Recent Advances in Polyphenol Research, Volume 8 E-Book
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- Herausgeber: John Wiley & Sons
- Kategorie: Wissenschaft und neue Technologien
- Sprache: Englisch
Plant polyphenols are specialized metabolites that constitute one of the most common and widespread groups of natural products. They are essential plant components for adaptation to the environment and possess a large and diverse range of biological functions that provide many benefits to both plants and humans. Polyphenols, from their structurally simplest forms to their oligo/polymeric versions (i.e. tannins and lignins), are phytoestrogens, plant pigments, antioxidants, and structural components of the plant cell wall. The interactions between tannins and proteins are involved in plant defense against predation, cause astringency in foods and beverages, and affect the nutritional and health properties of human and animal food plants. This eighth volume of the highly regarded Recent Advances in Polyphenol Research series is edited by Juha-Pekka Salminen, Kristiina Wähälä, Victor de Freitas, and Stéphane Quideau, and brings together chapters written by some of the leading experts working in the polyphenol sciences today. Topics covered include: * Structure, reactivity and synthesis * Bioactivity and bioavailability * Metabolomics, targeted analysis and big data * Quality control & standardization * Biogenesis and functions in plants and ecosystems * Biomaterials & applied sciences Distilling the most recent and illuminating data available, this new volume is an invaluable resource for chemists, biochemists, plant scientists, pharmacognosists and pharmacologists, biologists, ecologists, food scientists and nutritionists.
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Table of Contents
Cover
Series Page
Title Page
Copyright Page
Dedications
Contributors
Preface
Acknowledgments
1 Lignins and Lignification
1.1 Introduction
1.2 The Monolignol Pathway and Interacting Pathways – New Lignins
1.3 Lignin Conjugates, “Clip‐Offs’ – New Discoveries, and Enhancing Levels
1.4 Features of Lignification and the Possibility of New Polymerization Pathways
1.5 The Case for Model Studies and Synthesis
1.6 New or Improved Analytics
1.7 Conclusions and Opportunities
Acknowledgments
References
2 Synthesis of Epigallocatechin Gallate, Nobiletin, and Their Derivatives for Chemical‐Biological Studies
2.1 Synthetic Investigations of Catechin Derivatives
2.2 Synthesis and Application of Fluorescent Catechin Probes
2.3 Generation of Catechin Antibody
2.4 PET Imaging of Biodistribution of Catechin
2.5 Practical Synthesis of Nobiletin
2.6 Derivatization of Desmethyl Nobiletins
2.7 PET Imaging of Biodistribution of Nobiletin
2.8 Synthesis and Application of Fluorescent Nobiletin Probe
2.9 Conclusions
References
3 Procyanidins in the Onset and Progression of Colorectal Cancer
3.1 Introduction
3.2 Procyanidins: Chemistry and Metabolism
3.3 Procyanidins and CRC: Epidemiological Evidence
3.4 Procyanidins and CRC: Rodent Studies
3.5 Procyanidins and CRC: Mechanisms of Action
3.6 Conclusions and Open Questions
Acknowledgments
Conflict of Interest Disclosure
References
4 The Potential of Low Molecular Weight (Poly)phenol Metabolites for Attenuating Neuroinflammation and Treatment of Neurodegenerative Diseases
4.1 Introduction: Neurodegenerative Disorders, Dietary (Poly)phenols and Neuroinflammation
4.2 (Poly)phenols: Metabolism and Distribution
4.3 (Poly)phenol Metabolites and Their Brain Permeability
4.4 LMW (Poly)phenol Metabolites as Effectors for Attenuating Neuroinflammation
4.5 Concluding Remarks
Acknowledgments
References
5 Deciphering Complex Natural Mixtures through Metabolome Mining of Mass Spectrometry Data
5.1 Introduction
5.2 Materials and Methods
5.3 Results and Discussion
5.4 Current Limitations
5.5 Conclusions
5.6 Outlook
Acknowledgments
References
6 Application of MS‐Based Metabolomics to Investigate Biomarkers of Apple Consumption Resulting from Microbiota and Host Metabolism Interactions
6.1 Introduction
6.2 Materials and Methods
6.3 Results and Discussion
6.4 Conclusion
Acknowledgments
Funding
References
7 Non‐Extractable Polyphenols Should be Systematically Included in Polyphenol Analysis
7.1 Introduction: The Concept of Non‐Extractable Polyphenols
7.2 Analysis of Non‐Extractable Polyphenols
7.3 Why Should Non‐Extractable Polyphenols be Systematically Included in Polyphenol Analysis?
7.4 Relevance of the Determination of Non‐Extractable Polyphenols in Quality Control
7.5 Perspectives
References
8 Template‐Mediated Engineering of Functional Metal–Phenolic Complex Coatings
8.1 Introduction
8.2 Template‐Mediated Techniques to Deposit MPNs
8.3 MPN Film Properties
8.4 MPN Surface Interactions and Applications
8.5 Upscaling Considerations and Challenges
8.6 Method Automation: Possibilities and Outlook
8.7 Conclusions
References
9 Highly Efficient Production of Dihydroflavonol 4‐Reductases in Tobacco Cells and Refinement of the BuOH‐HCl Enzymatic Assay
9.1 Introduction
9.2 Results
9.3 Materials and Methods
9.4 Discussion
Acknowledgements
References
10 A Long and Winding Road
10.1 Introduction
10.2 The Importance of R2R3Myb Transcription Factors (TFs) in the Regulation of Phenylpropanoid Metabolism in Plants
10.3 The Role of bHLH Proteins in the Regulation of Phenylpropanoid Metabolism
10.4 The Role of the WDR in the MBW Complex in the Regulation of Polyphenol Metabolism
10.5 Additional Factors Regulating Transcriptional Controlof the MBW Complex
10.6 Conclusions
Acknowledgments
References
11 Analysis of Proanthocyanidins in Food Ingredients by the 4‐(Dimethylamino)cinnamaldehyde Reaction
11.1 Introduction
11.2 Background on the 4‐(Dimethylamino)cinnalmaldehyde (DMAC) Reaction with PACs
11.3 Mechanism of the Acid‐Catalyzed DMAC Reaction with PACs
11.4 Absorption and Emission Spectra of the DMAC Reaction Products
11.5 Standards for the DMAC Reaction and Accuracy of the Method
11.6 Interaction of PAC‐DMAC Reaction Products with Extra‐Intestinal Pathogenic
Escherichia coli
11.7 Conclusion
References
12 Reactions of Ellagitannins Related to Their Metabolism in Higher Plants
12.1 Introduction
12.2 Structural Variety of Ellagitannin Acyl Groups
12.3 Reactions of the DHHDP Group
12.4 Decomposition of 1,4‐DHHDP‐α‐d‐glucose
12.5 Amariin as a Precursor of Geraniin
12.6 Triterpenoid HHDP Esters in
Castanopsis sieboldii
12.7 Highly Oxidized Ellagitannins in
Carpinus japonica
12.8 Similarity of Catechin Oxidation to Oxidation of Methyl Gallate
12.9 Production Mechanism of DHHDP and HHDP
12.10 Oxidative Degradation of Ellagitannins
12.11 Conclusions
References
Index
End User License Agreement
List of Tables
Chapter 2
Table 2.1 Inhibition of influenza A viral infectivity toward MDCK cells.
Chapter 3
Table 3.1 Epidemiological studies evaluating the preventive effects of PC o...
