Lignin: Biosynthesis, Functions and Economic Significance [1 ed.] 1536147699, 9781536147698

Lignin is the main natural resource of aromatic structures on Earth. With the depletion of fossil oil and increased envi

319 95 15MB

English Pages 339 [320] Year 2019

Report DMCA / Copyright

DOWNLOAD FILE

Polecaj historie

Lignin: Biosynthesis, Functions and Economic Significance [1 ed.]
 1536147699, 9781536147698

Table of contents :
Contents
Preface
Acronyms
Chapter 1
Lignin Biosynthesis and Control of Lignin Composition
Abstract
Introduction
Lignin Composition and Monolignol Biosynthesis
Un-Conventional Lignin Monomers
C-Lignin from Natural Plants
Potential Pathway Leading to C-Lignin Biosynthesis
C-Lignin as a Novel Natural Polymer
Conclusion and Perspectives
Acknowledgments
References
Chapter 2
Monolignol Acylation in Lignin Biosynthesis
Abstract
Introduction
Occurence and Characterization of Naturally Acylated Lignin Units
Studies on Detection/Determination of Acylated Lignin Units
Identification of Biosynthetic Pathway to Acylated Lignin Units
Redesigning Lignin in Plants
Bioengineering of Zip-Lignin Hybrid Poplar
Zip-Lignin Strategy Facilitates Utilization of Plant Cell Walls
Recent Advances in Understanding of Monolignol Acylation
Conclusion
Acknowledgments
References
Chapter 3
Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation
Abstract
Introduction
Tricin as a Secondary Metabolite
Biosynthesis of Tricin
Tricin and Its Derivatives
Biological Functions and Potential Applications
Preparation of Tricin
Occurrence of Tricin in the Lignin Polymer
Characterization of Tricin
Quantitation of Tricin
Bioengineering of Tricin
Implications of Tricin’s Presence in Lignin
Conclusion
Acknowledgments
References
Chapter 4
Solution-State Multidimensional NMR of Lignins: Approaches and Applications
Abstract
Introduction
2D HSQC (Heteronuclear Single-Quantum Coherence)
Peak Assignments
Comparative Peak Quantification
Typical HSQC Profiles of Natural Lignins
Non-Canonical Lignin Substructures in Transgenic/Mutant Plants
HSQC Variants for Enhanced Quantitative Capability
2D HMBC (Heteronuclear Multiple-Bond Correlation)
2D HSQC-TOCSY (Heteronuclear Single-Quantum Coherence-Total Correlation Spectroscopy)
3D NMR Experiments
Sample Preparation Strategies for 2D and 3D NMR of Lignins
Direct Dissolution/Swelling of Whole Cell Walls
Derivatization of Whole Cell Walls for Complete Solubilization
Lignin-Enrichment via Cellulase Treatments
Isolation of Soluble Lignins
LCC Fractions
Conclusion
Acknowledgments
References
Chapter 5
Lignin Condensation and Lignin Depolymerization
Abstract
1. Introduction
2. Lignin Polymerization
3. Lignin Condensation
4. Lignin Depolymerization
4.1. Pyrolysis and Gasification
4.2. Catalytic Depolymerization
4.2.1. Reductive Depolymerization
4.2.2. Oxidative Depolymerization
4.3. Biological Degradation
Conclusion
References
Chapter 6
Challenges and Opportunities of Lignin Reductive Catalytic Depolymerization in the Bioethanol Refinary
Abstract
Introduction
Catalytic Reductive Depolymerization of Lignin
Depolymerization of Lignin Model Compounds
Depolymerization of Technical Lignins
Future Outlook
Activating C-C Bonds
Separating Monomeric Products
Catalysts Stability in Hot-Aqueous Environments
Conclusion
References
Chapter 7
The Potential Role of Enzymatic Catalysis and Metabolic Engineering in Lignin Valorization
Abstract
Introduction
The Bio-Degradation of Lignin-Derived Aromatic
The Enzymatic Degradation of β-Ethers
The Metabolic Pathways of Lignin Derivatives from G-/S-/H-Units
The Transportation of Lignin Derivatives from G-/S-/H-Units and Their Metabolic Regulation
Biotransformation for the Production of Chemicals from Depolymerized Lignin and Lignin Model Compounds
Enzymatic Conversion in Vitro
Metabolic Conversion in Vivo
Combining Enzymatic Catalysis and Metabolic Engineering with a Multi- Disciplinary Approach for Lignin Valorization
Conclusion
Acknowledgments
References
Chapter 8
Structural Elucidation of Lignin Macromolecules from p-Coumarate 3-Hydroxylase (C3H) Down-Regulated Transgenic Poplars (84 K)
Abstract
Introduction
Material and Methods
Plant Materials
Preparation of CELs
Structural Elucidation of CELs
Result and Discussion
The Fate of Lignin before and after C3H Down-Regulation
2D-HSQC NMR Analysis
13C NMR Spectra Analysis
31P-NMR Analysis
Molecular Weight Analysis
Conclusion
Acknowledgments
References
Chapter 9
Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres
Abstract
1. Introduction
2. Molecular Designs of Lignin Chain Structures
2.1. Click Reaction between Alkyne and Azide
2.2. Synthesis of Lignin Polymers via Click Reaction of Lignin-Alkyne with Lignin-Azide
2.3. Synthesis of Lignin Polymers via Click Reaction of Lignin-Alkyne with PEG-Azide
2.4. Synthesis of Lignin Polymers via Click Reaction of Lignin-PCL-Alkyne with Lignin-Azide
2.5. Synthesis of Lignin Polymers via the Click Reaction of Lignin- (PCL- co- PLA)-Alkyne with Lignin-Azide
2.6. DSC Analysis of Lignin Polymers Prepared by the Thermal Click Reactions
3. Fabrication of Size-Controlled Lignin Nanospheres
3.1. Size-Controlled Lignin Nanospheres
3.2. Morphology, Size and Yield of Lignin Nanospheres
3.3. The Chemical Characteristics of Lignin Nanospheres
3.4. The Composition Characteristics of Lignin Nanospheres
3.5. Size Influences of Lignin Nanospheres
3.6. The Formation Mechanism of Lignin Nanospheres
4. Fabrication of Hollow Nanospheres with a Single Hole
4.1. The Preparation of Lignin Hollow Nanospheres
4.2. Morphology and Size of Lignin Hollow Nanospheres
4.3. Surface Area, Pore Size and Distribution of Lignin Hollow Nanospheres
4.4. The Chemical Characteristics of Lignin Hollow Nanospheres
4.5. Effects of Stirring Rate, Dropping Speed of Water and pH on the Characteristics of Lignin Hollow Nanospheres
4.6. The Formation Mechanism of Lignin Hollow Nanospheres
Conclusion
Acknowledgment
References
Chapter 10
Thermal Characterization of Klason Lignins from Softwood and Hardwood Species
Abstract
Introduction
Theoretical Background
Kissinger Method
Flynn-Wall-Ozawa Method
Criado Method
Materials and Methods
Materials
Thermogravimetric Analysis (TGA)
Fourier Transform Infrared Spectroscopy (FTIR)
Results and Discussions
TGA Analysis
FTIR Analysis
Conclusion
Acknowledgments
References
Chapter 11
Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents: Applicability in Human Health Promotion
Abstract
1. Introduction
2. Structure and Sources of Lignin
3. Methods of Isolation of Lignin and Lignin Fractions
3.1. Acid Precipitation
3.2. Membrane Filtration
3.3. Solvents
4. Biological Properties of Lignin
4.1. Antioxidant Properties
4.2. Antiviral Effects of Lignin
5. Antimicrobial Films Applications
Conclusion
References
Chapter 12
Preparation of Dehydrogenation Polymer from Isoeugenol and Biological Activity Characterization
Abstract
Introduction
Experimental
Materials
Methods
Synthesis of DHP
Fractionation of the DHP
Molecular Weight Determination
Determination of Total Phenol Content
Analysis of the Structure of the DHP
Evaluation of Antimicrobial Activity
Evaluation of Anticancer Activity
Results and Discussion
Molecular Weight of the DHP
FTIR Analysis of the DHP
Analysis of the DHP by 13C-NMR Spectroscopy
Investigation of Antibacterial Activity
Investigation of Anticancer Activity of the DHP Fractions
Conclusion
Acknowledgments
References
About the Editors
Index
Blank Page
Blank Page

Citation preview

BIOCHEMISTRY RESEARCH TRENDS

LIGNIN BIOSYNTHESIS, FUNCTIONS AND ECONOMIC SIGNIFICANCE

No part of this digital document may be reproduced, stored in a retrieval system or transmitted in any form or by any means. The publisher has taken reasonable care in the preparation of this digital document, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained herein. This digital document is sold with the clear understanding that the publisher is not engaged in rendering legal, medical or any other professional services.

BIOCHEMISTRY RESEARCH TRENDS Additional books and e-books in this series can be found on Nova’s website under the Series tab.

BIOCHEMISTRY RESEARCH TRENDS

LIGNIN BIOSYNTHESIS, FUNCTIONS AND ECONOMIC SIGNIFICANCE

FACHUANG LU AND

FENGXIA YUE EDITORS

Copyright © 2019 by Nova Science Publishers, Inc. All rights reserved. No part of this book may be reproduced, stored in a retrieval system or transmitted in any form or by any means: electronic, electrostatic, magnetic, tape, mechanical photocopying, recording or otherwise without the written permission of the Publisher. We have partnered with Copyright Clearance Center to make it easy for you to obtain permissions to reuse content from this publication. Simply navigate to this publication’s page on Nova’s website and locate the “Get Permission” button below the title description. This button is linked directly to the title’s permission page on copyright.com. Alternatively, you can visit copyright.com and search by title, ISBN, or ISSN. For further questions about using the service on copyright.com, please contact: Copyright Clearance Center Phone: +1-(978) 750-8400 Fax: +1-(978) 750-4470 E-mail: [email protected] NOTICE TO THE READER The Publisher has taken reasonable care in the preparation of this book, but makes no expressed or implied warranty of any kind and assumes no responsibility for any errors or omissions. No liability is assumed for incidental or consequential damages in connection with or arising out of information contained in this book. The Publisher shall not be liable for any special, consequential, or exemplary damages resulting, in whole or in part, from the readers’ use of, or reliance upon, this material. Any parts of this book based on government reports are so indicated and copyright is claimed for those parts to the extent applicable to compilations of such works. Independent verification should be sought for any data, advice or recommendations contained in this book. In addition, no responsibility is assumed by the publisher for any injury and/or damage to persons or property arising from any methods, products, instructions, ideas or otherwise contained in this publication. This publication is designed to provide accurate and authoritative information with regard to the subject matter covered herein. It is sold with the clear understanding that the Publisher is not engaged in rendering legal or any other professional services. If legal or any other expert assistance is required, the services of a competent person should be sought. FROM A DECLARATION OF PARTICIPANTS JOINTLY ADOPTED BY A COMMITTEE OF THE AMERICAN BAR ASSOCIATION AND A COMMITTEE OF PUBLISHERS. Additional color graphics may be available in the e-book version of this book.

Library of Congress Cataloging-in-Publication Data ISBN:  H%RRN

Published by Nova Science Publishers, Inc. † New York

CONTENTS Preface

vii

Acronyms

xi

Chapter 1

Lignin Biosynthesis and Control of Lignin Composition Fang Chen

Chapter 2

Monolignol Acylation in Lignin Biosynthesis Fengxia Yue and Fachuang Lu

Chapter 3

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation Wu Lan, Jorge Rencoret, José Carlos del Río and John Ralph

Chapter 4

Solution-State Multidimensional NMR of Lignins: Approaches and Applications Yuki Tobimatsu, Toshiyuki Takano, Toshiaki Umezawa and John Ralph

1 25

51

79

Chapter 5

Lignin Condensation and Lignin Depolymerization Li Shuai

Chapter 6

Challenges and Opportunities of Lignin Reductive Catalytic Depolymerization in the Bioethanol Refinary J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

131

The Potential Role of Enzymatic Catalysis and Metabolic Engineering in Lignin Valorization Wenya Wang, Chen Shi and Robert J. Linhardt

165

Chapter 7

111

vi Chapter 8

Chapter 9

Chapter 10

Chapter 11

Chapter 12

Contents Structural Elucidation of Lignin Macromolecules from p-Coumarate 3-Hydroxylase (C3H) Down-Regulated Transgenic Poplars (84 K) Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang, Bing Wang, Jia-Long Wen, Feng-Xia Yue and Tong-Qi Yuan

191

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres Yanming Han, Fuquan Xiong, Gaiyun Li and Fuxiang Chu

207

Thermal Characterization of Klason Lignins from Softwood and Hardwood Species Matheus Poletto

243

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents: Applicability in Human Health Promotion Gabriela Vazquez-Olivo, Marilyn S. Criollo-Mendoza, Erick P. Gutiérrez-Grijalva, Manuel A. Picos-Salas and J. Basilio Heredia Preparation of Dehydrogenation Polymer from Isoeugenol and Biological Activity Characterization Yimin Xie, Xuekuan Chen, Houkuan Zhao, Chen Jiang, Hongfei Wu, Shuying Bi and Zhezi Ye

261

281

About the Editors

297

Index

299

PREFACE Fachuang Lu and Fengxia Yue State Key Laboratory of Pulp and Paper Engineering South China University of Technology, Guangzhou, China

Lignins recently became one of the hottest topics in biomass or lignocellulose conversions to fuels and chemicals with the increased concerns about issues associated with environmental pollution, global warming, and energy security. Relating to lignin’s biosynthesis, structures, properties and applications, thousands of research papers and many review articles or edited books have been published in the past 20 years because of tremulous effort from academic society and investment from government or private sectors. The main reasons for doing this are obviously correlated to the fact that lignin existing in plants represents great abundant natural polymer not fully or efficiently utilized due to its complexity in structure and composition. Compared to other natural polymers such as cellulose, hemicellulose and protein, lignin is the most unpopular one because it is often considered as a problem in many circumstances although it plays important roles in plant growth or development. Since the term “lignin” was coined to describe residues leftover from acidic hydrolysis of wood, the portrait of lignin has become clearer and closer to lignin itself thanks to the great advances in science and technology. In other words, achievements made in biology, genetics, chemistry, physics and computing science have greatly improved our understanding of lignin in terms of its biosynthetic pathways, chemical structures, functionalities, reactivities, and properties. There have been a few edited books published in the past, each of them focused and reviewed research results reported in one particular aspect about lignin. The book, Production of Biofuels and Chemicals from Lignin, edited by Zhen Fang and Richard L. Smith, provides updated reviews on technologies for lignin

viii

Fachuang Lu and Fengxia Yue

recovery/purification, lignin conversion to biofuels and chemicals; The 2015 book, Lignin and Lignans as Renewable Raw Materials, edited by Francisco G. Calvo-Flores, Jose A. Dbado, and Joaquin Isac-Garca, is more like a textbook that presents a fairly complete summary on the nature, structure, properties, and applications. This book can be seen as a continued effort from my previous book, Lignin: structural analysis, applications in biomaterials and ecological significance, to provide updated knowledge and development on lignin’s biosynthesis, functions and economic significance. This 12 chapter book begins with a brief review on the lignin biosynthesis pathway, recently discovered non-traditional lignin monomers and lignin composition, with the focus on the remarkable discovery of C-lignin and its potential applications; the following chapter 2 describes one of the particularly interesting aspects related to lignin acylation occurred in some hardwoods, dicots and monocots (grass) plants, with highlight on “zip-lignin” strategy aiming to bio-engineering (design) lignin for improved properties; and chapter 3 presents another interesting discovery of a novel constitute, Tricin, as initiator for lignification in some plant species including all grasses; chapter 4 summaries lignin’s structural features and functionalities revealed by solution-state multidimensional NMR, an advanced analytical techniques widely used for characterization of natural polymers including lignin; in chapter 5 two important aspects of reactions involved in processing lignin or lignocellulose, condensation and depolymerization, are discussed with attention to strategies for reducing the formation of condensed (C-C) linkages; chapter 6 provides a critical review on reductive catalytic depolymerization associated with bioethanol refinery, discussing the potential, opportunities, and challenges for implementing such techniques in biomass conversion; chapter 7 briefly reviews the recent developments on enzyme catalysis and metabolic engineering involved in lignin valorization, emphasizing non-radical lignolytic enzymes and their metabolic pathways for lignin conversion as well as their application in lignin biorefinery; the following chapter 8 gives a structural characterization of lignin from p-coumarate 3-hydroxylase (C3H) downregulated transgenic poplar revealing a slight increases in lignin content and S/G ratios of composition along with lower amount of β-5 linkage; chapter 9 discusses preparation, structures and performance of hollow nano materials made of lignin from agricultural residues via molecular design and controllable self-assembly; chapter 10 uses Fourier transform infrared spectroscopy (FTIR), and thermogravimetric analysis (TGA) to characterize Klason lignins of softwood and hardwood in order to evaluate the effects of lignin’s physical characteristics on their thermal properties and decomposition kinetics; chapter 11 reports information about the application of lignins in food, pharmaceutical and cosmetic industries, with focus on lignin’s antioxidant, antimicrobial and antiviral activity; finally chapter 12 evaluates a isoeugenol dehydrogenation polymer (DHP) synthesized by laccase-catalyzed bulk polymerization aiming to understand the relationship between structural characteristics and its antibacterial and anticancer activities, demonstrating that the low molecular weight fraction of the DHP has apparently anticancer activity.

Preface

ix

This book does not intend to cover all aspects of lignin; it offers brief updated information on selected topics relating to lignin, including lignin biosynthesis (chapters 13 and 8), structural characterization (chapters 4, 8, and 10), and applications (chapters 5-7, 9, and 11-12). It is expected to be of interest to students, scientists and engineers who are working in the field of biomass processing, environmental and chemical sciences, biofuels and biochemicals, and pulp & paper industries. We would like to thank all the contributing authors whose dedication and efforts made this book possible by providing the updated results and developments in their respective areas. Our kindest thanks are also conveyed to our colleagues Fang Chen (University of North Texas), Yuki Tobimatsu (Kyoto University), Li Shuai (Virginia Tech.), Wu Lan (École Polytechnique Fédérale de Lausanne), Yong Qian (South China University of Technology), and Xuebin Zhao (Tsinghua University) for their reviewing and proofreading. Our special thanks are given to John Ralph for his constructive discussion and encourage.

ACRONYMS 2D-NMR 4CL C4H CAD CCoAOMT CCR CEL CHI CHS COMT DFRC DHP DLS DMSO DTG F5H FMT FTIR G GC GC-MS GPC GST H HCT HMBC HPLC

2-dimensional nuclear magnetic resonance 4-coumarate:CoA ligase cinnamate 4-hydroxylase cinnamyl alcohol dehydrogenase caffeoyl-CoAO-methyltransferase cinnamoylCoA reductase cellulolytic enzyme lignin chalcone isomerase chalcone synthase caffeic acid-O-methyltransferase derivatization followed by reductive cleavage dehydrogenation polymer dynamic light scattering dimethylsulfoxide derivative thermogravimetric analysis ferulate 5-hydroxylase Feruloyl-CoA Monolignol Transferase gourier transform infrared spectroscopy guaiacyl gas chromatography gas chromatography-mass spectrometer gel permeation chromatography glutathione S-transferase p-hydroxyphenyl hydroxycinnamoyltransferase heteronuclear multiple-bond correlation high performance liquid chromatography

xii

Acronymns

HSQC heteronuclear single-quantum coherence HSQC-TOCSY heteronuclear single-quantum coherence-total correlation spectroscopy INEPT enhanced by polarization transfer IR infrared spectroscopy LCC lignin-carbohydrate complexes LC-MS liquid chromatography with mass spectrometric MRM multiple reaction monitoring MW molecular weight MWL milled wood lignin NCBI National Center for Biotechnology Information NMI N-methylimidazole NMR nuclear magnetic resonance Mn number-average molecular weights OMT O-methyltransferase PHA polyhydroxyalkanoate PLA polylactic acid PMT p-Coumaroyl-CoA Monolignol Transferase Q-HSQC quantitative HSQC S syringyl TBAF tetrabutylammonium fluoride TEM transmission Electron Microscope TGA thermogravimetric analysis THF tetrahydrofuran TIZ tosylation–iodination–zinc-metal treatment TMDP 2-chloro-4,4,5,5-tetramethyl-1,3,2-dioxaphospholane TMS trimethylsilyl TROSY transverse relaxation optimized spectroscopy UV ultraviolet XPS X-ray photoelectron spectroscopy

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 1

LIGNIN BIOSYNTHESIS AND CONTROL OF LIGNIN COMPOSITION Fang Chen* University of North Texas, Denton, Texas, US Center for Bioenergy Innovation, Oak Ridge, Tennessee, US

ABSTRACT Lignins are complex natural polymers resulting from oxidative coupling of 4hydroxycinnamyl alcohols. They are essential for the structural integrity of plant cell walls and are crucial for plant development. Their composition and structure vary from plant species, tissue and cell types, and developmental stages. The biosynthesis of lignin has been extensively studied and pathway of lignin biosynthesis has been under continuous revision. It becomes increasingly clear that lignin biosynthesis is more complex and flexible than it was previously thought. Many unconventional units have been identified in transgenic and mutant plants with perturbations in the monolignol biosynthetic pathway. This chapter give a briefly review on the recent studies on lignin biosynthesis pathway, unconventional lignin monomers and lignin composition, with the focus on the surprising discovery of a novel C-lignin derived only from caffeyl alcohol, the pathway leading to Clignin biosynthesis and potential C-lignin as a novel natural polymer for industrial utilization.

Keywords: lignin biosynthesis, unconventional lignin monomers, monolignols, caffeyl alcohol, C-lignin

*

Corresponding Author Email: [email protected]

2

Fang Chen

INTRODUCTION Lignins are abundant complex natural polymers resulting from the oxidative coupling of 4-hydroxycinnamyl alcohols. The primary monomers for lignification are the three p-hydroxycinnamyl alcohols - i.e., p-coumaryl, coniferyl, and sinapyl alcohols. These monolignols differ in their degree of methoxylation on the aromatic ring. When polymerized, they give rise to the three common lignin subunits, namely the p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) units (Figure 1). Besides the three primary monolignols, many other monomeric or dimeric compounds (e.g., hydroxycinnamic acids, hydroxycinnamaldehyde, cinnamoyl esters and flavonoids) can also serve as precursors for lignin polymerization [1-7]. In a lignin polymer, the lignin subunits are linked together by a series of characteristic linkages (β–O–4, β–5, β–β, etc.), formed by combinatorial phenoxy radial coupling reactions. As a result, the lignin molecules are heterogeneous in composition, molecular weight, cross-linking and functional groups [8-11]. In plant cell walls, lignin cross-links with carbohydrates, and some types of lignins are esterified with phenolic acids (e.g., p-coumaric acid and phydroxybenzoic acid) and acetic acid [3, 12-15].

Figure 1. The three primary monolignols and corresponding lignin subunits.

Lignin Biosynthesis and Control of Lignin Composition

3

Lignification is one of the prerequisites for terrestrial plant life [16-20]. A successful transition from aquatic conditions to a terrestrial environment requires the first land plant to develop rigid cell walls and hydrophobic extracellular biopolymers that can contribute to permeability control and water transport as well as provide UV protection. Lignification occurs mostly in the secondary cell wall of vessels, tracheids, and fibrous tissues of vascular plants. Lignin composition and structure in plants vary from species to species, and they also show a wide range of variability among tissues, cell types, and developmental stages [21-24]. Lignins are essential for the structural integrity of plant cell walls and are crucial for plant development [25, 26]. For instance, lignification of plant cell walls allows plants to grow upright and enables long distance transport of water and solutes in the vascular system [26, 27]. Lignins also play an important role in plant defense: they can establish mechanical barriers to pathogen invasion and provide a protection against pathogens, although the lignin biosynthesis may be differentially regulated in development of vascular tissues and plant defense responses [28-33]. Recent studies suggest that syringyl lignin may be more important than the guaiacyl unit to protect cells against fungal invasion [34, 35]. Furthermore, lignins limit the digestibility of plant matter by herbivores, reducing the nutritive value of the plant and presumably decreasing its desirability. It has been demonstrated that there is a negative correlation between lignin content and digestibility of forages by ruminants [36-39].

LIGNIN COMPOSITION AND MONOLIGNOL BIOSYNTHESIS Lignin composition varies considerably among plant species. Lignin in angiosperms is a heteropolymer of guaiacyl (G) and syringyl (S) units with small amounts of hydroxyphenyl (H) units, while lignin in gymnosperms lacks S units [23, 40-45]. It has been demonstrated that this lack of S units in gymnosperms is due to the absence of the gene which encodes the enzyme catalyzing the hydroxylation of 5-position at the aromatic ring [46, 47]. Since lignin polymerization is a random process governed by simple combinatorial chemistry, the lignin composition in polymers is likely controlled by the availability of the monolignol species at the location of lignification [8, 48]. However, the mechanisms controlling the relative abundance of the lignin subunits in angiosperms and the extent to which enzymatic specificity influences lignin composition in different plant species and cell types are still unknown. The biosynthesis of the monolignols starts with the deamination of phenylalanine, and then the monolignol precursors are functionalized by consequent aromatic hydroxylation and O-methylation and the simultaneous conversion of the side-chain carboxyl to an alcohol group [49]. The monolignol biosynthesis pathway has been extensively studied [44, 50-54]. It was thought that the hydroxylation and methylation reactions occur at the level of the cinnamic acids catalyzed by hydroxylase and methyltransferase. The resulting

4

Fang Chen

p-coumaric, ferulic, and sinapic acids are subsequently converted to the corresponding monolignols by the sequential action of 4-coumarate: CoA ligase (4CL), cinnamoyl CoA reductase (CCR), and cinnamyl alcohol dehydrogenase (CAD) [48, 53, 54] (Figure 2).

Figure 2. The biosynthesis pathways to monolignols. The figure shows the complete metabolic grid of reactions. The original proposed pathway was shown in blue and the later reversion was shown in other colors.

This pathway has been under continuous revision since the 1990s, driven by in vitro enzyme activity assay, functional genomic, and transgenic studies [46, 48, 55-58]. The discovery of caffeoyl-CoA-O-methyltransferase (CCoAOMT) suggested that the methylation of monolignol precursors could occur at the CoA level [59, 60]. The 13C labeled feeding experiments and re-evaluation of the substrate preference of ferulate 5-hydroxylase (F5H) and caffeic acid O-methyltransferase (COMT) demonstrated that the hydroxylation and methylation of the aromatic C5 positions may actually occur at the aldehyde or alcohol level [46, 56, 61, 62]. Most strikingly, the direct conversion of p-coumaric acid to caffeic acid had never been demonstrated; instead, the reaction responsible for hydroxylation at the 3-position of the p-hydroxyphenyl intermediates was shown to occur predominantly at the level of the shikimate ester of p-coumarate rather than at the free acid or its CoA level [63, 64]. This so-called ‘shikimate shunt’ includes a hydroxycinnamoyl CoA: shikimate/quinate hydroxycinnamoyl transferase (HCT) which

Lignin Biosynthesis and Control of Lignin Composition

5

converts p-hydroxycinnamoyl CoA to p-hydroxycinnamoyl shikimate as well as a coumaroyl shikimate 3’-hydroxylase (C3’H) which adds a hydroxyl group to the aromatic ring [65]. It also requires that HCT can act in the reverse direction to convert the caffeoyl shikimate to caffeoyl CoA (Figure 2). These revisions led to a widely accepted monolignol biosynthesis pathway that does not incorporate any p-hydroxycinnamic acids, except pcoumaric acid, as the intermediate precursors for monolignol biosynthesis. However, this pathway is overly simplified. Recent studies have demonstrated that in many plant species, the caffeoyl shikimate esterase (CSE) can catalyze caffeoyl shikimate to caffeic acid, playing an important role in monolignol biosynthesis [66-68] (Figure 2). These findings not only reintegrate caffeic acid into the pathway, but also raise the question about the role of ferulic acid in the monolignol biosynthesis, since COMT has been demonstrably involved in the methylation of both the C3 and the C5 positions and can convert caffeic acid to ferulic acid with high enzymatic activity. Identifying the in vivo substrate of COMT in relation to C3 methylation would further define the actual pathway of monolignol biosynthesis.

UN-CONVENTIONAL LIGNIN MONOMERS It becomes increasingly clear that lignin biosynthesis is more complex and flexible than was previously thought. Trace amounts of the units derived from monomers other than normal monolignols have been incorporated into lignin polymers [5, 69]. Many unconventional units have been identified in transgenic and mutant plants with perturbations in the monolignol biosynthetic pathway [1, 70]. Mutant defective in C3’H has a strongly reduced lignin content, changes in lignin composition, and developmental defects. The lignin of this mutant is almost entirely composed of H units [71-74]. It is very interesting that disruption of the transcriptional co-regulatory complex, Mediator, subunits MED5a and MED5b could rescue the stunted growth and lignin deficiency phenotype without restoring the synthesis of guaiacyl and syringyl lignin subunits [74]. Although cinnamyl aldehyde groups are always detected in lignins, higher levels of cinnamaldehydes in the lignin from plants down-regulated in cinnamyl alcohol dehydrogenase (CAD) indicate substantial incorporation of aldehyde monomers into the polymer [75-77]. In mutants of Medicago truncatula where the CAD1 gene has been disrupted, the plants contain lignin that is almost exclusively composed of hydroxycinnamaldehyde rather than hydroxycinnamyl alcohol units, suggesting that monolignol side chains can exhibit variation without seriously compromising the lignification process [27]. The wood of the pine mutant deficient in CAD showed brown color resembling that of maize brown midrib mutants in which xylem is red-brown due to a reduction in CAD activity [78]. Transgenic plants with a loss of function of caffeic acid 3-O-methyltransferase (COMT) produce less lignin, but the most striking effect of COMT deficiency is the reduction in S units and the

6

Fang Chen

incorporation of 5-hydroxyconiferyl alcohol into the lignin polymer [39, 76, 79, 80]. The incorporation of 5H units in lignin results in the novel benzodioxane structures in the lignin and indicates that COMT plays a major role in the C5 methylation of monolignol precursors and is essential for syringyl unit biosynthesis [81, 82]. In various cinnamoyl CoA reductase (CCR) deficient transgenic plants, higher amounts of ferulic acid are incorporated into lignin polymers [6, 83]. The transgenic plants display an orange brown coloration in xylem cell walls and an increase in the syringyl over guaiacyl (S/G) ratio. The most severe downregulated plants exhibited a strong reduction in lignin content with altered development and collapsed vessel cells. F5H catalyzes the C5 hydroxylation using coniferaldehyde and coniferyl alcohol as substrates [46, 56, 61]. F5H deficient mutant accumulates only guaiacyl lignin, while overexpression of F5H significantly increases the mole percentage of syringyl subunits in the lignin, generating a lignin that is almost entirely comprised of syringyl units [47, 84, 85]. More interestingly, over-expressing F5H in a COMT deficient mutant led to a new type of lignin significantly enriched in 5-hydroxyguaiacyl units [86]. The above results demonstrate that plants can tolerate large variations in lignin composition and that the copolymerization of uncommon monomers may result in novel lignin structures, indicating that the lignin polymer is extremely flexible in its composition. Another example is the recent discovery of tricin as a lignin subunit in many plant species [2, 3, 87]. Tricin is an O-methylated flavone derived from naringenin chalcone through the addition of malonyl CoA to p-coumaroyl CoA. It is widely distributed in many monocot species, including wheat, sorghum, and maize, in either free or conjugated form. Tricin was first identified as a lignin subunit in lignin preparations from wheat straw and subsequently detected in all examined monocots, as well as in some dicot plants [2, 3, 87]. Tricin was found to cross couple with monolignols to form tricin-(4’-O-β)-linkage and may function as a nucleation site for lignification in monocots. The fact of the incorporation of tricin in lignin polymers implies that the monomer exported to cell walls undergoing lignification could copolymerize with monolignols through radical coupling reactions and become part of the lignin polymer. Silencing chalcone synthase, a tricin biosynthesis pathway gene, in maize resulted in a strongly reduction of tricin incorporation in the lignin polymer [88]. More recently, it has been demonstrated that when the tricin biosynthesis pathway was blocked in a rice mutant, the tricin pathway intermediate precursor, naringenin, can be incorporated into the lignin polymer as a novel lignin component [89].

C-LIGNIN FROM NATURAL PLANTS p-Coumaryl, coniferyl, and sinapyl alcohols are the three main monolignols used for lignin biosynthesis. The other two theoretical monolignols, caffeyl alcohol and 5hydroxyconiferyl alcohol, were not thought to be “natural” monolignols until very recently

Lignin Biosynthesis and Control of Lignin Composition

7

(Figure 2). Although the 5H unit derived from 5-hydroxylconiferyl alcohol have been identified in several transgenic and mutant plants and C unit in Pinus radiata tracheary element cultures, the C unit has not been detected in the transgenic plants that downregulated in COMT or CCoAOMT [90, 91]. However, both the caffeyl alcohol and 5-hydroxyconiferyl alcohol were recently found to be the major building blocks of lignin in several natural plants [92-94].

Figure 3. A typical gas chromatograms showing thioacidolysis products from cured vanilla bean samples. The arrow points to the new peaks that late identifies as catechyl monomers. IS: internal standard; G: guaiacyl monomers; S: simapyl monomers. The peaks appear as doublets due to the formation of erythro and threo isomers of each thioacidolysis product.

Figure 4. C-lignin biosynthesis in developing vanilla seed. A: Cross section of fresh vanilla beans at various developmental stages and mature seeds. C-lignin can be detected from vanilla seed at 8 weeks after pollination. B: Gag chromatograms of vanilla samples after thioacidolysis. C-lignin can be detected in seed only. The stem and pod sample after seed removing are very similar, only G and S monomers can be detected thioacidolysis. IS: internal standard; C: catechyl monomers; G: guaiacyl monomers; S: simapyl monomers.

8

Fang Chen

Figure 5. A 2D 13C–1H correlation (HSQC) spectra of acetylated isolated lignins from V. planifolia: A: seed coat; B: a dehydrogenation polymer synthesized via peroxidase-catalyzed polymerization of caffeyl alcohol; C: stem lignin. (Reproduced from [92] with permission).

Figure 6. Gas chromatogram traces of the thioacidolysis products of lignin from seeds of the cactus species. IS: internal standard; C: catechyl monomers; G: guaiacyl monomers; S: simapyl monomers.

A novel lignin called C-lignin, derived only from caffeyl alcohol, was first discovered in vanilla seeds [92]. Initial studies on the lignin composition of mature vanilla beans by

Lignin Biosynthesis and Control of Lignin Composition

9

thioacidolysis revealed the expected presence of regular G and S type lignin, with an S/G ratio of 0.4 (Figure 3). A close examination of the gas chromatogram of a vanilla sample showed that there was a small peak with a mass spectra consistent with the α, β, γtrithioethylpropylcatechol derived from theoretical catechyl (C) units after thioacidolysis [92]. To identify the origin of this compound, we separated the vanilla seeds from other tissues of a vanilla bean and analyzed their lignin compositions with various vanilla tissue samples. The thioacidolysis analysis revealed that the catechyl units can only been detected in the isolated seeds. In contrast, thioacidolysis of the pod residue (after seed isolation), and the stem, leaf, and aerial roots released normal G and S monomers with essentially no C monomer. These data indicate that the lignins present in these tissues are typical G rich G/S-lignins. Importantly, the vanilla seed lignin was entirely composed of C units, with no G or S monomers that can be detected by thioacidolysis (Figure 4). The investigation of Clignin deposition in vanilla seeds at different developmental stages showed that C-lignin can first be detected in the seed coat at around 8 weeks after pollination, when the color of the seed coat turns from transparent white to brown. The C-lignin deposition increases with the seed maturation (Figure 4). At 5 months after pollination, mature seeds contain about 70 μmol C monomers per gram of dry seed. We estimated that the actual amount of the Clignin in the vanilla seed is substantially higher since thioacidolysis is known to be effective in cleavage of β-O-4 linkage of lignin, but not very efficient in breaking down the unusual structure of the C-lignin polymer. Klason analysis of the seed indicated a very high level (∼80%) of acid-insoluble lignin polymers in vanilla seeds, suggesting that C-lignin is a major component of the seed coat. The majority of the remaining material in the seed coat was crystalline cellulose. The chemical compositions of the pod residue and the stem were similar overall. These tissues were mostly rich in cellulose, with modest levels of hemicellulosic and pectic sugars, and Klason lignins [92]. To isolate the C-lignin, vanilla seeds were ball milled, treated with cellulase, and then extracted with dimethylsulfoxide (DMSO). Figure 5 shows the short-range 13C-1H correlation NMR spectra of the acetylated seed coat lignin. Volume integration of the contour signals confirmed that lignin from vanilla seeds is almost exclusively composed of C units. No typical G and S lignin signals were detected. High-field HSQC spectra of the side-chain (aliphatic) regions showed that benzodioxane linkage accounted for over 98% of the total identifiable dimeric units in vanilla seed coat lignin (Figure 5). The dominant benzodioxane linkages found in the C-lignin polymer indicate that catechyl units in Clignin are exclusively linked by one type of bond. It suggested that, after the β-O-4 radical coupling, internal trapping by the o-hydroxyl group in C units is extremely effective in the re-aromatization of quinone methide (QM) intermediates during the lignin polymerization process [95, 96]. The main benzodioxane backbones in the seed polymer have been determined to be trans/cis-isomeric mixtures. The trans/cis composition of the benzodioxane rings in the seed polymer is 97:3, very similar to that of the dehydrogenation polymer (DHP) made from caffeyl alcohol in vitro. These results suggest that the

10

Fang Chen

stereochemistry of re-aromatization of the QM is under simple kinetic chemical control. Therefore, it is most likely that caffeyl alcohol is enzymatically oxidized by plant oxidoreductases, such as peroxidases and laccases, but the cross coupling of free radicals onto the growing polymer is independent of enzymes or other proteins, in the same way that conventional monolignols are coupled during lignin polymerization. The normal β-O4 linkages, which are the predominant linkages in typical natural lignins, were absent in these polymers. In contrast, a typical angiosperm type lignin rich in β-aryl ether units, phenylcoumaran, and resinol units can be found in lignin isolated from vanilla stems. In summary, thioacidolysis and 2D-NMR data clearly indicate that the C-polymer is essentially a homopolymer synthesized purely from caffeyl alcohol, with benzodioxanes being essentially the only inter-monomer unit in the polymer. The molecular weight of vanilla C-lignin determined by GPC after acetylation showed broad distributions, ranging from a few hundred to 100K Da. The number-average degree of polymerization of the isolated lignins extracted with DMSO and 96% dioxane water solution were about18 and 13, respectively. It is very likely that the insoluble fractions left in seed coat after the extraction contains C-lignin with higher molecular masses. Compared to the G/S lignin isolated from vanilla stems, the C-lignin from the seed coat likewise has a lower average molecular mass. These results together with the in vitro DHPs indicate caffeyl alcohol may have a slightly lower polymerization capability both in vitro and in vivo. Furthermore, the lower molecular masses of C lignins may be attributable to the strong tendencies for the monomer toward the end-wise β-O-4 coupling reaction to make the linear polymer, excluding polymer-polymer coupling reactions via the other possible coupling modes such as 5-5 and 4-O-5 couplings that are typically seen in the polymerization of growing oligomers from the traditional monolignols. Table 1. Plants that accumulate C-lignin in their seed coats

Lignin Biosynthesis and Control of Lignin Composition

11

V. planifolia is a member of the Orchidaceae family in the order Asparagales. C-lignin can be detected in the seed coats of two other vanilla species, V. pompona and V. tahitensis. However, C units were not detected in the seed coats of Phalaenopsis orchid species, nor in seeds of Asparagus and Agave, two other members of the monocot order Asparagales. Interestingly, a recent study on seed coat chemistry shows that two orchids species, Neuwiedia veratrifolia and Cypripedium formosanum, contain C-lignin as well as G/S lignin in their seeds coats [97], indicating the widespread scope of C-lignin biosynthesis among the different plant families. Similar C-lignin is found in the seed coats of a number of cactus species in family of Cactaceae [93]. Interestingly, some cactus species possessed another novel natural lignin containing substantial of 5H units (Figure 6). Although Clignin is found coexisting with G/S lignin in some orchid plants, our data showed no coexistence of C-lignins with G/S lignin in all the cactus species we examined so far [93]. They contain either of the C- or G/S-types, but never both. Our analysis showed that Clignin or G/S-lignin types can be found in very closely related species (Figure 5). This finding suggests that the formation of C-lignin is not an ancient trait, but rather has occurred recently. It also suggest that C-lignin can completely replace the G/S lignin in the seed coat and provide new functions for seed coats. The diversity of lignin composition in these plants demonstrates that lignification in nature is even more flexible than we had thought before. Analysis of lignin composition by thioacidolysis revealed a high level of C-lignin in the seed coats of several members of the Cleomaceae and Euphorbiaceae families, including Cleome hassleriana (spider flower), Jatropha curcas (physic nut), Aleurites moluccana (candlenut), and Ricinus communis (castor bean) [94] (Table 1). The seed coats of these plants also contain significant amounts of G/S units derived from coniferyl and sinapyl alcohols. Klason analysis showed 50 to 70% acid-insoluble lignin in the mature seeds of these plants species, and about 40 to 60% of them were C-lignin. In cleome hassleriana, we found that the G lignin unit can be detected in the early seed development stages, a few days after pollination. The G lignin level increases rapidly and reaches a plateau at around 12 days after pollination. At this time, no C-lignin is detected. The Clignin signal first appears at 14 days after pollination, when the seed coat starts to darken, and then rapidly develops over the next few days, by which time the seed coats became dark and hard. Although the in vitro lignin polymerization showed the caffeyl alcohol was capable of cross-coupling with coniferyl and sinapyl alcohol, the NMR analysis of isolated pure lignin from cleome seed coats showed no evidence of direct linkages between Clignins and G-lignins [94]. The biosynthesis of C- and G- lignins is therefore at least temporally or spatially separated during C. hassleriana seed development, producing distinct C- and G- lignins within the seed coat. Analysis of cell wall preparations from stem, leaves, and roots of plant species from Cleomaceae and Euphorbiaceae indicated that

12

Fang Chen

the lignin in vegetative tissues was composed of G and S units without any trace of C units. The function C-lignin plays in seed coats is currently unknown.

POTENTIAL PATHWAY LEADING TO C-LIGNIN BIOSYNTHESIS The genetic and biological mechanisms that allow for the monolignol pathway to be redirected into the biosynthesis of caffeyl alcohol in seed coats are still not clear. To address potential pathways of C-lignin biosynthesis, we isolated RNA from different tissues of vanilla beans at different developmental stages, including the seeds, hairs, placental, and mesocarp tissues, and used next-generation sequencing technologies to generate very large RNA sequence datasets [98]. The annotated sequence data provide a foundation for understanding multiple aspects of the biochemistry and development of the vanilla bean and for identifying candidate genes involved in C-lignin biosynthesis. All the genes potentially involved in monolignol biosynthesis in V. planifolia were identified by comparing sequences with the annotated lignin pathway genes in Arabidopsis. These included genes encoding L-phenylalanine ammonia-lyase, coumaroyl shikimate 3’hydroxylase, cinnamate 4-hydroxylase, 4-coumarate:CoA ligase, cinnamoyl CoA reductase, hydroxycinnamoyl CoA:shikimate hydroxycinnamoyl transferase, caffeoylCoA 3-O-methyltransferase, caffeic acid 3-O-methyltransferase, cinnamyl alcohol dehydrogenase, and caffeoyl shikimate esterase. The vanilla seed transcriptome data clearly show that C-lignin formation in the seed coat involves coordinated expression of all monolignol biosynthetic genes, except the two O-methyltransferases, at C-lignin formation stages. Most of the predicted lignin-related genes shared similar expression patterns in vanilla bean tissues, with highest expression in seeds at 10 weeks postpollination [98]. CCoAOMT transcripts, required to convert caffeoyl moieties to feruloyl moieties, were virtually absent from developing seeds. The quantitative real-time RT-PCR analysis confirmed that PAL, 4CL, CCR, CAD, and C4H displayed very high transcript levels in 8 and 10 week old seeds, whereas CCoAOMT transcripts can be detected in other tissues but barely in seed tissues [98]. These data indicate that the low level of CCoAOMT transcripts is likely the biochemical basis for the production of C-lignin in the vanilla seed coats. It is currently unclear how the low expression of the O-methyltransferases specifically is regulated in vanilla seeds because the enzymes of the whole monolignol pathway are normally coordinately regulated. Deep mining of transcription factors in the transcriptome database should help us to answer this question. Cleome hassleriana is an ideal model system for studying the biosynthesis and deposition of C-lignin. C. hassleriana, closely related to Arabidopsis, is an ornamental plant with a short generation time. The whole genome of C. hassleriana and large transcriptomic sequences are publicly available. Our study shows that, in cleome seed, G lignin is deposited in the seed coat for the first 10-12 days after pollination, then, at middle

Lignin Biosynthesis and Control of Lignin Composition

13

of seed development stage, C-lignin starts to accumulate until the seed is fully matured. The switch from G-lignin to C-lignin deposition is associated with a strong reduction in extractable COMT and CCoAOMT enzyme activities in the seeds. Analysis of the molecular events occurring during the shift from G to C lignin production should allow us to understand the mechanisms by which the O-methyltransferases are downregulated and determine whether expression of new regulatory or biosynthetic genes is necessary for Clignin biosynthesis. These knowledge are critical for engineering C-lignin in other plants. It has been demonstrated that suppression of CCoAOMT expression in P. radiata tracheary element cultures resulted in the incorporation of caffeyl alcohol into lignin and produced a polymer containing catechyl units [99]. However, it remains to be seen if C-lignin can be engineered into xylem to replace the conventional lignin in transgenic plants by precise manipulation COMT and CCoAOMT in a tissue specific manner. Simultaneous mutations in COMT and CCoAOMT in Arabidopsis do not lead to substantial accumulation of C lignin in stems of transgenic plants, and the plants are dwarfed, fail to develop properly, and contain higher H lignin [100].

C-LIGNIN AS A NOVEL NATURAL POLYMER The presence of naturally high levels of C-lignin in the seed coats of plant species like Jatropha, candlenut, and castor bean suggests a near-term resource for exploitation of this polymer. For example, Jatropha is considered to be one of the best candidates for future biodiesel production because its seed kernel contains 43-59% oil. Jatropha today could produce nearly 2 MT of oil/ha per year (~500 gallons) and ~2 MT/ha per year of seed coat. About 45% of the lignin in the Jatropha seed coat is C-lignin. C-lignin has significant potential as a novel material for transformation into value-added chemicals and materials. The simplicity of C-lignin suggests that depolymerization of C-lignin would produce fewer products than that of conventional lignin, making the recovery of the desired products much easier and cheaper. Lignin extracted from candlenut nutshells contains high phenolic hydroxyl group content due to the existence of C-lignin in the nutshell [101]. The depolymerization of organosolv lignin extracted from candlenut over Cu-doped porous metal oxide catalyst showed that the lignin could be converted into simple mixtures of aromatic products in high yield, with the 4-(3-hydroxypropyl)-catechol being the major product, that could be isolated in high purity [101, 102]. C-lignin has a structure and properties that make it an ideal renewable polymer source for bio-based material manufacturing. For example, attempts have been made to use C-lignin isolated from vanilla seeds to make carbon fiber, a lightweight high performance material with broad industrial applications [103, 104]. The challenge now is to develop sources and efficient extraction

14

Fang Chen

and conversion processes for this natural biopolymer that can provide the necessary volumes for industrial processes.

CONCLUSION AND PERSPECTIVES Remarkable progress in the understanding of lignin biosynthesis has been made since the first establishment of the monolignol biosynthesis pathway. The first wave of lignin modification has been predominantly focused on reducing lignin content to improve biomass quality and overcome recalcitrance for bioenergy feedstocks through the lignin pathway gene downregulation [105, 106]. Now, the tailored in planta lignin structure and composition designs not only became possible, but emerged as favorable strategies to overcome biomass recalcitrance and provide high value chemicals and materials. The advent of next generation genomics with the high-throughput metabolomics methods will significantly accelerate the identification of the genes involved in the lignin biosynthetic pathway and regulation. However, some compelling questions remain. A major challenge in lignin biosynthesis is to understand how lignin composition is tightly regulated in different tissue and cell types. Many natural polyphenols, such as catechin, epicatechin and rosmarinic acid, has been identified or demonstrated to be able to copolymerize with normal monolignols, they are not found as acutal components in lignin polymers in plant cell walls [107, 108]. It is most likely that these compounds do not present in the lignifying cell wall. C-lignin has not been found in tissues other than the seed coats. It is not currently clear what prevents C-lignin biosynthesis and accumulation in stem tissues of natural plants. Understanding the molecular mechanisms that spatio-temporally regulate the tissue specific production of C-lignin will enable us to better regulate monolignol biosynthesis and to achieve a flexible design of cell wall lignin with desired structures. The flexibility in lignin monomer composition means that it is possible to redesign lignin in planta for reducing recalcitrance, facilitate lignin isolation, and develop lignin as a high-value coproduct [109]. A homogenous lignin structure is critical for the accelerated development and deployment of the advanced cellulosic biorefinery. The discovery of linear C-lignin homopolymers in nature is a significant breakthrough in lignin chemistry which drastically changes our view of lignification and represents important new opportunities for the redesign of lignin synthesis in plants for sustainable biomass and biomaterial production in the future. Ultimately, we need to demonstrate that dedicated crops like poplar and switchgrass can be engineered to produce homogeneous linear C-lignin or other specialty lignins in their stems. The plants engineered to produce lignin with desired structures and compositions should grow normally without major impacts on plant performance. Doing so will not only reduce the biomass recalcitrance to produce biofuels, but also facilitate the valorization of lignin for chemical and polymer production.

Lignin Biosynthesis and Control of Lignin Composition

15

ACKNOWLEDGMENTS This work was supported partially by the BioEnergy Science Center and the Center for Bioenergy Innovation (Oak Ridge National Laboratory), US Department of Energy (DOE) Bioenergy Research Centers supported by the Office of Biological and Environmental Research in the DOE Office of Science and National Science Foundation, USA.

REFERENCES [1]

Mottiar Y, Vanholme R, Boerjan W, Ralph J, Mansfield SD. (2016). Designer lignins: harnessing the plasticity of lignification. Current opinion in biotechnology 37: 190-200. [2] Lan W, Lu F, Regner M, Zhu Y, Rencoret J, et al. (2015). Tricin, a flavonoid monomer in monocot lignification. Plant physiology 167: 1284-1295. [3] Del Río JC, Rencoret J, Prinsen P, Martínez AnT, Ralph J, et al. (2012). Structural characterization of wheat straw lignin as revealed by analytical pyrolysis, 2D-NMR, and reductive cleavage methods. Journal of agricultural and food chemistry 60: 5922-5935. [4] Boerjan W, Ralph J, Baucher M. (2003). Lignin biosynthesis. Annual review of plant biology 54: 519-546. [5] Ralph J, MacKay JJ, Hatfield RD, O’malley DM, Whetten RW, et al. (1997). Abnormal lignin in a loblolly pine mutant. Science 277: 235-239. [6] Ralph J, Kim H, Lu F, Grabber JH, Leplé JC, et al. (2008). Identification of the structure and origin of a thioacidolysis marker compound for ferulic acid incorporation into angiosperm lignins (and an indicator for cinnamoyl CoA reductase deficiency). The Plant Journal 53: 368-379. [7] Grabber JH, Quideau S, Ralph J. (1996). p-Coumaroylated syringyl units in maize lignin: Implications for β-ether cleavage by thioacidolysis. Phytochemistry 43: 11891194. [8] Ralph J, Lundquist K, Brunow G, Lu F, Kim H, et al. (2004). Lignins: natural polymers from oxidative coupling of 4-hydroxyphenyl-propanoids. Phytochemistry Reviews 3: 29-60. [9] Davin LB, Lewis NG. (2005). Lignin primary structures and dirigent sites. Current opinion in biotechnology 16: 407-415. [10] Terashima N, Atalla R, Ralph S, Landucci LL, Lapierre C, et al. (1995). New preparations of lignin polymer models under conditions that approximate cell wall lignification. I. Synthesis of novel lignin polymer models and their structural

16

[11] [12] [13] [14]

[15]

[16] [17]

[18]

[19]

[20]

[21] [22] [23]

[24]

Fang Chen characterization by 13 C NMR. Holzforschung-International Journal of the Biology, Chemistry, Physics and Technology of Wood 49: 521-527. Bolker HI, Brenner HS. (1970). Polymeric structure of spruce lignin. Science 170: 173-176. Grabber JH. (2005). How do lignin composition, structure, and cross-linking affect degradability? A review of cell wall model studies. Crop Science 45: 820-831. Iiyama K, Lam TB-T, Stone BA. (1994). Covalent cross-links in the cell wall. Plant physiology 104: 315. Hoffmann L, Besseau S, Geoffroy P, Ritzenthaler C, Meyer D, et al. (2005). Acyltransferase-catalysed p-coumarate ester formation is a committed step of lignin biosynthesis. Plant Biosystems-An International Journal Dealing with all Aspects of Plant Biology 139: 50-53. Petrik DL, Karlen SD, Cass CL, Padmakshan D, Lu F, et al. (2014). p ‐ Coumaroyl‐CoA: monolignol transferase (PMT) acts specifically in the lignin biosynthetic pathway in Brachypodium distachyon. The Plant Journal 77: 713-726. Weng JK, Chapple C. (2010). The origin and evolution of lignin biosynthesis. New Phytologist 187: 273-285. Niklas KJ, Cobb ED, Matas AJ. (2017). The evolution of hydrophobic cell wall biopolymers: from algae to angiosperms. Journal of experimental botany 68: 52615269. Renault H, Alber A, Horst NA, Lopes AB, Fich EA, et al. (2017). A phenol-enriched cuticle is ancestral to lignin evolution in land plants. Nature communications 8: 14713. Rensing SA, Lang D, Zimmer AD, Terry A, Salamov A, et al. (2008). The Physcomitrella genome reveals evolutionary insights into the conquest of land by plants. Science 319: 64-69. Martone PT, Estevez JM, Lu F, Ruel K, Denny MW, et al. (2009). Discovery of lignin in seaweed reveals convergent evolution of cell-wall architecture. Current biology 19: 169-175. Zhong R, Ye Z-H. (2007). Regulation of cell wall biosynthesis. Current opinion in plant biology 10: 564-572. Donaldson LA. (2001). Lignification and lignin topochemistry—an ultrastructural view. Phytochemistry 57: 859-873. Xiao B, Sun X, Sun R. (2001). Chemical, structural, and thermal characterizations of alkali-soluble lignins and hemicelluloses, and cellulose from maize stems, rye straw, and rice straw. Polymer degradation and stability 74: 307-319. Sun J-X, Sun X-F, Sun R-C, Fowler P, Baird MS. (2003). Inhomogeneities in the chemical structure of sugarcane bagasse lignin. Journal of Agricultural and Food Chemistry 51: 6719-6725.

Lignin Biosynthesis and Control of Lignin Composition

17

[25] De Meester B, de Vries L, Özparpucu M, Gierlinger N, Corneillie S, et al. (2017). Vessel-specific reintroduction of Cinnamoyl Coa Reductase 1 (CCR1) in dwarfed ccr1 mutants restores vessel and xylary fiber integrity and increases biomass. Plant physiology: pp. 01462.02017. [26] Jones L, Ennos AR, Turner SR. (2001). Cloning and characterization of irregular xylem4 (irx4): a severely lignin‐deficient mutant of Arabidopsis. The Plant Journal 26: 205-216. [27] Zhao Q, Tobimatsu Y, Zhou R, Pattathil S, Gallego-Giraldo L, et al. (2013). Loss of function of cinnamyl alcohol dehydrogenase 1 leads to unconventional lignin and a temperature-sensitive growth defect in Medicago truncatula. Proceedings of the National Academy of Sciences 110: 13660-13665. [28] Vance C, Kirk T, Sherwood R. (1980). Lignification as a mechanism of disease resistance. Annual review of phytopathology 18: 259-288. [29] Tronchet M, Balague C, Kroj T, Jouanin L, Roby D. (2010). Cinnamyl alcohol dehydrogenases‐C and D, key enzymes in lignin biosynthesis, play an essential role in disease resistance in Arabidopsis. Molecular Plant Pathology 11: 83-92. [30] Shadle GL, Wesley SV, Korth KL, Chen F, Lamb C, et al. (2003). Phenylpropanoid compounds and disease resistance in transgenic tobacco with altered expression of L-phenylalanine ammonia-lyase. Phytochemistry 64: 153-161. [31] Gallego-Giraldo L, Escamilla-Trevino L, Jackson LA, Dixon RA. (2011). Salicylic acid mediates the reduced growth of lignin down-regulated plants. Proceedings of the National Academy of Sciences 108: 20814-20819. [32] Gallego‐Giraldo L, Jikumaru Y, Kamiya Y, Tang Y, Dixon RA. (2011). Selective lignin downregulation leads to constitutive defense response expression in alfalfa (Medicago sativa L.). New Phytologist 190: 627-639. [33] Kawasaki T, Koita H, Nakatsubo T, Hasegawa K, Wakabayashi K, et al. (2006). Cinnamoyl-CoA reductase, a key enzyme in lignin biosynthesis, is an effector of small GTPase Rac in defense signaling in rice. Proceedings of the National Academy of Sciences of the United States of America 103: 230-235. [34] Bhuiyan NH, Selvaraj G, Wei Y, King J. (2008). Gene expression profiling and silencing reveal that monolignol biosynthesis plays a critical role in penetration defence in wheat against powdery mildew invasion. Journal of Experimental Botany 60: 509-521. [35] Menden B, Kohlhoff M, Moerschbacher BM. (2007). Wheat cells accumulate a syringyl-rich lignin during the hypersensitive resistance response. Phytochemistry 68: 513-520. [36] Buxton D, Russell JR. (1988). Lignin constituents and cell-wall digestibility of grass and legume stems. Crop science 28: 553-558. [37] Jung H. (1989). Forage lignins and their effects on fiber digestibility. Agronomy Journal 81: 33-38.

18

Fang Chen

[38] Jung H, Vogel K. (1986). Influence of lignin on digestibility of forage cell wall material. Journal of Animal Science 62: 1703-1712. [39] Guo D, Chen F, Inoue K, Blount JW, Dixon RA. (2001). Downregulation of caffeic acid 3-O-methyltransferase and caffeoyl CoA 3-O-methyltransferase in transgenic alfalfa: impacts on lignin structure and implications for the biosynthesis of G and S lignin. The Plant Cell Online 13: 73-88. [40] Fergus B, Goring D. (1970). The location of guaiacyl and syringyl lignins in birch xylem tissue. Holzforschung-International Journal of the Biology, Chemistry, Physics and Technology of Wood 24: 113-117. [41] Fukushima K, Terashima N. (1991). Heterogeneity in formation of lignin. Wood Science and Technology 25: 371-381. [42] Terashima N, Fukushima K. (1988). Heterogeneity in formation of lignin—XI: an autoradiographic study of the heterogeneous formation and structure of pine lignin. Wood Science and Technology 22: 259-270. [43] Terashima N, Fukushima K, Takabe K. (1986). Heterogeneity in formation of lignin. VII. An autoradiographic study on the formation of guaiacyl and syringyl lignin in poplar. Journal of Wood Chemistry and Technology 6: 495-504. [44] Higuchi T. (1990). Lignin biochemistry: biosynthesis and biodegradation. Wood Science and Technology 24: 23-63. [45] Boudet A-M. (2000). Lignins and lignification: selected issues. Plant Physiology and Biochemistry 38: 81-96. [46] Humphreys JM, Hemm MR, Chapple C. (1999). New routes for lignin biosynthesis defined by biochemical characterization of recombinant ferulate 5-hydroxylase, a multifunctional cytochrome P450-dependent monooxygenase. Proceedings of the National Academy of Sciences 96: 10045-10050. [47] Meyer K, Shirley AM, Cusumano JC, Bell-Lelong DA, Chapple C. (1998). Lignin monomer composition is determined by the expression of a cytochrome P450dependent monooxygenase in Arabidopsis. Proceedings of the National Academy of Sciences 95: 6619-6623. [48] Campbell MM, Sederoff RR. (1996). Variation in Lignin Content and Composition (Mechanisms of Control and Implications for the Genetic Improvement of Plants). Plant physiology 110: 3. [49] Whetten R, Sederoff R. (1995). Lignin biosynthesis. The plant cell 7: 1001. [50] Brown SA, Neish A. (1955). Studies of lignin biosynthesis using isotopic carbon: IV. Formation from some aromatic monomers. Canadian journal of biochemistry and physiology 33: 948-962. [51] Neish A. (1960). Biosynthetic pathways of aromatic compounds. Annual review of plant physiology 11: 55-80.

Lignin Biosynthesis and Control of Lignin Composition

19

[52] Brown SA, Neish A. (1956). Studies of lignin biosynthesis using isotopic carbon: V. Comparative studies on different plant species. Canadian journal of biochemistry and physiology 34: 769-778. [53] Higuchi T. (1985). Biosynthesis of lignin. [54] Higuchi T, Brown SA. (1963). Studies of lignin biosynthesis using isotopic carbon: XIII. The phenylpropanoid system in lignification. Canadian journal of biochemistry and physiology 41: 621-628. [55] Humphreys JM, Chapple C. (2002). Rewriting the lignin roadmap. Current opinion in plant biology 5: 224-229. [56] Chen F, Yasuda S, Fukushima K. (1999). Evidence for a novel biosynthetic pathway that regulates the ratio of syringyl to guaiacyl residues in lignin in the differentiating xylem of Magnolia kobus DC. Planta 207: 597-603. [57] Dixon RA, Chen F, Guo D, Parvathi K. (2001). The biosynthesis of monolignols: a “metabolic grid,” or independent pathways to guaiacyl and syringyl units? Phytochemistry 57: 1069-1084. [58] Anterola AM, Lewis NG. (2002). Trends in lignin modification: a comprehensive analysis of the effects of genetic manipulations/mutations on lignification and vascular integrity. Phytochemistry 61: 221-294. [59] Ye Z-H, Kneusel RE, Matern U, Varner JE. (1994). An alternative methylation pathway in lignin biosynthesis in Zinnia. The Plant Cell 6: 1427-1439. [60] Zhong R, Morrison WH, Negrel J, Ye Z-H. (1998). Dual methylation pathways in lignin biosynthesis. The Plant Cell 10: 2033-2045. [61] Osakabe K, Tsao CC, Li L, Popko JL, Umezawa T, et al. (1999). Coniferyl aldehyde 5-hydroxylation and methylation direct syringyl lignin biosynthesis in angiosperms. Proceedings of the National Academy of Sciences 96: 8955-8960. [62] Parvathi K, Chen F, Guo D, Blount JW, Dixon RA. (2001). Substrate preferences of O‐methyltransferases in alfalfa suggest new pathways for 3‐O‐methylation of monolignols. The Plant Journal 25: 193-202. [63] Franke R, Hemm MR, Denault JW, Ruegger MO, Humphreys JM, et al. (2002). Changes in secondary metabolism and deposition of an unusual lignin in the ref8 mutant of Arabidopsis. The Plant Journal 30: 47-59. [64] Franke R, Humphreys JM, Hemm MR, Denault JW, Ruegger MO, et al. (2002). The Arabidopsis REF8 gene encodes the 3 ‐ hydroxylase of phenylpropanoid metabolism. The Plant Journal 30: 33-45. [65] Hoffmann L, Besseau S, Geoffroy P, Ritzenthaler C, Meyer D, et al. (2004). Silencing of hydroxycinnamoyl-coenzyme A shikimate/quinate hydroxycinnamoyltransferase affects phenylpropanoid biosynthesis. The Plant Cell 16: 1446-1465. [66] Vanholme R, Cesarino I, Rataj K, Xiao Y, Sundin L, et al. (2013). Caffeoyl shikimate esterase (CSE) is an enzyme in the lignin biosynthetic pathway in Arabidopsis. Science 341: 1103-1106.

20

Fang Chen

[67] Ha CM, Escamilla‐Trevino L, Serrani Yarce JC, Kim H, Ralph J, et al. (2016). An essential role of caffeoyl shikimate esterase in monolignol biosynthesis in Medicago truncatula. The Plant Journal. [68] Vargas L, Cesarino I, Vanholme R, Voorend W, Saleme MLS, et al. (2016). Improving total saccharification yield of Arabidopsis plants by vessel-specific complementation of caffeoyl shikimate esterase (cse) mutants. Biotechnology for Biofuels 9: 1. [69] Ralph J. (1996). An unusual lignin from kenaf. Journal of natural products 59: 341342. [70] Vanholme R, Demedts B, Morreel K, Ralph J, Boerjan W. (2010). Lignin biosynthesis and structure. Plant physiology 153: 895-905. [71] Ralph J, Akiyama T, Coleman HD, Mansfield SD. (2012). Effects on lignin structure of coumarate 3-hydroxylase downregulation in poplar. Bioenergy research 5: 10091019. [72] Ralph J, Akiyama T, Kim H, Lu F, Schatz PF, et al. (2006). Effects of coumarate 3hydroxylase down-regulation on lignin structure. Journal of Biological Chemistry 281: 8843-8853. [73] Abdulrazzak N, Pollet B, Ehlting J, Larsen K, Asnaghi C, et al. (2006). A coumaroylester-3-hydroxylase insertion mutant reveals the existence of nonredundant metahydroxylation pathways and essential roles for phenolic precursors in cell expansion and plant growth. Plant Physiology 140: 30-48. [74] Bonawitz ND, Im Kim J, Tobimatsu Y, Ciesielski PN, Anderson NA, et al. (2014). Disruption of Mediator rescues the stunted growth of a lignin-deficient Arabidopsis mutant. Nature 509: 376. [75] Baucher M, Chabbert B, Pilate G, Van Doorsselaere J, Tollier M-T, et al. (1996). Red xylem and higher lignin extractability by down-regulating a cinnamyl alcohol dehydrogenase in poplar. Plant Physiology 112: 1479-1490. [76] Ralph J, Lapierre C, Marita JM, Kim H, Lu F, et al. (2001). Elucidation of new structures in lignins of CAD-and COMT-deficient plants by NMR. Phytochemistry 57: 993-1003. [77] Halpin C, Knight ME, Foxon GA, Campbell MM, Boudet AM, et al. (1994). Manipulation of lignin quality by downregulation of cinnamyl alcohol dehydrogenase. The Plant Journal 6: 339-350. [78] MacKay JJ, O’Malley DM, Presnell T, Booker FL, Campbell MM, et al. (1997). Inheritance, gene expression, and lignin characterization in a mutant pine deficient in cinnamyl alcohol dehydrogenase. Proceedings of the National Academy of Sciences 94: 8255-8260. [79] Lapierre C, Rolando C. (1988). Thioacidolyses of pre-methylated lignin samples from pine compression and poplar woods. Holzforschung-International Journal of the Biology, Chemistry, Physics and Technology of Wood 42: 1-4.

Lignin Biosynthesis and Control of Lignin Composition

21

[80] Marita JM, Ralph J, Lapierre C, Jouanin L, Boerjan W. (2001). NMR characterization of lignins from transgenic poplars with suppressed caffeic acid Omethyltransferase activity. Journal of the Chemical Society, Perkin Transactions 1: 2939-2945. [81] Marita JM, Ralph J, Hatfield RD, Guo D, Chen F, et al. (2003). Structural and compositional modifications in lignin of transgenic alfalfa down-regulated in caffeic acid 3-O-methyltransferase and caffeoyl coenzyme A 3-O-methyltransferase. Phytochemistry 62: 53-65. [82] Chen F, Srinivasa Reddy MS, Temple S, Jackson L, Shadle G, et al. (2006). Multi‐ site genetic modulation of monolignol biosynthesis suggests new routes for formation of syringyl lignin and wall‐bound ferulic acid in alfalfa (Medicago sativa L.). The Plant Journal 48: 113-124. [83] Piquemal J, Lapierre C, Myton K, O’connell A, Schuch W, et al. (1998). Down‐

[84]

[85]

[86]

[87]

[88]

[89]

[90] [91]

regulation of Cinnamoyl‐CoA Reductase induces significant changes of lignin profiles in transgenic tobacco plants. The Plant Journal 13: 71-83. Huntley SK, Ellis D, Gilbert M, Chapple C, Mansfield SD. (2003). Significant increases in pulping efficiency in C4H-F5H-transformed poplars: improved chemical savings and reduced environmental toxins. Journal of Agricultural and Food Chemistry 51: 6178-6183. Meyer K, Cusumano JC, Somerville C, Chapple C. (1996). Ferulate-5-hydroxylase from Arabidopsis thaliana defines a new family of cytochrome P450-dependent monooxygenases. Proceedings of the National Academy of Sciences 93: 6869-6874. Weng JK, Mo H, Chapple C. (2010). Over‐expression of F5H in COMT‐deficient Arabidopsis leads to enrichment of an unusual lignin and disruption of pollen wall formation. The Plant Journal 64: 898-911. Lan W, Morreel K, Lu F, Rencoret J, del Río JC, et al. (2016). Maize TricinOligolignol Metabolites and Their Implications for Monocot Lignification. Plant physiology: pp. 02012.02016. Eloy N, Voorend W, Lan W, Saleme MdLS, Cesarino I, et al. (2016). Silencing chalcone synthase impedes the incorporation of tricin in lignin and increases lignin content. Plant physiology: pp. 01108.02016. Lam PY, Tobimatsu Y, Takeda Y, Suzuki S, Yamamura M, et al. (2017). Disrupting flavone synthase II alters lignin and improves biomass digestibility. Plant physiology 174: 972-985. Suzuki S, Lam TBT, Iiyama K. (1997). 5-Hydroxyguaiacyl nuclei as aromatic constituents of native lignin. Phytochemistry 46: 695-700. Ralph J, Lapierre C, Lu F, Marita JM, Pilate G, et al. (2001). NMR evidence for benzodioxane structures resulting from incorporation of 5-hydroxyconiferyl alcohol

22

[92]

[93]

[94]

[95] [96]

[97]

[98]

[99]

[100]

[101]

[102]

[103] [104]

Fang Chen into lignins of O-methyltransferase-deficient poplars. Journal of Agricultural and Food Chemistry 49: 86-91. Chen F, Tobimatsu Y, Havkin-Frenkel D, Dixon RA, Ralph J. (2012). A polymer of caffeyl alcohol in plant seeds. Proceedings of the National Academy of Sciences 109: 1772-1777. Chen F, Tobimatsu Y, Jackson L, Nakashima J, Ralph J, et al. (2013). Novel seed coat lignins in the Cactaceae: structure, distribution and implications for the evolution of lignin diversity. The Plant Journal 73: 201-211. Tobimatsu Y, Chen F, Nakashima J, Escamilla-Treviño LL, Jackson L, et al. (2013). Coexistence but independent biosynthesis of catechyl and guaiacyl/syringyl lignin polymers in seed coats. The Plant Cell Online 25: 2587-2600. Berstis L, Elder T, Crowley M, Beckham GT. (2016). Radical nature of C-lignin. ACS Sustainable Chemistry & Engineering 4: 5327-5335. Elder T, Berstis L, Beckham GT, Crowley MF. (2016). Coupling and Reactions of 5-Hydroxyconiferyl Alcohol in Lignin Formation. Journal of agricultural and food chemistry. Barsberg ST, Lee Y-I, Rasmussen HN. (2018). Development of C-lignin with G/Slignin and lipids in orchid seed coats–an unexpected diversity exposed by ATR-FTIR spectroscopy. Seed Science Research: 1-11. Rao X, Krom N, Tang Y, Widiez T, Havkin-Frenkel D, et al. (2014). A deep transcriptomic analysis of pod development in the vanilla orchid (Vanilla planifolia). BMC genomics 15: 964. Wagner A, Tobimatsu Y, Phillips L, Flint H, Torr K, et al. (2011). CCoAOMT suppression modifies lignin composition in Pinus radiata. The Plant Journal 67: 119129. Do CT, Pollet B, Thévenin J, Sibout R, Denoue D, et al. (2007). Both caffeoyl Coenzyme A 3-O-methyltransferase 1 and caffeic acid O-methyltransferase 1 are involved in redundant functions for lignin, flavonoids and sinapoyl malate biosynthesis in Arabidopsis. Planta 226: 1117-1129. Klein AP, Beach ES, Emerson JW, Zimmerman JB. (2010). Accelerated solvent extraction of lignin from Aleurites moluccana (Candlenut) nutshells. Journal of agricultural and food chemistry 58: 10045-10048. Barta K, Warner GR, Beach ES, Anastas PT. (2014). Depolymerization of organosolv lignin to aromatic compounds over Cu-doped porous metal oxides. Green Chemistry 16: 191-196. Nar M, Rizvi HR, Dixon RA, Chen F, Kovalcik A, et al. (2016). Superior plant based carbon fibers from electrospun poly-(caffeyl alcohol) lignin. Carbon 103: 372-383. Dixon R, D’souza N, Chen F, Nar M. (2018). Methods for producing carbon fibers from poly-(caffeyl alcohol). Google Patents.

