Forestry, Faculty of
DSpace
UBCV
Mottiar, Yaseen
Doctor of Philosophy - PhD
University of British Columbia
Lignin is a phenolic polymer found predominantly in the secondary cell walls of vascular plants where it contributes to water transport, mechanical support, and plant defence. The occurrence, composition, and structure of lignin vary widely between cell types, throughout development, in response to stress, and across plant lineages. In addition to the hydroxycinnamate-derived monolignols, an array of non-canonical monomers occurs in lignin, clearly illustrating that the molecular processes underpinning lignin formation are highly flexible. This innate plasticity enables the rational design of lignins with predetermined structures and physicochemical properties. Given the biological importance of lignin and the industrial significance of plant biomass, this dissertation explores several promising areas of research in cell wall biology, plant biochemistry, and lignin engineering. The shikimate and aromatic amino acid pathways are crucial metabolic gatekeepers that lie upstream of lignin biosynthesis. In this work, chorismate was studied as an important branchpoint metabolite and evidence was uncovered for redundancy of chorismate mutase isomers in poplar (Populus trichocarpa). Next, a lignin engineering strategy that exploits this important branchpoint was developed. Introduction of a bacterial chorismate pyruvate lyase into transgenic hybrid poplar (Populus alba × grandidentata) was used to divert carbon flux away from phenylpropanoid biosynthesis, leading to lignin with more alkali-labile p-hydroxybenzoate pendent groups. In commelinid monocots, p-coumarate occurs analogously as lignin acylations. The discovery of these moieties in the lignin of kenaf (Hibiscus cannabinus), a eudicot species, represents a new example of convergent evolution and provides a useful tool for lignin engineering. Finally, lignification was studied in eastern leatherwood (Dirca palustris) as the woody stems of this native shrub are exceptionally flexible. In addition to an unusual composition and distribution of lignin, this work uncovered a lignin-deficiency in the middle lamella that asks us to revisit unresolved questions in plant cell wall biology.
EXPLORING THE PLASTICITY OF LIGNIFICATION by Yaseen Mottiar B.Sc., University of Ottawa, 2008 B.A.Sc., University of Ottawa, 2008 M.Sc., University of Toronto, 2012 A thesis submitted in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Forestry) THE UNIVERSITY OF BRITISH COLUMBIA (Vancouver) August 2021 © Yaseen Mottiar, 2021 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the dissertation entitled: Exploring the plasticity of lignification submitted by Yaseen Mottiar in partial fulfillment of the requirements for the degree of Doctor of Philosophy in Forestry Examining Committee: Shawn D. Mansfield, Department of Wood Science Research Supervisor Robert D. Guy, Department of Forest and Conservation Sciences Supervisory Committee Member A. Lacey Samuels, Department of Botany Supervisory Committee Member Joerg C. Bohlmann, Michael Smith Laboratories University Examiner Michael K. Deyholos, University of British Columbia – Okanagan University Examiner Jacqueline Grima-Pettenati, University of Toulouse III, France External Examiner Additional Supervisory Committee Member: Simone D. Castellarin, Wine Research Centre Supervisory Committee Member iii Abstract Lignin is a phenolic polymer found predominantly in the secondary cell walls of vascular plants where it contributes to water transport, mechanical support, and plant defence. The occurrence, composition, and structure of lignin vary widely between cell types, throughout development, in response to stress, and across plant lineages. In addition to the hydroxycinnamate-derived monolignols, an array of non-canonical monomers occurs in lignin, clearly illustrating that the molecular processes underpinning lignin formation are highly flexible. This innate plasticity enables the rational design of lignins with predetermined structures and physicochemical properties. Given the biological importance of lignin and the industrial significance of plant biomass, this dissertation explores several promising areas of research in cell wall biology, plant biochemistry, and lignin engineering. The shikimate and aromatic amino acid pathways are crucial metabolic gatekeepers that lie upstream of lignin biosynthesis. In this work, chorismate was studied as an important branchpoint metabolite and evidence was uncovered for redundancy of chorismate mutase isomers in poplar (Populus trichocarpa). Next, a lignin engineering strategy that exploits this important branchpoint was developed. Introduction of a bacterial chorismate pyruvate lyase into transgenic hybrid poplar (Populus alba × grandidentata) was used to divert carbon flux away from phenylpropanoid biosynthesis, leading to lignin with more alkali-labile p-hydroxybenzoate pendent groups. In commelinid monocots, p-coumarate occurs analogously as lignin acylations. The discovery of these moieties in the lignin of kenaf (Hibiscus cannabinus), a eudicot species, represents a new example of convergent evolution and provides a useful tool for lignin engineering. Finally, lignification was studied in eastern leatherwood (Dirca palustris) as the woody stems of this native shrub are exceptionally flexible. In addition to an unusual composition and distribution of lignin, this work uncovered a lignin-deficiency in the middle lamella that asks us to revisit unresolved questions in plant cell wall biology. iv Résumé La lignine est un polymère phénolique trouvé principalement dans les parois cellulaires secondaires des plantes vasculaires. Elle contribue au transport de l'eau, au support mécanique et à la défense de la plante. L’apparition, la composition et la structure de la lignine varient considérablement entre les types de cellules tout au long de son développement, en réponse au stress et selon les lignages végétaux. En plus des monolignols dérivés de l’acide hydroxycinnamique, un éventail de monomères non canoniques se trouve dans la lignine, illustrant clairement que les processus moléculaires qui sous-tendent la formation des lignines sont très flexibles. Cette plasticité innée permet une conception rationnelle des lignines avec des structures et des propriétés physicochimiques prédéterminées. Compte tenu de l'importance biologique de la lignine et de l'importance industrielle de la biomasse végétale, cette thèse explore plusieurs domaines de recherche prometteurs dans la biologie des parois cellulaires, la biochimie végétale et l’ingénierie des lignines. Les voies de l’acide shikimique et des acides aminés aromatiques sont des gardiens métaboliques essentiels qui se trouvent en amont de la biosynthèse des lignines. Dans ce travail, l’acide chorismique a été étudié en tant qu’important métabolite au point de branchement en amont et des preuves ont été découvertes en faveur d’une redondance des isomères de la mutase de l’acide chorismique dans le peuplier (Populus trichocarpa). Ensuite, une stratégie d'ingénierie des lignines qui exploite cet important point de branchement a été développée. L'introduction d'une lyase d’acide chorismique et pyruvique bactérienne dans le peuplier hybride transgénique (Populus alba × grandidentata) a été utilisée pour détourner le flux de carbone de la biosynthèse des phénylpropanoïdes, conduisant à une lignine avec plusieurs des groupes d’acide parahydroxybenzoïque pendants qui sont labiles dans l’alcali. Dans les monocotylédones commélinidés, l’acide paracoumarique existe de manière analogue comme acylations de la lignine. La découverte de ces groupements dans la lignine du kénaf (Hibiscus cannabinus), une espèce d'eudicotylédones, fournit un nouvel exemple d'évolution convergente et un outil utile en ingénierie des lignines. Enfin, la lignification a été étudiée dans le bois de plomb (Dirca palustris) car les tiges ligneuses de cet arbuste indigène sont très flexibles. En plus d'une distribution atypique de lignine dans le xylème, ce travail a révélé une carence en lignine dans la lamelle moyenne, ce qui nous invite à revoir des questions non résolues dans la biologie des parois cellulaires des plantes. v Lay Summary Lignin is natural polymer found primarily in the secondary cell walls of vascular plants, including in the bulk of cells that comprise wood. Although lignin fulfils many essential roles in plants, there is also tremendous interest in manipulating lignin content, composition, and structure as the chemical stability of lignin presents a formidable challenge for the industrial processing of plant biomass. In this dissertation, the innate plasticity of lignin formation was examined, and complementary strategies for lignin engineering were developed to exploit the flexibility of lignification. This work reports new discoveries and prompts new questions in cell wall biology, plant biochemistry, and lignin engineering. These findings could inform future efforts to breed or genetically engineer plants for the sustainable industrial use of wood and plant biomass. vi Preface Chapter 2 is a reproduction of a published review article co-authored with colleagues at Ghent University (Belgium) and the University of Wisconsin–Madison (U.S.A.). Mottiar Y, Vanholme R, Boerjan W, Ralph J, Mansfield SD. Designer lignins: Harnessing the plasticity of lignification. Current Opinion in Biotechnology 2016, 37: 190–200. Publisher: Elsevier Ltd. Chapter 3 is a recently submitted manuscript co-authored with colleagues at the Oak Ridge National Laboratory (U.S.A.) and the University of Wisconsin–Madison. Dr. the performedTimothy Tschaplinski metabolites.MS analysis of the poplar -GC Dr. John Ralph helped with the NMR analysis. Chapter 4 is a recently submitted manuscript co-authored with colleagues at the University of Wisconsin–Madison and Niagara University (U.S.A.). Drs. Steven Karlen and John Ralph helped with the NMR, DFRC, GPC, and HRMS analysis. Dr. Robyn Goacher helped with the ToF-SIMS analysis. Chapter 5 is a collaborative study co-authored with colleagues at Michigan State University (U.S.A.) and the University of Wisconsin–Madison which will soon be submitted for publication. Sasha Peers and Dr. Curtis Wilkerson identified the gene, and Drs. Rebecca Smith, Steven Karlen, and John Ralph helped with NMR, DFRC, GPC, and enzyme assays. Chapter 6 is a reproduction of a published research article co-authored with colleagues at the University of Natural Resources and Life Sciences Vienna (Austria), Mississippi State University (U.S.A.), and the University of Toronto. Mottiar Y, Gierlinger N, Jeremic D, Master ER, Mansfield SD. Atypical lignification in eastern leatherwood (Dirca palustris). New Phytologist 2020, 226(3): 704–713. Publisher: John Wiley & Sons Ltd. A small portion of this dataset (FTIR, XRD, and TOF-SIMS) was previously submitted as part of my Master of Science degree at the University of Toronto. Finally, Appendix 3 is a reproduction of a published research article co-authored with a colleague at Niagara University, Dr. Robyn Goacher, who collected the ToF-SIMS data and helped with experimental design. Goacher RE*, Mottiar Y*, Mansfield SD. ToF-SIMS imaging reveals that p-hydroxybenzoate groups specifically decorate the lignin of fibres in the xylem of poplar and willow. Holzforschung 2020, 75(5): 452–462. *Equally contributing first-authors. Publisher: Walter de Gruyter GmbH. vii Table of Contents Abstract ........................................................................................................................................................ iii Résumé ......................................................................................................................................................... iv Lay Summary ................................................................................................................................................ v Preface .......................................................................................................................................................... vi Table of Contents ........................................................................................................................................ vii List of Tables ................................................................................................................................................ xi List of Figures ............................................................................................................................................. xii List of Abbreviations .................................................................................................................................. xiv Glossary ...................................................................................................................................................... xvi Acknowledgements ..................................................................................................................................... xx Dedication .................................................................................................................................................. xxi Chapter 1: Introduction 1.1 Lignin and plant cells walls ...................................................................................................... 1 1.2 Lignin biosynthesis and assembly ............................................................................................ 4 1.3 Evolution of lignin .................................................................................................................... 7 1.4 Biological importance of lignin ................................................................................................ 9 1.5 Industrial relevance of lignin .................................................................................................. 11 1.6 Research objectives................................................................................................................. 14 Chapter 2: Designer lignins: Harnessing the plasticity of lignification 2.1 Summary ................................................................................................................................. 16 2.2 Introduction ............................................................................................................................. 16 2.3 Lignin pathway engineering ................................................................................................... 19 2.4 Designer lignins ...................................................................................................................... 24 2.4.1 Shorter lignin chains ................................................................................................... 24 2.4.2 Less hydrophobic lignins ............................................................................................ 27 2.4.3 Lignins with less structural cross-linking to carbohydrates ........................................ 27 2.4.4 Lignins with novel chemically labile bonds ............................................................... 28 2.4.5 Value-added lignins .................................................................................................... 29 2.5 Perspectives on lignin engineering ......................................................................................... 30 2.6 Conclusions ............................................................................................................................. 32 Chapter 3: Suppression of chorismate mutase 1 in hybrid poplar reveals possible redundancy in the supply of lignin precursors 3.1 Summary ................................................................................................................................. 33 3.2 Introduction ............................................................................................................................. 33 3.3 Materials and Methods ............................................................................................................ 36 3.3.1 Phylogenetic analysis .................................................................................................. 36 3.3.2 RNAi-mediated suppression of chorismate mutase 1 in poplar .................................. 36 3.3.3 Gene expression .......................................................................................................... 38 3.3.4 Lignin content, composition, and histology ................................................................ 41 3.3.5 Metabolites ................................................................................................................. 42 3.4 Results and Discussion ........................................................................................................... 43 3.4.1 Identification of poplar CM isoforms ......................................................................... 43 3.4.2 Downregulation of CM1 in hybrid poplar .................................................................. 45 3.4.3 Compensatory lignification in CM1-suppressed poplar ............................................. 47 3.4.4 Systemic responses in plant metabolism and gene expression ................................... 52 3.4.5 Redundancy in the CM gene family ........................................................................... 56 viii Chapter 4: Metabolic engineering of p-hydroxybenzoate pendent groups in poplar lignin 4.1 Summary ................................................................................................................................. 60 4.2 Introduction ............................................................................................................................. 60 4.3 Materials and Methods ............................................................................................................ 63 4.3.1 Gene cloning and expression constructs ..................................................................... 63 4.3.2 Establishment of transgenic hybrid poplars ................................................................ 64 4.3.3 Expression of the ubiC transgene ............................................................................... 68 4.3.4 Lignin content and composition ................................................................................. 69 4.3.5 NMR analysis ............................................................................................................. 70 4.3.6 Metabolites and cell-wall-bound p-hydroxybenzoate ................................................. 71 4.3.7 Monolignol–p-hydroxybenzoate conjugates ............................................................... 74 4.3.8 Gel-permeation chromatography ................................................................................ 75 4.3.9 Lignin histology and ToF-SIMS imaging ................................................................... 75 4.3.10 Structural polysaccharides and saccharification potential ........................................ 76 4.3.11 Replicates and statistical analysis ............................................................................. 77 4.4 Results ..................................................................................................................................... 77 4.4.1 Transgenic poplars expressing bacterial CPL ............................................................. 77 4.4.2 Alterations in lignin .................................................................................................... 79 4.4.3 Elevated cell-wall-bound p-hydroxybenzoate ............................................................ 79 4.4.4 Accumulation of p-hydroxybenzoate-containing metabolites .................................... 82 4.4.5 Distribution of cell-wall-bound p-hydroxybenzoate ................................................... 85 4.4.6 Improved saccharification ........................................................................................... 86 4.5 Discussion ............................................................................................................................... 88 4.5.1 Ester-linked p-hydroxybenzoate pendent groups ........................................................ 88 4.5.2 Diverting carbon flux with CPL ................................................................................. 88 4.5.3 Strategies to optimise p-hydroxybenzoylation ............................................................ 91 4.5.4 Impacts of ester-linked monolignol conjugates .......................................................... 92 Chapter 5: Convergent evolution of lignin p-coumaroylation provides new tools for lignin engineering 5.1 Summary ................................................................................................................................. 94 5.2 Introduction ............................................................................................................................. 94 5.3 Materials and Methods ............................................................................................................ 97 5.3.1 Lignin analysis by NMR and DFRC ........................................................................... 97 5.3.2 Gene identification ...................................................................................................... 98 5.3.3 Recombinant protein expression ................................................................................. 98 5.3.4 Enzyme characterisation ........................................................................................... 101 5.3.5 Transgenic hybrid poplar .......................................................................................... 102 5.3.6 Transgene expression ................................................................................................ 103 5.3.7 Lignin content, composition, and molecular weight ................................................. 103 5.3.8 Cell-wall-bound phenolic acids and acetyl ............................................................... 104 5.3.9 Structural polysaccharides and saccharification potential ........................................ 104 5.3.10 Analysis of PMT sequences .................................................................................... 104 5.3.11 Replicates and statistics .......................................................................................... 105 5.4 Results and Discussion ......................................................................................................... 105 5.4.1 Kenaf lignin is naturally p-coumaroylated ............................................................... 105 5.4.2 Recombinant HcPMT produces monolignol–pCA conjugates ................................. 108 5.4.3 Transgenic poplar trees expressing HcPMT have altered lignin .............................. 111 5.4.4 Industrial relevance of pCA ...................................................................................... 116 5.4.5 HcPMT differs substantially from commelinid monocot PMT sequences ............... 119 5.4.6 p-Coumaroylation in kenaf may be the product of convergent evolution ................ 121 ix Chapter 6: Atypical lignification in eastern leatherwood (Dirca palustris) 6.1 Summary ............................................................................................................................... 124 6.2 Introduction ........................................................................................................................... 124 6.3 Materials and Methods .......................................................................................................... 126 6.3.1 Wood samples ........................................................................................................... 126 6.3.2 Analysis of lignin and structural polysaccharides .................................................... 126 6.3.3 Microscopy ............................................................................................................... 126 6.3.4 Confocal Raman imaging ......................................................................................... 127 6.3.5 Time-of-flight secondary ion mass spectrometry imaging ....................................... 128 6.4 Results ................................................................................................................................... 128 6.4.1 Fourier transform infrared spectroscopy ................................................................... 128 6.4.1 Lignin content and composition ............................................................................... 129 6.4.2 Structural polysaccharides ........................................................................................ 129 6.4.3 Cell wall histochemistry ........................................................................................... 131 6.4.4 Label-free chemical imaging .................................................................................... 133 6.4.5 Lignin-deficient middle lamellae .............................................................................. 135 6.5 Discussion ............................................................................................................................. 135 6.5.1 Unusual cell wall composition and lignin distribution ............................................. 137 6.5.2 Wood flexibility and lignin-deficient middle lamellae ............................................. 138 6.5.3 Challenging the model of lignification ..................................................................... 138 6.5.4 Implications of lignin-deficient middle lamellae ...................................................... 140 Chapter 7: Conclusions 7.1 Summary of major research contributions ............................................................................ 142 7.2 Diverting carbon flux away from lignin biosynthesis ........................................................... 144 7.3 Lignin acylations, monolignol transferases, and designer lignins ........................................ 145 7.4 Unresolved questions in lignin biology ................................................................................ 148 7.5 Outroduction ......................................................................................................................... 151 References ................................................................................................................................................. 152 Appendix 1. List of Concurrently Published Work ................................................................................... 187 Appendix 2. Supplementary Information Pertaining to Chapter 6 ............................................................ 188 A2.1 Supplementary Notes ............................................................................................................ 188 A2.1.1 The Dirca genus ..................................................................................................... 188 A2.1.2 FTIR analysis ......................................................................................................... 188 A2.1.3 Microfibril angle .................................................................................................... 189 A2.1.4 Additional Raman imaging .................................................................................... 190 A2.1.5 Lignin distribution in other Dirca species ............................................................. 190 A2.2 Supplementary Materials and Methods ................................................................................ 191 A2.2.1 Sample processing ................................................................................................. 191 A2.2.2 Fourier-transform infrared spectroscopy ............................................................... 191 A2.2.3 Lignin content ........................................................................................................ 192 A2.2.4 Structural polysaccharides composition ................................................................. 192 A2.2.5 Acetyl content ........................................................................................................ 193 A2.2.6 Lignin composition ................................................................................................ 193 A2.2.7 Cellulose content .................................................................................................... 194 A2.2.8 Crystallinity ........................................................................................................... 194 A2.2.9 Microfibril angle .................................................................................................... 195 A2.3 Supplementary Tables and Figures ....................................................................................... 196 x Appendix 3. ToF-SIMS imaging reveals that p-hydroxybenzoate groups specifically decorate the lignin of fibres in the xylem of poplar and willow .................................. 202 A3.1 Summary ............................................................................................................................... 202 A3.2 Introduction ........................................................................................................................... 202 A3.3 Materials and Methods .......................................................................................................... 205 A3.3.1 Plant samples ......................................................................................................... 205 A3.3.2 Chemical analysis of –pHB groups and lignin ....................................................... 206 A3.3.3 ToF-SIMS .............................................................................................................. 207 A3.4 Results and Discussion ......................................................................................................... 208 A3.4.1 Occurrence of –pHB groups in poplar and willow ................................................ 208 A3.4.2 ToF-SIMS detection of –pHB groups .................................................................... 210 A3.4.3 Distribution of –pHB groups in wood .................................................................... 212 A3.4.4 The importance of –pHB as lignin decorations ..................................................... 215 A3.5 Conclusions ........................................................................................................................... 217 A3.6 Supplementary Information .................................................................................................. 217 A3.6.1 Principal component analysis................................................................................. 217 xi List of Tables Table 3-1. Primers and PCR programmes for Chapter 3 .................................................................... 39 Table 3-2. Structural polysaccharides in Pa×gCM1 RNAi poplar trees ............................................. 49 Table 4-1. Primers and PCR programmes for Chapter 4 .................................................................... 65 Table 4-2. Structural polysaccharides in transgenic poplar expressing bacterial CPL ....................... 87 Table 5-1. Primers and PCR programmes for Chapter 5 .................................................................... 99 Table 5-2. Structural polysaccharides in transgenic poplars expressing HcPMT ............................. 113 Table 6-1. Cell wall analysis of leatherwood, aspen and spruce. ...................................................... 131 Table A2-1. Annotations of FTIR bands. ............................................................................................ 196 Table A3-1. Lignin analysis of poplar and willow samples analysed by ToF-SIMS .......................... 220 xii List of Figures Figure 1-1. Plant cell walls and the deposition of lignin ....................................................................... 2 Figure 1-2. Metabolic map of monolignol biosynthesis ........................................................................ 5 Figure 1-3. Diversification of plants and the evolution of lignin .......................................................... 8 Figure 1-4. Biological roles of lignin .................................................................................................. 10 Figure 1-5. Lignification produces an array of interunit linkages ....................................................... 12 Figure 2-1. Overview of plant cell wall lignification .......................................................................... 18 Figure 2-2. Lignin biosynthesis ........................................................................................................... 21 Figure 2-3. Lignin models and designer lignins .................................................................................. 26 Figure 3-1. Chorismate is a key branchpoint metabolite in plants ...................................................... 35 Figure 3-2. Chorismate mutase sequences .......................................................................................... 37 Figure 3-3. Phylogenetic analysis of the chorismate mutase gene family ........................................... 44 Figure 3-4. RNAi-mediated suppression of CM1 in hybrid poplar ..................................................... 46 Figure 3-5. Images of greenhouse-grown Pa×gCM1 RNAi poplar trees ............................................ 47 Figure 3-6. Lignin analysis of Pa×gCM1 RNAi poplar trees ............................................................. 48 Figure 3-7. Additional NMR spectra of Pa×gCM1 RNAi poplar trees .............................................. 50 Figure 3-8. Lignin histology of Pa×gCM1 RNAi poplar trees ........................................................... 51 Figure 3-9. Metabolite profiling of Pa×gCM1 RNAi poplar trees...................................................... 53 Figure 3-10. Total salicylic acid content is reduced in Pa×gCM1 RNAi poplar trees .......................... 54 Figure 3-11. Additional metabolite profiling of Pa×gCM1 RNAi poplar trees .................................... 55 Figure 3-12. RNA-seq analysis of Pa×gCM1 RNAi poplar trees ......................................................... 57 Figure 3-13. Expression profiles of poplar chorismate mutase homologues ........................................ 58 Figure 4-1. Lignin engineering with bacterial CPL ............................................................................. 61 Figure 4-2. DNA sequences for heterologous expression of bacterial CPL ........................................ 64 Figure 4-3. Expression of bacterial CPL in transgenic poplar ............................................................ 78 Figure 4-4. Lignin analysis of transgenic poplar expressing bacterial CPL ........................................ 80 Figure 4-5. Additional NMR spectra of transgenic poplar expressing bacterial CPL ......................... 81 Figure 4-6. Analysis of monolignol conjugates in transgenic poplar expressing bacterial CPL ......... 82 Figure 4-7. Analysis of soluble metabolites in transgenic poplars expressing bacterial CPL ............. 84 Figure 4-8. Lignin histology of transgenic poplar expressing bacterial CPL ...................................... 85 Figure 4-9. ToF-SIMS imaging of transgenic poplar expressing bacterial CPL ................................. 86 Figure 4-10. Saccharification potential of transgenic poplar expressing bacterial CPL. ...................... 87 Figure 5-1. Incorporation of ester-linked pendent groups in lignin .................................................... 