Table 3.2 Rodent studies evaluating the preventive effects of PC or PC‐rich...
Chapter 4
Table 4.1 LMW (poly)phenol metabolites resulting from dietary sources rich ...
Table 4.2 LMW (poly)phenol metabolites:
In silico
studies for passive perme...
Chapter 5
Table 5.1 Overview of key metabolome mining and annotation tools mentioned ...
Chapter 6
Table 6.1 List of metabolites found in biological fluids after both acute a...
Chapter 7
Table 7.1 Examples of chemical hydrolysis for NEPP release in vegetal mater...
Table 7.2 Examples of enzymatic hydrolysis for NEPP release in vegetal mate...
Table 7.3 Examples of chromatography conditions for NEPP hydrolyzates in a ...
Table 7.4 Effect of processing on NEPP content of some vegetable samples.
Chapter 8
Table 8.1 Summary of recent studies of MPN coatings on templates and bioint...
Chapter 9
Table 9.1 Primers.
Chapter 11
Table 11.1 Content of PACs determined using the 4‐(dimethylamino)cinnamalde...
List of Illustrations
Chapter 1
Figure 1.1 Biosynthetic pathways feeding into lignification. The primary bio...
Figure 1.2 Aldehyde‐containing lignins. (a) A model of a poplar lignin.(...
Figure 1.3 C‐lignin, derived from caffeyl alcohol. (a) Lignification using c...
Figure 1.4 Tricin and hydroxystilbenes in lignins. Tricin was first discover...
Figure 1.5 Zip‐Lignins from lignification with monolignol ferulate (ML‐FA) c...
Figure 1.6 Synthesis of acetaminophen (paracetamol, Tylenol®), from fossil
v
...
Figure 1.7 Composition‐extreme Arabidopsis mutants with introduced hydroxyci...
Figure 1.8 Aspects of radical generation, radical coupling, and lignificatio...
Figure 1.9 Radical coupling of catechols and catechol monomers
vs
Diels–Alde...
Figure 1.10 Lignin 4–
O
–5 units, appropriate model compounds, and their hydro...
Figure 1.11 DFRC and mild acidolysis, orthogonal methods for determining hyd...
Chapter 2
Figure 2.1 Structures of EGCg (
1
), DOEGCg (
2
), and APDOEGCg (
3
).
Scheme 2.1 Condensation of A and B rings and incorporation of amino linker m...
Scheme 2.2 Stereoselective construction of the
cis
‐benzopyran ring. Reagents...
Scheme 2.3 Conversion to probe precursor APDOEGCg (
3
). Reagents and conditio...
Scheme 2.4 Synthesis of the fluorescein probe
19
from APDOEGCg (
3
). Reagents...
Figure 2.2 Intracellular localization of APDOEGCg‐TG (
19
) in HUVEC. Conditio...
Scheme 2.5 Conjugation of
4
to HSA carrier protein
21
via glutaraldehyde lin...
Scheme 2.6 Rapid synthesis of PET probe
24
from EGCg (
1
). Reagents and condi...
Figure 2.3 Whole‐body PET imaging of the distribution of intravenously injec...
Figure 2.4 Structures of nobiletin (
25
) and sudachitin (
26
).
Scheme 2.7 Preparation of A‐ring acetophenone
33
and B‐ring acyl donor. Reag...
Scheme 2.8 Completion of total synthesis of nobiletin (
25
) and regio‐selecti...
Figure 2.5 Structure of nobiletin (
25
) and its desmethyl derivatives (
26
,
39
Scheme 2.9 Synthetic route to sudachitin (
26
). Reagents and conditions: (a) ...
Scheme 2.10 Rapid formation of PET probe 5‐[
11
C]nobiletin (
25a
) from
38
. Rea...
Figure 2.6 PET imaging of the accumulation of 5‐[
11
C]nobiletin (
25a
) in rat ...
Scheme 2.11 Synthesis of nobiletin probes
54
and
55
from the versatile precu...
Chapter 3
Figure 3.1 Chemical structure of PC: (a) monomeric flavan‐3‐ol (−)‐epicatech...
Figure 3.2 Potential mechanisms potentially involved in the anti‐CRC actions...
Chapter 4
Figure 4.1 (Poly)phenols route for absorption and metabolism in the human ga...
Figure 4.2 (Poly)phenol catabolism into low molecular weight (LMW) metabolit...
Figure 4.3 Different ways that LMW (poly)phenol metabolites could cross the ...
Figure 4.4 Possible molecular mechanisms of LMW (poly)phenols attenuation of...
Chapter 5
Figure 5.1 The chemical and structural annotation of complex natural mixture...
Figure 5.2 (a) Principal coordinate analysis (PCoA) plot based on Bray–Curti...
Figure 5.3 MolNetEnhancer provides direct insight into the non‐volatile chem...
Chapter 6
Figure 6.1 Kinetic curves of selected metabolites in plasma and urine, with ...
Figure 6.2 Chemical structures of key metabolites found in biological fluids...
Figure 6.3 Boxplot representing the cumulative excretion in urine (intensity...
Figure 6.4 Boxplot representing the cumulative excretion in urine (intensity...
Figure 6.5 Boxplot representing the cumulative excretion in urine (intensity...
Figure 6.6 Boxplot representing the cumulative excretion in urine (intensity...
Figure 6.7 Boxplot representing the cumulative excretion in urine (intensity...
Figure 6.8 Boxplot representing the cumulative excretion in urine (intensity...
Figure 6.9 A simplified diagram of the metabolic pathways for the main class...
Figure 6.10 Heat map of pairwise correlations between urine metabolites and ...
Chapter 7
Figure 7.1 Common procedure for polyphenol analysis.
Chapter 8
Figure 8.1 (a) Overview of MPN formation and deposition via discrete assembl...
Figure 8.2 (a) Overview of discrete assembly template‐mediated fabrication o...
Figure 8.3 Comparative overview of (a) discrete assembly and (b) multistep m...
Figure 8.4 Continuous MPN assembly involving rust from a nail as an iron sou...
Figure 8.5 (a) Oxidation‐mediated coordination (OMC) assembly employing ROS ...
Figure 8.6 (a) Overview of MPN formation via a spray‐coating process.(b)...
Figure 8.7 Examples of MPNs used as coating materials at biointerfaces: (a) ...
Figure 8.8 (a) Percentage of apoptotic HeLa and MB231 cells after treatment ...
Figure 8.9 (a) PET/computed tomography image of an
in vivo
model (smal...
Figure 8.10 (a) Extraction efficiency and mass of uranium from seawater by t...
Figure 8.11 Overview of the various steps involved in washing MPN particles ...
Figure 8.12 Template particle coating via immersive polymer assembly: (a) im...
Figure 8.13 Overview of various fabrication techniques involving microfluidi...
Figure 8.14 MPN formation via flash complexation, with complex formation fac...
Chapter 9
Figure 9.1 Crude extracts (a) of agroinfiltrated
N. benthamiana
leaves expre...
Figure 9.2 pH dependence of two gerbera DFRs (a), dependence on substrate co...
Figure 9.3 Linearity of the GDFR1‐3 reaction with 10 μL of tobacco extract (...
Figure 9.4 Correspondence of dihydroflavonol consumption (by HPLC) and butan...
Figure 9.5 Correspondence of the peak area sum of butanoylated Pg (a), Cy (b...
