

aromatic-derived carbon into central carbon metabolism via TCA cycle, and accumulating lipids

387

via lipid biosynthetic pathways. Our results suggest that R. jostii RHA1 VanA- could grow with

388

higher cell concentrations using lignin as sole carbon source compared to R. opacus PD630.

389

Furthermore, it was firstly reported that lignin was successfully utilized in co-fermentations by

390

these two Rhodococcus strains. GC-MS and NMR results showed that different lignin aromatics

391

were produced during the single-strain fermentation and co-fermentation. Finally, lignin to lipids

392

(with a yield of 0.39 g lipid/ g DCW) by co-fermentations has been achieved, yet the yield is still

393

low at the current stage. However, the results lead to support for our hypothesis: synthetic

394

reconstruction and balanced modification of key regulators and enzymes in lignin

395

depolymerization, aromatic compound catabolism, lipid biosynthesis, and other relevant

396

processes, will enable Rhodococcus strains to efficiently convert lignin to lipid.

397

■ AUTHOR INFORMATION Corresponding Author *Tel: +1-5093727640. E-mail:

398

[email protected]. Notes The authors declare no competing financial interest.

399 400

■ ACKNOWLEDGMENTS This work was supported by U.S. Department of Energy (DOE)

401

Award # DE-EE0006112 with the Bioproducts, Science & Engineering Laboratory and

402

Department of Biological Systems Engineering at Washington State University. This work was

403

performed in part at the William R. Wiley Environmental Molecular Science Laboratory

404

(EMSL), a national scientific user facility sponsored by the U.S. Department of Energy’s Office

405

of Biological and Environmental Research and located at the Pacific Northwest National

406

Laboratory, operated for the Department of Energy by Battelle. The authors would like to thank

407

Dr. Melvin Tucker from the National Renewable Energy Laboratory for kindly providing corn 18


Page 19 of 43

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408

stover for use in this work. Dr. YC He was partially supported by Jiangsu Government

409

Scholarship for Overseas Studies. We thank Mr. Peiyu Leu, Ms. Marie S. Switab, Dr. Daochen

410

Zhu and Drs. Hasan Bugra Coban for technical support. We also thank Dr. John Cort who helped

411

us to collect part of NMR data for this project and for insightful discussions.

412 413

■ LIST OF ABBREVIATIONS

414

GC-MS: Gas chromatography-mass spectroscopy;

415

GPC: Gel permeation chromatography;

416

HPLC: High performance liquid chromatography;

417

FAME: Fatty acid methyl ester.