96 Figure 5-2. Evidence that kenaf core lignin naturally contains pCA groups..................................... 106 Figure 5-3. Coomassie-stained SDS-PAGE gel of recombinat HcPMT ........................................... 108 Figure 5-4. Pairwise assays to screen HcPMT enzyme activity ........................................................ 109 Figure 5-5. Competitive enzyme activity assays testing HcPMT substrate preferences ................... 110 Figure 5-6. Transgenic hybrid poplar trees expressing HcPMT ....................................................... 111 Figure 5-7. Transgene expression in transgenic poplar expressing HcPMT ..................................... 112 Figure 5-8. Lignin analysis of transgenic poplar expressing HcPMT ............................................... 113 Figure 5-9. Analysis of lignin acylations in transgenic poplar expressing HcPMT .......................... 114 Figure 5-10. HPLC analysis of cell-wall-bound phenolics in transgenic poplar expressing HcPMT . 115 Figure 5-11. Additional NMR spectra of transgenic poplar expressing HcPMT ................................ 117 Figure 5-12. Saccharification potential of transgenic poplar expressing HcPMT ............................... 118 Figure 5-13. Analysis of PMT sequences and phylogeny ................................................................... 120 Figure 6-1. Semi-quantitative FTIR analysis of leatherwood, aspen and spruce .............................. 130 Figure 6-2. Lignin histochemical analysis of leatherwood, aspen and spruce. ................................. 132 Figure 6-3. Raman imaging of leatherwood. ..................................................................................... 134 Figure 6-4. Evidence of lignin-deficient middle lamellae in leatherwood. ....................................... 136 xiii Figure A2-1. The family Thymelaeaceae and the distribution of leatherwood .................................. 197 Figure A2-2. FTIR analysis of leatherwood, aspen and spruce ......................................................... 198 Figure A2-3. Microfibril angle measurements of leatherwood, aspen and spruce ............................. 199 Figure A2-4. Additional Raman imaging of leatherwood .................................................................. 200 Figure A2-5. Lignin-deficient middle lamellae across the genus Dirca ............................................ 201 Figure A2-6. Photos of leatherwood .................................................................................................. 201 Figure A3-1. Diagram of p-hydroxybenzoylated lignin ..................................................................... 203 Figure A3-2. Occurrence of ester-linked p-hydroxybenzoate groups in poplar tissues ..................... 208 Figure A3-3. Identification of p-hydroxybenzoate-specific ToF-SIMS fragment ............................. 211 Figure A3-4. ToF-SIMS imaging of poplar and willow .................................................................... 213 Figure A3-5. Multivariate curve resolution modelling of ToF-SIMS spectral images ...................... 214 Figure A3-6. Principal component analysis (PCA) of poplar and willow ToF-SIMS spectra ........... 218 Figure A3-7. Additional ToF-SIMS imaging of poplar and willow .................................................. 220 xiv List of Abbreviations 4CL 4-coumarate CoA ligase Ac acetyl (ester-linked form) ADH arogenate dehydrogenase ADT arogenate dehydratase ANOVA analysis of variance BA benzoate (ester-linked form) BADH a superfamily of acyltransferase enzymes, named after three enzymes: benzyl alcohol O-acetyltransferase, anthocyanin O-hydroxycinnamoyltransferase, N-hydroxycinnamoyl/benzoyltransferase, and deacetylvindoline 4-O-acetyltransferase C3ˈH p-coumaroyl shikimate/quinate 3ˈ-hydroxylase C3H p-coumarate 3-hydroxylase C4H cinnamate 4-hydroxylase CAD cinnamyl alcohol dehydrogenase CCoAOMT caffeoyl-CoA-O-methyltransferase CCR hydroxycinnamoyl-CoA reductase CesA cellulose synthase CM chorismate mutase CoA coenzyme A COMT caffeate/5-hydroxyconiferaldehyde-O-methyltransferase CPL chorismate pyruvate lyase CPR cytochrome P450 reductase CRISPR/Cas9 clustered regularly interspaced short palindromic repeats/CRISPR associated protein 9 CS chorismate synthase CSE caffeoyl shikimate esterase CTAB cetyl trimethylammonium bromide DAD diode-array detector DAHPS 3-deoxy-D-arabinoheptulosonate 7-phosphate synthase DE gene differentially expressed gene DFRC derivatisation followed by reductive cleavage DHQD-SDH 3-dehydroquinate dehydratase–shikimate dehydrogenase DHQS 3-dehydroquinate synthase EPSPS 5-enolpyruvylshikimate 3-phosphate synthase F5H ferulate/coniferaldehyde 5-hydroxylase FA ferulate (ester-linked form) FID flame ionisation detector FMT feruloyl-CoA:monolignol transferase FTIR Fourier-transform infrared spectroscopy G lignin guaiacyl lignin unit GC gas chromatography GO term gene ontology term H lignin p-hydroxyphenyl lignin unit HCT p-hydroxycinnamoyl-CoA:shikimate/quinate p-hydroxycinnamoyltransferase xv HPLC high-performance liquid chromatography HPPAT 4-hydroxyphenylpyruvate aminotransferase HRMS high-resolution mass spectrometry HSQC heteronuclear single quantum coherence KNAT a family of transcription factors, named after a leaf phenotype: knotted Arabidopsis thaliana MCR multivariate curve resolution MS mass spectrometry MYA million years ago MYB a family of transcription factors, named after avian myeloblastosis virus NAC a family of transcription factors, named after three genes: NAM (no apical meristem), ATAF1/2 (Arabidopsis thaliana transcription activation factor), and CUC2 (cup-shaped cotyledon) NADPH nicotinamide adenine dinucleotide phosphate NMR nuclear magnetic resonance PAL phenylalanine ammonia lyase PAT prephenate aminotransferase pCA p-coumarate (ester-linked form) PCA principal component analysis PCR polymerase chain reaction PDH prephenate dehydrogenase PDT prephenate dehydratase PET polyethylene terephthalate pHB p-hydroxybenzoate (ester-linked form) pHBA p-hydroxybenzoic acid (free acid form) PMT p-coumaroyl-CoA:monolignol transferase PPAT phenylprephenate aminotransferase PTFE polytetrafluoroethylene RNAi RNA interference RT-qPCR real-time quantitative polymerase chain reaction S lignin syringyl lignin unit SAH S-adenosylhomocysteine SAM S-adenosylmethionine SDS-PAGE sodium dodecyl sulphate-polyacrylamide gel electrophoresis SK shikimate kinase TAL tyrosine ammonia lyase ToF-SIMS time-of-flight secondary-ion mass spectrometry UDP uridine diphosphate UHPLC ultra high-pressure liquid chromatography UV ultraviolet VA vanillate WPM woody plant media WT wild-type XRD X-ray diffraction xvi Glossary acylation the condensation of a chemical moiety, an acyl group, onto another molecule whereby a carboxylic acid typically serves as the acyl donor and an alcohol serves as the acyl acceptor (examples of lignin acylations described in this dissertation include acetylation, p-coumaroylation, and p-hydroxybenzoylation) acyltransferase an enzyme that catalyses the condensation of an acyl donor and an acyl acceptor to form a conjugate molecule allostery a mechanism of biochemical regulation in which the binding of a molecule to an effector site causes activation or repression of enzyme activity angiosperms also known as flowering plants, an extremely large and diverse phylogenetic group of plant species that are characterised by having complex flowers and seeds with endosperms bioeconomy an economic system wherein biomass, specifically plant biomass, is valued as a source of food, energy, services, and diverse bioproducts biomass recalcitrance a complex set of chemical and physical properties of plant biomass that confers resistance to decomposition by mechanical, chemical and/or biological agents bioproducts any of a number of biochemicals, biofuels, or biomaterials that can practically or theoretically be produced using plant biomass biorefinery an emerging framework within the context of sustainability and the bioeconomy that describes an industrial facility which transforms plant biomass into a range of biofuels and bioproducts (in contrast to traditional refineries which transform and fractionate fossil-derived raw materials into fuels, chemicals, and other products) cell wall A polysaccharide-rich structural layer that surrounds the plasma membrane of most bacterial, fungal, algal, and plant cells to provide structural support, regulate permeability, and contribute to defence (in plants, nearly all cells have a primary cell wall and some cells also have a secondary cell wall) cellulose a polysaccharide consisting of parallel strands of β-(1,4)-linked D-glucose that occurs in the cell walls of plants, algae, and some bacteria chemically labile linkage a relative quality that refers to the ease with which a chemical bond can be broken, reflecting the inherent bond dissociation energy clip-offs a term that refers, in this dissertation, to chemically labile constituents that can be readily cleaved from lignin polymers, such as ester-linked pendent groups conjugate any chemical compound derived from the covalent linking of two or more molecules, particularly when these are joined via ether or ester bonds convergent evolution the process by which analogous traits arise independently through evolution dehydrogenative polymerisation the process by which lignin monomers are assembled into a polymer (this includes radical formation, radical coupling, and rearomatisation) delignification the process of removing lignin from lignocellulosic plant biomass xvii designer lignins a term, coined in this dissertation, that refers to lignin polymers which have been deliberately and rationally designed and realised by way of genetic manipulations dicots a large and diverse groups of angiosperm plant species that are characterised by the presence of two embryonic leaves (the eudicots form the largest clade within the dicots and are distinguished by the structure of their pollen grains) gene redundancy the occurrence of two or more genes whose expression can occur interchangeably with little or no phenotypic consequences since the gene products have sufficiently similar biochemical activities gymnosperms a large and diverse phylogenetic group of plant species that are characterised by having non-encased seeds hemicellulose any of a number of matrix polysaccharides other than cellulose and pectin found in the cell walls of plants (examples include xyloglucan, arabinoxylan, and glucomannan) heterologous expression the expression of a gene from one species in another species wherein it does not otherwise occur naturally homologous genes also known as homologues, two or more genes that share considerable sequence similarity due to a shared evolutionary history (compare with orthologues which are homologues that occur due to inheritance from a common ancestor, paralogues which are homologues that arise from duplication and neofunctionalisation following speciation events, and xenologues which are homologues resulting from horizontal gene transfer) lignin a complex phenolic polymer found predominantly in the cell walls of vascular plants that is assembled via the dehydrogenative polymerisation of, primarily, hydroxycinnamate derivatives known as monolignols lignin engineering the deliberate act of manipulating the content, composition, and/or structure of lignin by way of genetic manipulation lignin monomers monolignols and non-canonical lignin monomers that are compatible with the dehydrogenative polymerisation reactions that assemble lignin polymers lignin plasticity a term that refers to the inherent flexibility in the mechanisms of lignin formation including at the levels of biosynthesis and polymerisation lignification a term that refers either broadly to the biological processes of lignin biosynthesis and deposition, or more specifically to the dehydrogenative polymerisation reactions that assemble lignin polymers lignocellulose plant biomass which is composed predominantly of the polysaccharides cellulose, hemicellulose, and pectin, along with the phenolic polymer lignin metabolic flux the rate of turnover of metabolites and intermediates in a biosynthetic pathway metabolite a chemical compound that is either the end-product or an intermediate of a biosynthetic pathway middle lamella the space between the cell walls of two adjacent cells that is initially created when a cell plate is formed in a mother cell before it divides into two daughter cells, and which is typically rich in pectin but can also become lignified xviii monocot a large and diverse groups of angiosperm plant species that are characterised by the presence of just one embryonic leaf monolignol a term that refers to the classical lignin monomers that make-up the majority of lignin, namely p-hydroxyphenyl, coniferyl and sinapyl alcohol non-canonical lignin monomers a term that refers to lignin monomers other than the monolignols which are compatible with dehydrogenative polymerisation parenchyma a type of ground tissue in plants comprised of thin-walled cells (in the xylem of vascular plants, axial and ray parenchyma are produced by cell divisions of the vascular cambium and are important in storage and in the radial transport of water, minerals, nutrients, and other molecules) pectin any of a number of matrix polysaccharides other than cellulose and hemicellulose found in the cell walls of plants that contain D-galacturonic acid (examples include homogalacturonan and rhamnogalacturonan I and II) pendent groups a term that refers, in this dissertation, to ester-linked moieties in lignin that are pendent (i.e., that hang from the polymer) resulting from the participation of acylated monolignols in dehydrogenative polymerisation phenylpropanoids a large group of compounds that contain core structures comprised of a phenolic ring and a three-carbon propenyl sidechain phloem one of the major tissues that comprise the plant vasculature and which is primarily involved in the long-distance transport of photosynthate photosynthate the biochemical products of carbon fixation via photosynthesis, specifically the translocated sugars that are produced in source tissues and further metabolised in sink tissues physicochemical properties the suite of physical and chemical parameters that define molecular properties plant biomass the bodies of plants, particularly from so-called biomass feedstock species, that can be converted into a variety of bioproducts polymer a macromolecule composed of many repeating subunits (lignin, cellulose, hemicellulose, and pectin are examples of biopolymers that occur in plant cell walls) pretreatment a term that refers, in this dissertation, to the treatment of plant biomass with thermal, mechanical, and/or chemical processes to improve the accessibility of matrix polysaccharides for subsequent saccharification (pretreatment frequently involves delignification and potentially some degradation of polysaccharides) primary cell wall the relatively thin cell wall layer that surrounds nearly all plant cells and which is composed primarily of cellulose, hemicellulose, and pectin recombinant enzyme an enzyme produced in a protein production host by introducing an expression vector that harbours an exotic gene of interest saccharification a process by which constituent monomeric sugars are released from polysaccharides (in the case of lignocellulosic biomass, this involves the depolymerisation of cellulose, hemicellulose, and pectin into monosaccharides) xix saponification also known as alkaline hydrolysis, a chemical reaction in which an ester bond is cleaved resulting in an alcohol and an acid secondary cell wall the thickened cell wall layers that surround a subset of plant cells and which is composed primarily of cellulose, hemicellulose, and lignin secondary metabolites a suite of molecules that are not directly involved in normal growth, development, homeostasis, or reproduction, but which can nonetheless be highly consequential in diverse biological processes (contrast with primary metabolites which are involved in growth, development, homeostasis, or reproduction) spatiotemporal occurrence in space and time sustainability the degree to which a process or system can be maintained in a constant state without extraneous inputs or outputs (in the context of biomass, biorefineries, and the bioeconomy, this is largely aspirational and refers to technologies and practices which generate bioproducts with minimal non-renewable inputs and negligible waste) sylleptic branches branches that emerge from lateral buds without the apical meristem having experienced a rest period (contrast with proleptic branches which emerge following a cessation of growth, typically winter dormancy) tracheary elements specialised cells occurring in vascular plants that enable the long-distance transport of water and which feature lignified secondary cell walls at maturity (includes tracheids and xylem vessels) tracheid a type of tracheary element found in the xylem tissues of vascular plants that enable the long-distance transport of water, distinguished from xylem vessels by their smaller size, their shape, and the absence of perforation plates transgenic plant a plant in which a transgene, a foreign piece of DNA, has been transferred from another species (this could be another plant species, a bacterial species, etc.) value-added lignin a term, coined in this dissertation, that refers to designer lignin polymers which contain valuable chemical components beyond the canonical lignin monomers vascular plants also known as tracheophytes, a phylogenetic group of plant species that have vascular tissues which feature lignified tracheary elements that enable the long-distance transport of water wood the secondary xylem tissue that accrues from cell divisions of the vascular cambium in vascular plants xylem one of the major tissues that comprise the plant vasculature and which is involved in the long-distance transport of water, with primary xylem (including protoxylem and metaxylem) being formed from cell divisions of the procambium and secondary xylem being formed from the vascular cambium xylem fibre sclerenchyma cells that are dead at maturity and typically lignified to provide mechanical support to plant bodies xylem vessel a type of tracheary element found in the xylem tissues of angiosperms that enable the long-distance transport of water, distinguished from tracheids by their larger size, their shape, and the presence of perforation plates xx Acknowledgements I am grateful for the support and guidance of my supervisor Dr. Shawn Mansfield and my thesis advisory committee members Drs. Rob Guy, Lacey Samuels, Simone Castellarin, and the late Carl Douglas. Drs. John Ralph, Rebecca Smith, and Steven Karlen graciously hosted me in Madison and taught me about NMR, acyltransferases, and DFRC. This work would not have been possible without various other collaborators as well: Drs. Notburga Gierlinger, Emma Master, Dragica Jeremic, Timothy Tschaplinski, Robyn Goacher, Curtis Wilkerson, Wout Boerjan, and Ruben Vanholme. In addition, many scientific mentors inspired me and helped guide me down this path including Dr. Illimar Altosaar, Dr. Mohsin Zaidi, John Lant, Paul St. Louis, Brian Long, John Wills, Dr. Louis Jugnia, and Grant Dobson. I am also thankful for financial support from NSERC, UBC, and the Faculty of Forestry. I will always hold onto many happy memories and life-long friendships from my time at UBC. More than anyone else, Kaye Hare coached me through the most difficult days and celebrated with me on the happiest ones, and I will be forever grateful for this. I have met so many incredible and inspiring people during my time at UBC, and I would especially like to thank the following people for their intelligence, humour, moral support, and friendship: Amanda Johnson, Dr. Roli Wilhelm, Emily Murphy, Dr. Florien Heinkel, Natalie Hoffmann, Dr. Devra Waldman, Jonathan Gula, Marie-Pier Baril, Devin Harrington, Laura Peña, Ahmed Kenawy, Dr. Danyu Yao, Dr. Mabel Ting Wong, Arend Hoekstra, Dr. Alex Tsai, Dr. Richard Chandra, Dr. Lan Tran, Rohan Chandrasekhar, Matthew Doering, Jennifer Doering, Brenna Jansen, Dr. Annise Dobson, Dr. Mathias Scheutz, Dr. Krešimir Šola, Dr. Marie-Eve Bouchard, Dr. Fredrik Nielsen, Dr. Vera Novy, and so many more than I regrettably cannot mention in this space. During my time at St. John’s College and through my involvement with the FGSA and the GSS, I had the privilege of working with and learning from a great number of other truly remarkable people as well. Most of all, none of this would be possible without my family. I am endlessly grateful for their love and support. They taught me that the virtues of patience, empathy, and compassion are foundational for a happy life, and that hard work and perseverance always pay off in the end. xxi Dedication For my Mum and Dad. Down river lies the world… Marion Greene, 1946 1 Chapter 1: Introduction 1.1 Lignin and plant cells walls Cell walls are one of the defining features of plants (Hooke, 1665; von Mohl, 1851). They provide the mechanical strength and rigidity needed to resist turgor pressure that develops from osmotic flow into the large central vacuole (Pfeffer, 1877). They also enable plant morphogenesis, regulate permeability, and act as a barrier for defence (Stiles, 1923; Sampathkumar et al., 2014; Malinovsky et al., 2014). Plant cell walls originate from cytokinesis when the cell plate is laid down at the phragmoplast of a mother cell (Strasburger, 1880). As the callose-rich cell plate grows radially across the cell and becomes enriched in pectin and cellulose, it partitions the protoplasm and defines the two nascent daughter cells (Samuels et al., 1995; Drakakaki, 2015). The emergence of cell walls is underpinned by the cytoskeleton and relies upon the timely biosynthesis and delivery of cell wall polysaccharides. De novo cell wall formation also gives rise to the compound middle lamella, the space between the walls of adjacent cells (Allen, 1901). New cells originate in meristems (Nägeli, 1858). Following the germination of a seed and the polarisation of developing cells in the embryo, two apical meristems arise which drive growth in the root and in the shoot (Priestley, 1928). The ensuing period of primary growth produces a myriad of cell types, and continued elongation of roots and shoots is achieved by further divisions within the apical meristems. Through cell differentiation, this produces organs such as leaves, flowers, and lateral roots (Torrey, 1967). Secondary meristems, which also arise through this process, in turn drive the radial growth of stems and roots. For example, daughter cells of vascular cambial meristems undergo periclinal divisions to produce the secondary xylem and phloem cells that comprise the plant vascular system (Esau, 1943). Although many cells retain only primary walls at maturity, others undergo secondary cell wall formation, mostly after the cell has ceased expanding (Figure 1-1A; Albersheim et al., 2011; Zhong et al., 2018). The primary cell wall is relatively thin and is composed of cellulose, pectin, and hemicellulose, while the secondary wall is much thicker, consists of up to three layers formed by apposition and can become embedded with lignin (Cosgrove & Jarvis, 2012). Cellulose is produced by membrane-bound cellulose synthase enzyme complexes, and different complements of enzymes are involved in primary and 2 secondary cell wall formation (Watanabe et al., 2018). Cellulose microfibrils are largely disordered in the primary wall but occur in parallel strands in the secondary cell wall, and the relative angle of these micro- Figure 1-1. Plant cell walls and the deposition of lignin. A: In cells that undergo secondary cell wall formation, there are typically three secondary cell wall layers (S1, S2, and S3) laid down within the primary cell wall (P). The middle lamella (ML) is the region between the walls of adjacent cells. B: Lignin is assembled primarily from the monolignols p-coumaryl, coniferyl, and sinapyl alcohol which give rise to p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) lignin units in polymeric lignin. C: The formation of cell wall layers and the biosynthesis of the corresponding constituents occurs sequentially as depicted, adapted from Terashima (1990). Lignin is first deposited in the middle lamella and cell corners (ML + CC) concurrent to polysaccharide biosynthesis and continues within the other cell wall layers as the cell wall matures. The composition of monomers available for lignification also varies temporally. 3 fibrils along with the extent of cellulose crystallinity are important determinants of cell wall strength and flexibility (Frey-Wyssling, 1976; Barnett & Bonham, 2004). During cell wall formation, hemicelluloses and pectins are pre-assembled in the Golgi and then exported via the endomembrane system (Hoffmann et al., 2021). These polysaccharides can have a significant influence on cell wall properties by interacting with other cell wall components, primarily through non-covalent interactions (Braybrook et al., 2012; Berglund et al., 2020). Pectins also predominate in the middle lamella and help adjacent cells adhere (Zamil & Geitmann, 2017). In the primary wall, the hemicelluloses xyloglucan and arabinoxylan can occur along with the pectins homogalacturonan and rhamnogalacturonan I and II, as well as mixed-linkage glucans in the Poales, an order of grasses within the commelinid monocots (Keegstra et al., 1973; Carpita & Gibeaut, 1993; Smith & Harris, 1999). Secondary cell wall formation sees the production of glucuronoxylan and glucomannan in angiosperms, and galactoglucomannan and glucuronoarabinoxylan in gymnosperms (Scheller & Ulvskov, 2010; Kumar et al., 2016). It is within this polysaccharide-rich matrix that lignin is deposited. Extensive studies on lignin over the past 200 years have shown that it is a complex polymer assembled in muro by the polymerisation of various phenolic monomers, primarily the monolignols p-coumaryl, coniferyl, and sinapyl alcohol (Figure 1-1B; de Candolle, 1813; Adler, 1977; Boerjan et al., 2003). When incorporated into polymeric lignin, the monolignols give rise to p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) lignin units. Lignification ensues after the deposition of cell wall polysaccharides is largely complete (Figure 1-1C; Terashima, 1990). Lignin is first deposited in the cell corners and middle lamella, and while the majority is found in secondary cell walls, the highest concentrations exist in the middle lamella (Fergus et al., 1969; Donaldson, 2001). The deposition of lignin typically occurs as part of programmed cell death and leads to drastic changes in cell wall physicochemical properties (Bollhöner et al., 2012). Interestingly, xylem cells can also contribute lignin monomers to support the lignification of neighbouring cells in a non-cell autonomous manner (Hosokawa et al., 2001; Smith et al., 2013). Lignin has been particularly well studied in the cell walls of the plant vasculature, but it can also be found in the endodermis where it directs water into the symplastic continuum, in some pericarps for seed protection, and in various other cell types as a response to abiotic stress or attack by pests and pathogens 4 (Vance et al., 1980; Geldner, 2013; Landucci et al., 2020). Fibres and tracheary elements in secondary xylem have received the most attention since they form the bulk of wood, but the cell walls of xylem parenchyma and phloem fibres can also become lignified (Mellerowicz et al., 2001). 1.2 Lignin biosynthesis and assembly Due to its chemical recalcitrance, harsh deconstructive approaches are needed to study lignin, and this has often hampered progress and stoked confusion (Adler, 1977). Nonetheless, it has been established that lignin is a polymer resulting from the radical coupling of hydroxycinnamate derivatives (Klason, 1923; Erdtman, 1933; Freudenberg, 1959; Ralph et al., 2004a). Although there have been important recent revisions, the biosynthesis of lignin was largely mapped in the 1950s and 1960s by measuring the incorporation of 14C-labelled compounds (Freudenberg & Neish, 1968). For example, shikimate and phenylalanine were first implicated as lignin precursors using these techniques (Brown & Neish, 1955). Most of the ensuing enzymatic steps were initially demonstrated in vitro (Grushnikov & Shorygina, 1972; Lewis et al., 1999; Higuchi, 2006), and it was only in the latter part of the last century that mutant and transgenic plant lines were used to identify and validate the many biosynthetic genes (Campbell & Sederoff, 1996; Bonawitz & Chapple, 2010). The steps leading to monolignols exist as a metabolic grid rather than a linear pathway, and some routes are favoured over others in different plant taxa (Figure 1-2; Wang et al., 2019). The first enzyme in lignin biosynthesis, phenylalanine ammonia lyase (PAL), mediates the transition from primary metabolism by the deamination of phenylalanine, a product of the shikimate pathway (Koukal & Conn, 1961). In monocots, a tyrosine ammonia lyase also exists (TAL, Neish, 1961). Successive hydroxylations of the aromatic ring are then catalysed by the cytochrome P450-dependent monooxygenases: cinnamate 4-hydroxylase (C4H; Russell & Conn, 1967), p-coumaroyl shikimate/quinate 3-hydroxylase (C3ˈH; Schoch et al., 2001), and ferulate/coniferaldehyde 5-hydroxylase (F5H; Grand, 1984). Recently, a p-coumarate 3-hydroxylase (C3H) has been reported as well (Barros et al., 2019). Cytochrome P450 reductases (CPR) are also needed to reconstitute the active sites of these monooxygenases by transferring two electrons from NADPH (Benveniste et al., 1986). 5 Figure 1-2. Metabolic map of monolignol biosynthesis. The biosynthesis of lignin proceeds through a metabolic network that takes phenylalanine through a series of modifications to the phenolic ring (depicted vertically) and to the propenyl sidechain (depicted horizontally) to produce the three primary monolignols, p-coumaryl, coniferyl and sinapyl alcohol. Enzyme abbreviations, shown in orange, are defined in the text. The route highlighted in blue is believed to be the preferred pathway in poplar. 6 Side-chain activation is achieved by 4-coumarate CoA ligase (4CL; Walton & Butt, 1970), and p-hydroxycinnamoyl-CoA:shikimate/quinate p-hydroxycinnamoyltransferase (HCT) adds and then subsequently removes the shikimate or quinate ester (Stöckigt & Zenk, 1974). Caffeoyl shikimate esterase (CSE) can also remove this ester group, but yields caffeate instead of caffeoyl-CoA (Vanholme et al., 2013). Hydroxycinnamoyl-CoA reductase (CCR) removes the CoA thioesters (Gross et al., 1973). Caffeate/5-hydroxyconiferaldehyde-O-methyltransferase (COMT; Finkle & Nelson, 1963) and caffeoyl-CoA-O-methyltransferase (CCoAOMT; Pakusch et al., 1989) substitute ring hydroxyls for methoxy groups from S-adenosyl methionine (Byerrum et al., 1954). And finally, cinnamyl alcohol dehydrogenase (CAD) reduces the aldehyde groups to yield the monolignols (Gross et al., 1973). As a phenylpropanoid, lignin shares biosynthetic steps with various other secondary metabolites (Vogt, 2010). The phenylpropanoids derive from phenylalanine and tyrosine which are shuttled from aromatic amino acid biosynthesis and, further upstream, from the shikimate pathway which drives the production of chorismate via shikimate from phosphoenolpyruvate and erythrose 4-phosphate, both products of carbohydrate metabolism originating in photosynthesis (Maeda & Dudareva, 2012). As the shikimate and aromatic amino acid pathways are central to the production of a host of primary and secondary metabolites alike, these biosynthetic routes are subjected to extensive control at multiple levels including transcriptional regulation, post-translational modification, allosteric regulation, and the supply of substrates (Haslam, 1974; Gilchrist & Kosuge, 1980; Tzin et al., 2010; Tohge et al., 2013a). Following biosynthesis in the cytosol, monolignols are exported to the apoplast. Because free phenolics are inherently cytotoxic and insoluble, and since the biosynthetic enzymes could be feedback inhibited, monolignols may be glucosylated prior to export (Freudenberg, 1959; Liu et al., 2011; Terashima et al., 2016). In support of this hypothesis, the phenolic glucosides of coniferyl and sinapyl alcohol are abundant in some lignifying tissues, whereas free monolignols are not (Tiemann & Haarmann, 1874; Savidge, 1989; Tsuyama & Takabe, 2014). However, these could be mere storage forms and irrefutable evidence to the contrary has not been forthcoming. The mechanism driving lignin precursor export has been equally elusive. The leading hypotheses assert that monolignols transit the plasma membrane via passive diffusion down a concentration gradient (Vermaas et al., 2019; Perkins et al., 7 2019), through transporter proteins (Miao & Liu, 2010; Alejandro et al., 2012; Tsuyama et al., 2019; Väisänen et al., 2020), or by vesicle-mediated exocytosis (Pickett-Heaps, 1968), although this last mechanism has been largely debunked (Kaneda et al., 2008). Following export, secreted laccases and peroxidases situated in the apoplast catalyse the formation of monolignol radicals (Manskaya, 1948; Dean & Eriksson, 1994; Ros-Barceló, 1997; Marjamaa et al., 2009). The spatiotemporal distribution of these enzymes is a major determinant in lignin deposition (Schuetz et al., 2014; Hoffmann et al., 2020; Wang et al., 2020a; Blaschek et al., 2020; Hiraide et al., 2021). Resonance-stabilised radicals that result from abstraction of the hydrogen on the para hydroxyl group can be transferred to compatible molecules, or a pair of radicals can couple to form a new covalent bond (Harkin, 1967). In this way, a radical on a growing lignin chain can combine with that of an incoming monomer to drive end-wise polymer growth. A diffusible redox shuttle may also be involved in delivering radicals (Önnerud et al., 2002). As radicals are delocalised and can couple at various positions, different linkages are possible; however, β-aryl ether bonds are the most prevalent, particularly in S-lignin rich angiosperms. It has been established that dehydrogenative polymerisation of phenoxy radicals is controlled solely by chemical and thermodynamic factors, and that it largely proceeds in a stochastic manner such that only the supply of monomers and the availability of oxidative enzymes can actively control the composition and structure of the resulting polymer (Ralph et al., 2004a). An alternative hypothesis which proposed that lignin assembly may be template-driven and guided by dirigent proteins has been mostly discredited (Davin & Lewis, 2005; Ralph et al., 2008). However, it has been reported that a dirigent protein is required for proper lignin formation in the root endodermis of Arabidopsis (Hosmani et al., 2013). Although our understanding of lignin has progressed remarkably over the past few decades, under-explored areas of lignin biology persist and there may yet be surprises ahead. 1.3 Evolution of lignin The development of lignin was sine qua non for the diversification of land plants and the rise of tracheophytes. While many basal plant taxa produce diverse secondary metabolites including 8 phenylpropanoids, conventional wisdom holds that only vascular plants produce lignin and that the occurrence of S lignin is further confined to angiosperms (Linsbauer, 1899; Towers & Gibbs, 1953). However, explorations in lignin evolution have called into question this traditional paradigm. For instance, S lignin has been detected in various liverworts, lycophytes, in some but not all ferns, and in various cycads and gnetophytes (Baucher et al., 1998; Weng & Chapple, 2010; Novo Uzal et al., 2012). These sporadic occurrences point to a complex evolutionary history (Figure 1-3). Even more surprising, small amounts of lignin have been detected in the peripheral cells of the red alga Calliarthron cheilosporioides (Martone et al., 2009). There is also some evidence that the genes required for p-coumaryl alcohol biosynthesis may have evolved well before vascular plants emerged (Labeeuw et al., 2015). Figure 1-3. Diversification of plants and the evolution of lignin. The development of polymeric lignin (shown in orange) from simple phenolic compounds (shown in blue) occurred as plants evolved beyond their aquatic origins to predominate on land. The composition of H, G, and S-lignin units among the vascular plants is noted on each branch of the tree. The nodes are also labelled in millions of years ago (MYA). Orange asterisks indicate that S-lignin may have evolved repeatedly as there are reports in red algae and in some lycophytes, ferns, and basal gymnosperms. 