Figure 9.6 Patterns of substrate use by DFR enzymes heterologously produced ...
Chapter 10
Figure 10.1 Simplified overview of phenylpropanoid metabolic pathway in plan...
Figure 10.2 Phylogenetic relationships of phenylpropanoid‐related R2R3MYB TF...
Figure 10.3 Response of
Physcomitrella patens
wild‐type and knockout mutants...
Figure 10.4 Phylogenetic tree comparing the amino acid sequences of the DNA ...
Figure 10.5 Phylogenetic analysis of plant bHLH proteins belonging to subgro...
Figure 10.6 Muscle alignment of bHLH proteins. Gene IDs/accession numbers of...
Figure 10.7 The effect of mutations in the genes encoding bHLH‐1 (Delila) an...
Chapter 11
Figure 11.1 Chemical structures of 4‐(dimethylamino)cinnamaldehyde (a), cinn...
Figure 11.2 Proposed reaction mechanism for the formation of the open ring P...
Figure 11.3 MALDI‐TOF mass spectrum in positive reflectron mode for the reac...
Figure 11.4 MALDI‐TOF mass spectrum in positive reflectron mode for the reac...
Figure 11.5 MALDI‐TOF mass spectrum in positive reflectron mode for unreacte...
Figure 11.6 MALDI‐TOF mass spectrum in positive reflectron mode for unreacte...
Figure 11.7 MALDI‐TOF mass spectrum in positive reflectron mode for the tetr...
Figure 11.8 UV‐visible spectrum of isolated DMAC reaction products, showing ...
Figure 11.9 Change in absorbance at 559 and 640 nm during the DMAC reaction ...
Figure 11.10 Change in absorbance at 640 nm during the DMAC reaction of proc...
Figure 11.11 Absorbance at 640 nm over 60 minutes after reaction of 4‐(dimet...
Figure 11.12 Variation of time required to reach maximum absorbance (t
max
) u...
Figure 11.13 Calibration curves for catechin, procyanidin A2, procyanidin B2...
Chapter 12
Figure 12.1 Enzymatic production of an ellagitannin (
2
) from pentagalloyl gl...
Figure 12.2 Carboxylic acid forms of ellagitannin acyl groups. Comparisons b...
Figure 12.3 Chemical conversions of geraniin (
5
) to produce naturally occurr...
Figure 12.4 Mechanism of DHHDP redox disproportionation.
Figure 12.5 Production of ellagic acid (
3
) from the DHHDP group of furosin (
Figure 12.6 A potential mechanism describing the redox disproportionation of...
Figure 12.7 A potential mechanism for the reaction of ascorbic acid with an ...
Figure 12.8 Degradation of the DHHDP group of nupharanin (
8
) in pH 6 citrate...
Figure 12.9 Redox disproportionation of the 3,6‐DHHDP group of amariin (
9
) i...
Figure 12.10 Reactions of DHHDP esters of 28‐
O
‐glucosyl 2α,3β,23,24‐tetrahyd...
Figure 12.11 Reactions of carpinins F (
13
) in 1% H
2
SO
4
or H
2
O at elevated te...
Figure 12.12 Increased production of isocarpinins A (
15
) from carpinins F (
1
...
Figure 12.13 Oxidation of methyl gallate (
18
) with CuCl
2
, and subsequent rea...
Figure 12.14 Two possible routes for producing HHDP from galloyl groups. Das...
Figure 12.15 Enzymatic oxidation of pedunculagins (
22
) in the presence of ca...
Figure 12.16 Enzymatic oxidation of vescalagin (
25
) in the presence of catec...
Figure 12.17 Oxidative degradation of vescalagin (
25
) by
Lentinula edodes
.
Guide
Cover Page
Series Page
Title page
Copyright Page
Dedications
Contributors
Preface
Acknowledgments
Table of Contents
Begin Reading
Index
WILEY END USER LICENSE AGREEMENT
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Recent Advances in Polyphenol Research
A series for researchers and graduate students whose work is related to plant phenolics and polyphenols, as well as for individuals representing governments and industries with interest in this field. Each volume in this biennial series focuses on several important research topics in plant phenols and polyphenols, including chemistry, biosynthesis, metabolic engineering, ecology, physiology, food, nutrition, and health.
Volume 8 Editors:Juha‐Pekka Salminen (University of Turku, Finland), Kristiina Wähälä (University of Helsinki, Finland), Victor de Freitas (University of Porto, Portugal), and Stéphane Quideau (University of Bordeaux, France)
Series Editor‐in‐Chief:Stéphane Quideau (University of Bordeaux, France)
Series Editorial Board:Oyvind Andersen (University of Bergen, Norway)Denis Barron (Nestlé Research, Lausanne, Switzerland)Luc Bidel (INRAE, Montpellier, France)Véronique Cheynier (INRAE, Montpellier, France)Catherine Chèze (University of Bordeaux, France)Gilles Comte (University of Lyon, France)Fouad Daayf (University of Manitoba, Winnipeg, Canada)Olivier Dangles (University of Avignon, France)Kevin Davies (Plant & Food Research, Palmerston North, New Zealand)Maria Teresa Escribano‐Bailon (University of Salamanca, Spain)Sylvain Guyot (INRAE, Rennes, France)Ann E. Hagerman (Miami University, Oxford, Ohio, USA)Heidi Halbwirth (Vienna University of Technology, Austria)Amy Howell (Rutgers University, Chatsworth, New Jersey, USA)Victor de Freitas (University of Porto, Portugal)Johanna Lampe ((Fred Hutchinson Cancer Research Center, Seattle, Washington, USA)Vincenzo Lattanzio (University of Foggia, Italy)Stephan Martens (Fondazione Edmund Mach, IASMA, San Michele all'Adige, Italy)Nuno Mateus (University of Porto, Portugal)Fulvio Mattivi (University of Trento, Italy)Jess Reed (University of Wisconsin‐Madison, USA)Annalisa Romani (University of Florence, Italy)Erika Salas (Autonomous University of Chihuahua, Chihuahua, Mexico)Juha‐Pekka Salminen (University of Turku, Finland)Pascale Sarni‐Manchado (INRAE, Montpellier, France)Celestino Santos‐Buelga (University of Salamanca, Spain)Kathy Schwinn (Plant & Food Research, Palmerston North, New Zealand)Karl Stich (Vienna University of Technology, Austria)David Vauzour (University of East Anglia, Norwich, UK)Kristiina Wähälä (University of Helsinki, Finland)Kumi Yoshida (Nagoya University, Japan)Kazuhiko Fukushima (Nagoya University, Japan)
Recent Advances in Polyphenol Research
Volume 8
Edited by
Juha‐Pekka Salminen
Professor, Natural Compound Chemistry
Department of Chemistry
University of Turku, Finland
Kristiina Wähälä
Professor, Organic Chemistry
Faculty of Medicine and Faculty of Science
University of Helsinki, Finland
Victor de Freitas
Professor, Food Chemistry
Chemistry and Biochemistry Department, Faculty of Sciences
University of Porto, Portugal
Stéphane Quideau
Professor, Organic and Bioorganic Chemistry
Institut des Sciences Moléculaires, CNRS‐UMR 5255
University of Bordeaux, Talence, France & Institut Universitaire de France, Paris, France
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Library of Congress Cataloging‐in‐Publication Data applied for
ISBN: 9781119844761ISSN: 2474‐7696
Cover Design: WileyCover Images: The River Aura and the Turku Cathedral are the landmarks of the city of Turku, the venue for the XXX International Conference on Polyphenols, hosted by the Natural Chemistry Research Group at the University of Turku, Finland – The Onagraceae plant family produces the largest ellagitannins found to date in the plant kingdom: the Natural Chemistry Research Group found the undecameric ellagitannin first time in Oenothera biennis – The Island of Ruissalo in Turku harbors the most famous oak forest in Finland. Together with other deciduous tree species oaks produce a great color show with their autumn coloration. The two chromatograms on the top of the oak forest show how the group‐specific LC‐MS methodology developed by the Natural Chemistry Research group can visualize the chromatographic fingerprints of both terminal and extension units of procyanidin and prodelphinidin units present in plant proanthocyanidins.