418 419 420 421

■ REFERENCES

422 423 424 425 426 427 428 429 430 431

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Green Chemistry 2013, 15 (8), 2070-2074. 5. Pollegioni, L.; Tonin, F.; Rosini, E., Lignin-degrading enzymes. FEBS Journal 2015, 282 (7), 1190-1213. 6. Wang, H.; Tucker, M.; Ji, Y., Recent development in chemical depolymerization of lignin: a review. Journal of Applied Chemistry 2013, 2013. 7. Kosa, M.; Ragauskas, A. J., Bioconversion of lignin model compounds with oleaginous Rhodococci. Applied microbiology and biotechnology 2012, 93 (2), 891-900. 8. Tuomela, M.; Vikman, M.; Hatakka, A.; Itävaara, M., Biodegradation of lignin in a compost environment: a review. Bioresource Technology 2000, 72 (2), 169-183. 9. Bugg, T. D.; Ahmad, M.; Hardiman, E. M.; Rahmanpour, R., Pathways for degradation of lignin in bacteria and fungi. Natural product reports 2011, 28 (12), 1883-1896. 10. Chandra, R.; Bharagava, R. N., Bacterial degradation of synthetic and kraft lignin by axenic and mixed culture and their metabolic products. Journal of Environmental Biology 2013, 34 (6), 991. 11. Abd-Elsalam, H. E.; El-Hanafy, A. A., Lignin biodegradation with ligninolytic bacterial strain and comparison of Bacillus subtilis and Bacillus sp. isolated from Egyptian soil. Am Eurasian J Agric Environ Sci 2009, 5, 39-44. 12. Zhao, J.; Quan, C.; Fan, S., Role of Lignin in Bio-Ethanol Production from Lignocellulosic Biomass. Journal of Biobased Materials and Bioenergy 2013, 7 (5), 533-540. 13. Xie, S.; Sun, S.; Dai, S. Y.; Yuan, J. S., Efficient coagulation of microalgae in cultures with filamentous fungi. Algal Research 2013, 2 (1), 28-33. 14. Szőköl, J.; Rucká, L.; Šimčíková, M.; Halada, P.; Nešvera, J.; Pátek, M., Induction and carbon catabolite repression of phenol degradation genes in Rhodococcus erythropolis and Rhodococcus jostii. Applied microbiology and biotechnology 2014, 98 (19), 8267-8279. 15. Kurosawa, K.; Wewetzer, S. J.; Sinskey, A. J., Engineering xylose metabolism in triacylglycerol-producing Rhodococcus opacus for lignocellulosic fuel production. Biotechnology for biofuels 2013, 6 (1), 1. 16. Zhao, C.; Xie, S.; Pu, Y.; Zhang, R.; Huang, F.; Ragauskas, A. J.; Yuan, J. S., Synergistic enzymatic and microbial lignin conversion. Green Chemistry 2016, 18 (5), 1306-1312. 17. Ahmad, M.; Taylor, C. R.; Pink, D.; Burton, K.; Eastwood, D.; Bending, G. D.; Bugg, T. D., Development of novel assays for lignin degradation: comparative analysis of bacterial and fungal lignin degraders. Molecular Biosystems 2010, 6 (5), 815-821. 18. Chen, H.-P.; Chow, M.; Liu, C.-C.; Lau, A.; Liu, J.; Eltis, L. D., Vanillin catabolism in Rhodococcus jostii RHA1. Applied and environmental microbiology 2012, 78 (2), 586-588. 19. Sainsbury, P. D.; Hardiman, E. M.; Ahmad, M.; Otani, H.; Seghezzi, N.; Eltis, L. D.; Bugg, T. D., Breaking down lignin to high-value chemicals: the conversion of lignocellulose to vanillin in a gene deletion mutant of Rhodococcus jostii RHA1. ACS chemical biology 2013, 8 (10), 2151-2156. 20. Wei, Z.; Zeng, G.; Huang, F.; Kosa, M.; Huang, D.; Ragauskas, A. J., Bioconversion of oxygen-pretreated Kraft lignin to microbial lipid with oleaginous Rhodococcus opacus DSM 1069. Green Chemistry 2015, 17 (5), 2784-2789. 21. Ohta, Y.; Hasegawa, R.; Kurosawa, K.; Maeda, A. H.; Koizumi, T.; Nishimura, H.; Okada, H.; Qu, C.; Saito, K.; Watanabe, T.; Hatada, Y., Enzymatic Specific Production and Chemical Functionalization of Phenylpropanone Platform Monomers from Lignin. ChemSusChem 2016, Ahead of Print. 22. de Gonzalo, G.; Colpa, D. I.; Habib, M. H. M.; Fraaije, M. W., Bacterial enzymes 20