Lignin Biosynthesis and Control of Lignin Composition

23

[105] Chen F, Dixon RA. (2007). Lignin modification improves fermentable sugar yields for biofuel production. Nature biotechnology 25: 759-761. [106] Reddy MS, Chen F, Shadle G, Jackson L, Aljoe H, et al. (2005). Targeted downregulation of cytochrome P450 enzymes for forage quality improvement in alfalfa (Medicago sativa L.). Proceedings of the National Academy of Sciences of the United States of America 102: 16573-16578. [107] Grabber JH, Schatz PF, Kim H, Lu F, Ralph J. (2010). Identifying new lignin bioengineering targets: 1. Monolignol-substitute impacts on lignin formation and cell wall fermentability. BMC Plant Biology 10: 114. [108] Tobimatsu Y, Elumalai S, Grabber JH, Davidson CL, Pan X, et al. (2012). Hydroxycinnamate conjugates as potential monolignol replacements: in vitro lignification and cell wall studies with rosmarinic acid. ChemSusChem 5: 676-686. [109] Ragauskas AJ, Beckham GT, Biddy MJ, Chandra R, Chen F, et al. (2014). Lignin valorization: improving lignin processing in the biorefinery. Science 344: 1246843.

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 2

MONOLIGNOL ACYLATION IN LIGNIN BIOSYNTHESIS Fengxia Yue1 and Fachuang Lu1,2,3,* 1

State Key Laboratory of Pulp and Paper Engineering, South China University of Technology, Guangzhou, China 2 Guangdong Engineering Research Center for Green Fine Chemicals, Guangzhou, China 3 Department of Biochemistry, and DOE Great Lakes Bioenergy Research Center, Wisconsin Energy Institute, University of Wisconsin-Madison, WI, US

ABSTRACT Naturally acylated lignin units, the decoration of hydroxyls on lignin structural units with acyl groups, extensively exist in many plants species. More and more nontraditional monolignols have been found in wild-type and transgenic plants, including monolignol acetate, p-hydroxybenzoate, p-coumarate, and ferulate ester conjugate etc., which are now recognized as authentic lignin precursors for lignification. The flexibility of lignification to use monolignols, especially those naturally acylated lignin units, indicates a promising prospect of redesigning lignin to improve its properity for better utilization. One of the outstanding examples was the so called “zip-lignin,” which was bioengineered by introducing ester linkages into poplar lignin polymer backbone and therefore allowed less energy requirement for delignification/lignin depolymerization in downstream process. This chapter briefly reviews the occurrence and characterization of naturally acylated lignin units in plants, progress in zip-lignin engineering, as well as the recent advances in understanding of monolignol acylation.

Keywords: lignin, monolignol, acylation, zip-lignin, DFRC

*

Corresponding Author Email: [email protected]

26

Fengxia Yue and Fachuang Lu

INTRODUCTION Lignin is a major component of plant cell-wall complexes that plays an essential role in their development and strongly affects the utilization of sustainable lignocellulosic biomass. As a highly complex natural polymer, lignin is formed via combinatorial free radical coupling of hydroxycinnamyl alcohols, although various factors can dictate its ultimate structural features [1-3]. Lignification, the polymerization process that generates lignins, occurs mostly in the secondary cell wall of vascular plants. Lignin compositions and structures in plants vary from species and developmental stages [1, 4, 5]. In general, lignin is primarily comprised of three monomers, i.e., p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol, which differs in their degree of methoxylation on the aromatic ring (See Figure 1) [6]. Some of the selected lignins have long been known to be naturally acylated by the detection of various acids liberated from lignin samples after alkali treatment, which has drawn much research attention for decades[7, 8]. Due to the complexity and heterogeneity of lignin, it has been one of the most challenging problems for scientists to understand the composition, structure and biosynthetic pathway for various purposes ranging from plant breeding and genetic alteration to the biomass utilization [9, 10]. In recent years, significant advances in studies on the lignin biosynthesis and structures have been made with the development of analytical methods and techniques combined with appropriate lignin model compounds. In addition to the three primary monolignols, a host of nontraditional lignin monomers and dimers (other than H, G, and S units, Figure 2) have been found in wild-type and transgenic plants, such as monolignol acetate, phydroxybenzoate (pBA), p-coumarate (pCA), ferulate ester conjugates, tricin, caffeyl alcohol and 5-hydroxyconiferyl alcohol, which are now recognized as authentic lignin precursors of lignin polymers [11-24]. For instance, hydroxycinnamyl acetates are found in most hardwoods and are present in high levels in kenaf and palms; hydroxycinnamyl pBA are found in willows, poplars, palms, and aspens; hydroxycinnamyl pCAs are found in all grasses [25]. As a highly abundant natural and renewable polymer, lignin has wide potential applications due to its aromatic nature. The presence of so many different monomeric and dimeric lignin precursors implicates the flexibility of lignification, the diversity of lignin subunits and the chemical complexity of lignin polymer [11, 12, 26]. Specially, the widely occurrence of native acetylated lignin in a large set of vascular plants discloses a possible scenario for redesigning lignin for improved digestability or degradability, thereby the overall efficiency of biomass utilization. The successful introduction of monolignol ferulate conjugates into lignin polymer backbone (zip-lignin) in engineered poplar trees proved to be a promising way to produce lignin, as well as plants that are designed for deconstruction [27]. In this chapter, the following aspects of monolignol acylation in lignins will be reviewed: 1) Compositionally analysis and characterization of naturally acylated lignin units; 2) Applications and development of

Monolignol Acylation in Lignin Biosynthesis

27

acylated units in bioengineering of plants; 3) Recent advances in understanding of monolignol acylation.

Figure 1. Lignin biosynthesis of three traditional monomers (modified from [6]).

Figure 2. Typical nontraditional monolignols as lignin precursors (R 1, R2 = H or OMe).

28

Fengxia Yue and Fachuang Lu

OCCURENCE AND CHARACTERIZATION OF NATURALLY ACYLATED LIGNIN UNITS As a polymer having aryl ether and various other C–O–C and C–C interunit linkages, lignin’s composition and structure vary from different plant species [28-30]. Some lignins in certain plants have long been known to be naturally acylated [8]. Lignins in both C3and C4-grasses are partially p-coumaroylated [31]; In some hardwood trees, notably willow (Salix) and aspen (Populus), lignins are p-hydroxybenzoylated [32, 33]; and acetates have been also implicated in many hardwood lignins [25, 34]. Although there were no hints for how these esters are formed, it is now well-accepted that monolignols acylated by phenolic acids (especially pCA, ferulic acid (FA), and pBA) and acetic acid, are precusors or monomers of lignins in various species and that polymerization of these acylated monolignols result in lignins having various esters [35]. The first evidence for lignin acylation was reported in 1955 by Smith who detected phydroxybenzoic acids (pBA) in aspen (Populus temula) “native lignin” by mild alkaline hydrolysis [7]. Later in the same year he also found that sugarcane (Saccharum spp.) lignins contain p-hydroxycinnamate that contributes to the characteristic ultro-violet (UV) absorption behavior in aqueous alkali solution[8]. From the nature and distribution of ester residues it was considered that ester residues are attached to the pre-formed lignins [8]. Nakano et al. used hydrogenolysis and periodate oxidation to determine the distribution of pBA groups in aspenwood, and proposed that at least part of the pBA groups are located at the -position of the lignin side chain [32]. Regional chemistry (sites of acylation) was proposed for lignin acylation for the first time although no concrete evidence found as to how these ester distributions were resulted. For lignification, sites of acylation are cricial as the position of attachment of the acyl group suggests its biochemical incorporation pathway [31]. Three different pathways (Figure 3) were proposed for acylation of lignins by acids resulting in two different regiochemistries [25]: a) Acylation at the α-position of the lignin side chain indicates free acid (nucleophile) attacks on quinone methide lignin intermediates in a purely chemical controlled manner; b) Acylation at the γ-position suggests that the hydroxycinnamyl alcohol monomers (monolignols) are likely acylated first via certain acyltransferases (enzyme-mediated) and then incorporated into polymeric lignins by radical-coupling mechanisms [37-41]; c) Acylation by activated acids (e.g., p-coumaroyl-CoA) occurs post lignification, resulting in esters at α- (Lignin-I) and/or γ-positions (Lignin-III), depending on the selectivity of the possible transferases (Figure 3). However, the observation that pCAs in grass lignins are involved in many types of units (both isomers of β-O-4 units, β5 units, and even cinnamyl alcohol end-groups) suggests that enzyme-assisted acylation of lignin polymers (pathway c) is unlikely – the enzyme would have to be remarkably nonspecific [41].

Monolignol Acylation in Lignin Biosynthesis

29

Figure 3. Three possible pathways leading to the formation of acylated lignins, modified from [25, 36].

Studies on Detection/Determination of Acylated Lignin Units Determinations of regiochemistry in lignins are complicated by acyl migration known to occur with acetates and uronates but not with pCAs [31, 42, 43]. Various methods and techniques including Infrared Spectroscopy (IR), UV, Nuclear Magnetic Resonance (NMR), and the recent DFRC (Derivatization Followed by Reductive Cleavage) method were used to determine the site of acylation in lignins [28, 33, 36, 38, 41, 44-55]. Applying IR and UV combined with chemical degradation methods, Nakamura et al. used lignin model compounds, such as veratryl pCA and 3-(3,4-dimethoxyphenyl)propyl pCA, to establish the pattern of ester linkage between p-coumaric acid and phenylpropane moieties of bamboo lignin, as well as etherified p-hydroxybenzoic acid in poplar lignin [38]. Based on model study, it is suggested that the mode of ester linkage of phydroxybenzoic acid partially located at the -position in poplar and bamboo. Besides, it

30

Fengxia Yue and Fachuang Lu

is also suggested that parts of p-hydroxybenzoic acid are linked to coniferyl alcohol end group in poplar lignin [38]. Landucci et al. made assignments of p-acetoxybenzoate in Salix clones 13C NMR spectra by comparing with lignin model compounds, and NMR studies showed that pBA in willow lignins are free-phenolic. In addition, the disappearance of these signals in the spectrum of milled wood lignin (MWL) samples after mild alkaline hydrolysis indicated a similar reactivity as an ester linkage. No evidence for the existence of etherified pBA structures was found at that time [33]. Ralph unambiguously revealed pcoumaric acid is attached exclusively at the -position by using 2D HMQC NMR experiments combined with synthetic - and -p-coumaroylated lignin model compounds and a specifically isotope labeled coniferyl alcohol/coniferyl pCA dehydrogenation polymer (DHP) [41]. Such regiochemistry implicates involvement of pre-acylated phydroxycinnamyl alcohols as precursors in the lignification process, while the pathways leading to these acylated monolignols remains unexplained. This means that phydroxycinnamyl alcohols could be acylated enzymatically and subsequently incorporated into the lignin polymer via oxidative coupling [41]. Futhermore, pBAs were originally thought to be partially at the α-position in Populus and bamboo [37, 39, 40], but they may also be (exclusively) at the γ-position since the small amounts of pBAs released by hydrogenolysis of Populus lignins [32] and attributed to benzylic esters may easily come from benozates on the minor allylic cinnamyl alcohol end-groups (γ-position) – pCAs were found on such end-groups in maize lignins [41]. In addition, p-CAs on grasses were found exclusively esterified to the γ-hydroxyls of lignin side chains [25, 41, 56]. It was revealed that pCA units are attached primarily (ca 90%) to syringyl moiety in maize lignin by using thioacidolysis/desulphurization. However, recovery of γ-pCA was low due to ester cleavage by thioacidolysis, which was confirmed by model studies of thioacidolysis [57]. Since 1998, the invention of DFRC method has brought the revolutionary progress for the characterization of acylated units in lignin. In principle, DFRC method can selectively cleave the dominant β-O-4 linkages (and -ethers in lignins) in lignin to produce monomers and dimers for analysis by GC or GC-MS. Compared with other lignin analysis methods, one important advantage of DFRC is that lignin γ-esters (naturally occurred in many plant species) are not cleaved during the DFRC process. Therefore, native acylated units if attached on γ-position of side chain in β-aryl ether structures of lignin can be detected and determined by the DFRC method [36, 44-51]. For characterization of esters on lignins, the DFRC method is more sensitive and produces more informative data (S/G ratios of the released conjugated esters) compared with NMR. Lu et al. used the DFRC method detected and measured p-coumaroylated units in grass lignins, which have confirmed that pCAs are truly attached to the γ-position of lignin structural units [36]. As shown in Figure 4, pCA on lignins from grass plants such as maize and bamboo was detected and determined by the DFRC method. To date, it has

Monolignol Acylation in Lignin Biosynthesis

31

been found that S units of lignins from bamboo, corn, abaca, and curaua are acylated by pCA to a much greater degree than G units. In contrast, syringyl p-coumaroylation presents very low levels in some other plants, such as wheat [15]. Although that S units are more likely linked by labile β-aryl ethers could partially explain why the released S units involve more pCA, the main reason seems to be that preferentially acylation of sinapyl alcohol by p-coumaroyl transferase before its incorporation into lignin. Model compounds of β-aryl ether with γ-p-coumarate were used to test the efficiency of DFRC degradation to release hydroxycinnamyl pCA derivatives (G-pCA and S-pCA, Figure 4) [36]. It was showed that the β-aryl ether linkages in these models were cleaved in about 60-65% yield, much lower than the 95% yields from normal un-acylated β-aryl ether models [36, 44]. Thus, results from DFRC method may have underestimated the pCA on β-aryl ether of lignin. However, S/G ratios of the released acylated monomers seemingly to reflect the distributions of pCA between releasable S units and G units of lignin because studies with model compounds showed that acylation did not differentially affect the release of S or G units from β-aryl ethers [44]. Acetate groups have been recognized as being associated with lignin in kenaf [58] as well as in hardwoods [34] for several decades. However, it has been a challenging problem for scientist to clearly prove the attachment of acetate groups to lignins, especially the regiochemistry of such acetates [25, 34, 58]. Theoretically, information on natural lignin acetates could have been missed easily from NMR characterization because lignins are frequently acetylated for better NMR properties or purified using acetic acid before NMR characterization, which could have dramatically increased the content of acetylation [59, 60]. Besides, in some cases, the quite low abundance of acetates on lignin may also prevent NMR technique from being able to detect acetates. Ralph reported very high content of acetates in lignin from kenaf (Hibiscus cannabinus L.) bast fibers using 2D NMR [61]. However, determination of acylation sites is still limited by NMR. Later, a simple modified version of DFRC method (designated as DFRC′ method here), was developed for better characterization and determination of acetates naturally occurred on lignin. In DFRC′ method, all acetate-based reagents are replaced by their propionate analogues, which allows to detect naturally occurred acetates on lignin [50]. Using the completely acetate-free DFRC′ method, Ralph’s group has provided unambiguous evidence that kenaf bast fiber lignins are highly acetylated on the γ-position of side chain in lignin units and that such acetylation is disproportionately high on S units [25]. These results were consistent with those obtained from the previous NMR work [41, 50]. More importantly, identification of γ-acetylated β–β-linked S dimers in the DFRC′ degradation products of kenaf lignins implicated sinapyl acetate as a truly lignin precursor. Combined with NMR analysis, the DFRC′ method using advanced model compounds derived from sinapyl γ-acetate as substrates, has provided strong evidence that acetylation of kenaf lignin occurs on the γ-position through radical coupling reactions of acetylated monolignols [36, 52, 53].

32

Fengxia Yue and Fachuang Lu

To date, the DFRC′ method has provided the most efficient way for characterization of naturally aceylated lignins. Following the first demonstration of using the DFRC′ to detect naturally occurred acetates on lignin, del Rio et al. have carried out extensive studies on analysis of acetates on isolated lignin samples from various species of plants by using the DFRC′ method [62, 63]. From the results, acetylated lignin units were found in the MWL of all angiosperms investigated, including monocotyledons (sisal, palm tree, bamboo, and abaca) and eudicotyledons (beech, hornbeam, hemp, kenaf, jute, aspen, and eucalypt) but not in the gymnosperms analyzed (spruce and pine). In some plants (e.g., abaca, sisal, kenaf, or hornbeam), lignin acetylation is highly present, exceeding 45% of the uncondensed S lignin units. The analyses suggested that the structure of the lignins from these plants is highly remarkable, being extensively acylated at the γ-carbon of the lignin side chain (up to 80% acylation) with acetate and/or pCA groups and preferentially over S units. Whereas the lignins from sisal and kenaf are γ-acylated exclusively with acetate groups, the lignins from abaca and curaua are acylated by acetic and pCA groups [25, 64]. In addition, naturally acylated lignin units are found in more various plants, such as coconut coir fibers and elephant grass, by reexamination of lignin acylation [65, 66].

Figure 4. A reaction scheme showing how coniferyl/sinapyl dihydro-p-coumarates are released by the DFRC method rom maize lignin (the partial FID gas chromatogram confirsms the identity of GpCA and S-pCA in the DFRC products) [25].

Monolignol Acylation in Lignin Biosynthesis

33

Figure 5. Dehydrodimerization of sinapyl alcohol 1, and sinapyl γ-ester 2, and cross-coupling of 1 and 2. (The traditional monolignol 1 will dehydrodimerize initially, forming the –-coupled bis(quinone methide) intermediate QM1/1, which re-aromatizes by internal -OH attack on each quinone methide electrophilic -carbon to produce syringaresinol 3 as the overwhelmingly major product. When sinapyl acetate 2 dimerizes, it forms an analogous bis(quinone methide) intermediate QM2/2. However, QM2/2 cannot be re-aromatized by internal trapping; structures 5 arise from water attack on one quinone methide moiety with the resulting a-OH attacking the other quinone methide to form a tetrahydrofuran. When 1 and 2 radicals cross-couple, the intermediate bis(quinone methide) QM1/2 now has one quinone methide moiety which can be internally trapped by the single γ-OH to form a single tetrahydrofuran ring, but the other quinone methide can only be re-aromatized by attack of an external nucleophile. The –-bond formed via radical coupling is shown bolded.) [25, 53].

Identification of Biosynthetic Pathway to Acylated Lignin Units Based on the NMR evidence provided by Ralph et al, it was suggested that acylation by pCA exclusively on γ-positions of side chain of lignin units could be the results from the participation of the pre-acylated monolignol [41]. For a better understanding of lignification mechanisms in plants, free radical coupling of sinapyl γ-acetate 2 or cross coupling between sinapyl alcohol 1 were performed and the resulted β – β-linked dehydrodimers were analyzed by NMR and DFRC′ [53]. Compared with data from model compounds, It was found that the β–β coupling of monolignols can be significantly altered by pre-acetylation (or acylation in general) of the monolignols (Figure 5). It has been demonstrated that, with the γ-position acylated, β–β-coupling or cross-coupling can still

34

Fengxia Yue and Fachuang Lu

presumably happen but the re-aromatization reactions following the radical coupling step can no longer be driven by the internal attack of the γ-OH on the quinone methide intermediates QM1/1 (Figure 5, pathway a) – the γ-acetylation prevents such a reaction to produce resinol 3. For the generation of other products, such as compounds 4 and 5, other pathways should be involved (Figure 5, pathway c or bottom part of pathway b). The key point is that the acetyl group can remain attached in non-resinol β–β-coupling products, such as the novel tetrahydrofuran β–β-linked dehydrodimers, which could not have arisen from post-coupling acylation reactions [25]. Therefore, the discovering and identifying these unique non-resinol S structures (acetylated 4 and 5, Figure 6) became a milestone in recognzation of biosynthetic pathway for lignin acylations in Kenaf lignin providing concrete evidence that acetates on Kenaf lignins arise through incorporation of sinapyl acetate (the pre-acetylated sinapyl alcohol), as a lignin precursor, via enzyme-mediated radical coupling mechanisms [53]. Back to 2002, the detection of β–β-coupled dimeric DFRC′ products 7-8 with γ-acetate groups, in the previous study, which were expected to come from the non-resinol syringyl (tetrahydrofuran) structures, showed the preliminary evidence that acylation is at the monolignol stage [52]. Subsequently, all expected diagnostic β–β-coupled DFRC′ degradation products 7-8, 12 and 13, from those novel tetrahydrofuran structures 4 and 5 in Kenaf lignins were observed and identified using synthesized authentic compounds [53]. With the aid of the appropriated model compounds, 2D NMR analysis of the kenaf lignins confirmed the novel tetrahydrofuran structures, unique substructures that can be formed only from the dimerization of pre-acetylated sinapyl alcohol, in the polymer [53].

Figure 6. Major DFRC products from model compounds 3–5 that have been incorporated into lignin [52].

Monolignol Acylation in Lignin Biosynthesis

35

All these evidences above have pointed out that acetate groups in kenaf lignin arise from pre-acetylated sinapyl alcohol (pathway b, Figure 3). In other word, acylation of lignin happens at the monolignol stage. Furthermore, the other esters, i.e., pBA and pCA, on lignins are likely to be formed in this way [25]. Monolignols acylation in lignin implicates transferase enzymes (and, therefore, responsible genes) involved in the acylation, which have been identified in the following studies [53,67]. Access to the genes will open up new ways to alter lignification for expected plant properties that will be beneficial to the downstream utilization.

REDESIGNING LIGNIN IN PLANTS As a major component of lignocellulosic biomass, lignin is crucial to plant growth and development but is a major impediment to efficient biomass utilization by various processes [68]. Besides, lignin has very high energy density and potential commercial value due to its aromatic nature. Lignin valorization is becoming essential for the overall economics of plants biorefinery [69-72]. Modifying or redesigning lignin in plants has been made to reduce lignin levels or to genetically alter lignin biosynthesis for facilitating pulping, improving forage digestibility, or reducing the recalcitrance of lignocellulose feedstocks [70, 73-78]. Over the last decade, it becomes clear that lignification is more complex and flexible than it was previously thought. Many nontraditional units have been identified from lignins in transgenics, mutants as well as in various “natural” plants. The existence of unusual lignins points out the direction to design “ideal lignin” of plants for improving their utilization [68]. In recent years, researchers have been attempting to design plants to increase their degradability by incorporating monomers that lead to a lower degree of polymerization, reduced hydrophobicity, fewer bonds to other cell wall constituents, or novel chemically labile linkages in the polymer backbone [6]. Specifically, the plasticity of lignification is evidenced by the incorporation of γ-acylated monolignol conjugates (acetylated monolignols) in kenaf, sisal, palm and abaca, p-hydroxybenzoylated monolignols in poplar, willow and palm, and p-coumaroylated monolignols in the commelinid monocots [11]. Various potential alternative monomers have been proposed to be used for redesigning of lignins for various purposes. To date, the zip-lignin strategy represents a major breakthrough in designing lignins for better utilization of biomass [27].

Bioengineering of Zip-Lignin Hybrid Poplar Lignification shows considerable plasticity to incorporate nontraditional monomer units. Incorporating ester interunit linkages into lignin could facilitate fiber delignification and utilization [53, 79]. In previous studies, evidence has found that the acetate groups in

36

Fengxia Yue and Fachuang Lu

kenaf lignin stem from pre-acetylated sinapyl alcohol. The other esters including pBA and pCA on lignins are seemingly to be formed in the same manner. Therefore, the theory that one or several certain acyltransferases should be responsible for acylation of monlignols in such plants was proposed [41, 53]. The biological role of pCA in lignin has been highly speculative, and it is hypothesized that the pCA moieties may function as a radical booster [11, 28, 80-84]. In the presence of hydrogen peroxide, peroxidases isolated from corn cell wall can readily oxidize pCA to a radical but are poor oxidizers for sinapyl alcohol. Analysis of corn wall peroxidases did not find specific enzymes that would lead to the preferred incorporation of sinapyl alcohol as seen in other plants [83]. In 2012, Wither et al. successfully expressed the grass-specific acyltransferase enzyme OsPMT Escherichia coli that acylated monolignols with pCA. Although the role of lignin acylation in wall recalcitrance is unclear, the identification of enzymes responsible for monolignol acylation allows for the manipulation of lignin acylation in plants [67]. Later, the gene coding for a protein or enzyme capable of catalyzing the formation of the conjugate p-coumaroyl-CoA: monolignol transferase and feruloyl-CoA: monolignol transferase were identified in the Chinese medicinal dong quai or Chinese angelica (Angelica sinensis). The BAHD acyltransferase, FERULOYL-CoA MONOLIGNOL TRANSFERASE (FMT), was then successfully transformed into poplar (Populus alba × grandidentata), which naturally incorporates other acylated monolignol into its lignin [27]. Finally, a novel chemically labile ester bonds within the lignin backbone, called “zip-lignin” poplar, was developed, Figure 7. Plants that incorporate monolignol ferulates into their lignins have the potential to be particularly important economically, because their lignin backbones are permeated with a chemically labile ester linkage (Figure 7). However, determining the extent to which monolignol ferulates are incorporated into the lignin polymer is extremely difficult because of the diversity of products generated during the polymerization events [27]. In order to maximize the number of cells producing and incorporating the monolignol conjugates into the eudicot lignins, Smith et al. investigated the p-COUMAROYL-CoA MONOLIGNOL TRANSFERASE (PMT) that expressing in poplar and Arabidopsis [86]. Various analytical methods conclusively showed that the introduction of PMT transgenes from rice into Arabidopsis and poplar results in the production of monolignol-pCA conjugates at levels approaching those in monocots. However, the addition of these conjugates did not occur at the expense of the naturally incorporated monolignol pBAs, especially in poplar. Plants expressing the PMT transgene can therefore produce monolignol pCA conjugates essentially without competing with the formation of other acylated monolignols and without drastically impacting normal monolignol production. To date, improving efficiency of the monolignol ferulates incorporated into lignins is still challenging for the researchers, which is essential for increasing the digestibility of the cell wall. Therefore, viable methods to prompt the biosynthesis of monolignol ferulates levels are still undergoing [87, 88].

Monolignol Acylation in Lignin Biosynthesis

37

Figure 7. Model structure zip lignin and its potential behaviors under mild base treatment (A model of a high-zip lignin from 16 coniferyl alcohol monomers and 4 coniferyl ferulate conjugates (magenta) showing some of the ways that both the ferulate and the monolignol moiety may couple into the polymer to form a potentially more cross-linked polymer that nevertheless readily falls apart during pretreatment; readily cleavable ester bonds are shown hashed. Cleaving this oligomer containing 4 zipconjugates cleaves this model lignin into 5 fragments; in general, a polymer containing N zipconjugates will cleave into (N + 1) fragments) [6, 11, 85].

Zip-Lignin Strategy Facilitates Utilization of Plant Cell Walls Bioengineering of lignin in plants, including trees and agricultural crops, by genetically modification of its content or composition, is of great interest due to its impact on industry, agriculture and the environment [89]. One of the important aspects on genetic modification of lignins is to produce plants with desired properties to facilitate fiber delignification or improve its utilization [6, 69]. As early as 2008, Grabber et al. examined alkaline extractability of lignin and the enzymatic hydrolysis of structural polysaccharides after partial substitution of coniferyl alcohol with coniferyl ferulate (an ester conjugate from lignan biosynthesis) alters the formation of lignin models. As the results showed, incorporation of coniferyl ferulate increased lignin extractability by up to 2-fold in aqueous NaOH, providing an avenue for producing fiber with less non-cellulosic and lignin contamination or of delignifying at lower temperatures. Cell walls lignified with coniferyl ferulate were more readily hydrolyzed with fibrolytic enzymes under both with and without alkaline pretreatment. Base on the result, it was expected that zip-lignin strategy plants would greatly facilitate utilization of plant cell walls [79].

38

Fengxia Yue and Fachuang Lu

To date, the most successful transgenic modification of lignin is the zip-lignin strategy that introduced backbone esters by acyltransferases in lignin bioengineering [27]. The difference of zip-lignin poplar from wide type plants is that ester linkages were introduced to its lignin backbone, which is expected to decrease recalcitrance without altering either the function or the content of lignin in engineered plants. However, the report of utilization for zip-lignin poplar is very limited due to low availability of raw materials. After the successful modification of poplar with zip-lignin strategy, the resultant transgenic poplar was used for different treatments to evaluate its effects. Zhou et al. investigated the chemical pulping properties of zip-lignin hybrid poplar to determine if the strategic lignin modifications would enhance chemical pulping efficiencies. Both ziplignin hybrid poplar and wide-type poplar were used for kraft pulping under identical cooking conditions. Compared with the wild-type poplar, improved delignification efficiency confirmed the effect of zip-lignin. In addition, yield and carbohydrate content of the ensuing pulps were slightly elevated. The effect of the zip-lignin poplar compared to the control if both were pulped to the same end-of-cook kappa number were also estimated using statistical models, and the results indicated a gain of 1.41% in yield compared to the control (at a Kappa number range of 9–39). The results demonstrated that the zip-lignin engineered poplars showed better cooking efficiency than their wild-type counterparts under the alkaline kraft cooking conditions [90]. Besides the primarily pulping evaluation, three kind of different ionic liquids, including cholinium lysinate ([Ch][Lys]), 1 ethyl-3-methylimidazolium acetate ([C2C1Im][OAc]), and tetrabutylammonium hydroxide ([TBA][OH]), were applied for the pretreatment of zip-lignin hybrid poplar and wild-type poplar. By comparing with wildtype poplar, the transgenic poplars, especially those with relatively higher amounts of incorporated monolignol ferulate conjugates, yielded up to 23% higher sugar levels after pretreatment with any of these three different ionic liquids followed by enzymatic saccharification. Introducing of ester bonds into the lignin backbone resulted in increased pretreatment efficiency and more mono sugars released by enzymes with lower energy input. These results clearly indicate that zip-lignin strategy improved lignin’s inherent plasticity, demonstrating the potential for being applied to improve biomass conversion into liquid fuels. It has been demonstrated that the copper-catalyzed alkaline hydrogen peroxide (CuAHP) pretreatment method is effective at delignifying recalcitrant poplar biomass [91-94]. Bhalla et al. investigated the delignification efficiency of genetically modified (zip-lignin) poplar under mild alkaline and copper-catalyzed alkaline hydrogen peroxide (Cu-AHP) pretreatments. By comparing with the control (wild type) poplar, improved yields of glucose and xylose released by enzymatic hydrolysis were found for zip-lignin poplar following alkaline-only pretreatment (56% glucose yield from control poplar versus 67% glucose from zip-lignin poplar) and Cu-AHP pretreatment (77% glucose yield from control poplar versus 85% glucose from zip-lignin poplar). Additionally, glycome

Monolignol Acylation in Lignin Biosynthesis

39

profiling, microscopy, and compositional analysis consistently supported the hypothesis that the lignins in these engineered poplar trees are easier to be removed during pretreatment. Furthermore, fluorescence microscopy was applied to image the binding of a recombinantly expressed cellulose-specific carbohydrate binding module (CBM) from Clostridium thermocellum tagged with the green fluorescent protein (GFP), and CBM binding experiments confirmed that the crystalline cellulose is more accessible to hydrolytic enzymes in zip-lignin poplar than in wild type poplar. The introduction of ester bond to the lignin increased the chemical and microbial accessibility to cellulose, that is, the decreased recalcitrance of zip-lignin reduces the chemical inputs, severity of the CuAHP pretreatment process, and/or decreases the enzyme requirements to achieve comparable sugars yields [95]. Overall, these zip-lignin poplars having ester linkages incorporated into the lignin backbone, resulting in lignin that is more amenable to deconstruction methods, especially for alkaline pretreatments that are particularly effective at hydrolyzing ester bonds. However, current data is still inadequate to establish a completed evaluation system for the application of zip-lignin poplar due to the restriction of raw material. Based on these encouraging initial results, more comprehensive strategies, including methods to introduce higher monolignol ferulate content in the engineered plants, modifications of various plant species, techno-economic analysis of various alkali pretreatment methods, green solvent development, as well as pulping systems, are needed to maximize the economic impact of this technology.

RECENT ADVANCES IN UNDERSTANDING OF MONOLIGNOL ACYLATION As the continuous work of zip-lignin strategy, Karlen et al. re-examined the lignin of three plants known to produce monolignol ferulates in their extractives: Chinese angelica (A. sinensis, family Apiaceae), kenaf (Hibiscus cannabinus, family Malvaceae), and balsa (Ochroma pyramidale, family Malvaceae) [12, 96, 97]. With the new knowledge that plants naturally incorporate monolignol ferulates and the suitable analytical DFRC method to diagnostically detect their incorporation into the polymer in hand, Ralph’s group were able to show that these plants also used monolignol ferulates for their lignification. In addition, DFRC assay on a set of plants representing the spermatophytes or “seed plants,” including 13 gymnosperms and 54 angiosperms were also performed to determine whether the utilization of monolignol ferulates happens in lignification process. The results suggested that various angiosperm species might have convergently evolved to natively produce lignins that incorporate monolignol ferulate conjugates. Furthermore, they

40

Fengxia Yue and Fachuang Lu

proposed that such activities may be accomplished by a BAHD feruloyl–coenzyme A monolignol transferase, OsFMT1 (AT5), in rice and its orthologs in other monocots [96]. In addition to the well established monolignol biosynthetic pathway, various monolignol conjugates have also been shown to participate in lignification [11, 26, 53, 98100]. More and more new acylated units were found and confirmed in various plant species [11, 53, 98]. With the aid of NMR technique and appropriate model compounds, Lu et al. investigated naturally p-hydroxybenzoylated lignin in palms. In this study, the presence of both acetate and pBA in palms lignin was clearly revealed by NMR spectra. These results evidenced that pBA units originate from lignification of the preformed monolignol pBA conjugates in the way similar to the pCAs and acetates found acylating lignin units, and are almost entirely derived from sinapyl pBA at least in these palm lignins. It was also proposed that p-hydroxybenzoyl-CoA: monolignol transferases are involved in lignification in various willows (Salix spp.), poplars and aspen (Populus spp.), and palms (family Arecaceae) that have p-hydroxybenzoylated lignins. Identification of the genes involved will allow pBA levels on lignin to be manipulated as recently demonstrated for pCA [20, 99] Karlen et al. have also found that cell walls of leaf base tissues of the Canary island date palm (Phoenix canariensis) are highly decorated lignins by monolignol acetate, benzoate, and pBA etc. By examination of lignified cell wall preparations from 10 species of commelinid monocotyledons from nine families, it has been proved that commelinid monocotyledon lignins are acylated by pCA, and lignin-linked pCA occurs exclusively on the the γ-OH of lignin unit side chains, mostly on S units [26, 35]. Del Rio et al. also revealed by using 2D NMR and DFRC that the occurrence of associated ferulates together with traces of pCA acylating the γ-OH of lignin side chains, predominantly on S-lignin units from lignin in maize fibers [101]. Although functions of such complex and highly acylated lignin compositions are unknown, the presence of so many different monolignol conjugates has demonstrated the plasticity of lignification, the diversity of lignin precursors, as well as the chemical complexity of lignin polymers.

CONCLUSION Substantial evidence shows that many nontraditional monomers, especially naturally acylated units, can be incorporated into lignin in both wild type and genetically modified plants, resulting in diverse structural features in lignins. The flexibility in monomer compositions demonstrates the plasticity of lignification in plants, which is potentially useful for lignin bioengineering aimed to facilitate better utilization of lignin. Increasing research focus lignin’s recalcitrance and recent advances in our understanding of lignin biogenesis have led to novel approaches in plant biotechnology. Remarkable progress has been made in the study on lignin acylation since the first discovery of naturally acylated lignin units, which has provided significant new insight into the lignin biosynthesis. The

Monolignol Acylation in Lignin Biosynthesis

41

success of zip-lignin strategy by introducing backbone esters exemplifies the potential of monolignol acyltransferases in lignin engineering. Encouraged by the proven incorporation of monolignol ferulates conjugates into hybrid poplar, continuous development of additional alternative monomers should further be explored and evaluated for lignin bioengineering to improve the digestibility of lignocellulose biomass.

ACKNOWLEDGMENTS The authors are grateful for being members of Prof. Ralph’s group collaborating with Prof. Wilkerson C. G., and Prof. Mansfield, S. D. to develop the zip-lignin poplar. This work was funded by State Key Laboratory of Pulp and Paper Engineering [2018TS07, 201836, 2016TS03], the National Natural Science Foundation of China [31870560, 31770621], Guangdong Province Science Foundation for Cultivating National Engineering Research Center for Efficient Utilization of Plant Fibers (2017B090903003), and the DOE Great Lakes Bioenergy Research Center (DOE BER Office of Science, DE-FC0207ER64494).

REFERENCES [1]

[2]

[3] [4] [5] [6]

[7]

Ralph, J., Brunow, G., Harris, P. J., Dixon, R. A., Schatz, P. F., Boerjan, W. In Recent Advances in Polyphenol Research, Daayf, F., El Hadrami, A., Adam, L., Ballance, G. M., Eds., Wiley-Blackwell Publishing: Oxford, UK, 2008, Vol. 1. Ralph, J., Bunzel, M., Marita, J. M., Hatfield, R. D., Lu, F., Kim, H., Schatz, P. F., Grabber, J. H., Steinhart, H. 2004. “Peroxidase-dependent cross-linking reactions of p-hydroxycinnamates in plant cell walls.” Phytochemistry Reviews 3, 79-96. Yue, F., Lu, F., Ralph, S., Ralph, J. 2016. “Identification of 4–O–5-units in softwood lignins via definitive lignin models and NMR.” Biomacromolecules 17, 1909-20. Zhong, R. Q., Ye, Z. H. 2007. “Regulation of cell wall biosynthesis.” Current Opinion in Plant Biology 10, 564-72. Donaldson, L. A. 2001. “Lignification and lignin topochemistry – an ultrastructural view.” Phytochemistry 57, 859-73. Mottiar, Y., Vanholme, R., Boerjan, W., Ralph, J., Mansfield, S. D. 2016. “Designer lignins: Harnessing the plasticity of lignification.” Current Opinion in Biotechnology 37, 190-200. Smith, D. C. C. 1955. “p-Hydroxybenzoates groups in the lignin of Aspen (Populus tremula).” Journal of the Chemical Society, doi: 10.1039/JR9550002347 10.1039/ JR9550002347, 2347.

42 [8] [9]

[10]

[11] [12]

[13]

[14]

[15]

[16]

[17]

[18]

[19]

Fengxia Yue and Fachuang Lu Smith, D. C. C. 1955. “Ester groups in lignin.” Nature 176, 267-68. Fu, C., Mielenz, J. R., Xiao, X., Ge, Y., Hamilton, C. Y., Rodriguez, M., Jr., Chen, F., Foston, M., Ragauskas, A., Bouton, J. et al. 2011. “Genetic manipulation of lignin reduces recalcitrance and improves ethanol production from switchgrass.” Proceedings of the National Academies of Sciences 108, 3803-8. Yue, F., Lu, F., Sun, R. C., Ralph, J. 2012. “Syntheses of lignin-derived thioacidolysis monomers and their uses as quantitation standards.” Journal of Agricultural and Food Chemistry 60, 922-28. Ralph, J. 2010. “Hydroxycinnamates in lignification.” Phytochemistry Reviews 9, 65-83. Karlen, S. D., Zhang, C., Peck, M. L., Smith, R. A., Padmakshan, D., Helmich, K. E., Free, H. C. A., Lee, S., Smith, B. G., Lu, F. et al. 2016. “Monolignol ferulate conjugates are naturally incorporated into plant lignins.” Science Advances 2, e1600393: 1-9. Chen, F., Tobimatsu, Y., Jackson, L., Nakashima, J., Ralph, J., Dixon, R. A. 2013. “Novel seed coat lignins in the Cactaceae: structure, distribution and implications for the evolution of lignin diversity.” The Plant Journal 73, 201-11. Chen, F., Tobimatsu, Y., Havkin-Frenkel, D., Dixon, R. A., Ralph, J. 2012. “A polymer of caffeyl alcohol in plant seeds.” Proceedings of the National Academy of Sciences of the United States of America 109, 1772-77. del Río, J. C., Rencoret, J., Prinsen, P., Martínez, Á. T., Ralph, J., Gutiérrez, A. 2012. “Structural characterization of wheat straw lignin as revealed by analytical pyrolysis, 2D-NMR, and reductive cleavage methods.” Journal of Agricultural and Food Chemistry 60, 5922-35. Nakano, H., Kawada, N., Yoshida, M., Ono, H., Iwaura, R., Tonooka, T. 2011. “Isolation and identification of flavonoids accumulated in proanthocyanidin-free barley.” Journal of Agricultural and Food Chemistry 59, 9581-87. Bottcher, A., Cesarino, I., Santos, A. B., Vicentini, R., Mayer, J. L., Vanholme, R., Morreel, K., Goeminne, G., Moura, J. C., Nobile, P. M. et al. 2013. “Lignification in sugarcane: biochemical characterization, gene discovery and expression analysis in two genotypes contrasting for lignin content.” Plant Physiology 163. Zhao, Q., Tobimatsu, Y., Zhou, R., Pattathil, S., Gallego-Giraldo, L., Fu, C., Jackson, L. A., Hahn, M. G., Kim, H., Chen, F. et al. 2013. “Loss of function of Cinnamyl Alcohol Dehydrogenase 1 causes accumulation of an unconventional lignin and a temperature-sensitive growth defect in Medicago truncatula.” Proceedings of the National Academy of Sciences of the United States of America 110, 13660-65. Hoffmann, L., Besseau, S., Geoffroy, P., Ritzenthaler, C., Meyer, D., Lapierre, C., Pollet, B., Legrand, M. 2005. “Acyltransferase-catalysed p-coumarate ester formation is a committed step of lignin biosynthesis.” Plant Biosystems 139, 50-53.

Monolignol Acylation in Lignin Biosynthesis

43

[20] Petrik, D. L., Karlen, S. D., Cass, C. L., Padmakshan, D., Lu, F., Liu, S., Le Bris, P., Antelme, S., Santoro, N., Wilkerson, C. G. et al. 2014. “p-CoumaroylCoA:Monolignol Transferase (PMT) acts specifically in the lignin biosynthetic pathway in Brachypodium distachyon.” The Plant Journal 77, 713-26. [21] Lu, F., Marita, J. M., Lapierre, C., Jouanin, L., Morreel, K., Boerjan, W., Ralph, J. 2010. “Sequencing around 5-hydroxyconiferyl alcohol-derived units in caffeic acid O-methyltransferase-deficient poplar lignins.” Plant Physiology 153, 569-79. [22] Ralph, J., Lapierre, C., Marita, J., Kim, H., Lu, F., Hatfield, R. D., Ralph, S. A., Chapple, C., Franke, R., Hemm, M. R. et al. 2001. “Elucidation of new structures in lignins of CAD- and COMT-deficient plants by NMR.” Phytochemistry 57, 9931003. [23] Ralph, J., Lapierre, C., Lu, F., Marita, J. M., Pilate, G., Van Doorsselaere, J., Boerjan, W., Jouanin, L. 2001. “NMR evidence for benzodioxane structures resulting from incorporation of 5-hydroxyconiferyl alcohol into lignins of O-methyl-transferasedeficient poplars.” Journal of Agricultural and Food Chemistry 49, 86-91. [24] Lan, W., Lu, F., Regner, M., Zhu, Y., Rencoret, J., Ralph, S. A., Zakai, U. I., Morreel, K., Boerjan, W., Ralph, J. 2015. “Tricin, a flavonoid monomer in monocot lignification.” Plant Physiology 167, 1284-95. [25] Lu, F. Lignin: Structural Analysis, Applications in Biomaterials, and Ecological Significance, Nova Science Publishers, Inc: Hauppauge, New York, USA, 2014. [26] Karlen, S. D., Smith, R. A., Kim, H., Padmakshan, D., Bartuce, A., Mobley, J. K., Free, H. C. A., Smith, B. G., Harris, P. J., Ralph, J. 2017. “Highly decorated lignins occur in leaf base cell walls of the Canary Island date palm Phoenix canariensis.” Plant Physiology 175, 1058-67. [27] Wilkerson, C. G., Mansfield, S. D., Lu, F., Withers, S., Park, J. Y., Karlen, S. D., Gonzales-Vigil, E., Padmakshan, D., Unda, F., Rencoret, J. et al. 2014. “Monolignol ferulate transferase introduces chemically labile linkages into the lignin backbone.” Science 344, 90-93. [28] Ralph, J., Lundquist, K., Brunow, G., Lu, F., Kim, H., Schatz, P. F., Marita, J. M., Hatfield, R. D., Ralph, S. A., Christensen, J. H. et al. 2004. “Lignins: natural polymers from oxidative coupling of 4-hydroxyphenylpropanoids.” Phytochemistry Reviews 3, 29-60. [29] Boerjan, W., Ralph, J., Baucher, M. 2003. “Lignin biosynthesis.” Annual Reviews in Plant Biology 54, 519-46. [30] Vanholme, R., Morreel, K., Darrah, C., Oyarce, P., Grabber, J. H., Ralph, J., Boerjan, W. 2012. “Metabolic engineering of novel lignin in biomass crops.” New Phytologist 196, 978-1000. [31] Ralph, J., Helm, R. F. In Forage Cell Wall Structure and Digestibility, Jung, H. G., Buxton, D. R., Hatfield, R. D., Ralph, J., Eds., American Society of Agronomy, Crop Science Society of America, Soil Science Society of America: Madison, WI, 1993.

44

Fengxia Yue and Fachuang Lu

[32] Nakano, J., Ishizu, A., Migita, N. 1961. “Studies on lignin. XXXII. Ester groups of lignin.” Tappi 44, 30-32. [33] Landucci, L. L., Deka, G. C., Roy, D. N. 1992. “A 13C NMR study of milled wood lignins from hybrid Salix Clones.” Holzforschung 46, 505-11. [34] Sarkanen, K. V., Chang, H. M., Allan, G. G. 1967. “Species variation in lignins. III. Hardwood lignins.” Tappi 50, 587-90. [35] Karlen, S. D., Free, H. C. A., Padmakshan, D., Smith, B. G., Ralph, J., Harris, P. J. 2018. “Commelinid monocotyledon lignins are acylated by p-coumarate.” Plant Physiology 177, 513-21. [36] Lu, F., Ralph, J. 1999. “Detection and determination of p-coumaroylated units in lignins.” Journal of Agricultural and Food Chemistry 47, 1988-92. [37] Shimada, M., Fukuzuka, T., Higuchi, T. 1971. “Ester linkages of p-coumaric acid in bamboo and grass lignins.” Tappi 54, 72-78. [38] Nakamura, Y., Higuchi, T. 1976. “Ester linkage of p-coumaric acid in bamboo lignin.” Holzforschung 30, 187-91. [39] Nakamura, Y., Higuchi, T. 1978. “Ester linkage of p-coumaric acid in bamboo lignin. II. Syntheses of coniferyl p-hydroxybenzoate and coniferyl p-coumarate as possible precursors of aromatic acid esters in lignin.” Cellullose Chemistry and Technology 12, 199-208. [40] Nakamura, Y., Higuchi, T. 1978. “Ester linkage of p-coumaric acid in bamboo lignin. III. Dehydrogenative polymerization of coniferyl p-hydroxybenzoate and coniferyl p-coumarate.” Cellullose Chemistry and Technology 12, 209-21. [41] Ralph, J., Hatfield, R. D., Quideau, S., Helm, R. F., Grabber, J. H., Jung, H. J. G. 1994. “Pathway of p-coumaric acid incorporation into maize lignin as revealed by NMR.” Journal of the American Chemical Society 116, 9448-56. [42] Helm, R. F., Ralph, J. 1993. “Stereospecificity for zinc borohydride reduction of αaryloxy-β-hydroxy ketones.” Journal of Wood Chemistry and Technology 13, 593601. [43] Li, K., Helm, R. F. 1995. “Synthesis and rearrangement reactions of ester-linked lignin-carbohydrate model compounds.” Journal of Agricultural and Food Chemistry 43, 2098-103. [44] Lu, F., Ralph, J. 1997. “The DFRC method for lignin analysis. Part 1. A new method for β-aryl ether cleavage: lignin model studies.” Journal of Agricultural and Food Chemistry 45, 4655-60. [45] Lu, F., Ralph, J. 1997. “Derivatization followed by reductive cleavage (DFRC method), a new method for lignin analysis: Protocol for analysis of DFRC monomers.” Journal of Agricultural and Food Chemistry 45, 2590-92. [46] Lu, F., Ralph, J. 1998. “The DFRC method for lignin analysis. Part 3. NMR studies.” Journal of Wood Chemistry and Technology 18, 219-33.

Monolignol Acylation in Lignin Biosynthesis

45

[47] Lu, F., Ralph, J. In Lignin and Lignan Biosynthesis, Lewis, N. G., Sarkanen, S., Eds., American Chemical Society: Washington, DC, 1998. [48] Lu, F., Ralph, J. 1998. “The DFRC method for lignin analysis. Part 2. Monomers from isolated lignins.” Journal of Agricultural and Food Chemistry 46, 547-52. [49] Peng, J., Lu, F., Ralph, J. 1998. “The DFRC method for lignin analysis. Part 4. Lignin dimers isolated from DFRC-degraded loblolly pine wood.” Journal of Agricultural and Food Chemistry 46, 553-60. [50] Ralph, J., Lu, F. 1998. “The DFRC method for lignin analysis. Part 6. A modified method to determine acetate regiochemistry on native and isolated lignins.” Journal of Agricultural and Food Chemistry 46, 4616-19. [51] Lu, F., Ralph, J. 1999. “The DFRC method for lignin analysis. 7. The behavior of cinnamyl end-groups.” Journal of Agricultural and Food Chemistry 47, 1981-87. [52] Lu, F., Ralph, J. 2002. “Preliminary evidence for sinapyl acetate as a lignin monomer in kenaf.” Journal of the Chemical Society, Chemical Communications, 90-91. [53] Lu, F., Ralph, J. 2008. “Novel tetrahydrofuran structures derived from β–β-coupling reactions involving sinapyl acetate in Kenaf lignins.” Organic & Biomolecular Chemistry 6, 3681-94. [54] Lu, F., Ralph, J., Morreel, K., Messens, E., Boerjan, W. 2004. “Preparation and relevance of a cross-coupling product between sinapyl alcohol and sinapyl phydroxybenzoate.” Organic and Biomolecular Chemistry 2, 2888-90. [55] Goujon, T., Sibout, R., Pollet, B., Mabra, B., Nussaume, L., Bechtold, N., Lu, F., Ralph, J., Mila, I., Barrière, Y. et al. 2003. “A new Arabidopsis thaliana mutant deficient in the expression of O-methyltransferase impacts lignins and sinapoyl esters.” Plant Molecular Biology 51, 973-89. [56] Crestini, C., Argyropoulos, D. S. 1997. “Structural analysis of wheat straw lignin by quantitative 31P and 2D NMR spectroscopy. The occurrence of ester bonds and α-O4 substructures.” Journal of Agricultural and Food Chemistry 45, 1212-19. [57] Grabber, J. H., Quideau, S., Ralph, J. 1996. “p-Coumaroylated syringyl units in maize lignin, implications for β-ether cleavage by thioacidolysis.” Phytochemistry 43, 1189-94. [58] Das, N. N., Das, S. C., Sarkar, A. K., Mukherjee, A. K. 1984. “Lignin-xylan ester linkage in mesta fiber (Hibiscus cannabinus).” Carbohydrate. Research 129, 197207. [59] Björkman, A. 1956. “Studies on finely divided wood. Part I. Extraction of lignin with neutral solvents.” Svensk Papperstidnung 59, 477-85. [60] Lundquist, K. In Methods in Lignin Chemistry, Lin, S. Y., Dence, C. W., Eds., Springer-Verlag: Berlin, 1992. [61] Ralph, J. 1996. “An unusual lignin from Kenaf.” Journal of Natural Products 59, 341-42.

46

Fengxia Yue and Fachuang Lu

[62] del Río, J. C., Marques, G., Rencoret, J., Martinez, A. T., Gutierrez, A. 2007. “Occurrence of naturally acetylated lignin units.” Journal of Agricultural and Food Chemistry 55, 5461-68. [63] del Río, J. C., Gutierrez, A., Martinez, A. T. 2004. “Identifying acetylated lignin units in non-wood fibers using pyrolysis-gas chromatography/mass spectrometry.” Rapid Communications in Mass Spectrometry 18, 1181-85. [64] del Río, J. C., Rencoret, J., Marques, G., Gutierrez, A., Ibarra, D., Santos, J. I., Jimenez-Barbero, J., Zhang, L. M., Martinez, A. T. 2008. “Highly acylated (acetylated and/or p-coumaroylated) native lignins from diverse herbaceous plants.” Journal of Agricultural and Food Chemistry 56, 9525-34. [65] del Río, J. C., Prinsen, P., Rencoret, J., Nieto, L., Jiménez-Barbero, J., Ralph, J., Martínez, Á. T., Gutiérrez, A. 2012. “Structural characterization of the lignin in the cortex and pith of elephant grass (Pennisetum purpureum) stems.” Journal of Agricultural and Food Chemistry 60, 3619-34. [66] Rencoret, J., Ralph, J., Marques, G., Gutiérrez, A., Martínez, Á. T., del Rio, J. C. 2013. “Structural characterization of the lignin from coconut (Cocos nucifera) coir fibers.” Journal of Agricultural and Food Chemistry 61, 2434-45. [67] Withers, S., Lu, F., Kim, H., Zhu, Y., Ralph, J., Wilkerson, C. G. 2012. “Identification of a grass-specific enzyme that acylates monolignols with pcoumarate.” Journal of Biological Chemistry 287, 8347-55. [68] Li, Y., Shuai, L., Kim, H., Motagamwala, A. H., Mobley, J. K., Yue, F., Tobimatsu, Y., Havkin-Frenkel, D., Chen, F., Dixon, R. A. et al. 2018. ““Ideal lignin” facilitates full biomass utilization.” Science Advances, 4, eaau2968. [69] Rinaldi, R., Jastrzebshi, R., Clough, M. T., Ralph, J., Kennema, M., Bruijnincx, P. C. A., Weckhuysen, B. M. 2016. “Paving the way for lignin valorisation: Recent advances in bioengineering, biorefining and catalysis.” Angewandte Chemie (International Edition) 55, 8164-215. [70] Ragauskas, A. J., Beckham, G. T., Biddy, M. J., Chandra, R., Chen, F., Davis, M. F., Davison, B. H., Dixon, R. A., Gilna, P., Keller, M. et al. 2014. “Lignin valorization: Improving lignin processing in the biorefinery.” Science 344, 1246843. [71] Li, C. Z., Zhao, X. C., Wang, A. Q., Huber, G. W., Zhang, T. 2015. “Catalytic transformation of lignin for the production of chemicals and fuels.” Chemical Reviews 115, 11559-624. [72] Sun, Z., Fridrich, B., de Santi, A., Elangovan, S., Barta, K. 2018. “Bright side of lignin depolymerization: Toward new platform chemicals.” Chemical Reviews 118, 614-78. [73] Reddy, M. S. S., Chen, F., Shadle, G. L., Jackson, L., Aljoe, H., Dixon, R. A. 2005. “Targeted down-regulation of cytochrome P450 enzymes for forage quality improvement in alfalfa (Medicago sativa L.).” Proceedings of the National Academy of Sciences 102, 16573-78.

Monolignol Acylation in Lignin Biosynthesis

47

[74] Dien, B. S., Miller, D. J., Hector, R. E., Dixon, R. A., Chen, F., McCaslin, M., Reisen, P., Sarath, G., Cotta, M. A. 2011. “Enhancing alfalfa conversion efficiencies for sugar recovery and ethanol production by altering lignin composition.” Bioresource Technology 102, 6479-86. [75] Zhao, Q. 2016. “Lignification: Flexibility, Biosynthesis and Regulation.” Trends in Plant Science 21, 713-21. [76] Dien, B. S., Sarath, G., Pedersen, J. F., Sattler, S. E., Chen, H., Funnell-Harris, D. L., Nichols, N. N., Cotta, M. A. 2009. “Improved Sugar Conversion and Ethanol Yield for Forage Sorghum (Sorghum bicolor L. Moench) Lines with Reduced Lignin Contents.” Bioenergy Research 2, 153-64. [77] Vanholme, R., Ralph, J., Akiyama, T., Lu, F., Rencoret Pazo, J., Christensen, J., Rohde, A., Morreel, K., DeRycke, R., Kim, H. et al. 2010. “Engineering traditional monolignols out of lignins by concomitant up-regulation F5H1 and down-regulation of COMT in Arabidopsis.” The Plant Journal 64, 885-97. [78] Shuai, L., Amiri, M. T., Questell-Santiago, Y. M., Héroguel, F., Li, Y., Kim, H., Meilan, R., Chapple, C., Ralph, J., Luterbacher, J. S. 2016. “Formaldehyde stabilization facilitates lignin monomer production during biomass depolymerization.” Science 354, 329-33. [79] Grabber, J. H., Hatfield, R. D., Lu, F., Ralph, J. 2008. “Coniferyl ferulate incorporation into lignin enhances the alkaline delignification and enzymatic degradation of maize cell walls.” Biomacromolecules 9, 2510-16. [80] Takahama, U., Oniki, T., Shimokawa, H. 1996. “A possible mechanism for the oxidation of sinapyl alcohol by peroxidase-dependent reactions in the apoplast: Enhancement of the oxidation by hydroxycinnamic acids and components of the apoplast.” Plant and Cell Physiology 37, 499-504. [81] Takahama, U., Oniki, T. 1997. “Enhancement of peroxidase-dependent oxidation of sinapyl alcohol by an apoplastic component, 4-coumaric acid ester isolated from epicotyls of Vigna angularis L.” Plant and Cell Physiology 38, 456-62. [82] Takahama, U., Oniki, T. In Plant Peroxidases, Biochemistry and Physiology, Obinger, C., Burner, U., Ebermann, R., Penel, C., Greppin, H., Eds., Université de Genève: Genève, Switzerland, 1996. [83] Hatfield, R. D., Ralph, J., Grabber, J. H. 2008. “A potential role of sinapyl pcoumarate as a radical transfer mechanism in grass lignin formation.” Planta 228, 919-28. [84] Hatfield, R. D., Marita, J. M., Frost, K. 2008. “Characterization of p-coumarate accumulation, p-coumaroyl transferase, and cell wall changes during the development of corn stems.” Journal of the Science of Food and Agriculture 88, 2529-37.

48

Fengxia Yue and Fachuang Lu

[85] Bonawitz, N. D., Chapple, C. 2013. “Can genetic engineering of lignin deposition be accomplished without an unacceptable yield penalty?” Current Opinion in Biotechnology 24, 336-43. [86] Smith, R. A., Gonzales-Vigil, E., Karlen, S. D., Park, J. Y., Lu, F., Wilkerson, C. G., Samuels, L., Mansfield, S. D., Ralph, J. 2015. “Engineering monolignol p-coumarate conjugates into Poplar and Arabidopsis lignins.” Plant Physiology 169, 2992-3001. [87] Smith, R. A., Cass, C. L., Mazaheri, M., Sekhon, R. S., Heckwolf, M., Kaeppler, H., de Leon, N., Mansfield, S. D., Kaeppler, S. M., Sedbrook, J. C. et al. 2017. “Suppression of CINNAMOYL-CoA REDUCTASE increases the level of monolignol-ferulates incorporated into maize lignins.” Biotechnology for Biofuels 10, 109: 1-10. [88] Smith, R. A., Scheutz, M., Karlen, S. D., Bird, D., Tokunaga, N., Sato, Y., Mansfield, S. D., Ralph, J., Samuels, A. L. 2017. “Defining the diverse cell populations contributing to lignification in Arabidopsis stems.” Plant Physiology 174, 1028-36. [89] Weng, J. K., Li, X., Bonawitz, N. D., Chapple, C. 2008. “Emerging strategies of lignin engineering and degradation for cellulosic biofuel production.” Current Opinion in Biotechnology 19, 166-72. [90] Zhou, S., Runge, T., Karlen, S. D., Ralph, J., Gonzales-Vigil, E., Mansfield, S. D. 2017. “Chemical pulping advantages of Zip-lignin hybrid poplar.” ChemSusChem 10, 3565-73. [91] Bhalla, A., Bansal, N., Stoklosa, R. J., Fountain, M., Ralph, J., Hodge, D. B., Hegg, E. L. 2016. “Effective alkaline metal-catalyzed oxidative delignification of hybrid poplar.” Biotechnology for Biofuels 9, 1-10. [92] Li, Z., Chen, C. H., Hegg, E. L., Hodge, D. B. 2013. “Rapid and effective oxidative pretreatment of woody biomass at mild reaction conditions and low oxidant loadings.” Biotechnology for Biofuels 6, 119. [93] Li, Z., Chen, C. H., Liu, T., Mathrubootham, V., Hegg, E. L., Hodge, D. B. 2013. “Catalysis with Cu(II) (bpy) improves alkaline hydrogen peroxide pretreatment.” Biotechnology and Bioengineering 110, 1078-86. [94] Das, A., Rahimi, A., Ulbrich, A., Alherech, M., Motagamwala, A. H., Bhalla, A., da Costa Sousa, L., Balan, V., Dumesic, J. A., Hegg, E. L. et al. 2018. “Lignin conversion to low-molecular-weight aromatics via an aerobic oxidation-hydrolysis sequence: Comparison of different lignin sources.” ACS Sustainable Chemistry & Engineering 6, 3367-74. [95] Bhalla, A., Bansal, N., Pattathil, S., Li, M., Shen, W., Particka, C. A., Semaan, R., Gonzales-Vigil, E., Karlen, S. D., Ralph, J. et al. 2018. “Engineered lignin in poplar biomass facilitates Cu-AHP pretreatment.” ACS Sustainable Chemistry & Engineering 6, 2932-41. [96] Cass, C. L., Lavell, A. A., Santoro, N., Foster, C. E., Karlen, S. D., Smith, R. A., Ralph, J., Garvin, D. F., Sedbrook, J. C. 2016. “Cell wall composition and biomass

Monolignol Acylation in Lignin Biosynthesis

49

recalcitrance differences within a genotypically diverse set of Brachypodium distachyon inbred lines.” Frontiers in Plant Science 7, 708, 1-16. [97] Rencoret, J., Kim, H., Evaristo, A. B., Gutiérrez, A., Ralph, J., del Río, J. C. 2018. “Variability in lignin composition and structure in cell walls of different parts of macaúba (Acrocomia aculeata) palm fruit.” Journal of Agricultural and Food Chemistry 66, 138-53. [98] Ralph, J., Landucci, L. L. In Lignin and Lignans, Advances in Chemistry, Heitner, C., Dimmel, D. R., Schmidt, J. A., Eds., CRC Press (Taylor & Francis Group): Boca Raton, FL, 2010, doi: org/10.1201/EBK1574444865org/10.1201/EBK1574444865. [99] Lu, F., Karlen, S. D., Regner, M., Kim, H., Ralph, S. A., Sun, R. C., Kuroda, K. I., Augustin, M. A., Mawson, R., Sabarez, H. et al. 2015. “Naturally phydroxybenzoylated lignins in palms.” Bioenergy Research 8, 934-52. [100] Lapierre, C., Voxeur, A., Karlen, S. D., Helm, R. F., Ralph, J. 2018. “Evaluation of feruloylated and p-coumaroylated arabinosyl units in grass arabinoxylans by acidolysis in dioxane/methanol.” Journal of Agricultural and Food Chemistry 66, 5418-24. [101] del Río, J. C., Rencoret, J., Gutierrez, A., Kim, H., Ralph, J. 2018. “Structural characterization of lignin from maize (Zea mays L.) fibers: Evidence for diferuloylputrescine incorporated into the lignin polymer in maize kernels.” Journal of Agricultural & Food Chemistry 66, 4402-13.

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 3

TRICIN IN GRASS LIGNIN: BIOSYNTHESIS, CHARACTERIZATION, AND QUANTITATION Wu Lan*, Jorge Rencoret, José Carlos del Río and John Ralph Chemistry and Chemical Engineering, École polytechnique fédérale de Lausanne, Switzerland Instituto de Recursos Naturales y Agrobiología de Sevilla, Consejo Superior de Investigaciones Científicas, Seville, Spain Department of Biochemistry, and the Department of Energy’s Great Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin-Madison, Madison, WI, US

ABSTRACT Tricin [5,7-dihydroxy-2-(4-hydroxy-3,5-dimethoxyphenyl)-4H-chromen-4-one] is a member of the flavonoid family with significant biological roles in plant tissues. Even though tricin has been extensively studied as a flavonoid, the presence of tricin in the lignin polymer was only recently discovered. Differently from the other monomers, tricin is derived from a combination of the shikimate and polyketide biosynthetic pathways, and increasingly attracts attention from researchers. This chapter briefly introduces the occurrence of tricin in plants and the relevant biosynthetic pathway, discusses the identification and characterization of tricin that is incorporated into the lignin polymer, methods for its quantitation, as well as the implications of the tricin-lignin structure.

Keywords: flavonoid, flavonolignin, biosynthesis, NMR, HSQC, HMBC

*

Corresponding Author Email: [email protected]

52

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

INTRODUCTION Lignin is a complex phenylpropanoid polymer composed primarily of p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) units derived from the monolignols p-coumaryl, coniferyl, and sinapyl alcohols, respectively [1, 2]. But as our understanding of the biosynthesis and structure of lignin has escalated in the recent decade, several new lignin monomers have been discovered in wild-type and transgenic plants, such as monolignol acetate, p-hydroxybenzoate, p-coumarate, and ferulate ester conjugates that are now recognized as authentic lignin monomers [2-4], as well as components of incomplete monolignol biosynthesis such as caffeyl alcohol [5, 6], and hydroxycinnamaldehydes [7]. Recently, a special compound, tricin, was identified in the isolated lignin fraction from wheat straw [8]. Tricin [5,7-dihydroxy-2-(4-hydroxy-3,5-dimethoxyphenyl)-4H-chromen-4-one], containing benzoyl, cinnamoyl, and heterocyclic structures in its backbone, is a member of the flavone family. It has long been studied as an extractable compound (but was not known previously to be in the lignin polymer). As a secondary metabolite in cell walls, tricin is widely distributed in the leaves and stems of herbaceous and cereal plants including but not limited to bamboo, sugarcane, wheat, oat, and maize, and can be extracted from these plant tissues by organic solvents (methanol, ethanol, acetonitrile, methylene chloride) under ambient conditions or with heating [9-16].

TRICIN AS A SECONDARY METABOLITE Biosynthesis of Tricin Biosynthesis of flavones has been extensively studied, but the biosynthetic pathway of tricin remains mildly controversial. It is well established that tricin is derived from a combination of the shikimate and acetate/malonate-derived polyketide biosynthetic pathways [17] (Figure 1). The first committed step is catalyzed by chalcone synthase (CHS) using p-coumaroyl-CoA, a key intermediate on the general phenylpropanoid pathway, and malonyl-CoA (via the fatty acid pathway) as substrates. The generated naringenin chalcone is further converted to naringenin, a compound in the flavanone category, by chalcone isomerase (CHI). By dehydrogenation of the heterocyclic ring with the enzyme flavone synthase (FNSII), the flavanone is converted to a flavone, apigenin [17, 18]. Flavonoid 3',5'-hydroxylases (F3',5'H) were believed to generate tricetin from apigenin for 3',5'-O-methylation by an O-methyltransferase (FOMT) to form tricin [19-22]. However, another study suggested that chrysoeriol (a 3'-methoxylated flavone), instead of tricetin, was the intermediate for tricin synthesis in rice. Hence the biosynthetic pathway

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

53

leading to tricin should be reconstructed as being naringenin → apigenin → luteolin → chrysoeriol → selgin → tricin [23].

Figure 1. Proposed tricin biosynthetic pathway. The gray color indicates the pathway previously proposed [20,21], but that has been challenged recently [23]. PAL, pheammonialyase; C4H, cinnamate 4-hydroxylase; 4CL, 4-coumarate CoA ligase; HCT, p-hydroxycinnamoyl-CoA: quinate/shikimate p-hydroxycinnamoyltransferase; C3'H, p-coumaroyl ester 3-hydroxylase; CSE, caffeoyl shikimate esterase; CHS, chalcone synthase; CHI, chalcone isomerase; FNSII, flavone synthase II; F3'H, flavonoid 3'-hydroxylase; F5'H, flavonoid 5'-hydroxylase; FOMT, flavonoid O-methyltransferase; C5'H, chrysoeriol 5'-hydroxylase [18,24].

54

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

Tricin and Its Derivatives Tricin was first isolated in 1930 from the rust-resistant wheat (Triticum dicoccum) leaf [25]. In a later study, it was also identified in many monocotyledons species in families Poaceae [26, 27], Cyperaceae [28], Gramineae [29], and even in eudicotyledons such as alfalfa (Medicago sativa) [30]. The extractives from plants usually contained various conjugated forms, including tricin-glycosides, tricin-monolignols, and tricin-glycosidemonolignols. The natural presence of free tricin was mostly found in cereal plants, such as wheat, oat (Avena sativa), maize (Zea mays), rice (Oryza glaberrima), and barley (Hordeum vulgare). The isolated yield was significantly different from species to species. Njavara rice (Oryza sativa cv. Niavara) [31], a medicinal rice cultivated in India, contains the highest level of free tricin in the bran, with a concentration of 1931 mg/kg of whole cell wall. This content was considerably higher than in the non-medicinal rice cultivars Sujatha and Palakkadan Matta which contain only 49 and 120 mg/kg of free tricin [31]. Tricin primarily accumulates in the aerial part of the plant including straw, leaves, and husk, at different levels, and the accumulation of tricin in plants can be affected by season: a winter wheat variety contains higher level of free tricin in the husk than a spring variety [16]. Tricin-glycoside conjugates are the compounds with one or two carbohydrate units attached to tricin via either C–O ether bonds (mainly on 5-OH, 7-OH, and/or 4'-OH) or C– C bonds (on 6-C and/or 8-C) [32]. Similarly to free tricin, tricin–O–glycosides (Figure 2, compound b) are widespread across different grasses, whereas tricin-C-glycosides are not very common in plants [9]. The carbohydrates in tricin-glycosides are predominately glucose, but xylose [33], arabinose [34], rhamnose [9], and biovinose [35] are also found. Another form of tricin conjugates are the compounds that tricin links via its 4'-OH with a phenylpropanoid unit. The most common one is tricin 4'–O–(β-guaiacylglyceryl) ether (Figure 2, compound c) that is extracted with methanol along with tricin and tricinglycoside from Hyparrhenia hirta [10], oat [11], and rice [31]. These conjugates can also be in the form of a tricin-(4'–O–β)-p-coumaryl alcohol adduct [36] that has been detected in Aegilops ovate L. and maize. Additionally, coniferyl alcohol with γ-acylation by acetate or p-coumarate has also been reported in conjugation with tricin [37]. It is important to point out that, in previous studies, these conjugates were determined to be optically active and therefore termed as “flavonolignans” [11] (like their component lignan moieties which would logically be optically active [38]). However, a later study [39] demonstrated the racemic nature of these compounds. Therefore, now that tricin in lignin is known, these should be regarded as oligomers that are destined for the fully racemic lignins and should be termed as “flavonolignols”. Tricin-glycoside-phenylpropanoids are conjugates containing both carbohydrate and phenylpropanoid moieties (Figure 2, compounds d and e). Compared to tricin-glycosides and flavonolignols, these compounds have only been reported in a few plants such as

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

55

sugarcane (Saccharum officinarum) [12], rice (Zizania latifolia) [40], and Acacia nilotica [41], and only in trace amounts. The phenylpropanoid either links onto tricin directly via β–O–4' ether bonding, or onto a glucose moiety on a tricin-glycoside conjugate. Similarly, the carbohydrate etherifies with the 4'/7-OH on tricin.

Figure 2. Chemical structure of tricin a and examples of a tricin-O-glycoside b [13], tricin-Omonolignol c [10], and tricin-glycoside-phenylpropanoid d [12] and e [40]. (Modified from [39]).