All rights of the photos belong to Vesa Aaltonen (first photo), Marc Johnson and Kari Loikas (second photo), and Juha‐Pekka Salminen (third photo). In addition, the cover includes the official logo of the ICP2020TURKU, designed by Juha Harju.
Dedications
In memoriam
Prof. Takashi Yoshida (born in 1939, deceased on 1 May 2021) was a Professor of Medicinal Plants Chemistry at Okayama University, Japan, from 1993 to 2005. I worked with him as a research associate in the same laboratory. He also worked at the College of Pharmacy, Matsuyama University, Japan, as a Professor of Pharmacognosy from 2006 to 2012 after his departure from Okayama University in 2005. His research interests included the isolation and structural determination of ellagitannins and related polyphenols, as well as terpenoids in medicinal plants and foods in Japan, China, South‐East Asian, and South American countries and the examination of their physiological activities. His scientific activity is documented in 261 papers on the isolation and structure elucidation of tannins and terpenoids in many medicinal plants and foods, as well as their diverse pharmacological properties, such as anti‐cancer, anti‐Helicobacter pylori, antileishmanial, and anti‐methicillin‐resistant Staphylococcus aureus activities. He received the Tannin Award at the Phytochemical Society of North America Annual Meeting in 2008 (Philadelphia, USA) and the Groupe Polyphenol Medal at the 8th Tannin Conference in 2014 (Nagoya, Japan) for his achievements in the field of polyphenolic natural products. He worked with many wonderful collaborators worldwide, including pharmacologists, biochemists, and microbiologists. He was respected and loved around the world for his caring friendship, and I have the deepest respect for him as a researcher and educator.
Professor Hideyuki Ito
Okayama Prefectural University, Japan
Prof. Hidetoshi Yamada (born in Ehime, Japan, on 4 August 1962, deceased on 23 November 2019) studied chemistry at Osaka City University, obtaining a bachelor’s degree in 1985 and a master’s degree in 1987. He was appointed Assistant Professor at Tokushima Bunri University, working with the late Professor Mugio Nishizawa. He received his PhD degree for the synthesis and structural revision of osladin, a sweet saponin. In 1997, he moved to Kwansei Gakuin University as Associate Professor and was promoted Full Professor in 2004. The key motif of his research was curiosity, centering particular attention on the conformations of carbohydrates, which were his familiar molecules from his early career. Interest in oligosaccharides led him to develop several viable glycosylation protocols, including thermal glycosylation, which allowed him to achieve the synthesis of L‐rhamnose‐based cyclodextrin. He and Mugio‐sensei named the molecule “cycloawaodorin,” a facetious trivial name associated with Tokushima locality. Through these endeavors, he had a keen interest in the conformationally flipped glucosyl donors. An ingenious design of flipped donor by bridging allowed the β‐selective glycosylation without aid of the neighboring‐group participation and further led to the preparation of the smallest cyclodextrin. Motivated by the presence of inverted glucose motifs, he stumbled into the synthesis of ellagitannin‐class polyphenols, featuring D‐glucose as a core extensively esterified by digalloyl groups. Facing such a tremendous diversity, he was as ever positive to make a slogan, “Let us synthesize these natural products all.” To tackle this formidable challenge, he focused on two key motifs embedded within these complex structures: (1) hexahydroxydiphenoyl (HHDP) diesters based on two gallic acids directly C─C linked together, and (2) C─O linked digallates for which two gallic acids are linked through an oxygen atom. For the former HHDP motif, he exploited intramolecular oxidative biaryl coupling between gallates appended on a glucose scaffold, where CuCl2 and n‐BuNH2 were identified as effective reagents. For the latter O‐linked digallate motif, ortho‐quinone mono‐acetal was used as a platform to undergo an oxa‐Michael addition/elimination sequence. His chemical synthesis study opened a door to flexible, comprehensive access to the ellagitannin molecules as pure entities.
Prof. Toshiyuki Kan (born in Hokkaido, Japan, on 15 February 1964, deceased on 24 July 2021) studied chemistry at Hokkaido University, where he obtained his bachelor’s degree (1986) and master’s degree (1988). He received his PhD degree in 1993 under the guidance of Prof. Haruhisa Shirahama, working on the total synthesis of grayanotoxin III, a poisonous diterpene with a highly complex structure. After working as a researcher at Suntory Institute for Bioorganic Research with Prof. Yasufumi Ohfune (1993–1996), he was appointed Assistant Professor at University of Tokyo with Prof. Tohru Fukuyama in 1996 and was promoted as Associate Professor in 2003. In 2004, he moved to the University of Shizuoka as Full Professor. His research focus was complex natural product synthesis, as represented by grayanotoxin III (diterpenoid, skeletal complexity) and ecteinascidin 743 (alkaloid, skeletal/functional complexity). In addressing the synthesis of complex natural products, he was continuously exposed to care for the management of multiple functionalities, and he became able to come up with survival tactics or even strategies to achieve the total synthesis. Among others is the design of a new protecting group, “nosyl (Ns),” which served as bases for his alkaloid syntheses, and now one of the standard choices for protecting amino groups. Upon moving to his last destination, Shizuoka, one of the major tea and orange localities in Japan, he decided to study the synthesis and chemical biology of tea‐ and citrus‐derived polyphenols, including chafurosides A and B (black tea), epigallocatechin gallate (EGCg), and nobiletin. Here the nosyl group was proven to work for protecting phenols, endorsing a theanine synthesis via otherwise difficult biomimetic oxidative dimerization. He designed molecular probes for catechins and nobiletin, fluorescein, and PET probes, serving for in vivo imaging and antibody generation as well. He further addressed the syntheses of sesamin and sesaminol by an organocatalytic process and biomimetic construction of the furofuran skeleton and of hybrid‐type polyphenols, hedyotol A, princepin, and sophoraflavanone H by exploiting C─H insertion reactions. His study highlighted and gained insights into the traditional products indigenous to Shizuoka through chemical synthesis and chemical biology. He delivered a plenary lecture on “Total Synthesis of Hybrid Type Polyphenols” at the XXX International Conference on Polyphenols in Turku, Finland, in July 2021, which turned out to be his last lecture. The initial seven slides were a tribute to his friend, Hidetoshi Yamada. In many people’s memory, they both would be remembered by their energy, courage, spirit in research, determination and leadership in society, and friendship and warm‐hearted attitude toward everyone. They were intimate friends, sharing genuine scientific interests in complex natural product synthesis. Here they are (Hidetoshi on the left and Toshiyuki on the right) with their relaxed smiles like naughty kids, enjoying fine food and drinks!?! 合掌
Keisuke Suzuki
Professor Emeritus, Tokyo Institute of Technology, Japan
Contributors
Sara M. AbdouDepartment of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Helsinki, Finland
Sharmin AhamedDepartment of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Helsinki, Finland
Nick W. AlbertPlant & Food Research, Palmerston North, New Zealand
Emilia Alfaro‐ViquezReed Research Group, Department of Animal and Dairy Sciences, University of Wisconsin‐Madison, Madison, WI, USA
Daisuke AndoGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USACurrent address: Institute of Wood Technology, Akita Prefectural University, Noshiro, Japan
Tomohiro AsakawaDepartment of Fisheries‐Food Science, Tokai University, Shizuoka, Japan