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involved in lignin degradation. Journal of Biotechnology 2016, 236, 110-119. 23. Brown, M. E.; Chang, M. C. Y., Exploring bacterial lignin degradation. Curr. Opin. Chem. Biol. 2014, 19, 1-7. 24. He, Y.; Li, X.; Xue, X.; Swita, M. S.; Schmidt, A. J.; Yang, B., Biological conversion of the aqueous wastes from hydrothermal liquefaction of algae and pine wood by Rhodococci. Bioresour. Technol. 2017, 224, 457-464. 25. He, Y.-C.; Ding, Y.; Xue, Y.-F.; Yang, B.; Liu, F.; Wang, C.; Zhu, Z.-Z.; Qing, Q.; Wu, H.; Zhu, C., Enhancement of enzymatic saccharification of corn stover with sequential Fenton pretreatment and dilute NaOH extraction. Bioresource technology 2015, 193, 324-330. 26. Chen, X.; Shekiro, J.; Franden, M. A.; Wang, W.; Zhang, M.; Kuhn, E.; Johnson, D. K.; Tucker, M. P., The impacts of deacetylation prior to dilute acid pretreatment on the bioethanol process. Biotechnology for biofuels 2012, 5, 8-8. 27. Laskar, D. D.; Tucker, M. P.; Chen, X.; Helms, G. L.; Yang, B., Noble-metal catalyzed hydrodeoxygenation of biomass-derived lignin to aromatic hydrocarbons. Green Chemistry 2014, 16 (2), 897-910. 28. Wang, H.; Ruan, H.; Pei, H.; Wang, H.; Chen, X.; Tucker, M. P.; Cort, J. R.; Yang, B., Biomass-derived lignin to jet fuel range hydrocarbons via aqueous phase hydrodeoxygenation. Green Chemistry 2015, 17 (12), 5131-5135. 29. Kaur, B.; Chakraborty, D., Biotechnological and molecular approaches for vanillin production: a review. Applied biochemistry and biotechnology 2013, 169 (4), 1353-1372. 30. Kimura, N.; Kitagawa, W.; Mori, T.; Nakashima, N.; Tamura, T.; Kamagata, Y., Genetic and biochemical characterization of the dioxygenase involved in lateral dioxygenation of dibenzofuran from Rhodococcus opacus strain SAO101. Applied microbiology and biotechnology 2006, 73 (2), 474-484. 31. Huang, C.; Zong, M.-h.; Wu, H.; Liu, Q.-p., Microbial oil production from rice straw hydrolysate by Trichosporon fermentans. Bioresource Technology 2009, 100 (19), 4535-4538. 32. Lian, J.; Garcia-Perez, M.; Chen, S., Fermentation of levoglucosan with oleaginous yeasts for lipid production. Bioresource technology 2013, 133, 183-189. 33. Ruan, Z.; Zanotti, M.; Archer, S.; Liao, W.; Liu, Y., Oleaginous fungal lipid fermentation on combined acid-and alkali-pretreated corn stover hydrolysate for advanced biofuel production. Bioresource technology 2014, 163, 12-17. 34. Patel, A.; Sindhu, D. K.; Arora, N.; Singh, R. P.; Pruthi, V.; Pruthi, P. A., Biodiesel production from non-edible lignocellulosic biomass of Cassia fistula L. fruit pulp using oleaginous yeast Rhodosporidium kratochvilovae HIMPA1. Bioresource technology 2015, 197, 91-98. 35. Zeng, J.; Zheng, Y.; Yu, X.; Yu, L.; Gao, D.; Chen, S., Lignocellulosic biomass as a carbohydrate source for lipid production by Mortierella isabellina. Bioresource technology 2013, 128, 385-391. 36. Wang, B.; Rezenom, Y. H.; Cho, K.-C.; Tran, J. L.; Lee, D. G.; Russell, D. H.; Gill, J. J.; Young, R.; Chu, K.-H., Cultivation of lipid-producing bacteria with lignocellulosic biomass: Effects of inhibitory compounds of lignocellulosic hydrolysates. Bioresource technology 2014, 161, 162-170. 37. Kurosawa, K.; Laser, J.; Sinskey, A. J., Tolerance and adaptive evolution of triacylglycerol-producing Rhodococcus opacus to lignocellulose-derived inhibitors. Biotechnology for biofuels 2015, 8 (1), 1. 38. Holder, J. W.; Ulrich, J. C.; DeBono, A. C.; Godfrey, P. A.; Desjardins, C. A.; Zucker, J.; 21



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524 525 526 527 528 529 530 531 532 533 534 535 536 537