Biological Functions and Potential Applications Flavonoid compounds generally function as antioxidants, antimicrobial/ antiviral agents, allelochemicals, and photoreceptors that are involved in plant growth and development [22]. Similarly, as first isolated from rust-infected wheat leaves, tricin and its related conjugates also possess important bio-functions in plant growth. A study on flavones isolated from rice leaves showed that tricin acted as an allelochemical, defending the rice against weeds by inhibiting their growth. Tricin was able to act against fungal pathogens by inhibiting their spore germination [42]. It was also involved in plant-insect interactions showing high activity against insects [43] and mosquito larvae [9], and acting as an anti-feedant against boll weevils [44]. As it is found in most of our crop plants, tricin is an important part of the human diet and has several effects on human health. It is therefore an attractive candidate for pharmacological and medicinal studies [22]. One of the most prominent and well documented properties of tricin is its potential antitumor/anticancer activity. Tricin has

56

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

been shown to inhibit the proliferation of human hepatic stellate cells [45], breast tumor cells [46-48], colon cancer cells [49], and leukemia HL-60 cells [50]. An acylated tricinglycoside isolated from sugarcane (Saccharum officinarum) juice exhibited antiproliferative activity against several human cancer cell lines, with higher selectivity toward cells of the breast-resistant NIC/ADR line [12]. Additionally, tricin is able to interfere with inflammatory-related mouse colon carcinogenesis, suggesting the potential of tricin for clinical trials of colorectal cancer chemoprevention [51]. A preliminary study on the safety of applying tricin as a chemopreventive agent reported that tricin lacked genotoxic properties in the liver, lung, heart, and kidney tissues of mice, indicating that tricin could be safe enough for clinical development [52]. A structure-activity relationship study of the flavonoids suggested that the O-methylation and glucuronidation significantly increase the cytotoxicity [53]. Similarly, a study of flavones as colorectal cancer preventive agents indicated that the rank order of cancer chemopreventive efficacy is pentamethoxyflavone > tricin > apigenin [54]. Tricin has long been credited for its health-beneficial effects as an antioxidant [55] due to its ability to suppress lipoperoxidation. A previous study showed that the antioxidant activity of tricin was lower than that of luteolin and quercetin according to the rate of lipid peroxidation [55]. However, a later study on the reaction between 2,2-diphenyl-1picrylhydrazyl radical and tricin and its conjugates suggested that tricin exhibited higher radical scavenging activity than the commonly used compounds such as myricetin, quercetin, and catechin, but lower than the tricin-monolignol dimer [56]. An oxygen radical absorbance capacity assay showed that tricin-glycoside conjugates possessed high antioxidant capacity [13]. The antioxidant ability of tricin and its derivatives was believed to be one of the reasons for their potent anti-inflammatory activities [57, 58]. Shalini et al. extracted tricin from Njavara rice (a medicinal rice cultivated in India) bran and investigated its inflammatory suppression in human peripheral blood mononuclear cells. In this study tricin showed powerful anti-inflammatory activity [59, 60]. Another study also demonstrated the inhibitory activity of tricin on the generation of inflammatory mediators in a cell line of mouse macrophages stimulated with lipopolysaccharide [57].

Preparation of Tricin Unlike most flavonoid compounds, tricin is not readily available commercially and its isolation from plants is usually in extremely low yield; a total of 8 g of tricin was isolated from 40,000 kg of the Sasa albomarginata leaves in Oyama’s study [51]. In general, the phenolic compounds including tricin and its derivatives in plant tissues are extracted using a methanol/H2O mix solvent (50:50, 80:20, or 100:0, v/v) with or without dilute acid [13, 14, 40, 61]. Some studies also apply ethanol [9], hot water [51], dichloromethane [11], butanol [43], and acetonitrile [62] as the solvent to extract crude tricin extractives. A

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

57

dewaxing step removes lipids and chlorophyll pigments by diethyl ether or hexane and can be done on the whole cell wall before the tricin solvent extraction or on the crude tricin extractives after the solvent extraction. After distillation, the condensed products are extracted by different solvents (n-hexane, diethyl ether, chloroform, dichloromethane, and butanol) to remove the compounds unrelated to tricin and thereby enrich the concentration of tricin. Such a tricin-rich fraction is further purified mainly by chromatographic techniques to give the pure tricin. An alternative way to prepare tricin in suitable quantities for experimentation and pharmacological testing is through chemical synthesis [63-65] (Figure 3). Some of the methods were based on the formation of a 1,3-diketone and followed by an intra-molecular ketone-hydroxyl reaction to form a flavone backbone (Figure 3a). Another way to make the flavone backbone is the direct condensation of appropriately protected 2,4,6trihydroxyacetophenone and 4-hydroxy-3,5-dimethoxybenzaldehyde followed by dehydrogenation and cyclization by iodine and sodium acetate [66, 67] (Figure 3b).

Figure 3. Chemical synthesis of tricin. (a) Method based on the formation of a 1,3-diketone [64]; (2) method based on the direct condensation of a ketone and an aldehyde to form the chalcone backbone [67].

OCCURRENCE OF TRICIN IN THE LIGNIN POLYMER In 2012, tricin was disclosed to be present in the milled wood lignin isolated from wheat straw for the first time [8], primarily via its characteristic HSQC NMR correlations (Figure 4). The mechanism of tricin’s incorporation into grass lignin was further investigated in a later study (Figure 5) [67]. Radical coupling reactions catalyzed by

58

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

hydrogen peroxidase showed that even though reaction rate of cross coupling between tricin and monolignols was lower than those of simple dimerization of the monolignol, by limiting their concentrations, tricin was able to form a 4'–O–β linkage with monolignols. Furthermore, the long-range correlation of C4'–Hβ was identified in the HMBC spectrum of acetylated maize stover lignin, as shown in Figure 6. Tricin moieties were also found in the high molecular weight fraction of isolated lignin according to HSQC characterization [67]. All of these data together provided solid evidence that tricin is incorporated into lignin polymers via 4'–O–β coupling.

Figure 4. Aromatic region of the short range 1H–13C correlation (2D HSQC) NMR spectrum of isolated milled wood lignin (MWL) from wheat straw cell walls (A) and long range 1H–13C correlation (2D HMBC) NMR spectrum of MWL from wheat straw cell wall showing the main correlations of tricin units in lignin. (Modified from [8]).

Additional studies verified the presence of tricin in lignin preparations from various monocots, including Carex meyeriana [68], bamboo (Phyllostachys pubescens) [69], coconut (Cocos nucifera) coir [70], giant cane (Arundo donax) [71], rice [72], barley [73], sugarcane [74], and V. planifolia [6]. The pretreatment methods to isolate lignin could also affect the presence of tricin because tricin is not stable under harsh conditions.

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

Figure 5. Radical coupling reaction between tricin and a monolignol.

Figure 6. 2D HMBC spectrum of acetylated maize lignin [67]. Occurrence of the C4'–Hβ correlation demonstrates that tricin is incorporated into the lignin polymer via radical coupling and forming a 4'–O–β aryl ether bond.

59

60

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

Figure 7. Aromatic region of the 2D HSQC spectra of wheat straw lignin. Samples were pretreated with water at 160°C (A), 0.25 wt% H2SO4 aqueous solution at 160°C (B), and 1.0 wt% H2SO4 aqueous solution at 160°C (C) (modified from [75]). Tricin was completely depleted under higher acid concentration pretreatments (C).

For example, during dilute acid pretreatment of wheat straw, tricin was mostly retained under 160°C with 0.25% H2SO4 in water, whereas all of the tricin vanished when the acid concentration was increased to 1%, or when the pretreatment temperature was increased to 190°C (even without any acid) [75] (Figure 7). Similarly, steam explosion treatment at 200°C for only 10 minutes removed tricin from the lignin polymer significantly [76]. Alkaline conditions also affected the present of tricin even at low temperature. The content of tricin was much lower in the lignin isolated from alkaline pretreated wheat straw than that from the untreated sample [77]. On the other hand, some processes such as mild acid γ-valerolactone pretreatment [78, 79] and extractive-ammonia pretreatment [80] largely preserved the tricin moiety in the lignin polymer from corn stover. However, most of the above studies did not mention the reason for the absence of tricin. It is unclear whether it was cleaved from the polymer as an intact moiety or was degraded into other products. It is reported that the distribution of tricin in lignin polymer chains varied according to the molecular weight: the lower molecular weight fractions contained higher amounts of tricin, whereas in higher molecular weight fractions the tricin level was lower, and even no tricin units were found in the highest molecular weight fraction [81]. In contrast, our study showed that tricin moieties were approximately equally distributed in the lignin fractions of different molecular weights from corn stover [39]. Such differences might be due to the different lignin isolation methods applied in these two studies. Another study showed that different lignin-carbohydrate complex (LCC) fractions contained different levels of tricin [77]. The LCC is a lignin-cell wall cross-linked fraction in which ferulate (FA) presumably functions as a nucleation site. The FA acylates arabinose side-chains of arabinoxylans and radical coupling with monolignols can initiate the growth of lignin polymer chains. Zikeli et al. isolated two different LCC fractions; one was predominantly associated with glucan and the other one was mainly bound with xylan. It is surprising to find that the glucan-rich

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

61

LCC contained remarkably higher levels of tricin units than the xylan-rich LCC fraction did [77].

Characterization of Tricin NMR spectroscopy has enormously facilitated the investigations into structural aspects of complex lignin polymers [82, 83]. 1H NMR is the most widely applied NMR technique. But lignin appears broad and featureless in 1H NMR spectra in most of cases, not only because of lignin’s high molecular weight, but also the irregular bonding between monomers and, most importantly, the complexity caused by its stereochemical diversity. However, simple 1H NMR spectroscopy is able to identify the presence of tricin in lignin polymer because the tricin moiety shows a characteristic peak at 13.0 ppm, corresponding to 5-OH, that is very different from anything else in the lignin structure [84]. Compared to the 1D NMR techniques (1H and 13C NMR), the heteronuclear singlequantum coherence (HSQC) technique (a 2D heteronuclear NMR method) provides much more comprehensive information of inter-unit linkages and therefore is frequently applied for lignin characterization. The tricin moiety in the lignin polymer can be easily identified in HSQC spectroscopy because of its four characteristic correlations in the aromatic region, which do not overlap with the peaks from other lignin structures. The two correlation peaks at δC/δH 99.0/6.21 and 94.4/6.57 correspond to the C6/H6 and C8/H8 on aromatic ring (ring A) and the peak at δC/δH 104.9/7.05 is from the C3/H3 on the C = C double bond (heterogeneous ring C). The C2'/H2'(C6'/H6') correlation of the symmetric aromatic ring (ring B) raises a peak at δC/δH 104.0/7.30. After acetylation the peaks of C3/H3, C6/H6, and C8/H8 move to δC/δH 107.8/7.05, 110.2/7.71, and 114.3/7.07, whereas the peak of C2'/H2'(C6'/H6') does not change [67]. The chemical shift of C3/H3 indicates whether the tricin moiety is free or incorporated into the polymer via a 4'-O aryl ether bond. When the 4'-O-β bond is cleaved, the peak of C3/H3 slightly moves to downfield (δC/δH 103.5/7.04, Figure 8). All of the abovementioned chemical shifts are on the basis of using DMSO-d6 as solvent. When using CDCl3 or acetone-d6 as the solvent the chemical shifts of these peaks would be changed, especially in the 1H dimension. Table 1 summarizes the chemical shifts of the characteristic peaks of tricin (non-acetylated and acetylated) using different deuterated NMR solvents. 31 P NMR is another technique applied to characterize lignin, mainly focusing on qualitatively and quantitatively analyzing the various hydroxyl groups. It was also applied to characterize the tricin unit in lignin. The three hydroxyl groups (4'-OH, 5-OH, and 7OH) in tricin generate three peaks in the 31P NMR spectrum at δP 142.0, 137.6, and 136.4 ppm, respectively [76]. In the case of the 31P NMR spectrum of tricin-integrated lignin (enzymatic mild acidolysis lignin from wheat straw), two peaks at 137.6 and 136.4 ppm

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

62

originated from the 5-OH and 7-OH with different intensities indicating an overlap of the peaks from –OH on tricin and an –OH from another structure. Table 1. Selected diagnostic tricin unit chemical shifts from tricin-(4'–O–β)monolignol dimers in acetone-d6, CDCl3, and DMSO-d6. (Modified from [67])

Acetone-d6 CDCl3 DMSO-d6 Acetone-d6 CDCl3 DMSO-d6

Unacetylated (δC/δH) C3/H3 106.01/6.82 105.83/6.61 104.87/7.05 Acetylated (δC/δH) 108.67/6.78 108.36/6.60 107.82/7.05

C6/H6 99.79/6.27 99.75/6.33 98.98/6.21

C8/H8 94.98/6.56 94.34/6.49 94.37/6.57

110.29/7.49 109.08/7.39 110.18/7.71

114.80/6.95 113.65/6.84 114.33/7.07

Figure 8. Aromatic region of the 2D HSQC spectrum of enzymatic lignin from maize. The blue dashed ellipsoids indicate the chemical shifts of free tricin itself. The C3-H3 correlation is especially different between free tricin and 4'-O etherified tricin. (Modified from [67]).

There is no peak at 142.0 ppm from the 4'-OH on tricin because this hydroxyl group is etherified in the lignin. The spectrum of lignin isolated from a sample treated with steam explosion shows that the two sharp peaks from tricin disappear demonstrating that tricin was both cleaved and degraded during pretreatment [76]. Ragauskas et al. systematically investigated the application of 31P NMR to identify tricin. By comparing the 31P NMR spectrum of tricin-incorporated lignin from Zea mays and non-tricin-incorporated lignin from Populous trichocarpa with the spectra of flavonoids, it was found that 31P NMR did

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

63

provide diagnostic peaks for tricin and other flavonoids. However, again, cautions should be taken when applying such methods for quantitative studies due to peak overlap issues [85]. Another method to characterize tricin in lignin is using wet chemical degradative methods followed by chromatographic characterization. It should be noted that tricin is not able to be detected in gas chromatography (GC) because it is not sufficiently volatile, even when derivatized, under the temperature conditions applicable. Although the volatility of the compound can be improved by trimethylsilylation, the per-trimethylsilyl (TMS) tricin (with TMS on the 5, 7, and 4' hydroxyl groups) was not stable under the GC conditions and partially degraded into di- and mono-TMS tricin. Therefore, liquid chromatography with mass spectrometric (LC-MS) detection is more suitable for tricin-integrated lignin characterization. Tricin is able to be protonated and deprotonated, so it can be detected in both positive- and negative-ion mode, with mass to charge ratios (m/z) of 331 [M+H]+ and 329 [M-H]-, respectively. In tandem MS spectrometry (positive-ion mode), 331 [M+H]+ generates different ion fragments under specific collision energies. The fragments m/z 315 and 300 correspond to cleavage of one and two methoxyl groups. The cleavage of C2–C1' bond forms two ion fragments with m/z as 177 and 153 originating from the ring AC (chalcone backbone) and ring B (syringyl unit).

Figure 9. 31P NMR spectra of wheat straw enzymatic mild acidolysis lignin (EMAL, A), steam explosion-treated wheat straw EMAL (B), and tricin itself showing its three hydroxyl groups (C). (Adapted from [76]).

Quantitation of Tricin As tricin has been proven to be a monomer in lignin, quantitation of the tricin moiety in the lignin polymer became an important target. The proportions of tricin were roughly

64

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

estimated to be 10-15% of wheat straw lignin on the basis of volume integration of diagnostic contours in HSQC spectra. It was, however, thought to be excessive because it is well known that end-groups are over-quantified by such methods that are semiquantitative at best [86]. A suitable way to quantify tricin is to cleave tricin from the polymer by wet chemical degradative methods followed by LC(-MS) quantitation. Our group evaluated the efficiencies of cleaving the tricin-(4'–O–β)-ether bonds and the degradation of tricin under acidolysis, thioacidolysis, and derivatization followed by reductive cleavage (DFRC) methods [39]. To further increase the accuracy, a deuterated tricin (tricin-d6) was synthesized and used as an internal standard to correct not only sample variations during reaction and work up process, but to compensate for the variability in chromatographic separation, ionization, and MS detection [87, 88]. Thioacidolysis was found to be the best method as it produced a 96 mol% yield of tricin from tricin-(4'–O–β)coniferyl alcohol. We then screened various seed-plant species and, when present, quantified tricin content in lignin using the thioacidolysis method followed by LC-MS characterization. Oat, wheat, and brachypodium showed the highest amount of tricin as 33.1, 32.7, and 28.0 mg/g of lignin, respectively. All Poaceae samples examined in our study showed the presence of tricin. Some other species, such as Arecaceae, Orchidaceae, and even Fabaceae in Eudicotyledons contained some amount of tricin in their lignin polymers. Compared to the amount of extractable free tricin, the amounts of tricin in the lignins were higher. The bran of Njavara rice (O. sativa cv. Niavara), a medicinal rice cultivated in India, was reported to contain the highest extractable tricin level (1931 mg/kg) [31]. Wheat husk, a more abundant source, contained the second highest tricin level (777 mg/kg) among plants that have been examined [16]. The wheat sample used in our study contained only 376 mg/kg of extractable tricin, whereas the content of lignin-integrated tricin was 4841 mg/kg [39]. The difference was much more substantial when the tricin level was calculated on a lignin basis. Therefore, lignin residue from certain biomass pretreatment processes could be a potential source of large amounts of tricin if a feasible and industrially relevant method to liberate tricin is available.

Bioengineering of Tricin Lignin biosynthesis and bioengineering have attracted significant attention from researchers because it may play an important role in economically improving applications of agroindustrial biomass. Lignin has long been treated as an impediment to chemical pulping, forage digestion by livestock, and is a side-product of cellulosic biorefinery plants. But it has been increasingly recognized as potential source of aromatic bulk commodity chemicals. As tricin has been disclosed as one of the monomers in grass lignin, manipulation of the biosynthetic pathway responsible for flavone biosynthesis and

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

65

investigation of the consequences on cell wall composition, recalcitrance, and lignin structure has become an interesting research focus. Branching off from p-coumaroyl-CoA, flavonoids and monolignols are the two major downstream metabolite classes. The flux toward flavone biosynthesis and, thus, to the tricin monomer, is controlled by CHS. In maize, Colorless 2 (C2) is the gene encoding CHS and expressed in many parts of the plant including the pericarp, tassels, and vegetative organs. Therefore, disruption of C2 would cause the depletion of flavonoids, including tricin in stems and leaves of maize. In our previous study [89], CHS-deficient plants showed strikingly lower abundance of flavone metabolites compared to the C2 control plants and no tricin monomer was detected in the lignin polymer. The stems of control and mutant plants contained similar Klason lignin contents, whereas the leaf of C2 mutant maize showed higher lignin content than the control. Accordingly, the cell wall recalcitrance to enzymatic saccharification were more profound for the leaves of C2 mutants than that of controls, but quite similar for the stem samples. Table 2. Comparison of extractable tricin and lignin-integrated tricin (mg/kg) from non-extracted plant material. (Modified from [39]) Extractable tricin vs Lignin-integrated tricin Plant sample Extractable tricin Wheat straw 376.1 ± 69.5 Maize straw 90.9 ± 12.4 Oat straw 601.4 ± 48.4 Rice stem 64.1 ± 6.9 Extractable tricin reported in previous studies Plant species Part Njavara rice (Oryza sativa cv. Niavara) [31] Rice Oryza sativa cv. Sujatha [31] Rice Oryza sativa cv. Palakkadam Matta [31] Wheat (a winter cultivar) [16]

bran bran bran husk leaves bran

Extractable T-(4'–O–β)-G 1044.3 ± 149.9 299.7 ± 39.9 1270.0 ± 49.7 215.3 ± 28.1

Lignin-integrated tricin 4841.3 ± 217.8 1304.0 ± 39.6 5250.3 ± 121.9 979.7 ± 0.1

Extractable tricin 1930.5 ± 0.3 48.6 ± 0.1 119.8 ± 0.1 772 ± 31.8 253 ± 18.3 33 ± 15.9

Extractable T-(4'–O–β)-G 1217.7 ± 1.2 45.9 ± 0.9 Not detected Not reported Not reported Not reported

Flavone synthase II (FNSII) is the enzyme that catalyzes the direct conversion of flavanones to flavones. It is indispensable for the biosynthesis of both extractable tricinderived metabolites [90] and tricin monomers for lignification in rice vegetative tissues [24]. In the culm, sheath, and leaf tissue of rice, disrupting FNSII significantly altered the cell wall properties, such as reducing the lignin content and decreasing the S/G ratio in lignin. HSQC characterization of the lignin from FNSII mutant plants indicated the absence of tricin, and a small amount of naringenin units was integrated into the lignin polymer (Figure 11). Additionally, FNSII-knockout mutants exhibited better saccharification

66

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

efficiency than the control samples and did not display negative impacts on the growth and development of vegetative tissues [24].

Figure 10. Content of tricin in the lignins from different plant species. The error bars were calculated on the basis of standard deviation (Adapted from [39]).

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

67

Figure 11. Aromatic regions of 2D HSQC spectra of cell wall lignins from culm tissues of wild-type (WT, A) and FNSII-knockout mutant (B) rice plants. Lignin samples were prepared by enzymatic removal of cell wall polysaccharides with cellulase. In the lignin from fnsII mutant plants, tricin was completely depleted and naringenin units were evident. (Modified from [24]).

After the conversion of naringenin by FNSII to apigenin, substitutions on the B ring of the C6-C3-C6 flavan skeleton are catalyzed by flavonoid 3'-hydroxylases (F3'H) and Omethyltransferase (OMT) to produce flavonoids. CYP75B3 and CYP75B4 are the two enzymes in rice tissue with flavonoid 3'-hydroxylase activities [23,91,92]. By studying the metabolite profiles, cell wall properties, and lignin structure of the cyp75b3, cyp75b4 mutant and cyp75b3 cyp75b4 double-mutant plants, CYP75B3 was proved to be solely responsible for the biosynthesis of 3'-subsituted flavone C-glycosides. Disrupting CYP75B3 did not affect the lignin structure and cell wall properties. On the other hand, CYP75B4 was demonstrated to process both apigenin 3'-hydroxylation and chrysoeriol 5'hydroxylation activity corresponding to the production of lignin-bound tricin and tricin Oconjugates. Similarly to the FNSII mutant, a knockout in CYP75B4 decreased the S/G lignin unit composition and remarkably reduced the lignin levels, accordingly enhancing the carbohydrates digestibility. As expected, cyp75b4 mutant plants produced tricindepleted lignin. In culm tissue, apigenin, instead of tricin, was found to be integrated in lignin polymer [93].

68

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

Caffeoyl coenzyme A 3-O-methyltransferase (CCoAOMT) and caffeic acid-Omethyltransferase (COMT) were well documented as the important enzymes for the biosynthesis of monolignols. Downregulation or knockout of COMT or both enzymes enriched the G units in lignins and reduced the total lignin content [94-97]. A recent study also demonstrated that a comt mutant maize produced tricin-depleted lignin in stem tissue, whereas the tricin level was not significantly affected in the midrib. This observation suggested that COMT was also involved in the tricin biosynthetic pathway, at least in the stem tissue [98].

Implications of Tricin’s Presence in Lignin In maize metabolite profiling, coniferyl and sinapyl alcohol and their acetate and pcoumarate conjugates were all found to couple with tricin. Furthermore, using a chiral column, we were able to separate two enantiomers of the tricin-(4'–O–β)-coniferyl alcohol and tricin-(4'–O–β)-p-coumaryl alcohol using LC-MS/MS with multiple reaction monitoring (MRM). The identical peak areas of the two enantiomers indicated the racemic nature of the tricin conjugates [37]. In the case of the tricin-(4'–O–β)-coniferyl alcohol-(4'– O–β)-coniferyl alcohol trimer, 6 of the 8 isomers were separated and identified in LCMS/MS, suggesting the compounds were formed by simple radical reactions [37]. As in the established theory and as increasingly evidenced, lignins are the products of simple, but combinatorial, radical coupling chemistry [99,100]. However, such a concept was heatedly debated after notions of absolute proteinaceous control over lignin structure were championed for a period [101], but were eventually convincingly debunked [102]. The fact that tricin cross-couples with acetate and p-coumarate monolignol conjugates, the racemic nature of flavonolignols, and the diversity of the diastereomers support the combinatorial radical coupling theory, demonstrating that that lignins are racemic polymers, are characterized by being products with a huge number of possible isomers, and have no defined sequence or (repeating) structure. Biomimetic radical coupling reactions between tricin and monolignols and characterization of lignin polymer from nature products indicate that tricin only incorporated into the lignin polymer in the form of 4'–O–β-coupled products and their higher oligomers [67]. In this case, the tricin unit must be localized at one terminus of its lignin chain, and that terminus must be at the starting end of that chain. In other words, tricin acts as a nucleation site for lignin chain growth in monocots, a role that was proposed for ferulate on arabinoxylans [103]. Such an observation contributes to resolving a monocot lignin structural dilemma that has existed for decades: that monocot lignins (especially maize lignin), unlike other syringyl-guaiacyl lignins in dicots/hardwoods, have essentially no, or very low levels of resinols (β–β-coupled units) [67,96]. Such β–β units are produced only as the result of monolignol (sinapyl alcohol) dimerization and are the obvious

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

69

mechanism for starting a lignin chain. One of the reasons is that sinapyl p-coumarate homodimerization, instead of sinapyl alcohol coupling, is more preponderant to function as the starting point. Also, when the lignin chain is nucleated by another unit, such as tricin or ferulate, lignification does not need to start with a dimerization reaction.

CONCLUSION Tricin, a member in flavonoid family with a C6-C3-C6 backbone, is a novel monomer in lignin discovered in recent years. Not only does it arise from a different biosynthetic pathway from the traditional monolignols, it operates in a polymer chain nucleation function; its biological functions and potential pharmaceutical applications increasingly attract research interest. Studies related to the biosynthesis and bioengineering, identification, characterization, and quantification of tricin are providing new insight into the lignin structure. The lignin-bound tricin also provides a new and abundant source of such a high-market-priced chemical, encouraging research into its production methods from lignin polymers and its applications beyond its current roles.

ACKNOWLEDGMENTS The authors thank the China Scholarship Council, State Education Department, for supporting living expenses for Wu Lan’s PhD Program in the Department of Biological System Engineering, University of Wisconsin, Madison, USA. WL, and JRa were funded by the DOE Great Lakes Bioenergy Research Center (DOE Office of Science BER DEFC02-07ER64494 and DE-SC0018409). JRe and JdR was funded by the Spanish Project CTQ2014-60764-JIN (co-financed by FEDER funds).

REFERENCES [1] [2] [3]

Boerjan, W.; Ralph, J.; Baucher, M. 2003. “Lignin biosynthesis.” Annual Review of Plant Biology 54, 519-46. Ralph, J. 2010. “Hydroxycinnamates in lignification.” Phytochemistry Reviews 9, 65-83. Karlen, S. D.; Smith, R. A.; Kim, H.; Padmakshan, D.; Bartuce, A.; Mobley, J. K.; Free, H. C. A.; Smith, B. G.; Harris, P. J.; Ralph, J. 2017. “Highly decorated lignins in leaf tissues of the Canary Island date palm Phoenix canariensis.” Plant Physiology 175, 1058-67.

70 [4]

[5]

[6]

[7]

[8]

[9]

[10] [11]

[12]

[13]

[14]

[15]

Wu Lan, Jorge Rencoret, José Carlos del Río et al. Karlen, S. D.; Zhang, C. C.; Peck, M. L.; Smith, R. A.; Padmakshan, D.; Helmich, K. E.; Free, H. C. A.; Lee, S.; Smith, B. G.; Lu, F. C. et al. 2016. “Monolignol ferulate conjugates are naturally incorporated into plant lignins.” Science Advances 2. Chen, F.; Tobimatsu, Y.; Jackson, L.; Nakashima, J.; Ralph, J.; Dixon, R. A. 2013. “Novel seed coat lignins in the Cactaceae: structure, distribution and implications for the evolution of lignin diversity.” Plant Journal 73, 201-11. Chen, F.; Tobimatsu, Y.; Havkin-Frenkel, D.; Dixon, R. A.; Ralph, J. 2012. “A polymer of caffeyl alcohol in plant seeds.” Proceedings of the National Academy of Sciences of the United States of America 109, 1772-77. Zhao, Q.; Tobimatsu, Y.; Zhou, R.; Pattathil, S.; Gallego-Giraldo, L.; Fu, C.; Jackson, L. A.; Hahn, M. G.; Kim, H.; Chen, F. et al. 2013. “Loss of function of cinnamyl alcohol dehydrogenase 1 leads to unconventional lignin and a temperaturesensitive growth defect in Medicago truncatula.” Proceedings of the National Academy of Sciences of the United States of America 110, 13660-65. del Río, J. C.; Rencoret, J.; Prinsen, P.; Martinez, A. T.; Ralph, J.; Gutierrez, A. 2012. “Structural characterization of wheat straw lignin as revealed by analytical pyrolysis, 2D-NMR, and reductive cleavage methods.” Journal of Agricultural and Food Chemistry 60, 5922-35. Ju, Y.; Sacalis, J. N.; Still, C. C. 1998. “Bioactive flavonoids from endophyteinfected blue grass (Poa ampla).” Journal of Agricultural and Food Chemistry 46, 3785-88. Bouaziz, M.; Veitch, N. C.; Grayer, R. J.; Simmonds, M. S. J.; Damak, M. 2002. “Flavonolignans from Hyparrhenia hirta.” Phytochemistry 60: 515–520. Wenzig, E.; Kunert, O.; Ferreira, D.; Schmid, M.; Schuhly, W.; Bauer, R.; Hiermann, A. 2005. “Flavonolignans from Avena sativa.” Journal of Natural Products 68, 28992. Duarte-Almeida, J. M.; Negri, G.; Salatino, A.; de Carvalho, J. E.; Lajolo, F. M. 2007. “Antiproliferative and antioxidant activities of a tricin acylated glycoside from sugarcane (Saccharum officinarum) juice.” Phytochemistry 68, 1165-71. Van Hoyweghen, L.; Karalic, I.; Van Calenbergh, S.; Deforce, D.; Heyerick, A. 2010. “Antioxidant flavone glycosides from the leaves of Fargesia robusta.” Journal of Natural Products 73, 1573-77. Nakano, H.; Kawada, N.; Yoshida, M.; Ono, H.; Iwaura, R.; Tonooka, T. 2011. “Isolation and identification of flavonoids accumulated in proanthocyanidin-free barley.” Journal of Agricultural and Food Chemistry 59, 9581-87. Bottcher, A.; Cesarino, I.; dos Santos, A. B.; Vicentini, R.; Mayer, J. L. S.; Vanholme, R.; Morreel, K.; Goeminne, G.; Moura, J. C. M. S.; Nobile, P. M. et al. 2013. “Lignification in sugarcane: Biochemical characterization, gene discovery,

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

[16]

[17] [18]

[19]

[20]

[21]

[22] [23]

[24]

[25] [26] [27]

[28]

[29]

71

and expression analysis in two genotypes contrasting for lignin content.” Plant Physiology 163, 1539-57. Moheb, A.; Grondin, M.; Ibrahim, R. K.; Roy, R.; Sarhan, F. 2013. “Winter wheat hull (husk) is a valuable source for tricin, a potential selective cytotoxic agent.” Food Chemistry 138, 931-37. Winkel-Shirley, B. 2001. “Flavonoid biosynthesis. A colorful model for genetics, biochemistry, cell biology, and biotechnology.” Plant Physiology 126, 485-93. Morreel, K.; Goeminne, G.; Storme, V.; Sterck, L.; Ralph, J.; Coppieters, W.; Breyne, P.; Steenackers, M.; Georges, M.; Messens, E. et al. 2006. “Genetical metabolomics of flavonoid biosynthesis in Populus: a case study.” Plant Journal 47, 224-37. Zhou, J. M.; Fukushi, Y.; Wang, X. F.; Ibrahim, R. K. 2006. “Characterization of a novel flavone O-methyltransferase gene in rice.” Natural Product Communications 1, 981-84. Zhou, J. M.; Gold, N. D.; Martin, V. J. J.; Wollenweber, E.; Ibrahim, R. K. 2006. “Sequential O-methylation of tricetin by a single gene product in wheat.” Biochimica Et Biophysica Acta-General Subjects 1760, 1115-24. Zhou, J. M.; Fukushi, Y.; Wollenweber, E.; Ibrahim, R. K. 2008. “Characterization of two O-methyltransferase-like genes in barley and maize.” Pharmaceutical Biology 46, 26-34. Zhou, J.-M.; Ibrahim, R. K. 2010. “Tricin—a potential multifunctional neutraceutical.” Phytochemistry Reviews 9, 413-24. Lam, P. Y.; Liu, H. J.; Lo, C. 2015. “Completion of tricin biosynthesis pathway in rice: Cytochrome P450 75B4 is a unique chrysoeriol 5'-hydroxylase.” Plant Physiology 168, 1527-U760. Lam, P. Y.; Tobimatsu, Y.; Takeda, Y.; Suzuki, S.; Yamamura, M.; Umezawa, T.; Lo, C. 2017. “Disrupting flavone synthase II alters lignin and improves biomass digestibility.” Plant Physiology 174, 972-85. Anderson, J. A.; Perkin, A. G. 1931. “The yellow colouring matter of Khapli wheat, Triticum dicoccum.” Journal of Chemical Society [London] 1931, 2624-25. Harborne, J. B.; Hall, E. 1964. “Plant polyphenols. XII. The occurrence of tricin and of glycoflavones in grasses.” Phytochemistry 3, 421-28. Kuwazuka, S.; Oshima, Y. 1964. “Studies on polyphenols in rice plant. III. Isolation and determination of tricin-glycosides “glucotricin” and “tricinin”.” Agricultural and Biological Chemistry 28, A31-A31. Harborne, J. B. 1971. “Comparative biochemistry of flavonoids. 16. distribution and taxonomic significance of flavonoids in leaves of Cyperaceae.” Phytochemistry 10, 1569-&. Harborne, J. B.; Williams, C. A. 1976. “Flavonoid patterns in leaves of the Gramineae.” Biochemical Systematics and Ecology 4, 267-80.

72

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[30] Bickoff, E. M.; Livingston, A. L.; Booth, A. N. 1964. “Tricin from alfalfa: Isolation and physiological activity.” Journal of Pharmaceutical Sciences 53, 1411-12. [31] Mohanlal, S.; Parvathy, R.; Shalini, V.; Helen, A.; Jayalekshmy, A. 2011. “Isolation, characterization and quantification of tricin and flavonolignans in the medicinal rice Njavara (Oryza sativa L.), as compared to staple varieties.” Plant Foods for Human Nutrition 66, 91-96. [32] Ferreres, F.; Gil-Izquierdo, A.; Andrade, P. B.; Valentao, P.; Tomas-Barberan, F. A. 2007. “Characterization of C-glycosyl flavones O-glycosylated by liquid chromatography-tandem mass spectrometry.” Journal of Chromatography A 1161, 214-23. [33] Theodor, R.; Zinsmeister, H. D.; Mues, R.; Markham, K. R. 1981. “Flavone Cglycosides of 2 Metzgeria species.” Phytochemistry 20, 1851-52. [34] Parveen, M.; Khanam, Z.; Ali, A.; Ahmad, S. M. 2010. “A novel antimicrobial flavonoidic glycoside from the leaves of Alstonia macrophylla Wall ex A. DC (Apocynaceae).” Chinese Chemical Letters 21, 593-95. [35] Sun, J.; Yue, Y. D.; Tang, F.; Guo, X. F.; Wang, J.; Yao, X. 2013. “Flavonoids from the Leaves of Neosinocalamus affinis.” Chemistry of Natural Compounds 49, 82225. [36] Cooper, R.; Gottlieb, H. E.; Lavie, D. 1977. “New flavolignan of biogenetic interest from Aegilop Ovata L. 1.” Israel Journal of Chemistry 16, 12-15. [37] Lan, W.; Morreel, K.; Lu, F.; Rencoret, J.; del Río, J. C.; Voorend, W.; Vermerris, W.; Boerjan, W.; Ralph, J. 2016. “Maize tricin-oligolignol metabolites and their implications for monocot lignification.” Plant Physiology 171, 810-20. [38] Umezawa, T. 2004. “Diversity in lignan biosynthesis.” Phytochemistry Reviews 2, 371-90. [39] Lan, W.; Rencoret, J.; Lu, F. C.; Karlen, S. D.; Smith, B. G.; Harris, P. J.; del Río, J. C.; Ralph, J. 2016. “Tricin-lignins: occurrence and quantitation of tricin in relation to phylogeny.” Plant Journal 88, 1046-57. [40] Lee, S. S.; Baek, N. I.; Baek, Y. S.; Chung, D. K.; Song, M. C.; Bang, M. H. 2015. “New flavonolignan glycosides from the aerial parts of Zizania latifolia.” Molecules 20, 5616-24. [41] Khanam, Z.; Adam, F.; Singh, O.; Ahmad, J. 2011. “A novel acylated flavonoidic glycoside from the wood of cultivated Acacia nilotica (L.) Willd. Ex. Delile.” Bioresources 6, 2932-40. [42] Kong, C. H.; Xu, X. H.; Zhou, B.; Hu, F.; Zhang, C. X.; Zhang, M. X. 2004. “Two compounds from allelopathic rice accession and their inhibitory activity on weeds and fungal pathogens.” Phytochemistry 65, 1123-28. [43] Adjei-Afriyie, F.; Kim, C. S.; Takemura, M.; Ishikawa, M.; Horiike, M. 2000. “Isolation and identification of the probing stimulants in the rice plant for the white-

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

[44]

[45]

[46]

[47] [48]

[49]

[50]

[51]

[52]

[53]

73

back planthopper, Sogatella furcifera (Homoptera: Delphacidae).” Bioscience Biotechnology and Biochemistry 64, 443-46. Miles, D. H.; Tunsuwan, K.; Chittawong, V.; Kokpol, U.; Choudhary, M. I.; Clardy, J. 1993. “Boll weevil antifeedants from arundo donax.” Phytochemistry 34, 127779. Seki, N.; Toh, U.; Kawaguchi, K.; Ninomiya, M.; Koketsu, M.; Watanabe, K.; Aoki, M.; Fujii, T.; Nakamura, A.; Akagi, Y. et al. 2012. “Tricin inhibits proliferation of human hepatic stellate cells in vitro by blocking tyrosine phosphorylation of PDGF receptor and its signaling pathways.” Journal of Cellular Biochemistry 113, 234655. Cai, H.; Hudson, E. A.; Mann, P.; Verschoyle, R. D.; Greaves, P.; Manson, M. M.; Steward, W. P.; Gescher, A. J. 2004. “Growth-inhibitory and cell cycle-arresting properties of the rice bran constituent tricin in human-derived breast cancer cells in vitro and in nude mice in vivo.” British Journal of Cancer 91, 1364-71. Yan, J.; Sun, L. R.; Zhang, X. M.; Qiu, M. H. 2005. “A new flavone from Lycopodium japonicum.” Heterocycles 65, 661-66. Jeong, Y. H.; Chung, S. Y.; Han, A. R.; Sung, M. K.; Jang, D. S.; Lee, J.; Kwon, Y.; Lee, H. J.; Seo, E. K. 2007. “P-glycoprotein inhibitory activity of two phenolic compounds, (-)-syringaresinol and tricin from Sasa borealis.” Chemistry & Biodiversity 4, 12-16. Hudson, E. A.; Dinh, P. A.; Kokubun, T.; Simmonds, M. S. J.; Gescher, A. 2000. “Characterization of potentially chemopreventive phenols in extracts of brown rice that inhibit the growth of human breast and colon cancer cells.” Cancer Epidemiology Biomarkers & Prevention 9, 1163-70. Ninomiya, M.; Nishida, K.; Tanaka, K.; Watanabe, K.; Koketsu, M. 2013. “Structure-activity relationship studies of 5,7-dihydroxyflavones as naturally occurring inhibitors of cell proliferation in human leukemia HL-60 cells.” Journal of Natural Medicines 67, 460-67. Oyama, T.; Yasui, Y.; Sugie, S.; Koketsu, M.; Watanabe, K.; Tanaka, T. 2009. “Dietary tricin suppresses inflammation-related colon carcinogenesis in male Crj: CD-1 mice.” Cancer Prevention Research 2, 1031-38. Verschoyle, R. E.; Greaves, P.; Cai, H.; Arndt, B.; Broggini, M.; D'Incalci, M.; Riccio, E.; Doppalapudi, R.; Kapetanovic, I. M.; Steward, W. P. et al. 2006. “Preliminary safety evaluation of the putative cancer chemopreventive agent tricin, a naturally occurring flavone.” Cancer Chemotherapy and Pharmacology 57, 1-6. Plochmann, K.; Korte, G.; Koutsilieri, E.; Richling, E.; Riederer, P.; Rethwilm, A.; Schreier, P.; Scheller, C. 2007. “Structure-activity relationships of flavonoidinduced cytotoxicity on human leukemia cells.” Archives of Biochemistry and Biophysics 460, 1-9.

74

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[54] Cai, H.; Sale, S.; Schmid, R.; Britton, R. G.; Brown, K.; Steward, W. P.; Gescher, A. J. 2009. “Flavones as colorectal cancer chemopreventive agents-phenol-Omethylation enhances efficacy.” Cancer Prevention Research 2, 743-50. [55] Watanabe, M. 1999. “Antioxidative phenolic compounds from Japanese barnyard millet (Echinochloa utilis) grains.” Journal of Agricultural and Food Chemistry 47, 4500-05. [56] Ajitha, M. J.; Mohanlal, S.; Suresh, C. H.; Jayalekshmy, A. 2012. “DPPH radical scavenging activity of tricin and its conjugates isolated from “Njavara” rice bran: A density functional theory study.” Journal of Agricultural and Food Chemistry 60, 3693-99. [57] Moscatelli, V.; Hnatyszyn, O.; Acevedo, C.; Megias, J.; Alcaraz, M. J.; Ferraro, G. 2006. “Flavonoids from Artemisia copa with anti-inflammatory activity.” Planta Medica 72, 72-74. [58] Lee, S. S.; Baek, Y. S.; Eun, C. S.; Yu, M. H.; Baek, N. I.; Chung, D. K.; Bang, M. H.; Yang, S. A. 2015. “Tricin derivatives as anti-inflammatory and anti-allergic constituents from the aerial part of Zizania latifolia.” Bioscience Biotechnology and Biochemistry 79, 700-06. [59] Shalini, V.; Bhaskar, S.; Kumar, K. S.; Mohanlal, S.; Jayalekshmy, A.; Helen, A. 2012. “Molecular mechanisms of anti-inflammatory action of the flavonoid, tricin from Njavara rice (Oryza sativa L.) in human peripheral blood mononuclear cells: Possible role in the inflammatory signaling.” International Immunopharmacology 14, 32-38. [60] Shalini, V.; Jayalekshmi, A.; Helen, A. 2015. “Mechanism of anti-inflammatory effect of tricin, a flavonoid isolated from Njavara rice bran in LPS induced hPBMCs and carrageenan induced rats.” Molecular Immunology 66, 229-39. [61] Ogo, Y.; Ozawa, K.; Ishimaru, T.; Murayama, T.; Takaiwa, F. 2013. “Transgenic rice seed synthesizing diverse flavonoids at high levels: a new platform for flavonoid production with associated health benefits.” Plant Biotechnology Journal 11, 73446. [62] Norbaek, R.; Aaboer, D. B. F.; Bleeg, I. S.; Christensen, B. T.; Kondo, T.; Brandt, K. 2003. “Flavone C-glycoside, phenolic acid, and nitrogen contents in leaves of barley subject to organic fertilization treatments.” Journal of Agricultural and Food Chemistry 51, 809-13. [63] Owada, E.; Mieno, M. 1970. “Synthetic studies of flavonoid compounds. 2. improved synthesis of tricin.” Nippon Kagaku Zasshi 91, 1002-&. [64] Nagarathnam, D.; Cushman, M. 1991. “A short and facile synthetic route to hydroxylated flavones - new syntheses of apigenin, tricin, and luteolin.” Journal of Organic Chemistry 56, 4884-87.

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

75

[65] Pandurangan, N. 2014. “A new synthesis for acacetin, chrysoeriol, diosmetin, tricin and other hydroxylated flavones by modified Baker-Venkataraman transformation.” Letters in Organic Chemistry 11, 225-29. [66] Wang, J.; Zhou, R. G.; Wu, T.; Yang, T.; Qin, Q. X.; Li, I.; Yang, B.; Yang, J. 2012. “Total synthesis of apigenin.” Journal of Chemical Research, 121-22. [67] Lan, W.; Lu, F. C.; Regner, M.; Zhu, Y. M.; Rencoret, J.; Ralph, S. A.; Zakai, U. I.; Morreel, K.; Boerjan, W.; Ralph, J. 2015. “Tricin, a flavonoid monomer in monocot lignification.” Plant Physiology 167, 1284-95. [68] Mao, J. Z.; Zhang, X.; Li, M.-F.; Xu, F. 2013. “Effect of biological pretreatment with white-rot fungus trametes hirsuta C7784 on lignin structure in Carex meyeriana Kunth.” Bioresources 8, 3869-83. [69] Wen, J.; Sun, S.; Xue, B.; Sun, R. 2013. “Quantitative structural characterization of the lignins from the stem and pith of bamboo (Phyllostachys pubescens).” Holzforschung 67, 613-27. [70] Rencoret, J.; Ralph, J.; Marques, G.; Gutierrez, A.; Martinez, A. T.; del Río, J. C. 2013. “Structural characterization of lignin isolated from coconut (Cocos nucifera) coir fibers.” Journal of Agricultural and Food Chemistry 61, 2434-45. [71] You, T. T.; Mao, J. Z.; Yuan, T. Q.; Wen, J. L.; Xu, F. 2013. “Structural elucidation of the lignins from stems and foliage of Arundo donax Linn.” Journal of Agricultural and Food Chemistry 61, 5361-70. [72] Wu, M.; Pang, J.; Lu, F.; Zhang, X.; Che, L.; Xu, F.; Sun, R. 2013. “Application of new expansion pretreatment method on agricultural waste. Part I: Influence of pretreatment on the properties of lignin.” Industrial Crops and Products 50, 887-95. [73] Rencoret, J.; Prinsen, P.; Gutierrez, A.; Martinez, A. T.; del Río, J. C. 2015. “Isolation and structural characterization of the milled wood lignin, dioxane lignin, and cellulolytic lignin preparations from Brewer’s spent grain.” Journal of Agricultural and Food Chemistry 63, 603-13. [74] del Río, J. C.; Lino, A. G.; Colodette, J. L.; Lima, C. F.; Gutierrez, A.; Martinez, A. T.; Lu, F. C.; Ralph, J.; Rencoret, J. 2015. “Differences in the chemical structure of the lignins from sugarcane bagasse and straw.” Biomass & Bioenergy 81, 322-38. [75] Jensen, A.; Cabrera, Y.; Hsieh, C. W.; Nielsen, J.; Ralph, J.; Felby, C. 2017. “2D NMR characterization of wheat straw residual lignin after dilute acid pretreatment with different severities.” Holzforschung 71, 461-69. [76] Heikkinen, H.; Elder, T.; Maaheimo, H.; Rovio, S.; Rahikainen, J.; Kruus, K.; Tamminen, T. 2014. “Impact of steam explosion on the wheat straw lignin structure studied by solution-state nuclear magnetic resonance and density functional methods.” Journal of Agricultural and Food Chemistry 62, 10437-44.

76

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[77] Zikeli, F.; Ters, T.; Fackler, K.; Srebotnik, E.; Li, J. B. 2016. “Wheat straw lignin fractionation and characterization as lignin-carbohydrate complexes.” Industrial Crops and Products 85, 309-17. [78] Luterbacher, J. S.; Rand, J. M.; Alonso, D. M.; Han, J.; Youngquist, J. T.; Maravelias, C. T.; Pfleger, B. F.; Dumesic, J. A. 2014. “Nonenzymatic sugar production from biomass using biomass-derived γ-valerolactone.” Science 343, 27780. [79] Luterbacher, J. S.; Azarpira, A.; Motagamwala, A. H.; Lu, F.; Ralph, J.; Dumesic, J. A. 2015. “Aromatic monomer production integrated into the γ-valerolactone sugar platform.” Energy and Environmental Science 8, 2657-63. [80] Sousa, L. D.; Foston, M.; Bokade, V.; Azarpira, A.; Lu, F. C.; Ragauskas, A. J.; Ralph, J.; Dale, B.; Balan, V. 2016. “Isolation and characterization of new lignin streams derived from extractive-ammonia (EA) pretreatment.” Green Chemistry 18, 4205-15. [81] Zikeli, F.; Ters, T.; Fackler, K.; Srebotnik, E.; Li, J. B. 2016. “Fractionation of wheat straw Dioxane lignin reveals molar mass dependent structural differences.” Industrial Crops and Products 91, 186-93. [82] Lu, F. C.; Ralph, J. 2011. “Solution-state NMR of lignocellulosic biomass.” Journal of Biobased Materials and Bioenergy 5, 169-80. [83] Ralph, J.; Landucci, L. L. In Lignin and Lignans; Advances in Chemistry; Heitner, C.; Dimmel, D. R.; Schmidt, J. A., Eds.; CRC Press (Taylor & Francis Group): Boca Raton, FL, 2010, doi: org/10.1201/EBK1574444865 org/10.1201/EBK1574444865. [84] Kwon, Y. S.; Kim, C. M. 2003. “Antioxidant constituents from the stem of Sorghum bicolor.” Archives of Pharmaceutical Research 26, 535-39. [85] Li, M.; Pu, Y. Q.; Tschaplinski, T. J.; Ragauskas, A. J. 2017. “31P NMR Characterization of tricin and its structurally similar flavonoids.” Chemistry Select 2, 3557-61. [86] Mansfield, S. D.; Kim, H.; Lu, F. C.; Ralph, J. 2012. “Whole plant cell wall characterization using solution-state 2D NMR.” Nature Protocols 7, 1579-89. [87] Stokvis, E.; Rosing, H.; Beijnen, J. H. 2005. “Stable isotopically labeled internal standards in quantitative bioanalysis using liquid chromatography/mass spectrometry: necessity or not?” Rapid Communications in Mass Spectrometry 19, 401-07. [88] Schafer, J.; Urbat, F.; Rund, K.; Bunzel, M. 2015. “A stable-isotope dilution GC-MS approach for the analysis of DFRC (Derivatization Followed by Reductive Cleavage) monomers from low-lignin plant materials.” Journal of Agricultural and Food Chemistry 63, 2668-73. [89] Eloy, N. B.; Voorend, W.; Lan, W.; Saleme, M. D. S.; Cesarino, I.; Vanholme, R.; Smith, R. A.; Goeminne, G.; Pallidis, A.; Morreel, K. et al. 2017. “Silencing

Tricin in Grass Lignin: Biosynthesis, Characterization, and Quantitation

77

CHALCONE SYNTHASE in maize impedes the incorporation of tricin into lignin and increases lignin content.” Plant Physiology 173, 998-1016. [90] Lam, P. Y.; Zhu, F. Y.; Chan, W. L.; Liu, H. J.; Lo, C. 2014. “Cytochrome P450 93G1 is a flavone synthase II that channels flavanones to the biosynthesis of tricin O-linked conjugates in rice.” Plant Physiology 165, 1315-27. [91] Park, S.; Choi, M. J.; Lee, J. Y.; Kim, J. K.; Ha, S. H.; Lim, S. H. 2016. “Molecular and biochemical analysis of two rice flavonoid 3'-hydroxylase to evaluate their roles in flavonoid biosynthesis in rice grain.” International Journal of Molecular Sciences 17. [92] Shih, C. H.; Chu, H.; Tang, L. K.; Sakamoto, W.; Maekawa, M.; Chu, I. K.; Wang, M.; Lo, C. 2008. “Functional characterization of key structural genes in rice flavonoid biosynthesis.” Planta 228, 1043-54. [93] Lam, P. Y.; Tobimatsu, Y.; Lui, A. C.; Yamamura, M.; Wang, L.; Takeda, Y.; Suzuki, S.; Liu, H.; Zhu, F.; Chen, M. et al. 2018. “Recruitment of specific flavonoid B-ring hydroxylases for two independent biosynthesis pathways of flavone-derived metabolites in grasses.” New Phytologist, under revision. [94] Do, C. T.; Pollet, B.; Thevenin, J.; Sibout, R.; Denoue, D.; Barriere, Y.; Lapierre, C.; Jouanin, L. 2007. “Both caffeoyl Coenzyme A 3-O-methyltransferase 1 and caffeic acid O-methyltransferase 1 are involved in redundant functions for lignin, flavonoids and sinapoyl malate biosynthesis in Arabidopsis.” Planta 226, 1117-29. [95] Marita, J. M.; Ralph, J.; Hatfield, R. D.; Guo, D. J.; Chen, F.; Dixon, R. A. 2003. “Structural and compositional modifications in lignin of transgenic alfalfa downregulated in caffeic acid 3-O-methyltransferase and caffeoyl coenzyme A 3-Omethyltransferase.” Phytochemistry 62, 53-65. [96] Marita, J. M.; Vermerris, W.; Ralph, J.; Hatfield, R. D. 2003. “Variations in the cell wall composition of maize brown midrib mutants.” Journal of Agricultural and Food Chemistry 51, 1313-21. [97] Palmer, N. A.; Sattler, S. E.; Saathoff, A. J.; Funnell, D.; Pedersen, J. F.; Sarath, G. 2008. “Genetic background impacts soluble and cell wall-bound aromatics in brown midrib mutants of sorghum.” Planta 229, 115-27. [98] Fornalé, S.; Rencoret, J.; García-Calvo, L.; Encina, A.; Rigau, J.; Gutiérrez, A.; del Río, J. C.; Caparros-Ruiz, D. 2017. “Changes in cell wall polymers and degradability in maize mutants lacking 3' -and 5'-O-methyltransferases involved in lignin biosynthesis.” Plant and Cell Physiology 58, 240-55. [99] Ralph, J.; lundquist, K.; Brunow, G.; Lu, F.; Kim, H.; Schatz, P. F.; Marita, J. M.; Hatfield, R. D.; Ralph, S. A.; Christensen, J. H. 2004. “Lignins: natural polymers from oxidative coupling of 4-hydroxyphenylpropanoids.” Phytochemistry Reviews 3, 29-60. [100] Vanholme, R.; Demedts, B.; Morreel, K.; Ralph, J.; Boerjan, W. 2010. “Lignin biosynthesis and structure.” Plant Physiology 153, 895-905.

78

Wu Lan, Jorge Rencoret, José Carlos del Río et al.

[101] Davin, L. B.; Lewis, N. G. 2005. “Lignin primary structures and dirigent sites.” Current Opinion in Biotechnology 16, 407-15. [102] Ralph, J.; Brunow, G.; Harris, P. J.; Dixon, R. A.; Schatz, P. F.; Boerjan, W. In Recent Advances in Polyphenol Research; Daayf, F.; El Hadrami, A.; Adam, L.; Ballance, G. M., Eds.; Wiley-Blackwell Publishing: Oxford, UK, 2008; Vol. 1. [103] Ralph, J.; Grabber, J. H.; Hatfield, R. D. 1995. “Lignin-ferulate cross-links in grasses: Active incorporation of ferulate polysaccharide esters into ryegrass lignins.” Carbohydrate Research 275, 167-78.

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 4

SOLUTION-STATE MULTIDIMENSIONAL NMR OF LIGNINS: APPROACHES AND APPLICATIONS Yuki Tobimatsu1,*, Toshiyuki Takano2, Toshiaki Umezawa1 and John Ralph3 1

Research Institute for Sustainable Humanosphere, Kyoto University, Japan 2 Graduate School of Agriculture, Kyoto University, Japan 3 Department of Biochemistry, and the Department of Energy’s Great Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin-Madison, US

ABSTRACT Among numerous analytical techniques developed for lignins, solution-state multidimensional NMR provides unparalleled details of the polymers’ structural features to be elucidated. With an ability to diagnostically identify and approximate the diverse array of the structural elements in lignin polymers, the NMR techniques are becoming essential in numerous lignin research settings including those aiming to investigate the biosynthesis, bioengineering and biodegradation lignins, as well as the chemistry of lignins in various chemical and biochemical contexts in the light of the biorefinery concept. This chapter review will consider some basic and practical aspects of such NMR implementations in current lignin research. Specifically, a handful of key 2D and 3D NMR experiments for characterizing lignins and basic strategies for preparing suitable cell wall lignin samples to apply these NMR techniques are discussed.

Keywords: biomass, HMBC, HSQC, plant cell walls, quantitative HSQC, TOCSY

*

Corresponding Author Email: [email protected]

80

Yuki Tobimatsu, Toshiyuki Takano, Toshiaki Umezawa et al.

INTRODUCTION Lignin displays considerable structural heterogeneity. The chemical structure of lignin is complex and intricate within a single lignin polymer sample and also highly diverse and variable among practically any different lignin samples from different biomass origins, i.e., the substructure and bonding patterns of lignin polymers exhibit wide variability among plant lineages as well as tissue/cell types and are also influenced by developmental and environmental factors in plant growth. More than 10 different types of aromatic building units derived from different canonical lignin monomers beyond the three classical monolignols, i.e., coniferyl, sinapyl, and p-coumaryl alcohols, and more than 20 different types of side-chain linkages connecting these aromatic units or serving as the polymer endunits have been described, and it is highly probable that more exist in low proportions or in previously overlooked locations. In addition, genetic manipulation of the lignin biosynthetic pathway has generated “abnormal” lignins incorporating non-canonical aromatic units occasionally connected by novel inter-monomeric linkage types, thereby further expanding the array of lignin chemistry to be considered by researchers [1-6]. The structure of lignin has long been studied by chemical degradation of the polymer into smaller fragments that can be then identified and quantified to deduce the existence and abundance of the small elements in the original lignin polymer [7]. Indeed, much of our current understanding of lignin structure was derived from the pioneering degradative methods such as nitrobenzene oxidation, acidolysis, permanganate oxidation, thioacetolysis, and hydrogenolysis [8-12]. Newer chemical degradative methods, such as thioacidolysis [13, 14], derivatization followed by reductive cleavage (DFRC) [15, 16], ozonation [17, 18], tosylation–iodination–zinc-metal (TIZ) treatment [19], and γtosylation–thioetherification–sulfonylation–mild alkali hydrolysis (γ-TTSA) treatment [20], in combination with modern gas/liquid chromatographic and mass spectrometric measurements, remain indispensable for extensive analysis targeting specific features and interunit lignin linkage types. On another front, the advent of sophisticated nuclear magnetic resonance (NMR) techniques has greatly improved our capacity to study the complex lignin structure. In particular, multi-dimensional heteronuclear NMR, in which the chemical shift space available for resolution in conventional 1D NMR spectra is greatly expanded by addition of another spectral dimension, offers powerful tools to elucidate the complex array of the lignin subunit and linkage types. In fact, such NMR techniques were crucial in identifications of the dibenzodioxocin [21, 22] and spirodienone [23, 24] substructures, both of which are now well recognized as being essentially common linkage types in lignins of constitutive xylem tissues in vascular plants. In addition, NMR has been responsible for the recent discoveries of new types of lignin polymers biosynthesized

Solution-State Multidimensional NMR of Lignins: Approaches and Applications

81

from previously overlooked lignin monomers, such as caffeyl alcohol [25, 26], 5hydroxyconiferyl alcohol [27], tricin [28-30], hydroxystilbenes [31, 32], and diferuloylputrescine [33] in some specific plant lineages and tissues. Furthermore, the whole-cell-wall dissolution/solubilization methods now allow researchers to acquire 2D and 3D spectra that display well-resolved lignin resonances even without laborious isolation of lignins from biomass [34-43]. This review selects some basic and practical aspects of the current use of 2D and 3D NMR methods in lignin structural characterization. Specifically, we initially provide an overview of a handful of 2D and 3D NMR experiments most commonly used for recent lignin research, and subsequently consider strategies for preparing lignin samples to effectively implement these NMR techniques in various research settings, with highlighting some recent research reports – note that the literature highlighted here are just examples amongst many. Interested readers should refer to the book chapter by Ralph and Landucci [44] for expansive and comprehensive descriptions of the methodology and utility of NMR spectroscopy in lignin characterization and may also refer to other recent review articles addressing relevant subjects [45-48].

2D HSQC (HETERONUCLEAR SINGLE-QUANTUM COHERENCE) The 2D 1H–13C HSQC is the most popular NMR experiment currently used for many lignin research purposes. This NMR experiment correlates 13C and 1H nuclei by the evolution and transfer of single-quantum coherence via the one-bond C–H coupling (1JCH), and yields a 2D spectrum that has 13C chemical shifts on the “indirect” F1 dimension and 1 H chemical shifts on the “direct” F2 dimension with sharp cross-peaks representing the one-bond C–H relationships. An obvious advantage of HSQC as compared to 1D 1H and 13 C experiments is that it offers far better signal dispersion; overlapping protons may be isolated by their attachments to carbons with different chemical shifts, and likewise overlapping carbons may be distinguished by their proton shift differences. In addition, as it is an “inverse” proton-detected experiment, HSQC is much more sensitive and thus the acquisition time is shorter when compared with that of the 1D 13C experiments. Therefore, with recording an HSQC spectrum, time-consuming 1D 13C experiments can be often omitted in routine lignin characterization settings, unless the identification of quaternary carbons, which are not detected by HSQC, is crucial. An important acquisition parameter in the HSQC pulse sequence is the insensitive nuclei enhanced by polarization transfer (INEPT) delay which depends on the magnitude of 1JCH and consequently affects the relative intensities of correlation signals from various 1 H–13C pairs with different 1JCH values. For typical lignin samples, this delay time is set to maximize coherence transfer for an expected average value of 1JCH for major CHn

82

Yuki Tobimatsu, Toshiyuki Takano, Toshiaki Umezawa et al.

groups in lignins, which is typically 140-150 Hz. The phase-sensitive or multiplicity-edited pulse variants can be optionally chosen to make a clear distinction between carbons bearing an even (CH2) or odd number (CH or CH3) of directly bonded protons. The adiabatic pulse variants of HSQC experiments have been recommended as they seem to give less artifacts and more uniform correlation profiles relatively independently of 1JCH variations [39, 40].

Peak Assignments Extensive efforts by pioneering lignin researchers have established the 1H and 13C chemical shift assignments for many structural motifs found in lignins and associated polysaccharides in cell walls; these assignments were made based on the collection of rigorous NMR data of isolated lignins, synthetic lignins (dehydrogenation polymers, DHPs), and diverse monomeric and oligomeric lignin model compounds, as well as various carbohydrate standards. A useful NMR database listing 1H and 13C chemical shift assignments for over 450 lignin and carbohydrate model compounds in common NMR solvents such as acetone-d6, chloroform-d, and dimethylsulfoxide (DMSO)-d6 is available online [49]. Tables and datasets listing major chemical assignments for cell wall NMR analyses using DMSO-d6/pyridine-d5 [29, 50-54], perdeuterated ionic liquids (ILs)/DMSOd6 [38, 41, 42], and hexamethylphosphoramide (HMPA)-d18/DMSO-d6 [43] solvent systems can be also found. Such public resonance assignment data may allow researchers to annotate many of the previously established lignin and polysaccharide correlations. Assigning new or previously overlooked lignin resonances, however, is a non-trivial task that often requires synthesis of new model compounds for spectral comparison within HSQC sepctra and also, ideally, additional acquisition of other short-range and long-range correlation spectra to unambiguously authenticate the new signal assignments made.

Comparative Peak Quantification Semi-quantitative (comparative) analysis of integral ratios in HSQC spectra is possible especially between known 1H–13C pairs in similar chemical environments with the reasonable assumption that their 1JCH values fall within a similar, narrow range [39]. For example, C2–H2 correlations from guaiacyl (G) and C2–H2/C6–H6 correlations from symmetrical syringyl (S) and p-hydroxyphenyl (H) units have been commonly used to obtain estimates of the important H/G/S lignin compositional ratios. Likewise, Cα–Hα correlations from the phenylpropanoid side-chains are particularly useful for estimating the distribution of these inter-monomeric linkage types in lignins in various lignin samples.

Solution-State Multidimensional NMR of Lignins: Approaches and Applications

83

That said, it must be always kept in mind that HSQC is not absolutely quantitative, i.e., the simple proportionality of the volume integrations to the molar quantity of individual CHn groups in lignins can be perturbed considerably by many factors inherent in the common pulse program, including the mismatch of 1JCH-dependent delay parameters as mentioned above and several other factors, which are further considered in the section titled “HSQC Variants for Enhanced Quantitative Capability” below.

Typical HSQC Profiles of Natural Lignins Figures 1 and 2 show HSQC spectra of representative cell walls and isolated lignin samples, respectively, from a softwood (Pinus radiata), a hardwood/eudicot (Eucalyptus globulus), and a monocot grass (Oryza sativa; rice). The spectra of unfractionated cell walls display well-resolved lignin resonances from some of the major lignin substructures amongst overwhelming polysaccharide resonances, whereas those, along with other relatively minor lignin resonances, can be seen much more clearly in the spectra of the corresponding isolated lignin samples (further discussed in the section titled “Sample Preparation Strategies for 2D and 3D NMR of Lignins” below). The aromatic sub-regions resolve signals from typical guaiacyl (G), syringyl (S), and p-hydroxyphenyl (H) rings and volume integrals of the resolved C2–H2/C6–H6 contour signals (G2, S2/6, and H2/6) allow reasonable quantification of the H/G/S lignin unit composition (Figures 1A and 2A). The pine lignin follows the pattern of typical gymnosperm (softwood) lignins, composed of almost only G units, whereas the other angiosperm lignins from the hardwood eucalyptus and a grass rice are typical mixtures of G and S units. Regardless of the plant sources, H units are typically minor but may be slightly more prevalent, albeit still at low levels, in grass lignins as compared to in softwood and eudicot (including hardwood) lignins. As is typical for grass cell walls, the HSQC spectrum of rice cell walls additionally displays intense signals from p-coumarate (P) [55, 56] and flavone tricin (T) [28, 29] residues attached mainly on lignins, as well as signals from ferulates (F) mainly on hemicelluloses (Figure 1A) [55, 56]; the ferulate signals are therefore reasonably depleted in the spectra of the isolated rice lignin sample in which most of the polysaccharide components have been removed (Figure 2A). In softwood lignin spectra, the recently determined G2 correlations specifically from the 4–O–5 intermonomeric linkage type can be seen at δC/δH 105.0-106.5/6.8 at very low contour levels, whereas the identification of this particular linkage type is difficult in eudicot and grass lignin spectra as the correlations overlap with the main S2 correlations from other major linkage types (Figure 2A) [57, 58].

84

Yuki Tobimatsu, Toshiyuki Takano, Toshiaki Umezawa et al.

Figure 1. Partial 2D 1H–13C HSQC spectra of whole-cell-wall samples from a softwood (Pinus radiata), a hardwood (Eucalyptus globulus), and a monocot grass (Oryza sativa; rice). Aromatic (A), aliphatic (B), and polysaccharide anomeric (C) sub-regions are separately displayed. Contour coloring matches that of the lignin substructure units shown in Figure 2. Boxes labeled  2 and  4 indicate regions that are vertically scaled 2-fold and 4-fold, respectively.

Solution-State Multidimensional NMR of Lignins: Approaches and Applications

85

Figure 2. Partial 2D 1H–13C HSQC spectra of lignin-enriched cell wall or isolated lignin (milled-wood lignin, MWL) samples from a softwood (Pinus radiata), a hardwood (Eucalyptus globulus), and a monocot grass (Oryza sativa; rice). Aromatic (A) and aliphatic (B) are separately displayed. Volume integrals are given for the major lignin aromatic and side-chain structures that are color-coded to match their assignments in the spectrum. Boxes labeled 2, 4, and 8 indicate regions that are vertically scaled 2-fold, 4-fold, and 8-fold, respectively. n.d., not detected.

86

Yuki Tobimatsu, Toshiyuki Takano, Toshiaki Umezawa et al.

Figure 3. NMR observation of natural catechyl lignins. (A and B) Partial 2D 1H–13C short-range HSQC sepctra of acetylated whole seedcoat cell walls of Jatropha curcas (A) and Vernicia fordii (B). Volume integrals are given for the major lignin aromatic and side-chain structures that are color-coded to match their assignments in the spectrum. (C and D) Partial 2D 1H–13C long-range HMBC spectra of acetylated samples of isolated Euphorbiaceae and Cleomaceae seedcoat lignins and synthetic lignin polymer (GDHP) prepared from polymerization of caffeyl alcohol with coniferyl and sinapyl alcohol (G/S/C-DHP) and with coniferyl alcohol only (G/C-DHP). Subregions exhibiting aromatic correlations to benzodioxane α-protons (C) and correlations to β-aryl ether side-chain α-protons (D) are shown. Boxes with 2 or 4 indicate regions that were vertically scaled 2- or 4-fold. Adopted from Tobimatsu, Y., Chen, F., Nakashima, J., Jackson, L., Escamilla-Treviño, L. L., Dixon, R. A., and Ralph, J. 2013. Plant Cell 25:2587-2600 (www.plantcell.org; copyright American Society for Plant Biologists).

The aliphatic regions of the HSQC spectra resolve the correlations for the various lignin inter-monomeric linkage types and those for the polymer end-units and methoxyl

Solution-State Multidimensional NMR of Lignins: Approaches and Applications

87

groups (Figures 1B and 2B). As already mentioned, volume integrals of the well-resolved Cα–Hα correlations in the spectra of isolated lignin samples can be used to compare the distribution of lignin inter-monomeric linkage types between different lignin polymer samples (Figure 2B). The predominant signals are usually from the common α-free β–O– 4 units (β-aryl ethers, I), with lesser amounts of β–5 (phenylcoumarans, II), β–β (resinols, III), 5–5/4–O–β (dibenzodioxocins, IV), and β–1 (spirodienones, V) signals, and also signals arising from trace amounts of α-keto-β–O–4 (I″) and α–O–4/β–O–4 (diaryl ethers, I′′′) signals may be seen. The relative proportions of β–O–4 and β–1 signals are usually higher and those of β–5 and 5–5/4–O–β signals are conversely lower in hardwood (eudicot) and grass lignins when compared with those in softwood lignins. Such shifts of lignin linkage distributions are typical when lignin incorporates more S units than G or H units as has been demonstrated by NMR studies on transgenic plants with variously altered H/G/S unit composition [59-64]. The rice lignin spectrum additionally displays characteristic signals from acylated lignin units, notably tetrahydrofuran-type β–β units (III') [65, 66], affirming that this lignin, as is typical in grass lignins, is highly acylated at the γ-OH positions mainly by p-coumarates P (Figures 1B and 2B) [55, 67, 68]. In some taxa of vanilla orchid and cactaceae seedcoats, lignins are apparently biosynthesized solely from either of catechyl (C) or 5-hydroxyguaiacyl (5H)-type monolignols, i.e., caffeyl or 5-hydroxyconiferyl alcohols, producing C or 5H lignin homopolymers [25, 27], whereas lignins in many of the Euphorbiaceae and Cleomaceae seedcoats have been found to be mixtures of C lignins with conventional G or G/S lignins, although the two classes don’t appear to be co-synthesized/copolymerized [26]. HSQC spectra of these natural C and 5H lignins display characteristic aliphatic signals from trans/cis-benzodioxane units with linkages derived from β–O–4-type radical coupling that typifies lignification but following rearomatization of quinone methide intermediates via an atypical intramolecular trapping mechanism (Figure 3) [25, 69]. Likewise, as recently discovered, HSQC spectra of lignins from palm fruit endocarps resolve unique correlations from the cassigarol E- and aiphanol-type benzodioxane units, along with scirpusin B-type phenylcoumaran and kompasinol A-type resinol units, all of which have been derived from radical coupling of newly identified hydroxystilbene lignin monomers producing unique lignin polymers with traditional monolignols [31, 32].

Non-Canonical Lignin Substructures in Transgenic/Mutant Plants Extensive studies on the bioengineering of lignin have revealed the substantial plasticity of lignification in planta [1, 2, 4, 61, 70-72]. Manipulation of the cinnamate/ monolignol pathway has led to the incorporation of non-canonical lignin monomers to eventually generate new lignin substructures. For example, numerous CCoAOMT-deficient [73] and CAldOMT/COMT-deficient [74-76] plants have been found to produce

88

Yuki Tobimatsu, Toshiyuki Takano, Toshiaki Umezawa et al.

benzodioxane-type β–O–4 units via radical coupling of unusual caffeyl and 5hydroxyconiferyl alcohol monomers, respectively, in major xylem tissues where lignins are usually composed of G/S or G units derived from typical coniferyl alcohol with or without sinapyl alcohol monomers. In these plants, atypical caffeyl and 5-hydroxyconiferyl alcohol monomers are forced to participate in lignification by a truncation of the cinnamate/monolignol pathway due to the loss of the responsible OMT gene function. In a similar way, CCR-deficient plants produced novel bis-(β–O–4) units through incorporation of unusual ferulic acid monomer [77, 78], and numerous CAD-deficient plants produced various γ-aldehyde lignin substructures through incorporation of unusual p-hydroxycinnamaldehyde monomers [59, 79-83]. Such abnormal lignin substructures in various transgenic and mutant plants have been evidenced primarily by 2D and/or 3D NMR analysis of the lignins along with suitable model compounds such as in vitro synthetic lignins (DHPs) incorporating the anticipated new lignin monomers. As an example, Figure 4 shows HSQC spectra of lignins from a recently reported OsFNSII-deficient rice mutant (fnsII) that is defective in one of the key reaction steps, the conversion of naringenin to apigenin, requisite for the synthesis of the tricin lignin monomer in grasses [84]. In the aromatic sub-regions of the rice mutant spectra, the characteristic set of aromatic signals derived from the flavone aromatic system of tricin (T3, T6, T8, and T2´/6´) [28, 29] are essentially absent. Instead, diagnostic signals from the narigenin flavanone ring system (N8/6 and N2) are clearly visible in the spectra of the mutant lignins and exactly match with the signals appearing in the spectra of DHP prepared with naringenin and coniferyl alcohol (GN-DHP), demonstrating that the rice mutant has incorporated the non-canonical naringenin monomer instead of tricin into its lignin polymers [84].