Enrique Báez‐GarcíaInstitute of Food Science, Technology and Nutrition, Spanish Research Council (ICTAN‐CSIC), Madrid, SpainTecnológico Nacional de Mexico/Instituto Tecnológico de Tepic, Tepic, Nayarit, Mexico
Hany BashandyDepartment of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Helsinki, FinlandDepartment of Genetics, Cairo University, Giza, Egypt
Andrew BirminghamComplete Phytochemical Solutions LLC, Cambridge, WI, USA
Rafael CarechoNOVA Medical School, Faculdade de Ciências Médicas, Universidade NOVA de Lisboa, Lisboa, Portugal
Diogo CarregosaNOVA Medical School, Faculdade de Ciências Médicas, Universidade NOVA de Lisboa, Lisboa, Portugal
Frank CarusoDepartment of Chemical Engineering, The University of Melbourne, Parkville, VIC, Australia
Mingjie ChenGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USACurrent address: Institute of Microbiology, Guangdong Academy of Sciences, Guangzhou, China
José C. del RíoInstituto de Recursos Naturales y Agrobiología de Sevilla (IRNAS), CSIC, Seville, Spain
Pieter C. DorresteinCollaborative Mass Spectrometry Innovation Center, Skaggs School of Pharmacy and Pharmaceutical Sciences, University of California, San Diego, La Jolla, CA, USA
Cláudia Nunes dos SantosNOVA Medical School, Faculdade de Ciências Médicas, Universidade NOVA de Lisboa, Lisboa, Portugal
Madeleine ErnstSection for Clinical Mass Spectrometry, Department of Congenital Disorders, Danish Center for Neonatal Screening, Statens Serum Institut, Copenhagen, Denmark
Daniel Esquivel‐AlvaradoReed Research Group, Department of Animal and Dairy Sciences, University of Wisconsin‐Madison, Madison, WI, USA
Alexis EugeneGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Makoto InaiSchool of Pharmaceutical Sciences, University of Shizuoka, Shizuoka, Japan
Yi JuDepartment of Chemical Engineering, The University of Melbourne, Parkville, VIC, Australia
Toshiyuki KanSchool of Pharmaceutical Sciences, University of Shizuoka, Shizuoka, Japan
Kyo Bin KangCollege of Pharmacy, Sookmyung Women’s University, Seoul, Korea
Iris F. KappersLaboratory of Plant Physiology, Plant Sciences Group, Wageningen University and Research Wageningen, the Netherlands
Steven D. KarlenGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Hoon KimGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Abigail KramschusterComplete Phytochemical Solutions LLC, Cambridge, WI, USA
Christian G. KruegerReed Research Group, Department of Animal and Dairy Sciences, University of Wisconsin‐Madison, Madison, WI, USAComplete Phytochemical Solutions LLC, Cambridge, WI, USA
Wu LanGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USACurrent address: State Key Laboratory of Pulp and Paper Engineering, South China University of Technology, Guangzhou, China
Leta L. LanducciGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Jie LiDepartment of Biochemistry and Metabolism, John Innes Centre, Norwich Research Park, Colney, Norwich, UK
Yanding LiGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USACurrent address: BeiGene, Zhongguancun Life Science Park, Changping District, Beijing, China
Sarah LiuGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Fachuang LuGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Gerardo G. MackenzieDepartment of Nutrition, University of California‐Davis, Davis, CA, USA
Daniela MarquesNOVA Medical School, Faculdade de Ciências Médicas, Universidade NOVA de Lisboa, Lisboa, Portugal
Cathie MartinDepartment of Biochemistry and Metabolism, John Innes Centre, Norwich Research Park, Colney, Norwich, UK
Roosa MatomäkiDepartment of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Helsinki, Finland
Saku MattilaDepartment of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Helsinki, Finland
Fulvio MattiviDepartment of Cellular, Computational and Integrative Biology (CIBIO), University of Trento, Trento, ItalyDepartment of Food Quality and Nutrition, Fondazione Edmund Mach, Research and Innovation Centre, San Michele all’Adige, Italy
Marnix H. MedemaBioinformatics Group, Plant Sciences Group, Wageningen University and Research, Wageningen, the Netherlands
Lorenzo MolloDepartment of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Helsinki, FinlandDepartment of Environmental and Life Sciences, Laboratory of Plant and Algae Physiology, Università politecnica delle Marche, Ancona, Italy
NuoendagulaGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Patricia I. OteizaDepartment of Nutrition, University of California‐Davis, Davis, CA, USA
Daniel PapenbergBioinformatics Group, Plant Sciences Group, Wageningen University and Research, Wageningen, the NetherlandsLaboratory of Plant Physiology, Plant Sciences Group, Wageningen University and Research, Wageningen, the Netherlands
Jara Pérez‐JiménezInstitute of Food Science, Technology and Nutrition, Spanish Research Council (ICTAN‐CSIC), Madrid, Spain
John RalphGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USADepartment of Biochemistry, University of Wisconsin, Madison, WI, USA
Sally A. RalphThe US Forest Products Laboratory, One Gifford Pinchot Drive, Madison, WI, USA
Jess D. ReedReed Research Group, Department of Animal and Dairy Sciences, University of Wisconsin‐Madison, Madison, WI, USAComplete Phytochemical Solutions LLC, Cambridge, WI, USA
Jorge RencoretInstituto de Recursos Naturales y Agrobiología de Sevilla (IRNAS), CSIC, Seville, Spain
J.J. RichardsonDepartment of Chemical Engineering, The University of Melbourne, Parkville, VIC, Australia
Simon RogersSchool of Computing Science, University of Glasgow, Glasgow, UK
Sonia G. Sáyago‐AyerdiTecnológico Nacional de Mexico/Instituto Tecnológico de Tepic, Tepic, Nayarit, Mexico
Canan SenerGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Rebecca A. SmithGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Steve SpoljaricDepartment of Chemical Engineering, The University of Melbourne, Parkville, VIC, Australia
Takashi TanakaGraduate School of Biomedical Sciences, Nagasaki University, Nagasaki, Japan
Teemu H. TeeriDepartment of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Helsinki, Finland
Vitaliy I. TimokhinGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Maria M. UlaszewskaDepartment of Food Quality and Nutrition, Fondazione Edmund Mach, Research and Innovation Centre, San Michele all’Adige, Italy
PROMEFA Facility, San Raffaele Scientific Institute, Center for Omics Sciences, Milan, Italy
Justin J.J. van der HooftBioinformatics Group, Plant Sciences Group, Wageningen University and Research, Wageningen, the Netherlands
Koichi YoshiokaGreat Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
Wei ZhuDepartment of Nutrition, University of California‐Davis, Davis, CA, USA
Lingping ZhuDepartment of Agricultural Sciences, Viikki Plant Science Centre, University of Helsinki, Helsinki, Finland
Preface
Every 2 years, Groupe Polyphénols (GP) hosts the International Conference on Polyphenols (ICP). The anniversary XXX ICP was planned to be held in 2020 in Turku, Finland. Unfortunately, the COVID pandemic forced GP to postpone the conference by one year, although everything was already set and organized for the conference in 2020. After one year of pandemic, GP decided that the ICP2020TURKU should not be postponed further and it was successfully organized as a fully virtual conference from 13 to 15 July, 2021. This was the first ever virtual ICP hosted by GP since its foundation in 1972. Groupe Polyphénols is the world’s premier society of scientists in the fields of polyphenol chemistry, synthesis, bioactivity, nutrition, industrial applications, and ecology. Luckily, the great success of this virtual ICP encouraged GP to plan for new types of scientific activities for its members also in between the biannual ICPs. Since the ICP2020TURKU, GP has already organized the first Webinar in Polyphenols Research that will gather polyphenol scientists virtually three to four times a year to attend presentations of both established and young scientists. This approach thus also continues one of the main aims of the ICP2020TURKU by giving good opportunities to young scientists to present their recent research findings on polyphenols.