9 A long-standing hypothesis holds that evolution favoured the development of phenylpropanoids to withstand biotic and abiotic stresses, particularly as photoprotectants (Raven, 1984). More specifically, hydroxylation of the para position of aromatic rings bestows absorbance in the UV-B range (Weng & Chapple, 2010). However, some of these compounds have been mistaken for polymeric lignin in basal lineages. For example, although lignin does not exist in the cell walls of bryophytes, an hydroxycinnamate called sphagnum acid does occur in abundance (Rudolph & Samland, 1985). Interestingly, peroxidase enzymes capable of facilitating polymerisation appeared before lignin itself (Espiñeira et al., 2011). As lignin is biosynthetically related to suberin and sporopollenin, two other plant polymers that also contain phenolic constituents, there may be a common evolutionary origin (Niklas et al., 2017). For example, it has been shown that a cytochrome P450 monooxygenase implicated in moss cuticle formation is ancestral to enzymes involved in lignin biosynthesis (Renault et al., 2017). It is believed that some lignin biosynthetic enzymes evolved from core metabolic genes following genome duplication events and neo-functionalisation (Tohge et al., 2013b; Niklas et al., 2017). For example, 4CL enzymes bear some homology to the long‐chain fatty acyl‐CoA synthetases involved in the β‐oxidative catabolism of fatty acids (Ehlting et al., 2001). Similarly, HCT enzymes are members of the BAHD superfamily of acyltransferases which have diversified extensively in plants to fulfill a wide range of metabolic functions (D’Auria, 2006). On the other hand, PAL may have been acquired from bacteria via horizontal gene transfer to early land plants (Emiliani et al., 2009). Once the ability to lignify cell walls was fully developed, this trait was apparently highly selective and there are no extant vascular plants that are entirely devoid of lignin. 1.4 Biological importance of lignin Lignin plays diverse roles in plants (Figure 1-4). By rigidifying the cell walls of structural fibres, lignin contributes to compressive strength and reinforces plant bodies (Niklas, 1992). When polymeric lignin first appeared in early land plants, it was present only in peripheral tissues and not in the water-conducting cells, suggesting that it was only later that lignin was recruited to support water conduction (Boyce et al., 2003). Although evolution has harnessed lignin for various functions, it remains critical for mechanical 10 Figure 1-4. Biological roles of lignin. By fortifying the cell walls of vascular plants, lignin contributes to water conduction, mechanical support, and plant defence. strength in plants. For example, lignin has long been studied in the context of stem lodging in cereal crops (Pinthus, 1974). Micromechanical studies of transgenic plants have revealed that lignification of cell walls increases stiffness and decreases viscoelastic damping (Koehler & Telewski, 2006). Similarly, Arabidopsis with lignin-deficient interfascicular fibres have a pendent phenotype whereby the inflorescence stems cannot remain erect without support (Zhong et al., 1997; Smith et al., 2013). In the cell walls of tracheary elements, lignin provides waterproofing and facilitates long-distance water transport, and this too is best exemplified by lignin-deficient plants. For example, Arabidopsis mutants severely impaired in lignin biosynthesis often have an irregular xylem phenotype with misshapen tracheary elements (Jones et al., 2001; Berthet et al., 2011). Similarly, transgenic poplar deficient in lignin may exhibit collapsed xylem vessels, decreased hydraulic conductivity, and greater susceptibility to cavitation (Coleman et al., 2008a; Voelker et al., 2011; Awad et al., 2012). Reduced lignin is frequently associated with stunted plants and this has been attributed to weak xylem conductance; however, stunting could also be a pleiotropic effect of lignin perturbation caused by hyper- or hypo-accumulation of pathway intermediates and related metabolites (Bonawitz & Chapple, 2013; Ha et al., 2021). Lignified cell walls represent a formidable innate defence against pests and pathogens (Miedes et al., 2014). For example, transgenic plants deficient in lignin are more susceptible to biotic and abiotic stress (Zhao & Dixon, 2014; Cass et al., 2015). It is not surprising then that the formation of lignin and related 11 phenylpropanoids is upregulated in response to biological attack (Foster et al., 2015; Cesarino, 2019). Lignin biosynthesis is also activated by a variety of abiotic factors including light, temperature, drought, nitrogen stress, wounding, and exposure to heavy metals, salts, ozone, and radiation (Moura et al., 2010; Cabane et al., 2012). Evidence points to an elaborate and multifaceted sensing mechanism for cell wall integrity that can monitor changes in physicochemical properties and mobilise the requisite biosynthetic machinery to fortify cell walls (Rui & Dinneny, 2020). Lignin also plays an important role in the broader biosphere. The chemical recalcitrance of lignin leads to the formation of vast amounts of humic substances in soil as dead plant material breaks down (Thevenot et al., 2010; Lehmann & Kleber, 2015). In Nature, lignin degradation is the sole purview of certain clades of filamentous fungi and soil bacteria (Falck & Haag, 1927; Ayuso-Fernández, 2019; Wilhelm et al., 2019). Normal growth of plants, and in turn all the organisms and ecosystems that depend on plants, requires adequate soil structures with sufficient organic carbon. Moreover, as lignin decomposes slowly, it represents an immensely important global carbon sink and a regulator of carbon recycling dynamics (Zeikus, 1981; Whalen et al., 2014). Finally, the lignin content of forage crops can be detrimental for animal nutrition and feed conversion rates in agriculture (Besle et al., 1994). 1.5 Industrial relevance of lignin The industrial utility of plant biomass is inexorably linked to the recalcitrance of lignin (Calvaruso et al., 2017). This chemical durability results primarily from the stability of delocalised π bonds in the aromatic rings, and from the considerable enthalpy required to sever the covalent bonds that link lignin units together, particularly the strong carbon–carbon bonds (Vermerris & Nicholson, 2006; Parthasarathi et al., 2011). In addition to β-aryl ethers (β-O-4ˈ), lignin units are also frequently linked via phenylcoumaran (β-5ˈ[α-O-4ˈ]), resinol (β-βˈ[α-O-γˈ][γ-O-αˈ]), tetrahydrofuran (β-βˈ[α-O-αˈ]), and biphenyl ether (5-O-4ˈ) structures, and sometimes via dibenzodioxocin (β-O-4ˈ[α-O-4ˈˈ][5ˈ-5ˈˈ]) and spirodienone (β-1ˈ[α-O-αˈ]) (Ralph et al., 2004a; Figure 1-5). Long-standing assertions that lignin polymers have a large molecular weight and are highly cross-linked have recently been challenged (Crestini et al., 2011; Balakshin et al., 2020). Similarly, compelling evidence that native lignin is covalently bound to hemicellulose to form 12 Figure 1-5. Lignification produces an array of interunit linkages. A: Formation of β-aryl ether bonds occurs via the coupling of a radical at the β position of an incoming monolignol with a radical at the para position of a growing lignin chain. Radicals are produced by the action of peroxidase or laccase enzymes which consume hydrogen peroxide and molecular oxygen, respectively. Following radical coupling, the quinone methide intermediate is quenched, typically by the addition of water. B: Since radicals are delocalised across the aromatic ring and the conjugated propenyl sidechain, various other linkages are possible, as shown. lignin–carbohydrate complexes has been scant until only recently (Nishimura et al., 2018; Giummarella et al., 2019). However, despite its impressive complexity and stability, lignin is not chemically inert, and processes have been developed to modify, disassemble, and liberate lignin polymers. For as long as paper has been made from wood (Fenerty, 1844), there has been a push to improve the efficiency of industrial biomass processing. In pulping, the aim is to separate and disintegrate wood fibres, and this was traditionally accomplished solely by mechanical means (Sjöström, 1993). However, as cellulose in the cell wall is enveloped by lignin and adjacent cells adhere strongly due to lignin-rich 13 middle lamellae, various chemical treatments have been devised to reduce the mechanical requirements of pulping by removing or chemically altering lignin (Smook, 2002). Beyond pulp and paper, similar delignification approaches have been explored for the pretreatment of biomass with the aim of releasing polysaccharides for enzymatic digestion and fermentation into biofuels or upgrading into a variety of bioproducts (Chandra et al., 2007). Historically, lignin that was removed from biomass was valued primarily for its energy density and was often burned to provide heat and help offset the costs of biomass processing (Glasser, 2019). More recently, a market for lignin has emerged as it is increasingly being promoted for use in the production of everything from fine chemicals to bioplastics (Bajwa et al., 2019; de Vries et al., 2021). In the ‘lignin-first’ model of a biorefinery, the aim is to prevent lignin decomposition and condensation reactions during the fractionation of biomass such that lignin structures can be preserved and perhaps depolymerised later in a controlled manner (Abu-Omar et al., 2021). While this is firstly a challenge for chemists and chemical engineers alike, progress in lignin engineering of feedstock species could contribute as well. Fortuitously, efforts to elucidate lignin biosynthesis have inadvertently shown that lignin is more heterogenous than previously realised and that dehydrogenative polymerisation is a highly flexible process (Vanholme et al., 2012a). Beyond the three primary monolignols, there is a growing list of non-canonical monomers that are also compatible with lignification (Ralph et al., 2019a). For example, acylated monolignol conjugates generated by enzymes in the BAHD acyltransferase family can participate in lignification. This observation further implies that the mechanisms of monomer export and phenoxy radical generation are equally undiscriminating. In short, it appears that any chemically compatible molecules that can be synthesised and delivered to the site of lignin formation will become incorporated into lignin (Ralph, 2006). In this context, it is now apparent that feedstock engineering strategies can strive beyond merely reducing lignin content and perturbing the proportions of H, G, and S-lignin units. With this more expansive view on lignin biosynthesis and engineering, it has finally become possible to rationally design lignin polymers (Mansfield et al., 2012). 14 1.6 Research objectives This dissertation covers a wide range of topics in cell wall biology and lignin engineering. The five major research objectives are listed here and elaborated upon further below: 1. Review the published literature on lignin biosynthesis and biochemistry to identify new strategies and opportunities for lignin engineering (Chapter 2). 2. Examine the impacts on the biosynthesis of lignin ensuing from downregulation of chorismate mutase 1 in transgenic poplar (Chapter 3). 3. Evaluate the effects on cell wall chemistry of heterologous expression of a bacterial chorismate pyruvate lyase in transgenic poplar (Chapter 4). 4. Assess the potential of a new BAHD acyltransferase gene from kenaf for engineering the lignin of transgenic poplar (Chapter 5). 5. Characterise the physicochemical properties and the distribution of lignin in the xylem of leatherwood, a North American shrub with flexible woody stems (Chapter 6). Chapter 2 provides an overview of lignin biochemistry as it pertains to the goals of lignin engineering. Plant biotechnology could be used to rationally design lignin polymers in order to overcome industrial challenges and promote lignin valorisation. This chapter provides a summary of the progress and an outlook towards future work on “designer lignins”. The ensuing three research chapters report on three different strategies that were explored for lignin engineering in hybrid poplar (Populus alba × grandidentata; P39). As a fast-growing woody species, poplar is an excellent model for the study of lignin and phenylpropanoids (Douglas et al., 2011). A high-quality genome is available and genetic transformation techniques have been optimised. Moreover, strategies developed in poplar can be readily transferred to industrially relevant biomass genotypes. Chapter 3 describes efforts to perturb the metabolic flux directed into lignin biosynthesis using RNAi techniques to suppress the expression of chorismate mutase, an important biosynthetic enzyme that lies upstream of phenylpropanoid biosynthesis. Chapter 4 reports on the introduction of a bacterial 15 chorismate pyruvate lyase gene into poplar to divert carbon away from lignin and, at the same time, increase the production of ester-linked p-hydroxybenzoate pendent groups. This work highlights the importance of substrate supply in lignin biosynthesis and demonstrates the potential of engineering cell-wall-bound phenolics. Chapter 5 reports on the characterisation of a new BAHD monolignol transferase from kenaf (Hibiscus cannabinus). This enzyme is the first p-coumaroyl-CoA:monolignol transferase identified from dicots, and heterologous expression resulted in p-coumaroylated lignin in poplar. Finally, Chapter 6 describes a study on the lignin of eastern leatherwood (Dirca palustris), a slow-growing shrub with flexible woody branches. This work revealed that a highly unusual pattern of lignin deposition has evolved in this genus. This observation asks us to reconsider the classical model of cell wall organisation and revisit several unresolved questions on the roles of lignin in plants. 16 Chapter 2: Designer lignins: Harnessing the plasticity of lignification 2.1 Summary Lignin is a complex polyphenolic constituent of plant secondary cell walls. Inspired largely by the recalcitrance of lignin to biomass processing, plant engineering efforts have routinely sought to alter lignin quantity, composition, and structure by exploiting the inherent plasticity of lignin biosynthesis. More recently, researchers are attempting to strategically design plants for increased degradability by incorporating monomers that lead to a lower degree of polymerisation, reduced hydrophobicity, fewer bonds to other cell wall constituents, or novel chemically labile linkages in the polymer backbone. In addition, the incorporation of value-added structures could help valorise lignin. Designer lignins may satisfy the biological requirement for lignification in plants while improving the overall efficiency of biomass utilisation. 2.2 Introduction Lignin is a complex polyphenolic constituent of the secondary cell walls of vascular plants, accounting for 18–35% of the biomass by weight (Sarkanen & Ludwig, 1971). It is a crucial element of water conduction and plant defence systems in tracheophytes, and it contributes significantly to the compressive strength of secondary xylem tissues. Consequently, lignification of the plant vascular system represents an important evolutionary milestone for land plants. Lignin is one of the most abundant biopolymers on the planet and is an immensely important global carbon sink. However, the chemical recalcitrance of lignin poses a major challenge for industrial biomass processing, most notably in pulp and paper production and in the emerging cellulosic biofuels industry (Baucher et al., 2003; Zeng et al., 2014). In addition, the lignin content of forage crops is an important consideration in animal nutrition and feed conversion rates in agriculture (Besle et al., 1994). Polymeric lignin is constructed primarily from three 4-hydroxyphenylpropanoids known as monolignols — p-coumaryl alcohol, coniferyl alcohol and sinapyl alcohol — that differ only in the degree of aromatic ring methoxylation (Figure 2-1; Boerjan et al., 2003). Once incorporated into a lignin 17 18 Figure 2-1. Overview of plant cell wall lignification. A: Lignin is produced primarily in the secondary-thickened cell wall layers of xylem tissues; shown here are several xylem fibres and part of a vessel element, shown in purple. The blue-highlighted cell depicts the primary (P) and three secondary cell wall layers (S1, S2 and S3). Biosynthesis of the three canonical monolignols occurs in the cytoplasm, depicted in the green highlighted cell. Finally, three proposed models of monolignol export to the cell wall are shown in the brown-highlighted cell and polymerisation of monolignols occurs within the cell wall, highlighted in yellow. B: Laccase and peroxidase enzymes present in the cell wall generate monolignol radicals that are stabilised by electron delocalisation (shown for coniferyl alcohol) prior to radical coupling reactions. C: The major polymerisation reaction is the end-wise coupling of a monolignol radical, invariably at its β-position, with the radical of the phenolic end-unit of the growing polymer (shown for the β–O–4-coupling of coniferyl alcohol with a guaiacyl radical). D: An example of a typical polymer model, derived from 20 monolignols, for poplar lignin. For an explanation of colour coding, see the caption for Figure 2-3. 19 polymer, they produce p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) moieties. These canonical monolignols are synthesised in the cytoplasm prior to export to the site of polymerisation in the cell wall where laccase and peroxidase enzymes generate monolignol radicals by dehydrogenation, either through direct action on the monolignols or via a redox shuttle (Ros-Barceló et al., 2006). The incorporation of monolignols into a growing lignin polymer via the combinatorial coupling of radicals results in highly variable racemic polymers with different physicochemical features (Ralph et al., 2004a). For example, gymnosperm lignin is more resistant to degradation largely because it is composed primarily of G subunits that yield abundant β–5, 5–5, and β–β carbon–carbon bonds in addition to the less resilient ether-type β–O–4 linkages that feature more prominently in the S-rich lignins of angiosperms. The amount, composition, and structure of lignins are highly diverse across plant taxa, cell types, developmental stages, and across the cell wall layers. Lignin is embedded in the cell walls of the plant vasculature, notably in the xylem fibres and vessel elements that constitute the bulk of secondary xylem tissues, but also in the sclerenchyma fibres and sclereids in xylem and phloem, and in the cortex cells of the periderm. Lignin recalcitrance has received considerable research attention and recent advances in our understanding of lignin biogenesis have provoked novel approaches in plant biotechnology (Vanholme et al., 2008; Eudes et al., 2014; Vermerris et al., 2015). In this review, we summarise the progress in lignin engineering, highlight developments within the past three years in the area of designer lignins, and chart a course forward to producing less recalcitrant or more valuable lignins thereby highlighting the potential to enhance the overall utility of this abundant natural polymer. 2.3 Lignin pathway engineering Monolignol biosynthesis occurs via the shikimate and general phenylpropanoid pathways prior to export and deposition into the plant cell wall (see Figure 2-2 for a detailed overview of the biosynthetic pathways including enzyme abbreviations) (Boerjan et al., 2003). Seven enzyme-catalysed steps of the plastid-localised shikimate pathway convert photosynthate-derived phosphoenolpyruvate and erythrose 4-phospate into chorismate, and further transformations yield the aromatic amino acids phenylalanine, 20 21 Figure 2-2. Lignin biosynthesis. Biosynthesis of monolignols via the shikimate, aromatic amino acid, and phenylpropanoid pathways. Photosynthate, supplied as sucrose and stored transiently as starch, is metabolised in sink tissues via glycolysis and the oxidative pentose phosphate pathway to produce phosphoenolpyruvate and erythrose 4-phosphate. These in turn are converted via the shikimate pathway into chorismate. The aromatic amino acid pathway yields phenylalanine, tyrosine and tryptophan (not shown). Finally, the monolignols, primarily p-coumaryl alcohol, coniferyl alcohol and sinapyl alcohol, are produced through the general phenylpropanoid pathway and the monolignol-specific biosynthetic pathway. Enzymatic reactions and metabolite shuttling are shown in orange, the plastid as well as membrane transporters and the inner and outer plastid membranes are coloured in green, the ER and ER membranes are depicted in blue, and the plasma membrane and cell wall are highlighted in yellow. Note that not all routes shown have been demonstrated in all plants; for example: TAL has been found only in monocots, CSE activity has only been demonstrated in Arabidopsis so far, F5H is absent from most gymnosperms, and the route from p-coumarate directly to caffeate has only been demonstrated in poplar. The cascade of transcriptional regulation is shown in purple and includes proteins from the MYB, KNAT and NAC families of transcription factors as well as two subunits of the Mediator transcriptional co-regulator complex. Enzyme abbreviations: 3-deoxy-D-arabinoheptulosonate 7-phosphate synthase (DAHPS), 3-dehydroquinate synthase (DHQS), 3-dehydroquinate dehydratase–shikimate dehydrogenase (DHQD-SDH), shikimate kinase (SK), 5-enolpyruvylshikimate 3-phosphate synthase (EPSPS), chorismate synthase (CS), chorismate mutase (CM), prephenate aminotransferase (PAT), arogenate dehydrogenase (ADH), arogenate dehydratase (ADT), prephenate dehydrogenase (PDH), 4-hydroxyphenylpyruvate aminotransferase (HPPAT), prephenate dehydratase (PDT), phenylprephenate aminotransferase (PPAT), phenylalanine ammonia lyase (PAL), tyrosine ammonia lyase (TAL), cinnamate 4-hydroxylase (C4H), 4-coumarate CoA ligase (4CL), p-hydroxycinnamoyl-CoA:shikimate/quinate p-hydroxycinnamoyltransferase (HCT), p-coumaroyl-shikimate/quinate 3ˈ-hydroxylase (C3ˈH), caffeoyl shikimate esterase (CSE), caffeoyl-CoA O-methyltransferase (CCoAOMT), ferulate/coniferaldehyde 5-hydroxylase (F5H), caffeic acid/5-hydroxyconiferaldehyde O-methyltransferase (COMT), cinnamoyl-CoA reductase (CCR), and cinnamyl alcohol dehydrogenase (CAD). Other abbreviations: inorganic phosphate (Pi), S-adenosylmethionine (SAM), S-adenosylhomocysteine (SAH). 22 tyrosine and tryptophan. Phenylalanine is then shuttled to the cytosol where the first dedicated step of the general phenylpropanoid pathway is its deamination to produce cinnamic acid. Thereafter, an intricate succession of aromatic ring hydroxylations and catechol unit O-methylations interspersed with the activation of cinnamate as a CoA thioester and subsequent reduction via the aldehyde eventually yield the monolignols. There are numerous inter-species variations in the architecture of the lignin biosynthetic pathway, particularly in the preferred route through the metabolic grid. For example, grasses are uniquely capable of using tyrosine in addition to phenylalanine, and gymnosperms are generally unable to produce sinapyl alcohol because the requisite hydroxylase is absent (Wagner et al., 2015). Lignin engineering efforts were initially focused largely on the enzymes directly involved in the general phenylpropanoid and monolignol biosynthetic pathways. Mutants or transgenics with targeted downregulation of key biosynthetic genes in diverse plant species have shown varying levels of reduced lignin production (Vanholme et al., 2008). However, monolignol biosynthesis is highly plastic, allowing plants to substitute monolignols when one or more of the genes is disrupted or misregulated such that lignin composition and quantity are often concomitantly altered. For example, suppression of C3'H in hybrid poplar resulted in a 60% reduction in lignin as well as a shift toward H units (Coleman et al., 2008b), whereas knockdown of PAL in Brachypodium led to 43% less lignin and a relative increase in S and H units (Cass et al., 2015). On the other hand, the Arabidopsis fah1 mutant lacks a functional F5H, corresponding to a near-complete loss of S units but no significant change in total lignin content (Marita et al., 1999). As lignin structure is a reflection of the monomers available during polymerisation and the prevalence of different linkage types is a major determinant of chemical resilience, lignin composition represents an important parameter in biomass recalcitrance that rivals finite lignin content. For instance, the S to G lignin ratio in transgenic hybrid poplar lines has been positively correlated with cell-wall bioconversion rates (Mansfield et al., 2012). Non-canonical monolignols are also amenable to lignification, further emphasising the plasticity of lignin biogenesis. In transgenic poplar for example, downregulation of CCR has been shown to lead to the low-level incorporation of ferulic acid (Leplé et al., 2007), downregulation of CAD results in the incorporation of hydroxycinnamaldehydes (Ralph et al., 1997; Zhao et al., 2013), and a reduction in 23 COMT expression leads to a lignin derived, in part, from 5-hydroxyconiferyl alcohol (Marita et al., 2001). Similarly, suppression of CCoAOMT in transgenic pine cell cultures leads to the incorporation of caffeyl alcohol (Wagner et al., 2011). Interestingly, the seed coats of Vanilla planifolia and several Cactaceae species contain a lignin derived almost entirely from this monomer or the related 5-hydroxyconiferyl alcohol (Chen et al., 2012; Chen et al., 2013). More evidence of lignin plasticity is offered by the incorporation of γ-acylated monolignol conjugates, namely acetylated monolignols in kenaf, sisal, palm and abaca, p-hydroxybenzoylated monolignols in poplar, willow and palm, and p-coumaroylated monolignols in the commelinid monocots (Ralph, 2010). In effect, any compatible phenolic compounds present at the site of polymerisation in the cell wall are ‘candidates’ for radical transfer, radical–radical coupling, and lignin polymerisation (Ralph, 2006). Although covalent bonds have been shown to link lignin and some hemicelluloses, most notably via the dehydrodimerisation of ferulate on arabinoxylans in grasses (Ralph et al., 2004b), the direct attachment of lignin to hemicellulose remains difficult to authenticate (Du et al., 2014). Similarly, cell wall proteins could also become covalently bound to lignin, but this too remains largely unexplored (Cong et al., 2013). There has been remarkable progress in altering the amount and composition of lignin by targeting the general phenylpropanoid and monolignol biosynthetic pathways, but lignin-modified plants often exhibit developmental defects ranging from stem lodging to dwarfism. It may simply be that reduced lignin content alters structural integrity and impairs water transport, and such plants often have collapsed xylem cells (Voelker et al., 2011). However, perturbations to the biosynthetic pathway may also result in the overproduction of other phenylpropanoids and glycoside derivatives that provoke diverse pleiotropic effects (Bonawitz & Chapple, 2013). Besides lignin, the shikimate and phenylpropanoid pathways also give rise to an array of other primary and secondary metabolites. Moving forward, lignin engineering strategies should contemplate the effects of altered metabolic flux on related pathways and metabolites. Moreover, rewiring these metabolic networks will be an important element of innovative strategies that incorporate alternative monomers into designer lignins. 24 2.4 Designer lignins It is now eminently feasible to produce genetically engineered plants with severely reduced lignin levels delivering improved biomass processing; however, such plants are often less vigorous and agronomically inferior (Voelker et al., 2011; Bonawitz & Chapple, 2013). By engineering the chemical structure of lignin without drastically altering lignin content or functionality, it may be possible to satisfy the biological requirement for lignification while concomitantly reducing recalcitrance. Novel physicochemical properties could render lignin more easily extractable during processing and could even create new avenues in biomass utilisation (Vanholme et al., 2012a). At least five types of these designer lignins have been proposed: lignins with a lower degree of polymerisation, lignins that are less hydrophobic, lignins with fewer bonds to structural carbohydrates, lignins containing chemically labile bonds, and lignins designed to harbour value-added chemical moieties (Figure 2-3). Several examples of these designer lignins will be described here to illustrate the breadth of possibilities. 2.4.1 Shorter lignin chains Although conventional wisdom maintains that lignin polymers have a high molecular weight and are extensively cross-linked, recent evidence suggests that native lignin comprises relatively short oligomeric chains with only minimal branching (Crestini et al., 2011). Nonetheless, lignins with reduced degrees of polymerisation may be more readily extracted. Incorporation of monomers capable only of single coupling reactions that either initiate (‘starters’) or terminate (‘stoppers’) chain elongation could reduce average lignin polymer chain lengths. For example, dihydroconiferyl alcohol found in the lignin of gymnosperms (Ralph et al., 1997), as well as hydroxyphenylglycerols (Figure 2-3E) and hydroxybenzenoids detected in a variety of plants, can serve as lignin initiators (Ralph et al., 2004b; Eudes et al., 2012). A greater prevalence of these groups during active polymerisation may lead to more polymer initiation events resulting in more lignin chains that are shorter. Consistent with this approach, expression of the bacterial hydroxycinnamoyl-CoA hydratase-lyase gene in Arabidopsis lignifying cells did not alter plant growth or overall lignin content but did lead to sidechain shortening of monomers, the incorporation of p-hydroxybenzaldehyde and p-hydroxybenzoate groups, reduced lignin chain length, and improved 25 26 Figure 2-3. Lignin models and designer lignins. A: A poplar lignin model containing 20 units, S:G = 13:7. 4–O–5-Coupling was thought to produce branching, but this is now being questioned (Crestini et al., 2011); the unit (near the bottom of the structure) is shown here as a free-phenolic unit; p-hydroxybenzoates acylating some γ-OH groups are not shown. B: An all-S poplar lignin model, showing only the two types of primary units, β–β and β–O–4; such lignins are essentially linear and may have a low degree of polymerisation (i.e., are chain-shortened) (Stewart et al., 2009). C: Almost completely homogeneous (in terms of interunit linkage type) and linear lignins are produced, in vivo and in vitro, from atypical monolignols caffeyl alcohol (no 5-OMe) and 5-hydroxyconiferyl alcohol (Chen et al., 2012; Chen et al., 2013); such lignins are also devoid of lignin–polysaccharide cross-linking as rearomatisation of the quinone methide intermediate following β–O–4-coupling is via fast internal trapping by the o-phenol. D: 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 (Ralph, 2010; Wilkerson et al., 2014); readily cleavable ester bonds are shown hashed. Cleaving this oligomer containing 4 zip-conjugates cleaves this model lignin into 5 fragments; in general, a polymer containing n zip-conjugates will cleave into ≤(n + 1) fragments. E: A fragment of a chain-shortened polymer created by lignification using monomers, such as the guaiacylglycerol shown here, that can only start a lignin chain; this polymer is logically also more hydrophilic. F: A fragment of a particularly hydrophilic lignin polymer created in part using monolignol γ–O–1 β-glucosides in vitro (Tobimatsu et al., 2008). G: A fragment of lignin containing rosmarinic acid (purple) that has been incorporated into the chain via radical coupling (Tobimatsu et al., 2012). Such lignins would display various features, including being more hydrophilic (due to the acid group), having benzodioxane units (like the polymers in C) that preclude lignin-polysaccharide cross-linking at those sites, and zip-lignin signatures allowing the polymer to be readily cleaved by mild base (the cleavable ester linkage is shown hashed). H: An example of a high-value component in lignin–tricin end-units (magenta) occur in grass lignins (Lan et al., 2015). Tricin is a flavonoid and it is synthesised outside the monolignol biosynthetic pathway. I: Model of the extensive polysaccharide–polysaccharide (via arabinoxylan-bound ferulate dimerisation) and polysaccharide–lignin (via incorporation of the ferulates and diferulates (red, on arabinoxylan) into the lignin polymer) in all commelinid monocots (Ralph, 2010; Ralph et al., 2004b); again, readily cleavable ester bonds are shown hashed. The bonds formed by radical coupling reactions are bolded; bonds formed during post-coupling rearomatisation are grey, as are the OH groups from water addition (see Figure 2-1C). In all of the models, units derived from lignin monomers are in cyan (G) and green (S), whereas novel units are coloured uniquely. 27 saccharification (Eudes et al., 2012). An analogous tactic is to exploit the relative oxidation potential of different monomers in an effort to introduce more end-groups. For example, p-coumaryl alcohol favours radical transfer reactions and frequently occurs as free-phenolic endgroups such that H-rich lignins have a lower degree of polymerisation (Lapierre & Rolando, 1988). The S-rich lignin from transgenic poplar overexpressing F5H also appears to have a remarkably shorter chain length with an average degree of polymerisation of approximately 10 (Figure 2-3B; Stewart et al., 2009). 2.4.2 Less hydrophobic lignins Lignin is largely hydrophobic in nature and designer lignin polymers containing more hydrophilic constituents could be more soluble during delignification processes. Additionally, increased hydrophilicity could reduce hydrophobic interactions in the cell wall and improve enzyme accessibility during saccharification. Various alternative strategies are conceivable and candidate monomers may contain additional hydroxyl groups or conjugated hydrophilic moieties, such as carbohydrates (e.g., Figure 2-3E–G; Grabber et al., 2012). The opposite strategy could be favourable; molecular dynamics simulations of lignin with increased hydrophobicity predicted a reduction of noncovalent associations between lignin and hemicelluloses (Carmona et al., 2015) that could make the cell wall polysaccharides more accessible during saccharification. 