HSQC VARIANTS FOR ENHANCED QUANTITATIVE CAPABILITY The difficulty of performing “true” quantitative analysis is an important limitation of the conventional 2D 1H–13C HSQC NMR although, as mentioned above, it still offers valuable semi-quantitative information regarding the relative abundances of major lignin subunits and linkages that can be used for comparative analyses between samples under identical NMR conditions [see the subsection “Comparative peak quantification” in the section “2D HSQC (Heteronuclear Single-Quantum Coherence)” above]. Typical heteronuclear correlation NMR experiments, including HSQC, are not quantitative mainly because of resonance-specific signal attenuations during the coherence transfer steps. These signal attenuations can be arise from several factors such as variations in 1JCH, evolutions of 1H–1H homocouplings (JHH), carbon resonance offset effects, and also differential 1H/13C T2 relaxation effects. In 1D spectra, the influence of 1H T1 deviations can be easily suppressed by setting the relaxation delay in the pulse sequence properly (five

Solution-State Multidimensional NMR of Lignins: Approaches and Applications

89

times the longest T1 of interest) but the problem in HSQC experiments is that there is significant relaxation, and obviously more by the fastest relaxing components, during the pulse sequence itself, i.e., before that actual acquisition of each FID. This leads to a situation where, for example, the more mobile endgroups (with comparatively slow relaxation) become over-represented relative to the less mobile internal units of the lignin polymer chain [26, 30, 40]. The so-called quantitative HSQC (Q-HSQC) [85] is a variant developed to suppress 1 the JCH-dependence of coherence transfer by modulating the transfer delay within a single pulse program. The quantitative Carr-Purcell-Meiboom-Gill (CPMG)-adjusted HSQC (QCAHSQC) further suppresses resonance offset and JHH evolution effects by replacing the constant-time INEPT steps with the constant-time CPMG-INEPT steps [86]. In addition, as a powerful modification of Q-CAHSQC, Peterson and Loening [87] developed the socalled quick quantitative HSQC (QQ-HSQC) pulse scheme, in which signals corresponding to different INEPT delays are acquired simultaneously from different parts of the sample, resulting in greatly reduced number of scans per increment and therefore shorter total acquisition time to achieve the same spectral quality as Q-HSQC or QCAHSQC does. The utility of these Q-HSQC variants for quantitative analysis of lignin aromatic composition and linkage distribution in isolated lignin samples has been demonstrated by Crestini and co-workers [88-91] but only on quite clean isolated lignin samples that do not have the short relaxation times experienced with, for example, wholecell-wall samples. However, especially when analyzing polymer mixtures like typical lignin samples, peak integrals derived from the above Q-HSQC variants can still be skewed by large deviations in the T2 relaxation rate among different components in the sample mixture, e.g., high vs low molecular weight polymer fractions, and mobile end-groups vs rigid internal units even within a single polymer molecule. A protocol to acquire HSQC-derived quantitative data by using internal standard reference resonances and resonance-specific response factors derived from quantitative 1D 13C spectra has been proposed [92]. In addition, the so-called HSQC0 method, in which a quantitative time-zero HSQC0 spectrum is constructed by extrapolation of peak intensities from a series of constant time gradient HSQCn spectra, may offer a simpler way to cancel any factors contributing to the signal attenuations during the coherence transfer steps [93]. The HSQC0 method has been applied for quantitative analysis of lignin composition and linkage distributions in, for example, isolated spruce lignins [90], wheat organosolv lignins [94], and even unfractionated Miscanthus cell walls [42]. Okamura et al. (2016) present another dedicated quantitative method, the so-called ‘tolerant of any factor’ (TAF) method, which corrects for the loss of magnetization using calibration factors determined by additional transverse relaxation optimized spectroscopy (TROSY) experiments [95].

90

Yuki Tobimatsu, Toshiyuki Takano, Toshiaki Umezawa et al.

Figure 4. Partial 2D 1H–13C HSQC spectra of lignin-enriched cell wall samples from wild-type (WT) and FNSII-knockout mutant (fnsII) rice plants, and in vitro synthetic lignin polymers (DHPs). Aromatic (A-D) and aliphatic (E-H) sub-regions for lignin-enriched cell walls of WT (A and E) and fnsII mutant (B and F) plants and DHPs prepared from coniferyl alcohol only (G-DHP, C and G) and from coniferyl alcohol along with naringenin (GN-DHP, D and H) are separately displayed. Boxes labeled 2 indicate regions that are vertically scaled 2-fold. Contour coloration matches that of the lignin substructure units shown. Adopted from Lam, P. Y., Tobimatsu, Y., Takeda, Y., Suzuki, S., Yamamura, M., Umezawa, T., and Lo, C. 2017. 174:972-985 (www.plantphysiol.org; copyright American Society for Plant Biologists).

Solution-State Multidimensional NMR of Lignins: Approaches and Applications

91

2D HMBC (HETERONUCLEAR MULTIPLE-BOND CORRELATION) HMBC correlates protons with carbons via typically two- or three-bond coupling, providing more information about connectivities and more signal dispersion compared to 1D and 2D HSQC spectra. The coupling evolution time, or long-range coupling delay, must be carefully set to match with a long-range coupling constant (nJCH) of interest; for typical lignin polymer samples, this delay time is traditionally chosen in the range of 60-100 ms, which aims to maximize coherence transfer for nJCH of 5–8 Hz and while minimizing relaxation losses that occasionally become problematic especially when longer delay times (>100 ms corresponding to 10% Pd/C (which provided the same results as 30% Pd/C)>Pd(OH)2/C>Pd-PEPPSI-IPr>Pd(OAc)2. Notably, the authors indicated that Nafion SAC-13 activated the lignin aryl ether sites due to its activity as Brønsted acid, promoting hydrolysis over hydrogenation of aromatic rings, which facilitated the cleavage of the ether bonds to produce phenols. The combination of Brønsted acids and metal-supported catalysts has been reported to improve the extent of lignin reductive depolymerization. For instance, E. J. M. Hensen’s research group has shown that combining Pd/C catalysts with homogeneous HCl and Al(OTf)3 affords lignin monomer yields of 46 and 44%, respectively, during oak wood depolymerization [87]. Similar results were reported by B. Sels’ group when using acidic co-catalysts [88]. The authors showed that during biomass fractionation/ depolymerization in methanol using Pd/C as catalyst it is possible to favor the formation of lignin monomers and high-quality pulp when acidic (H3PO4) conditions are employed, because lignin depolymerization and alcoholysis of hemicellulose are promoted. This resulted in a lignin depolymerization efficiency of 65%. In contrast, the authors found that under alkaline reaction conditions (NaOH) lead to a broader product distribution, while reducing the extent of lignin depolymerization (~27%) and cellulose pulp yield. In addition to Cr, Cu, Ni, Ru, and Pd catalysts recent reports have shown that it is possible to depolymerize technical lignin and lignocellulosic biomass utilizing unsupported sulfided MoS2 catalyst [89]. In this case, the authors reported an increase from 6.65% to 18.47% of lignin monomer when the depolymerization reaction was performed

Challenges and Opportunities of Lignin …

149

in the presence of MoS2. This lignin monomer content was accompanied with a 75% fragmentation of the lignin, while more than 90% of the glucan was retained after reaction. This work provided promising results for improving the graminaceous plant lignin-first process with a more economic and sustainable catalyst formulation. These conservative yields were significantly improved when using MoOx supported on carbon nanotubes [90]. In this case, lignin derived from enzymatic mild acidolysis of lignocellulosic biomass was employed as feed-stream. The authors observed that monomer yields could be increased up to 47% when using this nanostructured catalyst under reductive conditions. In addition, by testing different catalysts it was observed that results were also affected by the catalyst used, being the best hydrogenolysis effect achieved with Ru/C catalyst. Regarding the lignin studied (corn stalk lignin and bamboo lignin), it was observed that the yield of products substantially varied from one to another, even though both belong to herbaceous lignin type. Nevertheless, some differences in the chemical structure of such materials seems to exist, specifically in the richness of the p-hydroxyl phenol units and the complexity of linkages, both factors being higher for corn stalk lignin, which may explain the higher yields of 4-ethylphenolics obtained from it. In addition, the stability of one of the catalysts was tested by reusing the Ru/C used in one catalytic run in a subsequent catalytic test at the same conditions. The catalyst seemed to maintain its activity within acceptable levels, though strict comparison is not possible due to differences in the reaction time which, as previously mentioned, is thought to play an important role in the yields obtained during the reaction. In addition, Song et al. [91] reported a method of nickelcatalyzed depolymerization of lignosulfonate into guaiacols at 200ºC using a wide range of solvents as reaction media and a H2 pressure of 5.0 MPa. When water was the reaction medium, no product was detected, whereas if the reaction was carried out in ethylene glycol (EG) conversions up to 91% were achieved. With regard to the second approach, water has also been investigated as reaction media, due to its environmentally friendly character and relatively simple scalability. In this sense, Zhang and coworkers [69] recently reported a series of experiments for hydrogenolysis of organosolv extracted birch lignin, using unsupported Ni, M and NiM catalysts (M: Ru, Pd and Rh). The reactions were carried out at relatively mild conditions (10 bar H2, 130ºC and aqueous medium), showing that higher yield to monomers was obtained when the bimetallic catalyst was used instead of any of the metals on its own. Nevertheless, performance of these catalysts in terms of yield of organosolv lignin to monomers is significantly lower when it is compared to the results obtained with the same catalysts and same operating conditions for the hydrogenolysis of dimeric model compounds. For instance, Ni85Ru15 catalyst is able to achieve ca. 100% of conversion with 58% yield to monomers when used in hydrogenolysis of 2-phenoxy-1-phenethanol, whereas when this catalyst is used with organosolv lignin, the yield to monomers decreases to 0.8%, though this yield is increased up to 6.8% when the reaction is maintained for 12 h instead of 1h.

150

J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

Notably, in that study the best results for extracted birch lignin were obtained with Ni85Rh15 when reaction was carried out for 1 h (yield to monomers = 2.6%), and with Ni85Ru15 when the reaction was carried out for 12 h (yield to monomers = 6.8%), being the conversion of lignin 50% and 44%, respectively. In this sense, the HDO of two technical lignin, one extracted with ethanol and the other by diluted alkali technique, was studied by Laskar et al. [92] using multifunctional catalyst matrix systems containing Ru, Rh or Pt supported either on Al2O3 or C, and acid zeolites. Both technical lignin selected as raw materials for the reaction presented a majority of uniformly distributed β-O-4’ linkages. Better yields were reported when using 5% Ru catalyst supported either on C or Al 2O3; specifically, the highest conversion of lignin (~ 61%) was achieved with 5% Ru/Al2O3 NH4+ Z-Y catalyst matrix for the ethanol extracted lignin, with a 74% of selectivity to aromatic compounds. Jie-wang et al.[93] investigated the hydrogenation of alkali lignin over Pd/C (5% Pd loading), CuO and Raney Ni catalysts. The reactions were carried out in basic aqueous media and, when compared at 4 MPa H2 and 390 K, CuO seemed to present poor catalytic activity. Raney Ni catalyzed the reaction to some extent, but the best results for reductive depolymerization were obtained with the Pd/C catalyst. GPC analysis showed that average molecular weight (MW) and number average molecular weight (Mn) decreased after the reaction, also with an increase of polydispersity index. Aromaticity was conserved to some extent, as analyses of the samples after reaction indicated the presence of some compounds with phenolic hydroxyl units. To summarize, the stability and accessibility of the catalysts and how to retain aromaticity are key factors to consider when studying the depolymerization of lignin. In general, most publications in this field lack of in-depth studies of these issues. Also, the heterogeneity and complexity of feedstocks causes a wide range of products obtained after depolymerization, generally with varying distributions of products depending on the lignin source. The latter, together with the multiple and different procedure for the recovery of products and their analysis complicate the systematic comparison of results. An overview of the most common heterogeneous catalyst systems used in HDO reaction, which presented best results preserving the benzene ring, is summarized in Table 1.

FUTURE OUTLOOK The simultaneous extraction and depolymerization of lignin from lignocellulosic biomass using reductive catalytic strategies has attracted significant research interest because it enables the selective deconstruction of complex biomass structure producing high-quality pulps and lignin streams with control molecular structure [94]. In these socalled “lignin-first” processes the lignin macromolecules are fragmented into highlyreactive aromatic dimers and monomers that in the presence of a reducing atmosphere and metal-catalysts can be readily passivated to form the corresponding saturated products. The

Challenges and Opportunities of Lignin …

151

challenge, however, is how to perform these reactions in large-scale continuous processes considering that initially biomass (solid) cannot directly interact with the catalyst. In a recent report, Bert Sels’ group reported that even when the catalyst is physically separated from the biomass it is possible to obtain lignin-monomers with well-defined structure and chemistry in high yields to monomers (~ 40%) [53, 95–97]. Similar results have been reported by Y. Roman-Leshkov’s group using flow-through reactors (Figure 7) [98, 99]. In this work, the first reactor containing the lignocellulosic biomass was exposed to a specific solvent under solvolysis conditions to perform the fragmentation and extraction of lignin. The effluent of this reactor was sent to a second reactor containing a reduction catalyst to perform the stabilization and hydrogenolysis of the lignin fragments. In another report, I. Kumaniaev et al. showed that it is possible to perform sequential solvolysis and hydrogenolysis using a flow-through system in which pulping and hydrogenolysis steps were separated in time and space [100]. In this case, it was possible to achieve a depolymerization strategy by which it was possible to reach 37% yield of lignin monomers. Notably, in these processes it was possible to generate a high-quality pulp that can be enzymatically hydrolyzed to glucose in yields above 80% without prior purification. These strategies relied on the solvolysis reaction of lignin macromolecules at high temperatures (>250C), which allows nearly complete depolymerization of the C-O bonds in the lignin. Followed by catalytic reduction in the presence of metal catalysts at relatively mild conditions (100-250C) to stabilize and further upgrade the lignin monomers to added-value products. Nevertheless, the high-reactivity of the lignin-fragments leads to losses via re-condensation reactions. To overcome this limitation the group led by J. S. Luterbacher developed a strategy in which formaldehyde is employed to hinder the C-C bond formation reactions between lignin fragments by forming; 1) stable six-member 1,3dioxane (acetal) with the 1,3 diols on lignin side-chains preventing the formation of benzylic cations, and 2) blocking the electron-rich ortho- and para- positions of the methoxyl groups on the aromatic rings. This stabilization strategy enables nearly theoretical yields of lignin monomers after solvolysis and hydrogenolysis under reductive conditions (47 mole% of Klason lignin for beech and 78 mole% for a high-syringyl transgenic poplar). Similar results were reported by X. Ouyang et al. using a two-step approach. In the first step, oak sawdust was delignified in acidic methanol (e.g., sulfuric acid, aluminium triflate, p-methylbenzenesulfonic acid, hydrochloride acid and phosphoric acid). The lignin-oil generated in this process was sent to a second reactor in which the phenolic-monomers were upgraded using Pd/C catalyst under reducing conditions yielding up to 25% to 4-n-propyl syringol/guaiacol. These results clearly indicate that it is possible to convert lignin-containing streams to monomer products in quantitative yields at industrially relevant conditions. However, in order to bring these paradigm-shifting strategies to large-scale it is necessary to address a number of important issues, including; 1) limited extent of C-C cleavage that limits the complete fragmentation of lignin into monomeric phenolics, 2) costly separation of the

152

J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

reaction solvent employed in the solvolysis step, 3) poor long-term catalyst stability due to the harsh reaction conditions that lead to carbon-deposition, metal-leaching, and porecollapse. In the next section we elaborate more on these issues and propose possible alternatives to overcome them.

Figure 7. Flow-through process for simultaneous lignocellulosic biomass fractionation and depolymerization using a tandem approach of solvent-extraction and reductive catalytic stabilization of lignin fragments.[98, 99, 101].

Activating C-C Bonds The majority of the research work performed on lignin depolymerization have been focused on the selective cleavage of C-O ether bonds, which can account up to ~50% of all the ether linkages in the lignin depending on the type of biomass [102]. As a result a large fraction of the lignin remains unaltered after solvation and hydrogenolysis, which limits greatly the total yield of monomeric products. To perform the C-C bond scission one could use acid catalysts that can activate protolytic cracking of these bonds, but the limitation of this approach is that high-temperatures are required, and low selectivity could be obtained as other recombination reactions can be catalyzed (e.g., condensation and oligomerization). Alternatively, hydrogenolysis of the C-C bond could be a more selective strategy. In this case, it will be necessary to develop catalysts with moderate interaction with C-C bonds and H-H in order to activate selectively the hydrogenolysis reaction, while preventing the full saturation of the aromatic ring.

Challenges and Opportunities of Lignin …

153

Separating Monomeric Products To facilitate the selective fragmentation/solvolysis of the lignin-containing streams into lignin monomers and dimers it is necessary to use organic solvents (e.g., methanol, ethanol, isopropanol, and butanol). While in many cases these C1-C4 alcohols can be produced from biomass via fermentative routes (e.g., ABE and bioethanol processes) the cost of solvent make-up, separation, and recirculation will greatly hinder the economic feasibility of these processes. Furthermore, the use of low mass ratios of lignin-containing feedstocks to solvent will lead to larger reactor volumes, recycling streams, and complex distillation trains. The importance of determining the “distillation resistance” has been recently highlighted by J. P. Lange [103]. In this report the analysis of 15 different industrial distillation sections indicated that in order to make a biomass conversion process economically feasible it is necessary that the “distillation resistance” and costs do not exceed 15–17C-1 and $100 per ton, respectively. Broadly speaking the “distillation resistance” corresponds to an energy requirement of approximately 10 GJ per ton of product, which is considered as the ceiling of the energy cost. As a result, any process with higher “distillation resistance” will be economically unfeasible. Furthermore, high energy requirements could lead to excessive CO2 footprint, hindering its qualification as advanced bio-fuel and/or bio-chemical. Thus, it is necessary to develop alternatives that can reduce the use of organic solvents, while maintaining the high yields to lignin monomers. Furthermore, stringent environmental regulations towards the usage of organic solvents will introduce new challenges for chemical companies in the upcoming years, which might also limit the use of extraction solvents in the lignin-first processes. Catalysts Stability in Hot-Aqueous Environments In liquid reaction environments new deactivation mechanisms are induced. In hot aqueous phase solid catalysts (e.g., zeolites, metal oxides, and supported metals) undergo fast deactivation due to the high-solvating and acidity of water molecules. At high temperatures chemical leaching, oxidation, dissolution-redeposition, and water-poisoning readily occurs [104–107]. An alternative to overcome these limitations is to functionalize the surface hydroxyl groups (-OH), associated to structural defects in microporous catalysts, with hydrophobic sylane molecules. This strategy greatly improves the hydrothermal stability of the catalyst in hot aqueous media (Figure 8) [107–109]. Preventing deactivation in water is major challenge in heterogeneous catalysis, but significant improvements are possible with the proper control of the support wettability and confinement of the catalyst [110–113]. In another report, it has been shown that surface wettability plays a crucial role also in the stabilization of noble metals clusters supported on reducible metal-oxides during reaction in aqueous media [114]. Control of surface wettability is especially relevant in conversion of biomass and green chemistry reactions, where water presence is unavoidable [38, 115–117]. Another alternative is to employ atomic layer deposition strategies to create highly confined metal clusters that are more

J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

154

resistant towards metal lixiviation and sintering in aqueous environments. The main limitation of this approach is the high cost associated with the synthesis procedure [113, 118]. Thus, the development of innovative catalysts capable of operating in corrosive aqueous media at high temperatures, “materials-gap,” is major hurdle for the readiness of the lignin-chemistry industry.

Before

After b#

a#

200#nm #

20#nm #

c#

HY

20#nm #

d#

HYOTS 20#nm #

20#nm #

Figure 8. Hydrophobized zeolite (HY-OTS) retained its crystallinity after reaction in aqueous environment.

The development of chemical processes that can transform lignin will be critical to newly commissioned second generation biorefineries and existing pulping industries. However, having a profitable and scalable process for lignin purification and refining will not be enough if lignin-derivatives are not competitive and fungible with existing petroleum-based counterparts. Notably, lignin catalytic depolymerization offers the possibility to activate the right bonds in the structure to create fragments with tailored chemistry, molecular size and structure. Industrial exploitation of lignin depolymerization catalysts will be linked to three key areas of knowledge (Figure 9), 1) physical chemistry, 2) process engineering, and 3) catalysis. The first part of the cycle begins with the development of experimental and computational tools that enable the proper characterization and modeling of lignincontaining feedstocks. This stage employs physico-chemical information to create an accurate model of the lignin structure, chemistry, and dynamics under real processing conditions that otherwise would be difficult to obtain using pure experimentation. Knowing the behavior of lignin polymers in a particular reaction environment supports the selection

Challenges and Opportunities of Lignin …

155

of the catalyst formulation and structure. This is particularly important, because the efficient utilization of catalysts is rather difficult when large macromolecules, like lignin, are employed as limiting reactants. Here, the ingenuity of scientists and engineers is critical to develop catalysts with the proper chemistry (basic, acid, and metal functionalities) and accessibility (aspect ratio and porosity) that allow selective cleavage of C-O linkages and intimate contact between lignin and the catalytic active site. Finally, the implementation of the catalyst requires the development of novel processes that can improve the accessibility of the catalyst to the lignin by increasing the solvation of the lignin globular structure.

Figure 9. Technological approach for the development of tailored-made catalytic materials for the depolymerization of lignin.

CONCLUSION A detailed review of the different reductive catalytic strategies for the depolymerization of lignin has been done to identify knowledge and technology gaps. The results reported in the literature showed that depending on the reaction conditions the depolymerization of lignin can take place via solvolysis/extraction followed by hydrogenation of the unsaturated monomers. During lignin depolymerization the selection of the solvent and reaction conditions plays a crucial role in determining the extent of

156

J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

fragmentation, while the second step of stabilization via hydrogenation can be readily accomplished using metal-catalysis sufficiently active towards C-O hydrogenolysis. Furthermore, the passivation of the lignin using short aldehydes (e.g., formaldehyde) has shown great potential as strategy to enhance the extent of lignin-depolymerization by preventing the recombination reactions. The main challenges that should be overcome in the future are primarily related to achieving complete depolymerization of lignin (i.e., activating C-C bond scission), reducing the usage of organic solvents for the solvolysis step, and improving the long-term stability of the metal-supported catalysts in hot-liquid environments. In this chapter we have identified some of the possible alternatives to overcome these limitations, but it is clear that significant research is required to bring these processes to large-scale lignin valorization.

REFERENCES [1]

[2] [3]

[4]

[5]

[6]

[7]

[8]

Tian, S. Q.; Zhao, R. Y.; Chen, Z. C. Review of the pretreatment and bioconversion of lignocellulosic biomass from wheat straw materials. Renew. Sustain. Energy Rev. 2018, 91 (June 2017), 483–489. Chen, H.; Fu, X. Industrial technologies for bioethanol production from lignocellulosic biomass. Renew. Sustain. Energy Rev. 2016, 57, 468–478. Aditiya, H. B.; Mahlia, T. M. I.; Chong, W. T.; Nur, H.; Sebayang, A. H. Second generation bioethanol production: A critical review. Renew. Sustain. Energy Rev. 2016, 66, 631–653. Zhao, Y.; Damgaard, A.; Christensen, T. H. Bioethanol from corn stover – a review and technical assessment of alternative biotechnologies. Prog. Energy Combust. Sci. 2018, 67, 275–291. Ragauskas, A. J.; Beckham, G. T.; Biddy, M. J.; Chandra, R.; Chen, F.; Davis, M. F.; Davison, B. H.; Dixon, R. a; Gilna, P.; Keller, M.; et al. Lignin valorization: improving lignin processing in the biorefinery. Science (80-. ). 2014, 344 (6185), 1246843. Rinaldi, R.; Jastrzebski, R.; Clough, M. T.; Ralph, J.; Kennema, M.; Bruijnincx, P. C. A.; Weckhuysen, B. M. Paving the Way for Lignin Valorisation: Recent Advances in Bioengineering, Biorefining and Catalysis. Angew. Chem., Int. Ed. 2016, pp 8164–8215. Gómez-Monedero, B.; Pilar Ruiz, M.; Bimbela, F.; Faria, J. Selective depolymerization of industrial lignin-containing stillage obtained from cellulosic bioethanol processing. Fuel Process. Technol. 2018, 173, 165–172. Naqvi, M.; Yan, J.; Dahlquist, E. Black liquor gasification integrated in pulp and paper mills: A critical review. Bioresour. Technol. 2010, 101 (21), 8001–8015.

Challenges and Opportunities of Lignin … [9] [10]

[11] [12] [13] [14]

[15] [16] [17] [18] [19] [20]

[21] [22]

[23]

[24] [25]

157

Hyvärinen, E. The downside of European Union emission trading - A view from the pulp and paper industry. Unasylva 2005, 56 (222), 39–41. Bahl, K.; Miyoshi, T.; Jana, S. C. Hybrid fillers of lignin and carbon black for lowering of viscoelastic loss in rubber compounds. Polymer (Guildf). 2014, 55, 3825–3835. Ataie, F. F.; Riding, K. A. Use of bioethanol byproduct for supplementary cementitious material production. Constr. Build. Mater. 2014, 51, 89–96. Crampton, E. W.; Maynard, L. A.; others. The relation of cellulose and lignin content to the nutritive value of animal feeds. J. Nutr. 1938, 15 (4), 383–395. Knudsen, K. E. B. Carbohydrate and lignin contents of plant materials used in animal feeding. Anim. Feed Sci. Technol. 1997, 67 (4), 319–338. Isahak, W. N. R. W.; Hisham, M. W. M.; Yarmo, M. A.; Yun Hin, T. A review on bio-oil production from biomass by using pyrolysis method. Renew. Sustain. Energy Rev. 2012, 16 (8), 5910–5923. Oasmaa, A.; Meier, D. Norms and standards for fast pyrolysis liquids. J. Anal. Appl. Pyrolysis 2005, 73 (2), 323–334. Mohan, D.; Pittman, C. U.; Steele, P. H. Pyrolysis of wood/biomass for bio-oil: A critical review. Energy and Fuels 2006, 20 (3), 848–889. Czernik, S.; Bridgwater, A. V. Overview of Applications of Biomass Fast Pyrolysis Oil. Energy & Fuels 2004, 18, 590–598. Bridgwater, A. V. Review of fast pyrolysis of biomass and product upgrading. Biomass and Bioenergy 2012, 38, 68–94. Yang, H.; Yan, R.; Chen, H.; Lee, D. H.; Zheng, C. Characteristics of hemicellulose, cellulose and lignin pyrolysis. Fuel 2007, 86 (12–13), 1781–1788. Meier, D.; van de Beld, B.; Bridgwater, A. V.; Elliott, D. C.; Oasmaa, A.; Preto, F. State-of-the-art of fast pyrolysis in IEA bioenergy member countries. Renew. Sustain. Energy Rev. 2013, 20, 619–641. Yoshida, T.; Matsumura, Y. Gasification of Cellulose, Xylan, and Lignin Mixtures in Supercritical Water. Ind. Eng. Chem. Res. 2001, 40 (23), 5469–5474. Pindoria, R. V; Megaritis, A.; Messenböck, R. C.; Dugwell, D. R.; Kandiyoti, R. Comparison of the pyrolysis and gasification of biomass: effect of reacting gas atmosphere and pressure on Eucalyptus wood. Fuel 1998, 77 (11), 1247–1251. Watanabe, M.; Inomata, H.; Osada, M.; Sato, T.; Adschiri, T. Catalytic effects of NaOH and ZrO 2 for partial oxidative gasification of n -hexadecane and lignin in supercritical water q. Fuel 2003, 82, 545–552. van Steen, E.; Claeys, M. Fischer-Tropsch catalysts for the biomass-to-liquid process. Chem. Eng. Technol. 2008, 31 (5), 655–666. Tijm, P. J.; Waller, F.; Brown, D.. Methanol technology developments for the new millennium. Appl. Catal. A Gen. 2001, 221, 275–282.

158

J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

[26] Faravelli, T.; Frassoldati, A.; Migliavacca, G.; Ranzi, E. Detailed kinetic modeling of the thermal degradation of lignins. Biomass and Bioenergy 2010, 34 (3), 290–301. [27] Millett, M. A.; Baker, A. J.; Satter, L. D. Pretreatments to Enhance Chemical, Enzymatic, and Microbiological Attack of Cellulosic Materials. Biotechnol. Bioeng. 1975, 5, 193–219. [28] Baldrian, P. Fungal laccases: occurrence and properties. FEMS Microb. Rev. 2005, 30, 215–242. [29] Tien, M.; Kirk, T. K. Lignin-degrading enzyme from Phanerochaete chrysosporium: Purification, characterization, and catalytic properties of a unique H(2)O(2)requiring oxygenase. Proc. Natl. Acad. Sci. USA. 1984, 81 (April), 2280–2284. [30] Rollag, M.; Niswender, G. D.; Reppert, S. M.; Perlow, M. J.; Tamarkin, L.; Klein, C.; Tamarkin, L.; Chen, H.; Mcnemar, a; Sidbury, J.; et al. Lignin-Degrading Enzyme from the Hymenomycete Phanerochaete chrysosporium Burds. 1982, 2 (16), 11–13. [31] Saper, N. I.; Hartwig, J. F. Mechanistic Investigations of the Hydrogenolysis of Diaryl Ethers Catalyzed by Nickel Complexes of N-Heterocyclic Carbene Ligands. J. Am. Chem. Soc. 2017, 139 (48), 17667–17676. [32] Zakzeski, J.; Bruijnincx, P. C. A.; Jongerius, A. L.; Weckhuysen, B. M. The Catalytic Valorization of Ligning for the Production of Renewable Chemicals. Chem. Rev. 2010, 110, 3552–3599. [33] Wang, H.; Tucker, M.; Ji, Y. Recent Development in Chemical Depolymerization of Lignin: A Review. J. Appl. Chem. 2013, 2013, 1–9. [34] Oconnell, J. P.; Boudart, M. Catalytic hydrogenation of cyclohexene. AIChE J. 1978, 24 (5), 904–911. [35] Madon, R. J.; Iglesia, E. Catalytic reaction rates in thermodynamically non-ideal systems. J. Mol. Catal. A Chem. 2000, 163 (1–2), 189–204. [36] Mukherjee, S.; Vannice, M. A. Solvent effects in liquid-phase reactions II. Kinetic modeling for citral hydrogenation. J. Catal. 2006, 243 (1), 131–148. [37] Mukherjee, S.; Vannice, M. A. Solvent effects in liquid-phase reactions. I. Activity and selectivity during citral hydrogenation on Pt/SiO2 and evaluation of mass transfer effects. J. Catal. 2006, 243 (1), 108–130. [38] Román-Leshkov, Y.; Davis, M. E. Activation of carbonyl-containing molecules with solid lewis acids in aqueous media. ACS Catal. 2011, 1 (11), 1566–1580. [39] Forchheim, D.; Gasson, J. R.; Hornung, U.; Kruse, A.; Barth, T. Modeling the lignin degradation kinetics in a ethanol/formic acid solvolysis approach. part 2. validation and transfer to variable conditions. Ind. Eng. Chem. Res. 2012, 51 (46), 15053– 15063. [40] Gasson, J. R.; Forchheim, D.; Sutter, T.; Hornung, U.; Kruse, A.; Barth, T. Modeling the lignin degradation kinetics in an ethanol/formic acid solvolysis approach. Part 1. Kinetic model development. Ind. Eng. Chem. Res. 2012, 51 (32), 10595–10606.

Challenges and Opportunities of Lignin …

159

[41] Sturgeon, M. R.; Kim, S.; Lawrence, K.; Paton, R. S.; Chmely, S. C.; Nimlos, M.; Foust, T. D.; Beckham, G. T. A Mechanistic investigation of acid-catalyzed cleavage of aryl-ether linkages: Implications for lignin depolymerization in acidic environments. ACS Sustain. Chem. Eng. 2014, 2 (3), 472–485. [42] Roberts, V. M.; Stein, V.; Reiner, T.; Lemonidou, A.; Li, X.; Lercher, J. A. Towards quantitative catalytic lignin depolymerization. Chem. - A Eur. J. 2011, 17 (21), 5939–5948. [43] Beauchet, R.; Monteil-Rivera, F.; Lavoie, J. M. Conversion of lignin to aromaticbased chemicals (L-chems) and biofuels (L-fuels). Bioresour. Technol. 2012, 121, 328–334. [44] Toledano, A.; Serrano, L.; Labidi, J. Organosolv lignin depolymerization with different base catalysts. J. Chem. Technol. Biotechnol. 2012, 87 (11), 1593–1599. [45] Song, Q.; Cai, J.; Zhang, J.; Yu, W.; Wang, F.; Xu, J. Hydrogenation and cleavage of the C-O bonds in the lignin model compound phenethyl phenyl ether over a nickelbased catalyst. Chinese J. Catal. 2013, 34 (4), 651–658. [46] Chang, J.; Danuthai, T.; Dewiyanti, S.; Borgna, A. Hydrodeoxygenation of Guaiacol over Carbon-Supported Metal Catalysts. Chem. Cat. Chem. 2013, 5 (10), 304–3049. [47] Ferrari, M.; Maggi, R.; Delmon, B.; Grange, P. Influences of the Hydrogen Sulfide Partial Pressure and of a Nitrogen Compound on the Hydrodeoxygenation Activity of a CoMo/Carbon Catalyst. J. Catal. 2001, 198 (1), 47–55. [48] Saidi, M.; Samimi, F.; Karimipourfard, D.; Nimmanwudipong, T.; Gates, B. C.; Rahimpour, M. R. Upgrading of lignin-derived bio-oils by catalytic hydrodeoxygenation. Energy Environ. Sci. 2014, 7 (1), 103–129. [49] Ferrari, M.; Bosmans, S.; Maggi, R.; Delmon, B.; Grange, P. CoMo/carbon hydrodeoxygenation catalysts: influence of the hydrogen sulfide partial pressure and of the sulfidation temperature. Catal. Today 2001, 65, 257–264. [50] Zakzeski, J.; Jongerius, A. L.; Bruijnincx, P. C. A.; Weckhuysen, B. M. Catalytic lignin valorization process for the production of aromatic chemicals and hydrogen. ChemSusChem 2012, 5 (8), 1602–1609. [51] Deuss, P. J.; Barta, K. From models to lignin: Transition metal catalysis for selective bond cleavage reactions. Coordination Chemistry Reviews. 2016. [52] Sun, Z.; Fridrich, B.; De Santi, A.; Elangovan, S.; Barta, K. Bright Side of Lignin Depolymerization: Toward New Platform Chemicals. Chem. Rev. 2018, 118 (2), 614–678. [53] Renders, T.; Van Den Bosch, S.; Koelewijn, S. F.; Schutyser, W.; Sels, B. F. Ligninfirst biomass fractionation: The advent of active stabilisation strategies. Energy Environ. Sci. 2017, 10 (7), 1551–1557. [54] Corma Canos, A.; Iborra, S.; Velty, A. Chemical routes for the transformation of biomass into chemicals. Chem. Rev. 2007, 107 (6), 2411–2502.

160

J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

[55] Zhao, C.; He, J.; Lemonidou, A. a.; Li, X.; Lercher, J. a. Aqueous-phase hydrodeoxygenation of bio-derived phenols to cycloalkanes. J. Catal. 2011, 280 (1), 8–16. [56] Zhao, C.; Kou, Y.; Lemonidou, A. A.; Li, X.; Lercher, J. A. Highly Selective Catalytic Conversion of Phenolic Bio-Oil to. Angew. Chem., Int. Ed. 2009, 48, 3987– 3990. [57] Strassberger, Z.; Alberts, A. H.; Louwerse, M. J.; Tanase, S.; Rothenberg, G. Catalytic cleavage of lignin β-O-4 link mimics using copper on alumina and magnesia–alumina. Green Chem. 2013, 15 (3), 768. [58] Aho, a.; Kumar, N.; Lashkul, a. V.; Eränen, K.; Ziolek, M.; Decyk, P.; Salmi, T.; Holmbom, B.; Hupa, M.; Murzin, D. Y. Catalytic upgrading of woody biomass derived pyrolysis vapours over iron modified zeolites in a dual-fluidized bed reactor. Fuel 2010, 89 (8), 1992–2000. [59] Cortés-Jácome, M. a.; Escobar, J.; Angeles Chávez, C.; López-Salinas, E.; Romero, E.; Ferrat, G.; Toledo-Antonio, J. a. Highly dispersed CoMoS phase on titania nanotubes as efficient HDS catalysts. Catal. Today 2008, 130 (1), 56–62. [60] Müller, C. A.; Maciejewski, M.; Mallat, T.; Baiker, A. Organically Modified Titania – Silica Aerogels for the Epoxidation of Olefins and Allylic Alcohols. J. Catal. 1999, 184, 280–293. [61] Müller, C. A.; Maciejewski, M.; Mallat, T.; Baiker, A. Titania – Silica Epoxidation Catalysts Modified by Polar Organic Functional Groups. J. Catal. 2000, 232, 221– 232. [62] Resasco, D. E.; Crossley, S. Molecular Engineering Approach in the Selection of Catalytic Strategies for Upgrading of Biofuels. AIChE J. 2009, 55 (5), 1082–1089. [63] Zhao, C.; Lercher, J. A. Catalytic Depolymerization and Deoxygenation of Lignin. In The Role of Catalysis for the Sustainable Production of Bio-Fuels and BioChemicals; Triantafyllidis, K. S., Lappas, A. A., Stöcker, M., Eds.; Elsevier: Amsterdam, 2013; pp 289–320. [64] Roberts, V.; Fendt, S.; Lemonidou, A. A.; Li, X.; Lercher, J. A. Influence of alkali carbonates on benzyl phenyl ether cleavage pathways in superheated water. Appl. Catal. B Environ. 2010, 95 (1–2), 71–77. [65] He, J.; Zhao, C.; Mei, D.; Lercher, J. A. Mechanisms of selective cleavage of C-O bonds in di-aryl ethers in aqueous phase. J. Catal. 2014, 309, 280–290. [66] Sergeev, A. G.; Webb, J. D.; Hartwig, J. F. A heterogeneous nickel catalyst for the hydrogenolysis of aryl ethers without arene hydrogenation. J. Am. Chem. Soc. 2012, 134 (50), 20226–20229. [67] Sergeev, A. G.; Hartwig, J. F. Selective, Nickel-Catalyzed Hydrogenolysis of Aryl Ethers. Science (80-. ). 2011, 332 (6028), 439–443. [68] He, J.; Zhao, C.; Lercher, J. A. Ni-catalyzed cleavage of aryl ethers in the aqueous phase. J. Am. Chem. Soc. 2012, 134 (51), 20768–20775.

Challenges and Opportunities of Lignin …

161

[69] Zhang, J.; Teo, J.; Chen, X.; Asakura, H.; Tanaka, T.; Teramura, K.; Yan, N. A series of NiM (M = Ru, Rh, and Pd) bimetallic catalysts for effective lignin hydrogenolysis in water. ACS Catal. 2014, 4 (5), 1574–1583. [70] Rensel, D. J.; Rouvimov, S.; Gin, M. E.; Hicks, J. C. Highly selective bimetallic FeMoP catalyst for C-O bond cleavage of aryl ethers. J. Catal. 2013, 305, 256–263. [71] Parsell, T. H.; Owen, B. C.; Klein, I.; Jarrell, T. M.; Marcum, C. L.; Haupert, L. J.; Amundson, L. M.; Kenttämaa, H. I.; Ribeiro, F.; Miller, J. T.; et al. Cleavage and hydrodeoxygenation (HDO) of C–O bonds relevant to lignin conversion using Pd/Zn synergistic catalysis. Chem. Sci. 2013, 4 (2), 806–813. [72] Chang, J.; Danuthai, T.; Borgna, A. Catalyst for Hydrodeoxygenation Reactions. WO 2013/191661 A1, 2013. [73] Gómez-Monedero, B.; Ruiz, M. P.; Bimbela, F.; Faria, J. Selective hydrogenolysis of α-O-4, β-O-4, 4-O-5 CO bonds of lignin-model compounds and lignin-containing stillage derived from cellulosic bioethanol processing. Appl. Catal. A Gen. 2017, 541 (February), 60–76. [74] Gomez-Monedero, B.; Faria, J.; Bimbela, F.; Ruiz, M. P. Catalytic hydroprocessing of lignin B-O-4 ether bond model compound phenethyl phenyl ether over ruthenium catalysts. Biomass Convers. Biorefinery 2017, 7 (3), 385–398. [75] Xu, W.; Miller, S. J.; Agrawal, P. K.; Jones, C. W. Depolymerization and hydrodeoxygenation of switchgrass lignin with formic acid. ChemSusChem 2012, 5 (4), 667–675. [76] Song, Q.; Wang, F.; Cai, J.; Wang, Y.; Zhang, J.; Yu, W.; Xu, J. Lignin depolymerization (LDP) in alcohol over nickel-based catalysts via a fragmentation hydrogenolysis process. Energy Environ. Sci. 2013, No. 6, 994–1007. [77] Sturgeon, M. R.; O’Brien, M. H.; Ciesielski, P. N.; Katahira, R.; Kruger, J. S.; Chmely, S. C.; Hamlin, J.; Lawrence, K.; Hunsinger, G. B.; Foust, T. D.; et al. Lignin depolymerisation by nickel supported layered-double hydroxide catalysts. Green Chem. 2014, 16 (2), 824–835. [78] Toledano, A.; Serrano, L.; Pineda, A.; Romero, A. a.; Luque, R.; Labidi, J. Microwave-assisted depolymerisation of organosolv lignin via mild hydrogen-free hydrogenolysis: Catalyst screening. Appl. Catal. B Environ. 2014, 145, 43–55. [79] Korányi, T. I.; Hensen, E. J. M. Preparative Aspects of Supported Ni2P Catalysts for Reductive Upgrading of Technical Lignin to Aromatics. Catal. Letters 2017, 147 (7), 1722–1731. [80] Warner, G.; Hansen, T. S.; Riisager, A.; Beach, E. S.; Barta, K.; Anastas, P. T. Depolymerization of organosolv lignin using doped porous metal oxides in supercritical methanol. Bioresour. Technol. 2014, 161C, 78–83. [81] Wang, X.; Rinaldi, R. A route for lignin and bio-oil conversion: Dehydroxylation of phenols into arenes by catalytic tandem reactions. Angew. Chemie - Int. Ed. 2013, 52 (44), 11499–11503.

162

J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

[82] Barta, K.; Matson, T. D.; Fettig, M. L.; Scott, S. L.; Iretskii, A. V.; Ford, P. C. Catalytic disassembly of an organosolv lignin via hydrogen transfer from supercritical methanol. Green Chem. 2010, 12 (9), 1640. [83] Matson, T. D.; Barta, K.; Iretskii, A. V; Ford, P. C. One-pot catalytic conversion of cellulose and of woody biomass solids to liquid fuels. J. Am. Chem. Soc. 2011, 133 (35), 14090–14097. [84] Barta, K.; Warner, G. R.; Beach, E. S.; Anastas, P. T. Depolymerization of organosolv lignin to aromatic compounds over Cu-doped porous metal oxides. Green Chem. 2014, 16 (1), 191–196. [85] Ye, Y.; Zhang, Y.; Fan, J.; Chang, J. Selective production of 4-ethylphenolics from lignin via mild hydrogenolysis. Bioresour. Technol. 2012, 118, 648–651. [86] Liguori, L.; Barth, T. Palladium-Nafion SAC-13 catalysed depolymerisation of lignin to phenols in formic acid and water. J. Anal. Appl. Pyrolysis 2011, 92 (2), 477–484. [87] Huang, X.; Ouyang, X.; Hendriks, B. M. S.; Gonzalez, O. M. M.; Zhu, J.; Korányi, T. I.; Boot, M. D.; Hensen, E. J. M. Selective production of mono-aromatics from lignocellulose over Pd/C catalyst: The influence of acid co-catalysts. Faraday Discuss. 2017, 202, 141–156. [88] Renders, T.; Schutyser, W.; Van Den Bosch, S.; Koelewijn, S. F.; Vangeel, T.; Courtin, C. M.; Sels, B. F. Influence of Acidic (H3PO4) and Alkaline (NaOH) Additives on the Catalytic Reductive Fractionation of Lignocellulose. ACS Catal. 2016, 6 (3). [89] Li, S.; Li, W.; Zhang, Q.; Shu, R.; Wang, H.; Xin, H.; Ma, L. Lignin-first depolymerization of native corn stover with an unsupported MoS2 catalyst. RSC Adv. 2018, 8 (3), 1361–1370. [90] Xiao, L. P.; Wang, S.; Li, H.; Li, Z.; Shi, Z. J.; Xiao, L.; Sun, R. C.; Fang, Y.; Song, G. Catalytic Hydrogenolysis of Lignins into Phenolic Compounds over Carbon Nanotube Supported Molybdenum Oxide. ACS Catal. 2017, 7 (11), 7535–7542. [91] Song, Q.; Wang, F.; Xu, J. Hydrogenolysis of lignosulfonate into phenols over heterogeneous nickel catalysts. Chem. Commun. 2012, 48 (56), 7019. [92] 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 Chem. 2014, 16 (2), 897. [93] Jie-wang, Y.; Gui-zhen, F.; Chun-de, J. Hydrogenation of Alkali Lignin Catalyzed by Pd/C. APCBEE Procedia 2012, 3 (May), 53–59. [94] Key, R. E.; Bozell, J. J. Progress toward Lignin Valorization via Selective Catalytic Technologies and the Tailoring of Biosynthetic Pathways. ACS Sustain. Chem. Eng. 2016, 4 (10), 5123–5135. [95] Van Den Bosch, S.; Renders, T.; Kennis, S.; Koelewijn, S. F.; Van Den Bossche, G.; Vangeel, T.; Deneyer, A.; Depuydt, D.; Courtin, C. M.; Thevelein, J. M.; et al.

Challenges and Opportunities of Lignin …

163

Integrating lignin valorization and bio-ethanol production: On the role of NiAl2O3catalyst pellets during lignin-first fractionation. Green Chem. 2017, 19 (14), 3313–3326. [96] Schutyser, W.; Van Den Bossche, G.; Raaffels, A.; Van Den Bosch, S.; Koelewijn, S. F.; Renders, T.; Sels, B. F. Selective Conversion of Lignin-Derivable 4Alkylguaiacols to 4-Alkylcyclohexanols over Noble and Non-Noble-Metal Catalysts. ACS Sustain. Chem. Eng. 2016, 4 (10), 5336–5346. [97] Van Den Bosch, S.; Renders, T.; Kennis, S.; Koelewijn, S. F.; Van Den Bossche, G.; Vangeel, T.; Deneyer, A.; Depuydt, D.; Courtin, C. M.; Thevelein, J. M.; et al. Integrating lignin valorization and bio-ethanol production: On the role of Ni-Al2O3 catalyst pellets during lignin-first fractionation. Green Chem. 2017, 19 (14), 3313– 3326. [98] Anderson, E. M.; Stone, M. L.; Katahira, R.; Reed, M.; Beckham, G. T.; RománLeshkov, Y. Flowthrough Reductive Catalytic Fractionation of Biomass. Joule 2017, 1 (3), 613–622. [99] Anderson, E. M.; Stone, M. L.; Hülsey, M. J.; Beckham, G. T.; Roman-Leshkov, Y. Kinetic Studies of Lignin Solvolysis and Reduction by Reductive Catalytic Fractionation Decoupled in Flow-Through Reactors. ACS Sustain. Chem. Eng. 2018, 6, 7951–7959. [100] Kumaniaev, I.; Subbotina, E.; Sävmarker, J.; Larhed, M.; Galkin, M. V.; Samec, J. S. M. Lignin depolymerization to monophenolic compounds in a flow-through system. Green Chem. 2017, 19 (24), 5767–5771. [101] Anderson, E. M.; Katahira, R.; Reed, M.; Resch, M. G.; Karp, E. M.; Beckham, G. T.; Roman-Leshkov, Y. Reductive Catalytic Fractionation of Corn Stover Lignin. ACS Sustain. Chem. Eng. 2016, 4, 6940–6950. [102] Sette, M.; Wechselberger, R.; Crestini, C. Elucidation of lignin structure by quantitative 2D NMR. Chem. Eur. J. 2011, 17 (34), 9529–9535. [103] Lange, J. P. Don’t Forget Product Recovery in Catalysis Research—Check the Distillation Resistance. ChemSusChem 2017, 10 (1), 245–252. [104] Lee, J.; Kim, Y. T.; Huber, G. W. Aqueous-phase hydrogenation and hydrodeoxygenation of biomass-derived oxygenates with bimetallic catalysts. Green Chem. 2014, 16 (2), 708. [105] Michel, C.; Gallezot, P. Why Is Ruthenium an Efficient Catalyst for the AqueousPhase Hydrogenation of Biosourced Carbonyl Compounds? ACS Catal. 2015, 5 (7), 4130–4132. [106] Ravenelle, R. M.; Schübler, F.; Damico, A.; Danilina, N.; Van Bokhoven, J. A.; Lercher, J. A.; Jones, C. W.; Sievers, C. Stability of zeolites in hot liquid water. J. Phys. Chem. C 2010, 114 (46), 19582–19595.

164

J. Faria, B. Gómez-Monedero and M. P. Ruíz-Ramiro

[107] Zhang, L.; Chen, K.; Chen, B.; White, J. L.; Resasco, D. E. Factors that Determine Zeolite Stability in Hot Liquid Water. J. Am. Chem. Soc. 2015, 137 (36), 11810– 11819. [108] Zapata, P. A.; Huang, Y.; Gonzalez-Borja, M. A.; Resasco, D. E. Silylated hydrophobic zeolites with enhanced tolerance to hot liquid water. J. Catal. 2013, 308, 82–97. [109] Zapata, P. A.; Faria, J.; Ruiz, M. P.; Jentoft, R. E.; Resasco, D. E. Hydrophobic zeolites for biofuel upgrading reactions at the liquid-liquid interface in water/oil emulsions. J. Am. Chem. Soc. 2012, 134, 8570–8578. [110] Shakeri, M.; Roiban, L.; Yazerski, V.; Prieto, G.; Klein Gebbink, R. J. M.; de Jongh, P. E.; de Jong, K. P. Engineering and Sizing Nanoreactors To Con fi ne Metal Complexes for Enhanced Catalytic Performance. Nat. Mater. 2014, 4, 3791–3796. [111] Prieto, G.; Shakeri, M.; De-Jong, K. P.; De-Jong, P. E. Quantitative Relationship between Support Porosity and the Stability of Pore-Confined Metal Nanoparticles Synthesis Catalysts. ACS Nano 2014, No. Xx, 2522–2531. [112] Wang, F.; Mielby, J.; Richter, F. H.; Wang, G.; Prieto, G.; Kasama, T.; Weidenthaler, C.; Bongard, H. J.; Kegnæs, S.; Fürstner, A.; et al. A polyphenylene support for pd catalysts with exceptional catalytic activity. Angew. Chemie - Int. Ed. 2014, 53 (33), 8645–8648. [113] Oneill, B. J.; Jackson, D. H. K.; Lee, J.; Canlas, C.; Stair, P. C.; Marshall, C. L.; Elam, J. W.; Kuech, T. F.; Dumesic, J. A.; Huber, G. W. Catalyst design with atomic layer deposition. ACS Catal. 2015, 5 (3), 1804–1825. [114] Aranda-Pérez, N.; Ruiz, M. P.; Echave, J.; Faria, J. Enhanced activity and stability of Ru-TiO2 rutile for liquid phase ketonization. Appl. Catal. A Gen. 2017, 531. [115] Dietrich, P. J.; Sollberger, F. G.; Akatay, M. C.; Stach, E. A.; Delgass, W. N.; Miller, J. T.; Ribeiro, F. H. Structural and catalytic differences in the effect of Co and Mo as promoters for Pt-based aqueous phase reforming catalysts. Appl. Catal. B Environ. 2014, 156–157, 236–248. [116] Gounder, R. Hydrophobic microporous and mesoporous oxides as Bronsted and Lewis acid catalysts for biomass conversion in liquid water. Catal. Sci. Technol. 2014, 4 (9), 2877–2886. [117] Resasco, D. E.; Sitthisa, S.; Faria, J.; Prasomsri, T.; Ruiz, M. P. Heterogeneous Catalysis in Biomass to Chemicals and Fuels; Kubicka, D., Kubicková, I., Eds.; Research Signpost: Kerala, 2011; Vol. 661. [118] Lu, J.; Elam, J.; Stair, P. Synthesis and stabilization of supported metal catalysts by atomic layer deposition. Acc. Chem. Res. 2013, 46 (8).

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 7

THE POTENTIAL ROLE OF ENZYMATIC CATALYSIS AND METABOLIC ENGINEERING IN LIGNIN VALORIZATION Wenya Wang1,2,*, Chen Shi1 and Robert J. Linhardt3 1

College of Life Science and Technology, Beijing University of Chemical Technology, Beijing, China 2 State Key Laboratory of Pulp and Paper Engineering, South China University of Technology, Guangzhou, China 3 Departments of Chemistry and Chemical Biology, Chemical and Biological Engineering, Center for Biotechnology and Interdisciplinary Studies, Rensselaer Polytechnic Institute, Troy, NY, US

ABSTRACT Lignin, the second richest biomass, is considered to be the potential resource of high value-added chemicals with the rapid development of lignocellulose biorefinery. During the past decades, physical and chemical methods have been used widely to degrade lignin into chemicals; however, they are lagged by several challenges, such as the fewer yields of low-weight molecules, higher energy consumption and non-specific cleavage. A large number of metabolic pathways in organism have been discovered for the lignin degradation and bioconversion, which was consist of lignolytic enzymes including radical lignolytic enzymes and non-radical lignolytic enzymes. The metabolic engineering of non-radical lignolytic enzymes shows a promising value for the conversion of lignin into aromatic chemicals or other high value-added chemicals. In the present mini-review, recent developments on enzyme catalysis and metabolic engineering of lignin valorizaiton will be *

Corresponding Author Email: [email protected]

166

Wenya Wang, Chen Shi and Robert J. Linhardt summarized and discussed, including already discovered non-radical lignolytic enzymes, their metabolic pathways and molecular mechanism for lignin conversion, their recent application in lignin biorefinery and the possible combination of bio-catalyst and physical/chemical methods for lignin refinery.

Keywords: lignin valorization, non-radical lignolytic enzymes, enzyme catalysis, metabolic engineering, high value-added chemicals

INTRODUCTION Lignin is a major component of lignocellulose and is also the most abundant aromatic polymer on earth. Lignin has a highly branched, three-dimensional, poly-phenolic structure that includes three phenylpropane units, namely p-coumaryl, coniferyl and sinapyl, joined by ether and C−C linkages. While lignin’s structural heterogeneity and poly-phenolic composition results in its inherent stability and recalcitrance, lignin can ultimately be converted into CO2 by microorganisms (bacteria, fungi, actinomycetes, etc.), maintaining the carbon balance in nature. The biological degradation of lignin by microorganisms is a complex process, involving the extracellular oxidative degradation of natural lignin into low-molecular weight aromatics, followed by their intracellular metabolic degradation, and their bioconversion as carbon and energy sources. The enzymes involved in these processes can be classified, on the basis of their reaction mechanism, as radical-dependent and nonradical lignolytic enzymes [1, 2], which constitute the metabolic pathways of lignin degradation in nature. Generally, the radical-dependent lignolytic enzymes are secreted extracellularly to produce the free radicals that degrade lignin into low-molecular-weight products. These radical-dependent lignolytic enzymes have been studied extensively over the past few decades, and have been thoroughly reviewed [3, 4]. With the rapid development of lignocellulose biorefinery, the intracellular metabolic degradation and bioconversion of lignins with a low-molecular weight aromatics has provided a new approach for the lignin volarization.

THE BIO-DEGRADATION OF LIGNIN-DERIVED AROMATIC Currently, the detailed metabolic pathways, transportation and regulation of ligninderived aromatics are mainly associated with bacterial resources. Corresponding research into fungi have lagged behind, and the metabolic pathways for aromatics in fungi have been mostly proposed on the basis of discovered intermediates. Early studies indicated that the production of ligninolytic enzymes in fungi occurred during secondary metabolism and were mainly triggered by limited nutrient levels, including carbon and nitrogen limitations

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

167

[5, 6]. These studies suggest that the large scale production of ligninolytic enzymes in fungi is not directly related to the catabolism of aromatics [7]. Thus, the metabolism and regulation of lignin-derived aromatics in fungi is far more complicated than those in bacteria. Lignin is a highly branched three-dimensional poly-phenolic structure, consisting of phydroxyphenyl (H), guaiacyl (G), and syringyl (S) units. In addition, β-O-4 linkages account for 40% to 70% of chemical bonds in natural lignin. Consequently, the metabolism of lignin-derived aromatics would necessarily involve enzymatic hydrolysis of β-O-4 linkages and the metabolic pathways of lignin derivatives from G-/S-/H-type units.

The Enzymatic Degradation of β-Ethers It has been discovered that enzymes acting on β-ethers can cleave β-O-4 bonds in lignin model compounds. The β-etherase system of Sphingobium sp. SYK-6 is most well-known among these enzymes. The degradation of β-O-4 bond by Sphingobium sp. SYK-6 includes Cα dehydrogenase (LigD, LigL, LigN, LigO), etherase enzyme (LigE, LigF, LigP) and glutathione-lyase (LigG) (Figure 1). The genes encoding these three kinds of enzymes have a number of family members, which have been reviewed by Wang [8] and Kamimura [9]. The mining of these three kinds of enzymes was undertaken for the National Center for Biotechnology Information (NCBI) by Kamimura and coworkers to identify additional βether enzymes [9]. The -etherase and C-dehydrogenase gene orthologs are mainly distributed in Sphingomonadaceae with the exception of LigE orthologs that are somewhat widely distributed in α-proteobacteria, in which Sphingomonadaceae represents 53%. The orthologs of the glutathione-removing enzyme gene were mainly distributed in αproteobacteria, however, the Nu class of Glutathione S-transferase (GST) from Novosphingobium sp. MBES04 (GST3), with stereospecificity for both R and S substrates, is distributed in α-proteobacteria, β-proteobacteria and γ-proteobacteria [9]. These results indicate that in nature Sphingomonadaceae is specialized for  -aryl ether catabolism, which is in agreement with the reports of the Bugg and coworkers [10]. The different family members of the  -etherase systems show a variety of pH optima, thermal stabilities and substrate specificities. In addition to their enantioselective differences, the chemical functionality of their substrates can significantly impact the activity of these enzymes. These functionalities include groups on the aromatic rings, linkages between two aromatic rings, groups at Cα, the side chain of Cβ, and the positions of methoxy groups on the aromatic rings. In particular, the presence of a hydroxyl group at the Cα position of lignin model compounds makes the β-O-4 bond resistant to the β-etherase. When the group at this Cα position is replaced by carbonyl, the β-etherase can effectively cleave the β-ether bond linking the aromatic groups in dimeric compounds [11, 12].

168

Wenya Wang, Chen Shi and Robert J. Linhardt

Figure 1. Reaction procedure of the LigDFG enzyme system. GGE, guaiacylglycerol-β-guaniacyl ether; MPHPV, α-(2-methoxyphenoxy)- β-hydroxypropiovanillone; GS-HPV, α-glutathionyl-βhydroxypropiovanillone; and HPV, β-hydroxypropiovanillone.

The large number of gene family members identified might have evolved to adapt to the environment and the intrinsic heterogeneity of lignin in order to effectively utilize lignin as a carbon and an energy source. However, from the point of industrial application, the presence of a large number of enzymes makes a process unduly complex. Thus, it is important to study the catalytic mechanisms of these enzymes to discover the enzymes that act on a broad range of substrates. Recently, the X-ray crystal structures of β-ether degrading enzymes have been reported [13-15] and these will undoubtedly be used in combination with gene mining and protein engineering to promote studies on β-ether degrading enzymes.

The Metabolic Pathways of Lignin Derivatives from G-/S-/H-Units In practice, natural lignin can be depolymerized into a large number of chemical products after physical, chemical or biological treatment. The type and number of these chemical products are dependent on the operating conditions. In the present paper, the metabolic pathways of depolymerizing products only focus on the aromatic slurry. The metabolic pathways of p-coumaric acid (pCA), ferulic acid (FA) and sinapic acid (SA) have been studied extensively. G-/S-/H- units and many lignin-derived aromatics share the common chemical structure with pCA, FA and SA. Consequently, the metabolic pathways of pCA, FA and SA have been chosen to introduce the catabolic pathways for ligninderived aromatics. pCA and FA can be catabolized to protocatechuate (PCA) or catechol (CA) through CoA-dependent β-oxidation and CoA-dependent non-β-oxidation pathways (Figure 2A). In the CoA-dependent β-oxidation pathway, CoA is first catalytically added, followed by double bond hydrolysis, further oxidation and sulfurolysis to remove the ethyl group. pCA

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

169

and FA are eventually hydrolyzed to p-hydroxybenzoic acid (pHBA) and vanillic acid, respectively [8, 16, 17] (Figure 2B1, B5). The CoA-dependent non-β-oxidation pathway is similar to the β-oxidation pathway with the exception that in the CoA-dependent βoxidation pathway further thiolysis and oxidation were not observed after the first removal of acetyl-CoA [16, 18] (Figure 2B2, B6). Additionally, there is also a side chain reduction pathway in the metabolism of pCA and FA. In the plants Glechoma sp. and Vanilla sp., a CoA-independent pathway produces pHBA from pHA, which proposed to involve one reductase (pHBALS) and one decarboxylase (pHBALS) (Figure 2B7) [16, 19]. In Corynebacterium glutamicum, FA is reduced by aromatic reductase (RE) to form dihydroferulic acid and decarboxylase (DCL) catalyzes dihydroferulic acid to produce acetic acid and vanillic acid. This pathway is generally present in bacteria and fungi grown under anaerobic conditions [8]. In addition, FA can also be reduced to coniferyl alcohol, which is further degraded to vanillic acid [20, 21] (Figure 2B4). Notably, FA can also be degraded by non-oxidative decarboxylation [8, 22, 23], which has not been observed in pCA catabolism. FA, in the non-oxidative decarboxylation pathway, is catalytically decarboxylated to vanillin and vanillic acid (Figure 2B3), which has been found in Fusarium solani (Mart) Sacc., [24] Bacillus coagulans [25], and Bacillus cereus strain PN24 [26]. Most catabolic microorganisms for pCA and FA cannot degrade SA, which indicates the adverse impact of aromatic methoxyl groups on SA catabolism.[27] The initial step from SA to syringic acid is proposed to be catalyzed by radical lignolytic enzymes (laccases) [27, 28] or non-radical lignolytic enzymes (decarboxylases) [29, 30] (Figure 2A). No genes corresponding to the decarboxylases involved in non-radical catalysis have yet been reported. SA is transformed into syringic acid through the removal of two carbon atoms from its side chain. In Sphingomonas paucimobilis SYK-6, syringic acid is Odemethylated by a tetrahydrofolate-dependent O-demethylase (DesA) to produce 3-Omethylgallate (3MGA), and then the 3MGA is O-demethylated by another O-demethylase (LigM) to generate gallic acid. The aromatic ring of 3MGA and gallic acid can be cleaved by dioxygenase, moving into the TCA cycle [31-33]. There is currently no evidence to support the transformation of syringic acid to PCA or CA prior to ring opening.

THE TRANSPORTATION OF LIGNIN DERIVATIVES FROM G-/S-/H-UNITS AND THEIR METABOLIC REGULATION The genes involved in aromatic metabolism are usually physically assembled in operons or in clusters within bacteria. Around the aromatic metabolizing genes there are always transporting genes responsible for the uptake of the aromatic substrates and

170

Wenya Wang, Chen Shi and Robert J. Linhardt

regulating genes, which act as trans-elements to regulate the aromatic metabolism. Metabolizing genes, transporting genes, and regulating genes have developed to efficiently assimilate the substrate and catabolize it for carbon and energy demands. In bacteria, there are four transporting systems for the uptake of lignin-derived aromatic compounds: ATP-binding cassette (ABC) transporters, major facilitator superfamily (MFS) transporters, a tripartite ATP-independent periplasmic (TRAP) transporter, and an ion transporter (IT) superfamily. The four transporting systems show signs of binding affinity towards pCA, FA, SA or their derivatives [9]. ABC transporters are widely distributed and are well-studied transporters. They can transport an enormous variety of substrates, ranging from small ions to large organic molecules [34]. When Enterobacter lignolyticus SCF1 and Bacillus ligniniphilus L1 was cultivated in media with lignin as sole carbon source, the production of ABC transporters increased [35], demonstrating that the ABC transporters were involved in the assimilation of lignin-derived aromatics. A fluorescence thermal shift-based assay indicated that the component of ABC transporters can interact with pCA, FA and SA with different binding affinities [36]. In Rhodopseudomonas palustris, the co-crystallization of ABC transporter (CouP) and FA indicates that H-bond interactions occur between the 4-OH group of the aromatic ring and His309/Gln305 and also between the carboxyl group on the FA side chain and Arg197, Ser222, and Thr102 [37]. CouT in R. jostii RHA1 is believed to be a MFS transporter for p-hydroxycinnamate. At the mRNA level, couT is upregulated in a strain cultivated on FA and pCA. The growth status indicates that pCA is better than FA as the growth substrate, reflecting the substrate specificity of couT [38]. In addition, there are also porins, substrate-specific channels or TonB-dependent receptors at the outer membrane of Gram-negative bacteria, involved in the uptake of aromatics [9]. There are not direct reports of their involvement in the transport of pCA, FA, and SA. The metabolizing genes of aromatics are usually clustered in the operon, around which regulating genes were recruited to manipulate the metabolic pathway and transportation for the effective catabolism. While the regulators of many aromatic monomers have been discovered, no regulators of lignin-derived biaryls have been reported. The FerC from Sphingobium sp. SYK-6 recognizes the thioester product of pCA, FA, and SA as effectors [9]. This suggests that FerC and its cognate FA catabolic regulon in SYK-6 are involved in the metabolism of lignin derivatives from G-/S-/H-type units. Other reported regulators only bind a portion of feruloyl-CoA, p-coumaroyl-CoA and sinapoyl-CoA. For example, CouR from R. jostii RHA1 and HcaR from Acinetobacter sp. ADP1 recognizes pcoumaroyl-CoA and feruloyl-CoA as effectors [39, 40], and CouR from R. palustris CGA009 and FerR from P. fluorescens BF13 bind p-coumaroyl-CoA and feruloyl-CoA, respectively [41, 42]. In addition to the regulators of the metabolic pathways of pCA, FA, and SA, the regulating genes for protocatechuate [9], vanillate [9], benzoate [43], and

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

171

catechol [44], the intermediates of pCA, FA, and SA catabolism, have also been discovered. This suggests that the regulation mechanism of lignin-derived aromatics is a complicated system involving substrate competition, the integration of different metabolic pathways, and environmental factors. In Pseudomonas putida, the catabolite repressor (Crc) binds to and inhibits the translation of benR mRNAs, and benR encodes the transcriptional activator inducing the expression of benzoate degradation genes. Further research indicates that Crc can also bind to the translation initiation regions of the mRNA of several structural genes in benzoate degradation and the benzoate transporter gene, which suggests that Crc may also control benzoate degradation and uptake. All this evidence demonstrates that Crc can regulate benzoate metabolism at multi-tier levels, including uptake, induction and degradation [45]. The catA2 gene in Pseudomonas putida mt-2 works as a enzymatic safety valve for excess of catechol, the intermediate of benzoate catabolism, to alleviate the toxicity of catechol. The catA2 gene is located downstream of the ben operon and the ben operon is regulated by the benR. CatA is in the cat operon for the normal catechol catabolism, which is activated by carR [44]. PcaY is an aromatic acid chemoreceptor in P. putida F1, which belongs to the pca operon under the control of PcaR. PcaK, a transporter protein, is also in the pca operon. Data suggests that PcaK facilitates the uptake of 4-hydroxybenzoate, resulting in the increase of pcaY expression. Consequently, the chemotaxis, transport, and metabolism of aromatic compounds are integrated by PcaR in P. putida [46]. Recent studies show the existence of an aerobicanaerobic metabolism switch [43] and a regulator coupled with dioxygenase [47] in aromatic catabolism. This provides many potential tools for increasing lignin volarization with microbiology by metabolic engineering.

BIOTRANSFORMATION FOR THE PRODUCTION OF CHEMICALS FROM DEPOLYMERIZED LIGNIN AND LIGNIN MODEL COMPOUNDS There are two approaches for the biotransformation of depolymerized lignin or lignin model compounds into chemicals. The first is in vitro enzymatic conversion and the second is in vivo metabolic conversion. Given that 40% to 70% of the chemical linkages in lignin are β-O-4, researchers have tried to degrade the lignin using a β-etherase system to obtain value-added chemicals or chemical precursors for the further metabolic engineering (Table 1). However, improving degradation efficiency is a big challenge for in vitro enzymatic catalysis. Moreover, although some value-added chemicals have been produced from lignin using microorganisms, most reports still rely on lignin model compounds to study the metabolic conversion in vivo.

Figure 2. Metabolic pathways of p-coumaric acid, ferulic acid and sinapic acid. (A) The brief outline of metabolic pathways; (B) Detailed metabolic pathways. BADH, benzaldehdye dehydrogenase; ECH, enoyl-CoA hydratase; HADH, 3-hydroxyacyl-CoA dehydrogenase; HDE, hydroxylase; pHBALS, phydroxybenzaldehydesynthase; pHCHL, p-hydroxycinnamoyl-CoA hydratase/lyase; pHBDC, p-hydroxybenzoate decarboxylase; pHB3H, p-hydroxybenzoate3-hydroxylase; pHCS, p-hydroxycinnamoyl-CoA synthetase; KAT, 3-ketoacyl-CoA thiolase; PCADC, protocatechuate deccarboxylase;DCL, decarboxylase; DHG, dehydrogenase; ECH, enoyl-CoA-hydratase; ECH/A, enoyl-CoA-hydratase/aldolase; FCS, feruloyl-CoA synthelase; FDC, ferulic acid decarboxylase; βKTE, β-keto thiolase; O-DML, O-demethylase; PCADC, protocatechuate deccarboxylase; POE, phenol oxidase; RE, reducase; TE, thiolase; VD, vanillin decarboxylase; VGDH, 4-vinylguaiaol dehydrogenase;3MGA, 3-O-methylgallate; PDC, pyrone-4,6-dicarboxylic acid.

Table 1. The enzymatic conversion in vitro of natural lignin Lignin resource

Involved enzymes

Process

The reacting conditions

Products and yield

softwood/hardwood alkali-lignin and bagasse organosolv-lignin

LigD/ligF/ligG/AVR

One pot

0.1mM ammounium acetate buffer (pH 9.0); NAD+; GSH

C-MWL and EMWLb

SDR3/ SDR55/ GST4/ GST5/ GST3

One pot

0.1M TAPS(pH 8.5); NAD+; GSH; 15℃

OrganoCat lignin from beech

Laccase/LigE/LigFNA/LigG-TD

Three steps

HP lignin and MCS ligninc

LigD/LigN/LigE/LigF /NaGSTNU (or LigG)/ AvGR

One pot

Setp I: violuric acid; 0.1M sodium acetate buffer (pH 5); [EMIM] [EtSO4]; room temperature; 250 rpm; 5 days. Setp II: 50 mM glycine/NaOH buffer (pH 9.5); DMSO (25%); GSH;25℃; overnight. Setp III: DMSO in step II was diluted to 10% with water; 20℃; overnight. 25 mM Tris buffer (pH 8.0); DMSO (2.0%); NAD+; GSH; room temperature; 4 hours

Guaiacol/Ferulic acid/Eugenol/ Acetovanillone/Vanillin (Softwood); Guaiacol/Vanillin (Hardwood); No products detected(Organosolv-lignin)a GHP from C-MWL (2.4 ± 0.005 wt % [24 ± 0.05 mg g1 lignin]); GHP/SHP from E- MWL (1.9 ± 0.012 and 4.7 ± 0.025 wt % [19 ± 0.12 and 47 ± 0.25 mg g-1 lignin], respectively) Oliy faction (coniferylaldehyde, guaiacyl unies, syringyl units and oligomeric species); 12.5 wt% yield.

a: Yield referred to Reiter[1]. b: Milled lignin from Cryptomeria japonica (C-MWL) and milled lignin from Eucalyptus globulus (E-MWL). c: HP - the high-syringyl hybrid poplar; MCS - maize corn stover. d: HPV- hydroxypropiovanillone; HPS- hydroxypropiosyringone.

1.0mM HPS from HP lignin (12.5 wt% yield); 0.4 mM HPV, 0.1 mM HPS and tricin from MCS lignin (5% yield (HPV+HPS))d.