The city of Turku is a city full of history. It was the first capital of Finland, before Helsinki, and it had the first Finnish‐speaking university in Finland. The main organizers of the ICP2020TURKU, the Natural Chemistry Research Group, had planned to organize the ICP at the main campus of the University of Turku, close to the River Aura and the Turku Cathedral (see the front cover). Other history‐oriented activities such as the gala dinner in the medieval Turku Castle were also planned and booked. However, now it remains for all the ICP participants to visit Turku on a later notice, once the COVID pandemic allows.
The XXX ICP was attended by 250 registrants from 36 countries, with 105 invited and contributed presentations. This great number of presentations was achieved by parallel sessions that maximized the opportunities given to young scientists to present their work. This eighth edition of Recent Advances in Polyphenol Research has 12 chapters that represent the work of the invited speakers at the XXX ICP and reflect the depth of science in this important field of natural product chemistry. The conference included 19 sessions on structure, reactivity, and synthesis; bioactivity and bioavailability; metabolomics, targeted analysis, and big data; quality control and standardization; biogenesis and functions in plants and ecosystems; and biomaterials and applied sciences.
We owe a special thanks to Tina Ahonen from the Aboa Congress and Event Services for her professional and excellent help in the organization of the conference. The great execution of the virtual ICP would not have been possible without the help of professionals of RajuLive Ltd. that made sure that all tiny details of the virtual conference, including the presentation recordings were of prime quality. The members and students of the Natural Chemistry Research Groups deserve our sincere thanks for their huge efforts in making the intermission activities in the form of entertaining videos that were also uploaded on the social media and for helping with the practical organization on site. Finally, we thank all the participants, who took active part in the conference sessions, and initiated a lot of scientific discussion in the conference chat, both after and between the presentations. You created a warm atmosphere for the conference and made it a really enjoyable event and a great learning experience during these otherwise difficult COVID times. We think that the ICP2020TURKU will always be remembered as a special conference, but luckily only for very good and positive reasons.
Juha‐Pekka Salminen
Kristiina Wähälä
Victor de Freitas
Stéphane Quideau
Acknowledgments
The editors wish to thank all members of the “Groupe Polyphénols” Board Committee (2018–2021) for their guidance and assistance throughout this project.
Dr. Denis BarronDr. Luc BidelDr. Catherine ChèzeDr. Peter ConstabelProf. Olivier DanglesDr. Kevin DaviesProf. M Teresa Escribano BailonProf. Victor de FreitasProf. Kazuhiko FukushimaDr. David GangDr. Sylvain GuyotProf. Ann E. HagermanDr. Irene Mueller‐HarveyProf. Stéphane QuideauProf. Jess ReedDr. Erika SalasProf. Juha‐Pekka SalminenProf. Kristiina Wähälä
1Lignins and Lignification: New Developments and Emerging Concepts
John Ralph1,2, Hoon Kim1, Fachuang Lu1, Rebecca A. Smith1, Steven D. Karlen1, Nuoendagula1, Koichi Yoshioka1, Alexis Eugene1, Sarah Liu1, Canan Sener1, Daisuke Ando1,*, Mingjie Chen1,**, Yanding Li1,***, Leta L. Landucci1, Sally A. Ralph3, Vitaliy I. Timokhin1, Wu Lan1,****, Jorge Rencoret4, and José C. del Río4
1 Great Lakes Bioenergy Research Center, The Wisconsin Energy Institute, University of Wisconsin, Madison, WI, USA
2 Department of Biochemistry, University of Wisconsin, Madison, WI, USA
3 The US Forest Products Laboratory, One Gifford Pinchot Drive, Madison, WI, USA
4 Instituto de Recursos Naturales y Agrobiología de Sevilla (IRNAS), CSIC, Seville, Spain
1.1 Introduction
In the previous volume in this series, Volume 7, Chapter 7 highlighted recent discoveries relating to the interactions between monolignol pathways with flavonoid and stilbenoid pathways producing monomers for lignification (del Río et al. 2021). The notion that lignification, the process of polymerization from monomers to the lignin polymer, may tolerate or even favor the use of monomers beyond the canonical monolignols (p‐coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol) is becoming mainstream even if there might not be universal agreement about what exactly constitutes lignification. Advances continue as ever more is revealed about the way cell wall polymers in various tissues in “natural plants” are derived and how the perturbations of genes, now in various interacting pathways, can affect lignification and the consequent composition and structure of the polymer. Although “structure” has little meaning for a polymer lacking defined repeating units of any length and possessing an overwhelming stereochemical complexity (Ralph et al. 2008), this newly revealed complexity to the composition and structure of lignin polymers, and the blurring of the definition of lignification, may seem alarming. It is worth emphasizing, however, that the process of lignification itself is a delightfully simple one involving a single, purely chemical mechanism and, accordingly, lignification is a much simpler process than the ones involved in the formation of hemicellulosic polysaccharides, for example.
Findings expanding the definition of lignin continue, even as the ramifications of the COVID‐19 pandemic have impeded research progress in general since 2020. To document this rapidly advancing field, we cover some new findings and a few of the emerging notions on lignification. So much is happening in this field that we cannot comprehensively cover even merely the work from our own labs; this chapter is best regarded as an update in which we concentrate on some of the research on a common theme that interests us. We also cover areas that often must be jettisoned from research papers, weaving in concepts that are in principle well‐known but occasionally need to be reemphasized because they have particular importance and/or may be distinctive to lignification and unfamiliar to researchers new to the field. Along with the evaluation and contemplation of “new” pathways and mechanisms, we also include minor subsections on the value and use of lignin models to understand reaction pathways, and the continued importance of developing diagnostic analytics to provide unambiguous new insight.
The chapter has been laid out in seven sections but keeping the ideas discretely under those headings has not been fully realized. Just as an example, we decided that observations on the use of monolignol conjugates in lignification needed their own section, yet much of the material could easily fit under sections preceding it. Similarly, model, synthetic, and analytical work often accompanies any discovery, but we have chosen to split one aspect out to provide some recognition for such crucial research components. Finally, there are some concepts that we wanted to convey here that simply do not fit well under the chosen headings. We trust that this will nevertheless be a readable and useful contribution despite these limitations.