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Zeng, Q.; Leach, A. L.; Ghiviriga, I.; Dancel, C., Comparative and functional genomics of Rhodococcus opacus PD630 for biofuels development. PLoS Genet 2011, 7 (9), e1002219. 39. Xie, S.; Qin, X.; Cheng, Y.; Laskar, D.; Qiao, W.; Sun, S.; Reyes, L. H.; Wang, X.; Dai, S. Y.; Sattler, S. E., Simultaneous conversion of all cell wall components by an oleaginous fungus without chemi-physical pretreatment. Green Chemistry 2015, 17 (3), 1657-1667. 40. Wältermann, M.; Luftmann, H.; Baumeister, D.; Kalscheuer, R.; Steinbüchel, A., Rhodococcus opacus strain PD630 as a new source of high-value single-cell oil? Isolation and characterization of triacylglycerols and other storage lipids. Microbiology 2000, 146 (5), 11431149. 41. Kurosawa, K.; Boccazzi, P.; de Almeida, N. M.; Sinskey, A. J., High-cell-density batch fermentation of Rhodococcus opacus PD630 using a high glucose concentration for triacylglycerol production. Journal of biotechnology 2010, 147 (3), 212-218.

22


Page 23 of 43

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538 539 540

Table 1. Changes of vanillin and its analogues in the fermentation broths of lignin Compound Vanillin

3-Hydroxy-4-methoxybenzaldehyde 2-Methoxy-4vinylphenol 2,3-Dihydro-benzofuran

541 542

Strain

Structure

Time

PD630 VanAPD630+VanAPD630 VanAPD630+VanAPD630 VanAPD630+VanAPD630 VanAPD630+VanA-

24 h + + + + + + + +

72 h + + + + + +

120 h + + + + + + +

168 h + + + + + + +

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

543

23



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Page 24 of 43

544

Table 2. The major aromatic compounds variation in R. opacus PD630 single fermentation on

545

lignin.

Compound

546

Time, h

Structure

2,3-Dihydro-benzofuran

24 +

72 -

120 -

168 -

2-Methoxy-4-vinylphenol

+

+

+

+

Vanillin

+

-

-

-

2-Ethoxy-4-anisaldehyde

+

-

-

-

2,3-Dimethoxybenzoic acid

+

-

-

-


24


Page 25 of 43

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547 548

Table 3. The major aromatic compounds variation in R. jostii RHA1 VanA- single fermentation

549

on lignin.

Compound

550 551

Time, h 72 120 + +

Structure


24 +

168 +

Vanillin

+

-

+

-

6-Methoxycoumaran-7-ol-3-one

+

-

-

-


+

+

+

+

3-Hydroxy-4-methoxy-benzaldehyde

-

+

-

+

3-(3-Hydroxyphenyl)-2-propenoic acid,

-

+

+

+


25



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Page 26 of 43

552 553

Table 4. The major aromatic compounds variation in co-fermentation with R. opacus PD630 and

554

R. jostii RHA1 VanA- on lignin.

Compound

555 556

Time, h 120 +

Structure


24 +

72 +

168 +


+

+

+

+

Vanillin

-

-

+

-

3-Hydroxy-4-methoxybenzaldehyde

-

-

-

+


26


Page 27 of 43

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


557

Legend for figures

558 559

Figure 1. 2D HSQC analysis of lignin samples (left two spectrums are ball milled lignin and

560

right two spectrums are alkali-extracted lignin).

561 562

Figure 2. Microbial growth of R. opacus PD630 and R. jostii RHA1 VanA- on lignin (10 g/L) as

563

the sole carbon source.

564 565

Figuer 3. Effects of single fermentation and co-fermentation of R. opacus PD630 and R. jostii

566

RHA1 VanA- on lignin degradation rate (a), pH of biodegradation media (b) and cell

567

concentrations (OD600) (c) (Initial lignin concentration was 10 g/L; 120 h).