The organic solution for lignin dissolution --------

Reference

N,Ndimethylformamide

[48]

DMSO

[49]

DMSO

[50]

[1]

174

Wenya Wang, Chen Shi and Robert J. Linhardt

Enzymatic Conversion in Vitro Given the complexity of lignin structures, the large scale depolymerization lignin, into value-added functional aromatic compounds, is difficult using chemical or physical treatment [2]. Enzymatic methods, particularly ones using non-radical lignolytic enzymes, represent an alternative approach for lignin valorization under environmentally-friendly and substrate-specific conditions [2]. The cleavage of natural lignin with the β-etherase system was first carried out by Reiter [1] (Table 1). The β-etherase system included LigD/F/G and AvGR, and softwood/hardwood alkali-lignin and bagasse organosolv-lignin were selected as substrates. GPC analysis showed that softwood and hardwood alkali-lignin were only slightly degraded, but this did not occur with bagasse-organosolv lignin even after 7 days of treatment [1]. Subsequently, Picart prepared a fluorescently labeled synthetic lignin model (DHP-MUAV), a multiple polymer of coniferyl alcohol and α-O-(βmethylumbelliferyl) acetovanillone (MUAV), to assess whether β-etherases could cleave β-O-4 aryl ether linkages present in lignin-like polymers. The results showed that DHPMUAV was converted into different fragments of smaller mass, suggesting that it was possible for the β-etherase to catalyze cleavage of lignin-like polymers [12]. Afterwards, three different research groups [48-50] reported the enzymatic degradation of natural lignin with improved β-etherase systems (Table 1). The highest yield of enzymatic degradation was 12.5 wt% and different depolymerization products were obtained (Table 1). Based on data from the in vitro enzymatic conversion, the efficiency of β-etherase systems towards natural lignin or lignin-like polymers was far lower than towards lignin model dimers. This might result from the problems of enzymatic enantioselectivity, substrate availability, or the inhibitory effect of lignin on enzyme activity [51]. Consequently, more research is still required to understand the mechanism of β-etherase catalysis and the interaction between these enzymes and lignin. In addition, from the view of industrial application, the yield of enzyme-hydrolyzing products still needs to be improved. Further studies need to be carried out on the β-etherase system, such as mining or engineering enzymes to broaden their range of substrates and improve their efficiency, simplifying the process, reducing its cost.

Metabolic Conversion in Vivo The “funneling pathway” includes both the upper and lower pathways, present in many microorganisms, for the metabolism of aromatic compounds [52] (Figure 3). In the upper pathways, aromatic molecules are catabolized into several conserved intermediates, including catechol (CA) in bacteria and protocatechuate (PCA) in most fungi and some bacteria [52]. In the lower pathways, the aromatic rings of these conserved intermediates

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

175

are cleaved by dioxygenases, producing ring-opened species that are metabolized through the β-ketoadipate pathway into the TCA cycle [52, 53]. Recently, researchers have used metabolic engineering to explore the biotransformation of lignin and lignin-derived monomers into different chemicals for the purpose of lignin valorization (Table 2). These target chemicals include aromatics, ringopened chemicals, TCA cycle chemicals, lipids and lipid derived polymers (polyhydroxyalkanoate, PHA). These chemicals are distributed within upper pathways, lower pathways, TCA cycle, and lipid synthesis pathways (Figure 3).

Figure 3. Lignin valorization by combining enzymatic catalysis and metabolic engineering and using a multi-disciplinary approach.

176

Wenya Wang, Chen Shi and Robert J. Linhardt Table 2. The reported chemicals biotranformed from lignin or lignin model compounds

Molecule Pyruvate, Lactate β-Ketoadipic acid, Muconolactone Muconic acid

Substrate benzoate and p-coumarate protocatechuic acid

Host P. pudita KT2440

Reference [74]

P. pudita KT2440

[58-59]

catechol, p-coumarate, benzoate, ferulate, guaiacol, vanillin, lignin

E. coli, E. coli XL-1 Blue, P. pudita KT2440, Sphingobium sp. SYK-6, Amycolatopsis sp. ATCC 39116 Pseudomonas putida (arvilla) mt-2 ATCC 23973 Escherichia coli BL21-Gold (DE3) Rhodopseudomonas palustris

[61-68,97]

Rhodococcus jostii RHA1

[71]

Aspergillus fumigatus, P. pudita KT2440 and A514, Cupriavidus basilensis B-8, Pandoraea sp. ISTKB Rhodococcus opacus DSM 1069 and PD630, Rhodococcus jostii RHA1 VanA-, Trichosporon oleaginosus, Cunninghamella echinulata FR3 E. coli, Pseudomonas putida, Rhodococcus, Pycnoporous

[53, 76, 83-85]

Picolinic acid, HMSa

catechol

3-Carboxy-muconate Benzoic/ phydroxybenzoic acid 2,5-PDCA, 2,4-PDCAb Fatty acid, PHA

vanillin corn stover hydrolysate

Lipid

4-hydroxybenzoic acid, resorcinol, vanillic acid, lignin, sorghum biomass

Vanillin

wheat straw lignocellulose alkaline pretreated lignin, organosolv lignin, kraft lignin

[70] [60] [57]

[77-82]

eugenol, isoeugenol, [55-56, 101-104] ferulic acid, vanillic acid, lignin PDCc protocatechuate P. putida PpY1100 [69] Succinic acid DHP (Synthetic Lignin) Phanerochaete chrysosporium [75] Note: a. 2-Hydroxymuconic semialdehyde (HMS); b. Pyridine 2,5-dicarboxylic acid (2,5-PDCA) Pyridine 2,4-dicarboxylic acid (2,4-PDCA); and c. 2-Pyrone-4,6-dicarboxylic acid (PDC).

Over the past several decades, various biotechnology-based approaches have been developed for the production of vanillin. A number of specialized microorganisms have been used to produce vanillin from aromatic molecules such as eugenol, isoeugenol, FA, vanillic acid [8, 54, 55]. Lignin has also been studied as a starting material for the biocatalytic production of vanillin. When a vanillin-dehydrogenase deletion strain of R. jostii RHA1 was grown for 144 h on pretreated wheat straw lignocellulose, it was found to accumulate vanillin with yields of up to 96 mg/L [56]. In addition, ferulate catabolic pathways and β-aryl ether cleavage produces vanillin as an intermediate metabolite within different lignin degradation pathways, providing valuable approaches for the bioproduction of vanillin from lignin. When Rhodopseudomonas palustris is cultivated in corn stover hydrolyzate, the bioconversion of aromatics can be observed. The deletion of badE gene of R. palustris, encoding benzoyl-CoA reductase, results in the accumulation of benzoic acid from aromatics in hydrolyzate and the deletion of hbaBCD gene, encoding 4hydroxybenzoyl-CoA reductase, accumulates 4-hydroxybenzoic acid. The concentration

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

177

of both benzoates was at more than 1 mM. It is interesting that the R. palustris can only utilize the aromatics in lignocellulose hydrolyzate, while leaving sugars unaltered. Thus, this approach provides an effective method to remove bio-toxic factors from biomass hydrolyzate before further biotransformation [57]. Dicarboxylic acid and its derivatives originate from the cleavage of aromatic ring in central intermediates. Currently dicarboxylic acid and its derivatives account for most kinds of microbial biotransformation products coming from lignin or lignin model compounds. These chemicals include β-ketoadipic acid [58], muconotactone [58, 59], 3carboxy-muconate [60], muconic acid [60-68], 2-pyrone-4,6-dicarboxylic acid (PDC) [69], picolinic acid [70], 2-hydroxymuconic semialdehyde [70], pyridine 2,5-dicarboxylic acid (2,5-PDCA) and 2,5-dicarboxylic acid (2,4-PDCA) [71]. All these chemicals share common structures (two or more carboxylic acid groups), making these compounds, or their derivatives, useful raw materials for the synthesis of bio-based polymers. The cleavage of aromatic ring has two modes: ortho ring-cleavage and meta ringcleavage, which was carried out by intradiol dioxygenases or extraiol dioxygenases, respectively [72]. Ring cleavage can be conducted by two enzymes, a 1,2-catechol dioxygenase or a 2,3-catechol dioxygenase in catechol, and when in protocatechuate can be cleaved by three enzymes in the 2,3 (meta), 4,5 (meta), and 3,4 (ortho) positions [73]. Muconic acid, the recent focus of dicarboxylic acids, can be converted into the dicarboxylic acid, adipic acid, through hydrogenation. Adipic acid is an important precursor of nylon, plasticizers, lubricants and polyester polyols. The production of muconic acid through the aromatic catabolic pathway results in higher atom efficiency than its production through the sugar pathway. When benzoate and glucose were fermented in a DO stat fed-batch culture of KT2440-CJ102 and the pH value was maintained at 7.0 for 124 h, muconic acid was produced at a titer of 34.5 g. L−1 [64]. By deleting Crc gene, encoding catabolite repression control protein, from P. pudita KT 2440, muconate production is enhanced and the yield of muconate produced from pCA after 36 h was increased nearly 70% and the yield from FA after 72 h was more than doubled [66]. Moreover, pCA and FA from alkaline pretreated lignin were converted by engineered P. putida KT2440 to 0.70 g L-1 of muconic acid in 24 h and the molar yield was 67% [64]. Barton [67] and Sonoki [68] developed an engineered strain that could produce muconic acid with lignin or lignin hydrolysate. Recent research indicates that insufficient protocatechuate decarboxylase activity is considered to be the bottleneck in muconic acid production. Studies on the increase of protocatechuate decarboxylase activity have been performed for the improvement of muconic acid production. The results indicate that the muconic acid production could be increased by 50% with pCA as substrate after the improvement of protocatechuate decarboxylase activity [65]. The “upper pathways” in aromatic-compounds degrading organisms are utilized to integrate or funnel the heterogeneous lignin-depolymerizing slurry into a few common intermediates, such as CA (1,2-dihydroxybenzene) or pCA (3,4-dihydroxybenzoate).

178

Wenya Wang, Chen Shi and Robert J. Linhardt

Cleaving the aromatic rings gave rise to different products through the ortho (intradiol) or meta (extradiol) patterns, which are funneled into TCA cycle. The metabolic pathways of various ring-opened chemicals enter the central metabolism with different carbon efficiency and redox balance [73]. The replacement of the protocatechuate (PCA) ortho pathway in P. pudita Kt2440 with a meta-cleavage pathway from Sphingobium sp. SYK-6 results in a nearly five-fold increase in the yield of pyruvate, which indicates that the catabolic pathway from ring-cleavage to TCA cycle could be selected to optimize the yield of a desired product [74]. With synthetic lignin, Phanerochaete chrysosporium (white rot fungus) was demonstrated to accumulate succinic acid by the short-cut TCA cycles, providing a potential strain for the future lignin biorefinery [75]. The recent and rapid progress on prokaryotic lignin depolymerizing enzymes suggests that lignin valorization, involving fatty acid metabolism, might result in potential applications in the preparation of biofuels and biodegradable materials. Utilizing lignin through the fatty acid and lipid synthesis pathway began with the study on lignin model compounds (4-hydroxybenzoic acid, resorcinol, vanillic acid), and then has led to use of natural lignin in different microorganisms, such as Aspergillus fumigatus [76], Cunninghamella echinulata FR3 [28], Rhodococcus opacus [77, 78], Rhodococcus jostii [79], and Trichosporon oleaginosus [80]. The synergy between laccase and microbial lignin conversion and co-fermentation of two R. jostii strains have been examined in lipid production [79, 81]. Lipid accumulation by Rhodococcus opacus DSM 1069 reached 26.99 ± 2.88% of its cellular dry weight by utilizing pine organosolv pretreatment effluent [82]. Linger [53] reported that the alkaline depolymerized lignin could be converted by Pseudomonas putida KT2440 into medium chain length (mcl)-PHAs through integrated biological funneling pathway, and another Pseudomonas putida strain A514 was engineered to improve the yield of PHA on the basis of genomic and proteomic analysis, in which PHA content reached 73% per cell dry weight [83]. Shi [84] and Kumar [85] reported that Cupriavidus basilensis B-8 and Pandoraea sp. ISTKB could also bioconvert kraft lignin into PHA. All these metabolic conversions provide a roadmap for the further research on lignin biotransformation and suggest that more value-added products will one day be produced from lignin.

COMBINING ENZYMATIC CATALYSIS AND METABOLIC ENGINEERING WITH A MULTI- DISCIPLINARY APPROACH FOR LIGNIN VALORIZATION Given the environmentally friendly and substrate-specific characterization, lignin valorization with enzymatic catalysis and metabolic engineering for the production of high value-added chemicals represent an attractive target. However, the economic feasibility

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

179

and efficiency of such processes remains a major challenge. Consequently, a multidisciplinary approach relying on biology, chemistry and physics will be needed to develop practical process for lignin biological valorization (Figure 3). The “funneling pathway” in microorganisms represents an practical approach for integrating different aromatic substances, obtained through lignin depolymerization, into some central intermediates for high-value chemical generation [52, 53]. The tools of synthetic biology and system biology, such as biosensors [86-89], “-omic” technologies [90], and tolerance engineering [91], can be applied to reconstruct the metabolic pathways to further improve the yield of target products. Next generation industrial biotechnologies (NGIB) represent a promising method for improving the economic feasibility of fermentation processes. Costs can be reduced by continuous bioprocessing under nonsterile (open) conditions using ceramic, cement or plastic bioreactors. Most lignin-utilizing microorganisms are aromatic-toxicity tolerant matching the NIGB requirements [92]. It is necessary to generate as many different lignin monomers and oligomers as possible using suitable pretreatment methods to improve the biotransformation efficiency of lignin. Unfortunately, the degradation of lignin using radical lignolytic enzymes requires weeks [93-94] and, thus, is unsuitable for the industrial applications. Chemical depolymerization usually results in the ring opening or can cause the competing repolymerization of lignin-depolymerized products, thus, it is inefficient for lignin monomer production. More recently, the formic acid [95] and formaldehyde [96] processes, developed to produce lignin monomer in high yield, have recently become available. These pretreatments or depolymerization technologies may afford promising processes for lignin valorization through the combination of chemical depolymerization and biotransformation. In addition to combining biotransformation and chemical catalysis in lignin depolymerization, industrial demands for saving energy and resources will also require the combination of the biotransformation and chemical or physical methods in other process steps. These combinations leverage the specificity of microorganisms and advantages of continuous reactions associated with chemical catalysis, thus, overcoming the drawbacks of either a purely chemical or biological approach for the industrial production of highvalue chemicals from lignin. Studies by Linger [53] and Vardon [97] provide successful examples for the application of combination technologies in lignin bio-valorization, in which mcl-PHA and cis, cis-muconic acid from lignin bioconversion were used to produce alkenoic acids and adipic acid by chemical catalysis. Furthermore, the synthesis of picolinic acid, 2,5-PDCA and 2,4-PDCA [71] using in vivo metabolic conversion have also involved in such combinational technologies. Furthermore, besides the application in microorganism, the “funneling pathway” strategy can also be used to pool the high-value alkylphenols from pyrolysis bio-oil, generated from lignocellulosic biomass, which is referred to as the chemical “funneling pathway” [98]. The biotransformation of cellulose sugar could also provide many valuable clues to new approaches for lignin biological volarization, such as one pot conversion or platform molecules [99-100].

180

Wenya Wang, Chen Shi and Robert J. Linhardt

CONCLUSION The insolubility and structural heterogeneity inherent to lignin and the complexity of its depolymerization provides a major challenge for lignin valorization. Recently, the enzymatic catalysis and metabolic engineering of lignin volarization provide some approaches for lignin bioconversion into high-value-added chemicals. These new approaches promise the generation of more varieties of lignin-derived high-value chemicals in the near future. However, enzymatic conversion in vitro shows its lowest efficiency towards natural lignin. This calls for studies on the mechanism of enzymatic catalysis and the identification of more β-etherases through protein engineering and gene mining. In addition, the chemical yields from natural lignin using metabolic conversion cannot meet the demands of industry. Thus, natural lignin will need to be depolymerized into more aromatic monomers or oligomers for the efficient bioconversion. Ultimately, the economic feasibility and effectiveness of industrial production will undoubtedly require the combination of biological, physical and chemical technologies for lignin biological valorization.

ACKNOWLEDGMENTS This work was supported by State Key Laboratory of Pulp and Paper Engineering (201760), Natural Science Foundation of China (NSFC 21576153), Beijing Natural Science Foundation (5162019).

REFERENCES [1]

[2]

[3] [4]

Reiter, J., Strittmatter, H., Wiemann, L., Schieder, D. & Sieber, V. (2013). “Enzymatic cleavage of lignin β-O-4 aryl ether bonds via net internal hydrogen transfer.” Green Chem, 15, 1373–1381. Picart, P., Dominguez de Maria, P. & Schallmey, A. (2015). “From gene to biorefinery: microbial β-etherases as promising biocatalysts for lignin valorization.” Front Microbiol, 6, 916-923. Wong, D. W. S. (2009). “Structure and action mechanism of ligninolytic enzymes.” Appl Biochem Biotech, 157, 174–209. Pollegioni, L., Tonin, F. & Rosini, E. (2015). “Lignin-degrading enzymes.” FEBS J, 282, 1190–1213.

The Potential Role of Enzymatic Catalysis and Metabolic Engineering … [5]

[6]

[7]

[8]

[9]

[10] [11]

[12]

[13]

[14]

[15]

181

Jeffries, T. W., Choi, S. & Kirk, T. K. (1981). “Nutritional regulation of lignin degradation by Phanerochaete chrysosporium.” Appl Environ Microbiol, 42(2), 290–296. Kirk, T. K., Croan, S., Tien, M., Murtagh, K. E. & Farrell, R. L. (1986). “Production of multiple ligninases by Phanerochaete chrysosporium: effect of selected growth conditions and use of a mutant strain.” Enzyme Microb Technol, 8(1), 27–32. Asina, F., Brzonova, I., Kozliak, E., Kubátová, A. & Ji, Y. (2017). “Microbial treatment of industrial lignin: successes, problems and challenges.” Renew Sust Energ Rev, 77, 1179-1205. Wang, W., Zhang, C., Sun, X., Su, S., Li, Q. & Linhardt, R. J. (2017). “Efficient, environmentally-friendly and specific valorization of lignin: promising role of nonradical lignolytic enzymes.” World J Microb Biot, 33(6), 125-139. Kamimura, N., Takahashi, K., Mori, K., Araki, T., Fujita, M., Higuchi, Y., et al. (2017). “Bacterial catabolism of lignin‐derived aromatics: new findings in a recent decade: update on bacterial lignin catabolism.” Environ Microbiol Rep, 9(6), 679705. Bugg, T. D., Ahmad, M., Hardiman, E. M. & Rahmanpour, R. (2011). “Pathways for degradation of lignin in bacteria and fungi.” Nat Prod Rep, 28, 1883-1896. Gall, D. L., Ralph, J., Donohue, T. J. & Noguera, D. R. (2014). “A group of sequence-related sphingomonad enzymes catalyzes cleavage of β-aryl ether linkages in lignin β-guaiacyl and β-syringyl ether dimers.” Environ Sci Technol, 48(20), 12454–12463. Picart, P., Mueller, C., Mottweiler, J., Wiermans, L., Bolm, C., Dominguez de Maria, P. & Schallmey, A. (2014). “From gene towards selective biomass valorization: bacterial beta-etherases with catalytic activity on lignin-like polymers.” Chemsuschem, 7, 3164–3171. Meux, E., Prosper, P., Masai, E., Mulliert, G., Dumarcay, S., Morel, M., Didierjean, C., Gelhaye, E. & Favier, F. (2012). “Sphingobium sp. SYK-6 Lig G involved in lignin degradation is structurally and biochemically related to the glutathione transferase omega class.” FEBS Lett, 586, 3944–3950. Helmich, K. E., Pereira, J. H., Gall, D. L., Heins, R. A., McAndrew, R. P., Bingman, C., Deng, K., Holland, K. C., Noguera, D. R., Simmons, B. A., Sale, K. L., Ralph, J., Donohue, T. J., Adams, P. D. & Phillips, G. N. Jr. (2016). “Structural basis of stereospecificity in the bacterial enzymatic cleavage of β-aryl ether bonds in lignin.” J Biol Chem, 291, 5234–5246. Pereira, J. H., Heins, R. A., Gall, D. L., McAndrew, R. P., Deng, K., Holland, K. C., Donohue, T. J., Noguera, D. R., Simmons, B. A., Sale, K. L., Ralph, J. & Adams, P. D. (2016). “Structural and biochemical characterization of the early and late enzymes in the lignin β-aryl ether cleavage pathway from Sphingobium sp. SYK-6.” J Biol Chem, 291, 10228–10238.

182

Wenya Wang, Chen Shi and Robert J. Linhardt

[16] Jung, D. H., Kim, E. J., Jung, E., Kazlauskas, R. J., Choi, K. Y. & Kim, B. G. (2016). “Production of p-hydroxybenzoic acid from p-coumaric acid by Burkholderia glumae BGR1.” Biotechnol Bioeng, 113(7), 1493–1503. [17] Priefert, H., Rabenhorst, J. & Steinbüchel, A. (2001). “Biotechnological production of vanillin.” Appl Microbiol Biotechnol, 56, 296–314. [18] Pan, C., Oda, Y., Lankford, P. K., Zhang, B., Samatova, F. N., Pelletier, A. D., Harwood, S. C. & Hettich, L. R. (2008). “Characterization of anaerobic catabolism of p-coumarate in Rhodopseudomonas palustris by integrating transcriptomics and quantitative proteomics.” Mol Cell Proteom, 7, 938–948. [19] Gallage, N. J., Hansen, E. H., Kannangara, R., Olsen, C. E., Motawia, M. S., Jørgensen, K., Holme, I., Hebelstrup, K., Grisoni, M. & Møller, B. L. (2014). “Vanillin formation from ferulic acid in Vanilla planifolia is catalyzed by a single enzyme.” Nat Commun, 5, 4037-4050. [20] Gupta, J. K., Hamp, S. G., Buswell, J. A. & Eriksson, K. E. (1981). “Metabolism of trans-ferulic acid by white-rot fungus Sporotrichum pulverulentum.” Arch Microbiol, 128, 349-354. [21] Falconnier, B., Lapierre, C., Lesage-Meessen, L., Yonnet, G., Brunerie, P., Ceccaldi, B. C., Corrieu, G. & Asther, M. (1994). “Vanillin as a product of ferulic acid biotransformation by the white rot fungus Pycnoporus cinnabarinus I-37: identification of metabolic pathways.” J Biotechnol, 37, 123–132. [22] Rosazza, J. P., Huang, Z., Dostal, L., Volm, T. & Rousseau, B. (1995). “Biocatalytic transformations of ferulic acid: an abundant aromatic natural product.” J Ind Microbiol, 15, 457–471. [23] Peng, X., Misawa, N. & Harayama, S. (2003). “Isolation and characterization of Thermophilic Bacilli degrading cinnamic, 4-coumaric, and ferulic acids.” Appl Environ Microb, 69, 1417–1427. [24] Nazareth, S. & Mavinkurve, S. (1986). “Degradation of ferulic acid via 4vinylguaiacol by Fusarium solani (Mart) Sacc.” Can J Microbiol, 32, 494–497. [25] Karmakar, B., Vohra, R. M., Nandanwar, H., Sharma, P., Gupta, K. G. & Sobti, R. C. (2000). “Rapid degradation of ferulic acid via 4-vinylguaiacol and vanillin by a newly isolated strain of Bacillus coagulans.” J Biotechnol, 80, 195–202. [26] Kadakol, C. & Kamanavalli, C. M. (2010). “Biodegradation of eugenol by Bacillus cereus strain PN24.” J Chem, 7, 474–480. [27] Xie, X. G., Huang, C. Y., Fu, W. Q. & Dai, C. C. (2016). “Potential of endophytic fungus Phomopsis liquidambari for transformation and degradation of recalcitrant pollutant sinapic acid.” Fungal Biol, 120, 402-413. [28] Xie, T., Liu, Z. C., Liu, Q. & Wang, G. G. (2015). “Structural insight into the oxidation of sinapic acid by CotA laccase.” J Struc Biol, 190, 155-161. [29] Jurkovfid, M. & Wurst, M. (1993). “Biodegradation of aromatic carboxylic acids by Pseudomonas mira,” FEMS Microbiol Lett, 111, 245-250.

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

183

[30] Mukherjee, G., Sachan, A., Ghosh, S. & Mitra, A. (2006). “Conversion of sinapic acid to syringic acid by a filamentous fungus Paecilomyces variotii.” J Gen Appl Microbiol, 52, 131-135. [31] Masai, E., Yamamoto, Y., Inoue, T., Takamura, K., Hara, H., Kasai, D., Katayama, Y. & Fukuda, M. (2007). “Characterization of ligV essential for catabolism of vanillin by Sphingomonas paucimobilis SYK-6.”.Biosci Biotechnol Biochem, 71, 2487-2892. [32] Kasai, D., Masai, E., Miyauchi, K., Katayama, Y. & Fukuda, M. (2004). “Characterization of the 3-O-Methylgallate dioxygenase gene and evidence of multiple 3-O-methylgallate catabolic pathways in Sphingomonas paucimobilis SYK-6.” J Bacteriol, 186, 4951–4959. [33] Abe, T., Masai, E., Miyauchi, K., Katayama, Y. & Fukuda, M. (2005). “A Tetrahydrofolate-dependent O-demethylase, LigM, is crucial for catabolism of vanillate and syringate in Sphingomonas paucimobilis SYK-6.” J Bacteriol, 187, 2030–2037. [34] Zhu, D., Zhang, P., Xie, C., Zhang, W., Sun, J., Qian, W. J., et al. (2017). “Bidegradation of alkaline lignin by Bacillus ligniniphilus L1.” Biotechnol Biofuels, 10(1), 44-57. [35] Deangelis, K. M., Sharma, D., Varney, R., Simmons, B., Isern, N. G., Markilllie, L. M., et al. (2013). “Evidence supporting dissimilatory and assimilatory lignin degradation in Enterobacter lignolyticus SCF1.” Front Microbiol, 4(4), 280-293. [36] Michalska, K., Chang, C., Mack, J. C., Zerbs, S., Joachimiak, A. & Collart, F. R. (2012). “Characterization of transport proteins for aromatic compounds derived from lignin: benzoate derivative binding proteins.” J Mol Biol, 423(4), 555-575. [37] Salmon, R. C., Cliff, M. J., Rafferty, J. B. & Kelly, D. J. (2013). “The CouPSTU and TarPQM transporters in Rhodopseudomonas palustris: redundant, promiscuous uptake systems for lignin-derived aromatic substrates.” Plos One, 8(3), e59844. [38] Otani, H., Lee, Y. E., Casabon, I. & Eltis, L. D. (2014). “Characterization of phydroxycinnamate catabolism in a soil Actinobacterium.” J Bacteriol, 196(24), 4293-303. [39] Otani, H., Stogios, P. J., Xu, X., Nocek, B., Li, S. N., Savchenko, A., et al. (2016). “The activity of couR, a marR family transcriptional regulator, is modulated through a novel molecular mechanism.” Nucleic Acids Res, 44(2), 595-607. [40] Parke, D. & Ornston, L. N. (2003). “Hydroxycinnamate (hca) catabolic genes from Acinetobacter sp. strain ADP1 are repressed by HcaR and are induced by hydroxycinnamoyl-coenzyme A thioesters.” Appl Environ Microbiol, 69(9), 53985409. [41] Calisti, C., Ficca, A. G., Barghini, P. & Ruzzi, M. (2008). “Regulation of ferulic catabolic genes in Pseudomonas fluorescens bf13: involvement of a marr family regulator.” Appl Microbiol Biotechnol, 80(3), 475-483.

184

Wenya Wang, Chen Shi and Robert J. Linhardt

[42] Hirakawa, H., Schaefer, A. L., Greenberg, E. P. & Harwood, C. S. (2012). “Anaerobic p-coumarate degradation by Rhodopseudomonas palustris and identification of CouR, a MarR repressor protein that binds p-coumaroyl coenzyme A.” J Bacteriol, 194(8), 1960. [43] Valderrama, J. A., Duranterodríguez, G., Blázquez, B., García, J. L., Carmona, M. & Díaz, E. (2012). “Bacterial degradation of benzoate: cross-regulation between aerobic and anaerobic pathways.” J Biol Chem, 287(13), 10494-10508. [44] Jiménez, J. I., Pérezpantoja, D., Chavarría, M., Díaz, E. & De, L. V. (2014). “A second chromosomal copy of the cata gene endows pseudomonas putida mt-2 with an enzymatic safety valve for excess of catechol.” Environ Microbiol, 16(6), 176778. [45] Hernándezarranz, S., Moreno, R. & Rojo, F. (2013). “The translational repressor Crc controls the Pseudomonas putida benzoate and alkane catabolic pathways using a multi-tier regulation strategy.” Environ Microbiol, 15(1), 227-241. [46] Luu, R. A., Kootstra, J. D., Nesteryuk, V., Brunton, C. N., Parales, J. V., Ditty, J. L., et al. (2015). “Integration of chemotaxis, transport and catabolism in Pseudomonas putida and identification of the aromatic acid chemoreceptor PcaY.” Mol Micicrobiol, 96(1), 134-147. [47] Garcı´a, L. L., Rivas-Marı´n, E., Floriano, B., Bernhardt, R., Ewen, K. M., ReyesRamı´rez, F., Santero, E., et al. (2011). “ThnY is a ferredoxin reductase-like ironsulfur flavoprotein that has evolved to function as a regulator of tetralin biodegradation gene expression.” J Biol Chem, 286(3), 1709-1718. [48] Ohta, Y., Hasegawa, R., Kurosawa, K., Koizumi, T., Maeda, Allyn H., Nishimura, H., et al. (2017). “Enzymatic specific production and chemical functionalization of phenylpropanone platform monomers from lignin.” Chemsuschem, 10(2), 425-433. [49] Picart, P., Liu, H., Grande, P. M., Anders, N., Zhu, L., Klankermayer, J., et al. (2017). “Multi-step biocatalytic depolymerization of lignin.” Appl Microbiol Biotec-hnol, 101(15), 6277-6287. [50] Gall, D. L., Kontur, W. S., Lan, W., Kim, H., Li, Y., Ralph, J., et al. (2018). “In vitro enzymatic depolymerization of lignin with release of syringyl, guaiacyl, and tricin units.” Appl Environ Microbiol, 84(3), AEM.02076-17. [51] Wang, C., Ouyang, X. H., Su, S. S., Liang, X., Zhang, C., Wang, W. Y., Yuan, Q. P. & Li, Q. (2016). “Effect of sulfonated lignin on enzymatic activity of the lignolytic enzymes Cα-dehydrogenase LigD and β-etherase LigF.” Enzyme Microb Technol, 93, 59–69. [52] Fuchs, G., Boll, M. & Heider, J. (2011). “Microbial degradation of aromatic compounds—from one strategy to four.” Nature Rev Microbiol, 9, 803-816. [53] Linger, J. G., Vardon, D. R., Guarnieri, M. T., Karp, E. M., Hunsinger, G. B., Franden, M. A., Johnson, C. W., Chupka, G., Strathmann, T. J., Pienkos, P. T. &

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

[54] [55]

[56]

[57]

[58]

[59]

[60]

[61]

[62]

[63]

[64]

185

Beckham, G. T. (2014). “Lignin valorization through integrated biological funneling and chemical catalysis.” PNAS (USA), 111, 12013-12018. Kaur, B. & Chakraborty, D. (2013). “Biotechnological and molecular approaches for vanillin production: a review.” Appl Biochem Biotechnol, 169, 1353–1372. Davis, K. M., Rover, M., Brown, R. C., Bai, X. L., Wen, Z. Y. & Jarboe, L. R. (2016). “Recovery and utilization of lignin monomers as part of the biorefinery approach.” Energies, 9, 808. Sainsbury, P. D., Hardiman, E. M., Ahmad, M., Otani, H., Seghezzi, N., Eltis, L. D. & Bugg, T. D. H. (2013). “Breaking down lignin to high-value chemicals: the conversion of lignocellulose to vanillin in a gene deletion mutant of Rhodococcus jostii RHA1.” ACS Chem Biol, 8, 2151−2156. Austin, S., Kontur, W. S., Ulbrich, A., Oshlag, J. Z., Zhang, W. P., Higbee, A., et al. (2015). “Metabolism of multiple aromatic compounds in corn stover hydrolysate by Rhodopseudomonas palustris.” Environ Sci Technol, 49(14), 8914-8922. Okamura-Abe, Y., Abe, T., Nishimura, K., Kawata, Y., Sato-Izawa, K., Otsuka, Y., Nakamura, M., Kajita, S., Masai, E., Sonoki, T. & Katayama, Y. (2016). “Betaketoadipic acid and muconolactone production from a lignin-related aromatic compound through the protocatechuate 3,4-metabolic pathway.” J Biosci Bioeng, 121, 652-658. Kondo, S., Sugimura, K., Okamura, Y., Mase, K. & Sato-Izawa, K. (2016). “Stable Chiral Carboxymuconolactone Production from a Lignin-Related Aromatic Compound, Protocatechuic Acid.” Ferment Technol, 5(135), 2-5. Gosling, A., Fowler, S. J., O’Shea, M. S., Straffon, M., Dumsday, G. & Zachariou, M. (2011). “Metabolic production of a novel polymer feedstock, 3-carboxy muconate, from vanillin.” Appl Microbiol Biotechnol, 90, 107–116. van Duuren, J. B., Wijte, D., Karge, B., dos Santos, V. A., Yang, Y., Mars, A. E. & Eggink, G. (2012). “pH-stat fed-batch process to enhance the production of cis, cismuconate from benzoate by Pseudomonas putida KT2440-JD1.” Biotechnol Prog, 28, 85-92. Sonoki, T., Morooka, M., Sakamoto, K., Otsuka, Y., Nakamura, M., Jellison, J. & Goodell, B. (2014). “Enhancement of protocatechuate decarboxylase activity for the effective production of muconate from lignin-related aromatic compounds.” J Biotechnol, 192, 71–77. Han, L., Liu, P., Sun, J., Wu, Y., Zhang, Y., Chen, W., Lin, J., Wang, Q. & Ma, Y. (2015). “Engineering catechol 1,2-dioxygenase by design for improving the performance of the cis, cis-muconic acid synthetic pathway in Escherichia coli.” Sci Rep-UK, 5, 13435. Vardon, D. R., Franden, M. A., Johnson, C. W., Karp, E. M., Guarnieri, M. T., Linger, J. G., Salm, M. J., Strathmann, T. J. & Beckham, G. T. (2015). “Adipic acid production from lignin.” Energy Environ Sci, 8, 617-628.

186

Wenya Wang, Chen Shi and Robert J. Linhardt

[65] Johnson, C. W., Salvachúa, D., Khanna, P., Smith, H., Peterson, D. J. & Beckham, G. T. (2016). “Enhancing muconic acid production from glucose and lignin-derived aromatic compounds via increased protocatechuate decarboxylase activity.” Metab Eng Commun, 3,111–119. [66] Johnson, C. W., Abraham, P. E., Linger, J. G., Khanna, P., Hettich, R. L. & Beckham, G. T. (2017). “Eliminating a global regulator of carbon catabolite repression enhances the conversion of aromatic lignin monomers to muconate in Pseudomonas putidakt 2440.” Metab Eng Commun, 5, 19-25. [67] Barton, N., Horbal, L., Starck, S., Kohlstedt, M., Luzhetskyy, A. & Wittmann, C. (2017). “Enabling the valorization of guaiacol-based lignin: integrated chemical and biochemical production of cis, cis-muconic acid using metabolically engineered amycolatopsis sp ATCC 39116.” Metab Eng, 45, 200-210. [68] Sonoki, T., Takahashi, K., Sugita, H., Hatamura, M., Azuma, Y., Sato, T., et al. (2017). “Glucose-free cis, cis-muconic acid production via new metabolic designs corresponding to the heterogeneity of lignin.” ACS Sustain Chem Eng, 6(1), 12561264. [69] Otsuka, Y., Nakamura, M., Shigehara, K., Sugimura, K., Masai, E., Ohara, S. & Katayama, Y. (2006). “Efficient production of 2-pyrone 4,6-dicarboxylic acid as a novel polymer-based material from protocatechuate by microbial function.” Appl Microbiol Biotechnol, 71, 608–614. [70] Asano, Y., Yamamoto, Y. & Yamada, H. (1994). “Catechol 2,3-dioxygenasecatalyzed synthesis of picolinic acids from catechols.” Biosci Biotechnol Biochem, 58, 2054-2056. [71] Mycroft, Z., Gomis, M., Mines, P., Law, P. & Bugg, T. D. (2015). “Biocatalytic conversion of lignin to aromatic dicarboxylic acids in Rhodococcus jostii RHA1 by re-routing aromatic degradation pathways.” Green Chem, 17, 4974-4979. [72] Vaillancourt, F. H., Bolin, J. T. & Eltis, L. D. (2006). “The ins and outs of ringcleaving dioxygenases.”Crit Rev Biochem Mol Biol, 41(4), 241-267. [73] Beckham, G. T., Johnson, C. W., Karp, E. M., Salvachúa, D. & Vardon, D. R. (2016). “Opportunities and challenges in biological lignin valorization.” Curr Opin Biotech, 42, 40-53. [74] Johnson, C. W. & Beckham, G. T. (2015). “Aromatic catabolic pathway selection for optimal production of pyruvate and lactate from lignin.” Metab Eng, 28, 240247. [75] Hong, C. Y., Ryu, S. H., Jeong, H., Lee, S. S., Kim, M. & Choi, I. G. (2017). “Phanerochaete chrysosporium multi-enzyme catabolic system for in vivo modification of synthetic lignin to succinic acid.” ACS Chemical Biology, 12(7), 1749-1759.

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

187

[76] Eduardo, B. T., Juan Manuel, S. Y., Otoniel, B. D. & Liliana, M. B. (2015). “Production of short-chain fatty acids from the biodegradation of wheat straw lignin by aspergillus fumigatus.” Bioresource Technol, 196(2), 418-425. [77] Kosa, M. & Ragauskas, A. J. (2012). “Bioconversion of lignin model compounds with oleaginous Rhodococci.” Appl Microbiol Biotechnol, 93, 891–900. [78] Kosa, M. & Ragauskas, A. J. (2013). “Lignin to lipid bioconversion by oleaginous Rhodococci.” Green Chem, 15, 2070-2074. [79] He, Y., Li, X., Ben, H., Xue, X. & Yang, B. (2017). “Lipid production from dilute alkali corn stover lignin by Rhodococcus strains.” ACS Sustain Chem Eng, 5(3), 2302-2311. [80] Yaguchi, A., Robinson, A., Mihealsick, E. & Blenner, M. (2017). “Metabolism of aromatics by Trichosporon oleaginosus, while remaining oleaginous.” Microb Cell Fact, 16(1), 206. [81] Zhao, C., Xie, S., Pu, Y., Zhang, R., Huang, F., Ragauskas, A. J. & Yuan, J. S. (2015). “Synergistic enzymatic and microbial lignin conversion.” Green Chem, 18, 1306-1312. [82] Wells, T. J., Wei, Z. & Ragauskas, A. (2015). “Bioconversion of lignocellulosic pretreatment effluent via oleaginous Rhodococcus opacus DSM 1069.”Biomass Bioenerg, 72(4), 200-205. [83] Lin, L., Cheng, Y., Pu, Y., Sun, S., Li, X. & Jin, M., et al. (2016). “Systems biologyguided biodesign of consolidated lignin conversion.” Green Chem, 18(20), 55365547. [84] Shi, Y., Yan, X., Li, Q., Wang, X., Liu, M., Xie, S., et al. (2016). “Directed bioconversion of kraft lignin to polyhydroxyalkanoate by Cupriavidus basilensis, B8 without any pretreatment.” Process Biochem, 52, 238-242. [85] Kumar, M., Singhal, A., Verma, P. K. & Thakur, I. S. (2017). “Production and characterization of Polyhydroxyalkanoate from lignin derivatives by Pandoraea sp. ISTKB.” ACS Omega, 2(12), 9156-9163. [86] Ho, J., Pawar, S. V., Hallam, S. J. & Yadav, V. G. (2018). “An improved whole-cell biosensor for the discovery of lignin-transforming enzymes in functional metagenomic screens.” ACS Synth Biol, 2(7), 392-398. [87] Ni, J., Gao, Y. Y., Tao, F., Liu, H. Y. & Xu, P. (2017). “Temperature‐directed biocatalysis for the sustainable production of aromatic aldehydes or alcohols.” Angew Chem Int Edit, 57(5), 1214-1217. [88] Delorenzo, D. M., Henson, W. R. & Moon, T. S. (2017). “Development of chemical and metabolite sensors for Rhodococcus opacus PD630.” ACS Synth Biol, 6(10), 1973-1978. [89] Nogales, J., García, J. L. & Díaz, E. (2017). “Degradation of Aromatic Compounds in Pseudomonas: A Systems Biology View.” In Aerobic Utilization of Hydrocarbons, Oils and Lipids. Springer International Publishing., 1-49.

188

Wenya Wang, Chen Shi and Robert J. Linhardt

[90] Xie, S. X., Syrenne, R., Sun, S. & Yuan, J. S. (2014). “Exploration of natural biomass utilization systems (NBUS) for advanced biofuel - from systems biology to synthetic design.” Curr Opin Biotech, 27(6), 195-203. [91] Mukhopadhyay, A. (2015). “Tolerance engineering in bacteria for the production of advanced biofuels and chemicals.” Trends Microbiol, 23(8), 498-508. [92] Chen, G. Q. & Jiang, X. R. (2018). “Next generation industrial biotechnology based on extremophilic bacteria.” Curr Opin Biotech, 50, 94-100. [93] Chi, Y., Hatakka, A. & Maijala, P. (2007). “Can co-culturing of two white-rot fungi increase lignin degradation and the production of lignin-degrading enzymes.” Int Biodeter Biodegrad, 59(1), 32-39. [94] Huang, D. L., Zeng, G. M., Feng, C. L., Hu, S., Lai, C., Zhao, M. H., Su, F. F., Tang, L. & Liu, H. L. (2010). “Changes of microbial population structure related to lignin degradation during lignocellulosic waste composting.” Bioresour Technol, 101(11), 4062-4067. [95] Rahimi, A., Ulbrich, A., Coon, J. J. & Stahl, S. S. (2014). “Formic-acid-induced depolymerization of oxidized lignin to aromatics.” Nature, 515, 249-252. [96] Li, S., Masoud, T. A., Ydna, M. Questell-Santiago., Florent, H., Li, Y. D., Kim, H., Richard, M., Clint, C., John, R. & Jeremy, S. L. (2016). “Formaldehyde stabilization facilitates lignin monomer production during biomass depolymerization.” Science, 354, 329-333. [97] Vardon, D. R., Rorrer, N. A., Salvachúa, D., Settle, A. E., Johnson, C. W., Menart, M. J., Cleveland, N. S., Ciesielski, P. N., Steirer, K. X., Dorgan, J. R. & Beckham, G. T. (2016). “cis, cis-Muconic acid: separation and catalysis to bio-adipic acid for nylon-6, 6 polymerization.” Green Chem, 18, 3397-3413. [98] Cao, Z., Engelhardt, J., Dierks, M., Clough, M. T., Wang, G. H., Heracleous, E., et al. (2017). “Catalysis meets nonthermal separation for the production of (Alkyl)phenols and hydrocarbons from Pyrolysis oil.” Angew Chem Int Edit, 129(9), 23342339. [99] Schwartz, T. J., O’Neill, B. J., Shanks, B. H. & Dumesic, J. A. (2014). “Bridging the chemical and biological catalysis gap: challenges and outlooks for producing sustainable chemicals.” ACS Catal, 4, 2060-2069. [100] Huo, Y., Zeng, H. & Zhang, Y. (2016). “Integrating metabolic engineering and heterogeneous chemocatalysis: New opportunities for biomass to chemicals.” ChemSusChem, 9, 1078–1080. [101] Plaggenborg, R., Overhage, J., Loos, A., Archer, J. A. C., Lessard, P., Sinskey, A. J., Steinbuchel, A. & Priefert, H. (2006). “Potential of Rhodococcus strains for biotechnological vanillin production from ferulic acid and eugenol.” Appl Microbiol Biotechnol, 72, 745–755.

The Potential Role of Enzymatic Catalysis and Metabolic Engineering …

189

[102] Barghini, P., Di Gioia, D., Fava, F. & Ruzzi, M. (2007). “Vanillin production using metabolically engineered Escherichia coli under non-growing conditions.” Microb Cell Fact, 6(1), 13-23. [103] Lee, E. G., Yoon, S. H., Das, A., Lee, S. H., Li, C., Kim, J. Y., Choi, M. S., Oh, D, K. & Kim, S. W. (2009). “Directing vanillin production from ferulic acid by increased acetyl-CoA consumption in recombinant Escherichia coli.” Biotechnol Bioeng, 102, 200–208. [104] Tilay, A., Bule, M. & Annapure, U. (2010). “Production of biovanillin by one-step biotransformation using fungus Pycnoporous cinnabarinus.” J Agric Food Chem, 58, 4401–4405.

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 8

STRUCTURAL ELUCIDATION OF LIGNIN MACROMOLECULES FROM P-COUMARATE 3-HYDROXYLASE (C3H) DOWN-REGULATED TRANSGENIC POPLARS (84 K) Tian-Ying Chen1, Zhi-Wen Li1, Han-Min Wang1, Bing Wang1, Jia-Long Wen1,*, Feng-Xia Yue2 and Tong-Qi Yuan1 1

Beijing Key Laboratory of Lignocellulosic Chemistry, Beijing Forestry University, Beijing, China 2 State Key Laboratory of Pulp and Paper Engineering, South China University of Technology, Guangzhou, China

ABSTRACT Elucidating the structural characteristics of lignin from p-coumarate 3-hydroxylase (C3H) down-regulated 84 K poplar is beneficial to the value-added utilization of the transgenic poplar. Herein, the cellulolytic enzyme lignin (CEL) preparations were extracted from the control and C3H down-regulated 84 K poplar. The major lignin linkages (β–O–4, β–5, β–β) were assigned and quantified by NMR techniques. It was found that the CEL from C3H down-regulated poplar has a higher S/G ratio, higher content of β–O–4 linkages, p-hydroxybenzoate (PB), and methoxy groups (OCH3). Interestingly, a small number of p-hydroxyphenyl (H) units were detected in the CEL samples extracted from C3H down-regulated 84 K poplars. In short, these results are essential for evaluating the effects of C3H down-regulation in poplar on the molecular characteristics of lignin macromolecules. Furthermore, the well-characterized structure characteristics of lignin from these poplars will be beneficial to the utilization of these bioresources in the current biorefinery scenario. *

Corresponding Author Email: [email protected]

192

Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang et al.

Keywords: C3H down-regulation, Lignin, 2D-HSQC NMR, S/G ratio, p-hydroxybenzoate

INTRODUCTION Lignin is an abundant aromatic polymer in lignocellulosic biomass, which attracts significant attentions due to its potential in converting into fuels, chemicals, and materials [1]. In nature, lignin is formed by oxidative coupling of three hydroxycinnamyl alcohols according to degree of methoxylation: p-coumaryl, coniferyl, and sinapyl alcohols via different ether and carbon-carbon linkages (β–O–4, β–5, β–β, etc.) or, characteristic endgroups (e.g., cinnamaldehyde units) [2]. Recently, Yue et al. has identified the 4–O–5 linkages in the lignin of softwood via NMR techniques, which proposed the new concept that lignin is rather “U type” linear structure than “highly branched” in conventional theory [3]. Meanwhile, lignin and carbohydrates (mainly hemicelluloses) are also linked by different chemical linkages (ether and ester bonds), forming three kinds of lignincarbohydrate complexes (LCC), such as phenyl glycoside, benzyl ether, and γ-ester [4]. These intricate connectivities pose a challenge for the deconstruction and value-added applications of lignin according to current cognitive level [5]. Although many pretreatment methods have been proposed to reduce “biomass recalcitrances” by removing lignin, pretreatment methods aiming at lignin removal are usually expensive and most pretreatments have negative effects on environments [6, 7]. Genetic engineering of biomass is a promising approach to reduce biomass recalcitrance during biofuels production. Genetic modification of specific enzymes related to lignin biosynthesis will change the chemical composition and structural characteristics of lignin in plants [8]. In general, p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol, are polymerized into lignin polymers during the process of lignification. The phenylpropanoid pathway starts with phenylalanine (Phe), buttyrosine (Tyr) may also be consumed in monocots, and then Phe was converted into precursor units of “H,” “G” and “S” units via phenylalanine ammonia-lyase (PAL) [9], cinnamate 4-hydroxylase (C4H) [10], tyrosineammonia-lyase (TAL) [11], 4-coumarate: CoA ligase (4CL) [1], p-coumarate 3-hydroxylase (C3H) [12], shikimate/quinatehydroxycinnamoyl transferase (HCT) [13], cinnamoylCoA reductase (CCR) [14], cinnamyl alcohol dehydrogenase (CAD) [15], caffeoyl shikimate esterase (CSE) [16], caffeic acid O-methyltransferase (COMT) [17], caffeoyl-CoAO-methyltransferase (CCoAOMT) [18], ferulate 5-hydroxylase (F5H) [19] and hydroxycinnamaldehyde dehydrogenase (HCALDH) [1]. During the biosynthesis process of lignin macromolecule, the up- and down-regulation of an array of enzymes could have an impact on the lignin structure and subsequent processability of the biomass. Understanding the chemical structures of lignin in the transgenic feedstock is beneficial to select optimal lignin characteristics that required for various applications of lignocellulosic materials. Chen et al. reported that seven genes-encoding enzymes in the

Structural Elucidation of Lignin Macromolecules …

193

monolignol pathway were independently down-regulated in alfalfa (Medicago sativa) using anti-sense and/or RNA interference technique [20]. They found that the downregulation of earlier enzymes rather than later enzymes in the pathway resulted in significant decrease in lignin content, irrespective of the degree of enzyme activity reduction. Subsequently, they analyzed the down-regulated alfalfa lines and demonstrated that the down-regulated plants have high yields of fermentable sugar without acid pretreatment as compared to that of wild-type (WT) plants [21]. These results suggested that genetical modification can overcome the recalcitrance of lignin and facilitate biofuel production. The early enzyme (e.g., C3H) in the pathway of lignin biosynthesis can provide entry to the synthesis of guaiacyl (G) and syringyl (S) lignin precursors from the nonmethoxylated p-hydroxyphenyl (H) branch of the pathway [22]. Ralph and coauthors [12] have demonstrated the effects of C3H down-regulation on structures of alfalfa lignin. It was reported that the proportion of p-hydroxyphenyl (H) units was strikingly increased after down-regulation of the C3H enzymes. Meanwhile, the authors have investigated the effects of C3H down-regulation on the structures of lignin in poplar by analyzing the acetylated CEL [23]. However, acetylation of CEL sample will lead to the signal loss of some substructures in the 2D-HSQC spectra, such as α, β-diaryl ethers and LCC linkages [24]. In the present study, CEL samples were extracted from control 84 K and C3H downregulated poplar with different C3H gene activities to illustrate the effects of C3H downregulation on the composition and structure of lignin macromolecules. The structural elucidation of non-acetylated CEL sample was achieved by the-state-of-art analytical techniques, such as 2D-HSQC NMR, 31P NMR and gel permeation chromatography (GPC) techniques. The results obtained from this study will provide theoretical basics for verification of lignin biosynthesis pathway in C3H down-regulated poplar sample and utilization of the genetically modified poplar wood in the current biorefinery process.

MATERIAL AND METHODS Plant Materials The transgenic 84 K poplar (Populusalba×P. glandulosa, 4 years) plants, which were C3H down-regulated poplars with different enzyme activity level (323, level 1, residual C3H gene activity ~18%; 325, level 2, residual C3H gene activity ~12%), were obtained as reported previously [25]. For the control experiment, the poplar (CK) was used as feedstock. Prior to using, all wood samples were debarked, dried, and smashed to pass a 20–40 mesh sieve. The extractives were removed after the treatment with ethanol/benzene (1:2, v/v) until the siphoned liquid colorless. The extract-free poplar powders were air-

194

Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang et al.

dried at 60oC for 16 h and stored at 5oC before use. The chemical composition of CK and genetically modified 84 K poplars (named 323 and 325) was determined by hydrolyzing the extractives-free poplar wood with dilute sulfuric acid according to the NREL standard analytical procedure [26]. The Klason lignins in CK, 323 and 325 poplar woods were determined to be 19.4%, 20.5% and 20.7%, respectively. Chemical composition of the poplar is shown in Table 1. All the chemicals used in this study were analytical grade and purchased from Sigma Chemical Co. (Beijing, China). Commercial cellulase ([email protected] CTec2, 100 FPU/mL) is provided from Novozymes (Beijing, China).

Preparation of CELs CEL preparation is one of the well-established methods to isolate lignin for structural elucidation of native lignin in lignocellulosic biomass [27]. Firstly, the dewaxed poplar wood was milled in a planetary ball-mill (Fritsch GmbH, Idar-Oberstein, Germany) at a fixed frequency of 450 rpm for 5 hunder N2 (room temperature) as previously [28]. The ball-milled materials were suspended in acetate buffer (0.05 M, pH4.8) with a loading of Cellic® CTec2 (50 FPU/g), and ensuring the concentration of substrate was 5%. The reaction mixtures were incubated at 50 oC in a rotary shaker (150 rpm) for 48 h. After enzymatic hydrolysis, the mixtures were centrifuged to remove the supernatant, and the residues were sequentially washed with acetate buffer and deionized water. The residues were freeze-dried and then extracted with 96% aqueous dioxane (v/v) with a solid toliquid ratio of 1:40 (g/mL) at room temperature for 24 h to extract lignin. The extracting liquid, which were concentrated and precipitated into 10 volumes of acid water (pH 2.0), then the precipitated lignin samples were freeze-dried to obtain crude CEL samples. The crude CEL samples were further dissolved in 90% acetic acid, and then regenerated into acidic water (pH = 2.0), after washing and freeze-drying, the purified CEL samples were obtained (labeled as CK-CEL, 323-CEL and 325-CEL, respectively).

Structural Elucidation of CELs The weight average (Mw) and number-average (Mn) molecular weights of the lignin preparations were determined by gel permeation chromatography (GPC, Agilent 1200, USA) according to a previous publication [29, 30]. In general, acetylation process is necessary for measuring molecular weights of the native lignin samples. Detailed, 50 mg of lignin sample was dissolved in a 3 mL solution of DMSO: N-Methylimidazole (2: 1, v/v). The dissolution was performed on a shaker at room temperature for 24 h in the dark. After that, 1 mL of acetic anhydride was added into the solution for an additional 1.5 h. At the end of the designated time, the solution was dropped slowly into 300 mL of acidic water

Structural Elucidation of Lignin Macromolecules …

195

(pH = 2.0, adjusted by HCl) to induce precipitation, and the precipitate was washed thoroughly with acidic water (pH = 2.0) until significant reducing of pungent odor, and then freeze-dried to obtain the acetylated lignin. 2D-HSQC spectra of lignin were aquired according to a previous publication [30]. In detail, 50 mg of lignin was dissolved in 0.5 mL of DMSO-d6, and the sequence “hsqcetgp” was selected from program library. The spectral widths were 5000 Hz and 20000 Hz for the 1H- and 13C-dimensions, respectively. The number of collected complex points was 1024 for 1H-dimension with a recycle delay of 1.5 s. The number of transients was 64, and 256 time increments wererecorded in the 13C-dimension.Prior to Fourier transformation, the data matrixes were zero filled up to 1024 points in the 13C-dimension. Data processing was performed using standard Bruker Topspin-2.1 software. Quantitative 31P NMR spectra of CELs were recorded on a Bruker 400 MHz spectrophotometer with 2048 scans, and 20 mg lignin was phosphitylated using 2-chloro-4,4,5,5-tetramethyl-1,3,2-dioxaphospholane (TMDP). A known amount of dry lignin (20 mg) was dissolved in 500 μL anhydrous pyridine and deuterated chloroform (1.6:1, v/v) under stirring. This was followed by the addition of 100 μL cyclohexanol (10.85 mg/mL) as an internal standard (IS), and 100 μL chromium (III) acetylacetonate solution (5 mg/mL in anhydrous pyridine and deuterated chloroform 1.6:1, v/v) as relaxation reagent. Finally, the mixture was treated with 100 μL phosphorylating reagent (2-chloro-1,3,2-dioxaphospholane) and was transferred into a 5 mm NMR tube for analysis according to the previous literature [31, 32]. All experiments in this study were performed in duplicate, and the data reported were the average values.

RESULT AND DISCUSSION For the purpose of biomass valorization, the “recalcitrance” of biomass can be reduced by modifying the phenylpropanoid biosynthesis via the enzymes in the pathway. Among these enzymes, coumarate 3-hydroxylase (C3H) is a crucial hydroxylase for the synthesis of the precursor of G and S units, which can be restrained via the up- and down-regulation of C3H. Some studies demonstrated that the C3H down-regulation can markedly increase the proportion of H unit relative to the normally dominant G and S units [12, 23]. Further study showed that C3H down-regulation facilitated the subsequent lignin extraction process [25]. However, the effects of C3H down-regulation in different levels on the structure of lignin are unknown yet. Therefore, two different C3H down-regulated poplar samples were used to indentify the effects of the C3H down-regulation on lignin structures.

The Fate of Lignin before and after C3H Down-Regulation Table 1 shows the chemical composition of the poplar samples with different degrees of C3H down-regulation. It was observed that the content of Klason lignin in CK poplar

196

Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang et al.

wood was 19.4%, while the content of lignin was slightly increased to 20.5% in 323 and 20.7% in 325 poplar sample, respectively, which is slightly inconsistent with the previous reports [12, 22]. The reason is that four-years growing in the field probably brought about the restored growth of transgenic poplar [25]. By contrast, the content of cellulose and hemicelluloses in the C3H down-regulated poplar woods slightly decreased as compared to that of the wild poplar (CK). In short, composition analysis indicated that downregulation of C3H resulted in the slight changes of chemical composition of the poplar wood. In general, changes of chemical composition will affect the subsequent processability of the lignocellulosic biomass. In this study, CEL was prepared to delineate the effects of various degrees of C3H down-regulation on the lignin structure of poplar wood. The yields of CK-CEL, 323-CEL and 325-CEL were 19.0, 21.5 and 23.0%, respectively. The improved yield of C3H-CEL was probably related to the reduced biomass recalcitrance and enhanced digestibility of the transgenic poplars. Table 1. Composition analysis of control and the poplar samples with various degrees of C3H down-regulation Sample CK 323 325 a

Yield (%)a 19.0 21.5 23.0

KLb 19.4 20.5 20.7

Cellulose 43.52 41.65 41.57

Hemicelluloses 22.56 21.38 21.36

The yield of lignin obtained by CEL preparation based on Klason lignin; KL, Klason lignin.

b

2D-HSQC NMR Analysis 2D-HSQC NMR spectra demonstrated the structural differences of lignin between wild-type 84 K poplar (CK) and various degrees of C3H down-regulated poplar samples. These differences could provide some fundamental basis for obtaining ideal lignin sources for subsequent lignin valorization. Figure 1 identified the inter-unit linkages (side-chain region) and chemical composition (aromatic region) in the 2D-HSQC spectra of the CKCEL, 323-CEL, and 325-CEL according to the previous signal assignments [24, 33-35]. In short, the identified linkages and basic compositions in the lignin macromolecules were depicted in Figure 2. In the side-chain regions (δC/δH 50-90/2.7-6.0) of the 2D-HSQC spectra (Figure 1, top) of CELs, the linkages, such as β–O–4 aryl ethers (A), resinols (B), and phenylcoumarans (C), could be obviously observed. It was found that all the lignin samples exhibited similar spectral patterns. Cross-signals of methoxyl groups (OCH3, δC/δH 55.6/3.72) and β–O–4 (A) were the prominent signals. The Cα–Hα correlations in the β–O–4 linkages were

Structural Elucidation of Lignin Macromolecules …

197

observed at δC/δH 71.8/4.86, while the Cβ–Hβ correlations corresponding to the β–O–4 linkages linked to G and S units can be distinguished at δC/δH 83.6/4.33 and 85.8/4.11, respectively.

Figure 1. 2D-HSQC spectra of the lignin fractions isolated from poplar.

The Cγ–Hγ correlations in the β–O–4 substructures were detected at δC/δH 59.4/3.70 and 3.39. Meanwhile, the content of β–O–4 linkages in C3H-CEL was slightly higher than that of CK-CEL, which is consistent with the results in a previous publication [25]. In addition, the Cγ-Hγ correlations in γ-acylated lignin units (A′) were also observed at δC/δH 63.2/4.38. This indicated that those lignin samples were partially acylated at the γ-carbon of side chains in β–O–4 aryl ether linkages [23]. Resinols (β–β, substructures B) can be easily identified in the spectra in conspicuous amounts as indicated by their Cα–Hα, Cβ–Hβ, and the double Cγ–Hγ correlations at δC/δH 84.9/4.66, 53.5/3.06, 71.0/3.81 and 4.18, respectively. The weak signal (δC/δH, 86.7/5.48) assigned to the Cα–Hα correlations of phenylcoumarans (β–5) suggested that the decreased content of β–5 linkages (from 0.90 to 0.28/100Ar). This phenomenon could be attributed to the reduction of G units (relative to per 100Ar) as compared to that of CK based on a publication [23], in which it was reported that phenylcoumarans are derived from coupling of a monolignol with G units.

198

Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang et al.

In the aromatic regions (δC/δH 100-135/6.0-8.0) of the spectra (Figure 1 bottom), the chemical composition in the lignin samples (CK-CEL, 323-CEL, and 325-CEL) can be clearly distinguished. The S-type lignin units showed a prominent signal for the C2,6–H2,6 correlation at δC/δH 103.9/6.69, whereas the signal for the Cα-oxidized S-units (S′) was observed at δC/δH 106.1/7.19. Additionally, the G-type lignin units showed different correlations for C2–H2 (δC/δH 110.8/6.98), C5–H5 (δC/δH 114.9/6.76), and C6–H6 (δC/δH 118.9/6.79). Meanwhile, there was a relative depletion in the levels of G units derived from methoxylated lignin monomers (coniferyl alcohols) in C3H down-regulated lignin, ranging from 26.77/100Ar in CK-CEL to 24.66/100Ar in 325-CEL. However, although minor H2,6 signals were detected at δC/δH 127.6/7.16, the obvious increase from 0.01 to 0.10/100Ar suggested that a striking elevation of phydroxyphenyl (H) units from the non-methoxylated monolignol, p-coumaryl alcohol. The quantification of 2D-HSQC spectra revealed that relative elevation of H units is at the expense of G units rather than S units, which is in agreement with the conclusion proposed by Ralph’s group [23]. Changes of S/G ratio from 2D-HSQC spectra can intuitively reflect the compositional change of lignin samples. Herein, the S/G ratio in the CK-CEL was 2.75, while S/G ratio for the C3H down-regulated transgenic lines was 2.83 and 3.05. In addition, an extremely important feature of poplar is p-hydroxybenzoate substructure (PB), which was observed as a strong signal at δC/δH 131.2/7.65 (C2,6–H2,6). Actually, in poplar wood, PB is the exclusively acylated the γ-position of lignin side chains [36]. Meanwhile, Ralph’s group also found that PB is almost exclusively linked with syringyl units in the natural poplar lignin [23]. Although the distribution of lignin precursor was changed (Reducing coniferyl alcohol and increasing p-coumaryl alcohol) induced by the C3H down-regulation, the effect of genetic modification on PB was not readily predictable. In this study, the spectra displayed in Figure 1 clearly showed the enhancement of PB content in the C3H-CELs. Precisely, the integral value of PB increased from 11.32 to 15.80/100Ar, as shown in Table 2. This suggested that the H units are more compatible with PB than G or S units, which is agreement with the conjecture proposed by John Ralph [23]. Table 2. Quantification of the lignin sampleswith various degrees of C3H down-regulation

a

H2,6

β–O–4

β–5

β–β

26.67

0.01

56.26 b

0.90 b

11.00 b

73.87

26.11

0.02

57.02

0.36

9.13

75.24

24.66

0.10

57.97

0.28

10.07

S/Ga

PB

CK-CEL

2.75

11.32b

73.32

323-CEL

2.83

15.80

325-CEL

3.05

14.12

Sample

S2,6

G2

S/G ratio obtained by the this equation: S/G ratio = 0.5I (S2,6)/I (G2). Results expressed per 100 Ar based on quantitative 2D-HSQC spectra and corresponding formula is listed in a previous publication [24].

b

Structural Elucidation of Lignin Macromolecules …

199

Figure 2. The identified linkages and aromatic units in the CELs from these poplar samples.

13C

NMR Spectra Analysis

The quantitative 13C NMR spectra of the CELs obtained from control and transgenic poplar wood are shown in Figure 3, which clearly showed the structural changes of CEL after the C3H down-regulation. The detailed signal assignments of lignin were achieved and marked in Figure 3 according to the previous publications [25, 28]. In detail, the obvious signals in the 13C-NMR spectra were detailedly assigned, such as, signal 1 (165.6 ppm, C-7 in PB units), signal 2 (162.1 ppm, C-4 in PB units), signal 3 (152.3 ppm, etherified S3,5 units), signal 4 (149.3 ppm, etherified G3), signal 5 (147.6 ppm, nonetherified S3,5 units, etherified G4 units), signal 6 (145.5 ppm, non-etherified G4), signal 7 (138.2 ppm, etherified S4),signal 8 (134.9 ppm, etherified S1 or G1), signal 9 (131.6 ppm, PB2,6), signal 10 (120.4 ppm, G6),signal 11 (115.4 ppm, G5), signal 12 (111.5 ppm, G2), signal 13 (104.4 ppm, S2,6), signal 14 (86.2 ppm, C-α in β–5 units), signal 15 (85.3 ppm, C-β in β–O–4 units), signal 16 (72.4 ppm, C-α in β–O–4 linkages, signal 17 (59.8 ppm, C-γ in β–O–4 units), signal 18 (56.0 ppm, OCH3). The quantitative results showed that the content of β–O–4 linkages increased from 0.62/Ar in CK-CEL to 0.63/Ar in 323CEL and 0.64/Ar in 325-CEL, which is accordance with the result of the corresponding 2D-HSQC NMR spectra. In addition, the PB was observed at 165.6 ppm and the corresponding signal intensity was also enhanced in the CELs from C3H down-regulated poplars. The quantitative data also demonstrated a slight elevation of PB content from 0.05/Ar in CK-CEL to 0.07/Ar in C3H-CEL. Moreover, the content of methoxyl (OCH3) was calculated to be increased from 2.10/Ar in CK to 2.19–2.20/Ar in C3H down-regulated poplar samples. Similarly, the content of methoxyl in the lignin was also in line with the corresponding S/G ratio calculated from 2D-HSQC spectra of the CELs.

200

Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang et al.

Figure 3. Quantitative 13C NMR spectra of CELs.

Structural Elucidation of Lignin Macromolecules … 31P-NMR

201

Analysis

To further evaluate the effects of C3H down-regulation on the functional group of lignin macromolecule, quantitative 31P-NMR spectra of CK-CEL, 323-CEL and 325-CEL (Figure 4) were performed. The aliphatic OH groups, phenolic OH groups, and COOH groups were determined by phosphatization of the different OH groups in lignin with 2chloro-4,4,5,5-tetramethyl-1,3,2-dioxaphospholane [37]. The various OH groups in the CELs were quantified using cyclohexanol as an internal standard, and the results are listed in Table 3. It was found that the CK-CEL contained more aliphatic OH groups than C3HCEL, suggesting that aliphatic OH groups in C3H-CEL was occupied by PB, which was in line with the content of PB from 2D-HSQC spectra. For phenolic OH groups, it was observed that the contents of syringyl and guaiacyl OH groups in CK-CEL were higher than those of C3H-CEL sample. Considering the difference in the content of β–O–4 linkages in the CELs, it can be speculated that the decrease of syringyl and guaiacyl type phenolic hydroxyl groups in C3H-CEL was related to the β–O–4 linkages.

Figure 4. 31P NMR spectra of the CK-CEL and C3H-CELs.

Table 3. Quantification of the functional groups by quantitative 31P NMR (mmol/g) Sample Aliphatic-OH S-OH C-G-OHa NC-G-OHb NC-H-OH COOH a

CK-CEL 4.78 0.33 0.03 0.31 0.27 0.03

323-CEL 4.60 0.27 0.01 0.26 0.31 0.03

325-CEL 4.69 0.30 0.02 0.29 0.35 0.02

C, condensed; NC, non-condensed.

b

In general, a high content of β–O–4 linkage in lignin means less phenolic hydroxyl in the lignin. However, the contents of H-type phenolic OH were increased from 0.27 to 0.31 and 0.35 mmol/g, corresponding to CK-CEL, 323-CEL and 325-CEL, respectively. The

202

Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang et al.

reason for this was ascribed to the elevation of phydroxyphenyl (H) units revealed by the aforementioned 2D-HSQC spectra analysis. Moreover, it is worth noting that the phenolic OH in H-type lignin structures (PB and H unit) mainly belonged to p-hydroxybenzoate (PB) substructures specifically linked to the lignin fractions. Furthermore, it was observed that the content of COOH groups in these lignin samples was similar.

Molecular Weight Analysis The average molecular weights (Mw and Mn) and polydispersity index (Mw/Mn) of CKCEL, 323-CEL and 325-CEL calculated from the GPC curves (relative values related to polystyrene) are displayed in Figure 5. The weight-average (Mw) molecular weight of CKCEL, 323-CEL and 325-CEL was 12990 g/mol, 12320 g/mol and 13170 g/mol, respectively. Although there are no significant differences in the molecular weight, C3HCELs (323-CEL and 325-CEL) had relatively narrow molecular weight distributions (2.63 for 323-CEL and 2.65 for 325-CEL) as compared to that of CK-CEL (2.75), suggesting that down-regulation of C3H genes facilitates the formation of homogeneous lignin fractions.

Figure 5. Weight-average (Mw), number-average molecular weights (Mn), and polydispersity (Mw/Mn) of the CELs.

CONCLUSION In this study, the CEL preparations were extracted from control and C3H downregulated 84 K poplar woods in relatively high yields, which can better characterize the structural variation of lignin from different feedstocks. Results showed that these CEL

Structural Elucidation of Lignin Macromolecules …

203

samples exhibited similar structural features although CEL samples from transgenic poplars had slightly higher contents of β–O–4 linkages and S/G ratios. Meanwhile, it was found that the CEL from transgenic plants had low content of β–5 linkages. Moreover, minor H unit was observed in lignin from C3H poplar with low residual C3H gene activity (12%). Furthermore, p-hydroxybenzoate (PB) content was increased in the lignin from C3H down-regulated poplar wood. In short, understanding the lignin structures of transgenic feedstock is beneficial for selecting optimal lignin characteristics required for downstream deconstruction and applications of lignocellulosic materials in a biorefinery scenario.

ACKNOWLEDGMENTS This work was supported by National Natural Science Foundation of China [31430092 and 31872698], Fundamental Research Funds for the Central Universities [2017JC13], Open Foundation of Guangxi Key Laboratory of Clean Pulp & Papermaking and Pollution Control [KF201714], Open Foundation of the State Key Laboratory of Pulp and Paper Engineering [201749 and 201836].

REFERENCES [1]

[2]

[3]

[4]

[5]

Rinaldi, R., Jastrzebski, R., Clough, M. T., Ralph, J., Kennema, M., Bruijnincx, P. C., Weckhuysen, B. M. 2016. “Paving the Way for Lignin Valorisation: Recent Advances in Bioengineering, Biorefining and Catalysis.” Angewandte ChemieInternational Edition 55, 8164-215. Ralph, J., Lundquist, K., Brunow, G., Lu, F., Kim, H., Schatz, F. P., Marita1, J. M., Hatfield, R. D., Ralph, S. A., Christensen, J. H., and Boerjan, W. 2004. “Lignins: Natural polymers from oxidative coupling of 4-hydroxyphenylpropanoids.” Phytochemistry Reviews 3, 29–60. Yue, F., Lu, F., Ralph, S. A., Ralph, J. 2016. “Identification of 4-O-5-units in Softwood Lignins via Definitive Lignin Models and NMR.” Biomacromolecules 17, 1909. Balakshin, M., Capanema, E., Gracz, H., Chang, H. M., Jameel, H. 2011. “Quantification of lignin–carbohydrate linkages with high-resolution NMR spectroscopy.” Planta 233, 1097-110. Ding, S. Y., Liu, Y. S., Zeng, Y., Himmel, M. E., Baker, J. O., Bayer, E. A. 2012. “How does plant cell wall nanoscale architecture correlate with enzymatic digestibility?” Science 338, 1055-60.