1.2 The Monolignol Pathway and Interacting Pathways – New Lignins
The monolignol biosynthetic pathway produces the three canonical monolignols for lignification, p‐coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol, differing in their degrees of methoxylation ortho to the phenol (Freudenberg and Neish 1968; Sarkanen and Ludwig 1971). Some of the enzymes are quite specific, whereas others are more general in nature. As summarized in Figure 1.1, and as has been reviewed (Boerjan et al. 2003; Ralph et al. 2004b; Shi et al. 2010; Weng and Chapple 2010), the major flux through the pathway was simplified from the full metabolic grid originally considered (Dixon et al. 2001; Higuchi 2006; Matsui et al. 2000) as favored substrates and pathways through the grid have been elucidated (Humphreys et al. 1999a, 1999b; Li et al. 2000; Osakabe et al. 1999), and new steps and their enzymes continue to be discovered, as reviewed (Bonawitz and Chapple 2010; Mottiar et al. 2016; Ralph et al. 2019; Vanholme et al. 2010a, 2019a). Figure 1.1 attempts to capture the modern notion of broader lignification, integrating in phenolic components beyond the strict monolignol pathway. Perturbing the various genes along the pathway from phenylalanine to the monolignols, or even from shikimic acid and further back, not only provides a rich source of insight into the pathway processes but is also capable of producing some striking lignins. With reference to the line in Michael Chrichton’s original Jurassic Park book (Chrichton 1990) and in the movie that “Life will find a way,” plants do not simply give up and die because they find themselves unable to synthesize a monolignol. Although the very notion was once considered heresy, plants can survive, even from “instantaneous” perturbations from which they do not have the luxury of evolving, by producing a functional polymer from other available phenolic components. This is most readily evident in plants utilizing “products of truncated monolignol biosynthesis,” a term we might have first introduced in print in 2003 (Boerjan et al. 2003), to produce the lignin polymer, sometimes quite successfully, but it is also clear that nature itself has, over time, explored options well beyond just utilizing pathway intermediates and still has plenty of surprises for us. The following recent examples are illustrative but are neither unique nor exhaustive.
1.2.1 Truncated Monolignol Biosynthesis
We will update just two examples here, those of plants deficient in the last enzyme of the pathway, cinnamyl alcohol dehydrogenase (CAD), and in one of the two primary O‐methyltransferases (OMT, i.e. CCoAOMT or COMT), Figures 1.1–1.3. The result in both cases is product monomers of incomplete monolignol biosynthesis, hydroxycinnamaldehydes in the former case, and the catecholic monomers caffeyl alcohol or 5‐hydroxyconiferyl alcohol in the latter. Both produce lignin polymers that function satisfactorily in the plant and, at least in the case of the catechols, are used as significant or even sole components to fabricate natural lignins in specialized tissues such as seedcoats.
1.2.1.1 CAD Deficiency
Lignins have long been known to contain low levels of hydroxycinnamaldehyde (and hydroxybenzaldehyde) units. The characteristic lignin stain, the Wiesner or phloroglucinol stain, is somewhat specific for hydroxycinnamaldehyde endgroups (Adler et al. 1948; Pomar et al. 2002); it has always been ironic that this common stain owes its utility to the incorporation of low levels of components that are often not even acknowledged as being involved in lignification. Cinnamaldehyde endgroups might be produced in the lignin in one of three ways. First, they could result from the oxidation of cinnamyl alcohol endgroups in lignin, themselves resulting from initial dimerization reactions of monolignols, particularly coniferyl alcohol. The conundrum here is that it is quite difficult to oxidize etherified hydroxycinnamyl alcohols, the decades that the lignin might remain in a tree notwithstanding. A second is for coniferaldehyde or sinapaldehyde to be produced by oxidizing the monolignols in the lignifying region of the cell wall by the action of H2O2, for example – such oxidation with H2O2 can be demonstrated, and “always” accompanies synthetic lignin preparations in which monolignols are mixed with peroxidase and H2O2 (Kim et al. 2003; Zhao et al. 2013). These hydroxycinnamaldehyde monomers can then couple and cross‐couple into the polymer following single‐electron oxidation analogously to the polymerization of the monolignols themselves (Kim et al. 2003). This possibility comes only with a conceptual problem for researchers wanting to assert the fidelity of lignification as resulting solely from monolignols, as it comes with an implicit recognition that monomer‐substitution (by something other than a monolignol, and in fact not an “ol” at all) is already occurring during “all” lignification. Third, it is possible that monolignol biosynthesis in the cytoplasm is not fully complete, even during “normal” lignification, and that such products of incomplete monolignol biosynthesis simply make it to the wall along with the monolignols, to polymerize purely via their ability to be single‐electron‐oxidized, and subject to their subsequent chemical coupling and cross‐coupling compatibility with the lignification reactions going on around them. As long as transporters required to transport these monomers across the plasma membrane are present, or if transporters are not required as recent in silico studies suggest (Vermaas et al. 2019), getting hydroxycinnamaldehydes into the lignin by this mechanism also suffers from no conceptual difficulties in a purely chemical, combinatorial process. We can conclude that a major mechanism is via incorporation of hydroxycinnamaldehyde monomers themselves, the latter two methods, into the polymerization process because, in “natural” wild‐type plants, NMR sensitivity is such that we can usually quite easily now detect hydroxycinnamaldehydes that have been incorporated into a growing lignin chain (and are therefore internal units) in addition to the endgroups noted above (Yamamoto et al. 2020), as seen by the S′G structure in Figure 1.2e. At least in the case of CAD‐deficient plants, the third mechanism is essentially required, as production of monolignols can be severely curtailed yet, as described below, the lignin is heavily derived from polymerization from hydroxycinnamaldehydes reaching the wall, obviously having originated in that form from the cytoplasm.