2.4.3 Lignins with less structural cross-linking to carbohydrates In addition to non-covalent interactions, lignin is conjectured to be linked to hemicelluloses through various types of covalent bonds. Whenever a monolignol couples via its β position to the growing polymer (or to another monomer), a quinone methide intermediate is produced and, in the case of the dominant β-ether units, this is overwhelmingly quenched and re-aromatised via nucleophilic attack from water (Figure 2-1C; Ralph et al., 2004a). Alcohol and carboxylic acid groups inherent to hemicelluloses may also theoretically serve as nucleophiles giving rise to the benzyl ether or benzyl ester bonds that contribute to recalcitrant lignin–carbohydrate complexes, but compelling evidence for this is still lacking. Potentially useful novel units containing o-diphenol groups, such as those arising from caffeyl alcohol or 5-hydroxyconiferyl alcohol, result in quinone methide intermediates that are rapidly internally trapped, 28 forming benzodioxane structures (Figure 2-3C) before any possibility of external nucleophilic attack can occur. A number of similar candidate monomers that could reduce polysaccharide–lignin cross-linking have recently been validated in biomimetic in vitro studies, including rosmarinic acid, epicatechin, ethyl gallate and epigallocatechin (Grabber et al., 2012; Tobimatsu et al., 2012). As noted above, monocots have a distinctive mechanism for cross-linking cell wall polymers to strengthen the wall; arabinoxylan is acylated with ferulate moieties that can radically cross-couple to adjacent feruloylated hemicellulose chains or to lignin resulting in extensive polysaccharide–polysaccharide and polysaccharide–lignin cross-linking (Figure 2-3I; Ralph, 2010). Several groups are currently working to identify the acyltransferase enzyme(s) responsible for acylating the arabinosyl units on arabinoxylans with ferulate (Molinari et al., 2013). 2.4.4 Lignins with novel chemically labile bonds As has been shown, various perturbations of the monolignol biosynthetic pathway result in a shift in monomer composition that may yield more chemically labile bonds (Boerjan et al., 2003; Vanholme et al., 2008; Vanholme et al., 2012b). For example, ferulic acid incorporates into the lignin of CCR-deficient plants producing acid-labile acetals (Leplé et al., 2007), and overexpression of F5H in hybrid poplar results in a lignin containing nearly 98% S units yielding more alkali-labile β-ethers (Figure 2-3B; Stewart et al., 2009). One of the most highly sought after objectives has been the introduction of S lignin units into conifer tree species that normally contain the more recalcitrant G-rich lignin. Recently, an important proof-of-principle milestone was achieved with the simultaneous introduction of COMT and F5H genes into transgenic pine cell cultures (Wagner et al., 2015). The ‘zip-lignin’ strategy has also been heralded as a major breakthrough in designer lignins (Ralph, 2010; Wilkerson et al., 2014). Inspired by the proven incorporation of ferulates integrally into grass lignins, an exotic feruloyl-CoA:monolignol transferase from Angelica sinensis was introduced into hybrid poplar and resulted in a lignin in which chemically labile ester bonds had been integrated into the polymer backbone (Figure 2-3D), improving cell wall digestibility after mild alkaline pretreatment (Wilkerson et al., 2014). Recently, a bacterial Cα-dehydrogenase was shown to oxidise the α-hydroxyl groups in lignin and, when introduced into 29 transgenic Arabidopsis, yielded low levels of novel chemically labile α-keto-β-ether units in lignin (Tsuji et al., 2015). Finally, as amino acids are also capable of cross-linking to lignin, an alternative approach is to direct tyrosine-rich or cysteine-rich peptides to the cell wall such that protease enzymes could digest these cross-links and accelerate lignin digestion (Liang et al., 2008). 2.4.5 Value-added lignins Lignin extracted during industrial biomass processing is frequently used for its calorific value, being burnt to provide process energy. But recently, aspirations of complete biomass utilisation within the modern biorefinery have inspired an array of lignin-derived high-value products (Ragauskas et al., 2014). Research in this area has primarily focused on optimising lignin recovery and developing products from lignin that can compete with existing petroleum-derived materials. However, moving forward, inherently valuable lignin polymers could be developed to facilitate the production of novel high-value products using industrial lignin waste streams. The abundant aldehyde groups in the above-mentioned hydroxycinnamaldehyde-derived lignins from CAD-deficient plants (Ralph et al., 1997; Zhao et al., 2013) create enormous potential for functionalisation in diverse applications. Lignins derived solely from caffeyl alcohol or 5-hydroxyconiferyl alcohol monomers produce homogeneous linear lignin chains of β–O–4-derived benzodioxane units (Figure 2-3C; Chen et al., 2012; Chen et al., 2013). Such regularity would likely be beneficial in applications such as the generation of lignin-derived carbon fibres, due to its homogeneous structure and lower complexity. However, as intriguing as the possibilities are, it is not yet clear that large, healthy plants can be produced with such lignins. Nature herself is revealing pathways by which valued products could naturally be produced in lignins and is even showing how they can be arranged to be on the end of a chain where they are, presumably, easiest to cleave off. For example, it was recently discovered that the flavonoid tricin is naturally incorporated into monocot lignins, as (starting) endgroups (Figure 2-3H; Lan et al., 2015). This discovery, further illustrating the inherent plasticity of lignification, was particularly remarkable because tricin is not produced via the monolignol biosynthetic pathway so it exemplifies how, through thoughtful metabolic engineering, it may be possible to incorporate other unique and valuable chemical constituents into lignin. Given the abundance of surplus 30 lignin available in industry, it is plausible that even low-level production of value-added lignins could help economise total biomass utilisation in biorefineries. 2.5 Perspectives on lignin engineering A number of potential alternative monomers has been proposed, many of which have overlapping functionalities in designer lignins (Vanholme et al., 2012a). For example, rosmarinic acid is a hydrophilic compound with a chemically labile ester linkage and two o-diphenol groups (Figure 2-3G; Tobimatsu et al., 2012). The resulting lignin polymers would therefore be less hydrophobic, possess fewer links to hemicelluloses, and have readily cleavable ester bonds within lignin chains. Before embarking on plant metabolic engineering with novel lignin monomers, in vitro experiments and biomimetic test systems can be used to validate design strategies (Grabber et al., 2012, Tobimatsu et al., 2012). For example, with rosmarinic acid, these tests revealed no barriers to radical formation or lignin polymerisation in maize cell walls and pointed to significant improvements in saccharification (Tobimatsu et al., 2012). A number of additional alternative monomers are currently being evaluated through a similar pipeline and plant engineering work with the most promising candidates will soon follow (Vanholme et al., 2012a). The ‘zip-lignin’ strategy to introduce backbone esters exemplifies the potential of monolignol acyltransferases in lignin engineering (Wilkerson et al., 2014). Presently, genes encoding p-coumaroyl-CoA:monolignol transferase and feruloyl-CoA:monolignol transferase have been discovered (Wilkerson et al., 2014; Petrik et al., 2014), whereas genes for acetyl-CoA and p-hydroxybenzoyl-CoA analogues remain elusive. Recent surprising discoveries in the chemical variability of natural lignins leave room to ponder what additional monolignol acylation possibilities may exist and whether corresponding acyltransferases might be found in Nature. Coupled with innovative metabolic pathway engineering to supply these alternative monomers, transformational changes in lignin engineering appear to be within reach. As the regulation of monolignol biosynthesis is largely enacted through transcriptional control, considerable attention has been devoted to demystifying these processes and harnessing them to actively switch biosynthesis on or off at will. In addition to the array of MYB family transcription factors that 31 activate monolignol biosynthetic genes and the NAC family transcription factors that serve as master switches for secondary cell wall biosynthesis, subunits of the transcriptional co-regulator Mediator have recently been identified as homeostatic repressors of monolignol biosynthesis and all of these show promise in lignin engineering (Schuetz et al., 2013; Bonawitz et al., 2014; Figure 2-2). Although many designer monomers may originate from the phenylpropanoid pathway directly, other monomers may have different metabolic origins and regulatory constraints. It also remains to be seen how designer lignins might ensure adequate cell wall properties in the context of recently proposed cell wall integrity models (Bonawitz & Chapple, 2013; Yang et al., 2013). Although monolignol transport mechanisms remain obscure, recent evidence points to active transport through membrane-bound transporter proteins (Alejandro et al., 2012). The realisation that alternative monomers are also translocated to the site of polymerisation in the cell wall casts some doubt on these concepts. It may be that monolignol transporters are exceptionally non-specific, or perhaps that monolignols are not exported by active transport whatsoever. Similarly, several peroxidase and laccase enzymes have been implicated in the dehydrogenation of monolignols (Lu et al., 2013); however, these proteins are also apparently non-selective as non-canonical monomers are routinely incorporated. Alternatively, it may be that these monomers are not accepted directly and rely solely on radical transfer reactions to enter polymerisation. Although there are no obvious barriers for export, radical formation, and incorporation of non-canonical monomers into lignin polymers, these mechanisms represent underexploited avenues in lignin engineering. Designer lignins invariably possess novel physicochemical properties that could potentially have adverse effects on plant physiology and development. Manipulations of the monolignol biosynthetic pathway have been known to produce plants with reduced growth, developmental defects, increased susceptibility to disease and water transport difficulties (Bonawitz & Chapple, 2013; Zhao & Dixon, 2014). Although not all lignin-modified plants are compromised and such widespread pleiotropic phenotypes may not be purely a consequence of reduced lignin content or altered composition, it emphasises the biological importance of lignin in plants. In an effort to produce agronomically viable plants, broader plant metabolism should be considered and it may be prudent to use tissue-specific 32 promoters to drive transgene expression. Spatiotemporal control may restrict designer lignins to xylem tissues, or xylem fibres more specifically (Eudes et al., 2014). Additionally, yield penalties could perhaps be overcome by vessel-specific complementation (Petersen et al., 2012). Given the diversity of lignin structures tolerated by plants in Nature, there is every reason to believe that some novel designer lignins will be compatible with normal plant growth. Moreover, it is imperative that all these novel transgenic strategies are appropriately field-tested to ensure that the observed trait modifications withstand normal growing conditions. 2.6 Conclusions Lignin engineering has evolved beyond simple perturbations of the general phenylpropanoid and monolignol biosynthetic pathways, culminating in a suite of designer lignins with novel physicochemical properties. Much like the Trojan horse used to covertly circumvent the wall defences of the ancient city of Troy, designer lignins may satisfy the biological requirement for lignin in plant cell walls while providing improved biomass utilisation efficiency. Moreover, high-value designer lignins have been conceived that could valorise lignin waste streams in biomass refineries, and could be targeted toward upgrading processing. Lignification shows remarkable plasticity, but the development of agronomically viable biomass feedstocks featuring designer lignins will require thoughtful selection of non-canonical monomers, validation in biomimetic systems, careful metabolic engineering, thorough assessment of pleiotropic effects that could potentially accrue from the incorporation of novel components, field trials, and the generation of sufficient volumes of material for industrially relevant bioconversion assessment. Although possibilities abound, maintaining plant health is paramount and, ultimately, the plants themselves will dictate which of these approaches can be tolerated. At the dawn of this new era in lignin engineering, we are limited only by the biological constraints of lignification and by our collective imagination. 33 Chapter 3: Suppression of chorismate mutase 1 in hybrid poplar reveals possible redundancy in the supply of lignin precursors 3.1 Summary Chorismate is an important branchpoint metabolite in the biosynthesis of lignin and a wide array of other metabolites in plants. Chorismate mutase (CM), the enzyme responsible for transforming chorismate into prephenate, is a key regulator of metabolic flux towards the aromatic amino acids and onward to lignin. We identified three CM genes in poplar (Populus spp.) and used RNA interference (RNAi) to suppress the expression of Pa×gCM1, the most highly expressed isoform in xylem. Although this strategy was successful in disrupting Pa×gCM1 transcripts, there was also an unanticipated increase in lignin content, a shift towards guaiacyl lignin units, and more xylem vessels with smaller lumen areas. This was accompanied by increased expression of the other two CM isoforms, Pa×gCM2 and Pa×gCM3, as well as widespread changes in gene expression and metabolism. This study provides evidence of possible redundancy within the CM gene family, at least in the developing xylem of poplar. 3.2 Introduction Lignin is a phenolic polymer found in vascular plants wherein it provides structural support and contributes to water conduction and defence. The deposition of lignin rigidifies secondary cell walls and helps adjoining cells adhere by reinforcing the middle lamella (Gibson, 2012; also see Chapter 6). Lignified cell walls in the plant vasculature have reduced permeability which enables long-distance water transport (Sperry, 2003). The insolubility and stability of lignin provides a physical and chemical barrier that helps guard against pests and pathogens (Miedes et al., 2014). However, the recalcitrance of lignin to biological and chemical degradation hinders efficient industrial processing of plant biomass (Li et al., 2016). As such, there is tremendous value in studying lignin biosynthesis. Lignin is assembled primarily from three hydroxycinnamate-derivatives known as monolignols – p-coumaryl, coniferyl, and sinapyl alcohol – that give rise to p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) units in lignin polymers (Boerjan et al., 2003). The monolignols are produced via the phenylpropanoid pathway from phenylalanine and, in grasses, from tyrosine as well (Cass et al., 2015; 34 also see Chapter 2). As lignin typically comprises 20–35% of the dry weight of plants, vast amounts of photosynthetic carbon are allocated to the phenylpropanoid pathway. Lignin formation is energetically costly, and it has been estimated that 0.975 – 1.724 moles of translocated sucrose are needed to produce every mole of monolignol, depending upon which specific biosynthetic route is followed (Amthor, 2003). Accordingly, lignin biosynthesis is tightly regulated at the levels of transcription, post-translational modification, allostery, and the supply of substrates for polymerisation (Whetten & Sederoff, 1995; Zhong & Ye, 2009; Wang et al., 2019). The shikimate and aromatic amino acid pathways, which lie upstream of phenylpropanoid biosynthesis, are responsible for the production of phenylalanine and tyrosine (Maeda & Dudareva, 2012). Through a series of enzymatic conversions, the shikimate pathway transforms phosphoenolpyruvate and erythrose 4-phosphate, products of carbohydrate metabolism via glycolysis and the pentose phosphate pathway, into chorismate (Bonner & Jensen, 1998). In addition to the phenylpropanoids, chorismate is also a precursor to folates, indoles, phylloquinone, anthranilate, and salicylic acid, as well as proteins via phenylalanine, tyrosine, and tryptophan (Herrmann, 1995; Mustafa & Verpoorte, 2005; Bowsher et al., 2008; Tzin & Galili, 2010; Tohge et al., 2013). Consequently, chorismate represents an important branchpoint at the junction of a plethora of metabolites in plants (Figure 3-1). Chorismate mutase (CM) catalyses the pericyclic Claisen rearrangement of chorismate to form prephenate, a precursor to both phenylalanine and tyrosine (Gorisch, 1978; Romero et al., 1995). Previous reports suggest that vascular plants typically have two or three CM isozymes with one of these occurring in the cytosol (Woodin et al., 1978; d’Amato et al., 1984; Poulsen & Verpoorte, 1991). In Arabidopsis, for example, CM1 and CM3 contain plastid-localisation signals and are allosterically regulated by aromatic amino acids, whereas CM2 is cytosolic and allosterically insensitive (Eberhard et al., 1996; Mobley et al., 1999). More recently, a complete cytosolic pathway for the biosynthesis of aromatic amino acids from chorismate has been uncovered (Qian et al., 2019), further illustrating the metabolic divarications that occur at chorismate. 35 Figure 3-1. Chorismate is a key branchpoint metabolite in plants. A biosynthetic map depicting the shikimate, aromatic amino acid (AAA), and phenylpropanoid biosynthetic pathways. Chorismate mutase (CM) is an important metabolic gatekeeper enzyme upstream of lignin and phenylpropanoid biosynthesis. The contributions of various CM isoforms to lignin biosynthesis have not been extensively characterised. However, CM1 is the most strongly expressed of the three CM genes in the stems of Arabidopsis (Ehlting et al., 2005) and poplar (Tsai et al., 2006), in which there exist large metabolic requirements for phenylalanine to supply lignin precursors. Conversely, CM2 and CM3 are not normally highly expressed during xylem differentiation. Herein, we used RNA interference (RNAi) to suppress the expression of CM1 in hybrid poplar (Populus alba × grandidentata; P39) in order to investigate the role of CM in lignification. 36 3.3 Materials and Methods 3.3.1 Phylogenetic analysis The three CM sequences from Arabidopsis (AtCM1, AT3G29200, Eberhard et al., 1993; AtCM2, AT5G10870, Eberhard et al., 1996; and AtCM3; AT1G69370, Mobley et al., 1999) were used as queries in BLAST searches to identify CM homologues from the reference genomes of Populus trichocarpa (JGI v4.1), Amborella trichopoda (AGP v1.0), Brachypodium distachyon (JGI v3.2), Brassica rapa (JGI FPsc v1.3), Eucalyptus grandis (JGI v2.0), Medicago truncatula (MTGD Mt4.0), Oryza sativa (RGAP v7.0), Phaseolus vulgaris (JGI v2.1), Physcomitrella patens (JGI v3.3), Picea abies (SGP v1.0), Selaginella moellendorffii (JGI v1.0), Solanum tuberosum (PGSC v4.03), Vitis vinifera (Genoscope 12X v2.1), and Zea mays (MGSP B73 RefGen_v4). These amino acid sequences were aligned using MEGA X with the MUSCLE algorithm and the default parameters (Kumar et al., 2018), and then used in the assembly of a maximum likelihood phylogenetic tree. CM sequences from the yeast Saccharomyces cerevisiae (ScCM, YPR060C, Ball et al., 1986), the red alga Porphyra umbilicalis (Pum2195s0001.1.p, JGI v1.5), and the green alga Chlamydomonas reinhardtii (Cre03.g155200.t1.1, JGI v5.6) were used as an outgroup, and the tree was resampled using 1000 bootstrap iterations. Pairwise alignments of the poplar and Arabidopsis homologues were then inspected more closely (Figure 3-2A). Sequence identity and similarity values were calculated using the SMS Ident and Sim tool (Stothard, 2000), and signal peptide predictions were performed using TargetP-2.0 (Armenteros et al., 2019). 3.3.2 RNAi-mediated suppression of chorismate mutase 1 in poplar The CM1 coding sequence was cloned from hybrid poplar (Populus alba × grandidentata; P39). RNA was isolated using a CTAB-based protocol (Kolosova et al., 2004) from the leaves of poplar plants which were flash-frozen in liquid nitrogen and ground into a powder with a mortar and pestle. This total RNA preparation was treated with DNase using the Turbo DNA-free kit (Life Technologies, Carlsbad, CA, U.S.A.) and then used in the synthesis of first-stand cDNA using the iScript cDNA synthesis kit (Bio-Rad Laboratories Inc., Hercules, CA, U.S.A). Primers designed to anneal within the upstream and downstream regions were initially used for gene amplification and were designed based on the Populus trichocarpa 37 Figure 3-2. Chorismate mutase sequences. A: Alignments of the three chorismate mutase (CM) isoforms from poplar and Arabidopsis, as labelled. Conserved amino acid residues are shown in bold. The active site and effector site residues identified by Westfall et al. (2014) are highlighted in green and purple, respectively. Percent identity and similarity are shown below each alignment. Predicted signal peptide cleavage sites for PtCM1 and PtCM3 are denoted with black arrowheads. B: The Pa×gCM1 nucleotide sequence cloned from hybrid poplar (Populus alba × grandidentata; P39). Polymorphisms compared to the reference genome of Populus trichocarpa are shown in blue. The region selected for the RNAi hairpin construct is highlighted in orange. 38 reference genome (see Table 3-1 for all primer sequences and reaction conditions). After sequencing this amplicon to identify any polymorphisms (Figure 3-2B), a 400-bp fragment was amplified with attB adaptor sequences to enable Gateway cloning, and then transferred into the pDON221 vector using BP Clonase (Life Technologies). Finally, an RNAi expression cassette was assembled by transferring this fragment from pDONR221 into pHELLSGATE12 (Helliwell & Waterhouse, 2003) using LR Clonase (Life Technologies). Chemically competent Agrobacterium tumefaciens EHA105 was transformed with the pHELLSGATE12 expression cassette following a freeze-thaw protocol (Wise et al., 2006). Next, leaf discs harvested from hybrid poplar plants grown in tissue culture were used for Agrobacterium-mediated transformation, as described previously (see section 4.3.2). Positive transformants were selected on media containing hygromycin and were confirmed using semi-quantitative RT-PCR. Of the 20 PCR-positive lines that were obtained, the most promising 3 lines were propagated in tissue culture. All biological replicates were derived from apical stem pieces after several rounds of propagation. Well-rooted plants were transferred into potting soil and grown in a greenhouse under natural lighting supplemented with high-pressure sodium lamps (minimum 500 µmol m-2s-1). The trees were supported with bamboo poles, watered daily with fertigated water, and monitored for any signs of pests or disease. After 20 weeks, the trees were harvested and the leaves and bark were carefully removed. Young leaves were collected and developing xylem tissue was scraped from the debarked stems, and then both were flash-frozen in liquid nitrogen and ground into a powder with a mortar and pestle. The debarked stems were air dried and subsequently comminuted into a powder using a Wiley mill fitted with a 40-mesh sieve. 3.3.3 Gene expression RNA was isolated from the frozen powder of young leaves and developing xylem as described above. The RNA was then treated with DNase and converted into first-strand cDNA. These cDNA preparations were diluted 8-fold and subjected to RT-qPCR analysis with gene-specific primers that had been tested over a wide range of template concentrations and found to have good priming efficiency. Thermal cycling, fluorescence detection, and melt-curve analysis was performed using a CFX 96 Real-Time PCR 39 Table 3-1. Primers and PCR programmes for Chapter 3. Details are provided for all PCR reactions used in construct assembly, transformant screening, and RT-qPCR. Description: Primer pair (5' → 3'): PCR reaction conditions: Amplification of the CM1 gene from hybrid poplar (Populus alba × grandidentata; P39) using primers that anneal in the UTR sequences Forward primer: GTC TAT TTG GCC GGT CAC TC Reverse primer: CAA CAC ATT TCC CAC TTT CC Primers designed based on the Populus trichocarpa reference genome. Amplicon size: 1305 bp 5 µL ...................................... 5× buffer 0.5 µL .............. 10 µM forward primer 0.5 µL ................ 10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ........................... template DNA 0.2 µL ......... iProof proofreading DNA polymerase (Biorad) Up to 25 µL ................ deionised water With thermal cycling as follows: 98°C for 30 s, 34 cycles of (98°C for 30 s, 55°C for 30 s, 72°C for 30 s), and then 72°C for 10 min. Note: template DNA was the cDNA from hybrid poplar leaves. Amplification of a 400-bp fragment of the CM1 gene from hybrid poplar, and addition of attB adaptor sequences to enable Gateway cloning Forward primer: GGG GAC AAG TTT GTA CAA AAA AGC AGG CTA TGG AGG CTA AGC TGT TAG Reverse primer: GGG GAC CAC TTT GTA CAA GAA AGC TGG GTG TAG CTT TTC AGT TTC CTT AAG GAT G Primers designed based on the consensus sequencing results of the CM1 gene isolated from hybrid poplar. Amplicon size: 400 bp 5 µL ...................................... 5× buffer 0.5 µL .............. 10 µM forward primer 0.5 µL ................ 10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ........................... template DNA 0.2 µL ......... iProof proofreading DNA polymerase (Biorad) Up to 25 µL ................ deionised water With thermal cycling as follows: 98°C for 30 s, 34 cycles of (98°C for 30 s, 55°C for 30 s, 72°C for 30 s), and then 72°C for 10 min. Note: template DNA was a plasmid containing the full-length CM1 gene plus parts of the 5' and 3' UTR sequences. Semi-quantitative RT-PCR used to select transgenic lines for further study Forward primer: GAA GGA GAT GAC GGC AAT TG Reverse primer: GGC AGC CTC ATA ATC ATC TG Primers designed based on the consensus sequencing results of the CM1 gene isolated from hybrid poplar. Amplicon size: 138 bp 2.5 µL ..................................10× buffer 0.5 µL ......................... 50 mM MgSO4 0.5 µL .............. 10 µM forward primer 0.5 µL ................ 10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ......................... template cDNA 0.2 µL .... Taq DNA polymerase (NEB) Up to 25 µL ................ deionised water With thermal cycling as follows: 94°C for 3 min, 34 cycles of (94°C for 30 s, 50°C for 30 s, 72°C for 30 s), and then 72°C for 5 min. Note: template cDNA was prepared with RNA isolated from the leaves of tissue-culture-grown poplars. Note: This table continues on the following page. 40 Note: Table continued from the preceding page. Description: Primer pair (5' → 3'): PCR reaction conditions: Quantification of expression levels using RT-qPCR For Pa×gCM1, Forward primer: GCT CAG CTT GGC AGA TAC AA Reverse primer: AAT GGG ATG TAG TAC CTG TGG Primers designed based on the consensus sequencing results of the CM1 gene isolated from hybrid poplar. Amplicon size: 108 bp Priming efficiency: 99.0% For elongation factor 1β (reference), Forward primer: GGC ATT AAG TTT TGT CGG TCT G Reverse primer: GCG GTT CAT CAT TTC ATC TGG Amplicon size: 97 bp Priming efficiency: 99.9% 5 µL .... BrightGreen qPCR master mix 0.3 µL .............. 10 µM forward primer 0.3 µL ................ 10 µM reverse primer 1 µL ............................ template cDNA Up to 10 µL ................ deionised water With qPCR parameters as follows: 95°C for 30 s, 39 cycles of (95°C for 5 s, 60°C for 15 s, plate read for fluorescence), and then followed by a melt-curve analysis as follows: ramp from 55°C to 95°C at 0.5°C per increment with hold-times of 5 s and a plate read for fluorescence. Note: template cDNA was prepared with RNA isolated from the young leaves and developing xylem of poplar. 41 instrument with the corresponding CFX Manager software (Bio-Rad Laboratories). Primers specific to the poplar elongation factor 1β gene were used as a reference for the calculation of relative expression. Prior to RNA-seq analysis, RNA preparations from developing xylem were assessed for yield and quality using an Agilent 2100 Bioanalyzer (Agilent Technologies) to ensure that all samples had RNA integrity numbers exceeding 9. Next, sequencing libraries were prepared using the TruSeq Stranded mRNA kit (Illumina Inc., San Diego, CA, U.S.A.) with 500 ng of purified RNA, and high-throughput sequencing was performed using a NextSeq 500 sequencer (Illumina Inc.). Sequence reads were trimmed and then aligned to the Populus trichocarpa reference genome. Differentially expressed genes were evaluated based on triplicate samples of line 3 and WT xylem RNA, and expressed as log2(fold change in expression). Finally, a gene ontology (GO) term enrichment analysis was performed with the PopGenIE tool using a false discovery rate correction for p-values (Sundell et al., 2015) with genes that were significantly upregulated (i.e., log2(fold change in expression) > 2) or significantly downregulated (i.e., log2(fold change in expression) < -2) in the xylem of line 3 compared to WT. Plots were assembled using Adobe Illustrator. 3.3.4 Lignin content, composition, and histology Extractive-free material was first obtained by extracting 40-mesh wood powder with hot acetone for 24 h in a Soxhlet apparatus. The total Klason lignin content was then determined using samples of extractive-free wood as the sum of acid-soluble and acid-insoluble fractions, and the composition of structural polysaccharides was measured by high performance anion-exchange chromatography of the hydrolysates, as described previously (see section 4.3.10). Thioacidolysis reactions were then performed to evaluate the lignin composition, as described previously (Robinson & Mansfield, 2009). Statistically significant differences between each line and the WT control were evaluated using a one-way ANOVA with a post-hoc Dunnett’s test conducted using SPSS Statistics 27 (IBM, Armonk, NY, U.S.A.). Samples for NMR analysis were first ball-milled and treated with Cellulysin cellulase (from Trichoderma viride, Calbiochem, MilliporeSigma, Burlington, MA, U.S.A.) to prepare enzyme-lignin, which was then dissolved in 4:1 dimethyl sulphoxide-d6:pyridine-d5. Two-dimensional 1H–13C 42 heteronuclear single-quantum coherence (2D-HSQC) NMR spectra were collected with a Biospin AVANCE 700 MHz spectrometer (Bruker Corp., Billerica, MA, U.S.A.), as described previously (Kim & Ralph, 2010). Transversal xylem cross-sections with a thickness of 10 µm were cut from the base of poplar stems using a sliding block microtome. Sections were stained with a 2:1 mixture of 3% phloroglucinol and concentrated hydrochloric acid, and then imaged using a Reichert-Jung Polyvar microscope (Leica Microsystems, Wetzlar, Germany) equipped with an Olympus UC90 camera (Olympus Corp., Tokyo, Japan). The average number of vessels per 1000 × 1000 µm area was evaluated by examining 23 non-overlapping images representing over 100,000 cells for line 3 (including more than 6000 vessels) and 31 images representing over 150,000 cells for WT (including more than 5000 vessels). Cell lumen areas were measured from the same images using an automated cell mapping and measurement pipeline with CellProfiler 4.1.3 (Carpenter et al., 2006). 3.3.5 Metabolites Frozen powders of developing xylem scrapings and young leaves were lyophilised overnight using a freeze-drier. Approximately 15 mg of dry material was extracted twice with 2.5 mL of 80% ethanol for 24 h. Prior to extraction, 75 µL of 1 mg mL-1 sorbitol was added as an internal standard. After combining the extracts, a 1 mL-aliquot was dried under a stream of nitrogen and then derivatised by silylation. Trimethylsilylated derivatives were obtained by adding 500 µL each of acetonitrile and N-methyl-N-(trimethylsilyl)trifluoroacetamide containing 1% trimethylchlorosilane to the dried residue, and incubating for 1 h at 70 °C. Samples were then analysed using an Agilent 7890A gas chromatograph (Agilent Technologies Inc., Santa Clara, CA, U.S.A.) equipped with an Rtx-5MS capillary column (5% diphenyl/95% dimethylpolysiloxane; 30 m × 250 μm × 0.25 μm film thickness) coupled to an Agilent 5975C inert XL mass spectrometer, as described previously (Tschaplinski et al., 2012). Metabolite concentrations were expressed as μg g-1 dry-weight material on a basis of sorbitol equivalents, and then used to calculate the fold difference relative to WT control plants. Heat maps were assembled using Adobe Illustrator. 43 Methanolic extracts were obtained by incubating 10-mg samples of developing xylem in 80% methanol containing 1% acetic acid at 50 °C in a hybridiser-incubator for 3 h. Aliquots were then transferred into a fresh tube, the solvent was removed by evaporation in a vacuum centrifuge, an equal volume of 2 M HCl was added, and acid hydrolysis was conducted at 90 °C for 2 h. The total amount of salicylic acid was then measured using an Agilent Infinity II 1290 UHPLC apparatus (Agilent Technologies Inc., Santa Clara, CA, U.S.A.) equipped with a Zorbax Eclipse Plus C-18 column (2.1 × 50 mm, 1.8 µm particle size, Agilent) and a diode array detector with the wavelength set to 304 nm. Good separation of peaks was achieved using an injection volume of 1 µL, a flow rate of 0.3 mL min-1, and a binary gradient of 5% to 30% of acetonitrile containing 0.01% trifluoroacetic acid (eluent A) in water containing 0.01% trifluoroacetic acid (eluent B) over 9 min. 3.4 Results and Discussion 3.4.1 Identification of poplar CM isoforms Amino acid sequences of the three CM isozymes from Arabidopsis were used to identify homologues in 14 diverse plant genomes. Among these species, there were as many as five isoforms per taxon (i.e., Brassica rapa) and as few as two (i.e., Solanum tuberosum, Eucalyptus grandis, and Picea abies). A maximum likelihood analysis of these putative CM sequences resulted in two major clades (Figure 3-3) – one containing AtCM2, the cytosolic CM from Arabidopsis (shown in orange), and the other containing the two plastidic forms, AtCM1 and AtCM3 (shown in blue). In accordance with previously published structure-functional analyses (Westfall et al., 2014), nearly all of the sequences in the plastidic cluster contained a glycine at position 148 which is necessary for allosteric regulation, with only one exception from Brachypodium distachyon (Bradi1g25640.1). Conversely, all sequences in the AtCM2-containing clade lacked this conserved glycine except for one sequence from Vitis vinifera (VIT 204s0008g06570.1). Interestingly, none of the CM sequences from Physcomitrella patens, Selaginella moellendorffii, Picea abies, or Amborella trichopoda contained this conserved residue, and these sequences predominantly clustered into a separate clade (shown in purple). 44 Figure 3-3. Phylogenetic analysis of the chorismate mutase gene family. A maximum likelihood tree showing the relationships among the amino acid sequences of chorismate mutase (CM) isoforms from diverse plant species including: Amborella trichopoda (AmTr), Arabidopsis thaliana (AT), Brachypodium distachyon (Bradi), Brassica rapa (Brara), Eucalyptus grandis (Eucgr), Medicago truncatula (Medtr), Oryza sativa (Os), Phaseolus vulgaris (Phvu), Physcomitrella patens (Pp), Picea abies (MA), Populus trichocarpa (Potri), Selaginella moellendorffii (Sm), Solanum tuberosum (PGSC), Vitis vinifera (VIT), and Zea mays (Zm). The CM sequences from Saccharomyces cerevisiae, Porphyra umbilicalis, and Chlamydomonas reinhardtii were included as an outgroup (coloured in green). Putative cytosolic CM sequences are coloured in orange, plastidic CM sequences are coloured in blue, and the sequences from S. moellendorffii, P. patens, P. abies, and one of the sequences from A. trichopoda are coloured in purple. The values at each node represent statistical confidence based on 1000 bootstrap iterations. The asterisk denotes PtCM1. Three isoforms of CM were identified in the reference genome of poplar and were assigned names based on phylogeny and sequence similarity with the Arabidopsis homologues. The poplar protein sequences were found to be 58–62% identical to Arabidopsis (Figure 3-2A). Both PtCM1 (Potri.018G019250) and PtCM3 (Potri.010G162300) contain the conserved effector site glycine residue at positions 202 and 199, respectively, which confers allosteric sensitivity. There is also strong predictive 45 support for the presence of plastid transit signals in both isoforms. In contrast, PtCM2 (Potri.018G019250) lacks this conserved residue and shows no evidence of a plastid signal. The occurrence of glycine residues at positions 138 and 135 in both PtCM1 and PtCM3 suggests that neither isoform exhibits the sensitivity to cysteine or histidine effectors which has been identified in AtCM3 due to an aspartate residue at this position (Westfall et al., 2014). In short, PtCM1 and PtCM3 appear to be plastidic isoforms that are most likely allosterically activated by tryptophan and repressed by phenylalanine and tyrosine, whereas PtCM2 appears to be cytosolic and unregulated. 