204 [6]

[7]

[8]

[9]

[10]

[11]

[12]

[13]

[14]

[15]

[16]

Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang et al. Zhao, X. B., Zhang, L. H., Liu, D. H. 2012. “Biomass recalcitrance. Part II: Fundamentals of different pre-treatments to increase the enzymatic digestibility of lignocellulose.” Biofuels Bioproducts Biorefining-Biofpr 6, 561-79. Vanholme, B., Cesarino, I., Goeminne, G., Kim, H., Marroni, F., Van, A. R., Vanholme, R., Morreel, K., Ivens, B., Pinosio, S. 2013. “Breeding with rare defective alleles (BRDA): a natural Populus nigra HCT mutant with modified lignin as a case study.” New Phytologist 198, 765-76. Vanholme, R., Morreel, K., Darrah, C., Oyarce, P., Grabber, J. H., Ralph, J., Boerjan, W. 2012. “Metabolic engineering of novel lignin in biomass crops.” New Phytologist 196, 978-1000. Jansen, F., Gillessen, B., Mueller, F., Commandeur, U., Fischer, R., Kreuzaler, F. 2014. “Metabolic engineering for p-coumaryl alcohol production in Escherichia coli by introducing an artificial phenylpropanoid pathway.” Biotechnology & Applied Biochemistry 61, 646-54. Millar, D. J., Long, M., Donovan, G., Fraser, P. D., Boudet, A. M., Danoun, S., Bramley, P. M., Bolwell, G. P. 2007. “Introduction of sense constructs of cinnamate 4-hydroxylase (CYP73A24) in transgenic tomato plants shows opposite effects on flux into stem lignin and fruit flavonoids.” Phytochemistry 68, 1497-509. Higuchi, T., Ito, Y., Kawamura, I. 1967. “p -hydroxyphenylpropane component of grass lignin and role of tyrosine-ammonia lyase in its formation.” Phytochemistry 6, 875-81. Ralph, J., Akiyama, T., Coleman, H. D., Mansfield, S. D. 2006. “Effects on Lignin Structure of Coumarate 3-Hydroxylase Downregulation in Poplar.” Journal of Biological Chemistry 281, 8843-53. Vanholme, B., Desmet, T., Ronsse, F., Rabaey, K., Breusegem, F. V., Mey, M. D., Soetaert, W., Boerjan, W. 2013. “Towards a carbon-negative sustainable bio-based economy.” Frontiers in Plant Science 4, 174. Van, A. R., Leplé, J. C., Aerts, D., Storme, V., Goeminne, G., Ivens, B., Légée, F., Lapierre, C., Piens, K., Van Montagu, M. C. 2014. “From the Cover: Improved saccharification and ethanol yield from field-grown transgenic poplar deficient in cinnamoyl-CoA reductase.” Proceedings of the National Academy of Sciences of the United States of America 111, 845-50. Fornalé, S., Capellades, M., Encina, A., Wang, K., Irar, S., Lapierre, C., Ruel, K., Joseleau, J. P., Berenguer, J., Puigdomènech, P. 2012. “Altered lignin biosynthesis improves cellulosic bioethanol production in transgenic maize plants down-regulated for cinnamyl alcohol dehydrogenase.” Molecular Plant 5, 817-30. Vanholme, R., Cesarino, I., Rataj, K., Xiao, Y., Sundin, L., Geert, G., Kim, H., Cross, J., Morreel, K., Araujo, P. 2013. “Caffeoyl shikimate esterase (CSE) is an enzyme in the lignin biosynthetic pathway in Arabidopsis.” Science 341, 1103-06.

Structural Elucidation of Lignin Macromolecules …

205

[17] Sattler, S. E., Palmer, N. A., Saballos, A., Greene, A. M., Xin, Z., Sarath, G., Vermerris, W., Pedersen, J. F. 2012. “Identification and Characterization of Four Missense Mutations in Brown midrib 12 (Bmr12 ), the Caffeic O -Methyltranferase (COMT) of Sorghum.” BioEnergy Research 5, 855-65. [18] Wagner, A., Tobimatsu, Y., Phillips, L., Flint, H., Torr, K., Donaldson, L., Pears, L., Ralph, J. 2011. “CCoAOMT suppression modifies lignin composition in Pinus radiata.” Plant Journal for Cell & Molecular Biology 67, 119-29. [19] Weng, J. K., Mo, H., Chapple, C. 2010. “Over‐expression of F5H in COMT‐ deficient Arabidopsis leads to enrichment of an unusual lignin and disruption of pollen wall formation.” Plant Journal 64, 898-911. [20] Chen, F., Srinivasa Reddy, M. S., Temple, S., Jackson, L., Shadle, G., Dixon, R. A. 2006. “Multi-site genetic modulation of monolignol biosynthesis suggests new routes for formation of syringyl lignin and wall-bound ferulic acid in alfalfa (Medicago sativa L.).” Plant Journal 48, 113-24. [21] Chen, F., Dixon, R. A. 2007. “Lignin modification improves fermentable sugar yields for biofuel production.” Nat Biotechnol 25, 759-61. [22] Pu, Y., Chen, F., Ziebell, A., Davison, B. H., Ragauskas, A. J. 2009. “NMR Characterization of C3H and HCT Down-Regulated Alfalfa Lignin.” BioEnergy Research 2, 198-208. [23] Ralph, J., Akiyama, T., Coleman, H. D., Mansfield, S. D. 2012. “Effects on Lignin Structure of Coumarate 3-Hydroxylase Downregulation in Poplar.” BioEnergy Research 5, 1009-19. [24] Wen, J. L., Sun, S. L., Xue, B. L., Sun, R. C. 2013. “Recent Advances in Characterization of Lignin Polymer by Solution-State Nuclear Magnetic Resonance (NMR) Methodology.” Materials 6, 359-91. [25] Peng, X. P., Wang, B., Wen, J. L., Yang, S. Z., Lu, M. Z., Sun, R. C. 2016. “Effects of Genetic Manipulation (Hct And C3h Down-Regulation) on Molecular Characteristics of Lignin and Its Bioconversion to Fermentable Sugars.” Cellulose Chemistry & Technology 50, 649-58. [26] Sluiter, A., Hames, B., Hyman, D., Payne, C., Ruiz, R., Scarlata, C., Sluiter, J., Templeton, D., Wolfe, J. 2008. “Determination of Total Solids in Biomass and Total Dissolved Solids in Liquid Process Samples.” Laboratory Analytical Procedure. [27] Chang, H., Cowling, E. B., Brown, W. 1975. “Comparative Studies on Cellulolytic Enzyme Lignin and Milled Wood Lignin of Sweetgum and Spruce.” Holzforschung 29, 153-59. [28] Wen, J. L., Xue, B. L., Xu, F., Sun, R. C., Pinkert, A. 2013. “Unmasking the structural features and property of lignin from bamboo.” Industrial Crops & Products 42, 33243.

206

Tian-Ying Chen, Zhi-Wen Li, Han-Min Wang et al.

[29] Chen, T. Y., Wen, J. L., Wang, B., Wang, H. M., Liu, C. F., Sun, R. C. 2017. “Assessment of integrated process based on autohydrolysis and robust delignification process for enzymatic saccharification of bamboo.” Bioresour Technol 244, 717-25. [30] Wen, J. L., Sun, S. L., Xue, B. L., Sun, R. C. 2013. “Quantitative structures and thermal properties of birch lignins after ionic liquid pretreatment.” Journal of Agricultural & Food Chemistry 61, 635-45. [31] Argyropoulos, D. S. 1995. “31P NMR in wood chemistry: A review of recent progress.” Research on Chemical Intermediates 21, 373-95. [32] Wen, J. L., Sun, S. L., Yuan, T. Q., Sun, R. C. 2015. “Structural elucidation of whole lignin from Eucalyptus based on preswelling and enzymatic hydrolysis.” Green Chemistry 17, 1589-96. [33] Yuan, T. Q., Sun, S. N., Xu, F., Sun, R. C. 2011. “Structural Characterization of Lignin from Triploid of Populus tomentosa Carr.” Journal of Agricultural & Food Chemistry 59, 6605-15. [34] Chen, T. Y., Wang, B., Wu, Y. Y., Wen, J. L., Liu, C. F., Yuan, T. Q., Sun, R. C. 2017. “Structural variations of lignin macromolecule from different growth years of Triploid of Populus tomentosa Carr.” Int J Biol Macromol 101, 747-57. [35] Chen, T. Y., Wang, B., Shen, X. J., Li, H. Y., Wu, Y. Y., Wen, J. L., Liu, Q. Y., Sun, R. C. 2017. “Assessment of structural characteristics of regenerated cellulolytic enzyme lignin based on a mild DMSO/ [Emim]OAc dissolution system from triploid of Populus tomentosa Carr.” RSC Advances 7, 3376-87. [36] Ralph, J., Lu, F. 1998. “The DFRC Method for Lignin Analysis. 6. A Simple Modification for Identifying Natural Acetates on Lignins.” Journal of Agricultural & Food Chemistry 46, 4616-19. [37] Jääskeläinen, A. S., Sun, Y., Argyropoulos, D. S., Tamminen, T., Hortling, B. 2003. “The effect of isolation method on the chemical structure of residual lignin.” Wood Science Technology 37, 91-102.

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 9

MOLECULAR DESIGN AND CONTROLLABLE SELF-ASSEMBLY OF LIGNIN HOLLOW NANOSPHERES Yanming Han1, Fuquan Xiong1,2, Gaiyun Li1 and Fuxiang Chu1* 1

Research Institute of Wood Industry, Chinese Academy of Forestry, Beijing, China 2 Central South University of Forestry and Technology, Changsha, China

ABSTRACT Designs and controllable fabrications of hollow nano materials from renewable natural resources have recently become an increasingly important research branch in materials area. In this chapter, preparation, structure and performance of hollow nano materials derived from agricultural lignin is discussed. The whole chapter is composed of five parts: (1) Introduction; (2) Molecular designs by applying click reactions between lignin and other polymer chains, i.e., Azide groups, polyethylene glycol chains and other polymer chains are grafted onto lignin, which make lignin amphiphilic molecular structures; (3) Fabrication of size-controlled lignin nanospheres; (4) Fabrication of the hollow nano materials by the interacting molecular chains in selected solvent system and characterization of morphology of lignin nano structure to study the effect of the molecular chains on the self-assembly of lignin nano structure; (5) Conclusion and Prospect. This chapter provides new methods for the fabrication of bio hollow nano-materials and some value-added applications of renewable lignin resources.

Keywords: lignin hollow nanospheres, molecular design, self-assembly

*

Corresponding Author Email: [email protected]

208

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

1. INTRODUCTION Considerable efforts have been directed toward the development of different kinds of materials derived from lignin, with attempts to take the advantage of the molecular structure of lignin and to find new alternatives to petroleum resources. Designs and controllable fabrications of hollow nanomaterials from renewable natural resources have recently become an increasingly important research branch in materials area. Lignin is an abundant biomass resource, which is defined as a random, amorphous, and threedimensional polymeric network. The major chemical functional groups in lignin include hydroxyl, methoxyl, carbonyl, and carboxyl, whose contents vary with the botanic origin and applied extraction processes. Hydroxyl groups on the aromatic ring of lignin are the most characteristic functional groups, which determine the reactivity and constitute the reactive sites to be exploited in lignin’s chemical reaction. In this chapter, preparation, structure and performance of hollow nano materials derived from lignin is discussed, which will provide a new method for the fabrication of bio hollow nano-materials and the valueadded application of renewable lignin resources.

2. MOLECULAR DESIGNS OF LIGNIN CHAIN STRUCTURES 2.1. Click Reaction between Alkyne and Azide The development of renewable polymers is driven by the reduction of the dependence on fossil fuels as organic material feedstocks. In recent years, polymers from natural and renewable resources have attracted a lot of attention, owing to their low cost, renewability, unique properties, and possible biodegradability [1-10]. Together with hemicelluloses, lignin, a natural polymer, acts as a cementing matrix of cellulose fibers within structure of plants and trees. It is one of the principal constituents of wood (around 25 - 35%), and it is the second abundant natural polymer. Lignin has an amorphous, highly branched macromolecular network structure with aromatic nature and complex compositions [11-14]. Currently, only a small portion of lignin waste from the pulping and paper industry is used as the feedstocks for low molecular weight chemicals and polymer applications [15-18]. Conversion of lignin into high valued products will have a positive impact on the biorefinery economics [19-21]. Several studies have explored the use of lignin as an inexpensive filler and additive in material applications. For instance, lignin was used in biobased polymeric composites to modify their properties such as hydrophobicity and stiffness [22-25]. Because the hydroxyl group is one of the most characteristic functional groups in lignin, the development of functional polymeric materials involving this type of

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 209 groups through esterification and etherification, have been investigated [26-29]. The modified lignin derivatives have broader applications in composite materials via grafting polymerization and cross-link coupling [30-32]. The Washburn’s group found that a polymer-grafted strategy significantly enhanced toughness of composites compared to homopolymers [31, 33]. We found that UV-absorbent thermoplastic elastomers could be prepared by polymer-grafted lignins [27]. Novel polyols derived from the lignin and fatty acids have been used for polyurethane fabrication [29]. The concept of “click” chemistry was first introduced by Sharpless and coworkers, defined as a reaction that is modular, wide in scope, and high in yield, with little side products which could be easily removed by non-chromatographic methods [34]. Click chemistry is a powerful approach with high fidelity in assembling specially designed building blocks [35-38]. Click chemistry could play an important role in the field of renewable polymers from natural products, through the choice of appropriate building blocks [39-41]. The most widely studied click reactions is Huisgen cycloaddition between azide and alkyne groups. In most cases, this reaction usually involves the use of copper catalysts [42-44]. The Bertozzi group introduced a novel strategy to avoid the use of copper, which is mostly based on strained cyclooctyne, however, the preparation of functional cyclooctynes is still challenging and limited in small scales, making it only applicable for some niche applications. Historically, huisgen cycloaddition was thermally promoted, meanwhile numerous materials have been prepared through the alkyne-azide cyclization at a higher temperature in the absence of any catalysts [45-47]. Propargylation of phenolic hydroxyl groups using propargyl bromide through nucleophilic substitution mechanism led to modified lignin product which could serve as the reactive building blocks for high molecular weight polymers because aryl propargyl ether moieties can be used as a prospective component for the click chemistry along with an organic azide [48-50]. Herein this section focus on a new approach to the functionalization of lignin via copper-free click chemistry [51-53]. This approach particularly caters the utilization of biomass toward the low-cost scalable renewable polymers and composites [54-56]. Specifically, we tried to utilize the concept of thermal click chemistry between azide and alkyne groups. As a result, a variety of lignin derivatives (Lignin-Azide, Lignin-Alkyne, Lignin-PCL-Alkyne and Lignin-(PCL-co-PLA)-Alkyne) were synthesized. Composites with high lignin content were prepared via this copper-free thermal click chemistry. This work also aims to display the application of click chemistry in lignin modifications, and the use of biodegradable polyesters can also maximize the sustainability. The synthesis of lignin polymers were carried out to use alkyne-azide thermal click reaction. Lignin-Lignin polymers were synthesized with Lignin-Alkyne with LigninAzide. Lignin-PEG-Lignin polymers were synthesized with Lignin-Alkyne and PEGAzide. Lignin-PCL-Lignin polymers were synthesized with Lignin-PCL-Alkyne and Lignin-Azide. Lignin-(PCL-co-PLA)-Lignin polymers were synthesized with Lignin(PCL-co-PLA)-Alkyne and Lignin-Azide.

210

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

2.2. Synthesis of Lignin Polymers via Click Reaction of Lignin-Alkyne with Lignin-Azide Lignin has a large amount of hydroxyl groups which can be functionalized through appropriate modifications such as esterification. Lignin-Br was synthesized through simple esterification reactions between bromobutyryl chloride and phenolic and aliphatic alcohol groups of lignin in the presence of TEA. Compared with the unmodified lignin, 1H NMR spectra (Figure 1) confirmed the formation of ester groups on lignin, as there are characteristic peaks at 2.2 ppm, 2.7 ppm and 3.5 ppm, corresponding to the methylene protons of bromobutyryl group. The bromo group in lignin was converted into azide (Lignin-Azide) subsequently, meanwhile, we observed shift of methylene –CH2Br (peak a) to –CH2N3 (peak d). On the other hand, lignin was reacted with propargyl bromide directly in the presence of potassium carbonate, yielding alkyne-modified lignin (LigninAlkyne). The incorporation of propargyl group onto lignin could be evidently seen from the broad peak of the attached methylene group (-O-CH2-) (4.3 to 4.9 ppm) and the proton on the alkyne group (2.4 to 2.5 ppm) in the 1H NMR spectrum.

Figure 1. 1H NMR spectra of lignin, Lignin-Br, Lignin-Azide, and Lignin-Alkyne [57], copyright with permission.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 211

Lignin-Lignin

Lignin-Azide

N3 aromatic ring of lignin

5000

4000

3000

2000

1000

0

Wavenumber (cm-1) Figure 2. FT-IR spectra of Lignin-Azide and lignin polymer after click reaction [57], copyright with permission.

FT-IR spectra of Lignin-Azide showed a strong absorption peak at ~ 2100 cm-1, corresponding to the azide absorption (Figure 2). The peaks in the range of 1406-1590 cm−1 correspond to the aromatic ring vibration. The azide absorption peak disappeared completely after thermal click reaction. The FT-IR spectra clearly demonstrated high fidelity of thermal click reaction between Lignin-Alkyne and Lignin-Azide. Besides, after the thermal treatment of mixed Lignin-Alkyne and Lignin-Azide, the product was no longer soluble in any organic solvents, such as THF, DCM, and toluene, which is attributed to the formation of crosslinked structures.

2.3. Synthesis of Lignin Polymers via Click Reaction of Lignin-Alkyne with PEG-Azide The reaction between Lignin-Azide and Lignin-Alkyne proved the possibility to make renewable polymer materials by a thermally-promoted click reaction. We therefore examined the integration of other components into lignin by introducing azide or alkyne functionalized polymers. Poly(ethylene glycol) (PEG) was functionalized with terminal azide groups. Tosylation and subsequent azide substitution were used for modification. 1H NMR spectra (Figure 3) confirmed completion of the tosylation and substitution reactions. FT-IR spectra of PEG-azide showed strong absorption at ~ 2100 cm-1, indicating the incorporation of the azide group (Figure 4).

212

Yanming Han, Fuquan Xiong, Gaiyun Li et al. f O

N3

N3

O

102

f

e

b

c

O O S O

d c

8

O

a O

S O

O

d

e

b

7

a

O

102

6

5

4

3

Chemical Shift (ppm) Figure 3. 1H NMR spectra of PEG-OTs and PEG-Azide [57], copyright with permission.

The di-functional PEG-Azide was used as a cross-linker to react with Lignin-Alkyne through a thermal click process. The azide peak at ~ 2100 cm-1 in the FT-IT spectra disappeared completely after thermal click reaction. The cured product also lost the solubility in common solvents.

Lignin-PEG-Lignin

PEG-Azide N3

4000

3000

2000

1000

Wavenumbers (cm-1) Figure 4. FT-IR spectra of PEG-Azide and Lignin-PEG-Lignin after click reaction [57], copyright with permission.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 213

Lignin-PCL-Lignin

Lignin-Azide

N3

5000

4000

3000

2000

1000

0

Wavenumbers (cm-1) Figure 5. FT-IR spectra of Lignin-Azide and Lignin- PCL- Lignin polymer after the click reaction [57], copyright with permission.

2.4. Synthesis of Lignin Polymers via Click Reaction of Lignin-PCL-Alkyne with Lignin-Azide The presence of hydroxyl groups on lignin would allow the preparation of grafted copolymers from the lignin via ring-opening polymerization (ROP). Incorporation of PCL could add biodegradability to the final product because polyesters were biodegradable polymers. We carried out ROP of CL using lignin as the initiator in the presence of stannous octoate. The obtained polymer Lignin-PCL was further propargylated with propargyl bromide in the presence of K2CO3. The resultant alkyne-containing polymer Lignin-PCL-Alkyne was reacted with Lignin-Azide to prepare a composite polymer Lignin-PCL-Lignin. FT-IR spectra also demonstrated the successful reaction between Lignin-Azide and Lignin-PCL-Alkyne. The strong azide absorption completely disappeared after the thermal click reaction (Figure 5).

2.5. Synthesis of Lignin Polymers via the Click Reaction of Lignin- (PCL- coPLA)-Alkyne with Lignin-Azide In order to demonstrate the robustness of this copper-free click reaction, we further applied it to prepare a copolymer grafted lignin. ROP of lactide and caprolactone from

214

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

lignin were catalyzed by stannous octoate, resulteing in a Lignin-(PCL-co-PLA) copolymer. Propargylation of the –OH end groups in Lignin-(PCL-co-PLA) was carried out by esterification with 5-hexynoic acid. Incorporation of the propargyl group onto Lignin-(PCL-co-PLA) could be evidently seen in the 1H NMR spectrum (Figure 6).

Figure 6. 1H NMR spectrum of Lignin-(PCL-co-PLA)-Alkyne [57], copyright with permission.

Lignin-(PCL-co-PLA)-Lignin

Lignin-Azide

N3

5000

4000

3000

2000

1000

0

Wavenumbers (cm-1)

Figure 7. FT-IR spectra of Lignin-Azide and Lignin-(PCL-co-PLA)-Lignin polymer after click reaction [57], copyright with permission.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 215 Lignin-(PCL-co-PLA)-Lignin was prepared by a thermal click reaction between Lignin-Azide and Lignin-(PCL-co-PLA)-Alkyne. FT-IR spectra demonstrated the reaction between alkyne groups and azide groups. The peak at ~2100 cm-1, corresponding to the azide absorption, disappeared completely after the thermal click reaction (Figure 7). Final product was found to be insoluble in organic solvents such as THF and DCM.

2.6. DSC Analysis of Lignin Polymers Prepared by the Thermal Click Reactions Lignin has a relatively high glass transition temperature (Tg) becausethe condensed rigid phenolic moieties and strong intermolecular hydrogen bonding interactions restrict mobility of lignin molecules. The grafting of PEG, PCL and PCL-co-PLA from lignin are expected to affect their thermal properties by their different chain structures. DSC analysis of the lignin polymers clearly shows that the incorporation of PEG, PCL and PCL-co-PLA chains significantly influences properties of lignin polymers (Figure 8). Depending on the grafted polymer structures, Tgs of lignin polymers varied in the range of - 34°C and 155°C with melting temperatures ranging from 38°C to 55°C.

Tg=-34 oC d

o

Tm=38 C c Tm=55 oC Tg=121 oC

b

Tm=51 oC Tg=155 oC a

-50

0

50

100

150

200

Temperature (oC) Figure 8. DSC curves of different lignin polymers: (a) Lignin-Lignin; (b) Lignin-PEG-Lignin; (c) Lignin-PCL-Lignin; (d) Lignin-(PCL-co-PLA)-Lignin [57], copyright with permission.

216

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

3. FABRICATION OF SIZE-CONTROLLED LIGNIN NANOSPHERES 3.1. Size-Controlled Lignin Nanospheres Self-assembly based on the spontaneous control of materials through noncovalent interactions, such as hydrogen bonding, van der Waals forces [58], electrostatic forces [59] and π-π interactions provides an effective method for application of materials. Recently, self-assembly are undergoing explosive development [60]. In order to prepare molecular aggregates with designed properties, geometries, and dimensions, various strategies for molecular self-assembly have been employed. Self-assembly of micelle may have potential applications in many fields, such as drug delivery [61], catalysis [62], medical diagnostics [63] and sensors [64]. Amphiphilic polymers with both hydrophobic and hydrophilic segments possess self-assembly behavior [65]. Different self-assembly micelle morphologies can be acquired such as spherical micelles, wormlike micelles, and vesicles by a precise control of the environmental conditions [66]. Spherical micelles are fit for using as carriers for agents [67]. In recent years, with rapid energy exhaustion and environmental awareness, increased attention has been paid to the development of spherical micelles obtained from natural renewable resources due to their inherent biodegradability and biocompatibility [65, 67-69]. Due to lignin’s complex structures, although it is an abundant renewable polyaromatic polymer composed of phenylpropanoid units, it could not be used extensively. In order to explore high-value-added application of lignin resources, scientists have made many attempts. Lignin nanoparticles can provide many opportunities for value-added applications of lignin [67, 70]. They may have potential applications in UV protection, antibacterial, nanofiller and bio-based carrier [71-74]. This suggests that lignin could play an important role as a new inexpensive and renewable starting material for propagating new products [75]. So far many lignin nanoparticles preparation methodswere reported, the nanoparticles obtained by methods, such as a precipitation method [76], a mechanical method [70] and polyaddition [77], usually possess irregular shape while spherical lignin nanoparticles can be prepared by self-assembly. Qian [67] produced uniform colloidal spheres using acetylated lignin via self-assembly; however, this step required environmentally unfriendly chemicals such as acetyl bromide. Theoretically, lignin exhibits self-assembly behavior because it is a natural amphiphilic polymer composed of hydrophilic hydroxyl groups and hydrophobic aromatic rings [78]. Lievonen alobtained spherical lignin nanoparticles with compact structures using the unmodified lignin through dialysis [69]. Enzymatic hydrolysis lignin shows better chemical activity, applicability and solubility in ubiquitous organic solvents than lignosulfonate or kraft lignin [57, 79]. However, lignin

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 217 has not been efficiently utilized [80], therefore, the development of value-added ligninbased materials, chemicals or fuels could greatly improve the biorefinery viability through an energy-effective and cost-effective route [82]. Application of lignin in phenolformaldehyde resin [81], polyurethane foams [82] and UV-absorbent coatings [83, 84] has been reported in the literature. Meanwhile, in our recent research, a kind of lignin-based nanoparticles fluorescently labeled by pyrene has been prepared, which have potential applications in nano sensors due to its oxygen-responsive property [75]. Herein this section mainly focused on a simple self-assembly method for fabricating uniform lignin nanospheres through π-π interactions without the need of chemical modification of lignin. Lignin was dissolved in tetrahydrofuran at different initial concentrations and subsequently self-assembled with adding water under magnetic stirring for fabricating the nanospheres [85].

3.2. Morphology, Size and Yield of Lignin Nanospheres Typical TEM images for fabricating lignin nanospheres at four different pre-dropping concentrations were presented in Figure 9. Highly ordered and uniform lignin nanospheres could be seen. Increasing the pre-dropping lignin concentration resulted in an increase in the diameter of the lignin nanospheres. This may be due to the fact that higher pre-dropping lignin concentration leads to the participation of more lignin molecules in the formation of each nanosphere, thereby larger diameter of the nanospheres [69]. The change rule of diameter of the nanospheres with pre-dropping lignin concentration was further confirmed by dynamic light scattering (DLS), which measured the particle sizes of the nanospheres in deionized water. Figure 10A shows that the nanospheres size increased from 327 nm to 588 nm with increasing of the pre-dropping lignin concentration from 0.5 mg/ml to 2 mg/ml. It is worth noting that the yield of the nanospheres rised with the increase of the pre-dropping lignin concentration. The highest yield of the nanospheres was 72.15% at a pre-dropping lignin concentration of 2 mg/ml. The stability of the nanospheres at different pre-dropping concentrations was studied through tracking the size change of the nanospheres with time. Figure 10B shows that average diameters of the four kinds of lignin nanospheres had no significant change after 30 days, due to the effect of electrical double layer repulsion [69]. When lignin nanospheres are dispersed in deionized water, a surface charge, which is produced by the hydroxyl groups and carboxyl groups of lignin molecules, promotes formation of electrical double layers and thereby stabilize nanospheres dispersion. Meanwhile, the typical TEM images of four kinds of nanospheres after 30 days showed that the average diameter of the lignin nanospheres had no aggregation or degradation, compared with the initial nanospheres.

218

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

Figure 9. TEM images of lignin nanospheres obtained at different pre-dropping lignin concentrations. Pre-dropping lignin concentration: (A) 0.5 mg/ml, (B) 1 mg/ml, (C) 1.5 mg/ml, and (D) 2 mg/ml. The sample used here was prepared at a stirring rate of 600 rpm and at a dropping speed of 4 ml/min [85], copyright with permission.

Figure 10. The yield and size of lignin nanospheres obtained at different pre-dropping lignin concentrations (A) and the size change of the nanospheres with time (B). The sample used here was prepared at a stirring rate of 600 rpm and at a dropping speed of 4 ml/min [85], copyright with permission.

3.3. The Chemical Characteristics of Lignin Nanospheres In order to compare chemical structural features of lignin nanospheres in the preparation process, FTIR spectra of lignin and the nanospheres were analyzed and the results were showed in Figure 11. The lignin exhibited bands at 3400 cm-1 (the stretching vibrations of the hydroxyl group), 1700 cm-1 (C=O stretching), 1598 cm-1 (aromatic skeletal vibration), 1514 cm-1 (C-C stretching of aromatic skeletal), 1460 cm-1 (C-H stretching of aromatic skeletal), and 1425 cm-1 (aromatic skeletal vibrations combined with C-H in-plane deformation) [75, 86, 87]. Compared to the raw lignin, no significant change

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 219 was observed, suggesting that the preparation process of the nanospheres had no impact on the structures of lignin.

Figure 11. FTIR spectra of lignin and lignin nanospheres obtained at different pre-dropping lignin concentration. The sample used here was prepared at the stirring rate of 600 rpm and at a dropping speed of 4 ml/min [85], copyright with permission.

3.4. The Composition Characteristics of Lignin Nanospheres Further analysis of TEM images of lignin nanospheres obtained at different predropping concentration shows that a few nanospheres existed gaps at surface, especially for the sample obtained at initial concentration of 0.5 mg/ml. It is due to the effect of volatile speed of solvent. To prove this, we contrasted morphologies of the nanospheres at two different preparation temperatures. The sample used in this part was prepared at an initial concentration of 0.5 mg/ml, a stirring rate of 600 rpm and a dropping speed of 4 ml/min. The preparation temperatures of samples were 40°C and 15°C, respectively. Then samples were dropped on the copper grids for SEM and TEM. Figure 12A is the typical SEM image of sample obtained at the preparation temperature of 40°C. It can be observed that the surface of almost every nanosphere had gaps. Compared with a higher preparation temperature, the nanospheres obtained at the preparation temperature of 15°C were nearly without gaps (Figure 12B). Figure 12C and D are TEM images for the nanospheres obtained at the preparation temperature of 40°C and 15°C, respectively. THF and water are miscible at room temperature [88]. Lignin has a better solubility in THF than in water. When deionized water was gradually added into lignin-tetrahydrofuran solution, the nanospheres slowly formed through hydrophobic aggregation. In the preparation process, a higher temperature would lead to quicker evaporation of residual

220

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

THF in the nanospheres, which breaks through the surface of the nanospheres, leading to the formation of the gaps.

Figure 12. Typical SEM and TEM images of lignin nanospheres obtained at two different preparation temperatures: (A, C) 40°C and (B, D) 15°C. The sample used here was prepared at an initial lignin concentration of 0.5 mg/ml, at a stirring rate of 600 rpm and at a dropping speed of 4 ml/min [85], copyright with permission.

3.5. Size Influences of Lignin Nanospheres Effects of the stirring rate and the dropping speed of water on lignin nanospheres size were studied at the fixed initial lignin concentration of 0.5 mg/ml and the preparation temperature of 15°C. The acquiescent stirring rate and dropping speed of water were 600 rpm and 4 ml/min, respectively. Figure 13A exhibits the relationship between the average diameter and the stirring rate. The average diameter of the nanospheres decreased with the increase of the stirring rate because high stirring rate may improve the mixing performance of the aqueous phase and the organic phase. Moreover, the insets showed the typical TEM images of lignin nanospheres obtained at stirring rates of 500 rpm and 800 rpm, respectively. The average diameter of the lignin nanospheres obtained at the stirring rate of 500 rpm was bigger than that of 800 rpm. The relationship between average diameter and dropping speed of water are showed in Figure 13B. The average diameter of the nanospheres decreased with the increasing of the dropping speed of water. Correspondingly, typical TEM images of the lignin nanospheres are illustrated at dropping speeds of water, 2 ml/min and 8 ml/min, respectively. Compared with the lignin nanospheres obtained at the dropping speed of water of 2 ml/min, the

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 221 average diameter of the lignin nanospheres would decrease at the dropping speed of water of 8 ml/min, which is attributed to the quick transformation of lignin molecules in “frozen” state for formation of the lignin nanospheres [89].

3.6. The Formation Mechanism of Lignin Nanospheres The formation process of the lignin nanospheres is presented in Figure 14 (A-C). In state A, lignin was completely dissolved in THF, so no obvious particles could be seen on corresponding TEM images (Figure 14A). In state B, when water was added to the solution, some lignin molecules with stronger hydrophobic ability formed the cores of the nanospheres at the water content of 50 vol% (Figure 14B). When water content increased to 80 vol% in the system, the THF in the solution was removed by a dialysis process. The stable lignin nanospheres could be seen in the Figure 14C. The formation rule is consistent with acetylated lignin colloidal spheres reported in the literature [67].

Figure 13. Effects of the stirring rate (A) and the dropping speed of water (B) on lignin nanospheres size. The initial lignin concentration in THF and preparation temperature were fixed at 0.5 mg/ml and 15°C, respectively. The acquiescent stirring rate and the dropping speed of water are 600 rpm and 4 ml/min, respectively. Insets are TEM images of lignin nanospheres corresponding to the stirring rate and the dropping speed of water [85], copyright with permission.

Figure 14. TEM images of the samples obtained from the dispersions at different water contents. The pre-dropping lignin concentration was 1.5 mg/ml. Water content: (A) 0 vol%, (B) 50 vol%, (E) > 80 vol%, after removing THF [85], copyright with permission.

222

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

Previous studies have suggested that lignin in organic solvents would tend to form aggregates through π-π interaction of the aromatic rings [90-92]. The driving force for formation of lignin nanospheres is considered to be π-π interactions between lignin molecules. This can be confirmed by the examination of absorption spectra. Figure 15 shows the UV-Vis absorption spectra of lignin and lignin nanospheres. The inset is the enlarged part of Figure 15. The absorption peaks at around 280 nm belongs to π-π transition of the characteristics of guaiacyl structural units in lignin [93]. Raw lignin exhibits one strong absorption band at 275 nm in deionized water and THF, respectively, suggesting that blank changes without affecting peak positions of the absorption spectra. However, the absorption band of lignin nanospheres in water is red-shifted to 280 nm. After it was redissolved in THF, the red-shifted absorption band returns to 274 nm. These results confirm that the existence of π-π interactions between lignin molecules in the process of forming the nanospheres and the tendency of the lignin molecules growing into a spherical shape in order to minimize their surface energy. Based on the above controlled experiments, a possible formation mechanism for the nanospheres is proposed. Lignin nanospheres were formed through a layer-by-layer selfassembly approach from inside to outside based on π-π interactions. With increasing of initial lignin concentration, more lignin molecules participate in the formation of the nanospheres, leading to an increase in the diameter of the nanospheres. The average diameter of the nanospheres decreased with an increase of stirring rate or the dropping speed of water. Moreover, the preparation temperature has an effect on regularity of the nanospheres.

Figure 15. UV-Vis absorption spectra of lignin and lignin nanospheres. The nanospheres were obtained at an initial lignin concentration of 0.5 mg/ml, at a stirring rate of 600 rpm and at a dropping speed of 4 ml/min [85], copyright with permission.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 223 Compared to the previous preparation method for lignin nanospheres [67], the method presented in this work does not need chemical modification of lignin. Meanwhile, the typical morphology features of particles obtained by the straightforward dialysis method [69] and the method described in this work were compared. Lignin nanospheres were prepared by dissolving lignin in tetrahydrofuran (0.5 mg/ml) and subsequently introducing water into the system through dialysis (15°C). The results showed that the particles obtained by the straightforward dialysis method were also spherical but exhibited distinct gaps. The nanospheres this work exhibited tunable size and regularity without any gaps, and their size could be controlled by the initial lignin concentration, stirring rate, the dropping speed of water and the preparation temperature.

4. FABRICATION OF HOLLOW NANOSPHERES WITH A SINGLE HOLE 4.1. The Preparation of Lignin Hollow Nanospheres The hollow nano or macrospheres usually show substantially different properties in terms of density, specific surface area and surface permeability [94, 95]. They are expected to be applied in many fields such as coating technology, catalysis, delivery systems, and composite materials [96]. When hollow spheres are used in carrying medicine, which usually have higher efficiency drug loading than spheres with compact structures. Tremendous efforts have been made in fabrication of hollow spheres with controlled composition, tailored structure, and unique properties [97, 98]. Recently, as a new class of hollow structure, a hollow capsule with a single hole, that is also known as open-mouthed hollow capsule, has stimulated great interests because the enhanced uptake capacity, diffusivity, and catalytic performance [99-102]. Moreover, hollow particles with controllable holes have potential value in the field of selective encapsulation [103]. The hollow spheres with an opening hole have been reported using various types of organic polymer and inorganic oxide, respectively, such as chiral phenylacrylonitrile tartaric acids [104], polystyrene [105], poly(acryamide-ethylene glycol dimeth acrylate) [106], polystyrene/poly(divinylbenzene) [107], polystyrene/poly(3,4- ethylenedioxythiophene) [108], poly(methylsilsesquioxane) [109], titanium dioxide [102], and silica [110]. The reported lignin nanoparticles preparation methods include a mechanical method, self-assembly, a precipitation method and poly addition. Yiamsawas prepared hollow lignin nanocapsules by interfacial polyaddition and the resultant lignin nanoparticles were of irregular shape [77]. The formation mechanism of the hollow spheres is still unknown. Recently, the increased demand for bio-ethanolfrom lignocellulosic biomass was driven by depletion of fossil, leading to the production of lignin in large quantities as a by-product [111]. If lignin could be transformed to value-added materials, chemicals or fuels through

224

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

an energy-effective and cost-effective route, it could greatly improve the biorefinery viability. In light of the potential value of the hollow nanospheres with a single hole and renewable enzymatic hydrolysis lignin [103], this section mainly focused on a simple onestep method for fabricating lignin hollow nanospheres with a single hole, without the aid of surfactants and a template removal step during the preparation [112].

4.2. Morphology and Size of Lignin Hollow Nanospheres Typical SEM and TEM images of lignin hollow nanospheres at different initial lignin concentration are showed in Figure 16. The SEM images exhibited hollow structure and a single hole on the surface of particle, the particle was spherical (when the initial lignin concentration was fixed at 0.2 mg/ml, the particle was irregular). It is worth noting that the diameter of the single hole of the nanospheres decreased with increasing the initial lignin concentration. Their hollowness was further supported by using TEM. The TEM images displayed a clear contrast between the center and the shell, indicatingthe presence of a cavity. Furthermore, the thickness of shell wall increased with the increase of the predropping lignin concentration. The reason can be explained that a higher pre-dropping lignin concentration means more lignin available for the formation of shell wall. This indicates that the size of single hole and the thickness of shell wall can be adjusted by employing different pre-dropping lignin concentrations. The particle sizes of the hollow nanospheres in deionized water were tracked by using DLS. As the increase of the pre-dropping lignin concentration, an increase of the mean nanospheres size (from 419 nm to 566 nm) was observed in Figure 17A. A higher predropping lignin concentration means more lignin available for the formation of shell in the initial phase. An increase in particle size as a function of the initial polymer concentration has been observed in the synthesis of lignin nanospheres with compact structures [113]. Compared with conventional polymer hollow spheres, the lignin hollow spheres have smaller diameters [105, 107, 114]. Figure 17B showed that the stability of lignin hollow nanospheres at pre-dropping lignin concentrations of 0.5 mg/ml and 2 mg/ml, respectively. Apparent changes in the average diameter of two kinds of lignin hollow nanospheres were not observed after 15 days while the nanospheres aggregation occurred after 45 days, with an increase in the average diameter. The nanosphere edges became obscure after sixty days. When lignin hollow nanospheres are dispersed in water, the phenolic hydroxyl groups and carboxyl groups provide the nanospheres a surface charge that promotes the formation of electrical double layers, which can stabilize the nanospheres dispersion via electrical double layer repulsion. Lignin hollow nanospheres with asymmetric physical chemistry properties will

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 225 affect electrical double layer structure, leading to the nanospheres aggregation. It is worth noting that the increased ratio of the diameter of the nanospheres at a pre-dropping lignin concentration of 0.5 mg/ml was higher than that of 2 mg/ml after 45 days, which may be attributed to the larger hole of the former

4.3. Surface Area, Pore Size and Distribution of Lignin Hollow Nanospheres Nitrogen adsorption-desorption isotherms of lignin and lignin hollow nanospheres is presented respectively (Figure 18). The lower portion of loop is traced out on adsorption, and the upper portion on desorption. A hysteresis loop was found in the adsorptiondesorption isotherm of all samples, which resembles the type IV of Brunauer’s classification [115]. The very low amount of nitrogen adsorbed at low relative pressure indicates the nearly absence of the microporosity. When pre-dropping lignin concentration was 2 mg/ml, the hollow nanospheres exhibited pore radius distribution in 1.5 to 14 nm range, supporting that the sample was mesopores material. As for the hollow nanospheres at a pre-dropping lignin concentration of 0.5 mg/ml, it showed the pore radius distribution from 15 to 60 nm, demonstrating the presence of mesopores and macropores. The surface areas of lignin hollow nanospheres were 25.4 m2 g-1 and 5.79 m2 g-1 at predropping lignin concentrations of 0.5 mg/ml and 2 mg/ml, respectively, which were higher than 0.824 m2 g-1 of the raw lignin sample. Under the same conditions, a higher predropping lignin concentration leaded to a higher thickness of shell wall, leading to the decrease of the surface area. The results are in agreement with the results obtained from the corresponding TEM images. Moreover, the changing trend of total pore volume was consistent with the surface area.

Figure 16. SEM (A-D) and TEM (E-H) images of lignin hollow nanospheres at different pre-dropping lignin concentration. Pre-dropping lignin concentration: (A, E) 0.5 mg/ml, (B, F) 1 mg/ml, (C, G) 1.5 mg/ml, (D, H) 2 mg/ml [112], copyright with permission.

226

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

Figure 17. Particle size analysis of lignin hollow nanospheres. (A) Effect of pre-dropping lignin concentration on lignin hollow nanospheres size (inset are TEM images of lignin hollow nanospheres). (B) Effect of time after preparation on lignin hollow nanospheres size (inset are TEM images of lignin hollow nanospheres, and the nanosphere edges become obscure after sixty days) [112], copyright with permission.

Figure 18. Nitrogen sorption isotherms of lignin and lignin hollow nanospheres at different predropping lignin concentrations (0.5 mg/ml and 2 mg/ml) [112], copyright with permission.

4.4. The Chemical Characteristics of Lignin Hollow Nanospheres The lignin and lignin hollow nanospheres were characterized by FTIR analysis and the results are showed in Figure 19A. The spectra are normalized to 1 at 1510-1514 cm-1 that is assigned to maximum absorbance peak of aromatic skeleton stretch in the studied samples. The lignin exhibited bands at 3400 cm-1 (the stretching vibrations of the hydroxyl group), 2900 cm-1 (C-H stretching of methyl or methylene groups), 1700 cm-1 (C=O stretching), 1598 cm-1 (aromatic skeletal vibration), 1514 cm-1 (C-C stretching of aromatic skeletal), 1460 cm-1 (C-H stretching of aromatic skeletal), 1425 cm-1 (aromatic skeletal vibrations combined with C-H in-plane deformation), 1332 cm-1 (syringyl units vibration), 1118 cm-1 (aromatic C-H deformation in syringyl unit), and 1031 cm-1 (OH stretching of

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 227 primary alcohol) [75, 87, 116]. In comparison to the raw lignin, lignin hollow nanospheres showed the same bands, hinting that the chemical structure of lignin did not change during the preparation of the lignin hollow nanospheres. It is worth noting that the characteristic peak of the lignin hollow nanospheres at 1700 cm-1 was weakened compared with the raw material. Moreover, the semi-quantitative analysis of FTIR spectra showed that the A1700/A1514 ratio of lignin was 0.5528, which was higher than that of lignin hollow nanospheres (0.5058 and 0.4828) at pre-dropping lignin concentrations of 0.5 mg/ml and 2 mg/ml, respectively. This may be because lignin molecules more carboxy groups may not produce aggregation during the preparation of lignin hollow nanospheres. The yields of lignin hollow nanospheres were 70.8% and 69.9% (w/w) at pre-dropping lignin concentration of 0.5 mg/ml and 2 mg/ml, respectively, and suggested that some lignin molecules were not involved in the formation of lignin hollow nanospheres. 31 P NMR spectra of samples may be used to determine the amount of carboxyl, phenolic hydroxy groups and aliphatic hydroxy groups in lignin [117]. Lignin is marked with reagents containing phosphorus and subjected to NMR analysis [118]. Figure 19B summarizes the 31P NMR analysis results of the lignin and lignin hollow nanospheres. Compared with lignin raw material, contents of aliphatic -OHs, carboxyl groups and phenolic -OHs decreased except that aliphatic -OHs of lignin hollow nanospheres at the pre-dropping lignin concentration for 0.5 mg/ml increased slightly. It suggested that some hydrophilic lignin molecules might not produce aggregation during the preparation of the lignin hollow nanospheres, leading to that the lower concentration of hydrophilic groups in the lignin hollow nanospheres than that in the raw lignin. Furthermore, it is highlighted that the content of carboxyl groups of the lignin hollow nanospheres was decreased distinctly, hinting that it had significant influences on the hydrophilicity of lignin, consistent with the result of the FTIR.

Figure 19. FTIR spectra (A) and structural characteristics calculated from the 31P NMR spectra (B) of lignin and lignin hollow nanospheres at different pre-dropping lignin concentrations (0.5 mg/ml and 2 mg/ml). (S) syringyl units; (G) guaiacyl units; (H) p-hydroxyphenyl units [112], copyright with permission.

228

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

XPS can be used to measure elemental composition for top 1-5 nm depth of the surface region [119]. The O/C ratio on the nanospheres surface region was measured by XPS (the typical measurement error is about 10%). Meanwhile the O/C ratio of the nanospheres was measured by using elemental analysis (the typical measurement error is about 0.1%). Since the hydrophilic groups of the lignin hollow nanospheres are hydroxyl and carboxyl groups, the more hydrophilic chains must have a higher O/C ratio. The O/C ratio on the nanospheres surface region is 0.19, and the O/C ratio of the nanospheres is 0.46. It is because the hollow nanospheres have a relatively hydrophobic external surface and a relatively hydrophilic internal surface, probably due to a layer of hydrophobic lignin membrane formed at two-phase interface between water and THF and the gradual aggregation of lignin molecules in internal surface of membrane.

4.5. Effects of Stirring Rate, Dropping Speed of Water and pH on the Characteristics of Lignin Hollow Nanospheres Effects of stirring rates, dropping speeds of water and pH values on the characteristics of lignin hollow nanospheres were studied at a fixed initial lignin concentration of 0.5 mg/ml. The acquiescent stirring rate and dropping speed of water were 600 rpm and 2 ml/min, respectively. Without stirring, the hollow nanospheres exhibited a smaller diameter in average, but the uniformity is less satisfying. Figure 20A shows that an increase of the stirring rate from 500 rpm to 800 rpm resulted in a decrease of the mean hollow nanospheres size from 480.4 nm to 353.5 nm. It is demonstrated that a higher stirring rate causes a greater shear force, leading to forming a smaller size of the hollow nanospheres [120]. The typical SEM and TEM images of lignin hollow nanospheres showed that the diameter of the hollow nanospheres was bigger at the stirring rate of 500 rpm than at the stirring rate of 800 rpm, consistent with the results of the DLS. Moreover, the SEM images displayed that the diameter of single hole was bigger at the stirring rate of 500 rpm than at the stirring rate of 800 rpm.

Figure 20. Effects of stirring rate (A), dropping speed of water (B) and pH (C) on the characteristics of lignin hollow nanospheres. Insets are TEM images of lignin hollow nanospheres at pH values of 1.8 (a) and 12.3 (b), respectively [112], copyright with permission.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 229 Figure 20B shows the relationship between the average diameter and the dropping speed of water. With increase of the dropping speed of water, athedecrease of mean hollow nanospheres sizthee (from 435.8 nm to 340.1 nm) was observed. When the dropping speed of water was increased to 20 ml/min, the average diameter decreased further. This is mainly because the preparation process of hollow nanospheres via self-assembly approach exhibits competition of thermodynamics and kinetics [89]. When the dropping speed of water increase, the ratio of kinetics in preparation process increases, leading to the transformation of lignin molecules in “frozen” state for the formation of lignin hollow nanospheres. Correspondingly, the typical SEM and TEM images of the lignin hollow nanospheres are illustrated at dropping speeds of water of 1 ml/min and 4 ml/min, respectively, consistent with the results of the DLS. Effect of pH value of suspension on characteristics of lignin hollow nanospheres is shown in Figure 20C. It can be seen that the nanosphere dispersions were stable at pH values between 3.5 and 12. When the pH values of dispersions were less than 3.5, the average particle sizes of nanospheres gradually increased. This may be attributed to the aggregation caused by the decreased negative charge of the nanospheres surface at pH values less than 3.5 [69]. TEM images of the nanospheres at pH 1.8 exhibited blocking aggregation (Figure 20C-a). Furthermore, the average particle size of nanospheres was almost zero at pH values above 12, probably attributed to the dissolution of the nanospheres in alkali solution above pH 12. As can be seen from the TEM morphology, the sample at pH of 12.3 was dissolved.

4.6. The Formation Mechanism of Lignin Hollow Nanospheres The nanospheres form hollow structure due to the effect of THF levels. Normally THFwater is miscible. However, we found that phase separation existed between THF of analytical grade purity (AR-THF) and water. AR-THF-water have Tyndall phenomenon compared with THF of chromatographic grade (HPLC-THF), indicating that phase separation existed between AR-THF and water due to the formation of nano-emulsion [121]. It is due to the fact that AR-THF may contain a small amount of impurity compared with HPLC-THF so that phase separation existed between THF and water. In order to prove this, we compared the morphologies of the nanospheres prepared with two different THF solvents, i.e., HPLC-THF and THF of adding the same volume of HPLC-THF after ARTHF at natural volatilize. Figure 21A is the typical TEM image of nanospheres prepared with HPLC-THF. It can be seen that the nanospheres exhibited the compact structure. However, the nanospheres prepared with THF of adding the same volume of HPLC-THF after AR-THF exhibited the hollow structure (Figure 21B), suggesting that a small amount of impurity in AR-THF is the reason for forming the hollow structure. Moreover, Tyndall phenomenon was found in the mixture between THF of adding the same volume of HPLC-

230

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

THF after AR-THF at natural volatilize and water. By further determination of the compositions of AR-THF by GC-MS, we found that AR-THF contained some impurities, such as toluene, trimethylphosphine oxide, butylated hydroxytoluene and triethyl citrate, compared with HPLC-THF. They may lead to the less polarity of AR-THF so that phase separationexisted between AR-THF and water, thereby forming a nano-emulsion system. At last, the nanospheres with hollow structure are produced by means of nano-emulsion soft template. The formation process of lignin hollow nanospheres is proposed in Figure 22 (A-E). In state A, lignin was dissolved in THF completely, so no obvious particles could be seen on the corresponding TEM images (Figure 22A). When deionized water was dropped to the lignin/tetrahydrofuran solution, phase separation took place, that is, continuous phase (THF) and dispersed phase (water). In state B, when water content reached 20 vol%, some lignin molecules with a stronger hydrophobic ability formed a layer of membrane at the two-phase interface between t water and THF, leading to the wrapping of water. An dynamic equilibrium between continuous phase and dispersed phase was formed [122]. When the sample was dropped onto copper grids, the hollow spheres were formed with evaporation of water (Figure 22B). Further increasing water content would bring about an increase of the pressure gradient between the inside and outside of the membrane until the dynamic equilibrium was destroyed [123]. When water content reached 40% (v/v), the phase inversion took place and the pressure gradient decreased resulting in the breaking of the membrane (Figure 22C). The phase that was previously continuous became dispersed, leading to the reformation of the relatively hydrophobic membrane. Therefore, lignin-THF solution was wrapped by the membrane. As water content further increased, more and more water permeated through the membrane, and led to more lignin molecules aggregating in the internal surface. Furthermore, its pressure gradient inside and outside gradually increased so that a single hole could appear at the thinner side [124]. The lignin hollow nanospheres with single holes could be seen at a water content of 50 vol% in Figure 22D. The lignin hollow nanospheres formed completely at a water content of 80 vol%, and then the THF in the solution was removed by the dialysis process. The stable lignin hollow nanospheres could be seen in Figure 22E. The driving force of lignin molecules assembly is considered π-π interactions, which is demonstrated by UV-Vis absorption spectra (Figure 23). The absorption peak at 275 nm which belongs to π-π transition of the characteristics of guaiacyl structural [93], was observed for the raw lignin in deionized water and THF, respectively. It is shown that the peak position of the absorption spectra is not affected when the blank changes. The absorption peak of the lignin hollow nanospheres in water is red-shifted to 284 nm, and the red-shifted absorption peak returns to 280 nm after redissolving in THF. This demonstrated the existence of π-π interactions between lignin molecules during the formation of the lignin hollow nanospheres.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 231

Figure 21. TEM images of lignin nanospheres prepared with two different THF, (A) HPLC-THF and (B) THF of adding the same volume of HPLC-THF after AR-THF at natural volatilize. The sample used here was prepared at a pre-dropping lignin concentration of 0.5 mg/ml, at a stirring rate of 600 rpm and at a dropping speed of 4 ml/min [112], copyright with permission.

Figure 22. TEM images of the samples obtained from the dispersions at different water contents. The pre-dropping lignin concentration was 0.5 mg/ml. Water content: (A) 0 vol%, (B) 20 vol%, (C) 40 vol%, (D) 50 vol%, (E) > 80 vol%, after removing THF. (G) Schematic representation of formation process of the lignin hollow nanosphere in a THF-H2O [112], copyright with permission.

Based on the above experiments, a possible formation mechanism for the lignin hollow nanospheres is proposed, as shown schematically in Figure 22. Lignin can be completely dissolved in THF. By adding water, lignin molecules with a stronger hydrophobic will form a layer of membrane at the two-phase interface between water and THF, leading to the wrapping of water. Increasing water content will bring about an increase of pressure gradient between the inside and outside of the membrane until phase inversion takes place. Lignin-THF solution is wrapped by the membrane. As water content further increases, more and more water molecules permeate through the membrane, leading to that the aggregation of more and more lignin molecules at the internal surface of the membrane by a layer-by-layer self-assembly based on π-π interactions. Finally, the thinner side of the membrane will fracture for maintaining stable pressure inside and outside. Therefore, it is reasonable to believe that it is the pressure difference between the inside and outside of the membrane that leads to formation of hole in the shell of each hollow nanosphere. The primary holes are formed at the thinnest part of micelle walls. With the increasing of predripping lignin concentrations, more lignin molecules participate during the formation of the micelle walls so that the thickness of micelle walls increase and therefore the diameter of single holes decreases.

232

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

Figure 23. UV-Vis absorption spectra of lignin and lignin hollow nanospheres (concentration 0.1 mg/5 ml). The nanospheres were obtained at an initial lignin concentration of 0.5 mg/ml, at a stirring rate of 600 rpm and at a dropping speed of 2 ml/min [112], copyright with permission.

Compared to the template method, i.e., direct polymerization and cross-linking reactions occurring at the surface of the template and the complete removal of the template, the simple and convenient self-assembly method avoids the introduction of impurities, therefore is suitable for modern chemical preparation. We hope this novel method could contribute to the development of new strategies to fabricate nanolignin materials with novel structures and functions.

CONCLUSION This chapter provides new methods for the fabrication of bio-based hollow nanomaterials and for value-added applications of renewable lignin resources. Reactive lignin derivatives with either alkyne or azide groups, including Lignin-Azide, Lignin-Alkyne, Lignin-PCL-Alkyne or Lignin- (PCL-co-PLA) -Alkyne, were prepared through facile chemical modifications from lignin. Molecular design and rearrangement of lignin were carried out by application of click reactions between lignin and other polymer chains. Azide groups, polyethylene glycol chains and other polymer chains are grafted onto lignin, which make lignin amphiphilic molecular structures. Fabrication of the ordered hollow nano structure is achieved by the interaction between molecular chains and selective solvent system. The morphology of lignin hollow nano structure is characterized. The effect of the molecular chains on the self-assembly of lignin hollow nano structure is studied in detail, as well as other fabrication factors. The proposed lignin nanospheres are

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 233 eco-friendly, cost-effective and therefore a promising candidate for biomass-based nanocarriers. The morphology and surface properties of lignin nanospheres can affect their biomedical applications. For the morphology of lignin nanospheres, the diameters of both the microspheres and the internal cavity, the sizes of holes and their distribution in the shell are expected to be controlled precisely. The shell breaking method is mainly suitable for the preparation of polymer nanospheres. In some cases, the materials used in this method must be capable of swelling or shrinkage. The inhomogeneous diameters of nanospheres could also limit their biomedical applications with special demands. In future research, versatile and bio-friendly methods for the fabrication of lignin nanospheres with controllable compositions and morphologies are desired [95]. For example, we should pay more attention to the following research areas of lignin nanospheres: (1) lignin nanospheres with hybrid resources to meet holistic function requirements should be more focused; (2) for diagnosis and therapy, controlled release of functional materials at the destination site is very important. Surface modifications of lignin nanospheres may be an efficient way to achieve the controlled release; (3) in some biomedical applications, lignin nanospheres with larger holes are preferable while in some cases they may cause the leakage problem. Hence, precise control of the morphology of lignin nanospheres is still challenging. We hope that the general principles of lignin nanospheres will provide the readers with the useful knowledge and ideas to develop more effective methods for the fabrication of lignin nanospheres.

ACKNOWLEDGMENT This work was supported by National Natural Science Foundation of China (Grant No. 31770610).

REFERENCES [1] [2] [3] [4]

Coates, G. W. & Hillmyer, M. A. (2015). “A Virtual Issue of Macromolecules: “Polymers from Renewable Resources”.” Macromolecules, 42, 7987-89. Hillmyer, M. A. & Tolman, W. B. (2014). “Aliphatic polyester block polymers: renewable, degradable, and sustainable.” Acc Chem Res, 47, 2390-96. Sharma, V. & Kundu, P. P. (2008). “Condensation polymers from natural oils.” Progress in Polymer Science, 33, 1199-215. Xia, Y. & Larock, R. C. (2010). “Vegetable oil-based polymeric materials: synthesis, properties, and applications.” Green Chemistry, 12, 1893-909.

234 [5] [6]

[7]

[8]

[9] [10]

[11] [12] [13] [14]

[15] [16]

[17] [18]

[19]

Yanming Han, Fuquan Xiong, Gaiyun Li et al. Yao, K. & Tang, C. (2013). “Controlled Polymerization of Next-Generation Renewable Monomers and Beyond.” Macromolecules, 46, 1689-712. Wilbon, P. A., Chu, F. & Tang, C. (2013). “Progress in renewable polymers from natural terpenes, terpenoids, and rosin.” Macromolecular Rapid Communications, 34, 8-37. Mathers, R. T. (2011). “How well can renewable resources mimic commodity monomers and polymers?” Journal of Polymer Science Part A Polymer Chemistry, 50, 1-15. Meier, M. A. R., Metzger, J. O. & Schubert, U. S. (2007). “ChemInform Abstract: Plant Oil Renewable Resources as Green Alternatives in Polymer Science.” Chemical Society Reviews, 36, 1788-802. Miller, S. A. (2013). “Sustainable Polymers: Opportunities for the Next Decade.” Acs Macro Letters, 2, 550-54. Wang, J., Yao, K., Wang, C., Tang, C. & Jiang, X. (2013). “Synthesis and drug delivery of novel amphiphilic block copolymers containing hydrophobic dehydroabietic moiety.” Journal of Materials Chemistry B, 1, 2324-32. Chung, H. & Washburn, N. R. (2012). “Chemistry of lignin-based materials.” Green Materials, 1, 137-60. Calvo-Flores, F. G. & Dobado, J. A. (2010). “Lignin as Renewable Raw Material.” Chemsuschem, 3, 1227-35. Laurichesse, S. & Avérous, L. (2014). “Chemical modification of lignins: Towards biobased polymers.” Progress in Polymer Science, 39, 1266-90. Saito, T., Brown, R. H., Hunt, M. A., Pickel, D. L., Pickel, J. M., Messman, J. M., Baker, F. S., Keller, M. & Naskar, A. K. (2012). “Turning renewable resources into value-added polymer: development of lignin-based thermoplastic.” Green Chemistry, 14, 3295-303. Holmberg, A. L., Reno, K. H. & Wool, R. P. (2014). “Biobased building blocks for the rational design of renewable block polymers.” Soft Matter, 10, 7405-24. Pandey, M. P. & Kim, C. S. (2011). “Lignin Depolymerization and Conversion: A Review of Thermochemical Methods.” Chemical Engineering & Technology, 34, 29-41. Wang, J., Manley, R. S. J. & Feldman, D. (1992). “Synthetic polymer-lignin copolymers and blends.” Progress in Polymer Science, 17, 611-46. Zakzeski, J., Bruijnincx, P. C., Jongerius, A. L. & Weckhuysen, B. M. (2013). “The catalytic valorization of lignin for the production of renewable chemicals.” Chemical Reviews, 110, 3552-99. Uihlein, N. & Schebek, L. (2009). “Environmental impacts of a lignocellulose feedstock biorefinery system: an assessment.” Biomass & Bioenergy, 33, 793-802.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 235 [20] Quan, B., Lei, H., Zacher, A. H., Lu, W., Ren, S., Jing, L., Yi, W., Liu, Y., Tang, J., Qin, Z. (2012). “A review of catalytic hydrodeoxygenation of lignin-derived phenols from biomass pyrolysis.” Bioresource Technology, 124, 470-77. [21] Zhang, X., Tu, M. & Paice, M. G. (2011). “Routes to Potential Bioproducts from Lignocellulosic Biomass Lignin and Hemicelluloses.” Bioenergy Research, 4, 24657. [22] Blinkovsky, A. M. & Dordick, J. S. Peroxidase-catalyzed synthesis of lignin, phenol copolymers. [23] Avérous, L. (2004). “Biodegradable Multiphase Systems Based on Plasticized Starch: A Review.” Journal of Macromolecular Science Part C, 44, 231-74. [24] Wos, D., Mousavioun, P. & Fellows, C. M. (2011). “Value-adding to cellulosic ethanol: Lignin polymers.” Industrial Crops & Products, 33, 259-76. [25] Wang, J., Yao, K., Korich, A. L., Li, S., Ma, S., Ploehn, H. J., Iovine, P. M., Wang, C., Chu, F. & Tang, C. (2011). “Combining renewable gum rosin and lignin: Towards hydrophobic polymer composites by controlled polymerization.” Journal of Polymer Science Part A Polymer Chemistry, 49, 3728-38. [26] Korich, A. L., Fleming, A. B., Walker, A. R., Wang, J., Tang, C. & Iovine, P. M. (2012). “Chemical modification of organosolv lignin using boronic acid-containing reagents.” Polymer 53, 87-93. [27] Yu, J., Wang, J., Wang, C., Liu, Y., Xu, Y., Tang, C. & Chu, F. (2015). “UVabsorbent lignin-based multi-arm star thermoplastic elastomers.” Macromolecular Rapid Communications, 36, 398. [28] Nemoto, T., Konishi, G. I., Tojo, Y., An, Y. C. & Funaoka, M. (2011). “Functionalization of lignin: Synthesis of lignophenol-graft-poly(2-ethyl-2oxazoline) and its application to polymer blends with commodity polymers.” Journal of Applied Polymer Science, 123, 2636-42. [29] Laurichesse, S., Huillet, C. & Averous, L. (2014). “Original polyols based on organosolv lignin and fatty acids: New bio-based building blocks for segmented polyurethane synthesis.” Green Chemistry, 16, 3958-70. [30] Jiang, S., Kai, D., Dou, Q. Q. & Xian, J. L. (2015). “Multi-arm carriers composed of antioxidant lignin core and poly(glycidyl methacrylate-co-poly(ethylene glycol) methacrylate) derivative arms for highly efficient gene delivery.” Journal of Materials Chemistry B, 3, 6897-904. [31] Gupta, C. & Washburn, N. R. (2014). “Polymer-Grafted Lignin Surfactants Prepared via Reversible Addition–Fragmentation Chain-Transfer Polymerization.” Langmuir the Acs Journal of Surfaces & Colloids, 30, 9303-12. [32] Zhang, J., Chen, Y., Sewell, P. & Brook, M. A. (2015). “Utilization of softwood lignin as both crosslinker and reinforcing agent in silicone elastomers.” Green Chemistry, 17, 3176-76.

236

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

[33] Hilburg, S. L., Elder, A. N., Chung, H., Ferebee, R. L., Bockstaller, M. R. & Washburn, N. R. (2014). “A universal route towards thermoplastic lignin composites with improved mechanical properties.” Polymer, 55, 995-1003. [34] Kolb, H. C., Finn, M. G. & Sharpless, K. B. (2010). “Click Chemistry: Diverse Chemical Function from a Few Good Reactions.” Cheminform, 32, no-no. [35] Wu, P., Feldman, A. K., Nugent, A. K., Hawker, C. J., Scheel, A., Voit, B., Pyun, J., Fréchet, J. M., Sharpless, K. B. & Fokin, V. V. (2004). “Efficiency and fidelity in a click-chemistry route to triazole dendrimers by the copper(i)-catalyzed ligation of azides and alkynes.” Angewandte Chemie, 43, 3928-32. [36] Brett Helms, †, J. L. M., And, C. J. H., ‡, & †, J. M. J. F. (2004). “Dendronized Linear Polymers via “Click Chemistry”.” Journal of the American Chemical Society, 126, 15020. [37] Iha, R. K., Wooley, K. L., Nyström, A. M., Burke, D. J., Kade, M. J. & Hawker, C. J. (2009). “Applications of Orthogonal, “Click” Chemistries in the Synthesis of Functional Soft Materials.” Chemical Reviews, 109, 5620-86. [38] Hawker, C. J. & Wooley, K. L. (2005). “The convergence of synthetic organic and polymer chemistries.” Science, 309, 1200-05. [39] June, S. M., Bissel, P. & Long, T. E. (2012). “Segmented block copolyesters using click chemistry.” Journal of Polymer Science Part A Polymer Chemistry, 50, 3797805. [40] Yuan, L., Hamidi, N., Smith, S., Clemons, F., Hamidi, A. & Tang, C. (2015). “Molecular characterization of biodegradable natural resin acid-substituted polycaprolactone.” European Polymer Journal, 62, 43-50. [41] Yao, K., Wang, J., Zhang, W., Lee, J. S., Wang, C., Chu, F., He, X. & Tang, C. (2011). “Degradable rosin-ester-caprolactone graft copolymers.” Biomacromolecules, 12, 2171-7. [42] Tunca, U. Orthogonal multiple click reactions in synthetic polymer chemistry, 2014. [43] Sumerlin, B. S., Tsarevsky, N. V., Louche, G., And, R. Y. L. & Matyjaszewski, K. (2005). “Highly Efficient “Click” Functionalization of Poly(3-azidopropyl methacrylate) Prepared by ATRP.” Macromolecules, 38, 7540-45. [44] Golas, P. L. & Matyjaszewski, K. (2010). “Marrying click chemistry with polymerization: expanding the scope of polymeric materials.” Chemical Society Reviews, 39, 1338-54. [45] Lutz, J. F. “Copper-free azide-alkyne cycloadditions.” [46] Besset, C., Binauld, S., Ibert, M., Fuertes, P., Pascault, J., Fleury, E., Bernard, J. & Drockenmuller, E. (2011). “Copper-Catalyzed vs Thermal Step Growth Polymerization of Starch-Derived α-Azide−ω-Alkyne Dianhydrohexitol Stereoisomers: To Click or Not To Click?” Macromolecules, 43, 17-19. [47] And, G. O. J. & Houk, K. N. (2008). “Predictions of Substituent Effects in Thermal Azide 1,3-Dipolar Cycloadditions:  Implications for Dynamic Combinatorial

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 237

[48]

[49]

[50] [51]

[52] [53]

[54]

[55]

[56]

[57] [58]

[59]

[60]

(Reversible) and Click (Irreversible) Chemistry.” Journal of Organic Chemistry, 73, 1333-42. Huaming Li, Fuyong Cheng, Andy M. Duft, a. & Alex Adronov, ‡. (2005). “Functionalization of Single-Walled Carbon Nanotubes with Well-Defined Polystyrene by “Click” Coupling.” Journal of the American Chemical Society, 127, 14518-24. Wei, P., Yan, X., Li, J., Ma, Y., Yao, Y. & Huang, F. (2012). “Novel [2]rotaxanes based on the recognition of pillar[5]arenes to an alkane functionalized with triazole moieties.” Tetrahedron, 68, 9179-85. Nair, C. P. R., Bindu, R. L. & Ninan, K. N. (2001). “Thermal characteristics of addition-cure phenolic resins.” Polymer Degradation & Stability, 73, 251-57. Rambarran, T., Gonzaga, F. & Brook, M. A. (2013). “Multifunctional amphiphilic siloxane architectures using sequential, metal-free click ligations.” Journal of Polymer Science Part A Polymer Chemistry, 51, 855-64. Canalle, L. A., Berkel, S. S. V., Haan, L. T. D. & Hest, J. C. M. V. (2010). “CopperFree Clickable Coatings.” Advanced Functional Materials, 19, 3464-70. Spruell, J. M., Wolffs, M., Leibfarth, F. A., Stahl, B. C., Heo, J., Connal, L. A., Hu, J. & Hawker, C. J. (2011). “Reactive, multifunctional polymer films through thermal cross-linking of orthogonal click groups.” Journal of the American Chemical Society, 133, 16698-706. Hong, J., Luo, Q. & Shah, B. K. (2010). “Catalyst- and Solvent-Free “Click” Chemistry: A Facile Approach to Obtain Cross-Linked Biopolymers from Soybean Oil.” Biomacromolecules, 11, 2960-5. Hong, J., Luo, Q., Wan, X., Petrović, Z. S. & Shah, B. K. (2012). “Biopolymers from vegetable oils via catalyst- and solvent-free “click” chemistry: effects of crosslinking density.” Biomacromolecules, 13, 261-6. Peng, P., Cao, X., Feng, P., Jing, B., Xu, F. & Sun, R. (2012). “Binding cellulose and chitosan via click chemistry: Synthesis, characterization, and formation of some hollow tubes.” Journal of Polymer Science Part A Polymer Chemistry, 50, 5201-10. Han, Y., Yuan, L., Li, G., Huang, L., Qin, T., Chu, F. & Tang, C. (2016). “Renewable polymers from lignin via copper-free thermal click chemistry.” Polymer, 83, 92-100. Wu, Z., Liu, J., Li, Y., Cheng, Z., Li, T., Zhang, H., Lu, Z. & Yang, B. (2015). “SelfAssembly of Nanoclusters into Mono-, Few-, and Multilayered Sheets via DipoleInduced Asymmetric van der Waals Attraction.” Acs Nano, 9, 6315-23. Zheng, Y., Rosa, L., Thai, T., Ng, S., Gomez, D., Ohshima, H. & Bach, U. (2014). “Asymmetric gold nanodimer arrays: electrostatic self-assembly and SERS activity.” Journal of Materials Chemistry A, 3, 240-49. Zhang, K., Jiang, M. & Chen, D. (2012). “Self-assembly of particles—The regulatory role of particle flexibility.” Progress in Polymer Science, 37, 445-86.