568 569

Figure 4. The major aromatic compounds variation in R. opacus PD630 single fermentation on

570

lignin at 24h, 72h, 120h, and 168h, respectively (a). The major aromatic compounds variation in

571

R. jostii RHA1 VanA- single fermentation on lignin 24h, 72h, 120h, and 168h, respectively (b).

572

The major aromatic compounds variation in co-fermentation with R. opacus PD630 and R. jostii

573

RHA1 VanA- on lignin 24h, 72h, 120h, and 168h, respectively (c).

574 27



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Page 28 of 43

575

Figure 5. 1H NMR for assaying the 24 h- and 72 h-fermentation samples from the co-

576

fermentation of R. opacus PD630 and R. jostii RHA1 VanA- using lignin (a) and glucose (b) as

577

carbon source. All the spectrums have been normalized by using the same intensity for the H2O

578

peak (~ 4.8 ppm).

579 580

Figure 6. Effects of different substrate lignin loading on the incremental lipid yield for co-

581

fermentation fermentation of R. opacus PD630 and R. jostii RHA1 VanA- (a); Distribution of

582

fatty acids of the accumulated in cells of PD630 and VanA- grown on lignin as carbon source (b).

583 584 585 586 587 588 589 590 591 592 593 594 595 596 597 598 28


Page 29 of 43

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

599 600 601 602


Figure 1

603 604 605 606 607 608 609 610 611 612 613 614 615 616 617 618 619 620 621 29



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

622 623 624 625 626 627

Page 30 of 43

Figure 2

628 629 630

30


Page 31 of 43

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631


Figure 3

632 633 634

31



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635

Page 32 of 43

Figure 4

636 637 638

32


Page 33 of 43

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639


Figure 5

640 641 642 643 33



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

644 645 646 647

Page 34 of 43

Figure 6

648 649 650 651 652 34


Page 35 of 43

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653 654 655 656 657 658 659




660

Engineering, Washington State University, 2710 Crimson Way, Richland, WA 99354. Tel: 509-

661

372-7640, Fax: 509-372-7690, E-mail: [email protected]

662

663 664 665

Synopsis

666 667 668 669

A series of Rhodococcus strains were selected to enable a platform for bioconversion of alkali corn stover lignin to lipids.

35



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

2D HSQC analysis of lignin samples (left two spectrums are ball milled lignin and right two spectrums are alkali-extracted lignin) 254x190mm (96 x 96 DPI)


Page 36 of 43

Page ACS 37Sustainable of 43 Chemistry & Engineering

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ACS Sustainable Chemistry & Engineering Page 38 of 43

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1 ACS Paragon Plus Environment

Page 39 of 43


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

1H NMR for assaying the 24 h- and 72 h-fermentation samples from the co-fermentation of R. opacus PD630 and R. jostii RHA1 VanA- using lignin (a) and glucose (b) as carbon source. All the spectrums have been normalized by using the same intensity for the H2O peak (~ 4.8 ppm). 254x190mm (96 x 96 DPI)


Page 40 of 43

Page 41 of 43

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1H NMR for assaying the 24 h- and 72 h-fermentation samples from the co-fermentation of R. opacus PD630 and R. jostii RHA1 VanA- using lignin (a) and glucose (b) as carbon source. All the spectrums have been normalized by using the same intensity for the H2O peak (~ 4.8 ppm). 254x190mm (96 x 96 DPI)


ACS Sustainable Chemistry & Engineering Page 42 of 43

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Page 43 of 43



Engineering, Washington State University, Richland, WA 99354. Tel: 509-372-7640, Fax: 509372-7690, E-mail: [email protected]

1. Lignin deploymerization

Lignin-derived compounds R. jostii RHA1 vanA- & R. opacus PD630

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


2. Aromatic degradation

e.g. β-ketoadipate pathway R. jostii RHA1 vanA- & R. opacus PD630

TCA Cycle

3. Lipid biosynthesis R. jostii RHA1 vanA-

R. jostii RHA1 vanA& R. opacus PD630

FAS II

(36%) (48%)

Synopsis A series of Rhodococcus strains were selected to enable a platform for bioconversion of alkali corn stover lignin to lipids.


Lipid Production from Dilute Alkali Corn Stover Lignin by

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