3.4.2 Downregulation of CM1 in hybrid poplar An RNAi approach was used to suppress the expression of Pa×gCM1, the orthologue of PtCM1, in hybrid poplar (Populus grandidentata × Populus alba; P39). Three independent transformants were selected for in-depth study (Figure 3-4A). As the three CM genes have relatively low sequence similarity to one another, it was possible to design a hairpin target specific for CM1 transcripts (Figure 3-2B). This strategy was successful in misregulation of Pa×gCM1. Although lines 1 and 2 exhibited no obvious growth or developmental defects, line 3 poplars were extremely dwarfed compared to wild-type (WT) control trees (Figure 3-4B). All biological replicates of this line were consistently shorter in stature and thinner in stem diameter. Stunting is frequently observed as a response to perturbations in lignin production, often due to compromised vascular systems (Muro-Villanueva et al., 2019). However, dwarfing can also result from myriad other metabolic or hormonal imbalances in plants (Bonawitz & Chapple, 2013; Hollender & Dardick, 2015). The leaves of line 3 poplars were also small and showed obvious signs of chlorosis (Figure 3-5), even when grown under optimal conditions in tissue culture. Gene expression analysis by RT-qPCR confirmed reduced expression of Pa×gCM1 transcripts in all three lines (Figure 3-4C). However, line 3 poplars were evidently the most downregulated in Pa×gCM1. Surprisingly, the expression of Pa×gCM2 and Pa×gCM3 was also altered. This was most pronounced in the developing xylem of line 3, wherein the expression of Pa×gCM2 was more than doubled and Pa×gCM3 expression was increased by more than 10-fold. These unexpected observations suggest that CM gene expression can be modulated in compensation, at least in xylem. 46 Figure 3-4. RNAi-mediated suppression of CM1 in hybrid poplar. A: Representative photos of each line. B: Heights and diameters at the time of harvest. Error bars show standard deviation. The values marked with an asterisk are significantly different from the wild-type (WT) control, depicted with horizontal dashed lines (one-way ANOVA with Dunnett’s test, n=5 for each line with technical triplicates, p-value < 0.05). C: Fold change in expression of CM1, CM2, and CM3 in developing xylem and leaves relative to WT. The horizontal dashed line represents no change in expression. The black arrowheads denote that error bars are not shown for the xylem of line 3 because the biological replicates were pooled and analysed in triplicate due to limited samples. Otherwise, three biological replicates were analysed in triplicate for each line. 47 Figure 3-5. Images of greenhouse-grown Pa×gCM1 RNAi poplar trees. A: The stunted tree at the front is one of the biological replicates of line 3. B: A closer view showing the chlorotic leaves of line 3. 3.4.3 Compensatory lignification in CM1-suppressed poplar To examine how reduced expression of Pa×gCM1 affected xylem lignification, the lignin content and composition of mature wood were examined next. Rather than a decrease in lignin content, line 3 exhibited a 10% increase in the total amount of lignin compared to WT (Figure 3-6A), likely resulting from the compensatory expression of Pa×gCM2 and Pa×gCM3. This change in lignin content was accompanied by an increase in the proportion of G-lignin units measured by thioacidolysis (i.e., decreased S:G lignin ratio; Figure 3-6A). Although there were no significant differences in the total lignin contents of lines 1 and 2 compared to the WT control, these lines also had increased G-lignin content. The shift towards coniferyl alcohol-derived G-lignin units could perhaps be a result of altered flux through the phenylpropanoid pathway, consistent with previous reports on transgenic poplars (Wang et al., 2018a). An analysis of the structural polysaccharides also revealed differences in composition (Table 3-2), perhaps reflecting broader changes in cell wall biosynthesis and assembly. 48 Figure 3-6. Lignin analysis of Pa×gCM1 RNAi poplar trees. A: Klason lignin content was measured as the sum of acid-insoluble and acid-soluble fractions, and is plotted on the left axis in blue. The ratio of syringyl (S): guaiacyl (G) lignin monomers was evaluated by thioacidolysis, and is plotted on the right axis in orange. Five biological replicates were analysed in triplicate for each line and the error bars show standard deviation, except for line 3 wherein the biological replicates were pooled and analysed in triplicate due to limited samples (denoted with a black arrowhead). The values marked with an asterisk are significantly different from the wild-type (WT) control, depicted with horizontal dashed lines (one-way ANOVA with Dunnett’s test, n=5 for each line with technical triplicates, p-value < 0.05). B: Partial two-dimensional 1H–13C HSQC NMR spectra for enzyme-lignin samples of line 3 and the WT control, as labelled, showing the aromatics region. The colour-coding and peak annotations for p-hydroxyphenyl (H, light purple), guaiacyl (G, blue), and syringyl units (S and S', dark and light magenta), as well as p-hydroxybenzoate pendent groups (yellow, pHB) are elaborated with the structures shown on the side. Percentages are expressed on a basis where H+G+S = 100%. 49 Table 3-2. Structural polysaccharides in Pa×gCM1 RNAi poplar trees. The values shown in bold are significantly different from the wild-type (WT) control (one-way ANOVA with post-hoc Dunnett’s test, n=5 for each line with technical triplicates, p-value < 0.05). The data corresponding to line 3 was not included in the ANOVA analysis since the biological replicates were pooled due to limited samples. These values represent mean ± standard error for technical replicates. Otherwise, mean ± standard deviation is provided for three biological replicates. Sugar (% w/w) Line 1 Line 2 Line 3 WT Glucose 42.14 ± 1.18 43.73 ± 1.28 41.21 ± 0.48 46.33 ± 0.79 Xylose 16.22 ± 0.34 17.04 ± 0.95 14.83 ± 0.15 17.48 ± 0.28 Mannose 1.93 ± 0.09 1.69 ± 0.16 0.96 ± 0.01 1.55 ± 0.06 Galactose 0.61 ± 0.04 0.68 ± 0.04 0.90 ± 0.01 0.74 ± 0.03 Rhamnose 0.44 ± 0.01 0.40 ± 0.01 0.44 ± 0.01 0.43 ± 0.01 Arabinose 0.28 ± 0.01 0.32 ± 0.03 0.37 ± 0.01 0.34 ± 0.02 The observation that lignin content was in fact elevated in line 3 relative to WT shows that compensatory expression of Pa×gCM2 and Pa×gCM3 was more than sufficient to satisfy the demand for chorismate during xylem lignification. However, the apparent over-supply of lignin precursors may be a sign that chorismate formation driven by Pa×gCM2 and Pa×gCM3 was not adequately controlled. This could be related to differences in allosteric regulation or subcellular localisation, or might simply be a consequence of timing. Typically, lignin formation is tightly controlled at multiple levels so that monolignol biosynthesis, export, and polymerisation occur at the right time and in the right place as part of programmed cell death (Donaldson, 2001; Boerjan et al., 2003; Bollhöner et al., 2012; Behr et al., 2019). Analysis by 2D-HSQC NMR confirmed the shift in lignin composition towards G-lignin units and revealed a small decrease in H-lignin (Figure 3-6B). Despite the change in monomer composition, there was no significant difference in the proportion of p-hydroxybenzoate groups. These ester-linked pendent moieties occur naturally in the lignin of poplar, and their abundance can readily change in response to altered flux through the phenylpropanoid pathway (see Chapter 4 and Appendix 3). Further examination of the NMR spectra showed that there were also minor decreases in the proportions of ꞵ-aryl ether, resinol, and tetrahydrofuran linkages, as well as an increase in phenylcoumaran structures (Figure 3-7), likely due to the greater abundance of G-lignin units. 50 Figure 3-7. Additional NMR spectra of Pa×gCM1 RNAi poplar trees. Partial two-dimensional 1H–13C HSQC NMR spectra for enzyme-lignin samples of line 3 and the WT control, as labelled, showing the aliphatics region. The colour-coding and peak annotations for ꞵ-aryl ether (A, cyan), phenylcoumaran (B, green), resinol (C, magenta), and tetrahydrofuran units (C', magenta) are elaborated with the structures shown below. Relative levels of units are based on the sum of the identified units and is from uncorrected volume integrals. 51 Next, transversal stem cross-sections were examined by microscopy to ascertain whether there were changes in xylem development. Staining with phloroglucinol-HCl revealed marked differences in xylem morphology between line 3 and the WT control, namely an increase in the number of vessels and a decrease in the average vessel lumen area (Figure 3-8). Despite these differences in vessel density and lumen size, misshapen vessels were not observed, and the vascular system appeared to be properly formed and functional. Accordingly, stunting of line 3 is not likely a direct consequence of lignin perturbation but could instead be a sign of more far-reaching metabolic stresses. Figure 3-8. Lignin histology of Pa×gCM1 RNAi poplar trees. Transverse xylem cross-sections from line 3 (A) and WT (B) poplar stems, as labelled, stained with phloroglucinol-HCl. The black scale bars represent 100 µm. Histograms for line 3 (C) and WT (D) showing the distribution of cell lumen areas with fibres shown in blue and vessels in yellow. 52 Increased vessel density may partially help explain the elevated levels of guaiacyl units observed in line 3 xylem as poplar vessels are known to be enriched in G lignin (Musha & Goring, 1975; also see Chapter 6). However, a corresponding decrease in lignin-associated p-hydroxybenzoate would also have been expected as these groups are normally restricted to fibre cell walls (see Appendix 3). Therefore, the observation that p-hydroxybenzoate levels remained unchanged in line 3 could be further evidence of altered carbon flux through the phenylpropanoid pathway. 3.4.4 Systemic responses in plant metabolism and gene expression As chorismate is a key precursor to a wide range of metabolites in plants, we next examined whether the perturbations in lignin biosynthesis were accompanied by more widespread changes in metabolism. Gas chromatography coupled with mass spectrometry was used to quantify an array of metabolites including amino acids and soluble phenolics. The levels of all amino acids detected were significantly reduced in the developing xylem of all lines compared to WT with line 3 again being the most drastically altered (Figure 3-9A). Histidine, aspartate, and tryptophan were especially deficient in the xylem of line 3. Metabolic profiling of young leaves also showed reductions in various amino acids, with the exception of glycine which was more abundant in line 3 compared to WT (Figure 3-9C). Free amino acids are broadly involved in modulating stress responses, membrane permeability, and ion uptake (Barneix & Causin, 1996; Rai, 2002). The reduced levels in Pa×gCM1-suppressed poplar could be an indication that one or more of these processes have been affected. The abundance of many phenolic compounds was also altered in the developing xylem and young leaves of line 3 (Figure 3-9B, D). For example, syringin, a glucoside of sinapyl alcohol, was reduced below detection levels in xylem. This observation is consistent with the observed reduction in S-lignin units evidenced by thioacidolysis and NMR, and could be related to the increased proportion of G-lignin-rich vessel elements. Several salicylic acid derivatives were also drastically reduced, perhaps a result of perturbations in the pool of salicylic acid, which is also derived from chorismate. Interestingly, several of these compounds were actually more abundant in lines 1 and 2 wherein the compensatory expression of Pa×gCM2 and Pa×gCM3 was lower and there were no obvious impacts on growth and 53 Figure 3-9. Metabolite profiling of Pa×gCM1 RNAi poplar trees. A, B: Heat maps depicting the log of the fold change in the levels of 19 amino acids and 31 phenolic compounds in the developing xylem of lines 1–3 relative to the wild-type (WT) control, as labelled. C, D: Heat maps depicting the log of the fold change in the levels of 29 phenolic compounds and 16 amino acids in the young leaves of lines 1–3 relative to WT, as labelled. The colour gradient bar shows that red and orange correspond to negative values (i.e., a decrease relative to WT), whereas blue corresponds to positive values (i.e., an increase relative to WT). Grey squares labelled ND indicate that several metabolites were not detected in line 3. The black arrowheads denote that biological replicates of developing xylem were pooled for line 3 due to limited samples. Otherwise, five biological replicates were analysed for each line. 54 development. In the Salicaceae family, which includes poplar, salicinoid metabolites are known to occur naturally as defensive compounds (Boeckler et al., 2011). More broadly, free salicylic acid plays a role in regulating plant growth and development, photosynthesis, ion uptake and transport, and responses to biotic and abiotic stress (Vicente & Plasencia, 2011). Further analysis of the soluble phenolics by UHPLC confirmed that the developing xylem of line 3 poplars contained drastically reduced total amounts of salicylic acid (Figure 3-10). In addition to free amino acids and phenolics, metabolite profiling revealed widespread changes in the occurrence of various organic acids (Figure 3-11A, C). Several of these metabolites were actually decreased in lines 1 and 2 but elevated in line 3. Organic acids play an important role in various processes, but perhaps most notably they help maintain the redox balance of reactive oxygen species in plants that results from photosynthesis and cellular respiration (Igamberdiev & Bykova, 2018). Other metabolites including some sugars, sugar alcohols, and sterols were also decreased in the Pa×gCM1-suppressed lines (Figure 3-11B, D). These observations show that widespread changes in metabolic processes occurred as a result of Pa×gCM1 misregulation and may provide some additional clues as to how line 3 was so severely stunted and chlorotic. Figure 3-10. Total salicylic acid content is reduced in Pa×gCM1 RNAi poplar trees. UHPLC-DAD chromatograms of acid-hydrolysed methanolic extracts from the developing xylem of line 3, in orange, and from the wild-type (WT) control, in blue, along with a salicylic acid standard, in green. The DAD detector was set to 304 nm, the UV maximum of salicylic acid. 55 Figure 3-11. Additional metabolite profiling of Pa×gCM1 RNAi poplar trees. A, B: log of the fold change in the levels of 22 organic acids and 28 other miscellaneous metabolites in the developing xylem of lines 1–3 relative to the wild-type (WT) control, as labelled. C, D: Heat maps depicting the log of the fold change in the levels of 25 organic acids and 38 other miscellaneous metabolites in the young leaves of lines 1–3 relative to WT, as labelled. The colour gradient bar shows that red and orange correspond to negative values (i.e., a decrease relative to WT) whereas blue corresponds to positive values (i.e., an increase relative to WT). Grey squares labelled ND indicate that several metabolites were not detected in line 3. The black arrowheads denote that the biological replicates of developing xylem were pooled for line 3 due to limited samples. Otherwise, five biological replicates were analysed for each line. 56 Interestingly, many of the metabolite classes that were depleted in line 3 were reportedly elevated in the aerial tissues of Arabidopsis when an allosterically insensitive CM gene was overexpressed (Tzin et al., 2009). In order to further resolve the systemic responses in Pa×gCM1-suppressed poplar, RNA-seq experiments were performed to compare the gene expression profiles of the developing xylem in line 3 to the WT control. An enrichment analysis of GO terms revealed that many of the differentially expressed genes are involved in photosynthesis, ion binding, and redox processes (Figure 3-12). These results provide further evidence that suppression of Pa×gCM1 had widespread consequences beyond merely affecting the biosynthesis of lignin and phenylpropanoids. 3.4.5 Redundancy in the CM gene family Few studies have considered whether CM isoforms have distinct or overlapping roles in plants despite the importance of this gene family in both primary and secondary metabolism. Transcriptome databases for Arabidopsis and poplar show that AtCM2 and PtCM2, the cytosolic and allosterically insensitive isoforms, are constitutively expressed, whereas the other isoforms have partially overlapping expression profiles in various organs and tissues (Waese et al., 2017; Sundell et al., 2015; Sundell et al., 2017; Figure 3-13). The observation that RNAi-mediated suppression of Pa×gCM1 did not result in decreased lignin biosynthesis suggests that at least partial redundancy occurs within the poplar CM gene family. As an alternative to this hypothesis, it is also possible that other CM homologues are differentially expressed as a stress response. Interestingly, both the unregulated and regulated isoforms (Pa×gCM2 and Pa×gCM3) showed elevated expression in response to the suppression of Pa×gCM1. At present, it remains unclear whether such apparent functional redundancy extends beyond the metabolic requirements for chorismate to supply monolignol biosynthesis in developing xylem tissues. It would be a worthwhile future experiment to specifically knock down the other two CM isoforms in poplar to see if Pa×gCM1 is upregulated and to ascertain whether compensatory expression can be reciprocal. It is also unknown how broadly CM gene redundancy might occur across different plant taxa. In petunia, for example, RNAi-mediated suppression of CM1 was accompanied by a reduction in the biosynthesis of volatile phenolic compounds in flowers, but was not associated with increased expression of any other 57 Figure 3-12. RNA-seq analysis of Pa×gCM1 RNAi poplar trees. A: Enrichment of gene ontology (GO) terms for differentially expressed (DE) genes that were upregulated in the developing xylem of line 3 relative to the wild-type (WT) control. B: Enrichment of GO terms for DE genes that were downregulated in the developing xylem of line 3 relative to WT. The colours correspond to p-values, as shown in the legend. Three biological replicates were analysed for each. 58 Figure 3-13. Expression profiles of poplar chorismate mutase homologues. Gene expression for PtCM1 (A), PtCM2 (B), and PtCM3 (C) from the online databases PopGenIE (Sundell et al., 2015) and AspWood (Sundell et al., 2017) showing various tissues of poplar as labelled (expanding buds, flowers, seeds, wood, suckers, roots, and various leaves) on the left, and throughout differentiating phloem and xylem on the right. 59 isoforms (Colquhoun et al., 2010), suggesting that CM redundancy may not be universal. Thus, further mutant and knockout studies in diverse plant species will be needed to resolve this question. As the phylogenetic analysis showed that some taxa evidently contain as many as five CM isoforms, these may be the most informative species for future studies. The severe stunting of line 3 was not merely a consequence of changes in lignin biosynthesis or phenylpropanoid metabolism. Instead, it appears that allocation of carbon towards lignin biosynthesis via Pa×gCM2 and Pa×gCM3 may have led to perturbations in the pool of salicylic acid. A severe dwarfing phenotype was reported when salicylic acid hyperaccumulated in transgenic Arabidopsis (Mauch et al., 2001). On the other hand, salicylic acid accumulation led to oxidative stress but not stunting in poplar (Xue et al., 2013). Additional studies have illustrated that metabolic connections exist between CM activity and salicylic acid levels. For example, CM was downregulated in tobacco plants when salicylic acid levels were constitutively elevated (Nugroho et al., 2002). Incidentally, various fungal pathogens and parasitic nematodes have exploited this connection by secreting a CM enzyme that compromises plant defence systems by altering salicylic acid content in host tissues (Lambert et al., 1999; Djamei et al., 2011). Clearly, CM occupies a critically important metabolic position in plants. In summary, this work highlights the central role of chorismate as a key precursor to a wide range of important plant metabolites. Compensatory expression of apparently redundant CM isoforms in Pa×gCM1-misregulated hybrid poplar was accompanied by widespread changes in gene expression and metabolism which affected photosynthesis, ion transport, and redox processes, as well as plant growth and xylem development. These systemic responses may have been the result of perturbations in salicylic acid biosynthesis, as this too derives from chorismate. 60 Chapter 4: Metabolic engineering of p-hydroxybenzoate pendent groups in poplar lignin 4.1 Summary Ester-linked p-hydroxybenzoate occurs naturally in poplar lignin as pendent groups that can be released by mild alkaline hydrolysis. These ‘clip-off’ phenolics can be separated from biomass and upgraded into diverse high-value bioproducts. We introduced a bacterial chorismate pyruvate lyase gene into transgenic poplar trees with the aim of producing more p-hydroxybenzoate from chorismate, itself a metabolic precursor to lignin. By driving heterologous expression specifically in the plastids of cells undergoing secondary wall formation, this strategy achieved a 50% increase in cell-wall-bound p-hydroxybenzoate in mature wood and nearly 10 times more in developing xylem relative to control trees. Comparable amounts also remained as soluble p-hydroxybenzoate-containing metabolites, pointing to even greater engineering potential. Mass spectrometry imaging showed that the elevated p-hydroxybenzoylation was largely restricted to the cell walls of fibres. Finally, transgenic lines outperformed control trees in assays of saccharification potential. This study highlights the potential of augmenting cell-wall-bound phenolate esters and shows the importance of substrate supply in lignin engineering. 4.2 Introduction Lignin is a complex phenolic biopolymer found primarily in the secondary cell walls of vascular plants. The development of lignin was a milestone in plant evolution and the lignified plant vasculature is a defining feature of tracheophytes (Weng & Chapple, 2010). Lignin is a critical component of water conduction and plant defence systems, and it contributes greatly to the compressive strength of stems and branches (Dixon & Barros, 2019). Although the chemical complexity and recalcitrance of lignin presents a major challenge for industrial biomass processing, it also paves the way for diverse biochemicals and bioproducts to be made from lignin (Rinaldi et al., 2016). To that end, there is considerable interest in developing biomass feedstocks with engineered lignin polymers (see Chapter 2). Lignin is assembled primarily from three monolignol precursors – p-coumaryl, coniferyl, and sinapyl alcohol (Figure 4-1; Freudenberg & Neish, 1968). Once incorporated into a lignin polymer, these become 61 Figure 4-1. Lignin engineering with bacterial CPL. Metabolic map depicting the shikimate and general phenylpropanoid pathways in the plastid and cytosol, as well as the export of lignin monomers and polymerisation in poplar cell walls. The novel route from chorismate catalysed by bacterial chorismate pyruvate lyase (CPL) that leads to the incorporation of more p-hydroxybenzoate pendent groups in lignin is shown in blue. p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) lignin units, respectively. The monolignols are synthesised in the cytosol prior to export into the apoplast where laccase and peroxidase enzymes generate monolignol radicals that undergo coupling and cross-coupling reactions (Ralph et al., 2004a). This process of dehydrogenative polymerisation proceeds in a combinatorial manner such that the structure of the resulting polymer reflects the complement of monomers delivered to the site of lignification. Consequently, a wide variety of lignin composition and structure exists in Nature. The flexibility of lignification is perhaps best exemplified by loss/gain-of-function experiments. For example, downregulation of p-coumaroyl-CoA 3'-hydroxylase (C3'H) in transgenic poplar led to dramatically increased levels of H units (Coleman et al., 2008b), whereas overexpression of ferulate 5-hydroxylase 62 (F5H) resulted in a polymer built predominantly with S units (Huntley et al., 2003; Stewart et al., 2009). The inherent plasticity of lignification extends beyond the three primary monolignols, as demonstrated by the growing number of non-canonical monomers that have been reported (Ralph et al., 2019a). Among these, the γ-linked monolignol conjugates are the most prevalent and best studied. For example, the lignin found in kenaf bast fibres is naturally acetylated as a result of the incorporation of pre-acylated monolignols (Ralph, 1996; Lu & Ralph, 2008). Similarly, the lignin of commelinid monocots is decorated with p-coumarate groups derived from monolignol–p-coumarate conjugates produced via p-coumaroyl-CoA:monolignol transferases, members of the BAHD superfamily of acyltransferases (Ralph et al., 1994; Hatfield et al., 2008a; Withers et al., 2012). Analogously in poplars and willows, p-hydroxybenzoate groups (henceforth denoted as pHB for the ester-linked pendent form, and pHBA for the free acid) comprise up to 10% of the lignin (Smith, 1955b; also see Appendix 3). These too derive from monolignol conjugates (Morreel et al., 2004), but no p-hydroxybenzoyl-CoA:monolignol transferases have yet been reported. Cell-wall-bound pHB has also been detected throughout the family Arecaceae (palms), in Japanese spikenard (Aralia cordata), in the roots of carrot (Daucus carota), in the stems of purple mountain saxifrage (Saxifraga oppositifolia), and at especially high levels in the Mediterranean seagrass Posidonia oceanica (Pearl et al., 1959; Hibino et al., 1994; Parr et al., 1997; Faleva et al., 2020; Rencoret et al., 2020). As pHB groups are ester-linked to lignin polymers, they can be released by mild alkaline hydrolysis. Once separated from the biomass, these ‘clip-off’ phenolics may be used directly as platform chemicals or upgraded into a variety of other biochemicals and bioproducts (Rinaldi et al., 2016; Wang et al., 2018b; Becker & Whittmann, 2019). For example, derivatives of pHBA are used in cosmetics (as paraben preservatives), in polyester plastics (as liquid crystal copolymers), and as precursors to a wide range of pharmaceutical compounds (Aalto et al., 1953; Jackson & Kuhfuss, 1976; Manuja et al., 2013). Alternatively, engineered microbes have been developed that catabolise and funnel phenolics including pHBA into metabolic pathways leading to high-value compounds (Beckham et al., 2016; Kamimura et al., 2017). However, to economise the use of clip-offs for such applications, the innate levels in feedstock plants must be substantially improved through cell wall engineering (Karlen et al., 2020). 63 Herein, we report a novel strategy to increase the levels of pHB groups by bolstering the supply of pHBA precursors. Although it has been established that pHBA derives from phenylpropanoid biosynthesis (Figure 4-1; El-Basyouni et al., 1964; Zenk & Müller, 1964), the endogenous pathway has not been fully elucidated and remains an inaccessible engineering target. Fortuitously, an alternate route has evolved in bacteria: chorismate pyruvate lyase (CPL) cleaves chorismate, a product of the shikimate pathway, to produce pHBA and pyruvate (Siebert et al., 1994). Previous studies on the heterologous expression of this enzyme in planta have reported hyperaccumulation of soluble pHBA-containing metabolites (Siebert et al., 1996; Li et al., 1997). As poplar can further deploy pHBA to the cell wall, we asked whether expression of bacterial CPL would lead to increased p-hydroxybenzoylation of poplar lignin. 4.3 Materials and Methods 4.3.1 Gene cloning and expression constructs The ubiC gene encoding CPL was cloned from Escherichia coli K12 (Figure 4-2; Nichols and Green, 1992; Siebert et al., 1992). A simplified phenol–chloroform DNA purification protocol was followed (Chachaty & Saulnier, 2000) using an overnight culture grown in liquid LB media. The full-length coding sequence of the ubiC gene was amplified by PCR with primers designed using the published bacterial genome (Blattner et al., 1997; Table 4-1). A synthetic plastid localisation signal sequence was used which includes the chimeric transit signals from the RuBisCO genes of sunflower and maize with an intervening spacer sequence, and which has previously been successfully used for chloroplast targeting (Eudes et al., 2012). Standard PCR and overlap extension PCR techniques were used to amplify the plastid localisation signal (381 bp) followed by the ubiC coding sequence (498 bp) separately at first, and then together as a fusion product with attB adaptor sequences added to the 5' and 3' ends to enable Gateway-mediated cloning (Life Technologies, Carlsbad, CA, U.S.A.). This fragment was then transferred into the pDONR/Zeo plasmid using a BP Clonase reaction and subsequently, using an LR Clonase reaction, into a modified version of the plant expression vector pH7WG2 (Karimi et al., 2002) in which the constitutive promoter sequence was replaced with a cellulose synthase promoter from Populus trichocarpa for 64 >synthetic chloroplast transit signal sequence ATGGCTTCGATCTCCTCCTCAGTCGCGACCGTTAGCCGGACCGCCCCTGCTCAGGCCAACATGGTGGCTCCGTTCACCGGCCTTAAGTCCAACGCCGCCTTCCCCACCACCAAGAAGGCTAACGACTTCTCCACCCTTCCCAGCAACGGTGGAAGAGTTCAATGCATGCAGGTGTGGCCGGCCTACGGCAACAAGAAGTTCGAGACGCTGTCGTACCTGCCGCCGCTGTCGACGATGGCGCCCACCGTGATGATGGCCTCGTCGGCCACCGCCGTCGCTCCGTTCCAGGGGCTCAAGTCCACCGCCAGCCTCCCCGTCGCCCGCCGCTCCTCCAGAAGCCTCGGCAACGTCAGCAACGGCGGAAGGATCCGGTGCATGCAG >ubiC cds from Escherichia coli K12 ATGTCACACCCCGCGTTAACGCAACTGCGTGCGCTGCGCTATTGTAAAGAGATCCCTGCCCTGGATCCGCAACTGCTCGACTGGCTGTTGCTGGAGGATTCCATGACAAAACGTTTTGAACAGCAGGGAAAAACGGTAAGCGTGACGATGATCCGCGAAGGGTTTGTCGAGCAGAATGAAATCCCCGAAGAACTGCCGCTGCTGCCGAAAGAGTCTCGTTACTGGTTACGTGAAATTTTGTTATGTGCCGATGGTGAACCGTGGCTTGCCGGTCGTACCGTCGTTCCTGTGTCAACGTTAAGCGGGCCGGAGCTGGCGTTACAAAAATTGGGTAAAACGCCGTTAGGACGCTATCTGTTCACATCATCGACATTAACCCGGGACTTTATTGAGATAGGCCGTGATGCCGGGCTGTGGGGGCGACGTTCCCGCCTGCGATTAAGCGGTAAACCGCTGTTGCTAACAGAACTGTTTTTACCGGCGTCACCGTTGTACTAA Figure 4-2. DNA sequences for heterologous expression of bacterial CPL. The synthetic chloroplast transit signal and the ubiC gene encoding bacterial chorismate pyruvate lyase from Escherichia coli. spatiotemporally-controlled expression. This modified plasmid was prepared with restriction cloning by excising the 35S promoter from pH7WG2 using the SacI and SbfI sites through partial digestion and then ligating a PCR-amplified CesA8 promoter sequence (1985 bp) from poplar (see Wilkerson et al., 2014) with adaptors containing the appropriate restriction enzyme recognition sequences. Standard molecular biology techniques were followed as described in Sambrook & Russell (2006). All PCR reactions were performed using recombinant Taq DNA polymerase or Platinum Pfx proofreading DNA polymerase (Thermo Scientific) and all PCR products were confirmed using agarose gel electrophoresis and DNA sequencing. Sub-cloning efficiency E. coli DH5α (Life Technologies) was used for transformations and cultures were grown in a shaking water bath incubator at 37 °C and 200 rpm. 4.3.2 Establishment of transgenic hybrid poplars The CPL expression construct was transferred to Agrobacterium tumefaciens EHA105 using a freeze-thaw transformation protocol (Wise et al., 2006). A single colony, which was confirmed by DNA sequencing, was then used for Agrobacterium-mediated transformation of hybrid poplar, Populus alba × grandidentata; P39. Wild-type (WT) poplar trees were maintained on woody plant media (WPM; Lloyd & McCown, 1981) with 10 µg L-1 1-naphthaleneacetic acid and 1 mL L-1 Plant Preservative Mixture (Plant 65 Table 4-1. Primers and PCR programmes for Chapter 4. Details are provided for all PCR reactions used in construct assembly, transformant screening, and RT-qPCR. Description: Primer pair (5' → 3'): PCR reaction conditions: Amplification of the synthetic chloroplast transit signal and the ubiC gene For transit signal (381 bp), Forward primer: ATG GCT TCG ATC TCC TCC TC Reverse primer: CTG CAT GCA CCG GAT CC For ubiC (498 bp), Forward primer: ATG TCA CAC CCC GCG TTA AC Reverse primer: TTA GTA CAA CGG TGA CGC C 2.5 µL ................................. 10× buffer 0.5 µL ......................... 50 mM MgSO4 0.5 µL .............. 10 µM forward primer 0.5 µL ................10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ......................... template DNA* 0.2 µL ................ Pfx DNA polymerase Up to 25 µL ................ deionised water With thermal cycling as follows: 94 °C for 30 s, 34 cycles of (94 °C for 15 s, 60 °C for 30 s, 68 °C for 90 s), and then 68 °C for 10 min. * Note: template DNA was a purified plasmid in the case of the transit signal, and a sample of genomic DNA from E. coli K12 for the ubiC gene. Construct assembly (Step 1: Addition of overlap extensions) For transit signal, Forward primer: ATG GCT TCG ATC TCC TCC TC Reverse primer: GTT AAC GCG GGG TGT GAC ATC TGC ATG CAC CGG ATC C For ubiC, Forward primer: GGA TCC GGT GCA TGC AGA TGT CAC ACC CCG CGT TAA C Reverse primer: TTA GTA CAA CGG TGA CGC C 2.5 µL ................................. 10× buffer 0.5 µL ......................... 50 mM MgSO4 0.5 µL .............. 10 µM forward primer 0.5 µL ................10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ......................... template DNA* 0.2 µL ................ Pfx DNA polymerase Up to 25 µL ................ deionised water With thermal cycling as follows: 94 °C for 30 s, 34 cycles of (94 °C for 15 s, 60 °C for 30 s, 68 °C for 90 s), and then 68 °C for 10 min. * Note: template DNA was a plasmid containing either the transit signal or the ubiC sequences as cloned above. Construct assembly (Step 2: Amplification of a chimeric product with attB sequences to enable Gateway cloning) Forward primer: GGG GAC AAG TTT GTA CAA AAA AGC AGG CTA TGG CTT CGA TCT CCT CCT C Reverse primer: GGG GAC CAC TTT GTA CAA GAA AGC TGG GTT TAG TAC AAC GGT GAC Chimeric product size: 936 bp 2.5 µL ................................. 10× buffer 0.5 µL ......................... 50 mM MgSO4 0.5 µL .............. 10 µM forward primer 0.5 µL ................10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ................ template with signal* 0.5 µL .................. template with ubiC* 0.2 µL ................ Pfx DNA polymerase Up to 25 µL ................ deionised water With thermal cycling as follows: 94 °C for 30 s, 34 cycles of (94 °C for 15 s, 60 °C for 30 s, 68 °C for 90 s), and then 68 °C for 10 min. * Note: template DNA was purified PCR products obtained in Step 1 above. Note: Table continues on the following page. 66 Note: Table continued from the preceding page. Description: Primer pair (5' → 3'): PCR reaction conditions: PCR-screening to identify positive transformants Forward primer*: GTA CCG TCG TTC CTG TGT C Reverse primer: TTA GTA CAA CGG TGA CGC C Amplicon size: 224 bp 2.5 µL ................................. 