238

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

[61] Tyrrell, Z. L., Shen, Y. & Radosz, M. (2010). “Fabrication of micellar nanoparticles for drug delivery through the self-assembly of block copolymers.” Progress in Polymer Science, 35, 1128-43. [62] Polshettiwar, V., Baruwati, B. & Varma, R. S. (2009). “Self-Assembly of Metal Oxides into Three-Dimensional Nanostructures: Synthesis and Application in Catalysis.” Acs Nano, 3, 728. [63] Sandhu, A., Handa, H. & Abe, M. (2010). “Synthesis and applications of magnetic nanoparticles for biorecognition and point of care medical diagnostics.” Nanotechnology, 21, 442001. [64] Chen, X., Jing, X., Wang, J., Liu, J., Song, D. & Liu, L. (2013). “Self-assembly of ZnO nanoparticles into hollow microspheres via a facile solvothermal route and their application as gas sensor.” Crystengcomm, 15, 7243-49. [65] Guo, Y., Zhang, L., Li, H., Han, Y., Zhou, J. & Wang, X. (2016). “Self-assembly and paclitaxel loading capacity of α-tocopherol succinate-conjugated hydroxyethyl cellulose nanomicelle.” Colloids polymer Science, 294, 135-43. [66] Jain, S. & Bates, F. S. (2003). “On the origins of morphological complexity in block copolymer surfactants.” Science, 300, 460-64. [67] Qian, Y., Deng, Y., Qiu, X., Li, H. & Yang, D. (2014). “Formation of uniform colloidal spheres from lignin, a renewable resource recovered from pulping spent liquor.” Green Chemistry, 16, 2156-63. [68] Guo, Yanzhu., Chen, Hui., Wang, Xiaohui., Shen, Zuguang. & Shu, Xuancai. (2013). “Direct grafting modification of pulp in ionic liquids and self-assembly; behavior of the graft copolymers.” Cellulose, 20, 873-84. [69] Lievonen, M., Valle-Delgado, J. J., Mattinen, M. L., Hult, E. L., Lintinen, K., Kostiainen, M. A., Paananen, A., Szilvay, G. R., Setälä, H. & Österberg, M. (2016). “Simple process for lignin nanoparticle preparation.” Green Chemistry, 18, 1416-22. [70] Nair, S. S., Sharma, S., Pu, Y., Sun, Q., Pan, S., Zhu, J. Y., Deng, Y. & Ragauskas, A. J. (2015). “High Shear Homogenization of Lignin to Nanolignin and Thermal Stability of Nanolignin‐Polyvinyl Alcohol Blends.” Chemsuschem, 7, 3513-20. [71] Gîlcǎ, I. A., Cǎpraru, A. M., Grama, S. & Popa, V. I. (2011). “Agents for wood bioprotection based on natural aromatic compounds and their complexes with copper and zinc.” Cellulose Chemistry & Technology, 45, 227-31. [72] Richter, A. P., Brown, J. S., Bharti, B., Wang, A., Gangwal, S., Houck, K., Cohen Hubal, E. A., Paunov, V. N., Stoyanov, S. D. & Velev, O. D. (2015). “An environmentally benign antimicrobial nanoparticle based on a silver-infused lignin core.” Nature Nanotechnology, 10, 817-23. [73] Yang, W., Fortunati, E., Dominici, F., Kenny, J. M. & Puglia, D. (2015). “Effect of processing conditions and lignin content on thermal, mechanical and degradative behavior of lignin nanoparticles/polylactic (acid) bionanocomposites prepared by melt extrusion and solvent casting.” European Polymer Journal, 71, 126-39.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 239 [74] Zimniewska, M., Kozłowski, R. & Batog, J. (2008). “Nanolignin Modified Linen Fabric as a Multifunctional Product.” Molecular Crystals & Liquid Crystals, 484, 43/[409]-50/[16]. [75] Xiong, F., Han, Y., Li, G., Qin, T., Wang, S. & Chu, F. (2016). “Synthesis and characterization of renewable woody nanoparticles fluorescently labeled by pyrene.” Industrial Crops & Products, 83, 663-69. [76] Frangville, C., Rutkevi, M., ius, d. x., Richter, A. P., Velev, O. D., Stoyanov, S. D. & Paunov, V. N. (2013). “Fabrication of Environmentally Biodegradable Lignin Nanoparticles.” Chemphyschem, 13, 4235-43. [77] Yiamsawas, D., Baier, G., Thines, E., Landfester, K. & Wurm, F. R. (2014). “Biodegradable lignin nanocontainers.” Rsc Advances, 4, 11661-63. [78] Bartzoka, E. D., Lange, H., Thiel, K. & Crestini, C. (2016). “Coordination Complexes and One-Step Assembly of Lignin for Versatile Nanocapsule Engineering.” Acs Sustainable Chemistry & Engineering, 4. [79] Jin, Y., Cheng, X. & Zheng, Z. (2010). “Preparation and characterization of phenolformaldehyde adhesives modified with enzymatic hydrolysis lignin.” Bioresource Technology, 101, 2046-48. [80] Ragauskas, A. J., Beckham, G. T., Biddy, M. J., Chandra, R., Chen, F., Davis, M. F., Davison, B. H., Dixon, R. A., Gilna, P. & Keller, M. (2014). “Lignin valorization: improving lignin processing in the biorefinery.” Science, 344, 1246843. [81] Chu, Fuxiang., Zhang, Ma. & Wang, Li Zhang. (2013). “Preparation and properties of lignin-phenol-formaldehyde resins based on; different biorefinery residues of agricultural biomass.” Industrial Crops & Products, 43, 326-33. [82] Li, Y., Han, Y., Qin, T. & Chu, F. (2012). “Preparation of Polyurethane Foams Based on Liquefied Corn Stalk Enzymatic Hydrolysis Lignin.” Journal of Biobased Materials & Bioenergy, 6, 51-58. [83] Liu, X., Zong, E., Jiang, J., Fu, S., Wang, J., Xu, B., Li, W., Lin, X., Xu, Y. & Wang, C. (2015). “Preparation and characterization of Lignin-graft-poly (ε-caprolactone) copolymers based on lignocellulosic butanol residue.” International Journal of Biological Macromolecules, 81, 521-29. [84] Zong, E., Jiang, J., Liu, X., Fu, S., Xu, Y. & Chu, F. (2016). “Combination of lignin and l-lactide towards grafted copolymers from lignocellulosic butanol residue.” International Journal of Biological Macromolecules, 86, 80-88. [85] Xiong, F., Han, Y., Wang, S., Li, G., Qin, T., Chen, Y. & Chu, F. (2017). “Preparation and formation mechanism of size-controlled lignin nanospheres by selfassembly.” Industrial Crops & Products, 100, 146-52. [86] Casas, A., Alonso, M. V., Oliet, M., Rojo, E. & Rodríguez, F. (2012). “FTIR analysis of lignin regenerated from Pinus radiata and Eucalyptus globulus woods dissolved in imidazolium-based ionic liquids.” Journal of Chemical Technology & Biotechnology, 87, 472-80.

240

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

[87] Xiong, F., Zhou, L., Qian, L. & Liu, S. (2014). “Effects of Pretreatment Methods Using Various 1,4-Dioxane Concentrations on the Performance of Lignocellulosic Films of Eucalyptus citriodora.” Bioresources, 10. [88] Lejček, P., Matouš, J., Novák, J. P. & Pick, J. (1975). “Phase equilibria and excess molar volumes of tetrahydrofuran (1) + deuterium oxide (2).” Journal of Chemical Thermodynamics, 7, 927-35. [89] Li, H., Deng, Y., Liu, B., Ren, Y., Liang, J., Qian, Y., Qiu, X., Li, C. & Zheng, D. (2016). “Preparation of Nanocapsules via the Self-Assembly of Kraft Lignin: A Totally Green Process with Renewable Resources.” Acs Sustainable Chemistry & Engineering, 4. [90] Deng, Y., Feng, X., Zhou, M., Qian, Y., Yu, H. & Qiu, X. (2011). “Investigation of Aggregation and Assembly of Alkali Lignin Using Iodine as a Probe.” Biomacromolecules, 12, 1116-25. [91] Sarkanen, S., Teller, D. C., Abramowski, E. & Mccarthy, J. L. (1982). “Lignin. 19. Kraft lignin component conformation and associated complex configuration in aqueous alkaline solution.” Macromolecules, 15, 1098-104. [92] Sarkanen, S., Teller, D. C., Hall, J. & Mccarthy, J. L. (1981). “Lignin. 18. Associative effects among organosolv lignin components.” Macromolecules, 14, 426-34. [93] Ouyang, X., Deng, Y., Qian, Y., Zhang, P. & Qiu, X. (2011). “Adsorption characteristics of lignosulfonates in salt-free and salt-added aqueous solutions.” Biomacromolecules, 12, 3313. [94] Rafati, A. A., Borujeni, A. R. A., Najafi, M. & Bagheri, A. (2011). “Ultrasonic/surfactant assisted of CdS nano hollow sphere synthesis and characterization.” Materials Characterization, 62, 94-98. [95] Si, Y., Chen, M. & Wu, L. (2016). “Syntheses and biomedical applications of hollow micro-/nano-spheres with large-through-holes.” Chemical Society Reviews, 45, 690714. [96] Hah, H. J., Kim, J. S., Jeon, B. J., Sang, M. K. & Yong, E. L. (2003). “Simple Preparation of Monodisperse Hollow Silica Particles Without Using Templates.” Chemical Communications, 34, 1712-13. [97] Deng, Z., Chen, M., Zhou, S., You, B. & Wu, L. (2006). “A Novel Method for the Fabrication of Monodisperse Hollow Silica Spheres.” Langmuir the Acs Journal of Surfaces & Colloids, 22, 6403-7. [98] Zhang, Q., Wang, W., Goebl, J. & Yin, Y. (2009). “Self-templated synthesis of hollow nanostructures.” Nano Today, 4, 494-507. [99] Shi, Q., Zhang, P., Li, Y., Xia, H., Wang, D. & Tao, X. (2015). “Synthesis of openmouthed, yolk-shell [email protected] nanoparticles with access to interior surfaces for enhanced electrocatalysis.” Chemical Science, 6, 4350-57.

Molecular Design and Controllable Self-Assembly of Lignin Hollow Nanospheres 241 [100] Li, X., Zhou, L., Wei, Y., Eltoni, A. M., Zhang, F. & Zhao, D. (2015). “Anisotropic Encapsulation-Induced Synthesis of Asymmetric Single-Hole Mesoporous Nanocages.” Journal of the American Chemical Society, 137, 5903. [101] Xu, J., Ma, A., Xu, Z., Liu, X., Chu, D. & Xu, H. (2016). “Synthesis of Au and Pt Hollow Capsules with Single Holes via Pickering Emulsion Strategy.” Journal of Physical Chemistry C, 119. [102] Ding, S., Lin, T., Wang, Y., Lü, X. & Huang, F. (2013). “New facile synthesis of TiO2 hollow sphere with an opening hole and its enhanced rate performance in lithium-ion batteries.” New Journal of Chemistry, 37, 784-89. [103] Hyuk, I. S., Jeong, U. & Xia, Y. (2005). “Polymer hollow particles with controllable holes in their surfaces.” Nature Materials, 4, 671. [104] Li, D. M. & Zheng, Y. S. (2011). “Single-hole hollow nanospheres from enantioselective self-assembly of chiral AIE carboxylic acid and amine.” Journal of Organic Chemistry, 76, 1100-8. [105] Hyun, D. C., Lu, P., Choi, S. I., Jeong, U. & Xia, Y. (2013). “Microscale Polymer Bottles Corked with a Phase-Change Material for Temperature-Controlled Release.” Angewandte Chemie International Edition, 52, 10468-71. [106] Guan, G., Zhang, Z., Wang, Z., Liu, B., Gao, D. & Xie, C. (2010). “Single-Hole Hollow Polymer Microspheres toward Specific High‐Capacity Uptake of Target Species.” Advanced Materials, 19, 2370-74. [107] Minami, H., Kobayashi, H. & Okubo, M. (2005). “Preparation of hollow polymer particles with a single hole in the shell by SaPSeP.” Langmuir, 21, 5655-58. [108] Luo, S. C., Jiang, J., Liour, S. S., Gao, S., Ying, J. Y. & Yu, H. H. (2009). “Magnetic PEDOT hollow capsules with single holes.” Chemical Communications, 21, 266466. [109] Chang, M. W., Stride, E. & Edirisinghe, M. (2010). “A new method for the preparation of monoporous hollow microspheres.” Langmuir the Acs Journal of Surfaces & Colloids, 26, 5115-21. [110] Li, D. M., Chen, Y. C., Zhang, C., Song, S. & Zheng, Y. S. (2013). “Different morphologies of silica synthesized using organic templates from the same class of chiral compounds.” Journal of Materials Chemistry B, 1, 1622-27. [111] Liu, X., Wang, J., Li, S., Zhuang, X., Xu, Y., Wang, C. & Chu, F. (2014). “Preparation and properties of UV-absorbent lignin graft copolymer films from lignocellulosic butanol residue.” Industrial Crops & Products, 52, 633-41. [112] Xiong, F., Han, Y., Wang, S., Li, G., Qin, T., Chen, Y. & Chu, F. (2017). “Preparation and Formation Mechanism of Renewable Lignin Hollow Nanospheres with a Single Hole by Self-Assembly.” Acs Sustainable Chemistry & Engineering, 5.

242

Yanming Han, Fuquan Xiong, Gaiyun Li et al.

[113] Qiu, X. Q., Li, H., Deng, Y. H., Qian, Y. & Yi, C. H. (2014). “The Acetylation of Alkali Lignin and Its Use for Spherical Micelles Preparation.” Acta Polymerica Sinica, 1458-64. [114] Han, J., Song, G. & Guo, R. (2010). “A Facile Solution Route for Polymeric Hollow Spheres with Controllable Size.” Advanced Materials, 18, 3140-44. [115] Brunauer, S., Emmett, P. H. & Teller, E. (1938). “Adsorption of Gases in Multimolecular Layers.” Journal of the American Chemical Society, 60, 309-19. [116] Casas, A., Oliet, M., Alonso, M. V. & Rodríguez, F. (2012). “Dissolution of Pinus radiata and Eucalyptus globulus woods in ionic liquids under microwave radiation: Lignin regeneration and characterization.” Separation & Purification Technology, 97, 115-22. [117] Granata, A. & Argyopoulos, D. S. (1995). “2-chloro-4,4,5,5-tetramethyl-1,3,2dioxaphospholane, a reagent for the accurate determination of the uncondensed and condensed phenolic moieties in lignins.” Journal of Agricultural & Food Chemistry, 43, 1538-44. [118] Gilca, I. A., Popa, V. I. & Crestini, C. (2015). “Obtaining lignin nanoparticles by sonication.” Ultrasonics – Sonochemistry, 23, 369-75. [119] Tang, C. Y., Kwon, Y. N. & Leckie, J. O. (2007). “Probing the nano- and microscales of reverse osmosis membranes—A comprehensive characterization of physiochemical properties of uncoated and coated membranes by XPS, TEM, ATRFTIR, and streaming potential measurements.” Journal of Membrane Science, 287, 146-56. [120] Mateovic, T., Kriznar, B., Bogataj, M. & Mrhar, A. (2002). “The influence of stirring rate on biopharmaceutical properties of Eudragit RS microspheres.” Journal of Microencapsulation, 19, 29-36. [121] Zhang, C., Zhu, Y., Zhang, R., Xie, Y., Wang, K. & Liu, X. (2015). “Pickering emulsions stabilized by composite nanoparticles prepared from lysozyme and dopamine modified poly (γ-glutamic acid): effects of pH value on the stability of the emulsion and the activity of lysozyme.” Rsc Advances, 5, 90651-58. [122] Norato, M. A., Tavlarides, L. L. & Tsouris, C. (2010). “Phase inversion studies in liquid‐liquid dispersions.” Canadian Journal of Chemical Engineering, 76, 486-94. [123] Ioannou, K., Nydal, O. J. & Angeli, P. (2005). “Phase inversion in dispersed liquid– liquid flows.” Experimental Thermal & Fluid Science, 29, 331-39. [124] Lv, H., Lin, Q., Zhang, K., Yu, K., Yao, T., Zhang, X., Zhang, J. & Yang, B. (2008). “Facile fabrication of monodisperse polymer hollow spheres.” Langmuir the Acs Journal of Surfaces & Colloids, 24, 13736.

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 10

THERMAL CHARACTERIZATION OF KLASON LIGNINS FROM SOFTWOOD AND HARDWOOD SPECIES Matheus Poletto* Post-Graduate Program in Engineering of Processes and Technologies (PGEPROTEC), University of Caxias do Sul (UCS), Caxias do Sul, Brazil

ABSTRACT Lignin is a natural polymer presented in wood, being one of the most abundant biomacromolecules, second only to cellulose in natural abundance. The wood species affects the properties of lignin. Depending on the extraction method used its physicalchemical and thermal characteristics also differ considerably. This study investigates Klason lignins from softwood and hardwood species of Pinus taeda (LPIT) and Eucalyptus grandis (LEUG), respectively. The work evaluates the influence of its physical characteristics on the thermal properties and decomposition kinetics of the lignins analyzed. The lignins were characterized using Fourier transform infrared spectroscopy (FTIR), and thermogravimetric analysis (TGA). The results demonstrated that differences between thermal stability and thermal degradation mechanisms in the samples studied may be associated with differences in hydrogen bond density in lignin. LPIT presented higher intermolecular interactions between hydroxyl groups than LEUG. This result might indicate a higher strength of the intermolecular interactions in LPIT than in LEUG. The thermal degradation processes of both lignins initiate by diffusion process, however, at higher conversion values the degradation mechanism is a complex procedure due to the degradation of highly condensed aromatic structures presented in lignin.

Keywords: lignin, thermal stability, degradation mechanism

*

Corresponding Author Email: [email protected]

244

Matheus Poletto

INTRODUCTION The increased interest in the conversion of wood and its components for producing alternative fuels and chemicals necessitates a fundamental understanding of the main characteristics, composition, thermal behavior and processes involving pyrolysis of biomass [1-3]. Knowledge of the kinetics of thermal reactions is vital for predicting thermal behavior of biomass materials such as lignin [1, 3]. Lignin is a complex three-dimensional network natural polymer that serves as a continuous matrix component in plant cell walls [4-6]. Lignin is a polyaromatic polyol that is readily available and relative inexpensive, because can be obtained from lignocellulosic wastes. Depending on the extraction method used and its original source the physicalchemical characteristics differ considerably [4, 7]. Several methods have been developed to separate lignin from the other constituents of the lignocellulosic materials [2, 3, 8]. Each separation method modifies to some degree the chemical structure of the naturally occurring lignin. The Klason method is one of the most typical lignin determination procedures [8, 9]. The method separates lignin as an insoluble material by depolymerization of cellulose and hemicellulose in 72% sulfuric acid followed by hydrolysis of the dissolved polysaccharides in boiling 3% sulfuric acid [9]. The wood species also affects the properties of lignin [4, 7]. Chemically, softwood lignins consist largely of guaiacylpropane units, while hardwood lignins are basically composed of both guaiacyl- and syringylpropane units [4, 7, 10]. Figure 1 presents the chemical structure of some compounds found in lignin. Thus, the types and amounts of intra- and intermolecular hydrogen bonding present in lignin from softwoods or hardwoods may vary and influence its physical and chemical properties [4]. Several techniques have been used to characterize the structural aspects and functional groups present in lignin [4, 7, 11]. Therefore, infrared spectroscopy has been proven to be a highly effective means of investigating specific interactions within and between molecules [12-14]. FTIR can be used to qualitatively and quantitatively study the mechanism of intermolecular interactions through hydrogen bonding [12, 14]. On the other hand, fast pyrolysis is one of the lignin conversion processes that will potentially lead to more valuable phenol and phenolic compounds [8, 15]. Wang et al. performed pyrolysis of lignins isolated from hardwood and softwood, and found that methoxy group content has a significant influence on the pyrolysis of lignin [7]. Bährle et al. used Klason lignin for comparing the pyrolysis behavior of hardwood and softwood lignins [16]. They suggested that radical formation during hardwood Klason lignin pyrolysis was more frequent than that of softwood Klason lignin pyrolysis, and was mainly due to high methoxy group content in hardwood Klason lignin [16]. Understand the pyrolysis kinetics of lignin and its correlation with physical and chemical properties of wood is important to better understand wood thermal degradation

Thermal Characterization of Klason Lignins from Softwood …

245

and provide useful information for rational design and scaling-up of pyrolysis reactors [8, 15].

Figure 1. Chemical structure of several lignin model compounds.

In order to better understand the thermal degradation and pyrolysis kinetics of lignin and its correlation with their properties, this study investigates the differences in thermal behavior and degradation kinetics of two Klason lignins obtained from different tree species. The pyrolysis characteristics of Klason lignin isolated from hardwood specie (Eucalyptus grandis) and from softwood specie (Pinus taeda) were compared by thermogravimetry. The influence of hydrogen bonding intensity on the thermal properties and decomposition kinetics of both lignins was evaluated. To better understand the pyrolysis process, the degradation mechanism of both lignins was also evaluated.

Theoretical Background The fundamental equation used in all kinetic studies is generally described as [17]:

d  k (T ) f ( ) dt

(1)

where k is the rate constant and f (α) is the reaction model, a function dependent on the reaction mechanism. Eq. (1) expresses the rate of conversion, dα/dt, at a constant temperature as a function of the rate constant and the reduction in the reactant concentration. In this study, the conversion rate α is defined as:



m 0  mt m0  m f

(2)

Matheus Poletto

246

where m0 is the initial weight of the sample, mf is the final weight and mt is the sample’s weight at time (t). The rate constant k is generally given by the Arrhenius equation:

k (T )  Ae

 Ea RT

(3)

where Ea is the apparent activation energy (kJ.mol-1), R is the gas constant (8.314 kJ.mol1 -1 K ), A is the pre-exponential factor (min-1) and, T is the absolute temperature (K). Combing Eqs. (1) and (3) gives the following relationship:  Ea d  Ae RT f ( ) dt

(4)

For a dynamic thermogravimetric analysis (TGA) in a non-isothermal experiment, introducing the heating rate β=dT/dt into Eq. (4), Eq. (5) is obtained as:

d  A   Ea RT   e f ( ) dT   

(5)

Equations (4) and (5) are the fundamental expressions of analytical methods used to calculate kinetic parameters on the basis of the TGA data [13, 18].

Kissinger Method Generally, the Kissinger method is used to calculate the activation energy of the solid state reaction. The Kissinger method derives the activation energy using the peak temperature at which the maximum reaction rate occurs and the order of reaction is obtained using the shape of the mass loss-time curve [19]. The Kissinger method [19] adopts the following Eq. (6):   ln  2  Tmax





  AR Ea n 1    ln  ln n1   max     RTmax   Ea

(6)

where Tmax corresponds to the temperature at the maximum reaction rate of the TG curve and αmax is the conversion degree at the temperature at the maximum reaction rate. A plot of ln

 T  against  1 T 2 max

slope of this straight line [13, 19].

max

 produces a fitted straight line. According to the

Ea R the apparent activation energy

Ea

can be calculated

Thermal Characterization of Klason Lignins from Softwood …

247

Flynn-Wall-Ozawa Method The activation energy values for the degradation process were determined by the isoconversional Flynn-Wall-Ozawa (FWO) method. This method can be used for determination of the Ea values without any knowledge of the reaction mechanisms. This is defined by Eq. (7) [20, 21]:  AE a  E log   log   2.315  0.4567 a  RT  g  R 

(7)

where β is the heating rate, A is the pre-exponential factor, g(α) is a function of the conversion, Ea is the activation energy and R is the gas constant. Therefore, for different heating rates (β) and a given degree of conversion (α), a linear relationship is observed by plotting log β vs. 1/T, and the Ea is obtained from the slope of the straight line [20, 21].

Criado Method The degradation reaction mechanism can be determined using the Criado method [22], which can accurately determine the reaction mechanism in a solid reaction process. This is defined by a Z(α) type function: Z   

d dt  x T 

(8)

where x = Ea/RT and π(x) is an approximation of the temperature integral that cannot be expressed in a simple analytical form. In this study, were used the fourth rational expression of Senum and Yang [23], which gives errors lower than 10-5% when x > 20. The master curves as a function of the conversion degree corresponding to the different models listed in Table 1 [17] were obtained according to Eq. (9): Z    f  g  

(9)

From Eq. (5) and Eq. (8), the following relationship can be derived: Z   

d Ea RTa e P x  dT R E

(10)

Eq. (9) is used to plot the Z(α) versus α curves for the different models listed in Table 1, whereas Eq. (10) is used to represent the experimental curve. By comparing these two curves, the type of mechanism involved in the thermal degradation can be identified. The activation energy values obtained using FWO method were used to determine the

Matheus Poletto

248

degradation mechanisms proposed by Criado [22]. This method uses references theoretical curves obtained from Eq. (9) that are derivatives of f(α) and g(α)functions represented in Table 1, called master curves, which are compared with experimental data for determination of the mechanism of a solid-state reaction process. Table 1. Algebraic expressions of functions of the most common reaction mechanisms operating in solid-phase reactions Mechanisms/ Process in the solid state R1– Phase boundary controlled reaction (one-dimensional movement) R2 – Phase boundary controlled reaction (contracting area) R3 – Phase boundary controlled reaction (contracting volume) D1 – One-dimensional diffusion D2 – Two-dimensional diffusion (Valensi’s equation) D3 – Three-dimensional diffusion (Jander’s equation)

f(α) – reaction mode 1

g(α) – fraction mode α

2(1 – α) ½ 3(1 – α) 2/3 (1/2) α -1 [ -ln (1- α)]-1

[1-ln (1-α)]1/2 [1-ln (1-α)]1/3 α2 (1- α) ln (1-α) + α

(3/2) [ 1 - (1- α)1/3]-1 (1- α)2/3 1-α

[1-(1-α)1/3]2

F1 (First-order) – Random nucleation with one nucleus on the individual particle F2 (Second-order) – Random nucleation with two nuclei on the (1 - α)2 individual particle F3 (Third–order) – Random nucleation with three nuclei on the (1/2) (1 - α)3 individual particle

-ln (1- α) 1/(1- α) 1/(1- α)2

MATERIALS AND METHODS Materials Softwood Klason lignin from Pinus taeda and hardwood Klason lignin from Eucalyptus grandis were prepared as follows. Firstly the extractives were eliminated from wood via Soxhlet extraction in triplicate using: ethanol/benzene; ethanol; and hot water, in accordance with the Tappi T204 cm-97 standard. Afterwards the Klason lignin was obtained in accordance with the Tappi T222om-02 standard. The lignin purification procedure occurs after extensively washed the samples with deionized water until obtain a filtrate with pH equal to 7. The samples were dried in vacuum oven at 105°C for 4h before the tests.

Thermogravimetric Analysis (TGA) The thermogravimetric analysis (TGA50 – Shimadzu) was carried out under N2 atmosphere with a purge gas flow of 50 cm3.min-1 and at temperature from 25 to 800ºC.

Thermal Characterization of Klason Lignins from Softwood …

249

Approximately 10 mg of each sample was used. To calculate the kinetics parameters the TGA was carried out at four different heating rates (5, 10, 20 and 40°C.min-1).

Fourier Transform Infrared Spectroscopy (FTIR) FTIR spectra were obtained using a Nicolet IS10- Thermo Scientific spectrometer. Samples with 5 mg were dispersed in 100 mg of KBr followed by compression to form pellets. The sample collection was obtained using 32 scans, from 4000 cm-1 to 400 cm-1, at a resolution of 4 cm-1. Second derivative spectra were obtained by applying the SavtzkyGolay function [12] in FTIR spectra. Care was taken to ensure all samples remained dry during sample preparation and FTIR analysis, the sample for FTIR analysis were removed from the vacuum oven only in the moment of the test.

RESULTS AND DISCUSSIONS TGA Analysis Figure 2 shows the TGA and DTG curves for pyrolysis of the lignins studied. As shown in the thermogravimetric curve, the thermal degradation of both lignins proceeded over a wide temperature range from approximately 180°C to 800°C. This can be explained by the fact that lignin contains many aromatic rings with various branches with different activities of its chemical bonds and functional groups [7]. In the first degradation stage between 30100ºC the weight loss was less than 4% and can be mainly attributed to the loss of moisture [7, 24] present in the samples. After the first weight loss, the degradation process is slower between 110-180°C and for LPIT a plateau can be seen in this temperature range. As shown in Figure 2, the LPIT thermal stability was higher than that of LEUG. The initial weight loss temperature (Ti) considered in this work as the temperature at which the sample loses 10% of its weight is 8°C higher in LPIT than in LEUG, as presented in Table 2. Table 2. Thermal degradation temperatures, DTG peak and % residue for the lignins studied Lignin samples LPIT LEUG

Ti (°C) 10 wt% loss 300 292

DTG peak (°C) 389 378

Residue at 800°C (%) 46.3 48.8

250

Matheus Poletto

Figure 2. Thermogravimetric curves for both lignins studied.

Figure 3. Plot of the straight lines using the Kissinger method.

It is in the second stage, between 180-500°C, the main lignin degradation process occurs. The DTG peak occurs at 378°C for LEUG, while for LPIT the peak is centered at 389ºC. Pyrolytic degradation in this region involves fragmentation of inter-unit linkages, releasing monomeric phenols into the vapor phase [25, 26], that may accelerate the degradation process. Above 500°C the degradation process is possibly related to the slow decomposition of some aromatic rings in lignin [24, 26]. At 800°C almost 50% of all lignin samples still remain unvolatilized due to the formation of highly condensed aromatic structures [26]. The kinetic parameters for the lignins studied were calculated using the Kissinger’s method. The maximum temperature (T m) at different heating rates was obtained from the DTG curves. In addition, ln(β/ Tm) was plotted against 1/T for each lignin sample for obtained the activation energy and frequency factor, as can be seen in Figure 3.

Thermal Characterization of Klason Lignins from Softwood …

251

From Table 3 it can be seen that the activation energy for LPIT is higher than that for LEUG. On the other hand, it is interesting to compare the activation energy obtained in this study with the values found in the literature. In this study the activation energy values for the Klason lignin were in the range of 158-166 kJ/mol, while the values found in the literature for the Klason lignin varied from 12.5-89 kJ/mol [27-29]. This great difference may be caused by erroneous assumptions of the first order reaction by other authors. As presented in Table 2 the order of reaction for the Klason lignin for both the softwood and hardwood species studied was 1.5. Jiang et al. in a recent study also obtained an order of reaction of 1.5 for different wood lignins isolated using the Klason method [8]. The frequency factor is higher in LPIT than in LEUG. Frequency factor is a reflection of the frequency of the collisions between the reacting molecules [8]. Table 3. Kinetic parameters obtained for both lignins studied Lignin samples LPIT LEUG

Ea (kJ/mol) 165.69 158.43

n 1.42 ± 0.07 1.48 ± 0.02

Frequency factor 8.26 x 1011 2.86 x 1011

Figure 4. Activation energy values for lignins studied.

Figure 4 shows the activation energy values obtained through the FWO method for both lignins studied. It can be observed that LEUG present higher Ea values than LPIT. The evaluation of conversion values also reveals a different behavior for lignins studied. For LEUG when α is equal to 0.1 only the loss of moisture occurs, because the lower value of Ea. For LPIT the degradation rate is slower than in LEUG and the loss of moisture occurs between conversion values range of 0.1-0.2. For LEUG the main lignin degradation occurs between 0.2-0.6, while for LPIT the main lignin degradation occurs between 0.3-0.6. In this stage several degradation reactions occur, involved different Ea values.

252

Matheus Poletto

According to Faravelli et al. (2010) [30] the first relevant radical initiation reaction in lignin involves the scission of C-O bond in the β-O-4 lignin structure forming phenoxy radicals. A bond energy of about 242.8 kJ.mol-1, as proposed by Back (1989) [31], allows the activation energy for this reaction in the gas phase. However, Ea values between 176188 kJ.mol-1 can also be obtained [8]. For LEUG the Ea values are very close to the maximum value (242.8 kJ.mol-1) when for LPIT between 0.3-0.4 the Ea values are more close to the minimum range values. The scission of phenylether compounds present in lignin generally involves activation energy values at around 300 kJ.mol-1 [15]. For LEUG this step occurs between 0.4-0.5, however for LPIT this step only occurs at conversion value equal to 0.6, probably because the slower conversion rate in this sample, which may be associated with the higher thermal stability of the guaiacyl units. When the conversion values are between 0.6-0.7 the degradation process initiate a different pathway. In this stage the degradation of the highly condensed aromatic structures formed takes place in both lignins and both Ea values and frequency factor are very different from the previous stages. Additionally, the conversion range between 0.7-0.9 may be associated with degradation occurring in a narrow and accelerated range of degradation and another degradation mechanism could be occurs at higher conversion values [32, 33]. The Ea values obtained using the FWO method were used to determine the thermal degradation mechanisms proposed by [22]. This method uses reference theoretical curves obtained from Eq. (8) that are derivatives of the f(α) and g(α) functions represented in Table 1; called master curves, they are compared to experimental data to determine the mechanism of the solid-state degradation process [22]. As can be seen in Table 1, the algebraic expressions that represent the theoretical mechanisms were separated into three main groups: Rn, Dn and Fn. Respectively, these mechanisms describe: phase boundary controlled reaction; diffusion processes that are related to the heat transfer capacity along the material structure; and the random degradation of nuclei. The determination of the Z(α) values was carried out using a heating rate (β) of 10ºC.min-1, and the calculated Z(α) values were determined by applying the Ea values obtained using the FWO method. Figure 5 presents the master curves as well as the results of the experimental data obtained. When α = 0.1 the experimental value are clored to the diffusion curves. These degradation mechanisms refer to a diffusion process in one, two or three dimensions, respectively [22]. When the conversion values are in the range of α = 0.2-0.6, the experimental values did not overlap any theoretical curve. The experimental values are more closely related with a D3 mechanism and the shape of the experimental curve is also similar to a D3 mechanism in this α interval, but the experimental values did not overlap the D3 theoretical curve. This result indicate that in the range of α = 0.2-0.6 the lignin degradation process did not follow any degradation mechanism related to Criado method. When the conversion values are in the range of 0.7-0.9 the shape of the experimental curve for both lignins studied present a different trend. The experimental points are furthest from the theoretical values indicating a different thermal degradation behavior at this interval.

Thermal Characterization of Klason Lignins from Softwood …

253

As discussed earlier, in this stage the degradation of the highly condensed aromatic structures formed during the previous stages takes place leading to slow decomposition reactions of these aromatic compounds.

Figure 5. Master curves and experimental curves for the lignins studied.

In summary, the differences in the structures and chemical natures of lignin from different tree species could account for the diversity in their thermal degradation behaviors. The syringyl as well as the guaiacyl units are built into the lignin macromolecule mainly by ether bonds, and the ether bonds between syringyl units are easier to split than those between guaiacyl units [34]. This may contribute to higher thermal stability and higher activation energy for softwood lignin that consists largely of guaiacylpropane units (G units) than for hardwood lignin that is basically composed of both guaiacyl and syringylpropane units (S units) [7]. Noncovalent interactions, such as hydrogen bonds, should also affect the thermal properties of lignin [4].

FTIR Analysis Figure 6 shows the FTIR spectra of the lignins. The peak positions of the several bands in the FTIR spectra are summarized in Table 4. The spectral differences between both lignin samples are observed in the fingerprint region between 1800 and 900 cm-1. The LPIT sample showed typical bands of softwood lignins. A prominent band at 1264 cm-1 assigned to C-O of the guaiacyl ring, a band at 1143 cm-1 related to aromatic C-H in-plane deformation in the guaiacyl ring, a 1088 cm-1 band assigned to C-O deformations of secondary alcohols and aliphatic ethers and a band at 871 cm-1 for LPIT and 912 cm-1 for LEUG assigned to C-H out of plane in positions 2, 5 and 6 (G units) [4, 7, 26]. Softwood lignin, often referred to as guaiacyl lignin is mainly composed of coniferyl alcohol units, which make up more than 95% of the structural units

Matheus Poletto

254

in this lignin, the remainder consisting mainly of p-coumaryl alcohol type units [7]. On the other hand, the LEUG sample presented similar bands for hardwood lignins at 1309 cm-1 related to C-O of the syringyl ring and a band at 1108 cm-1 assigned to aromatic C-H deformation in the syringyl ring [4, 7]. Hardwood lignin is composed of coniferyl alcohol and sinapyl alcohol derived units in varying proportions, commonly referred to as guaiacylsyringyl lignin [7].

Figure 6. FTIR spectra of the lignin samples evaluated.

Table 4. Bands observed for the lignins samples LPIT Band position (cm-1) 3396 2935 2843 1706 1663 1605 1497 1464 1427 ----1264 1211 1143 ----1088 1028 871

LEUG Band position (cm-1) 3405 2938 2841 1708 1664 1608 1497 1463 1425 1309 ----1217 ----1108 ----1023 912

Assignment O-H Stretching C-H Stretching C-H Stretching C=O Stretching Unconjugated C=O Stretching Conjugated Aromatic skeletal vibration plus C=O Stretching Aromatic skeletal vibration C-H Deformation C-H in-plane deformation with aromatic ring stretching C-O of the syringyl ring C-O of the guaiacyl ring C-C plus C-O Stretching typical of G units Aromatic C-H in plane deformation in the guaiacyl ring Aromatic C-H deformation in the syringyl ring C-O deformations secondary alcohols and aliphatic ethers Aromatic C-H in plane deformation (G>S) C-H out of plane in positions 2,5 and 6 (G units)

Thermal Characterization of Klason Lignins from Softwood …

255

The ratio between the bands intensities at 2935, 1264 and 871 cm-1 for LPIT and between the bands intensities at 2938, 1309 and 912 for LEUG were used for determined differences in the guaiacyl and syringyl groups quantities presents in the two lignins studied. The results are presented in Table 5. The prominent differences in the quantities of guaiacyl and syringyl groups can be found in the both lignins. In LPIT that is mainly composed of coniferyl alcohol units has larger quantity of G units when compared with LEUG, as can be seen in Table 5. On the other hand, LEUG is composed by a mixture of coniferyl alcohol and sinapyl alcohol, however with several S units in its structure. In addition, a prominent shoulder for both lignins at around 3600 cm-1 can be seen in Figure 5. According to Kubo and Kadla (2005) [4], in lignin the hydroxyl groups in alcoholic and phenolic compounds form several intermolecular and intramolecular hydrogen bonds between 3640-3520 cm-1 [4]. However, these bands can be better identified from the second derivative spectra of the FTIR spectra [4]. Two bands assigned to free hydroxyl groups in alcoholic groups occur in the region between 3640-3616 cm-1; between 3560-3550 cm-1 other bands related to intramolecular hydrogen bond in phenolic groups occur and at 3520-3480 cm-1 a band assigned to intermolecular hydrogen bond also occurs [4, 35]. Table 5. Differences in the guaiacyl and syringyl groups quantities presents in the both lignins studied LPIT I1264/I2935 1.42 ± 0.06 G units

I871/I2935 0.26 ± 0.03 G units

LEUG I1309/I2938 1.70 ± 0.02 S units

I912/I2938 0.01 ± 0.003 G units

Table 6. Bands position and wavenumber shift for the lignin bands obtained from the second derivative of the FTIR spectra Band position (cm-1) LPIT 3633 ± 3 3616 ± 2 3567 ± 7 3549 ± 2 3528 ± 4

LEUG 3630 ± 1 3612 ± 1 3563 ± 1 3547 ± 1 3521 ± 1

ΔυO-H (cm-1) LPIT 239 ± 5 222 ± 2 176 ± 6 155 ± 2 130 ± 2

LEUG 225 ± 3 209 ± 3 160 ± 3 144 ± 3 118 ± 2

According to Purcell and Drago (1967) [36] the wavenumber shift (ΔυO-H) between the free hydroxyl-stretching vibration and that of hydrogen-bonded species, related to the center of the hydroxyl band, yields a measure of the average strength of the intermolecular interactions [36]. The wavenumber shift between the free hydroxyl-stretching vibration

Matheus Poletto

256

and that of hydrogen-bonded species for the bands obtained from the second derivative curve are listed in Table 6. LPIT has higher ΔυO-H than LEUG for all bands evaluated. This result confirms that the intermolecular and intramolecular interactions between hydroxyl groups in LPIT are stronger than in LEUG. This behaviour may be associated with the higher thermal stability and higher Ea observed for LPIT by thermogravimetric analysis. Therefore, more energy is necessary to break down the hydrogen bonds formed between the lignin groups in LPIT than in LEUG and consequently there is an increase in thermal stability of LPIT sample. For phenol groups, an enthalpy-OH wavenumber shift relationship has been developed [37] in which a linear relationship between the enthalpy of hydrogen bond formation and OH wavenumber shift exists and can be expressed by Eq. 11: -ΔH (kcal/mol) = 0.016 ΔυO-H + 0.63

(11)

The ΔυO-H values obtained from the bands assigned to phenolic groups in lignin at 3565 cm and 3550 cm-1 were used to determine the enthalpy of hydrogen bonded. The hydrogen bond distances R for the two bands described above are obtained according Eq. 12, proposed by Pimentel and Sederholm (1956) [38]: -1

Δν = 4430 × (2.84 – R)

(12)

where Δν = νo – ν, νo is the monomeric stretching frequency, which is taken to be 3600 cm1 , and ν is the stretching frequency observed in the FTIR spectra of the sample. The obtained results for ΔH and R are presented in Table 7. Table 7. Enthalpy-OH wavenumber shift and hydrogen bond distances values for the lignins studied Lignin samples LPIT LEUG

-ΔH (kcal/mol) 3565 cm-1 3.45 ± 0.09 3.18 ± 0.05

3550 cm-1 3.11 ± 0.03 2.93 ± 0.05

R (Å) 3565 cm-1 2.846 2.848

3550 cm-1 2.851 2.852

The enthalpy of hydrogen bond formation between the phenolic groups is higher in LPIT than in LEUG. In addition, the hydrogen bond distances in LPIT are lower than in LEUG. The donor and acceptor atoms in the O-H ∙∙∙ O hydrogen bond in LPIT are nearer than in LEUG and so the interactions between both atoms are more intense which results in stronger intermolecular interactions. This may be better explained by the highest wavenumber shift and enthalpy-OH wavenumber found for the LPIT sample. In summary, the above result suggests that the hydrogen bond interactions between the phenolic groups in LPIT are stronger than the ones formed in LEUG and may probably cause an increase

Thermal Characterization of Klason Lignins from Softwood …

257

in the LPIT thermal stability. Other parameters were not evaluated in this work, such as molecular weight and methoxyl groups can also influence the thermal properties of the lignins. However, the effects of hydrogen bonding are an important aspect of the thermal stability of the lignins studied.

CONCLUSION Thermogravimetric results showed that LPIT is more thermally stable than LEUG. In addition, FTIR results confirm that the intermolecular and intramolecular interactions between hydroxyl groups in LPIT are stronger than in LEUG. This behaviour corroborates the higher thermal stability observed for LPIT by thermogravimetric analysis. The degradation of both lignins initiate by a diffusion process. However, when the conversion values are higher than 0.1 the results showed that the lignin degradation mechanism is a complex procedure and involves the degradation of a highly condensed aromatic structure. From the obtained results, it is possible to state that higher hydrogen bonding between lignin compounds contribute to higher thermal stability and higher activation energy of the lignins studied.

ACKNOWLEDGMENTS The authors wish to thank Cambará S/A, Madeireira Gold Martini for supplying the samples of Pinus taeda and Eucalyptus grandis, respectively.

REFERENCES [1] [2]

[3] [4]

Ferdous, D., Dalai, A. K., Bej, S. K., Thring, R. W. 2002. “Pyrolysis of lignins: experimental and kinetics studies.” Energy & Fuels 16: 1405-1412. Brosse, N., El Hage, R., Chaouch, M., Pétrissans, M., Dumarçay, S. 2010. “Investigation of the chemical modifications of beech wood lignin during heat treatment.” Polymer Degradation and Stability 95: 1721-1726. Wang, S., Ru, B., Lin, H., Sun, W., Luo, Z. 2015. “Pyrolysis behaviors of four lignin polymers isolated from the same pine wood.” Bioresource Technology 182: 120-127. Kubo, S., Kadla, J. 2005. “Hydrogen bonding in lignin: a Fourier transform infrared model compound study.” Biomacromolecules 6: 2815-2821.

258 [5]

[6]

[7]

[8] [9]

[10]

[11]

[12]

[13]

[14]

[15] [16]

[17]

[18]

Matheus Poletto Godoy, E. A., Rodrigues, J. C. C., Alves, A. M. M., Lazo, D. A. 2007. “Content and quality study of the lignin by analytical pyrolysis in Pinus Caribaea.” Maderas Ciência y tecnología 9: 179-188. Kiaei, M., Kord, B., Vaysi, R. 2014. “Influence of residual lignin content on physical and mechanical properties of kraft pulp/PP composites.” Maderas Ciência y tecnología 16: 495-503. Wang, S., Wang, K., Liu, Q., Gu, Y., Luo, Z., Cen, K., Fransson, T. 2009. “Comparison of the pyrolysis behavior of lignins from different tree species.” Biotechnology Advances 27: 562-567. Jiang, G., Nowakowski, D. J., Bridgwater, A. V. 2010. “A systematic study of the kinetics of lignin pyrolysis.” Thermochimica Acta 498: 61-66. Yasuda, S., Fukushima, K., Kakehi, A. 2001. “Formation and chemical structures of acid-soluble lignin I: sulfuric acid treatment time and acid-soluble lignin content in hardwood.” Journal of Wood Science 47: 69-72. Buranov, A. U., Ross, K. A., Mazza, G. 2010. “Isolation and characterization of lignins extracted from flax shives using pressurized aqueous ethanol.” Bioresource Technology 101: 7446-7455. El Hage, R, Brosse, N., Chrusciel, L., Sanchez, C., Sannigrahi, P., Ragauskas, A. 2009. “Characterization of milled wood lignin and ethanol organosolv lignin from miscanthus.” Polymer Degradation and Stability 94: 1632-1638. Popescu, M-C, Popescu, C-M, Lisa, G., Sakata, Y. 2011. “Evaluation of morphological and chemical aspects of different wood species by spectroscopy and thermal methods.” Journal of Molecular Structure 988: 65-72. Poletto, M., Dettenborn, J., Pistor, V., Zeni, M., Zattera, A. J. 2010. “Materials Produced from Plant Biomass. Part I: Evaluation of Thermal Stability and Pyrolysis of Wood.” Materials Research 13: 375-379. Poletto, M., Zattera, A. J. Santana, R. M. C. 2012. “Strucutral differences between wood species: evidence from chemical composition, FTIR spectroscopy, and thermogravimetric analysis.” Journal of Applied Polymer Science 126: E336-E343. Klein, M. T., Virk, P. S. 2008. “Modeling of lignin thermolysis.” Energy & Fuels 22: 2175-2182. Bährle, C., Custodis, V., Jeschke, G., van Bokhoven, J., Vogel, F. 2014. “In situ observation of radicals and molecular products during lignin pyrolysis.” ChemSusChem 7: 2022-2029. Bianchi, O., Martins, J. De N., Fiorio, R., Oliveira, R. V. B., Canto, L. B. 2011. “Changes in activation energy and kinetic mechanism during EVA crosslinking.” Polymer Testing 30: 616-624. Sanchez-Silva, L., López-González, D., Villaseñor, J., Sánchez, P., Valverde, J. L. 2012. “Thermogravimetric-mass spectrometric analysis of lignocellulosic and marine biomass pyrolysis.” Bioresource Technology 109: 163-172.

Thermal Characterization of Klason Lignins from Softwood …

259

[19] Kissinger, H. E. 1956. “Variation of peak temperature with heating rate in differential thermal analysis.” Journal of Research of the National Bureau of Standards 57: 217221. [20] Flynn, J. H., Wall, L. A. 1966. “General treatment of the thermogravimetry of polymers.” Journal of Research of the National Bureau of Standards 70A: 487-523. [21] Ozawa T. 1965. “A new method of analyzing thermogravimetric data.” Bulletin of the Chemical Society of Japan 38: 1881-1886. [22] Criado, J. M., Málek, J., Ortega, A. 1989. “Applicability of the master plots in kinetic analysis of non-isothermal data.” Thermochimica Acta 147: 377-385. [23] Pérez-Maqueda, L. A., Criado, J. M. 2000. “The accuracy of Senum and Yang’s approximations to the Arrhenius integral.” Journal of Thermal Analytical and Calorimetry 60: 909-915. [24] Zhao, X., Liu, D. 2010. “Chemical and thermal characteristics of lignins isolated from Siam weed stem by acetic acid and formic acid delignification.” Industrial Crops and Products 32: 284-291. [25] Marcovich, N. E., Reboredo, M. M., Aranguren, M. I. 2001. “Modified woodflour as thermoset fillers II. Thermal degradation of woodflours and composites.” Thermochimica Acta 372: 45-47. [26] Tejado, A., Peña, C., Labidi, J., Echeverria, J. M., Mondragon, I. 2007. “Physicochemical characterization of lignins from different sources for use in phenolformaldehyde resin synthesis.” Bioresource Technology 98: 1655-1663. [27] Domburg, G. E., Sergeeva, V. N. 1969. “Thermal degradation of sulphuric acid lignins of hard wood.” Journal of Thermal Analysis 1: 53-62. [28] López Pasquali, C. E., Herrera, H. 1997. “Pyrolysis of lignin and IR analysis of residues.” Thermochimica Acta 293: 39-46. [29] Caballero, J. A., Font, R., Marcilla, A., Garcia, A. N. 1993. “Flash pyrolysis of klason lignin in a pyroprobe 1000.” Journal of Analytical and Applied Pyrolysis 27: 221– 244. [30] Faravelli, T., Frassoldati, A., Migliavacca, G., Ranzi, E. 2010. “Detailed kinetic modeling of the thermal degradation of lignins.” Biomass and Bioenergy 34: 290301. [31] Back, M. H. 1989. “Comment on the thermal decomposition of anisole and the heat of formation of the phenoxy radical.” Journal of Physical Chemistry 93: 6880-6881. [32] Yao, F., Wu, Q., Lei, Y., Guo, W., Xu, Y. 2008. “Thermal decomposition kinetics of natural fibers: activation energy with dynamic thermogravimetric analysis.” Polymer Degradation and Stability 93: 90-98. [33] Ornaghi Jr., Poletto, M.; Zattera, A. J., Amico, S. C. 2014. “Correlation of the thermal stability and the decomposition kinetics of six different vegetal fibers.” Cellulose 21: 177-188.

260

Matheus Poletto

[34] Jakab, E., Faix, O., Till, F., Székely, T. 1995. “Thermogravimetry/mass spectrometry study of six lignins within the scope of an international round robin test.” Journal of Analytical and Applied Pyrolysis 35: 167–179. [35] Kadla, J., Kubo, S. 2003. “Miscibility and hydrogen bonding in blends of poly(ethylene oxide) and kraft lignin.” Macromolecules 36: 7803-7811. [36] Purcell K. F., Drago, R. S. 1967. “Theoretical aspects of the linear enthalpy wavenumber shift relation for hydrogen-bonded phenols.” Journal of the American Chemical Society 89: 2874-2879. [37] Joesten, M. D., Drago, R. S. 1962. “The validity of frequency shift-enthalpy correlations. I. Adducts of phenol with nitrogen and oxygen donors.” Journal of the American Chemical Society 84: 3817-3821. [38] Pimentel, G. C., Sederholm, C. H. 1956. “Correlation of infrared stretching frequencies and hydrogen bond distances in crystals.” Journal of Chemical Physics 24: 639-641.

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 11

LIGNIN AND ITS DERIVATIVES AS ANTIOXIDANT, ANTIVIRAL AND ANTIMICROBIAL AGENTS: APPLICABILITY IN HUMAN HEALTH PROMOTION Gabriela Vazquez-Olivo, Marilyn S. Criollo-Mendoza, Erick P. Gutiérrez-Grijalva, Manuel A. Picos-Salas and J. Basilio Heredia* CONACYT - Centro de Investigación en Alimentación y Desarrollo A.C., Carretera a Eldorado Km, Col. Campo El Diez, Culiacán, Sinaloa, México.

ABSTRACT Lignin is the second major component of vegetal biomass after cellulose, and mainly produced as a by-product of paper industry, which makes it relatively cheap to obtain. Generally, this biopolymer is used to produce energy and to recover inorganic chemicals, however, many efforts has been made to find new applications of lignins, in order to add extra value to them. Lignin is a heterogeneous and complex phenolic polymer, and its utilization can be challenging since its chemical structure is difficult to characterize, ergo, various methodologies have been employed to valorized lignins and their derivatives. Depending on the method of isolation, different kinds of lignins can be obtained, and they have been reported to have antioxidant, antimicrobial and antiviral activity. Additionally, recent reports provide evidence of its protective role against different diseases, which is mainly due to the antioxidant properties, and has been demonstrated to be nontoxic for humans. These reports provide information for the application of lignins in food, pharmaceutical and cosmetic industries.

*

Corresponding Author Email: [email protected]

262

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al.

Keywords: lignin, antioxidant, antimicrobial, antiviral, lignin-carbohydrate complex, lignophenols, alkali lignin, lignosulphonate

1. INTRODUCTION Lignin is the second most abundant biopolymer in nature, and it represents about 1530% (w/w) of the total lignocellulosic biomass [1]. Obtained as main by-product from pulp and paper industries, the current applications of lignin are of low added value, mainly being burnt for energy production [2-4]. Moreover, less than 2% of industrial lignin is recovered for usage as a chemical product [5]. Nevertheless, the increasing interest in the efficient use of natural resources as raw material for different applications has aroused the investigation on the development of various processes, in order to fully utilize lignocellulosic biomass, particularly lignin[6]. Lignin plays an important role in plants, acting as the ‘cement’ that holds together the lignocellulosic fibers and providing rigidity to the plant cell wall. Furthermore, it has a phenolic structure bearing one or more hydroxyl groups [7]. This phenolic structure confers to the lignin many properties, such as antioxidant acting as scavenger of free radicals [8], as well as its antimicrobial activity [9] and more recently studied its antiviral activity [10]. Moreover, the usage of lignin is challenging because of the heterogenicity of its structure and to the fact that the biological properties that are affected by the botanical origin, molecular weight and, by technical factors such as the method of extraction and isolation. In this context, several investigations have tried to overcome these drawbacks, particularly, there is an increasing interest in the potential application of lignins to improve human health [11]. Despite the fact that the use of lignin has been challenging, it is still a promising biopolymer due to its natural origin, low cost and abundance, and can be applied in food, pharmaceutical and cosmetic industries [12], besides the traditional applications, its benefit to human health and low toxicity has been proved in in vitro chemical assays, as well as in cellular and animal models. The present chapter makes a compilation from 2007 to 2017 studies concerning the application and possible utilizations of lignin and its derived compounds in food, pharmaceutical and cosmetic industries.

2. STRUCTURE AND SOURCES OF LIGNIN Lignin is one of the main constituent of dietary fiber. It is a complex hydrophobic molecule that exists in most cell walls along with heteroxylans. Lignin posses different

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents

263

chemical funcional groups such as hydroxyl, methoxyl, carbonyl and carboxyl groups in different proportions, depending on the genetic origin and the method of extraction [11]. In nature, lignin is derived from the polymerization of substituted phenyl propylene units. The building blocks of lignin are called monolignols, which in turn make the lignin subunits. The primary monolignols are p-coumaryl alcohol (4-hydroxycinnamyl), coniferyl alcohol (3-methoxy 4-hydroxycinnamyl), and sinapyl alcohol (3, 5-dimethoxy 4hydroxycinnamyl); and, hydroxyphenyl (H), guaiacyl (G), and syringyl (S) are the main lignin subunits. These molecules are synthesized as the result of the enzymatic conversion of L-phenylalanine, a biosynthetic process that is still not well understood [13-15]. Additionally, plants contain lignin subunits derived from ferulic acid, ferulates, coniferaldehyde, sinapaldehyde, 5-hydroxyconiferyl alcohol, and acylated monolignols containing acetate, p-hydroxybenzoate, or p-coumarate moieties [13, 16]. Due to the diversity of monolignols and their radical coupling with phenolic compounds, lignin structure is remarkable complex, and is nowadays still not fully known [17]. The structure and composition of lignin depends strongly on the type of biomass and even on the part of the plant, and they are relatively insoluble [16, 18]. Also, lignin structure can vary according to the taxonomical classification of their plant origin. For example, lignin extracted from gymnosperms lacks of S units whilst angiosperms are rich in G/Sunits [14, 16, 19]. Moreover, lignin composition can also be affected by the phenological stages of plants. For instance, at early stages of lignification, coniferyl alcohol with small amounts of p-coumaryl alcohols is copolymerized into the primary wall to form mixed G and H units [19]. However, during secondary wall development, coniferyl alcohol and increasing amounts of sinapyl alcohol are copolymerized to form mixed G and S units. Interestingly, it seems that lignin composition does not alter its interactions with the other components of the matrix [19-21]. Moreover, due to the distribution of the H, G and S units, lignin polymers can be classified as type-G (softwood lignin), type-G-S (hardwood lignin), type H-G-S (grass lignin), and type-H-G (compression wood lignin) [16]. Apart of the wood, lignin is present in different vegetative wastes such as straws of rice, wheat and corn, sugarcane bagasse, tea residues and bamboo culm, among others.[22, 23]. According to the extraction process, different types of lignins are identified, however, due to the heterogenicity of the raw material, until now, there is no standardized method for the extraction of lignin without causing modifications to its structure. Some examples of industrially obtained lignins are: lignosulfonates, which are the by-products of sulfite pulping; Kraft lignins or sulfate lignins, obtained from the Kraft pulping process; soda lignin, extracted from the black liquor of soda pulping process; Klason lignin, which is the component of wood or pulp that is insoluble in 72% sulfuric acid; organosolv lignin, obtained as a by-product from the process that uses organic solvents to solubilize lignin [11, 24, 25].

264

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al.

3. METHODS OF ISOLATION OF LIGNIN AND LIGNIN FRACTIONS 3.1. Acid Precipitation Acid precipitation is the most widely method used to fractionate and isolate lignin. Commonly, acid precipitation of lignin is carried using different inorganic acids like phosphoric acid, hydrochloric acid and nitric acid, and, sulphuric acid, but, the latter is the most used for this purpose [26]. The precipitation process consists of adding concentrate acid to a lignin solution (liquor) obtained from the pulping process until the desired pH is reached (pH around 1-2). After that, it is necessary to wait until lignin sedimentation, following centrifugation and washing with distilled water to remove impurities (at this point, some researchers perform a second centrifugation but it is not always necessary), and, finally the lignin dries either by air-dry and/or oven dry [27-30]. Furthermore, vacuum filtration could be used instead of centrifugation [26] and the obtained lignin can be dissolved and precipitated once again by the same procedure of acid precipitation [31].

3.2. Membrane Filtration Membrane filtration is a very common method of isolation and fractionation of lignin and this technique includes ultrafiltration and nanofiltration. This process uses membranes with different molecular weight cut-off (MWCO) to separate and fractionate lignin according to its molecular weight (MW). As compared to others methods, ultrafiltration yield lignins with higher purity. For instance, high molecular weight lignin-carbohydrate complex with high purity, has been previously isolated from sodium based spent sulfite liquor by ultrafiltration [32]. Moreover, lignin fractions obtained by membranes with MWCO bigger than 15 kDa tend to be highly contaminated, probably with lignincarbohydrate complexes, compared to low MWCO [33]. Additionally, the fractionation of Kraft lignin showed that fractions with less molecular weight are prone to contain fewer aliphatic hydroxyl groups and more phenolic hydroxyl groups [34]. On the other hand, ultrafiltration allows to achieve fractions of lignin with fewer contamination compared to differential precipitation, which usually yield lignin with hemicellulose and silicates. Likewise, lignin fractions obtained by membranes with MWCO bigger than 15 kDa are highly contaminated, probably with lignin-carbohydrate complex [35].

3.3. Solvents Lignin of high purity level can be obtained using dioxane as an extraction solvent [36], furthermore, lignin can be fractionate using other organic solvents. After enzymatic

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents

265

hydrolysis of the lignin material, extraction with aqueous dioxane and subsequent extraction with dichloromethane, acetic ether and n-butyl alcohol, showed that the former fraction had the lowest MW lignin, while the later fraction had the highest MW lignin. Additionally, these methods produces lignin with more than 90% of purity, removing most of the glucose in the matrix. However, the method do not remove other sugars like xylose and arabinose, which are components from hemicellulose, and are very difficult to eliminate [37]. Furthermore, to eliminate hemicellulose is common to use aqueous ethanol precipitation yielding a more pure lignin, notwithstanding, this method have the drawback that some carbohydrates can be remaining [38, 39]. Besides, the use ethanol and sulfuric acid on organosolv lignin, enhanced with microwave irradiation, produce lignin with high purity (low in ash, sugars and sulfur) [40]. Similarly, fractionation of Kraft lignin by sequential precipitation using organic solvents with decreasing solubility displayed structural differences among them. Furthermore, high MW lignin and polydispersity was shown in the first fraction and low MW and polydispersity, along with high free hydroxyl and methoxyl groups content were shown in the subsequent fractions [41]. Another way to isolate lignin fractions is dissolving softwood Kraft lignin in a polar solvent, like acetone, and adding a miscible-non polar solvent to precipitate the lignin like hexane [42]. In addition, it has been demonstrated that organosolv lignin solubility in acetone depends on its concentration; for instance, with acetone at a concentration curan 100 (46%) > steam explosion (25%) > lignosulfonate (11%). The authors suggested that lignins from different sources possessed potential health benefits which opens a new industrial application for them. The same researchers investigated the antioxidant capacity of lignins from different sources (bagasse, lignosulfonate, curan 100, and steam explosion) by measuring the inhibition of human erythrocyte hemolysis induced by AAPH (2,2’azobis (2-amidinopropane)) [24]. The inhibition was concentration dependent for all lignins. Furthermore, the lignin obtained from bagasse showed the highest inhibitory effect (100%) at the concentration of 100 μg/mL, also, the IC50 was similar to the positive control epicatechin with values of 44.9 and 42.27 μg/mL respectively. Moreover, the lignins did not shown any irritation when applied on eyes and skin (0.1 and 0.5 g, respectively). The authors concluded that additional studies are needed in order to elucidate the association between the chemical structure of lignins and their antioxidant activity. Toh, Yokoyama, Noda and Yuguchi [23], analyzed the antioxidant capacity of lignin obtained from the residues of green tea leaf in terms of its effect on the autoxidation of linoleic acid. Compared to the commercial antioxidants α-Tocopherol and t-butylhydroxyanisole (BHA), green tea lignin showed low effect by suppressing oxidation near to 50% relative to linoleic acid alone at a concentration of 1.25 x 10-3 (w/w). The effect on the antioxidant properties of lignin from different botanical origins obtained by the use of different solvents has been demonstrated. Bhat, Khalil and Karim [48], studied lignin obtained from black liquor generated from the extraction of palm oil. The radical scavenging activity against the well-known scavenger power of 1,1-diphenyl2-picrylhydrazyl (DPPH) was tested in lignins dissolved in two solvents (DMSO and 2methoxy ethanol). Lignin dissolved in 2-methoxy ethanol showed the highest percentage of inhibition (84.2%) at a concentration of 50 μg/mL. The authors suggested that the lignin

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents

267

used in this study was more easily soluble in polar solvents which influence its antioxidant capacity. The lignin obtained from Acanthopanax senticosus remainder extracted by two different organosolv methods was investigated [49]. The results showed that the acetic acid–water method was the most effective for the free radical scavenging activity, and the radical scavenging capacity against DPPH had a value of IC50 = 0.6587 mg/ml. García, Toledano, Andrés and Labidi [7], used lignin fractions from black liquor resulting from different fractionation and autohydrolysis process of Miscanthus sinensis (organosolv, autohydrolysis and alkaline lignin). The antioxidant capacity was measured by DPPH method, and the values obtained were in the following descending order: organosolv lignin > autohydrolysis lignin > alkaline lignin. The authors concluded that the presence of nonlignin components such as hemicelluloses, can decrease the antioxidant capacity of lignins, because carbohydrate can interfere in its antioxidant potential by generating hydrogen bonds with lignin phenolic groups. Also, the authors stated that the process of extraction can influence the antioxidant activity of the lignin obtained from the same origin. Likewise, Aadil, Barapatre, Sahu, Jha and Tiwary [8] analyzed the radical scavenging activity against DPPH, 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS), the ferric reducing power (FRAP), the antioxidant power (phosphomolybdate assay), the reducing power and the scavenging activity of H2O2 (HPS-CUPRAC assay) of fractions of lignin from Acacia wood extracted by alkali, hot water and organosolv (by different solvents) methods. The results showed that lignin extracted with acetone possessed the highest values in most assays. This extract displayed a radical scavenging activity against DPPH value of 93.85%, a FRAP of 294.16 μmol, an antioxidant power value of 342.22 mmol/100 mg of ascorbic acid equivalents, and a reducing power value of 196 μg/mL at a concentration of 250 μg/mL. On the contrary, in ABTS assay, ethanol extracted lignin showed the highest inhibition (95.35%) after the commercial lignin and in HPS-CUPRAC assay, propanol extracted lignin showed the highest value (EC50 = 101.8 μg/mL) at a concentration of 200 μg/mL. The authors concluded that the organosolv lignin fractions possess more free phenolic hydroxyl groups and low MW which act as electron donators. Salleh, Ismail and Ibrahim [50] studied soda oil palm empty fruit bunch (EFB) lignin, the results showed that EFB lignin strongly inhibited 4- methylumbelliferone glucuronidation in rat liver microsome through mixed-type inhibition. Sa’don, Rahim, Ibrahim, Brosse and Hussin [51], compared the antioxidant properties of modified autohydrolyzed ethanol organosolv lignin (AHC EOL), and unmodified autohydrolyzed ethanol organosolv lignin (AH EOL) in terms of reducing power, this assay describes the reduction of Fe3+ to Fe2+ owing to the presence of reducers (antioxidants). The results showed that the modification by incorporating m-cresol in lignin matrix improved its antioxidant properties. The authors implied that modified lignin had more reactive sites which increased the scavenging potential of their phenolic structures on oxygen-containing reactive free radicals. Also, the authors suggested that a high content of syringyl units conferred a better reducing power ability to lignin.

268

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al.

Since lignin is a very complex heteropolymer, and its antioxidant activity varies according to several factors, some authors have attempted to improve its antioxidant properties through nanotechnology. Lu, Liu, Yang, Zu, Zu, Zhu, Zhang, Zhang, Zhang, Sun, Huang, Zhang and Li [49], evaluated the free radical scavenging activity, the reducing power and the superoxide radical scavenging activity of nanoparticles of organosolv lignin, also they compared the values with non-nanoscale lignin. The nanoscale lignin showed a remarkable better free radical scavenging activity than the non-nanoscale (IC50 = 2.70 and 32.21 mg/mL respectively). Also, the reducing power was stronger with IC50 values of 1.41 and 15.49 mg/mL for nano-scale and non-nanoscale lignin respectively. Likewise, the superoxide radical scavenging activity was stronger. The authors suggested that the improving in the antioxidant potential was due to the solubility of nanoscale lignin, which was 12.4 times higher than non-nanoscale lignin. Also, some studies have reported the potential of the incorporation of lignin in nanoparticles and poly (lactide) films as a way in which it can be used in food, pharmaceutical and cosmetic industries. Interestingly, this has resulted in a higher antioxidant activity than its parent molecules [47, 52, 53]. Alkali lignin is a “co-product of biofuel and the paper industry which is separated from fibers by chemical pulping but is less valuable than the primary products” [17]. Aiming to use lignin antioxidant properties in food, pharmaceutical and cosmetic industries, some studies have focused on the incorporation of alkali lignin into nanoparticles with promising results, as it has been shown lignin nanoparticles have higher antioxidant capacity and better UV protection than its parent molecules [47, 52, 53]. Sun, Cao, Xu, Sun and Jones [54], have studied the antioxidant potential of Phyllostachys pubescens lignin fractions obtained by steam explosion combined with alkali and alkaline ethanol delignification. All the extracts showed antiradical activity by DPPH and ABTS assays, and ferric reducing power by FRAP assay. Nonetheless, there were a lower response by the DPPH assay in lignins extracted with methanol; and the highest antioxidant potential was shown in alkali extracted lignins, the authors suggested that the components that contribute to antioxidant capacity might not be soluble in methanol, also, the high response in alkali extracted lignins could be due to the higher content of phenolic OH in its structure. Suzuki, Matsushita, Imai, Sakurai, Henriques de Jesus, Ozaki, Finger and Fukushima [55], evaluated the antioxidant activity in terms of radical scavenging activity against DPPH and superoxide dismutase-like activity in methanol and alkali extracts from three Amazonian woods (from heartwood, and sapwood). The levels of antioxidant activity were related to the phenolic content, where heartwood showed the highest phenolic content as well as a high radical scavenging activity. Also, the superoxide dismutase-like activity was higher in heartwood samples. Structural and functional modifications in lignin may lead to significant changes in its antioxidant capacity. For example, alkali lignin depolymerization may produce compounds with different reduction properties, limiting their free radical scavenging properties [56]. Also, alkali lignin with high content of p-coumaric to ferulic acid ratios and syringyl to

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents

269

guaiacyl ratios has been associated with higher antioxidant capacity by in vitro chemical methods [54, 55]. Additionally, during the extraction process of alkali lignin, hydroxycinnamic acid derivatives like p-coumaric and ferulic acids can be released, thus reflecting a high antioxidant capacity, due to the phenolic hydroxyl groups in those molecules [54, 57]. In conclusion, the different types of lignin have a potential antioxidant capacity on human health that is worth to keep studying; moreover, lignin can exert its antioxidant capacity due to the free phenolic hydroxyl groups which act as electron donators, also the modification of lignin through the incorporation of m-cresol in lignin matrix enhance its antioxidant property. Additionally, lignins with high content of syringyl units have been reported to have a better reducing power. Overall, the use of lignin and its derivatives as antioxidant additives is of economic importance due to the added value to these molecules obtained as by-products. Further in vitro and in vivo studies are needed to evaluate the bioavailability, pharmacokinetics and pharmacological properties of these molecules on human health and its mechanisms of action.

4.2. Antiviral Effects of Lignin Bioactive compounds from plant origin have shown promising antiviral activities. Lignin, as a natural product, has been proposed as a natural source of antiviral drugs and a adjuvant in the treatment of viral infections. This biopolymer has been reported mainly for its anti- human immunodeficiency virus (HIV), anti-herpes simplex virus (HSV)-1 and anti influenza activity. The HIV is a virus that damages the immune system and weakens defense systems against infections and some types of cancer, causing that infected person acquired immune deficiency syndrome. There are two types of HIV virus: HIV-1 and HIV-2, the main difference is their pathogenicity, being the most infectious the HIV-1 [58, 59]. As stated earlier, lignin is present in many natural sources. Studies on the extract rich in lignin from Sasa senanensis Rehder, found that it has a protective effect against HIV-1 and influenza infection towards cultured cells. The authors suggested that this effect might be due to the lignin fraction present in the extract [60]. Likewise, commercial products from Sasa senanensis Rehder showed a good anti-HIV-1 activity. Product A containing 100% pure extract of Sasa senanensis Rehder with Fe (II) –chlorophylline, while products B and C consisted of Cu (II) - chlorophyllin and a minor amount of lignin-carbohydrate complexes (LCC), besides, product C was supplemented with ginseng extract and pine leaf extract. The Product A demonstrated to have 4 to 5-fold higher anti-HIV activity that the other commercial products, which was attributed to the LCC and Fe (II)-chlorophyllin complex present in the product [61]. Lignin fractions from mulberry juice showed good anti-HIV-1 activity, although the lignin shows lower value than the commercial anti-HIV agents

270

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al.

(dextran sulfate, curdlan sulfate, 3′-azido-2′, 3′-dideoxythymidine and dideoxy-cytidine). The authors suggested that the LCC was responsible for the anti-HIV-1 activity [62]. Some mechanisms have been proposed for the antiviral property of lignin, such as the suppression of the HIV-1 gene expression via inhibition of the NF-kB activity, and also, the β-5 units have shown to be more effective in this inhibition than the other units present in lignin [63]. Moreover, it has been demonstrated that the antiviral effect of lignin is modified by the method of extraction and the MW of lignin, likewise it has been demonstrated to be component specific [63]. Although, the effect of lignin in the inhibition of virus infection has been studied, there is a lack of research regarding the mechanism of action of lignin against viruses. HSV is another virus whose infection has been treated with lignin rich extracts. This virus is a common human pathogen that causes highly contagious infections affecting between 60% -95% of human adults, moreover, it is the main cause of mortality in transplant recipients by combination of the direct cytopathic effect of viral replication and indirect immunopathological mechanism dependent on the host [64]. The in vitro and in vivo anti-HSV activity has been reported for Prunella vulgaris. A fraction rich in LCC was isolated from this natural source. The results showed activity against HSV-1 and HSV-2. The effects observed could be due to the prevention of the fraction to the HSV-1 binding to Vero cells. Moreover, this fraction also demonstrated to inhibit viral penetration into Vero cells, which are key steps on the HSV infection cycle. Additionally, in the in vivo assay, the LCC fraction showed good results for skin and genital infection, however, the isolation needs to be optimized to be economically viable [65]. Raghuraman, Tiwari, Zhao, Shukla, Debnath and Desai [66] reported the anti-HSV-1 activity of sulfate lignin and found that the inhibition was proportional to the MW of this polymer. Additionally, the authors suggested that the antiviral effect of lignin sulfate may be due to the resemblance of this polymer to the heparin sulfate, a cellular receptor which recognize viral glycoproteins. The antiviral activity of lignin-carbohydrate protein complexes isolated from the seeds of Pimpinella anisum was tested against eight viruses (HSV-1 and -2, measles, mumps, poliovirus, coxsackie, human coronavirus, and human cytomegalovirus). All the isolated fractions showed potent antiviral activity. The authors suggested that the complexes might be exerting its effect at the virus adsorption and penetration into host cells which are early stages of virus replication. Additionally, high MW lignins appear to be more effective at interfering with virus adsorption and/or penetration [10]. Thakkar, Tiwari and Desai [67], suggested that the synthesized non-sulfated carboxylated lignin is a more efficient inhibitor of HSV-1 than sulfate lignin. In this study, the synthesized non-sulfated carboxylated lignin inhibited the HSV-1 entry into HeLa cells.

Table 1. Review of biological activities of lignins an its derivatives Lignin source/Lignin derivative Bagasse, lignosulfonate, curan 100, and steam explosion lignin Bagasse, lignosulfonate, curan 100, and steam explosion lignin

Biological Activity Cytotoxic

Effect Cytotoxicity at very high concentrations to HaCaT and 3T3 mouse fibroblast cell lines

Reference [47]

Antioxidant

Inhibited lipid peroxidation in RBCs Inhibited human erytrocyte hemolysis induced by AAPH in a concentration dependent manner Suppress oxidation of linoleic acid Radical scavenging activity against DPPH Radical scavenging activity against DPPH Radical scavenging activity against DPPH Radical scavenging activity against DPPH, ABTS and H2O2, ferric reducing power, antioxidant and reducing power Inhibited 4- methylumbelliferone glucuronidation in rat liver microsome through mixed-type inhibition Reducing power Radical scavenging activity against DPPH Radical scavenging activity against DPPH, ABTS, and ferric reducing power Radical scavenging activity against DPPH and, super oxide dismutase-like activity

[11] [24]

Radical scavenging activity against DPPH, reducing power and superoxide radical scavenging activity Protected the T-cell leukemia cells (MT-4) from de cytopathic effect of HIV infection Anti-influenza activity by protecting MDCK cells against influenza virus infection Inhibited transcription from HIV-1 5’-long terminal repeat (LTR). Anti-HIV activity Anti-HIV activity Inhibited the binding of HSV-1 to Vero cells and the penetration. Inhibited the cellular entry of HSV-1 and HSV-2 in a concentration dependent manner Showed antiviral activity against four viruses (HSV-1, HSV-2, human cytomegalovirus and measles virus) Inhibited HSV-1 entry into HeLa cells.