Figure 1.1 Biosynthetic pathways feeding into lignification. The primary biosynthetic pathway leading to the monolignols and the two related hydroxycinnamyl alcohols, bolded and highlighted in yellow (in the colored version of this figure), is to the right and is shown as a metabolic grid as in the old days (Dixon et al. 2001; Higuchi 2006; Matsui et al. 2000); unfortunately, since modifications to the pathway flux have been discovered, it is not possible to order the intermediates in a linear fashion vertically – conversions jump rows in a way that is simply not pleasing (and, were this a circuit‐board, would represent bad design but seems unavoidable here). Where possible, the primary pathways are shown with black arrows, and the major pathways are slightly bolder, whereas minor pathways, or those that might not be completely demonstrated are in a light gray; dashed arrows represent pathways (and genes/enzymes) that are not yet known. Note that we do not try to show the pathway from p‐hydroxybenzoate, via its CoA thioester (not shown) to the monolignol p‐hydroxybenzoate (and other benzoate) conjugates in the bottom row; the long‐anticipated (Ralph 2010; Ralph et al. 2019) transferase enzyme/gene required to produce monolignol p‐hydroxybenzoates has now been identified by two groups (de Vries et al. 2022; Zhao et al. 2021). For clarity, the five types of aromatic nuclei, i.e. with the aromatic ring substitution at the p‐hydroxyphenyl, catechyl, guaiacyl, 5‐hydroxyguaiacyl, and syringyl level, in their columns are color‐coded in common. To the left are the pathways to the hydroxystilbenes and flavonoids that may also be monomers used in lignification. The gray‐ringed structures to the left of center at the bottom are the primary inputs into these pathways with the aromatics deriving from shikimic acid, via the amino‐acids phenylalanine and tyrosine (Yoo et al. 2013). A major branchpoint for all three pathways is p‐coumaroyl‐CoA, also bolded, and highlighted in pink. Enzyme abbreviations: Monolignol pathway: PAL, phenylalanine ammonia‐lyase; TAL, tyrosine ammonia‐lyase (in monocots); C4H, cinnamate 4‐hydroxylase; C3H, p‐coumarate (or p‐coumaroyl‐CoA) 3‐hydroxylase; C3′H, p‐coumaroyl shikimate 3′‐hydroxylase; CCoAOMT, caffeoyl‐CoA O‐methyltransferase; 4CL, 4‐coumarate: CoA ligase; CCR, cinnamoyl‐CoA reductase; HCT, hydroxycinnamoyl‐CoA: quinate/shikimate hydroxycinnamoyl transferase; CSE, caffeoyl shikimate esterase; F5H, ferulate 5‐hydroxylase; COMT, caffeic acid O‐methyltransferase; CAD, cinnamyl alcohol dehydrogenase; HCALDH, hydroxycinnamaldehyde dehydrogenase; PMT, p‐coumaroyl‐CoA: monolignol transferase; FMT, feruloyl‐CoA: monolignol transferase; XMT, a general hydroxycinnamoyl‐CoA: monolignol transferase; POD, peroxidase (or laccase, not shown except in flavonoid pathway); Hydroxystilbene pathway: STS, stilbene synthase; R3′H, resveratrol 3′‐hydroxylase; COMT?, the monolignol pathway COMT or another OMT; Flavonoid pathway: CHS, chalcone synthase; CHI, chalcone isomerase; FNS, flavone synthase; F3′H, flavonoid 3′‐hydroxylase; F5′H, flavonoid 5′‐hydroxylase or a dual‐function 3′/5′‐hydroxylase; FOMT, flavonoid‐O‐methyltransferase; C5′H, chrysoeriol 5′‐hydroxylase. We heartily thank Ruben Vanholme (VIB) for helping clarify the nuances of the monolignol pathway.
Figure 1.2 Aldehyde‐containing lignins. (a) A model of a poplar lignin.
Source: Ralph et al. (2019)/Elsevier.
(b) A model of a poplar CAD‐deficient lignin that incorporated mainly sinapaldehyde units.
Source: Ralph et al. (2019)/Elsevier.
(c) The aromatic regions of HSQC NMR spectra from lignins isolated from WT Medicago truncatula and (d) a CAD mutant in the same background showing that the lignin is heavily (~95%) hydroxycinnamaldehyde‐derived, i.e. that this viable plant has very little of its lignin derived from any of the canonical monolignols.
Source: Zhao et al. (2013)/with permission of Proceedings of the National Academy of Sciences.
(e) A mulberry variety (Nezumigaeshi) with normal‐colored wood (inset) and in which the NMR aldehyde region shows the typical hydroxycinnamaldehyde X and benzaldehyde SA and V) endgroups along with, as we often detect now (Van Acker et al. 2017; Yan et al. 2019), low levels of the product in which sinapaldehyde has 8–O–4‐cross‐coupled with a G‐unit in the growing polymer to become part of the internal backbone of the lignin, structure S′G, not just an endgroup.
Source: Yamamoto et al. (2020)/Oxford University Press.
(f) The same aldehyde region from the NMR of the lignin isolated from a wild mutant mulberry (Sekizaisou) discovered to have a red‐colored wood and leaves that are particularly suited for silkworm feeding that was recently revealed to be a CAD‐deficient mutant, the first reported in a hardwood.
Source: Yamamoto et al. (2020)/Oxford University Press.
Units are derived from the 8–O–4‐cross‐coupling of both sinapaldehyde and coniferaldehyde into the polymer (S′G and G′S, but not G′G); by separate experiments, it is possible to determine that sinapaldehyde cross‐couples with either guaiacyl or syringyl units, whereas coniferaldehyde cross‐couples with only syringyl units (as is also the case in vitro) (Kim et al. 2003). The green peak labeled “Z (new)” has been observed previously in spectra from our labs from aldehyde‐rich lignins and synthetic lignins incorporating coniferaldehyde but had not previously been structurally assigned. (g) The new product has recently been determined to be an unusual benzofuran Z that is clearly not a primary coupling product but arises from 8–5‐dimerization of coniferaldehyde followed by further single‐electron oxidation and then radical disproportionation to a quinone methide intermediate followed by rearomatization (K. Yoshioka, in preparation). Note: Here and in Figure 1.3, we use the standard convention by which the 3‐carbon sidechains on monolignol and lignin units are labeled α, β, and γ (from the aromatic C1 attachment), whereas the benzaldehydes and cinnamaldehydes, as well as other “oxidized units” such as ferulate and p‐coumarate, use 7, 8, and 9.
Figure 1.3 C‐lignin, derived from caffeyl alcohol. (a) Lignification using caffeyl alcohol as the sole monomer results in an almost pure homopolymer of benzodioxane units arising from β–O–4‐coupling; although C‐units can, in principle, couple with all the options open to G‐units, the β–O–4‐coupling propensity is simply overwhelming such that only low levels of other structures can be found in the polymer.
Source: Chen et al. (2012)/National Academy of Sciences.
(b) Thioacidolysis of vanilla seedcoats releases the typical isomer pair of a sole C‐monomer, with no contributions from S‐ or G‐units derived from the canonical monolignols.
Source: Chen et al. (2012)/with permission of Proceedings of the National Academy of Sciences.
(c–e) The stem, pod, and leaf tissues show S/G profiles typical of an angiosperm, with no evidence of C‐units.
Source: Chen et al. (2012)/Proceedings of the National Academy of Sciences.
(f) A caffeyl alcohol synthetic lignin (dehydrogenation polymer, DHP) is rather simple, with notable end‐units in its HSQC NMR spectrum because of its low molecular weight. The main units are benzodioxanes.
Source: Adapted from Li et al. (2018).
(g) An isolated enzyme lignin (EL) from vanilla seedcoats has a particularly simple HSQC NMR spectrum, showing essentially only benzodioxane units from β–O–4‐coupling (along with peaks from residual cellulose, C); from the original data used in figure 1a and S1C in Li et al. (2018). The inset is a picture of a sliced‐though vanilla seedpod at 10 days post‐pollination, showing the black seedcoats containing the C‐lignin.
Source: Chen et al. (2012)/with permission of National Academy of Science.
(h) Yield and selectivity of Pd/C‐catalyzed hydrogenolysis to produce arylpropanol, arylpropane, and other monomers, for the vanilla seedcoat C‐lignin here (in which the arylpropanol monomer is catechylpropanol and the minor arylpropane monomer is catechylpropane) vs spruce, birch, and high‐S poplar. The C‐lignin delivers almost 90% yields of monomers in a product that is 90% one compound, dihydrocaffeyl alcohol (= catechylpropanol); the prior record monomer yield had been from the high‐S lignin (Lan et al. 2018a; Shuai et al. 2016). Conditions: 5% Pd/C, 473 K, 15 h, 4 MPa H2.
Source: Li et al. (2019)/Royal Society of Chemistry.
Picture in the bottom right is part of the picture prepared for a cover issue (Li et al. 2018). Note: Here and in Figure 1.2