10× buffer 0.5 µL ......................... 50 mM MgSO4 0.5 µL .............. 10 µM forward primer 0.5 µL ................10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ........................... template DNA 0.5 µL ............... Taq DNA polymerase Up to 25 µL ................ deionised water With thermal cycling as follows: 94 °C for 3 min, 34 cycles of (94 °C for 30 s, 55 °C for 30 s, 72 °C for 30 s), and then 72 °C for 5 min. * Note: template DNA was genomic DNA preparations from young poplar leaves. Quantification of expression levels using RT-qPCR For the signal–ubiC sequence, Forward primer*: CCA GGG GCT CAA GTC CAC Reverse primer: CAT GCA CCG GAT CCT TCC Amplicon size: 94 bp Priming efficiency: 99.5% For elongation factor 1β (reference), Forward primer: GGC ATT AAG TTT TGT CGG TCT G Reverse primer: GCG GTT CAT CAT TTC ATC TGG Amplicon size: 97 bp Priming efficiency: 99.9% For actin (reference), Forward primer: ACC AGT GTG TCT TGG TCT ACC C Reverse primer: CGA TGC CGA GGA TAT TCA AC Amplicon size: 127 bp Priming efficiency: 99.2% 5 µL .... BrightGreen qPCR master mix 0.3 µL .............. 10 µM forward primer 0.3 µL ................10 µM reverse primer 1 µL ............................ template cDNA Up to 10 µL ................ deionised water With qPCR parameters as follows: 95 °C for 30 s, 39 cycles of (95 °C for 5 s, 60 °C for 15 s, plate read for fluorescence), and then followed by a melt-curve analysis as follows: 95 °C for 10 s, ramp from 55 °C to 95 °C at 0.5 °C per increment with hold times of 5 s and a plate read for fluorescence. * Note: template DNA was cDNA preparations synthesised with RNA extracted from developing poplar xylem. 67 Cell Technology, Washington, DC, U.S.A.) in sterile Magenta GA-7 containers at room temperature with 16 h daylight under a photon flux of 100–200 µmol m-2 s-1. On Day 1 of the transformation procedure, a 5-mL aliquot of liquid YEP media was inoculated with the A. tumefaciens clone and incubated at 30ºC overnight with shaking. On Day 2, small discs were cut from the leaves of WT poplar trees using a hand-held paper hole punch and left floating in liquid WPM media overnight. Concomitantly on Day 2, the overnight culture of A. tumefaciens was used to inoculate 50 mL of liquid WPM containing 100 µM acetosyringone which was incubated for a second overnight period at 30ºC. On Day 3, the leaf discs were transferred into the A. tumefaciens culture and, after 30 min of vigorous mixing, the discs were placed abaxial-side-up onto co-cultivation media (WPM with 0.1 µM 1-naphthaleneacetic acid, 0.1 µM 6-benzylaminopurine, and 0.1 µM thidiazuron) and incubated at room temperature in darkness. On Day 6, the leaf discs were transferred to decontamination media (WPM with 0.1 µM 1-naphthaleneacetic acid, 0.1 µM 6-benzylaminopurine, 0.1 µM thidiazuron, 500 µg/mL carbenicillin and 250 µg/mL cefotaxime) to kill the A. tumefaciens, and then again maintained in darkness. On Day 9, the leaf discs were moved to regeneration media (WPM with 0.1 µM 1-naphthaleneacetic acid, 0.1 µM 6-benzylaminopurine, 0.1 µM thidiazuron, 500 µg mL-1 carbenicillin, 250 µg mL-1 cefotaxime, and 25 µg mL-1 hygromycin) and placed under subdued lighting. Once small shoots appeared after 4–8 weeks, healthy leaf discs were transferred to shooting media (WPM with 0.01 µM 6-benzylaminopurine, 500 µg mL-1 carbenicillin, 250 µg mL-1 cefotaximen and 25 µg mL-1 hygromycin). After another 2–4 weeks, the elongated shoots were transferred to rooting media (WPM with 0.01 µM 1-naphthaleneacetic acid, 500 µg mL-1 carbenicillin, 250 µg mL-1 cefotaxime, and 25 µg mL-1 hygromycin) in Magenta GA-7 containers under normal lighting. Finally, once shoots had successfully rooted on this selection media, excised stem pieces were transferred to maintenance media without antibiotics. WT trees used as controls were also regenerated from leaf discs following a parallel procedure without the use of antibiotics for selection. Aseptic techniques were used for all tissue culture manipulations. Semi-quantitative RT-PCR was used to screen and select transgenic lines harbouring the ubiC expression construct. For this, RNA was prepared from leaf tissue using the Trizol reagent (Life Technologies). The RNA was treated with DNase using the Turbo DNA-free kit (Life Technologies) and then used to 68 synthesise first-stand cDNA using the OneScript Plus cDNA synthesis kit (Applied Biological Materials Inc., Richmond, BC, Canada). PCR reactions with primers designed to anneal within the ubiC gene were used to screen for transgenic plants and select the most promising lines for further study (Table 4-1). Primers specific to the poplar elongation factor 1β gene were used as a reference for semi-quantitative comparisons. Prior to planting in soil, all trees were propagated once more from apical stem pieces to ensure even initial growth. Out of 25 independent transformation events, the five best lines were chosen for further study and five biological replicates of each line were planted in a peat-based soil in the UBC horticultural greenhouse. Natural light was supplemented with high-pressure sodium lamps (minimum photon flux of 500 µmol m-2s-1) and the trees were irrigated on flooding tables as needed with water containing Hoagland’s solution (Hoagland & Arnon, 1938). Immediately after planting and for the subsequent two weeks, trees were covered with transparent plastic cups to maintain high humidity levels during acclimation. Regular inspections were performed for any signs of pests or pathogens, and the stems were supported by staking with bamboo poles. After 18 weeks in the greenhouse, the heights of the trees were measured from the apex to the root collar, and the stem diameters were measured at 10 cm above the root collar. The leaves and bark were removed from the stems and the developing xylem was scraped from the debarked stems using a sterile razor blade. The developing xylem tissue was then flash-frozen in liquid nitrogen, ground in a mortar and pestle to a fine powder, and stored at -80 °C until further use. After air drying the stems, a 10 cm-long piece was removed from the bottom of each tree and cut into small pieces, the pith was removed, and the wood was disintegrated using a Wiley mill to pass a 40-mesh sieve for further analysis. 4.3.3 Expression of the ubiC transgene RT-qPCR was used to assess the relative expression levels of the CPL expression construct in transgenic poplar xylem. RNA was prepared using a CTAB-based workflow (Kolosova et al., 2004) with developing xylem tissue which had been ground in a mortar and pestle with liquid nitrogen and stored at -80 °C. The RNA was treated with DNase and then used to synthesise first-stand cDNA as described above. After 69 diluting the cDNA eight-fold, qPCR reactions were prepared using BrightGreen qPCR Master Mix (Applied Biological Materials Inc.) with gene-specific primers that had been tested over a range of template concentrations and found to be highly specific and have good priming efficiency (Table 4-1). Manufacturers’ instructions were followed for all kits and reagents. Thermal cycling, fluorescence detection, melt-curve analysis, and quantification was performed using a CFX 96 Real-Time PCR instrument and the corresponding CFX Manager software (Bio-Rad Laboratories Inc., Hercules, CA, U.S.A.). Primers specific to the poplar elongation factor 1β and actin genes were used as reference genes for the calculation of relative expression. 4.3.4 Lignin content and composition The Klason lignin method was used to evaluate lignin content as the sum of acid-insoluble and acid-soluble fractions. Wood powder which had been processed in a Wiley mill to pass a 40-mesh sieve was exhaustively Soxhlet-extracted for 24 h using hot acetone. The extractive-free wood powder was dried overnight at 50 °C and subsamples of 200 mg were accurately weighed into 125-mL glass serum bottles. Following the addition of 3 mL of cold 72% sulphuric acid, each sample was stirred with a glass rod every 10 min for 2 h prior to the addition of 112 mL of water. Once capped and sealed, the bottles were autoclaved for 1 h at 121 °C and 15 psi. Pre-weighed and oven-dried medium-coarseness sintered glass filtering crucibles were used to collect the remaining residues which were then rinsed with 150 mL of water and dried overnight at 105 °C before the acid-insoluble lignin was determined gravimetrically. The acid-soluble lignin component was evaluated by UV spectrophotometry of acid-hydrolysis filtrates using the Beer-Lambert law with an extinction coefficient of 110 L g-1 cm-1 at 205 nm. The lignin composition was evaluated by quantifying lignin monomers released with thioacidolysis (Robinson & Mansfield, 2009). Briefly, 10-mg samples of extractive-free wood powder were subjected to thioacidolysis using 1 mL of a freshly distilled dioxane containing 2.5% boron trifluoride etherate and 10% ethanethiol for 4 h at 100 °C with periodic mixing. The reactions were then incubated at -20 °C for 5 min, and quenched with 0.3 mL of 0.4 M sodium bicarbonate. As an internal standard, 0.2 mL of 5 mg mL-1 tetracosane was added. The reaction products were collected following phase separation using 1 mL 70 dichloromethane and 2 mL deionised water. The organic phase was then dried by passing through a loosely packed column of anhydrous sodium sulphate, and the solvent was evaporated using a vacuum centrifuge. Samples were resuspended in 0.7 mL dichloromethane and derivatised by adding 20 μL pyridine and 100 μL N,O-bis(trimethylsilyl) acetamide to a 20-μL volume of sample. Derivatised lignin monomers were then separated using a Trace 1310 GC apparatus (Thermo Scientific, Waltham, MA, U.S.A.) equipped with a TraceGOLD 5MS column (30 m × 0.32 mm, 0.25 µm thickness) and a flame-ionisation detector. Helium was used as the carrier gas with a constant-velocity flow rate of 1 mL min-1, the injection port was heated to 250 °C, and the column oven temperature was controlled as follows: hold at 130 °C for 3 min, then ramp at 6.5 °C min-1 to 230 °C and hold for 6 min, and then finally ramp at 2.5 °C min-1 to 250 °C and hold for 10 min. Peaks corresponding to the main diastereoisomers of trithioethylated G and S-lignin monomers were identified with reference to synthetic standards and were integrated using the Chromeleon 7 software package (Thermo Scientific). Finally, the peak areas were corrected with previously determined response factors and then used to calculate the monomer ratio. 4.3.5 NMR analysis Two-dimensional 1H–13C HSQC NMR spectra were collected using enzyme-lignin samples prepared by enzymatic digestion of ball-milled wood tissue (Kim & Ralph, 2010). Wood powder which had been processed in a Wiley mill to pass a 40-mesh sieve was subjected to sequential rounds of solvent extraction in an ultrasonic water bath as follows: 3 × 20 min with water, 3 × 20 min with 80% ethanol, 3 × 20 min with acetone. After drying in an oven at 50 °C, extractive-free wood powder was processed in a Fritsch Pulverisette 7 planetary ball mill using 20-mL agate jars containing 600 mg of wood powder and ten 10-mm agate balls at 600 rpm with 30 cycles consisting of 10 min grinding and 5 min rest. After ball-milling, the wood powder was quantitatively transferred to 50-mL Falcon tubes using 45 mL of sodium acetate buffer (pH 5.0) and treated with 30 mg of Cellulysin cellulase (from Trichoderma viride, Calbiochem, MilliporeSigma, Burlington, MA, U.S.A.). After three days of incubation at 37 °C on a rotary shaker, the powder was rinsed twice with acetate buffer and fresh enzyme was added. After another three days of incubation, the powder was rinsed twice with distilled water and then freeze-dried. 71 Approximately 10 mg of enzyme-lignin powder was dissolved in 0.5 mL of 4:1 dimethyl sulphoxide-d6:pyridine-d5 in a 5-mm outer diameter NMR tube. Spectra were collected using a standard adiabatic-pulse programme (hsqcetgpsisp2.2) on a Biospin AVANCE 700 MHz spectrometer (Bruker Corp., Billerica, MA, U.S.A.) equipped with a 5-mm QCI 1H/31P/13C/15N cryoprobe with the proton coils closest to the sample. The dimethyl-sulphoxide peak at δH 2.5 ppm and δC 39.5 ppm was used for calibration, and volume integrations for the peaks of interest were performed using TopSpin 4.08 (Bruker Corp.). Proportions were expressed on a basis where % pHB = ½ pHB2/6 / 100% aromatics, 100% aromatics = ½ (H2/6) + G2 + ½ (S2/6 + S'2/6), and 100% aliphatics = Aα + Bα + Cα + C'α. Peak annotations followed from previous reports (Kim & Ralph, 2010). 4.3.6 Metabolites and cell-wall-bound p-hydroxybenzoate Developing xylem powder that had been ground with a mortar and pestle and stored at -80 °C was freeze-dried and used for the quantification of soluble and cell-wall-bound pHBA. 10-mg portions of wood powder were weighed into 2-mL screw-cap vials and 950 μL of 80% methanol was added along with 50 μL of 1 mg mL-1 o-anisic acid as an internal standard. Once firmly sealed, the vials were incubated for 3 h at 50 °C in a hybridiser-incubator rotating at 40 rpm. After cooling for 5 min at -20 °C, the phenolic extracts were centrifuged for 5 min at 16,000 g and filtered using 0.45-μm nylon syringe filters. The remaining wood powder was set aside for further analysis of cell-wall-bound pHB, whereas 200-µL aliquots of the methanolic extracts were used for the analysis of free pHBA. Saponification was then used to cleave ester-linked conjugates of pHBA in the methanolic extracts. Aliquots (200 µL) of the extracts were transferred into new vials, 10 µL of 0.2 M NaOH was added to reduce the volatility by converting to sodium salts, and the methanol was evaporated using a vacuum centrifuge. To the dried residue, 300 µL of 2 M NaOH was added and the vials were incubated for 24 h at 30 °C in a hybridiser-incubator. Aliquots (180 µL) of the saponified extracts were used for quantification of ester-linked pHBA metabolites after acidifying with 18 µL of 72% H2SO4. To cleave ether-linked conjugates, the saponified methanolic extracts were subjected to acid hydrolysis. Aliquots (120 µL) of the saponified extracts were transferred into new vials and 120 µL of 72 3 M HCl was added. The vials were incubated at 95 °C for 3 h with mixing every 15 min, then cooled at -20 °C for 5 min and used for the quantification of ether-linked pHBA metabolites. The wood powder remaining after extraction with methanol was used for the quantification of cell-wall-bound pHB in developing xylem. To ensure that all soluble metabolites were removed, three additional extractions were performed with 1 mL acetone each for 24 h in a hybridiser-incubator at 30 °C. Residual acetone was removed by evaporation in a vacuum centrifuge and the dried wood powder was accurately weighed into fresh vials. To release cell-wall-bound pHB groups, 500 µL of 2 M NaOH was added which contained 50 µg mL-1 of 3,4,5-trimethoxycinnamic acid as an internal standard. After incubating for 24 h at 30 °C in a hybridiser-incubator, the saponification reactions were acidified with 50 µL of 72% H2SO4, and the remaining wood powder was removed by centrifugation at 16,000 g for 5 min. The amount of cell-wall-bound pHB in mature xylem was determined in the same manner except that acetone-extracted 40-mesh wood powder was used directly. Quantification of pHBA was performed using an Agilent Infinity II 1290 UHPLC apparatus (Agilent Technologies Inc., Santa Clara, CA, U.S.A.) equipped with a Zorbax Eclipse Plus C-18 column (2.1 × 50 mm, 1.8 µm particle size, Agilent) and a diode array detector. Good peak separation was achieved using an injection volume of 1 µL, a flow rate of 0.5 mL min-1, and a binary gradient of 5% to 25% of acetonitrile containing 0.01% trifluoroacetic acid (eluent A) and water containing 0.01% trifluoroacetic acid (eluent B) over 6 min. Integration of pHBA, o-anisic acid and 3,4,5-trimethoxycinnamic acid peaks was performed at the UV maxima of 255, 296, and 304 nm, respectively. Subtraction of the amount of free pHBA in methanolic extracts from the total following saponification yielded the amount of alkali-labile pHBA. Similarly, subtraction of the amounts of free and alkali-labile pHBA from the total following acid hydrolysis yielded the amount of pHBA released by alkali and acid. A five-point calibration curve prepared with authentic standards enabled quantification. Identification of metabolites was performed by high-resolution mass spectrometry (HRMS) using a Nexara X2 UHPLC apparatus (Shimadzu Corp., Kyoto, Japan) equipped with a diode array detector and an Impact II Ultra-High Resolution Qq-Time-of-Flight mass spectrometer (Bruker Corp.). Methanolic extracts were prepared using freeze-dried samples of developing xylem as described above and then 73 analysed using two different chromatographic conditions. First, 10-µL samples were injected onto a Kinetex XB-C18 column (1.7 µm × 100 mm × 2.1 mm, 100 Å, Phenomenex, Torrance, CA, U.S.A.). The mobile phase was a binary gradient of 0.1% formic acid in water (eluent A) and acetonitrile (eluent B) with a flow rate of 0.6 mL min-1. The gradient programme was applied over 26 min as follows: 1% B for 1.5 min, then ramp to 20% B at 10 min, continue ramping to 100% B at 17 min, hold until 20 min, then return to 1% B at 21 min and hold for 5 min to recondition the column. UV detection was performed from 250−600 nm. HRMS chromatograms were acquired from m/z 50−1000 with the electrospray-ionisation source operating in negative-ion mode. The MS configuration was as follows: capillary voltage, +3.5 kV; nebulising gas, nitrogen at 4 bar; temperature, 210 °C; drying gas, nitrogen at 8 L min-1; collision energy, 20 eV. The HRMS chromatograms were analysed using MetaboScape 4.0 (Bruker Corp.). Calibration of the MS detector was performed independently for each sample using 10 mM sodium formate in 50:50 isopropanol/water, which eluted between 0.7–0.72 min. The bucket table selection criteria were as follows: intensity threshold, 1000 counts; minimum peak length, 5 spectra; time range, 1.2–17 min; mass range, m/z 50−1000. The peaks were validated by assessing the extracted ion chromatograms, the MSMS fragments, and the average mass spectra over the elution peak. Chemical formulae were calculated using the SmartFormula algorithm for compounds containing CHONa with a scoring tolerance of mSigma <50. Putative metabolite identities were deduced from these chemical formulae using CompoundCrawler to search the PubChem and Chemical Entities of Biological Interest (ChEBI) databases. Finally, the putative identities were further scrutinised using MetFrag to examine the predicted fragment ions. To separate the glucosides of pHBA, 5-µL samples were injected onto a Luna Omega SUGAR column (3 × µm × 100 mm × 4.6 mm, 100 Å, Phenomenex, Torrance, CA, U.S.A.). The mobile phase was a low-pressure quaternary gradient consisting of 0.1% formic acid in water (eluent A), 0.2% ammonium hydroxide in water (eluent B), methanol (eluent C), and 0.1% formic acid in acetonitrile (eluent D), at a flow rate of 0.56 mL min-1. The gradient programme was applied over 30 min as follows: 99:0:0:1 A/B/C/D for 1 min, then ramp to 50:0:0:50 A/B/C/D until 8 min and then ramp to 36.7:0:63.3:0 A/B/C/D, then continue to ramp to 5:0:95:0 A/B/C/D at 14 min and hold until 15.5 min, then ramp to 0:5:95:0 74 A/B/C/D at 16 min, hold until 18 min, and then ramp to 99:0:0:1 A/B/C/D by 20 min and hold until 30 min to equilibrate the column. The UV-vis and HRMS chromatograms were acquired using the same parameters described above. Under these chromatographic conditions, good separation was achieved between the acid glucoside [1-O-(4-hydroxybenzoyl)-β-D-glucopyranoside] and phenolic glucoside [4-(β-D-glucosyloxy)-benzoic acid] of pHBA. Authentic standards were used for peak validation. 4.3.7 Monolignol–p-hydroxybenzoate conjugates Derivatisation followed by reductive cleavage (DFRC) was used to identify ester-linked conjugate groups in lignin (Regner et al., 2018). To a 50-mg sample of extractive-free 40-mesh wood powder in a small glass vial, 3 mL of brominating solution was added containing 20% acetyl bromide in acetic acid. After incubating at 50 °C for 2 h with stirring, the acetyl bromide was evaporated in a vacuum centrifuge. A 0.5-mL portion of ethanol was added to quench any residual acetyl bromide, and this too was then evaporated. Next, 100–150 mg of nano-powdered zinc was added along with 5 mL of a 5:4:1 mixture of 1,4-dioxane/acetic acid/water and the vial was incubated for 1 h in an ultrasonic water bath. After spiking with an internal standard solution containing deuterated synthetic standards, the reaction products were extracted in a separatory funnel charged with 10 mL of saturated ammonium chloride solution using 3 × 10 mL of dichloromethane. The combined organic phases were dried with anhydrous sodium sulphate, filtered into a round-bottom flask through filter paper, and evaporated using a rotary evaporator apparatus. The dried residue was acetylated by adding 1 mL each of acetic anhydride and pyridine, which was then also removed by evaporation. The residue was dissolved in 5 mL of dichloromethane and quantitatively transferred to a Supelclean LC-SI filter (500 mg silicon dioxide, Supelco Analytical, MilliporeSigma) and then eluted using 10 mL of a 1:1 mixture of ethyl acetate/hexanes. After drying with a rotary evaporator, the residue was dissolved in 1 mL of dichloromethane and the reaction products were analysed using a TQ8030 GC-MS (Shimadzu Corp.) equipped with an RXi-5Sil MS column (Restek Corp., Bellefonte, PA, U.S.A.) with helium as the carrier gas. Ions corresponding to cis and trans 4-acetoxysinapyl p-acetoxybenzoate were detected using multiple reaction monitoring mass detection as described previously (Regner et al., 2018). No peaks were detected for 4-acetoxyconiferyl p-acetoxybenzoate. 75 4.3.8 Gel-permeation chromatography The same enzyme-lignin samples prepared for NMR were used to evaluate lignin molecular weight by gel-permeation chromatography. First, 1 mL each of pyridine and acetic anhydride was added to 5-mg portions of enzyme-lignin powder in small glass vials, and acetylation reactions were allowed to proceed for 16 h at 50 °C. After the solvents were removed by evaporation in a vacuum centrifuge, 2 mL of tetrahydrofuran was added, and the vials were placed in a sonicating water bath for 1 hr. The dissolved lignin samples were then filtered through 0.2-µm PTFE syringe filters and analysed by gel-permeation chromatography using a Shimadzu Prominence HPLC (Shimadzu Corp.) equipped with a photodiode array detector and a series of three columns (TOSOH Bioscience, Tokyo, Japan): a TSKgel guard column with an Alpha stationary phase (6.0 mm × 4 cm, 13 μm), a TSKgel Alpha‐2500 column (7.8 mm × 30 cm, 13 μm), and a TSKgel Alpha‐M column (7.8 mm × 30 cm, 13 μm). Tetrahydrofuran was used as an eluent with a flow rate of 0.5 mL min-1 at 40 °C. Detection was performed at 270 nm and the molecular weights were calibrated using low molecular weight polystyrene ReadyCal Standards (MilliporeSigma) ranging from 266 to 66,000 Da. 4.3.9 Lignin histology and ToF-SIMS imaging Transversal xylem cross-sections were cut from air-dried stems to a thickness of roughly 10 µm using a sliding-block microtome. A drop of water on the blade prevented sample curling while sectioning. Phloroglucinol–hydrochloric acid (freshly prepared 2:1 mixture of 3% dye with concentrated hydrochloric acid) was used to stain lignin (Adler et al., 1948). Visualisation of stained sections was performed using a Reichert-Jung Polyvar microscope equipped with an Olympus UC90 full-colour camera (Olympus Corp., Tokyo, Japan). Images were cropped and scale bars were added using Adobe Illustrator (Adobe Systems Inc., Mountain View, CA, U.S.A.). ToF-SIMS imaging of pHB was performed as described previously (see section A3.3.3). Briefly, xylem cross-sections were cut to a thickness of roughly 100 µm and extracted in a Soxhlet apparatus for 4 h in 95% ethanol, 8 hours in ethanol/toluene (70/30), and finally 4 h in water. Sections were then dried fully in a desiccator before being affixed to glass slides using double-sided tape. ToF-SIMS spectra were 76 acquired with a ToF-SIMS 5 instrument (ION-TOF GmbH, Münster, Germany) using 60 keV Bi3++, 100 μs cycle time, and 512 × 512 pixels rastered randomly over an area of either 200 × 200 μm2 or 250 × 250 μm2. Delayed extraction and electron flooding were used during data acquisition, and the spectra were calibrated with peaks corresponding to C3H5+, C4H7+, C7H7+, and C5H5O2+ ions. A semi-quantitative index was calculated for pHB / S-lignin as a ratio of the ion counts for pHB (m/z 121.03) and S-lignin (sum of m/z 167.07 and 181.05) for regions of interest that either excluded vessel cell walls and included fibres, or vice versa. SurfaceLab 6.6 (ION-TOF GmbH, Münster, Germany) was used for data analysis, and false-colour ion image overlays were assembled using Adobe Photoshop and Adobe Illustrator (Adobe Systems Inc.). 4.3.10 Structural polysaccharides and saccharification potential A 10-mL aliquot of acid hydrolysate was retained for each sample from the Klason lignin determination procedure. The composition of structural polysaccharides was measured by high-performance anion-exchange chromatography. Good separation and detection of the neutral structural sugars was obtained using a CarboPac PA1 anion-exchange column (Dionex Corporation, Sunnyvale, CA, U.S.A.) and pulsed amperometric detection with a Dionex DX-600 HPLC apparatus fitted with a gold detector. An aliquot of 15 μL of 1:10 diluted acid hydrolysate was injected with water as the eluent at a flow rate of 1 mL min-1. A column clean-up was performed between samples using 200 mM sodium hydroxide, and detection was enabled by the post-column addition of 200 mM NaOH at a flow rate of 0.5 mL min-1. Fucose was used as an internal standard and calibration standards of known sugar concentrations enabled quantification and correction for mass increase upon hydrolysis. The saccharification potential was evaluated following pretreatment and enzymatic hydrolysis. Wood powder which had been processed in a Wiley mill to pass a 40-mesh sieve was subjected to either acidic pretreatment, alkaline pretreatment, or no pretreatment. Following the addition of 1 mL of either 2% H2SO4 or 62.5 mM NaOH to 20-mg samples of wood powder into screw-cap vials, pretreatment reactions were incubated at 90 °C for 3 h with frequent mixing by inversion. After cooling, the reactions were neutralised as required with either 4 M NaOH or 2 M H2SO4. Following centrifugation, the wood powder 77 was rinsed three times with deionised water and once with acetone, and then dried overnight at 50 °C. No-pretreatment controls were also washed in the same manner. Next, 10-mg samples were weighed into fresh vials and 1 mL of 100 mM sodium acetate buffer containing 0.2 fpu mL-1 of Cellic CTec3 HS enzyme cocktail (Novozymes A/S, Bagsværd, Denmark) was added and mixed by inversion. Enzymatic hydrolysis reactions were performed at 50 °C in a hybridiser-incubator rotating at 40 rpm. Reactions were terminated after 72 h by incubating at 90 °C for 10 min, and the amounts of liberated glucose and xylose were measured as described above. 4.3.11 Replicates and statistical analysis For the measurement of transgene expression, lignin content, lignin composition, cell-wall-bound pHB, soluble pHBA metabolites, and structural polysaccharides, five biological replicates were analysed per line and triplicate samples were processed for each. The DFRC reactions and the saccharification analysis used three biological replicates per line measured in triplicate. Three biological replicates were analysed using 2D-NMR and gel-permeation chromatography. HRMS of metabolites was performed using five biological replicates each of line 5 and the WT control. And finally, individual representative images were selected for lignin histology and ToF-SIMS imaging. For the HRMS analysis, a Student’s t-test was performed using MetaboScape 4.0 (Bruker Corp.) with any missing data imputed using the sample group mean values. Metabolite differences with a p-value of 0.01 and a four-fold change in signal intensity were deemed to be statistically significant. A Student’s t-test with equal variances was also used for the ToF-SIMS image analysis. For all other analyses, statistically significant differences between transgenic lines and the WT control were evaluated using a one-way ANOVA with a post-hoc Dunnett’s test performed using SPSS Statistics 27 (IBM, Armonk, NY, U.S.A.). 4.4 Results 4.4.1 Transgenic poplars expressing bacterial CPL Transgenic hybrid poplar lines were generated by Agrobacterium-mediated transformation to evaluate the effects of heterologous expression of bacterial CPL (Figure 4-2, Table 4-1). Five lines chosen for in-depth analysis were grown in a greenhouse alongside wild-type (WT) control trees for 18 weeks. Other 78 than a small yield penalty and the occurrence of sylleptic branching, which is characteristic of fast-growing hybrid poplar and could be a response to metabolic stress (Ceulemans et al., 1990), no major developmental defects were observed (Figure 4-3A). In comparison to control trees, transgenic lines were as much as 10% shorter and 12% thinner in stem diameter (Figure 4-3B). RT-qPCR was used to measure the transgene expression in developing xylem tissue and revealed that line 5 had the highest expression (Figure 4-3C). Figure 4-3. Expression of bacterial CPL in transgenic poplar. A: Representative photos of each transgenic line and wild-type (WT) control trees. B: Heights and stem diameters measured at the time of harvest are shown in blue and orange on the left and right axes, respectively. Those values marked with an asterisk are significantly different from the WT control, depicted with horizontal dashed lines (one-way ANOVA with Dunnett’s test, n=5 for each line with technical triplicates, p-value < 0.05). C: Relative expression of the ubiC gene, shown in green, was measured by RT-qPCR. No expression was detected for the WT control. Five biological replicates were analysed for each line using technical triplicates. 79 4.4.2 Alterations in lignin The xylem cell wall composition was analysed using extractive-free wood powder. The total Klason lignin content was slightly, but significantly, reduced in all lines compared to the WT control (Figure 4-4A). Of these, line 5 had the lowest lignin content with a 5% reduction in acid-insoluble lignin and a 15% reduction in acid-soluble lignin. Thioacidolysis was then used to assess whether there was a change in lignin composition. Analysis of liberated monomers revealed a significant albeit small decrease in S lignin in all transgenic lines except line 1 (Figure 4-4A). Two-dimensional 1H–13C heteronuclear single-quantum coherence nuclear magnetic resonance (HSQC NMR) spectroscopy was used to further examine changes in the composition and structure of lignin. Spectra of enzyme-lignin preparations confirmed the small shift in monomer composition (Figure 4-4B). Moreover, this analysis revealed an increase in cell-wall-bound pHB groups. Volume integrations showed a 50% increase in the amount of pHB in the mature xylem of line 5 compared to WT. Furthermore, the proportion of β-aryl ethers was slightly increased at the expense of resinol structures (Figure 4-5). Resinol units arise from the dimerisation of monolignol radicals when the γ-OH traps the quinone methide intermediate (Ralph et al., 2004a). However, the occurrence of γ-linked pHB groups precludes this mechanism so that other structures including β–β-linked tetrahydrofuran units as well as β-aryl ethers prevail in highly γ-acylated lignins (Lu & Ralph, 2008). Next, gel-permeation chromatography was used to ascertain whether increases in p-hydroxybenzoylation affected lignin molecular weight (Figure 4-4C). This analysis showed that there were no statistically significant differences on either a weight- (MW) or number-average basis (MN). 4.4.3 Elevated cell-wall-bound p-hydroxybenzoate To corroborate the NMR results, we quantified the amount of pHBA released from extractive-free cell wall material using mild alkaline hydrolysis (Figure 4-6A). As pHB groups are ester linked, they can be liberated by saponification. The transgenic poplars all contained significantly more pHB than WT control trees, with line 5 again being the most altered. In developing xylem, line 5 had at least 9 times greater levels of pHB groups than the control trees. However, the differences were less pronounced in mature 80 Figure 4-4. Lignin analysis of transgenic poplar expressing bacterial CPL. Lignin content, composition, and linkage prevalence were altered in transgenic poplars due to the incorporation of p-hydroxybenzoate. A: The Klason lignin content is plotted on the left axis with the acid-insoluble fraction shown in dark blue and the acid-soluble fraction in light blue, and the ratio of syringyl (S): guaiacyl (G) lignin units is plotted on the right axis in orange. Those values marked with an asterisk are significantly different from the wild-type (WT) control, depicted with horizontal dashed lines (one-way ANOVA with Dunnett’s test, n=5 for each line with technical triplicates, p-value < 0.05). B: Two-dimensional 1H–13C HSQC NMR spectra for enzyme-lignin samples of line 5 and the WT control, as labelled, showing the aromatics region. The colour-coding and peak annotations for p-hydroxyphenyl (H, light purple), guaiacyl (G, blue) and syringyl units (S and S', dark and light purple), as well as p-hydroxybenzoate pendent groups (yellow, pHB) are elaborated with the structures shown on the side. Proportions based on integrated peak volumes are provided as the mean ± standard error for three biological replicates. C: The molecular weight distribution is plotted on a weight-average basis (MW) in blue and on a number-average basis (MN) in orange. 81 Figure 4-5. Additional NMR spectra of transgenic poplar expressing bacterial CPL. Two-dimensional 1H–13C HSQC NMR spectra for enzyme-lignin samples of line 5 and the WT control, as labelled, showing the aliphatics region. The colour-coding and peak annotations are elaborated with the structures shown below for ꞵ-aryl ether (A, cyan), phenylcoumaran (B, green), resinol (C, magenta), and tetrahydrofuran units (C', magenta). Proportions based on integrated peak volumes are provided as the mean ± standard error for three biological replicates. 82 Figure 4-6. Analysis of monolignol conjugates in transgenic poplar expressing bacterial CPL. A: The amount of p-hydroxybenzoate released by mild alkaline hydrolysis is shown for developing xylem (blue) and mature xylem (orange). B: The amount of sinapyl p-hydroxybenzoate conjugates released by DFRC (sum of cis and trans isomers of 4-acetoxysinapyl p-acetoxybenzoate) is shown in green for mature xylem. No conjugates of coniferyl p-hydroxybenzoate were detected. Those values marked with an asterisk are significantly different from the wild-type (WT) control (one-way ANOVA with Dunnett’s test, n=5 for each line with technical triplicates, p-value < 0.05). xylem which is comprised mostly of cells that have undergone programmed cell death. In mature xylem, line 5 had 50% more pHB than WT, a value that conforms with the NMR spectra of the same samples. Next, the derivatisation followed by reductive cleavage (DFRC) method was used to validate the findings from alkaline hydrolysis and to confirm that pHB groups originated from monolignol–pHB conjugates. Since reductive cleavage releases monomers bound via β-aryl ether linkages but does not cleave ester bonds, DFRC yields a diagnostic product of S–pHB conjugates, namely 4-acetoxysinapyl p-acetoxybenzoate. Analysis of mature xylem showed significantly higher levels of S–pHB conjugates in all transgenic lines except line 1, with line 5 having 70% more than the WT control (Figure 4-6B). 