[68]

Residues of green tea leaf Black liquor from oil palm extraction Acanthopanax senticosus reminder Miscanthus sinensis Acacia wood Soda oil palm EFB Autohydrolyzed ethanol organosolv lignin Leucaena leucocephala stems Phyllostachys pubescens Amazonian woods (methanol and alkali extracts) Nanoscale lignin Sasa senanensis Rehder Organosolv and high-boiling solvent lignin Mulberry juice Leaves from Sasa senanensis (LCC) Prunella vulgaris (LCC) Sulfated lignin Lignin-carbohydrate protein complexes isolated from the seeds of Pimpinella anisum Non-sulfated carboxylated lignins

Inhibition of drug metabolizing enzymes Antioxidant

Antiviral

[23] [48] [49] [7] [8] [50] [51] [52] [54] [55]

[60] [60] [63] [62] [61] [65] [66] [10] [67]

272

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al.

From the above studies, it can be concluded that LCC seems to be an important structure in the antiviral property of lignin, also, lignin has shown to be potent inhibitor of enveloped viruses and act through different mechanisms, such as inhibition of adsorption and/or penetration of the virus into the cells and the MW and linkages between the polymer are also important. Future studies are needed regarding the structural characteristics of the LCC to exert the antiviral activity. The summary of the biological activities mentioned previously of lignins from different origins, is shown in Table 1.

5. ANTIMICROBIAL FILMS APPLICATIONS One of the major problems that food industry has to overcome is food preservation. For this reason, the search for materials with characteristics such as durability and bioactive properties become important [69, 70]. Nowadays, several researches have focused on the search for compounds with antimicrobial properties in order to avoid premature food spoilage. Furthermore, the environmental concerns have increased the interest in natural compounds in order to replace the chemical additives. In this context, lignin, which is a rich source of phenolic antioxidants, has demonstrated to be a good inhibitor of the growth of some bacteria. Lignin has been combined with some compounds such as chitosan or alginate, in order to develop composite films with greater antimicrobial properties. For instance, Rai, Dutta and Mehrotra [9] developed a biodegradable antimicrobial lignin-chitosan composite film, and 200 µg of this material demonstrated antimicrobial activity against the food pathogens Bacillus subtilis and Pseudomonas aeruginosa. Furthermore, the antibacterial activity of the composite was greater than the lignin or chitosan tested alone. The authors suggested that the inclusion of lignin in the composite improved its antimicrobial properties due to the hydrophobicity, lipophilicity and capability of this polymer to bind with proteins in an acid environment, which is an important characteristic since the bacterial cell walls are rich in lipoproteins and phospholipids. Lignin has shown to enhance the light barrier and antimicrobial properties of alginate when mixed with this hydrophilic polysaccharide in a ratio of lignin: alginate of 30:70, which could be useful for preventing lipid oxidation in food system. Moreover, the antimicrobial properties are dependent of the origin and extraction procedure of lignin. For instance, the lignin extracted from pine cone has displayed a good antimicrobial property due to the alkenic double bonds, methoxy groups and a coumaryl type lignin structure [30]. Due to the properties observed in lignin, it has been added as a nanoparticle in different blends. For instance, the addition of lignin nanoparticles (LNP) to polyvinyl alcohol (PVA) and polylactic acid (PLA), chitosan (CH) and cellulose nanocrystals (CNC) to develop antimicrobial film has been reported to have antimicrobial properties. The gram-negative

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents

273

bacteria Xanthomonas arboricola pv. pruni and Pectobacterium carotovorum subsp. odoriferum are susceptible to the lignin nanoparticles; particularly, CH-LNP (97-3 wt%) film showed a good antimicrobial activity against P. carotovorum subsp. Odoriferum after 3 h with values of 6.8x105 CFU/mL, compared to the initial inoculum of 1x106 CFU/mL by the liquid medium test, and the activity was effective even after 12 h; meanwhile, PVACH-LNP (87.3-9.7-3 wt%) reduced the growth of X. arboricola pv. pruni after 1, 3, 12 and 24 h with values of 4.1 × 105, 9.8 × 105, 2.6 × 107 and 9.7 × 107 CFU/mL, respectively [71]. Similarly, the film composed by PLA, LNP and CNC had activity against X. arboricola pv. pruni even after 12 h, furthermore, Xanthomonas axonopodis pv. vesicatoriaz growth was inhibited by this film composite, according to the broth medium test [72]. Likewise, Pseudomonas syringae pv. tomato growth was inhibited by films composed of PLA-LNPCNC and PLA-LNP, additionally, the CNC mitigated the effect by means of increasing the crystallinity value, reducing the diffusion mechanism, and, consequently, the antimicrobial activity. The mechanism proposed for the antibacterial activity of lignin is associated to the presence of phenolic compounds, which are molecules known for its biological properties, and the different functional groups containing oxygen in lignin structure such as –OH, -CO and -COOH [73]. Overall, it has been established, that the antimicrobial effect of the lignin depends on its origin, and it is usually related to its phenolic composition, however, these molecules has to be protected in order to maintain the antimicrobial activity, therefore, blends with other materials such as chitosan or alginate must be developed. Additionally, lignin has the capacity to damage the cell membrane causing lysis and emptying the cell interiors. For this reasons, lignin can be a good alternative to replace petrochemical based products, and its incorporation may add antioxidant and antimicrobial properties and improve UV absorption, thereby enhancing shelf life of food products [74].

CONCLUSION Understanding the biological properties of lignin is challenging, and requires a lot of resources to make its extraction with desirable characteristics. However, lignin as part of the dietary fiber complex might be more valuable than previously considered. Lignin can be obtained from by-products of food industry and from industrial crops. It is a renewable and abundant raw material. Some of its functional groups such as hydroxyl, methoxy and carboxyl groups are account for its antioxidant properties which makes it an ideal source of natural antioxidants in order to replace synthetic ones, also, its components may contribute to the prevention of viral infection. Additionally, because of the interaction of LCC with drugs, it can be developed into an effective antiviral drug. Likewise, because of the antimicrobial properties, the interest on the application of lignin in food packaging

274

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al.

has been increasing. The potential health benefits and industrial applications of lignins make them a promising research resource. Studies on the antioxidant, antimicrobial, and antiviral properties of lignin from different sources have been addressed, however, future research needs to be developed. There is a need of a better comprehensive understanding of the mechanisms of action and structural requirements of lignin and its derivatives, in order to fully understand their biological effects; also, it is imperative to complete the detailed characterization of the structures and properties of lignins from different botanical origins, which will allow to determine the behavior of lignins in different applications, and the ways in which their structures influence their biological activities. Furthermore, although, some researchers have reported the low toxicity of lignins, the study of their biological properties needs to be carried out in animal models and, in clinical studies to evaluate the in vivo pharmacokinetic behavior. Finally, many researches had opened the possibility of new applications of lignins. Certainly, the use of lignin will allow reduce the negative environmental impact caused by the industrial wastes and, also the extraction of high value-added substances.

REFERENCES [1] [2]

[3]

[4]

[5] [6]

[7]

Xu, C.; Ferdosian, F. In Conversion of lignin into bio-based chemicals and materials; Xu, C.; Ferdosian, F., Eds.; Springer: Berlin, Germany, 2017. García, A.; Spigno, G.; Labidi, J. 2017. “Antioxidant and biocide behavior of lignin fractions from apple tree pruning residues.” Industrial Crops and Products 104, 24252. Yang, A.-l.; Jiang, W.-j. 2007. “Studies on a Cationically Modified Quaternary Ammonium Salt of Lignin.” Chemical Research in Chinese Universities 23, 479-82. Dabral, S.; Hernández, J.; Kamer, P.; Bolm, C. 2017. “Organocatalytic Chemoselective Primary Alcohol Oxidation and Subsequent Cleavage of Lignin Model Compounds and Lignin.” ChemSusChem. Lora, J.; Naceur, B.; Gandini, A. Industrial commercial lignins: sources, properties and applications; Elsevier, Amsterdam, 2008. Martinez, V.; Mitjans, M.; Vinardell, M. P. 2012. “Pharmacological Applications of Lignins and Lignins Related Compounds: An Overview.” Current Organic Chemistry 16, 1863-70. García, A.; Toledano, A.; Andrés, M. Á.; Labidi, J. 2010. “Study of the antioxidant capacity of Miscanthus sinensis lignins.” Process Biochemistry 45, 935-40.

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents [8]

[9]

[10]

[11]

[12]

[13]

[14]

[15] [16]

[17]

[18]

[19]

[20]

275

Aadil, K. R.; Barapatre, A.; Sahu, S.; Jha, H.; Tiwary, B. N. 2014. “Free radical scavenging activity and reducing power of Acacia nilotica wood lignin.” International Journal of Biological Macromolecules 67, 220-27. Rai, S.; Dutta, P. K.; Mehrotra, G. K. 2017. “Lignin Incorporated Antimicrobial Chitosan Film for Food Packaging Application.” Journal of Polymer Materials 34, 171-83. Lee, J. B.; Yamagishi, C.; Hayashi, K.; Hayashi, T. 2011. “Antiviral and immunostimulating effects of lignin-carbohydrate-protein complexes from Pimpinella anisum.” Biosci Biotechnol Biochem 75, 459-65. Ugartondo, V.; Mitjans, M.; Vinardell, M. P. 2009. “Applicability of lignins from different sources as antioxidants based on the protective effects on lipid peroxidation induced by oxygen radicals.” Industrial crops and products 30, 184-87. Liu, M.; Wen, Y.; Qi, J.; Zhang, S.; Li, G. 2017. “Fine Chemicals Prepared by Bamboo Lignin Degradation through Electrocatalytic Redox between Cu Cathode and Pb/PbO2 Anode in Alkali Solution.” Chemistry Select 2, 4956-62. Vanholme, R.; Cesarino, I.; Rataj, K.; Xiao, Y.; Sundin, L.; Goeminne, G.; Kim, H.; Cross, J.; Morreel, K.; Araujo, P. et al. 2013. “Caffeoyl shikimate esterase (CSE) is an enzyme in the lignin biosynthetic pathway in Arabidopsis.” Science 341, 1103-06. Ragauskas, A. J.; Beckham, G. T.; Biddy, M. J.; Chandra, R.; Chen, F.; Davis, M. F.; Davison, B. H.; Dixon, R. A.; Gilna, P.; Keller, M. et al. 2014. “Lignin Valorization: Improving Lignin Processing in the Biorefinery.” Science 344. Buranov, A. U.; Mazza, G. 2008. “Lignin in straw of herbaceous crops.” Industrial Crops and Products 28, 237-59. Li, C.; Zhao, X.; Wang, A.; Huber, G. W.; Zhang, T. 2015. “Catalytic Transformation of Lignin for the Production of Chemicals and Fuels.” Chemical Reviews 115, 11559624. Carpita, N. C.; Ralph, J.; McCann, M. C. In Biochemistry & Molecular Biology of Plants; Buchanan, B. B.; Gruissem, W.; Jones, R. L., Eds.; Wiley Blackwell: Hoboken, NJ, USA, 2015. Vazquez-Olivo, G.; López-Martínez, L. X.; Contreras-Angulo, L.; Heredia, J. B. 2017. “Antioxidant Capacity of Lignin and Phenolic Compounds from Corn Stover.” Waste and Biomass Valorization, DOI: 10.1007/s12649-017-0028-510.1007/s12 649-017-0028-5. Grabber, J. H. 2005. “How Do Lignin Composition, Structure, and Cross-Linking Affect Degradability? A Review of Cell Wall Model Studies This paper was originally presented at the Lignin and Forage Digestibility Symposium, 2003 CSSA Annual Meeting, Denver, CO.” Crop Science 45, 820-31. He, L.; Terashima, N. 1990. “Formation and Structure of Lignin in Monocotyledons. III. Heterogeneity of Sugarcane (Saccharum officinarum L.) Lignin with Respect to

276

[21] [22]

[23] [24]

[25] [26]

[27]

[28]

[29]

[30]

[31]

[32]

[33]

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al. the Composition of Structural Units in Different Morphological Regions.” Journal of Wood Chemistry and Technology 10, 435-59. He, L.; Terashima, N. In Holzforschung - International Journal of the Biology, Chemistry, Physics and Technology of Wood, 1991; Vol. 45. Li, M. F.; Sun, S. N.; Xu, F.; Sun, R. C. 2012. “Microwave-assisted organic acid extraction of lignin from bamboo: Structure and antioxidant activity investigation.” Food Chemistry 134, 1392-98. Toh, K.; Yokoyama, H.; Noda, H.; Yuguchi, Y. 2010. “Antioxidant capacity of lignin from green tea waste.” Journal of Food Biochemistry 34, 192-206. Vinardell, M. P.; Ugartondo, V.; Mitjans, M. 2008. “Potential applications of antioxidant lignins from different sources.” Industrial Crops and Products 27, 220-23. Tappi, T. 2006. “222 om-02: Acid-insoluble lignin in wood and pulp.” TAPPI Test Methods. Minu, K.; Jiby, K. K.; Kishore, V. V. N. 2012. “Isolation and purification of lignin and silica from the black liquor generated during the production of bioethanol from rice straw.” Biomass and Bioenergy 39, 210-17. Domínguez-Robles, J.; Sánchez, R.; Díaz-Carrasco, P.; Espinosa, E.; GarcíaDomínguez, M. T.; Rodríguez, A. 2017. “Isolation and characterization of lignins from wheat straw: Application as binder in lithium batteries.” International Journal of Biological Macromolecules 104, 909-18. Domínguez-Robles, J.; Sánchez, R.; Espinosa, E.; Savy, D.; Mazzei, P.; Piccolo, A.; Rodríguez, A. 2017. “Isolation and Characterization of Gramineae and Fabaceae Soda Lignins.” International Journal of Molecular Sciences 18, 327. Nitsos, C.; Stoklosa, R.; Karnaouri, A.; Voros, D.; Lange, H.; Hodge, D.; Crestini, C.; Rova, U.; Christakopoulos, P. 2016. “Isolation and Characterization of Organosolv and Alkaline Lignins from Hardwood and Softwood Biomass.” Acs Sustainable Chemistry & Engineering 4, 5181-93. Aadil, K. R.; Prajapati, D.; Jha, H. 2016. “Improvement of physcio-chemical and functional properties of alginate film by Acacia lignin.” Food Packaging and Shelf Life 10, 25-33. Ibrahim, M. M.; Agblevor, F. A.; El-Zawawy, W. K. 2010. “Isolation and Characterization of Cellulose and Lignin from Steam-Exploded Lignocellulosic Biomass.” BioResources 5, 397-418. Al-Rudainy, B.; Galbe, M.; Wallberg, O. 2017. “Influence of prefiltration on membrane performance during isolation of lignin-carbohydrate complexes from spent sulfite liquor.” Separation and Purification Technology 187, 380-. Toledano, A.; García, A.; Mondragon, I.; Labidi, J. 2010. “Lignin separation and fractionation by ultrafiltration.” Separation and Purification Technology 71, 38-43.

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents

277

[34] Sevastyanova, O.; Helander, M.; Chowdhury, S.; Lange, H.; Wedin, H.; Zhang, L. M.; Ek, M.; Kadla, J. F.; Crestini, C.; Lindstrom, M. E. 2014. “Tailoring the Molecular and Thermo-Mechanical Properties of Kraft Lignin by Ultrafiltration.” Journal of Applied Polymer Science 131, 11. [35] Toledano, A.; Serrano, L.; Garcia, A.; Mondragon, I.; Labidi, J. 2010. “Comparative study of lignin fractionation by ultrafiltration and selective precipitation.” Chemical Engineering Journal 157, 93-99. [36] Liu, Z.; Meng, L.; Chen, J.; Cao, Y.; Wang, Z.; Ren, H. 2016. “The utilization of soybean straw III: Isolation and characterization of lignin from soybean straw.” Biomass and Bioenergy 94, 12-20. [37] Espinosa, E.; Sánchez, R.; Otero, R.; Domínguez-Robles, J.; Rodríguez, A. 2017. “A comparative study of the suitability of different cereal straws for lignocellulose nanofibers isolation.” International Journal of Biological Macromolecules 103, 99099. [38] Zhang, A.; Lu, F.; Liu, C.; Sun, R.-C. 2010. “Isolation and Characterization of Lignins from Eucalyptus tereticornis (12ABL).” Journal of Agricultural and Food Chemistry 58, 11287-93. [39] Wang, G.; Chen, H. 2013. “Fractionation and characterization of lignin from steamexploded corn stalk by sequential dissolution in ethanol–water solvent.” Separation and Purification Technology 120, 402-09. [40] Monteil-Rivera, F.; Phuong, M.; Ye, M.; Halasz, A.; Hawari, J. 2013. “Isolation and characterization of herbaceous lignins for applications in biomaterials.” Industrial Crops and Products 41, 356-64. [41] Jiang, X.; Savithri, D.; Du, X. Y.; Pawar, S.; Jameel, H.; Chang, H. M.; Zhou, X. F. 2017. “Fractionation and Characterization of Kraft Lignin by Sequential Precipitation with Various Organic Solvents.” Acs Sustainable Chemistry & Engineering 5, 83542. [42] Cui, C. Z.; Sun, R. K.; Argyropoulos, D. S. 2014. “Fractional Precipitation of Softwood Kraft Lignin: Isolation of Narrow Fractions Common to a Variety of Lignins.” Acs Sustainable Chemistry & Engineering 2, 959-68. [43] Sadeghifar, H.; Wells, T.; Le, R. K.; Sadeghifar, F.; Yuan, J. S.; Ragauskas, A. J. 2017. “Fractionation of Organosolv Lignin Using Acetone:Water and Properties of the Obtained Fractions.” Acs Sustainable Chemistry & Engineering 5, 580-87. [44] Guerra, A.; Lucia, L. A.; Argyropoulos, D. S. 2008. “Isolation and characterization of lignins from Eucalyptus grandis Hill ex Maiden and Eucalyptus globulus Labill. by enzymatic mild acidolysis (EMAL).” Holzforschung 62, 24-30. [45] Zoia, L.; Orlandi, M.; Argyropoulos, D. S. 2008. “Microwave-Assisted Lignin Isolation Using the Enzymatic Mild Acidolysis (EMAL) Protocol.” Journal of Agricultural and Food Chemistry 56, 10115-22.

278

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al.

[46] Saha, K.; Dasgupta, J.; Chakraborty, S.; Antunes, F. A. F.; Sikder, J.; Curcio, S.; dos Santos, J. C.; Arafat, H. A.; da Silva, S. S. 2017. “Optimization of lignin recovery from sugarcane bagasse using ionic liquid aided pretreatment.” Cellulose 24, 3191207. [47] Ugartondo, V.; Mitjans, M.; Vinardell, M. P. 2008. “Comparative antioxidant and cytotoxic effects of lignins from different sources.” Bioresource Technology 99, 6683-87. [48] Bhat, R.; Khalil, H. P. S. A.; Karim, A. A. 2009. “Exploring the antioxidant potential of lignin isolated from black liquor of oil palm waste.” Comptes Rendus Biologies 332, 827-31. [49] Lu, Q.; Liu, W.; Yang, L.; Zu, Y.; Zu, B.; Zhu, M.; Zhang, Y.; Zhang, X.; Zhang, R.; Sun, Z. et al. 2012. “Investigation of the effects of different organosolv pulping methods on antioxidant capacity and extraction efficiency of lignin.” Food Chemistry 131, 313-17. [50] Salleh, N. M.; Ismail, S.; Ibrahim, M. N. M. 2017. “The inhibition of hepatic and renal glucuronidation of p-Nitrophenol and 4-Methylumbelliferone by oil palm empty fruit bunch lignin and its main oxidation compounds.” Pharmacognosy magazine 13, S102. [51] Sa’don, N. A.; Rahim, A. A.; Ibrahim, M. N. M.; Brosse, N.; Hussin, M. H. 2017. “Modification of oil palm fronds lignin by incorporation of m-cresol for improving structural and antioxidant properties.” International Journal of Biological Macromolecules 104, 251-60. [52] Yearla, S. R.; Padmasree, K. 2016. “Preparation and characterization of lignin nanoparticles: evaluation of their potential as antioxidants and UV protectants.” Journal of Experimental Nanoscience 11, 289-302. [53] Domenek, S.; Louaifi, A.; Guinault, A.; Baumberger, S. 2013. “Potential of Lignins as Antioxidant Additive in Active Biodegradable Packaging Materials.” Journal of Polymers and the Environment 21, 692-701. [54] Sun, S.-N.; Cao, X.-F.; Xu, F.; Sun, R.-C.; Jones, G. L. 2014. “Structural Features and Antioxidant Activities of Lignins from Steam-Exploded Bamboo (Phyllostachys pubescens).” Journal of Agricultural and Food Chemistry 62, 5939-47. [55] Suzuki, R.; Matsushita, Y.; Imai, T.; Sakurai, M.; Henriques de Jesus, J. M.; Ozaki, S. K.; Finger, Z.; Fukushima, K. 2008. “Characterization and antioxidant activity of Amazonian woods.” Journal of Wood Science 54, 174-78. [56] Barapatre, A.; Aadil, K. R.; Tiwary, B. N.; Jha, H. 2015. “In vitro antioxidant and antidiabetic activities of biomodified lignin from Acacia nilotica wood.” International Journal of Biological Macromolecules 75, 81-89. [57] Wen, J. L.; Xue, B. L.; Xu, F.; Sun, R. C.; Pinkert, A. 2013. “Unmasking the structural features and property of lignin from bamboo.” Industrial Crops and Products 42, 332-43.

Lignin and Its Derivatives as Antioxidant, Antiviral and Antimicrobial Agents

279

[58] De Clercq, E. 2009. “Anti-HIV drugs: 25 compounds approved within 25 years after the discovery of HIV.” Int J Antimicrob Agents 33, 307-20. [59] WHO; World Health Organization, 2017. [60] Sakagami, H.; Amano, S.; Kikuchi, H.; Nakamura, Y.; Kuroshita, R.; Watanabe, S.; Satoh, K.; Hasegawa, H.; Nomura, A.; Kanamoto, T. et al. 2008. “Antiviral, antibacterial and vitamin C-synergized radical-scavenging activity of Sasa senanensis Rehder extract.” In Vivo 22, 471-6. [61] Sakagami, H.; Iwamoto, S.; Matsuta, T.; Satoh, K.; Shimada, C.; Kanamoto, T.; Terakubo, S.; Nakashima, H.; Morita, Y.; Ohkubo, A. et al. 2012. “Comparative study of biological activity of three commercial products of Sasa senanensis Rehder leaf extract.” In Vivo 26, 259-64. [62] Sakagami, H.; Asano, K.; Satoh, K.; Takahashi, K.; Kobayashi, M.; Koga, N.; Takahashi, H.; Tachikawa, R.; Tashiro, T.; Hasegawa, A. et al. 2007. “Anti-stress, anti-HIV and vitamin C-synergized radical scavenging activity of mulberry juice fractions.” In Vivo 21, 499-505. [63] Mitsuhashi, S.; Kishimoto, T.; Uraki, Y.; Okamoto, T.; Ubukata, M. 2008. “Low molecular weight lignin suppresses activation of NF-kappaB and HIV-1 promoter.” Bioorg Med Chem 16, 2645-50. [64] Lischka, P.; Zimmermann, H. 2008. “Antiviral strategies to combat cytomegalovirus infections in transplant recipients.” Curr Opin Pharmacol 8, 541-8. [65] Zhang, Y.; But, P. P.; Ooi, V. E.; Xu, H. X.; Delaney, G. D.; Lee, S. H.; Lee, S. F. 2007. “Chemical properties, mode of action, and in vivo anti-herpes activities of a lignin-carbohydrate complex from Prunella vulgaris.” Antiviral Res 75, 242-9. [66] Raghuraman, A.; Tiwari, V.; Zhao, Q.; Shukla, D.; Debnath, A. K.; Desai, U. R. 2007. “Viral inhibition studies on sulfated lignin, a chemically modified biopolymer and a potential mimic of heparan sulfate.” Biomacromolecules 8, 1759-63. [67] Thakkar, J. N.; Tiwari, V.; Desai, U. R. 2010. “Non-sulfated, cinnamic acid-based lignins are potent antagonists of HSV-1 entry into cells.” Biomacromolecules 11, 1412-16. [68] Lu, Q.; Zhu, M.; Zu, Y.; Liu, W.; Yang, L.; Zhang, Y.; Zhao, X.; Zhang, X.; Zhang, X.; Li, W. 2012. “Comparative antioxidant activity of nanoscale lignin prepared by a supercritical antisolvent (SAS) process with non-nanoscale lignin.” Food Chemistry 135, 63-67. [69] Peng, Y.; Wu, Y.; Li, Y. 2013. “Development of tea extracts and chitosan composite films for active packaging materials.” International Journal of Biological Macromolecules 59, 282-89. [70] Rojas-Graü, M. A.; Soliva-Fortuny, R.; Martín-Belloso, O. 2009. “Edible coatings to incorporate active ingredients to fresh-cut fruits: a review.” Trends in Food Science & Technology 20, 438-47.

280

G. Vazquez-Olivo, M. S. Criollo-Mendoza, E.k P. Gutiérrez-Grijalva et al.

[71] Yang, W.; Owczarek, J. S.; Fortunati, E.; Kozanecki, M.; Mazzaglia, A.; Balestra, G. M.; Kenny, J. M.; Torre, L.; Puglia, D. 2016. “Antioxidant and antibacterial lignin nanoparticles in polyvinyl alcohol/chitosan films for active packaging.” Industrial Crops and Products 94, 800-11. [72] Yang, W.; Fortunati, E.; Dominici, F.; Giovanale, G.; Mazzaglia, A.; Balestra, G. M.; Kenny, J. M.; Puglia, D. 2016. “Effect of cellulose and lignin on disintegration, antimicrobial and antioxidant properties of PLA active films.” International Journal of Biological Macromolecules 89, 360-68. [73] Yang, W.; Fortunati, E.; Dominici, F.; Giovanale, G.; Mazzaglia, A.; Balestra, G. M.; Kenny, J. M.; Puglia, D. 2016. “Synergic effect of cellulose and lignin nanostructures in PLA based systems for food antibacterial packaging.” European Polymer Journal 79, 1-12. [74] Zemek, J.; Košíková, B.; Augustín, J.; Joniak, D. 1979. “Antibiotic properties of lignin components.” Folia Microbiologica 24, 483-86.

In: Lignin Editors: Fachuang Lu and Fengxia Yue

ISBN: 978-1-53614-769-8 © 2019 Nova Science Publishers, Inc.

Chapter 12

PREPARATION OF DEHYDROGENATION POLYMER FROM ISOEUGENOL AND BIOLOGICAL ACTIVITY CHARACTERIZATION Yimin Xie1,2,*, Xuekuan Chen1, Houkuan Zhao1, Chen Jiang1, Hongfei Wu1, Shuying Bi1 and Zhezi Ye1 1

Research Institute of Pulp and Paper Engineering, Hubei University of Technology, Wuhan, China 2 Hubei Provincial Key Laboratory for Green Light Industry, Hubei University of Technology, Wuhan, China

ABSTRACT To understand the relationship between the chemical structural characteristics of dehydrogenation polymer (DHP) and its antibacterial and anticancer activities, DHP with low molecular weight was prepared in vitro with isoeugenol (IEG) as a precursor of lignin biosynthesis using laccases as the catalyst by a bulk method. The chemical structure of DHP was analyzed with Fourier-transform infrared spectroscopy (FTIR) and carbon-13 nuclear magnetic resonance (13C-NMR) spectroscopy. The antibacterial and anticancer activities of the four DHP fractions with low molecular weight extracted by petroleum ether, diethyl ether, ethanol and acetone were also investigated. The FTIR spectra suggested that the isoeugenol could be polymerized to guaiacyl-type DHP. Analysis of the 13 C-NMR spectra indicated that the DHP was mainly composed of β-5, β-O-4, β-1, β-β and 5-5 subunits. The results of the determination of antibacterial activity showed that the four DHP fractions had a high content of phenolic hydroxyl group, showed strong antimicrobial activity against Escherichia coli and Staphylococcus aureus. The cytotoxicity assay demonstrated that the low molecular weight fraction extracted by ether had obvious anticancer activity. These results showed that DHP with low degree of polymerization had good biological activity. *

Corresponding Author Email: [email protected]

282

Yimin Xie, Xuekuan Chen, Houkuan Zhao et al.

Keywords: dehydrogenation polymer, isoeugenol, 13C-NMR, antimicrobial activity, anticancer activity

INTRODUCTION Lignin, one of the three main components that occurs naturally in higher plants, is a high-molecular weight polymer of phenolic compounds with a complex structure [1]. Deposited in the cell wall, lignin is necessary for both structural support as well as preventing microbial attack [2]. Lignin is polymerized by three monolignols, i.e., pcoumaryl, coniferyl and sinapyl alcohols, with lignin oxidase as the catalyst [3, 4]. As one of the possible phenolic monomers of native wood lignin, isoeugenol is a compound that has a guaiacyl structure [5]. To investigate the occurrence of lignin, dehydrogenation polymer (DHP) synthesized with isoeugenol has been used as a lignin model to explore the lignin structures [6-9]. The biosynthesis of isoeugenol has been elucidated recently in some plants where isoeugenol is produced at a high level [10, 11]. It has been shown that isoeugenol can be synthesized from the monolignol of coniferyl alcohol by a specific enzyme, which results in acylation of side-chains [12]. It has been proposed that isoeugenol biosynthesis is involved in the lignin biosynthetic pathway. The structure of DHP synthesized from isoeugenol has been studied only rarely. Salanti [9] analyzed the lignin-like polymer resulting from isoeugenol radical coupling and found that the characteristic signals of intermonomeric bonds correlated to the β-carbon and αcarbon of β-O-4 and β-5 moieties, respectively, by carbon-13 nuclear magnetic resonance (13C-NMR). Ye [13] studied the polymerization of isoeugenol in the presence of polysaccharides and found that the generated DHP contained β-1, β-β, β-5 and β-O-4 structures. Hunay [14] performed mild oxidative degradation of the isoeugenol polymer and Björkman’s milled wood lignin (MWL) and he found that guaiacyl compounds were produced in both of the samples. Combined with further experimental results, it was confirmed that the isoeugenol polymer was revealed to be of striking similarities to natural lignin [15]. Plant polyphenols are natural products and have high antioxidant, antibacterial and other biological activities [16]. Lignin as a type of plant polyphenolic compound contains a large number of chemical functional groups, such as phenolic hydroxyl, aliphatical hydroxyl, carboxyl, carbonyl and methoxy group. It has been suggested that phenolic hydroxyl and methoxy groups have certain biological activities [17]. Pan [18] analyzed 21 kinds of solvent lignin from the same hybrid poplar and found that the ethanol soluble fractions had the strongest antioxidant properties. Compared to other fractions, these fractions contained more phenolic hydroxyl groups and less alcoholic hydroxyl groups with lower molecular weight and narrower polydispersity. A similar result was found in the studies by Hage [19] and Ma [20]. They obtained the low molecular weight fraction of

Preparation of Dehydrogenation Polymer …

283

MWL in the solvent phase by organic solvent extraction of wood meal, while the fraction with higher molecular weight was rich in precipitate [21]. Compared to a series of guaiacyland syringyl-type lignin and lignin derivatives, isoeugenol was also proved as notable antimicrobial [22, 23]. It has been proposed that the bacteriostatic actions of isoeugenol operate against a variety of bacteria, such as Escherichia coli, Bacillus licheniformis and Staphylococcus aureus [22, 24]. Paper packaging has been endowed with antimicrobial properties by modification with DHPs obtained from isoeugenol [25]. Espinozaacosta [26] summarized the research reports concerning the biological activities of technical lignin and found that lignin exerts potential antimutagenic properties. However, there are few detailed studies on the biological activity of the DHP fractions with different molecule weights obtained from isoeugenol. In the present study, isoeugenol was polymerized to DHP using a bulk method and catalyzed by laccase. The obtained DHP was further fractionated with different organic solvents into fractions with different molecular weight. The chemical structures of the products were characterized with FTIR and 13C-NMR spectroscopy. The antimicrobial and anticancer activities of the obtained DHP were also investigated.

EXPERIMENTAL Materials Isoeugenol (98%) was purchased from Sigma Co. Ltd. Laccase (No. 51003, 1000 IU/mL, determined by the methods of Fukushima and Kirk [27]) was obtained from Novazyme Co. Ltd. All other chemicals were of analytical grade. Gram-negative bacterium, E. coli ATCC 25922 and gram-positive bacterium, S. aureus CMCC(B) 26003 were obtained from Shanghai Luwei Technology Co. Ltd. and used as test organisms. The bacterial inoculums were prepared to obtain a bacterial suspension in 5 mL of nutrient broth. The common nutritional agar culture medium was purchased from Aobox Biotechnology Company and used for the agar plates. HeLa cells were obtained from Shanghai Xinyu Biotechnology Co. Ltd. and used to detect the antitumour properties of the fractions. An autoclave was used for sterilization and media preparation at 121°C for 20 min.

Methods Synthesis of DHP Five grams of isoeugenol was dissolved in 50 mL mixture (1:1, v/v) of ethanol and acetate buffer (0.1 M, pH = 5.0), and mixed with 1 mL laccase (1000 IU/mL). During the

284

Yimin Xie, Xuekuan Chen, Houkuan Zhao et al.

reaction, the mixture was bubbled with sterile air for the required oxygen and maintained in a water bath at 30°C. After 24 h, the crude product was collected by centrifugation and washed with distilled water. After being freeze dried, the crude product was dissolved in 60 mL dichloroethane/ethanol (2:1, v/v) by stirring at 20°C for 6 h in order to remove the contaminant of the enzyme. The supernatant was collected by centrifugation and DHP was obtained after removing the solvent by rotary evaporation in vacuo.

Fractionation of the DHP As shown in Figure 1, the DHP was extracted by petroleum ether, diethyl ether, ethanol and acetone. Four grams of the DHP was suspended in 200 mL of petroleum ether with magnetic stirring. By centrifugation, the petroleum ether solution fraction (F 1) was obtained as the supernatant followed by rotary evaporation with a yield of 25.6%. The above precipitate was further fractionated with diethyl ether, ethanol and acetone using the same method. Fractions F2, F3 and F4 were obtained with yields of 63.9%, 2.5% and 1.0% respectively.

Figure 1. Fractionation procedure of the DHP.

Molecular Weight Determination Two milligrams of the F1, F2, F3 and F4 fractions were dissolved in high-performance liquid chromatography grade tetrahydrofuran and filtrated by 0.22 µm membrane filter.

Preparation of Dehydrogenation Polymer …

285

The molecular weights were determined by gel permeation chromatography (GPC) with a column of Shim-pack GPC-803D (Size: 300 mm × 8 mm ID). Tetrahydrofuran was applied as mobile phase with a flow rate of 0.6 mL/min. Column temperature was 30°C and injection volume was 25 µL.

Determination of Total Phenol Content Based on the method by Christel Quettier-Deleu [28], the total phenol content of each fraction was determined. Seven milliliters of distilled water, 0.5 mL phenolic phenol reagent and 0.5 mL ethanol solution of catechol were mixed and shaken for 3 min. After 2 mL 20% Na2CO3 solution was added, the mixture was heated for 1 min in a 100°C water bath and cooled naturally to 20°C. Then, its absorbance was determined with a UV-Vis spectrometer at 685 nm. A standard curve was plotted for the concentration of catechol and absorbance. By repeating the above steps, F1, F2, F3, F4 and isoeugenol (IEG) were tested to determine the total phenol content. Analysis of the Structure of the DHP The FTIR spectrum of the DHP was obtained with a Thermofisher Nicolet 6700 FTIR spectrometer using the KBr pellet technique. The samples were scanned 32 times in the range of 4000 to 500 cm-1. The liquid 13C-NMR spectrum of the sample was determined at 100.6 MHz on a Varian oneProbe 400 NMR spectrometer. The sample was placed in a φ5 mm determining tube and dissolved in 0.6 mL dimethyl sulfoxide (DMSO)-d6 solvent. Pulse delay was 1.75 s and acquisition time was 0.9 s. The sample was scanned approximately 20000 times. Evaluation of Antimicrobial Activity Antimicrobial activities of the DHP fractions were tested using the filter paper agar diffusion method [29]. Antimicrobial activities were determined by their zone of inhibition. The bacterial suspension was diluted with sterile saline to 1.5 × 108 CFU/mL. Two hundred microliters of bacterial suspension was uniformly coated onto every agar plate. The four DHP fractions were dissolved in DMSO to obtain a series of solutions with concentration of 5 mg/mL to 2.5 mg/mL. Dried sterile filter papers of 6 mm in diameter that had been prepared previously were immersed in the above solutions for 6 h. Then, the filter papers were removed and attached onto the agar plates, followed by the addition of 10 µL of the corresponding sample solutions onto the surface of the filter papers. After culture at 37°C for 16 h, the zone of inhibition was observed and measured. Evaluation of Anticancer Activity Anticancer activity of the four DHP fractions were detected by the 3- (4,5- dimethyl2- thiazolyl) - 2,5-diphenyl-2-H-tetrazolium bromide (MTT) method [30]. HeLa cells were cultured in Roswell Park Memorial Institute 1640 medium, which was supplemented with

Yimin Xie, Xuekuan Chen, Houkuan Zhao et al.

286

100 U/mL penicillin and 100 µg/mL streptomycin. The cells were cultured at 37°C and 5% CO2 in a humidified incubator. HeLa cells (1 × 106 cells/mL) were pre-treated with various concentrations of the four DHP fraction solutions or solution medium (1% ethanol) for 24 h. The cells were rinsed and incubated with MTT (5 mg/mL) for 4 h. The culture solution was discarded and the formazan blue formed in the cells was dissolved with DMSO. The absorbance at 490 nm was measured in a microplate reader. The absorbance of the formazan formed in the intact cell was taken as the controls. Finally, the semi-inhibition concentration (IC50%) was calculated via the curves.

RESULTS AND DISCUSSION Molecular Weight of the DHP The molecular weights of the four DHP fractions, i.e., F1, F2, F3 and F4 are shown in Table 1. The average molecular weights (Mw) of the F1, F2, F3 and F4 fractions were 330, 621, 1211 and 3670, respectively. Table 1. Molecular weight of the fractions of the DHP Fractions F1 F2 F3 F4

Mw 330 621 1211 3670

Mn 126 237 449 1579

Mw/Mn 2.6 2.6 2.7 2.3

Note: TPC: Total phenol content.

Figure 2. FTIR spectra of isoeugenol (a) and the DHP (b).

TPC(mg catechol/g sample) 149 104 93.5 80

Preparation of Dehydrogenation Polymer …

287

The molecular weight of the fractions increased with the solubility of the solvent. Because the molecular weight of monomeric compound IEG was 162, the main components of the four fractions can be calculated to be dimer, tetramer, heptamer and dodecamer. The dispersion coefficient (Mw/Mn) of the four components of F1, F2, F3 and F4 were approximately 2.6, which means there were a lot of low molecular weight substances in the DHP fractions. The total phenol contents of the F1, F2, F3 and F4 were 149, 104, 93.5 and 80 mg catechol/g sample, respectively, which indicated that the total phenol content decreased with the increase of molecular weight.

FTIR Analysis of the DHP As shown in Figure 2, FTIR spectroscopy was used to analyze the DHP from isoeugenol to reveal the chemical structure in the region between 2200 cm-1 and 600 cm-1. Based on the assignment reported previously [6, 13, 14], the specific absorption associated with lignin assigned to aromatic skeletal vibrations were located at 1509 cm-1 and 1600 cm1 , as shown in Figure 2. Furthermore, the stretching vibration of the guaiacyl rings was at a lower wavenumber range at 1267 cm-1. However, a strong band at 1740 cm-1 which was identified as unconjugated carboxyl stretching only appeared in the DHP spectrum. This indicated the possibility of the formation of carboxyl groups by oxidation of isoeugenol during polymerization catalyzed by laccase. Moreover, the peak at 963 cm-1 was characteristic of a conjugated double bond in dehydrodiisoeugenol, which could not be found in the spectrum of MWL [6].

Analysis of the DHP by 13C-NMR Spectroscopy The 13C-NMR spectrum of DHP was shown in Figure 3. The assignment of signal absorption peaks was shown in Table 2, and the structural units of DHP were shown in Figure 4. The signal at 172.2 ppm (No. 2) could be assigned to γ-COOH (I, Figure 4) of cinnamic acid. The peak at 169.8 ppm (No. 3) was Cα = O (IV and V, Figure 4) of vanillic acid and its derivative. These two different carbonyl groups indicated that DHP had a different degree of oxidation in the side chain during the polymerization process [8]. The signal at 152.4 ppm (No. 4) was from C3 in etherified 5-5 structure (X, Figure 4). In addition, signals at 130 to 134 ppm (No. 9-11) were mainly from C1 in β-O-4, β-5 and 55. At 128.4 ppm (No. 12) there was a weak signal, which came from C α and Cβ in -Cα = Cβ- structure in the coniferyl alcohol (II, Figure 4) and showed that a small amount of γCH3 of isoeugenol was oxidised to γ-CH2OH. From 110 ppm to 125 ppm, there was a large amount of signal, which was mainly from C5 and C6 of the aromatic ring of lignin [31, 32].

Yimin Xie, Xuekuan Chen, Houkuan Zhao et al.

288

Figure 3. 13C-NMR spectrum of the DHP. COOH

CH3

CH2OH

O

O

L

COOH

O

O

O L

O L

I

II

O L

O L

V

IV

III

R1

O

O

O O L

R1

O

O

O

O

O

R1

O

O

O

OH

O

HO

OH O

O

O

HO O

O

O

L

O

L

VI

L

L

VIII

VII

IX O

L

L

O

O O

L

OH HO

OH

OH

O

HO

O OH O

L R1

X

O

O

O

XI

XII

L

Note: R 1=H or alkyl; L=polylignol

Figure 4. Substructures of the DHP.

R1

Preparation of Dehydrogenation Polymer …

289

The obvious signal at 93.0 ppm (No. 24) was the resonance of Cα in β-5 with -CγH3 (VII, Figure 4). This demonstrated that the DHP contained large amount of β-5 structure. The signal at 88.7 ppm (No. 25) was the resonance of Cβ in β-1 (XI, Figure 4), which indicated that the DHP contained only small amount of β-1. It was suggested that the DHP contained β-β (XII, Figure 4) as Cβ in β-β appeared at 85.7 ppm (No. 26). The signal at 78.1 ppm (No. 29) was Cα in β-O-4, which demonstrated that DHP contained a large amount of β-O-4 with γ-CH3 (XIII, Figure 4). This meant that a lot of β-O-4 could be synthesized by dehydrogenation polymerization of isoeugenol even with the bulk method [9, 15]. Signal No. 30 can be assign to Cα in β-O-4 with γ-CH2OH (IX, Figure 4). A weak signal at 55.0ppm (No. 34) was from Cβ of β-5 with γ-CH2OH (VI, Figure 4). In addition, there was a strong signal from Cγ of β-5 with -CγH3 at 45.1 ppm (No. 35). This proved that the content of β-5 in DHP was high. The strong signals at 18.4 ppm (No. 39) and 15.6 ppm (No. 40) could be assigned to the γ-methyl groups of β-5 and β-O-4 in the DHP. From the spectral analysis, it could be concluded that the main structural units in the DHP were β-5, β-O-4, β-1, β-β and 5-5. These were very similar to the structural units of protolignin.

Investigation of Antibacterial Activity The DHP obtained from isoeugenol was hydrophobic, i.e., could only be dissolved in organic solvents such as DMSO. Therefore, the antibacterial activity of DMSO was first determined. It was found that DMSO could not inhibit the growth of E. coli and S. aureus. Therefore, DMSO was suitable to dissolve the DHP samples in the antibacterial experiment. The zones of inhibition of bacteria were shown in Figure 5. The antibacterial activities of the four DHP fractions (F1 ~ F4) were presented in Table 3 and Table 4. The F1 and F2 fractions with concentration of 5 mg/mL and 2.5 mg/mL, respectively, inhibited the growth of E. coli and S. aureus, resulting in an obvious zone of inhibition. However, the experiments with the F3 and F4 fractions did not show clear inhibition zone. Table 4 showed that the F1 and F2 fractions produced zones of inhibition on gram-positive S. aureus (8 mm and 9 mm, respectively), whereas zones of inhibition of 7–8 mm appeared on gramnegative E. coli as shown in Table 3. These results indicated that the antibacterial compounds of the DHP were mainly low molecular weight fractions that were soluble in petroleum ether and diethyl ether. The structures of the side chain containing a double bond at the α and β positions, and/or a methyl group at γ might play a considerable role in antimicrobial activity [22]. Most of these structures might remained in the F1 and F2 fractions. The antibacterial experiments proved that the DHP fractions with low molecular weight had strong antibacterial activity

290

Yimin Xie, Xuekuan Chen, Houkuan Zhao et al.

against E. coli and S. aureus. Moreover, it provided a theoretical basis for lignin modification and application. Table 2. Assignments of 13C NMR spectrum of the DHP Signal 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

 (ppm) 180.4 172.2 169.8 152.4 150.5 147.8 146.7 144.1 133.8 131.4 129.9 128.4 125.1 123.1 121.7 120.4 119.0 116.8 115.8 113.9 111.7 110.6 109.4 93.0 88.7 85.7 84.0 79.2 78.1 74.8 69.8 64.4 56.5 55.0 45.1 40.5 29.4 21.2 18.4 15.6

Assignments O-C=O in carboxylic acid Cα in cinnamic acid Cα=O in benzoic acid C3 (etherified 5-5) C3(β-O-4) C3(β-5) C4(β-5) or C3(β-O-4) C4(5-5) C1(β-O-4) C1(β-5) C1(5-5) Cα and Cβ (coniferyl alcohol) C5(5-5) C6(β-β) C6(5-5) C1(β-1) or C1(etherified β-O-4) C6(β-5) or C6(β-O-4) C5(β-β) C4(β-β) C5(β-O-4) C2(β-O-4) C2(β-5) C2(5-5) Cα(β-5) Cβ(β-1) Cβ (β-β with -CγCH3) Cα(etherified β-O-4) Cβ(etherified β-O-4) or Cα(β-β) Cβ(β-O-4 with -CγCH3) Cα(β-O-4 with Cγ-CH2OH) unknown Cγ(β-5 with Cγ-CH2OH) -OCH3 Cβ(β-5 with Cγ-CH2OH) Cγ (β-5 with -CγCH3) DMSO -CH3 -CH3 Cγ(β-5 with -CγH3) Cγ(β-O-4 with -CγH3)

Preparation of Dehydrogenation Polymer …

291

Table 3. Activities of the DHP fractions against E. coli Concentration gradient(mg/ml) 2.5 5

Zone of inhibition(mm) F1 F2 F3 8 7 -8 7 --

F4 ---

Legend: F1: Figure 5a and Figure 5b; F2: Figure 5c and Figure 5d. The 5 and 2.5 represent concentration of solution in mg/mL. Figure 5. The inhibition zone of the F1 and F2 fractions of the DHP.

Table 4. Activities of the DHP fractions against S. aureus Concentration gradient(mg/ml) 2.5 5

Zone of inhibition(mm) F1 F2 F3 9 8 -9 8 --

F4 ---

Investigation of Anticancer Activity of the DHP Fractions As shown in Figure 6 and Table 5, the four DHP fractions (F1 ~ F4) had an inhibitory effect on the proliferation of HeLa cancer cells. With an increase in concentration, the effect of growth inhibition was more obvious. The lower molecular weight fractions extracted by petroleum (F1) and diethyl ether (F2) showed excellent anticancer activity, with semi-inhibitory concentrations (IC50%) of 59.764 µg/mL and 16.146 µg/mL, respectively. The inhibition effect of high molecular weight fractions was not significant. The anticancer activities were relevant to the phenolic hydroxyl structure in DHP fractions with low molecular weight. Lignin is one of the polyphenolic compounds in plants and

Yimin Xie, Xuekuan Chen, Houkuan Zhao et al.

292

phenolic hydroxyl has a strong anti-free radical capacity. The oligomers with low molecular weight from DHP polymerized by isoeugenol led to a high content of free phenolic hydroxyl groups [26, 33].

Figure 6. Relationship between concentrations of the DHP fractions and inhibition rates.

Table 5. IC50% values of the four DHP fractions Fractions of DHP IC50% (μg/ml)

F1 59.764

F2 16.146

F3 317.697

F4 822.731

CONCLUSION 1. The FTIR analysis showed that the isoeugenol precursor was polymerized to be DHP. By analyzing the 13C-NMR spectrum of the DHP, it was found that the phenylpropane units of the DHP were mainly connected by β-O-4, β-β, β-5 and β1 linkages with the existence of minor subunits as ferulic acid, isoeugenol and coniferyl alcohol. These results proved that the association between phenylpropane units in the DHP from isoeugenol was similar to that of protolignin. 2. The antibacterial experiments showed that the DHP fractions with low molecular weight had strong antibacterial activities against Escherichia coli and Staphylococcus aureus. This result provided a theoretical basis for lignin modification and application. 3. The four DHP fractions classified by petroleum ether, diethyl ether, ethanol and acetone obviously inhibited the proliferation of Hela cancer cells, especially for

Preparation of Dehydrogenation Polymer …

293

the diethyl ether extracted fraction F2. This effect may be related to the high content of free phenolic hydroxyl in its structure.

ACKNOWLEDGMENTS The authors are grateful for the support of the Nature Science Foundation of China (grant No. 21878070) and Hubei Provincial Key Laboratory of Green Materials for Light Industry (Q20131402). Author Contributions: Yimin Xie conceived and designed the experiments; Xuekuan Chen, Houkuan Zhao, Chen Jiang and Hongfei Wu performed the experiments; Shuying Bi, Zhezi Ye analyzed the data; Yimin Xie and Xuekuan Chen wrote the paper. The authors declare no conflict of interest.

REFERENCES [1]

[2]

[3] [4] [5] [6]

[7]

[8]

Baurhoo, B., Ruiz-Feria, C. A. & Zhao, X. (2008). “Purified lignin: Nutritional and health impacts on farm animals-A review.” Animal Feed Science and Technology, 144(3), 175-184. Upton, B. M. & Kasko, A. M. (2015). “Strategies for the Conversion of Lignin to High-Value Polymeric Materials: Review and Perspective.” Chemical reviews, 116(4), 2275-2306. Holmgren, A., Norgren, M., Zhang, L., et al. (2009). “On the role of the monolignol γ-carbon functionality in lignin biopolymerization.” Phytochemistry, 70(1), 147-155. Freudenberg, K. (1965). “Lignin: its constitution and formation from p-hydroxycinnamyl alcohols.” Science, 148(3670), 595-600. Zemek, J., Košíková, B., Augustin, J., et al. (1979). “Antibiotic properties of lignin components.” Folia microbiologica, 24(6), 483-486. Evliya, H. & Olcay, A. (1974). “Oxidative polymerisation of isoeugenol and mild oxidation of synthetic polymers with alkaline cupric hydroxide.” Holzforschung, 28(4), 130-135. Schneider, S., Rolando, C., Lapierre, C., et al. (1994). “Thioacidolysis of dehydropolymers of isoeugenol and 4-(1-hydroxybutyl)-2-methoxyphenol prepared under oxidative conditions.” Holzforschung, 48(4), 313-317. Shiba, T., Xiao, L, Miyakoshi, T., et al. (2000). “Oxidation of isoeugenol and coniferyl alcohol catalyzed by laccases isolated from Rhus vernicifera Stokes and Pycnoporus coccineus.” Journal of Molecular Catalysis B: Enzymatic, 10(6), 605615.

294 [9]

[10]

[11]

[12]

[13]

[14] [15] [16]

[17]

[18]

[19]

[20] [21] [22]

Yimin Xie, Xuekuan Chen, Houkuan Zhao et al. Salanti, A., Orlandi, M., Tolppa, E. L., et al. (2010). “Oxidation of isoeugenol by salen complexes with bulky substituents.” International Journal of Molecular Sciences, 11(3), 912-926. Hoballah, M. E., Stuurman, J., Turlings, T. C. J., et al. (2005). “The composition and timing of flower odour emission by wild Petunia axillaris coincide with the antennal perception and nocturnal activity of the pollinator Manduca sexta.” Planta, 222(1), 141-150. Koeduka, T., Fridman, E., Gang, D. R., et al. (2006). “Eugenol and isoeugenol, characteristic aromatic constituents of spices, are biosynthesized via reduction of a coniferyl alcohol ester.” Proceedings of the National Academy of Sciences, 103(26), 10128-10133. Muhlemann, J. K., Woodworth, B. D., Morgan, J. A., et al. (2014). “The monolignol pathway contributes to the biosynthesis of volatile phenylpropenes in flowers.” New Phytologist, 204(3), 661-670. Ye, Z, Xie, Y., et al. (2016). “Dehydrogenation Polymerization of Isoeugenol and Formation of Lignin-carbohydrate Complexes with Presence of Polysaccharide.” Chemistry and Industry of Forest Products, 36(2), 45-50. Hunay, E. (1989). “C-13 NMR Studies of a Dehydropolymer (DHP) from Isoeugenol; Comparison with Spruce Lignin.” Holzforschung, 43(1), 61-64. Nakano, J. (1988). Chemistry of Lignin Basic and Application. Tokyo: Uni Press. (Nakano 1988, 545). Jamshidian, M., Tehrany, E. A. & Desobry, S. (2012). “Release of synthetic phenolic antioxidants from extruded poly lactic acid (PLA) film.” Food Control, 28(2), 445455. Arshanitsa, A. (2013). “Fractionation of technical lignins as a tool for improvement of their antioxidant properties.” Journal of Analytical & Applied Pyrolysis, 103(9), 78-85. Pan, X., Kadla, J. F., Ehara, K., et al. (2006). “Organosolv ethanol lignin from hybrid poplar as a radical scavenger: relationship between lignin structure, extraction conditions, and antioxidant activity.” Journal of Agricultural & Food Chemistry, 54(16), 5806-5813. Hage, R. E., Perrin, D. & Brosse, N. (2012). “Effect of the Pre-Treatment Severity on the Antioxidant Properties of Ethanol Organosolv Miscanthus x giganteus Lignin.” Natural Resources, 03(2), 29-34. Ma, P., Gao, Y. & Zhai, H. (2013). “Fractionated Wheat Straw Lignin and Its Application as Antioxidant.” Bioresources, 8(4), 5581-5595. Ropponen, J., Räsänen, L., Rovio, S., et al. (2011). “Solvent extraction as a means of preparing homogeneous lignin fractions.” Holzforschung, 65(4), 543-549. Zemek, J., Košíková, B., Augustin, J., et al. (1979). “Antibiotic properties of lignin components.” Folia microbiologica, 24(6), 483-486.

Preparation of Dehydrogenation Polymer …

295

[23] Zemek, J., Valent, M., Pódová, M., et al. (1987). “Antimicrobiai properties of aromatic compounds of plant origin.” Folia microbiologica, 32(5), 421-425. [24] Laekeman, G. M., Van Hoof, L., Haemers, A., et al. (1990). “Eugenol a valuable compound for in vitro experimental research and worthwhile for further in vivo investigation.” Phytotherapy Research, 4(3), 90-96. [25] Elegir, G., Kindl, A., Sadocco, P., et al. (2008). “Development of antimicrobial cellulose packaging through laccase-mediated grafting of phenolic compounds.” Enzyme and Microbial Technology, 43(2), 84-92. [26] Espinozaacosta, J. L., Torreschávez, P. I., Ramírezwong, B., et al. (2016). “Antioxidant, antimicrobial, and antimutagenic properties of technical lignins and their applications.” Bioresources, 11(2), 5452-5481. [27] Fukushima, Y. & Kirk, T. K. (1995). “Laccase component of the Ceriporiopsis subvermispora lignin-degrading system.” Applied and Environmental Microbiology, 61(3), 872-876. [28] Quettierdeleu, C., Gressier, B., Vasseur, J., et al. (2000). “Phenolic compounds and antioxidant activities of buckwheat (Fagopyrum esculentum Moench) hulls and flour.” Journal of Ethnopharmacology, 72(1–2), 35-42. [29] Muniandy, K., Hassan, Z. & Isa, M. H. M. (2014). “Effect of heat and filter sterilization on the efficiency of coleus aromaticus as an antibacterial agent against diabetic wound pathogens.” International Journal of Pharmacy and Pharmaceutical Sciences, 6(10), 438-443. [30] Choi, C. Y., Park, K. R., Lee, J. H., et al. (2007). “Isoeugenol suppression of inducible nitric oxide synthase expression is mediated by down-regulation of NF-kappaB, ERK1/2, and p38 kinase.” European Journal of Pharmacology, 576(1–3), 151-159. [31] Lüdemann, H. D. & Nimz, H. (1973). “Carbon-13 nuclear magnetic resonance spectra of lignins.” Biochemical and biophysical research communications, 52(4), 1162-1169. [32] Xie, Y., Yasuda, S., Wu, H., et al. (2000). “Analysis of the structure of lignincarbohydrate complexes by the specific 13C tracer method.” Journal of wood science, 46(2), 130-136. [33] Munk, L., Sitarz, A. K. & Kalyani, D. C., et al. (2015). “Can laccases catalyze bond cleavage in lignin?.” Biotechnology Advances, 33(1), 13-24.

ABOUT THE EDITORS Fachuang Lu, PhD, Professor, graduated from University of Wisconsin-Madison in 1998. Lu obtained his PhD in plant cell wall chemistry. He had worked as a senior scientist in the biochemistry department at UW-Madison for over 8 years before he took a position as a professor/director of State Key Laboratory of pulp and paper engineering at South China University of Technology (SCUT) in 2015. His research interests include analytical methodology on plants from various origins, biosynthesis of plant cell wall components, organic synthesis of plant cell wall model compounds, and plant derived bioactive molecules for various applications, etc. He has published over 120 papers in peer-reviewed journals, and co-authored in 6 book chapters. His current research projects are focusing on structural elucidation of lignin from genetic modified plants, identifying potential transgenic as feedstock for biofuel/biochemical productions, and converting lignin into platform chemicals.

Fengxia Yue, PhD, Associate Professor. Dr. Yue received her PhD degree in Pulping and Paper Engineering from South China University of Technology. She joined John Ralph’s Group, as a research associate at University of Wisconsin-Madison till August, 2017 when she was appointed as an associate professor in State Key Laboratory of Pulp and Paper Engineering, South China University of Technology. Her research interests include lignin biochemistry and biosynthesis, lignin characterization, lignin model compounds synthesis, NMR structural analysis, and green products from renewable resources.

INDEX # 1,2-dihydroxybenzene, 177 2-chloro-4,4,5,5-tetramethyl-1,3,2dioxaphospholane, xii, 195, 201, 242 3-methoxy, 139, 143, 263 4-hydroxybenzoic acid, 176, 178 4-hydroxycinnamyl, 1, 2, 263 5-hydroxyconiferyl alcohol, 6, 21, 22, 26, 43, 81, 87, 88, 105, 263 5-hydroxylase, xi, 4, 18, 21, 104, 114, 126, 192

A acidolysis, 49, 61, 63, 64, 80, 101, 104, 117, 127, 149, 265, 277 amphiphilic, 207, 216, 232, 234, 237 angiosperm, 10, 15, 39, 83, 106 anticancer, viii, 55, 281, 282, 283, 285, 291 anti-free radical capacity, 292 antimicrobial, vi, viii, 55, 72, 238, 261, 262, 272, 273, 274, 275, 280, 281, 282, 283, 285, 289, 295 antioxidant, vi, viii, 56, 70, 76, 235, 261, 262, 265, 266, 268, 269, 271, 273, 274, 275, 276, 278, 279, 280, 282, 294, 295 antiviral activity, viii, 261, 270, 271, 272 aromatic rings, 116, 120, 122, 123, 138, 147, 148, 151, 167, 174, 178, 216, 222, 249, 250 azide groups, 207, 211, 215, 232

B benzoate, 40, 170, 176, 177, 183, 184, 185

benzodioxane, 6, 9, 21, 43, 86, 87, 88, 92, 105 bioengineering, 23, 27, 35, 37, 38, 40, 46, 48, 64, 69, 79, 87, 99, 100, 125, 156, 203 bioethanol, v, viii, 131, 132, 133, 153, 156, 157, 161, 204, 276 bonding patterns, 80

C cactaceae seedcoats, 87 caffeic acid, xi, 4, 5, 12, 18, 21, 22, 43, 68, 77, 104, 192 caffeoyl shikimate esterase, 5, 12, 19, 20, 53, 192, 204, 275 catechol, 13, 120, 168, 171, 174, 176, 177, 184, 185, 186, 285, 286, 287 cellulolytic enzyme lignin, xi, 98, 109, 191, 205, 206 cement, 179, 262 chalcone, xi, 6, 21, 52, 53, 57, 63 cinnamic acid(s), 3, 279, 287, 290 cinnamoyl CoA reductase, 4, 6, 12, 15, 17 cinnamyl alcohol, xi, 4, 5, 12, 17, 20, 28, 30, 42, 70, 106, 192, 204, 293 cinnamyl alcohol dehydrogenase, xi, 4, 5, 12, 17, 20, 42, 70, 106, 192, 204 coniferyl alcohol, 6, 26, 30, 37, 54, 64, 68, 86, 88, 90, 92, 93, 113, 169, 174, 192, 198, 253, 255, 263, 282, 287, 290, 292, 293, 294 correlation spectroscopy, xii, 92, 93 cytotoxicity, 56, 73, 265, 266, 271, 281

D decarboxylase(s), 169, 172, 177, 185, 186 decarboxylation, 169

Index

300 dehydrodiisoeugenol, 287 dehydrogenase, xi, 4, 5, 12, 17, 20, 42, 70, 106, 167, 172, 176, 184, 192, 204 derivatization followed by reductive cleavage, xi, 44, 64, 80, 101 diaryl, 87, 104, 114, 138, 158, 193 dibenzodioxocin(s), 80, 87, 101, 114 dicarboxylic acid, 172, 176, 177, 186 diferuloylputrescine, 49, 81, 102 dihydroferulic, 169 dimethylsulfoxide, xi, 9, 82, 103 dioxygenase, 169, 171, 177, 183, 185, 186 downstream, 25, 35, 65, 121, 135, 171, 203

E ester linkage, 25, 29, 36, 38, 39, 44, 45, 114

F F5H, xi, 4, 6, 21, 192, 205 fabrication, 207, 208, 209, 216, 223, 232, 233, 238, 239, 240, 242 ferulate ester conjugates, 26, 52 ferulic acid, 5, 6, 15, 21, 28, 88, 106, 168, 172, 173, 176, 182, 188, 189, 205, 263, 268, 292 flavonoid, 15, 43, 51, 52, 53, 55, 56, 67, 69, 71, 73, 74, 75, 77, 102 flavonolignans, 54, 70, 72 flavonolignin, 51 flavonolignols, 54, 68 formaldehyde, 47, 116, 117, 118, 119, 122, 126, 151, 156, 179, 188, 217, 239, 259 Fourier transform infrared spectroscopy, viii, 243, 249

G genetic manipulation, 19, 42, 80, 126, 205 glucomannans, 99 guaiacol, 119, 120, 136, 139, 140, 141, 144, 148, 151, 159, 173, 176, 186 guaiacylpropane, 244, 253 gymnosperms, 3, 32, 39, 263

H hardwood, vi, viii, 28, 44, 83, 84, 85, 87, 113, 114, 121, 122, 123, 127, 130, 173, 174, 243, 244, 245, 248, 251, 253, 254, 258, 263, 276 heterogeneity, 18, 26, 80, 108, 150, 166, 168, 180, 186, 265, 275 hybrid poplar, 35, 38, 41, 48, 104, 126, 173, 282, 294 hydrogenation, 121, 129, 136, 137, 138, 144, 146, 147, 148, 150, 155, 158, 159, 160, 162, 163, 177 hydrogenolysis, 28, 30, 80, 119, 121, 122, 129, 136, 138, 140, 141, 142, 147, 149, 151, 152, 156, 158, 160, 161, 162 hydrolysate, 176, 177, 185 hydrolysis, vii, 28, 30, 37, 38, 48, 80, 98, 100, 132, 138, 144, 148, 167, 168, 194, 206, 216, 224, 239, 244, 265 hydroxycinnamaldehyde, 2, 5, 192 hydroxycinnamaldehyde dehydrogenase, 192 hydroxycinnamyl, 1, 2, 5, 26, 28, 30, 31, 106, 192, 263 hydroxylase, vi, viii, xi, 3, 4, 12, 18, 19, 20, 21, 53, 67, 71, 77, 104, 107, 114, 126, 172, 191, 192, 195, 204, 205

K Klason lignin, vi, viii, 9, 65, 151, 194, 195, 196, 243, 244, 245, 248, 251, 263

L laccase, viii, 124, 173, 178, 182, 283, 287, 295 ligase, xi, 4, 12, 53, 192 lignification, viii, 2, 3, 5, 6, 11, 14, 15, 16, 17, 18, 19, 21, 23, 25, 26, 28, 30, 33, 35, 39, 40, 41, 42, 43, 47, 48, 65, 69, 70, 72, 75, 87, 100, 102, 104, 105, 126, 192, 263 lignin biosynthesis, v, viii, ix, 1, 3, 5, 6, 7, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 25, 26, 27, 35, 40, 42, 43, 64, 69, 77, 100, 101, 105, 106, 111, 113, 114, 115, 192, 193, 204, 281 lignin-carbohydrate complex, xii, 60, 76, 91, 99, 108, 109, 192, 262, 264, 269, 276, 279, 295 lignin-carbohydrate protein complexes, 270, 271 lignin-first, 121, 132, 149, 150, 153, 159, 162, 163 lignolytic, viii, 165, 166, 169, 174, 179, 181, 184

301

Index low-weight molecules, 165

M magnetization, 89, 91, 92, 96 maize, 5, 6, 15, 16, 21, 30, 32, 40, 44, 45, 47, 48, 49, 52, 54, 58, 59, 62, 65, 68, 71, 72, 77, 102, 105, 173, 204 manganese peroxidase, 124 metabolic, v, viii, 4, 19, 43, 165, 166, 167, 168, 169, 170, 171, 172, 174, 175, 178, 179, 180, 182, 185, 186, 188, 204 methylation, 3, 4, 5, 6, 19, 52, 56, 71, 74, 96 methylene, 52, 116, 117, 118, 122, 210, 226 methyltransferase, xi, xii, 3, 4, 5, 12, 18, 21, 22, 43, 45, 52, 53, 67, 68, 71, 77, 104, 106, 192 microorganisms, 130, 166, 169, 171, 174, 176, 178, 179 milled wood lignin, xii, 30, 44, 57, 58, 75, 98, 101, 107, 109, 205, 258, 282 Monocot, 21 monolignol acetate, 25, 26, 40, 52 monophenolic, 120, 163 multi-dimensional heteronuclear (NMR), v, viii, xi, xii, 9, 11, 15, 16, 20, 21, 29, 30, 31, 33, 34, 40, 41, 42, 43, 44, 45, 51, 57, 58, 61, 62, 63, 70, 75, 76, 79, 80, 81, 82, 83, 86, 87, 88, 91, 93, 94, 95, 96, 97, 98, 99, 101, 102, 103, 104, 105, 107, 108, 109, 116, 117, 127, 128, 163, 191,192, 193, 195, 196, 199, 200, 201, 203, 205, 206, 210, 211, 212, 214, 227, 281, 282, 283, 285, 287, 288, 290, 292, 294, 297

N nitrobenzene oxidation, 80, 123, 130 nuclear magnetic resonance, xi, xii, 29, 75, 80, 103, 108, 109, 205, 281, 282, 295 number-average, xii, 10, 194, 202, 266

P p-acetoxybenzoate, 30 p-coumarate, viii, 4, 16, 25, 26, 31, 32, 42, 44, 46, 47, 48, 52, 54, 68, 69, 83, 87, 95, 98, 176, 182, 184, 191, 192, 263 p-coumaroylation, 31 permanganate oxidation, 80 perturbations, 1, 5 phenolic, 2, 13, 20, 28, 30, 56, 73, 74, 113, 116, 117, 118, 119, 122, 124, 129, 130, 138, 141, 144, 148, 150, 151, 160, 162, 166, 167, 201, 209, 210, 215, 224, 227, 237, 242, 244, 255, 256, 261, 262, 263, 264, 267, 268, 269, 272, 273, 275, 281, 282, 285, 291, 293, 294, 295 phenylalanine, 3, 12, 17, 98, 192, 263 phenylcoumaran, 10, 87, 114 phenylpropanoid, 17, 19, 52, 54, 55, 82, 91, 92, 192, 195, 204, 216 p-hydroxybenzoate, 25, 26, 44, 45, 52, 95, 172, 191, 192, 198, 202, 203, 263 p-hydroxybenzoic acid, 2, 28, 29, 169, 176, 182 p-hydroxyphenyl, xi, 2, 4, 52, 82, 83, 113, 167, 191, 193, 227 picolinic acid, 176, 177, 179, 186 platinum, 142 PMT transgenes, 36 Poaceae, 54, 64 polyethylene glycol, 207, 232 polymer-grafted, 209, 235 pre-acetylation, 33 protocatechuate, 168, 170, 172, 174, 176, 177, 178, 185, 186 protolignin, 108, 289, 292

Q quinone methide, 9, 28, 33, 34, 87, 116

R O o-methyltransferase, xi, xii, 4, 5, 12, 13, 18, 21, 22, 43, 45, 52, 53, 67, 68, 71, 77, 104, 106, 192 oxidation, 28, 47, 48, 80, 120, 123, 130, 153, 168, 182, 266, 271, 272, 274, 278, 287, 293, 294

radical coupling, 6, 9, 26, 31, 33, 34, 57, 59, 60, 68, 87, 88, 113, 116, 263, 282 radical scavenging activity, 56, 74, 265, 266, 268, 271, 275, 279 regiochemistry, 29, 30, 31, 45 repolymerization, 111, 112, 116, 117, 119, 121

Index

302 S Salix clones, 30, 44 self-assembly, vi, viii, 207, 216, 217, 222, 223, 229, 231, 232, 237, 238, 239, 240, 241 semi-quantitative, 64, 82, 88, 141, 227 shikimate and polyketide biosynthetic pathways, 51 side-chain linkages, 80 sinapic acid, 4, 114, 168, 172, 182, 183 sinapyl alcohol, 2, 6, 11, 26, 31, 33, 34, 35, 36, 45, 47, 52, 68, 86, 88, 92, 113, 114, 192, 254, 255, 263, 282 softwood, vi, viii, 41, 83, 84, 85, 87, 91, 93, 101, 104, 107, 113, 114, 119, 121, 122, 123, 127, 128, 173, 174, 192, 203, 235, 243, 244, 245, 248, 251, 253, 263, 265, 276, 277 spirodienone, 80, 98, 101, 114 stabilize, 122, 151, 217, 224

T thermogravimetry, 245, 259, 260 thioacetolysis, 80 thioacidolysis, 7, 8, 9, 10, 11, 15, 30, 42, 45, 64, 80, 101, 106, 293 total, xii, 9, 20, 56, 68, 75, 89, 92, 113, 114, 129, 137, 139, 140, 148, 152, 205, 225, 262, 285, 286, 287

transgenic, vi, viii, 1, 4, 5, 7, 13, 17, 18, 21, 25, 26, 38, 52, 74, 77, 87, 88, 91, 104, 114, 151, 191, 192, 193, 196, 198, 199, 203, 204, 297 tricin, v, viii, 6, 15, 21, 26, 43, 51, 52, 53, 54, 55, 56, 57, 58, 59, 60, 61, 62, 63, 64, 65, 66, 67, 68, 69, 70, 71, 72, 73, 74, 75, 76, 77, 81, 83, 88, 91, 102, 173, 184

U uncondensed, 32, 122, 123, 242

V valorization, v, viii, 14, 23, 35, 46, 121, 156, 158, 159, 162, 163, 165, 166, 174, 175, 178, 179, 180, 181, 185, 186, 195, 196, 234, 239, 275 value-added, 13, 112, 128, 130, 165, 166, 171, 174, 178, 180, 191, 192, 207, 208, 216, 217, 223, 232, 234, 274 vanilla seeds, 8, 9, 12, 13 vanillic acid, 123, 169, 176, 178, 287 vascular, 3, 19, 26, 80, 126

Z zip-lignin, viii, 25, 26, 35, 36, 37, 38, 39, 41, 48