4.4.4 Accumulation of p-hydroxybenzoate-containing metabolites As CPL produces pHBA, the key precursor to pHB groups, it was reasonable to anticipate elevated levels in transgenic trees. However, in order to be deployed to the cell wall, pHBA is presumably first converted 83 into p-hydroxybenzoyl-CoA, then conjugated with a monolignol, and finally exported from the cell to the apoplast (see Figure 4-1). In this light, the levels of free pHBA were measured next. The developing xylem of all transgenic lines contained significantly more than control trees (Figure 4-7A). At the same time, much more pHBA occurred as conjugates – in both alkali-labile and acid-labile forms. Compared to control trees, line 5 had 20 times more total pHBA as metabolites in developing xylem. Moreover, the level of pHBA in soluble metabolites was comparable to the amount of cell-wall-bound pHBA released by alkaline hydrolysis. UHPLC coupled with HRMS was then applied to methanolic extracts of line 5 and WT control samples to separate and identify the various pHBA-containing metabolites. This analysis revealed numerous compounds that were over-represented in line 5 compared to control trees (Figure 4-7B). Many of these metabolites evidently contained p-hydroxybenzoate moieties as shown by the presence of a fragment ion at m/z 137.024 (i.e., peaks 1–6, 8). Putative compound identities were deduced by examining the fragment patterns and comparing to library databases (Figure 4-7C). The most abundant peak in line 5 samples (peak 1) resulted from the coelution of more than one glucoside of pHBA. In order to resolve these compounds, the methanolic extracts were subjected to further chromatographic analysis (Figure 4-7D). Authentic standards were used to confirm the presence of both the acid glucoside and the phenolic glucoside of pHBA in the developing xylem of line 5 poplars. Both compounds have also previously been detected at low levels in the extracts of a variety of plant taxa (Herrmann, 1978). 84 Figure 4-7. Analysis of soluble metabolites in transgenic poplars expressing bacterial CPL. A: The amounts of free p-hydroxybenzoate (dark blue) as well as p-hydroxybenzoate released by alkali treatment (medium blue) and p-hydroxybenzoate released by alkali and acid treatments (light blue) are shown for methanolic extracts of developing xylem. Those values marked with an asterisk are significantly different from the wild-type (WT) control (one-way ANOVA with Dunnett’s test, n=5 for each line, p-value < 0.05). B: Volcano plot showing xylem metabolites in methanolic extracts with an abundance significantly different in line 5 compared to the WT control. The numbered metabolites in the shaded areas have a fold change in signal intensity greater than 4 or less than -4 with a p-value less than 0.01 (Student’s t-test, n=12 for each line). C: Chemical formulae and putative metabolite identities were assigned to metabolites by examining the m/z values and fragmentation patterns. Metabolites related to p-hydroxybenzoate produced a fragment with m/z 137.024, highlighted in blue. D: UHPLC-DAD traces for methanolic extracts of line 5 and the WT control at 255 nm showing the presence of the acid glucoside and the phenolic glucoside of p-hydroxybenzoate, confirmed using authentic standards. 85 4.4.5 Distribution of cell-wall-bound p-hydroxybenzoate Lignin staining with phloroglucinol showed no obvious differences in xylem cell morphology nor lignin distribution (Figure 4-8). Time-of-flight secondary-ion mass spectrometry (ToF-SIMS) was then used to evaluate the distribution of pHB groups in xylem cross-sections. The peak at m/z 121.03, which has previously been attributed to cell-wall-bound pHB (see Appendix 3), was used for mapping purposes and to prepare false-colour overlay images (Figure 4-9). This analysis revealed that pHB groups were predominantly limited to the cell walls of fibres in the xylem of both line 5 and WT. An index calculated for pHB ion counts normalised to those of S-lignin structures confirmed that the fibres of line 5 contained significantly more pHB groups compared to the WT control. When calculated for vessel cell walls, the index was again higher for line 5, but the values were lower due to the weak signal intensity of pHB in vessels. Figure 4-8. Lignin histology of transgenic poplar expressing bacterial CPL. Transverse xylem cross-sections of line 5 (A) and the wild-type control (B) stained with phloroglucinol-HCl. Scale bars represent 100 µm. 86 Figure 4-9. ToF-SIMS imaging of transgenic poplar expressing bacterial CPL. False-colour ion images with p-hydroxybenzoate groups shown in magenta (m/z 121.03) and lignin shown in green (sum of m/z 77.04 and 91.05) for extractive-free transverse xylem cross-sections of line 5 and the wild-type (WT) control, as labelled. The gradient scales on the right show ion intensities, and the white scale bars represent 50 µm. The pHB / S index was calculated as a ratio of ion counts for p-hydroxybenzoate at m/z 121.03 divided by the sum of S-lignin at m/z 167.07 and 181.05, and the values shown are the mean ± standard error (n = 5 for line 5, n = 3 for WT). 4.4.6 Improved saccharification Finally, the sugar released following various pretreatments and enzymatic hydrolysis was evaluated to assess the saccharification potential of the woody biomass. Transgenic poplar yielded greater amounts of glucose and xylose after 72 hours of enzymatic hydrolysis following either acidic or alkaline pretreatments (Figure 4-10), commensurate with the observed increases in p-hydroxybenzoylation. For 87 both glucose and xylose release, alkaline pretreatment performed the best. Compared to the WT control, line 5 released 10% more glucose and 50% more xylose following alkaline pretreatment. As the total amount of cell wall polysaccharides was actually slightly lower in line 5 compared to the control samples (Table 4-2), this increase in sugar release represents a bone fide improvement in saccharification. Figure 4-10. Saccharification potential of transgenic poplar expressing bacterial CPL. The release of glucose (shades of blue) and xylose (shades of green) is shown following various pretreatments (alkaline, acidic, and no pretreatment as labelled) and 72 h of enzymatic hydrolysis. Those values marked with an asterisk are significantly different from the wild-type (WT) control (one-way ANOVA with Dunnett’s test, n=3 for each line, p-value < 0.05). Table 4-2. Structural polysaccharides in transgenic poplar expressing bacterial CPL. Those values shown in bold are significantly different from the wild-type (WT) control (one-way ANOVA with Dunnett’s test, n=5 for each line, p-value < 0.05). Sugar (% w/w) Line 1 Line 2 Line 3 Line 4 Line 5 WT Glucose 43.88 ± 0.98 42.64 ± 1.72 44.97 ± 0.72 43.04 ± 1.51 44.08 ± 1.35 45.73 ± 0.87 Xylose 17.21 ± 0.32 16.67 ± 0.28 17.13 ± 0.24 17.05 ± 0.32 16.70 ± 0.93 18.81 ± 0.74 Mannose 1.69 ± 0.15 1.39 ± 0.07 1.58 ± 0.08 1.53 ± 0.08 1.42 ± 0.09 1.86 ± 0.09 Galactose 0.74 ± 0.03 0.71 ± 0.03 0.73 ± 0.05 0.74 ± 0.04 0.74 ± 0.02 0.79 ± 0.08 Rhamnose 0.42 ± 0.02 0.42 ± 0.02 0.38 ± 0.02 0.43 ± 0.04 0.39 ± 0.02 0.46 ± 0.01 Arabinose 0.31 ± 0.01 0.31 ± 0.02 0.30 ± 0.01 0.33 ± 0.01 0.31 ± 0.03 0.33 ± 0.04 88 4.5 Discussion 4.5.1 Ester-linked p-hydroxybenzoate pendent groups Although the occurrence of cell-wall-bound phenolics has long been recognised, there have only recently been concerted efforts to engineer these constituents for industrial gain using plant biotechnology (Ralph, 2010). In many ways, progress was stymied by confusion surrounding the nature of the covalent bond between pHB and lignin, with earlier studies favouring a linkage via the ⍺ position (Nakano et al., 1961; Okabe & Kratzl, 1965). Furthermore, various authors have suggested that pHB groups could be bound to lignin via ether linkages (Terashima et al., 1979; Kim et al., 1995; Sun et al., 2002). Similarly, the mechanism of incorporation was rather elusive. Although free pHBA can be incorporated into synthetic lignin in vitro, the efficiency is much improved with coniferyl–pHB (Okabe & Kratzl, 1965; Nakamura & Higuchi, 1978). Most likely, this is because free pHBA preferentially transfers radicals whereas the conjugates contain monolignol moieties that can sustain radicals and readily participate in coupling reactions (Ralph, 2010). Moreover, cross-coupling products of sinapyl alcohol and sinapyl–pHB have been detected in poplar, and diagnostic products of S–pHB can be released by DFRC (Morreel et al., 2004; Lu et al., 2004), providing strong evidence that pHB occurs as γ-linked esters arising from the incorporation of monolignol conjugates during lignification. Once the relevant BAHD monolignol transferase genes are identified, these will be obvious engineering targets. However, simply increasing the expression of monolignol transferases may not achieve the desired titres of pHBA. For example, heterologous expression of p-coumaroyl-CoA:monolignol transferases in poplar and Arabidopsis achieved at most 0.35 and 1.3% w/w cell-wall-bound p-coumarate, respectively (Smith et al., 2015; Sibout et al., 2016). Accordingly, we developed a complementary strategy to bolster the supply of the acylating phenolic components. 4.5.2 Diverting carbon flux with CPL By expressing bacterial CPL in plastids, we sought to divert carbon away from chorismate and the metabolic pathway leading to lignin. In this way, the addition of more pendent groups could simultaneously result in reduced lignin content. Although recent work has identified a cytosolic supply of 89 chorismate (Qian et al., 2019), it is generally thought that chorismate which is destined for lignin predominates in plastids. As chorismate is also an essential precursor in protein biosynthesis, a cellulose synthase promoter was selected in order to drive expression only in cells undergoing secondary cell wall formation. This strategy was successful in diverting carbon away from chorismate and general phenylpropanoid biosynthesis, and led to modest reductions in lignin content and a small shift in composition away from S lignin. Significant increases in cell-wall-bound pHB were also observed, particularly during xylem development where the best performing transgenic line had nearly 10-fold more pHB than WT trees. This study is the first to deliberately engineer pHB, but hardly the first to appreciate that pHB content can vary given the inherent plasticity of lignification. It has been shown that pHBA is synthesised via p-coumarate and a metabolic branch of the phenylpropanoid pathway, and recent evidence favours a β-oxidation route (Terashima et al., 1975; Yazaki et al., 1991; Widhalm & Dudareva, 2015). In this context, it is not surprising that metabolic flux through the lignin pathway can affect pHB content. For example, elevated pHB levels were observed in transgenic poplar deficient in the lignin biosynthetic enzymes caffeoyl-CoA O-methyltransferase, cinnamoyl-CoA reductase, C3'H, p-hydroxycinnamoyl-CoA:shikimate p-hydroxycinnamoyl transferase, cinnamate 4-hydroxylase (C4H), and 4-coumarate:CoA ligase (Meyermans et al., 2000; Zhong et al., 2000; Leplé et al., 2007; Coleman et al., 2008b; Peng et al., 2014; Wang et al., 2018a; Tsai et al., 2020). Conversely, pHB was reduced in F5H-overexpressing poplar (Stewart et al., 2009). Despite the increases in p-hydroxybenzoylation observed in CPL poplar, a large pool of pHBA remained unbound to the cell wall and accumulated as soluble xylem metabolites. Plants possess endogenous glycosyltransferase enzymes that can act on free pHBA to produce both the acid glucoside and the phenolic glucoside (Katsuṁata et al., 1989; Lim et al., 2002; Nishizaki et al., 2014). These same compounds accumulated when CPL was expressed in tobacco, potato and sugarcane (Siebert et al., 1996; Köhle et al., 2003; Viitanen et al., 2004; McQualter et al., 2005). Furthermore, it has been reported that glucosylation of pHBA occurs cytosolically in Lithospermum and that these glucosides are sequestered in the vacuole (Yazaki et al., 1995). 90 As conjugation of pHBA with monolignols is also believed to occur in the cytosol, the implication is that glucosylation potentially competes with monolignol acylation. This could be problematic because metabolites that have been transported into the vacuole are presumably inaccessible to β-glucosidase enzymes which could release the phenolic aglycones. However, when radiolabelled pHBA was provided to tobacco cell cultures, both the phenolic glucoside and the acid glucoside initially accumulated before a portion of the latter was subsequently incorporated into the cell wall (Li et al., 1997). Although a fulsome cell wall analysis was not provided in that study, this observation suggests that the acid glucoside can be cleaved and deployed to the apoplast, perhaps as cells mature, undergo cell death, and release their vacuolar contents. Glucosylation is not the only fate for surplus pHBA in the developing xylem of CPL poplar. Various derivatives of the glucoside salicin occur naturally in poplar and willow (Boeckler et al., 2011). These metabolites contain phenolics, typically benzoate or cinnamate, that acylate either the sugar, as in populin and tremuloidin, or the salicyl alcohol group, as in populoside (Braconnot, 1830; Pearl & Darling, 1959; Erickson et al., 1970). Although pHBA-containing versions have been synthesised (Stepanova et al., 2014), these apparently have not been reported as naturally occurring. But based on mass fragment patterns, we have detected peaks that may correspond to pHBA-containing derivatives of salicin in the developing xylem of CPL poplar (peaks 3 and 4, Figure 4-7B). Glucosides of gentisyl alcohol also occur in poplar, and these too can be acylated with phenolic acids as in salireposide and nigracin (Wattiez, 1931; Thieme & Benecke, 1967). One compound in the pool of line 5 xylem metabolites may be a p-hydroxybenozylated homologue of these (peak 6, Figure 4-7B). And finally, possible pHBA-containing versions were detected of grandidentatin (peaks 5 and 8, Figure 4-7B), a glucoside of 1,2-cyclohexanediol acylated with p-coumarate which has been identified in the bark of Populus grandidentata (Pearl & Darling, 1962) and in the xylem of RNAi-suppressed C3'H transgenic poplar (Coleman et al., 2008b). The occurrence of pHB groups in these metabolites in place of other phenolics underscores the effectiveness of bacterial CPL in planta, and also reveals substantial plasticity in the biosynthesis of these compounds in poplar xylem. 91 4.5.3 Strategies to optimise p-hydroxybenzoylation Realising even greater pHB levels may ultimately require combined expression of CPL with a natural or engineered p-hydroxybenzoyl-CoA:monolignol transferase. Ongoing work is focused on identifying candidate genes, but additional strategies may also be contemplated. One of the most impactful interventions would be blocking the formation of pHBA-containing metabolites. UDP-glycosyltransferases play a central role in detoxifying and modulating the accumulation of phenolics (Le Roy et al., 2016). Unfortunately, the specific enzymes responsible for glucosylation of pHBA in poplar are currently unknown. Although the biosynthesis of salicinoids also remains largely obscured (Babst et al., 2010), recent work has uncovered some of the steps involved in the formation of phenolic glycosides in poplar so these may soon be accessible engineering targets (Fellenberg et al., 2020). Alternatively, if pHBA could be rapidly conjugated with monolignols and then deployed to the cell wall, there would be no need for detoxification and less pHBA would become sequestered as metabolites in vacuoles. Phenolic metabolites in the xylem of poplar could also represent an untapped fount of valuable compounds (Devappa et al., 2015). Many salicinoid metabolites are active as defence compounds in poplar (Boeckler et al., 2011), and some of these may also have medicinal value. Furthermore, as most of the pHBA-containing compounds in the xylem of CPL poplar were ester-linked conjugates, treatment with alkali could be used to liberate the pHBA aglycones. In many ways, the accumulation of readily extractable pHBA as xylem metabolites in CPL poplar could be highly advantageous. We have previously shown that pHB groups occur predominantly in the cell walls of xylem fibres in poplar and willow (see Appendix 3). While there was a small but proportionate increase in cell-wall-bound pHB groups in the vessels of CPL poplar, the levels remained much lower than in fibres suggesting that pHBA may not be compatible with lignification in G-lignin rich vessels. Although the selection of the cellulose synthase promoter likely ensured an ample supply of pHBA for all xylem cell types, monolignol–pHB conjugates were still not efficiently produced in vessels. This could be the result of limited expression of p-hydroxybenzoyl-CoA:monolignol transferases, or perhaps related to the poor availability of sinapyl alcohol in lignifying vessels as acyltransferases are known to prefer this monolignol 92 as an acyl acceptor (Wither et al., 2012). In order to achieve p-hydroxybenzoylation in the G-rich xylem vessels of poplar, future engineering work may need to contemplate sinapyl alcohol availability or use non-selective transferases that also act upon coniferyl alcohol. 4.5.4 Impacts of ester-linked monolignol conjugates Although the biological role of ester-linked pendent groups remains unclear, it has been suggested that one result may be increased polymerisation rates for S-rich lignin (Takahama et al., 1996). There is also some evidence that such groups may promote plant defence as carrot cells respond to a fungal elicitor by incorporating cell-wall-bound pHB groups (Schnitzler & Seitz, 1989). Although plant growth was slightly impaired in CPL poplar, no irregular xylem phenotypes were observed and there was no indication that changes in lignin structure had other discernible effects on development. Aside from any potential biological implications, elevated pHB levels led to improved biomass digestibility. Previous lignin engineering efforts have achieved improved saccharification by significantly altering the lignin content, composition, and/or molecular weight (see Chapter 2). In one of the few reports on the effects of pendent groups, overexpression of a p-coumaroyl:CoA–monolignol transferase in Arabidopsis led to novel p-coumarate groups and significantly improved saccharification, although this was also accompanied by a reduction in lignin content (Sibout et al., 2016). In contrast, CPL-expressing poplar had only modest reductions in lignin (less than 5%), a small increase in β-aryl ether units, and no change in average molecular weight. These observations suggest that the gains realised in saccharification could be specifically related to elevated pHB levels. It may be that these pendent groups alter the interactions between cell wall components or otherwise improve substrate access during pretreatment and/or enzymatic hydrolysis. It has also previously been suggested that released pHBA acts as a blocking agent to prevent lignin condensation reactions and effectively enhance depolymerisation rates (Chua & Wayman, 1979). Beyond improvements in biomass processing, cell-wall-bound pHB groups are themselves potentially valuable as phenolic compounds. However, the industrial use of such clip-offs will only be economical if the recovery processes are cost effective (Karlen et al., 2020). In this light, CPL poplar may be an ideal 93 feedstock candidate for biorefineries as alkaline hydrolysis would provide a pure stream of phenolics without the need for further separations unlike, for example, bioenergy grasses which would release heterogenous mixtures of p-coumarate and ferulate. In general terms, the lignin engineering strategy promoted herein could be applied to the manipulation of other clip-offs as well. For example, increased p-coumaroylation and ferulolyation could be facilitated by enriching the pools of those respective phenolic acyl groups. As support for this approach, the levels of cell-wall-bound benzoate, which was only recently found to be compatible with lignification in poplar, were elevated in plants deficient in C4H and C3'H (Kim et al., 2020). In summary, expression of bacterial CPL in transgenic poplar led to increased p-hydroxybenzoylation of lignin and enhanced saccharification potential of the wood, but no change in the distribution of pHB groups in the xylem. This work demonstrates the importance of substrate supply in engineering cell-wall-bound phenolics. However, abundant pHBA-containing metabolites in these transgenic poplars also represent untapped potential. Accordingly, pHB groups warrant further research consideration in the context of lignin valorisation as these ester-linked pendent groups can be clipped-off for use in the production of a wide variety of biochemicals, biomaterials, and bioproducts. 94 Chapter 5: Convergent evolution of lignin p-coumaroylation provides new tools for lignin engineering 5.1 Summary Ester-linked p-coumarate (pCA) is a hallmark feature of the secondary cell walls in commelinid monocot plants. It has been shown that pCA groups arise during lignification from the participation of monolignol conjugates assembled by p-coumaroyl-CoA:monolignol transferase (PMT) enzymes, members of the BAHD superfamily of acyltransferases. Herein we report that kenaf (Hibiscus cannabinus), a eudicot species, naturally contains p-coumaroylated lignin in the core tissues of stems. Moreover, we identified a novel acyltransferase, HcPMT, that shares less than 30% amino acid identity with monocot PMT sequences. Recombinant HcPMT showed a preference in enzyme assays for p-coumaroyl-CoA and benzoyl-CoA as acyl donors, and sinapyl alcohol as an acyl acceptor. Heterologous expression of HcPMT in hybrid poplar trees led to the incorporation of pCA, an increase in the average molecular weight of lignin, and no change in saccharification potential. The occurrence of pCA outside of monocot lineages provides a new example of convergent evolution in lignin. Furthermore, this work shows the value in mining diverse plant taxa for new monolignol acyltransferases and highlights the potential of pCA in lignin engineering. 5.2 Introduction Lignin is a phenolic polymer found predominantly in the secondary cell walls of vascular plants. The deposition of lignin alters the physicochemical properties of cellulosic cell walls and thereby contributes to mechanical strength, water conduction, and plant defence (Barros et al., 2015; Mottiar et al., 2020). At the same time, lignin presents a formidable challenge for the industrial use of plant biomass due to its considerable chemical recalcitrance (Yoo et al., 2020). An improved understanding of lignin biosynthesis and formation could be harnessed to enhance feedstock species and thereby economise the sustainable use of plant biomass for the production of biochemicals, biofuels, and an array of other bioproducts and biomaterials. 95 Lignin is assembled primarily from three monolignol precursors – p-coumaryl, coniferyl, and sinapyl alcohols – which form p-hydroxyphenyl (H), guaiacyl (G), and syringyl (S) lignin units once incorporated into a polymer (Boerjan et al., 2003). The biosynthesis of monolignols occurs in the cytosol via the general phenylpropanoid pathway. Following export of monolignols from the cell, lignin polymerisation occurs in muro via the coupling of monolignol radicals generated by peroxidase and laccase enzymes. As this assembly process is stochastic and dependent solely upon the supply and chemical propensity of monomers for radical coupling (Ralph et al., 2004a), the composition and structure of lignin varies widely across different cell types and plant lineages. Aside from the three primary lignin monomers, other constituents can be found in many species. For example, the lignin of commelinid monocots features ester-linked p-coumarate pendent groups (pCA; Smith, 1955a; Higuchi et al., 1967). Similarly, p-hydroxybenzoate moieties (pHB) decorate the lignins of poplar and willow (Smith, 1955b; see Appendix 3). The lignin found in the bast fibres of kenaf is naturally acetylated (Ralph, 1996). And perhaps most remarkable, the leaves of Canary Island date palms have lignin with ester-linked pCA, pHB, benzoate (BA), ferulate (FA), vanillate (VA), and acetate (Ac) groups (Karlen et al., 2017). Although the biological role(s) of lignin acylation remains largely unknown, there is considerable interest in harnessing such biochemistry to create designer lignins (see Chapter 6). Ester-linked pendent groups arise from the incorporation of γ-acylated monolignol conjugates that participate in radical coupling (Lu & Ralph, 2002; Lu & Ralph, 2008; Figure 5-1A). Members of the BAHD superfamily of acyltransferases catalyse the assembly of these conjugates prior to their export and polymerisation (Hatfield et al., 2008a; Withers et al., 2012; Petrik et al., 2014; Marita et al., 2014). Moreover, due to the inherent plasticity of lignification, acyltransferases have been successfully used in lignin engineering. For example, heterologous expression of p-coumaroyl-CoA:monolignol transferase (PMT) genes from rice, Brachypodium, and maize led to the incorporation of novel pCA groups into the lignin of transgenic plants, including in Arabidopsis, poplar, and alfalfa (Smith et al., 2015; Sibout et al., 2016; Marita et al., 2016). Conjugates of pCA are predominantly assembled in monocots with sinapyl alcohol (Grabber et al., 1996), suggesting that PMT enzymes may have a bias in monolignol substrate preference. One role of 96 Figure 5-1. Incorporation of ester-linked pendent groups in lignin. A: Conjugates formed by p-coumaroyl-CoA:monolignol transferase (PMT) enzymes can participate in radical coupling and lignification, as shown. B: A hypothetical model of lignin from kenaf core tissue depicting p-coumarate groups in orange and ferulate groups in magenta. C: A hypothetical model of lignin from kenaf bast fibres depicting acetate groups in green. D: A hypothetical model of lignin from transgenic poplar expressing HcPMT depicting novel p-coumarate groups in orange, and native p-hydroxybenzoate groups in blue. 97 acylated monolignols may be to enhance the incorporation of S lignin units during lignification through radical transfer mechanisms (Takahama et al., 1996). Although pCA is a good substrate for peroxidase and laccase enzymes and can become integrated into synthetic dehydrogenative polymers formed in vitro, pCA occurs primarily as free-phenolic pendent groups in lignin (Ralph et al., 1994). This is because phenoxy radicals of pCA preferentially undergo radical transfer rather than coupling reactions during lignification (Hatfield et al., 2008b; Ralph, 2010; Gani et al., 2019), meaning that only the monolignol moieties of conjugates participate in polymerisation. This contrasts with monolignol–FA conjugates, for example, that engage in radical coupling efficiently at both ends and may thereby become integrated into lignin polymer backbones (Wilkerson et al., 2014). Cell-wall-bound pCA groups have traditionally been considered a unique feature in the cell walls of commelinid monocots (Harris & Hartley, 1980; Karlen et al., 2018). However, herein we report that the lignin of kenaf (Hibiscus cannabinus), a eudicot species, naturally contains ester-linked pCA pendent groups in the core tissue of stems. This was unequivocally shown by NMR and using derivatisation followed by reductive cleavage (DFRC) methods. We further characterised a BAHD acyltransferase from kenaf which showed activity as a PMT in assays of recombinant enzyme and when expressed heterologously in transgenic hybrid poplar trees. These observations support a growing consensus that lignin acylation, including by pCA groups, is not phylogenetically limited to a few select clades but rather occurs more broadly in diverse plant taxa. 5.3 Materials and Methods 5.3.1 Lignin analysis by NMR and DFRC Two-dimensional 1H–13C HSQC NMR spectra were collected using enzyme-lignin samples prepared by enzymatic digestion with Cellulysin cellulase (from Trichoderma viride, Calbiochem, MilliporeSigma, Burlington, MA, U.S.A.) following ball-milling of extractive-free cell wall material, as described previously (Kim & Ralph, 2010). HSQC spectra were collected using a Biospin AVANCE 700 MHz spectrometer (Bruker Corp., Billerica, MA, U.S.A.) with an adiabatic-pulse programme (hsqcetgpsisp2.2). The dimethyl sulphoxide peak at δH 2.5 ppm and δC 39.5 ppm was used for spectral calibration, and 98 volume integrations were performed using TopSpin 4.08 (Bruker Corp.). Proportions were expressed on a basis where % pCA = ½ pCA2/6 / 100% aromatics, % pHB = ½ pHB2/6 / 100% aromatics, 100% aromatics = ½ (H2/6) + G2 + ½ (S2/6 + S'2/6), and 100% aliphatics = Aα + Bα + Cα + C'α. Peak identifications were based on previous reports (Kim & Ralph, 2010). Ester-linked conjugates in lignin were released by DFRC, as described previously (Regner et al., 2018). For kenaf samples, products corresponding to the peracetates of cis- and trans-isomers of coniferyl 7,8-dihydro-p-coumarate (G–DHpCA), sinapyl 7,8-dihydro-p-coumarate (S–DHpCA), and coniferyl 7,8-dihydro-ferulate (G–DHFA) were detected using multiple reaction monitoring mass detection. For poplar samples, the peracetates of cis- and trans-sinapyl p-hydroxybenzoate (S–pHB) peaks were also measured although coniferyl p-hydroxybenzoate (G–pHB) was not detected. 5.3.2 Gene identification Total RNA was isolated from the stems of kenaf using the TRIzol reagent (Invitrogen, ThermoFisher Scientific, Carlsbad, CA, U.S.A.). cDNA was synthesised using the Creator SMART cDNA library construction kit (Clontech, Mountain View, CA, U.S.A.) and was subjected to 454 sequencing using a 454 GSFLX Titanium Sequencer (Roche, Basel, Switzerland). Sequence contigs were then aligned and compiled into a database which was queried for BAHD acyltransferase genes as described previously (Peers, 2012). A single BADH gene, denoted HcPMT, was chosen based on expression patterns and the presence of conserved motifs. A synthetic version of the 1335-bp HcPMT gene was procured from GeneArt (ThermoFisher Scientific, Waltham, MA, U.S.A.). 5.3.3 Recombinant protein expression The HcPMT coding sequence was transferred into the pDONR221 plasmid and then, using the Gateway cloning system (Invitrogen), into the pDEST17 expression vector which contains an N-terminal His6 tag to enable immobilised metal affinity chromatography (see Table 5-1 for primers and PCR conditions). This plasmid was transformed into chemically competent BL21 (DE3) pLysS E. coli cells (Invitrogen) for protein expression. 99 Table 5-1. Primers and PCR programmes for Chapter 5. Details are provided for all PCR reactions used in construct assembly, transformant screening, and RT-qPCR. Description: Primer pair (5' → 3'): PCR reaction conditions: Amplification of the synthetic HcPMT coding sequence (1335 bp) Forward primer: ATG GCA CTG CTA CGA CCG Reverse primer: CTA AGC ACC AGT TGC TTC ATC 2.5 µL ..................................10× buffer 0.5 µL .......................... 50 mM MgSO4 0.5 µL .............. 10 µM forward primer 0.5 µL ................ 10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ......................... template DNA* 0.2 µL ................ Pfx DNA polymerase Up to 25 µL ................ deionised water With thermal cycling as follows: 94°C for 30 s, 34 cycles of (94°C for 15 s, 60°C for 30 s, 68°C for 90 s), and then 68°C for 10 min. * Note: template DNA was the synthetic HcPMT coding sequence. Addition of the attB adaptor sequences to enable Gateway cloning Forward primer: GGG GAC AAG TTT GTA CAA AAA AGC AGG CTA TGG CAC TGC TAC GAC CG Reverse primer: GGG GAC CAC TTT GTA CAA GAA AGC TGG GTC TAA GCA CCA GTT GCT TCA TC 2.5 µL ..................................10× buffer 0.5 µL .......................... 50 mM MgSO4 0.5 µL .............. 10 µM forward primer 0.5 µL ................ 10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ......................... template DNA* 0.2 µL ................ Pfx DNA polymerase Up to 25 µL ................ deionised water With thermal cycling as follows: 94°C for 30 s, 34 cycles of (94°C for 15 s, 60°C for 30 s, 68°C for 90 s), and then 68°C for 10 min. * Note: template DNA was a plasmid containing the HcPMT coding sequence. PCR-screening to identify positive transformants Forward primer: CTG ACC GCA TGT GTT TGG Reverse primer: CTA AGC ACC AGT TGC TTC ATC Amplicon size: 543 bp 2.5 µL ..................................10× buffer 0.5 µL .......................... 50 mM MgSO4 0.5 µL .............. 10 µM forward primer 0.5 µL ................ 10 µM reverse primer 0.5 µL ............. 10 mM dNTPs mixture 0.5 µL ........................... template DNA 0.5 µL ............... Taq DNA polymerase Up to 25 µL ................ deionised water With thermal cycling as follows: 94°C for 3 min, 34 cycles of (94°C for 30 s, 55°C for 30 s, 72°C for 30 s), and then 72°C for 5 min. * Note: template DNA was genomic DNA preparations from young poplar leaves. Note: This table continues on the following page. 100 Note: Table continued from the preceding page. Description: Primer pair (5' → 3'): PCR reaction conditions: Quantification of expression levels using RT-qPCR For HcPMT transgene, Forward primer: GCC TGC AAG CGT TAT GGA AA Reverse primer: GCT TCA TCA TCT GCC AGG GT Amplicon size: 63 bp Priming efficiency: 99.9% For elongation factor 1β (reference), Forward primer: GGC ATT AAG TTT TGT CGG TCT G Reverse primer: GCG GTT CAT CAT TTC ATC TGG Amplicon size: 97 bp Priming efficiency: 99.9% For actin (reference), Forward primer: ACC AGT GTG TCT TGG TCT ACC C Reverse primer: CGA TGC CGA GGA TAT TCA AC Amplicon size: 127 bp Priming efficiency: 99.2% 5 µL .... BrightGreen qPCR master mix 0.3 µL .............. 10 µM forward primer 0.3 µL ................ 10 µM reverse primer 1 µL ............................ template cDNA Up to 10 µL ................ deionised water With qPCR parameters as follows: 95°C for 30 s, 39 cycles of (95°C for 5 s, 60°C for 15 s, plate read for fluorescence), and then followed by a melt-curve analysis as follows: 95°C for 10 s, ramp from 55°C to 95°C at 0.5°C per increment with hold times of 5 s and a plate read for fluorescence. * Note: template DNA was cDNA preparations synthesised with RNA extracted from developing poplar xylem. 101 A starter culture was inoculated in 10 mL of TB-Amp media (12 g L-1 tryptone, 24 g L-1 yeast extract, 2.3 g L-1 KH2PO4, 12.5 g L-1 K2HPO4, 4 mL L-1 glycerol, 50 µg mL-1 ampicillin). After 16 h incubation at 37 °C on an orbital shaker at 200 rpm, the cells were transferred into 250 mL of fresh TB-Amp media in a 1-L baffled flask to a starting OD600 of 0.1. After 6 h of additional cultivation, the OD600 reached 0.8 and the temperature was decreased to 16 °C. Protein expression was then induced by adding isopropyl-β-D-thiogalactopyranoside to a concentration of 10 mM. After 16 h of further cultivation at 16 °C, bacterial cells were harvested by centrifugation at 4000 g and 4 °C. The cell pellet was resuspended in an equal volume of BugBuster protein extraction reagent containing 1 U mL-1 benzonase nuclease (MilliporeSigma, Burlington, MA, U.S.A.) and 0.5 mM phenylmethylsulphonyl fluoride, and slowly agitated at room temperature for 20 min. Cell debris was then pelleted by centrifugation at 16,000 g and 4 °C, and the supernatant was slowly concentrated and buffer-exchanged 10,000 fold with 20 mM sodium phosphate buffer, pH 7.4 using an Amicon Ultra 15 mL 10K centrifugal concentrator unit (MilliporeSigma). The 5 mL-volume of crude protein was then loaded onto a His GraviTrap flow-through column (GE Healthcare, Chicago, IL, U.S.A.). After washing the column with 8 mL of 20 mM sodium phosphate buffer containing 500 mM sodium chloride, elution was performed using 2-mL portions of the same buffer, but containing increasing concentrations of imidazole (50 mM, 100 mM, 150 mM, 200 mM, 250 mM, 300 mM and 600 mM). SDS-PAGE analysis revealed that the fraction eluted using 150 mM contained a protein with an apparent molecular weight close to the theoretical size of HcPMT (~50 kDa). This fraction was buffer-exchanged 10,000 fold with 25 mM sodium phosphate buffer (pH 5.8), and the protein concentration was determined using the Bio-Rad protein assay (Bio-Rad Laboratories, Hercules, CA, U.S.A.) Finally, the protein preparation was flash-frozen by pipetting droplets into liquid nitrogen and then stored at -80 °C until further use. 5.3.4 Enzyme characterisation The purified enzyme was tested for activity with pCA-CoA, BA-CoA, pHB-CoA, FA-CoA and Ac-CoA as acyl donors and p-coumaryl, coniferyl, and sinapyl alcohols as monolignol acyl acceptors. BA-CoA and Ac-CoA were commercially available (Sigma-Aldrich, St. Louis, MO, U.S.A.) whereas the other CoAs 102 were enzymatically synthesised using Nt4CL1, as described previously (Beuerle & Pichersky, 2002). The monolignol conjugates used as authentic standards were produced following established chemical synthetic methodology (Zhu et al., 2013a). Enzyme assays were performed as described previously (Withers et al., 2012). Briefly, enzyme assays were set up as 50-μL reactions containing 50 mM phosphate buffer (pH 6), 1 mM dithiothreitol, 1 mM of each CoA thioester, 1 mM of each monolignol, and 5 μg of purified HcPMT enzyme (1 μg μL-1). The reactions were incubated at room temperature for 60 min at which point they were terminated by adding 50 μL of 100 mM hydrochloric acid, solubilised by adding 900 μL methanol, and then filtered through 0.2-μm nylon syringe filters. Reaction products were analysed by liquid chromatography coupled with mass spectrometry (LC-MS, LCMS8040, Shimadzu Corp., Kyoto, Japan) using a Kinetex XB-C18 column (250 × 4.60 mm, 100 Å, Phenomenex, Torrance, CA, U.S.A.) with water (A) and methanol (B) as eluents at a flow rate of 1 mL min-1. The binary gradient programme was ramped as follows: initial condition of 5% eluent B in eluent A, a linear increase to 100% eluent B over 30 min, hold for 4 min, then return to 5% eluent B over 1 min, and hold for 10 min to re-equilibrate the column. Reaction products were monitored using a photo diode array detector (λ = 250–400 nm) along with mass spectrometry in both positive and negative ion modes (Q3 scans between m/z 120–600). Products were verified by comparing retention times and mass fragmentation patterns to authentic standards. 5.3.5 Transgenic hybrid poplar The Gateway cloning system (Invitrogen) was used to transfer the synthetic HcPMT sequence from pDONR221 into the plant expression vector pMDC32 which contains a 35S promoter (Curtis & Grossniklaus, 2003). This binary vector was transferred into Agrobacterium tumefaciens EHA105 by freeze-thaw transformation of chemically competent cells (Wise et al., 2006). Agrobacterium-mediated transformation of hybrid poplar (Populus alba × grandidentata P39) was then performed as described previously (see section 4.3.2). Five transgenic lines were selected by PCR and multiplied in tissue culture. Prior to planting in the greenhouse, all plants were propagated from apical stem pieces to ensure even initial growth. For each 103 transgenic line, five biological replicates were planted with a peat-based soil in two-gallon pots, grown in the greenhouse under an 18-h daylight regime, and watered as needed with fertigated water. Immediately upon planting and for the subsequent two weeks, plants were covered with transparent plastic cups to maintain humidity during acclimatisation. Trees were supported with bamboo stakes and monitored regularly for any signs of disease or attack. Poplar trees were harvested after 18 weeks of growth. The plant heights and stem diameters were recorded, and the leaves and branches were removed. The bark was peeled from the stems, and developing xylem was scraped from the wood and flash-frozen in liquid nitrogen. Debarked stems were air-dried and then later cut into small pieces and comminuted to a powder using a Wiley mill fitted with a 40-mesh sieve. Extractive-free samples were obtained by extracting wood powder for 24 h with hot acetone in a Soxhlet extraction apparatus. 5.3.6 Transgene expression Xylem scrapings were ground to a fine powder in a mortar and pestle with liquid nitrogen, and total RNA was isolated using a CTAB-based procedure (Kolosova et al., 2004). Following DNase-treatment with the Turbo DNA-free kit (Ambion, Life Technologies), RNA was used in the synthesis of first-strand cDNA with the OneScript Plus cDNA synthesis kit (Applied Biological Materials Inc., Richmond, BC, Canada). Next, the cDNA was diluted 16-fold and used in qPCR reactions prepared with BrightGreen qPCR Master Mix (Applied Biological Materials Inc.). Gene-specific primers validated over a wide range of template concentrations were used (Table 5-1). Reference reactions were performed using primers designed to amplify the poplar actin and elongation factor 1β sequences. Thermal cycling, fluorescence detection, melt-curve analysis, and the calculation of relative expression was performed using a CFX 96 Real-Time PCR instrument (Bio-Rad Laboratories). 5.3.7 Lignin content, composition, and molecular weight The lignin content and composition were evaluated using established methodology for Klason lignin determination and thioacidolysis, as described previously (Huntley et al., 2003; Robinson & Mansfield, 2009). Lignin molecular weight was analysed for enzyme-lignin preparations using gel-permeation 104 chromatography, as described elsewhere (see section 4.3.8). 5.3.8 Cell-wall-bound phenolic acids and acetyl Cell-wall-bound phenolic acids were released by mild alkaline hydrolysis (saponification) and quantified using HPLC analysis, as described previously (see section A3.3.2). Cell-wall-bound acetyl was measured in a similar manner, as described previously (see section A2.2.5). 5.3.9 Structural polysaccharides and saccharification potential The composition of structural polysaccharides was measured by high-performance anion-exchange chromatography. Briefly, 15 μL of 1:10 diluted acid hydrolysate from the Klason lignin hydrolysis was injected onto a CarboPac PA1 anion-exchange column (Dionex Corporation, Sunnyvale, CA, U.S.A.) and the six major neutral sugars were separated using 1 mL min-1 water as the eluent. Pulsed amperometric detection was enabled by the post-column addition of 200 mM NaOH at 0.5 mL min-1. Saccharification potential was evaluated by pretreating wood powder with either 2% sulphuric acid or 62.5 mM sodium hydroxide for 3 h at 90 °C, and then subjecting it to enzymatic hydrolysis using 0.2 fpu mL-1 of Cellic CTec3 HS enzyme cocktail in 100 mM sodium acetate buffer at 50 °C. Reactions were terminated after 0.5, 1, 2, 6, 12, 24 or 72 h by heating to 95 °C for 10 min. Finally, the release of glucose and xylose was measured by anion-exchange chromatography. 5.3.10 Analysis of PMT sequences The amino acid sequence of HcPMT was aligned with the three commelinid monocot PMT sequences, OsPMT (Os01g18744), BdPMT (Bradi2g36910) and ZmPMT (GRMZM2G028104), using the Clustal Omega algorithm (Sievers et al., 2011). Sequence identity and similarity was calculated using the SMS Ident and Sim tool (Stothard, 2000). Next, putative homologues of HcPMT were identified from the following sequenced eudicot genomes using BLAST with a cut-off E-value of 10-100: Populus trichocarpa (JGI v3.0), Arabidopsis thaliana (TAIR 10), Vitis vinifera (Genoscope 12X), Medicago truncatula (MTGD Mt4.0), and Solanum lycopersicum (iTAG 4.0). Amino acid sequences were aligned using MEGA X with the MUSCLE algorithm and the default parameters (Kumar et al., 2018). This alignment was then subjected to a maximum likelihood analysis to construct a phylogenetic tree. The OsPMT, 105 BdPMT, and ZmPMT sequences were included as an outgroup, and the phylogeny was resampled using 1000 bootstrap iterations. 5.3.11 Replicates and statistics All enzyme assays were performed in triplicate. For the measurement of tree heights, stem diameters, transgene expression, lignin content, lignin composition, cell-wall-bound pCA, pHB, and acetyl, structural polysaccharides, and saccharification potential, five biological replicates were analysed per line and triplicate technical samples were processed for each. The DFRC reactions for transgenic poplars were conducted with three biological replicates per line measured in triplicate. Three biological replicates were analysed by NMR and gel-permeation chromatography for transgenic poplars. One sample each of kenaf bast and kenaf core tissue was used for NMR and DFRC. Statistically significant differences between the transgenic poplar lines and the WT control were evaluated using a one-way ANOVA with a post-hoc Dunnett’s test performed using SPSS Statistics 27 (IBM, Armonk, NY, U.S.A.). 5.4 Results and Discussion 5.4.1 Kenaf lignin is naturally p-coumaroylated The bast fibres of kenaf contain an S-rich lignin that is extensively acetylated (Ralph, 1996). It has been shown that acetyl groups predominantly acylate the γ-OH of S-lignin units, and that they derive from the incorporation of pre-acylated monolignols during lignification (Lu & Ralph, 2002). In contrast to the bast fibres, the core tissues of kenaf stems have a lignin that is less acetylated and has lower levels of S units (Seca et al., 1998). In addition, alkaline hydrolysis showed that mature kenaf stems may have cell-wall-bound pCA and FA (Geronikaki et al., 1979; Morrison et al., 1999). Previously, pCA groups may have been mistaken for H-lignin units (Neto et al., 1996). However, pyrolysis GC/MS of kenaf uncovered only trace amounts of hydroxycinnamic acids (Gutiérrez et al., 2004). To examine this further, we analysed kenaf bast and core tissues using solution-state two-dimensional 1H–13C heteronuclear single-quantum coherence (HSQC) NMR. Spectra collected from enzyme lignin preparations showed well-resolved aromatic signatures corresponding to H, G, and S lignin in both bast 106 Figure 5-2. Evidence that kenaf core lignin naturally contains pCA groups. A: 2D-NMR 1H–13C HSQC spectra for enzyme-lignin preparations of kenaf core and bast tissues, as labelled, showing the aromatics region. The colour-coding and peak annotations for p-hydroxyphenyl (H, mauve), guaiacyl (G, blue), and syringyl units (S and S', dark and light purple), as well as p-coumarate (pCA, orange) and ferulate groups (FA, pink) are elaborated with the structures shown. Also shown, in grey, are peaks corresponding to phenylalanine (Phe), tyrosine (Tyr), cinnamyl alcohol endgroups (I), cinnamaldehyde endgroups (J), and spirodienone units (D). B: Multiple reaction monitoring traces from GC-MS analysis of DFRC reaction products corresponding to kenaf core and bast tissues, as labelled. The spectrum for kenaf bast is delayed by 15 seconds. Peaks corresponding to cis and trans isomers of monolignols (H, mauve; G, blue; S, dark purple) and monolignols conjugates with p-coumarate (coniferyl 7,8-dihydro-p-coumarate, G–DHpCA and sinapyl 7,8-dihydro-p-coumarate, S–DHpCA; yellow) and ferulate (coniferyl 7,8-dihydro-ferulate, G–DHFA, pink) are labelled and colour-coded with reference to the structures shown. 107 and core tissues, with bast fibres having a higher ratio of S:G lignin units (Figure 5-2A). This analysis also revealed the presence of pCA and FA groups in the core tissues but not in bast fibres. DFRC reactions, which cleave β-aryl ether bonds in lignin while leaving ester-linkages intact, were performed next (Figure 5-2B). Diagnostic dihydro-p-coumarate (DHpCA) products derived from pCA conjugates with S-lignin (S–DHpCA) and G-lignin units (G–DHpCA) were detected from kenaf core tissue, but not bast fibres. Products corresponding to monolignol–FA conjugates were also observed, as has been reported previously (Karlen et al., 2016). Notably, pCA was associated with both G and S units, whereas FA was exclusively conjugated to G units. Our analysis confirms the presence of pCA in the core tissues of kenaf and provides a more fulsome view of kenaf lignin structures (Figure 5-1B, C). Recently, pCA was observed (but not remarked upon) in mulberry, a member of the eudicot family Moraceae (Yamamoto et al., 2020). These examples illustrate that p-coumaroylation is not restricted to monocot lineages, as has long been the prevailing view. In an extensive survey of dicots, pCA was released from the cell walls of diverse taxa including members of the Bignoniaceae, Caryophyllaceae, Gesneriaceae, Meliaceae, Rubiaceae, and Styracaceae families (Hartley & Harris, 1981). There have even been reports that small amounts of pCA in Populus spp., particularly in the bark (Pearl & Bayer, 1960; Sun et al., 2001). However, it remains to be seen how widely pCA occurs within the eudicots and especially whether it represents a substantial fraction of the total lignin in any of these taxa. Whether or not p-coumaroylation, or any lignin acylation for that matter, has an important biological function remains an open question. It is becoming clear, however, that acylated lignin moieties are far more common than had been previously appreciated (Ralph et al., 2019a). To date, only three PMT enzymes have been identified and characterised, and all occur in commelinid monocots: rice (OsPMT; Withers et al., 2012), Brachypodium (BdPMT; Petrik et al., 2014), and maize (ZmPMT; Marita et al., 2014). In fact, the only monolignol acyltransferase reported thus far from eudicots is the feruloyl-CoA:monolignol transferase (FMT) from Angelica sinensis (Wilkerson et al., 2014). In order to shed light on the occurrence of p-coumaroylation within the eudicots and potentially develop a new resource for lignin engineering, we set out to characterise a novel BAHD acyltransferase from kenaf. 108 5.4.2 Recombinant HcPMT produces monolignol–pCA conjugates HcPMT protein was expressed in E. coli and purified by immobilised metal affinity chromatography (Figure 5-3). The recombinant enzyme was first tested in a series of pairwise assays with individual acyl donor substrates (p-coumaroyl-CoA, pCA-CoA; benzoyl-CoA, BA-CoA; p-hydroxybenzoyl-CoA, pHB-CoA; feruloyl-CoA, FA-CoA; and acetyl-CoA, Ac-CoA) and monolignol acyl acceptors (p-coumaryl, coniferyl, and sinapyl alcohols). When provided with pCA-CoA and any of the three monolignols, recombinant HcPMT produced monolignol–pCA conjugates (Figure 5-4A–C). HcPMT also reacted with BA-CoA to produce coniferyl–BA and sinapyl–BA, but not p-coumaryl–BA (Figure 5-4D–F). Similarly, coniferyl–pHB and sinapyl–pHB were produced with pHB-CoA (Figure 5-4G–I). However, feruloyl-CoA yielded only sinapyl–FA as a product (Figure 5-4J–L). As FA is linked only to G units in kenaf lignin, this suggests that HcPMT is not responsible for feruloylation in kenaf. And finally, HcPMT produced sinapyl–Ac and coniferyl–Ac when provided with Ac-CoA (Figure 5-4M–O). In order to determine the preferred acyl donor, HcPMT was subjected to a series of substrate competition assays with all five acyl-CoA substrates along with each monolignol provided separately. In the presence of the five acyl donors and p-coumaryl alcohol, recombinant HcPMT produced only p-coumaryl–pCA (Figure 5-5A). With coniferyl alcohol as the acyl acceptor, coniferyl–pCA was produced along with a smaller amount of coniferyl–BA (Figure 5-5B). And similarly, with sinapyl Figure 5-3. Coomassie-stained SDS-PAGE gel of recombinat HcPMT. Recombinant HcPMT enzyme was purified from E. coli by immobilised metal affinity chromatography. The sizes of molecular weight standards (Precision-Plus Dual Colour Standards, Bio-Rad Labs) are labelled on the left in kDa. 109 Figure 5-4. Pairwise assays to screen HcPMT enzyme activity. A-C: Assays of pCA-CoA with the three monolignols produced p-coumaryl–pCA, coniferyl–pCA, and sinapyl–pCA. D-F: Assays with BA-CoA yielded coniferyl–BA and sinapyl–BA. G-I: Assays with pHB-CoA yielded coniferyl–pHB and sinapyl–pHB. J-L: Assays with FA-CoA yielded only sinapyl–FA. M-O: Assays with Ac-CoA yielded coniferyl–Ac and sinapyl–Ac. Chromatographic traces show the absorbance at 273 nm. Insets show mass fragmentation patterns for monolignol conjugates, which were validated using authentic standards. All reactions were performed at room temperature for 60 min with 1 µg purified recombinant enzyme, 1 mM dithiothreitol, 1 mM of each CoA thioester, and 1 mM of each monolignol in 50 mM phosphate buffer (pH 6). 110 Figure 5-5. Competitive enzyme activity assays testing HcPMT substrate preferences. A: In the presence of p-coumaryl alcohol and five CoA thioesters (pCA-CoA, BA-CoA, pHBA-CoA, FA-CoA, and Ac-CoA), HcPMT produces only p-coumaryl–pCA. B: In the presence of coniferyl alcohol and all five CoA thioesters, HcPMT produces coniferyl–pCA and coniferyl–BA. C: In the presence of sinapyl alcohol and all five CoA thioesters, HcPMT produces sinapyl–pCA and sinapyl–BA. Chromatographic traces show the absorbance at 273 nm. The insets show mass spectra for monolignol–pCA conjugates in blue and monolignol–BA conjugates in orange with m/z values labelled. All reactions were performed at room temperature for 60 min with 1 µg purified recombinant enzyme, 1 mM dithiothreitol, 1 mM of each CoA thioester, and 1 mM of each monolignol in 50 mM phosphate buffer (pH 6). alcohol and all five acyl donors provided, sinapyl–pCA and sinapyl–BA were produced (Figure 5-5C). The identities of the conjugation products were confirmed by mass spectrometry and with reference to authentic standards. Thus, from the five acyl donor substrates tested, recombinant HcPMT was most active with pCA-CoA and BA-CoA. Since acetyl-CoA was not a preferred substrate, it is unlikely that HcPMT is a contributor to lignin acetylation in kenaf. 111 5.4.3 Transgenic poplar trees expressing HcPMT have altered lignin Although in vitro assays are invaluable for determining substrate preferences and catalytic attributes, they do not necessarily reflect the true enzyme activity in vivo as enzymes that are apparently bifunctional may be catalytically limited by the availability of substrates. We therefore generated transgenic hybrid poplar trees to test the effects of HcPMT in plants. Five transgenic poplar lines representing independent stable transformation events were selected for in-depth analysis and were grown under controlled conditions in a greenhouse (Figure 5-6A). After 16 weeks of cultivation, some of the transgenic lines appeared to be stunted compared to wild-type (WT) control trees. Lines 4 and 5 were significantly shorter and line 5 had a smaller average stem diameter (Figure 5-6B). These lines also showed the highest levels of transgene expression, measured by RT-qPCR (Figure 5-7). In addition, line 5 exhibited extensive sylleptic branching which can be a sign of metabolic stress in hybrid poplars. Figure 5-6. Transgenic hybrid poplar trees expressing HcPMT. A: Representative photos of each transgenic line and the wild-type (WT) control. B: Heights and stem diameters measured at the time of harvest are plotted on the left and right axes, respectively, in blue and orange. Values marked with an asterisk are significantly different from the WT control (one-way ANOVA with Dunnett’s test, n=5 for each line using technical triplicates, p-value < 0.05). 112 Figure 5-7. Transgene expression in transgenic poplar expressing HcPMT. Relative expression of HcPMT, shown in green, was measured by RT-qPCR. No expression was detected for the wild-type (WT) control. Five biological replicates were analysed for each line using technical triplicates. To ascertain whether expression of HcPMT affected the biosynthesis and assembly of cell wall components, the Klason lignin content was measured using samples of extractive-free powdered wood. Compared to WT control trees, there were no significant differences in total lignin content nor in the proportions of acid-insoluble or acid-soluble lignin (Figure 5-8A). Similarly, no consistent differences were noted in the composition of structural polysaccharides (Table 5-2). There were, however, significant differences in lignin composition with lines 1 and 5 having a greater proportion of S lignin compared to WT control trees (Figure 5-8A). Next, the molecular weights of enzyme-lignin preparations were assessed using gel-permeation chromatography. Compared to WT control trees, lignin from lines 1, 3, 4, and 5 had greater weight-averaged molecular weight, MW, as well as an increased dispersity index, ĐM (Figure 5-8B). Alkaline hydrolysis reactions were then performed to release ester-linked cell-wall-bound components (Figure 5-9A; Figure 5-10). Substantial quantities of pCA were released from extractive-free wood powder for all five transgenic lines, whereas WT control trees yielded only trace amounts (Figure 5-1D). Poplar lignin is naturally p-hydroxybenzoylated and slight reductions in pHB groups were observed as a consequence of pCA incorporation, although these differences were only significant for line 1. When OsPMT was expressed in poplar, it was observed that the biosynthesis and incorporation of pCA groups did not come at the expense of pHB (Smith et al., 2015). Although recombinant HcPMT was also active in vitro on BA-CoA, and it is known that BA is compatible with lignification in poplar (Kim et al., 2020), only trace amounts of BA groups were detected (Figure 5-10). There was also no change in cell-wall-bound Ac groups confirming that HcPMT does not produce acetylated monolignols in planta. 113 Figure 5-8. Lignin analysis of transgenic poplar expressing HcPMT. A: The Klason lignin content is plotted on the left axis with the acid-insoluble and acid-soluble fractions shown in dark and light blue, respectively. The ratio of syringyl (S): guaiacyl (G) lignin monomers is plotted on the right axis in orange. B: The molecular weight is plotted on the left axis on a weight basis (MW) in dark green and on a number basis (MN) in light green. The dispersity (ĐM=MW/MN) is plotted on the right axis in magenta. Values marked with an asterisk are significantly different from the wild-type (WT) control (one-way ANOVA with Dunnett’s test, n=5 for each line using triplicates with p-value < 0.05 in A, and n=3 for each line with p-value < 0.1 in B). Table 5-2. Structural polysaccharides in transgenic poplars expressing HcPMT. The values shown in bold are significantly different from the wild-type (WT) control (one-way ANOVA with post-hoc Dunnett’s test, n=5 for each line, p-value < 0.05). Sugar (% w/w) Line 1 Line 2 Line 3 Line 4 Line 5 WT Glucose 41.25 ± 3.13 44.30 ± 2.97 44.30 ± 0.90 44.62 ± 0.93 46.60 ± 3.41 45.21 ± 0.79 Xylose 15.76 ± 3.53 17.78 ± 0.65 18.26 ± 0.30 18.18 ± 0.22 18.23 ± 0.34 17.64 ± 0.30 Mannose 1.45 ± 0.19 1.67 ± 0.16 1.60 ± 0.09 1.58 ± 0.06 1.80 ± 0.08 1.57 ± 0.10 Galactose 0.79 ± 0.05 0.74 ± 0.02 0.75 ± 0.05 0.74 ± 0.07 0.70 ± 0.09 0.77 ± 0.08 Rhamnose 0.44 ± 0.02 0.44 ± 0.01 0.46 ± 0.01 0.43 ± 0.02 0.43 ± 0.01 0.43 ± 0.01 Arabinose 0.38 ± 0.03 0.33 ± 0.01 0.36 ± 0.04 0.35 ± 0.02 0.27 ± 0.03 0.34 ± 0.02 114 Figure 5-9. Analysis of lignin acylations in transgenic poplar expressing HcPMT. A: The amounts of ester-linked p-coumarate (pCA) and p-hydroxybenzoate (pHB) released by alkaline hydrolysis are plotted on the left axis in orange and blue, respectively. Cell-wall-bound acetyl is plotted on the right axis in green. B: The amounts of the peracetates of sinapyl dihydro-p-coumarate (S–DHpCA) and coniferyl dihydro-p-coumarate (G–DHpCA) conjugates released by DFRC are plotted on the left axis in light orange and dark orange, respectively. Sinapyl p-hydroxybenzoate (S–pHB) is plotted on the right axis in blue. Values marked with an asterisk are significantly different from the wild-type (WT) control (one-way ANOVA with Dunnett’s test, n=5 in A and n=3 in B for each line using technical triplicates, p-value < 0.05). C: Partial 2D-NMR 1H–13C HSQC spectra for enzyme-lignin samples of line 4 and the WT control, as labelled, showing the aromatics region. The peak annotations for p-hydroxyphenyl (H, light purple), guaiacyl (G, blue) and syringyl units (S and S', dark and light purple), as well as p-coumarate (orange) and p-hydroxybenzoate pendent groups (yellow) are elaborated with the structures shown at the side. Proportions for integrated peak volumes are provided as the mean ± standard error for three biological replicates. 115 Figure 5-10. HPLC analysis of cell-wall-bound phenolics in transgenic poplar expressing HcPMT. A. Alkaline hydrolysates of extractive-free wood released substantial amounts of p-coumarate (pCA) from line 4 (green) and only trace amounts from the wild-type (WT) control (yellow). Detection was performed at 315 nm, the λmax for pCA. B. The same as in A, but with detection at 255 nm, the λmax for p-hydroxybenzoate (pHB), showing the slight decrease in pHB released from line 4 relative to the WT control. The shaded inset in B provides a zoomed-in view of 26–28 min and shows that only trace amounts of benzoate (BA) were detected. Chromatograms are also presented for authentic pCA (orange), pHB (blue), BA (red), as well as o-anisic acid (purple) which was used as an internal standard. 116 DFRC reactions were performed next to release intact ester-linked conjugates (Figure 5-9B). This work revealed that pCA predominantly occurred as conjugates with S-lignin units (S–DHpCA) since only minor quantities of G conjugates (G–DHpCA) were detected. The activity of HcPMT in transgenic poplar was therefore largely consistent with the native activity in kenaf core tissue. As has been observed previously, endogenous pHB was conjugated only to S units (see Chapter 4). As a final confirmation of pCA incorporation, enzyme lignin preparations from line 4 and the WT control were examined by NMR (Figure 5-9C). HSQC spectra corroborated the thioacidolysis and alkaline hydrolysis results and showed that the incorporation of pCA in poplar did not drastically alter the S:G lignin ratio and resulted in only a slight decrease in pHB groups. Again, no BA groups were observed indicating that HcPMT was unable to generate monolignol–BA conjugates in poplar likely because the preferred pCA substrate was readily available during lignification or perhaps because BA-CoA is limiting in poplar xylem tissues. Volume integrations indicated that pCA groups represented roughly 6.6% of the total lignin in transgenic poplar trees expressing HcPMT, much greater than the endogenous levels observed in kenaf core tissue. Inspection of the aliphatics region of the spectra showed that incorporation of additional γ-acylated monolignol conjugates also led to a small increase in the proportion of β-aryl ether linkages at the expense of phenylcoumaran and resinol structures (Figure 5-11). Such changes in lignin composition, molecular weight, and structure could potentially impact cell wall recalcitrance. 5.4.4 Industrial relevance of pCA The occurrence of pCA in the cell walls of commelinid monocots enhances lignin solubility due to the abundance of free phenolic groups (Lapierre et al., 1989). Similarly, lignin from transgenic Arabidopsis and poplar with novel pCA groups resulting from expression of BdPMT was significantly more soluble in alkali (Sibout et al., 2016; Lapierre et al., 2021). However, it has been reported that lignin interacts with hemicelluloses in plant cell walls through electrostatic and other non-covalent forces (Kang et al., 2019). A greater abundance of pCA pendent groups decorating lignin polymers could conceivably promote such interactions and thereby restrict access to structural polysaccharides. As such, the question of whether p-coumaroylation enhances or impedes cell wall digestibility remains largely unresolved. 117 Figure 5-11. Additional NMR spectra of transgenic poplar expressing HcPMT. Two-dimensional 1H–13C HSQC NMR spectra for enzyme-lignin samples of line 4 and the wild-type (WT) control, as labelled, showing the aliphatics region. The colour-coding and peak annotations are elaborated with the structures shown below for ꞵ-aryl ether (A, cyan), phenylcoumaran (B, green), resinol (C, magenta), and tetrahydrofuran units (C', magenta). Proportions for integrated peak volumes are provided as the mean ± standard error for three biological replicates. 118 To evaluate whether expression of HcPMT in poplar affected biomass recalcitrance, powdered wood was subjected to various pretreatments (alkaline, acidic, and no pretreatment) followed by 72-h of enzymatic hydrolysis (Figure 5-12). Regardless of pretreatment regime, no improvement in saccharification potential was observed among any of the transgenic lines. In fact, several lines showed significant reductions in digestibility compared to the WT control. This contrasts with the increased sugar release observed in transgenic Arabidopsis expressing BdPMT (Sibout et al., 2016). However, no change in cell wall digestibility was reported for alfalfa lines with elevated pCA levels resulting from expression of ZmPMT (Marita et al., 2016). In a study of inbred maize lines with varying contents of Figure 5-12. Saccharification potential of transgenic poplar expressing HcPMT. A: The release of glucose (shades of blue) and xylose (shades of green) is shown following various pretreatments (alkaline, acidic, and no pretreatment as labelled) and a 72-h enzymatic hydrolysis. Those values marked with an asterisk are significantly different from the wild-type (WT) control (one-way ANOVA with Dunnett’s test, n=3 for each line, p-value < 0.05). B: Enzymatic hydrolysis time course showing the release of glucose for line 5 (orange) and the WT control (magenta) following alkaline pretreatment. Error bars show standard deviation for three biological replicates, each measured in triplicate. C: The same as B but showing xylose release. 119 hydroxycinnamates, pCA was negatively correlated to cell wall degradability (Zhang et al., 2011). A possible explanation for these contrasting observations is that the lignin content was reduced in the transgenic BdPMT Arabidopsis, but unchanged in HcPMT-expressing poplar and in these other examples. The observed increases in average molecular weight of lignin in HcPMT transgenic poplar could offer another explanation for reduced digestibility as lignin polymer size and structure plays an important role in chemical recalcitrance (Tolbert et al., 2014). Although p-coumaroylation does not necessarily improve biomass digestibility, pCA itself may be a lucrative engineering target in its own right. As pCA groups are ester-bound to the lignin backbone, treatment with mild alkali can be used to efficiently release these pendent groups. In some applications involving fermentation, such free phenolics could be cytotoxic and detrimental to growth and conversion rates (Palmqvist & Hahn-Hägerdal, 2000). However, these ‘clip-offs’ would be commercially valuable if they could be economically separated from the residual biomass (Karlen et al., 2020; Timokhin et al., 2020). Liberated pCA could be used directly as there are well-established applications in foods, pharmaceuticals, and cosmetics (Boz, 2015; Pei et al., 2016; Boo, 2019). Alternatively, pCA could be converted into platform chemicals and then upgraded into various high-value bioproducts and biomaterials. For example, work is ongoing to develop enhanced strains of Pseudomonas and Rhodococcus capable of efficient catabolism of pCA to produce organic acids that can be readily upgraded into an array of valuable chemicals (Otani et al., 2014; Becker et al., 2015; Mohamed et al., 2020). In this way, the discovery and development of monolignol acyltransferases may provide valuable new molecular tools for lignin engineering and the pursuit of designer lignins (see Chapter 2). 5.4.5 HcPMT differs substantially from commelinid monocot PMT sequences A comparison of PMT amino acid sequences reveals noteworthy differences and similarities (Figure 5-13A). Although OsPMT, BdPMT, and ZmPMT all share 60% sequence identity, HcPMT is less than 30% identical. When amino acid similarities are considered, HcPMT is at most 43% similar to the other PMTs. All four sequences bear the universally conserved HXXXD and DXGWG motifs that are hallmarks of BAHD acyltransferases (St. Pierre & De Luca, 2000). The conserved histidine in the former sequence is known to 120 Figure 5-13. Analysis of PMT sequences and phylogeny. A: An alignment of the HcPMT amino acid sequence with those of the previously characterised PMTs from rice (OsPMT), Brachypodium (BdPMT), and maize (ZmPMT). Fully conserved residues are shown in bold, and residues conserved only among the commelinid monocot sequences are shown in blue. The universal motifs characteristic of BAHD enzymes are highlighted in purple, clade-specific motifs are highlighted in orange, and additional residues discussed in the text are highlighted in green (labelled as Note 1 and Note 2). B: A phylogenetic tree prepared by maximum likelihood analysis showing the relationship of HcPMT with putative homologues identified in Populus trichocarpa (Potri), Arabidopsis thaliana (AT), Vitis vinifera (GSVIV), Medicago truncatula (Medtr), and Solanum lycopersicum (Solyc). Sequences for OsPMT, BdPMT and ZmPMT were included as an outgroup. The values at each node represent statistical confidence based on 1000 bootstrap iterations. The scale bar corresponds to 0.2 substitutions per site. Sequences shown in green are discussed in the text. 121 deprotonate the hydroxyl group of acyl acceptors, whereas the latter motif is apparently structurally important (Ma et al., 2005). All the PMT sequences also have conserved motifs characteristic of clade Va, namely YPFAGR, QVTXXXCGG, and GXYGN (Tuominen et al., 2011). The rice FMT is also a member of this group, but the FMT from Angelica sinensis (the only other monolignol transferase reported from eudicots) falls into clade III (Karlen et al., 2016). Although no crystal structures are currently available for any PMT or FMT enzyme, comparisons with other BAHD acyltransferases may be insightful. For example, structure-functional analysis of the hydroxycinnamoyl-CoA:shikimate hydroxycinnamoyltransferases (HCT) enzymes from Coffea canephora and Sorghum bicolor revealed that glycine and tryptophan residues guard the entrance to the substrate binding pocket and thereby influence the selectivity toward various acyl donors (Lallemand et al., 2012; Walker et al., 2013). Although these residues are conserved in the monocot PMTs, alanine and leucine occur instead in HcPMT, perhaps resulting in a larger active site opening (see Note 1 in Figure 5-13A). Similarly, comparing the HCT structures with an anthocyanin malonyltransferase from Chrysanthemum × morifolium (Unno et al., 2007) points to a series of residues that may comprise the acyl acceptor binding pocket (see Note 2 in Figure 5-13A). Only some of these positions are conserved among all PMT sequences, however, so further interpretation will require a PMT structural model. 5.4.6 p-Coumaroylation in kenaf may be the product of convergent evolution Putative homologues of HcPMT were identified in the genomes of five eudicot species by amino acid homology and then used in the assembly of a phylogenetic tree (Figure 5-13B). As has been observed previously with BAHD acyltransferases (Tuominen et al., 2011), these sequences predominantly clustered by taxon, pointing to gene duplication and neofunctionalisation events that occurred after the common ancestors diverged. This pattern of taxon-specific expansion is typical of genes involved in secondary metabolism (Ober, 2005), and likely reflects the versatility in the active site of BAHD enzymes, which are known to accept a wide range of acyl donors and acceptors. Included among the putative homologues of HcPMT are several previously characterised BAHD enzymes that exhibit diverse activities. For example, AT3G03480 encodes an acetyl-CoA:(Z)-3-hexen-1- 122 ol acetyltransferase involved in the biosynthesis of leaf volatiles released upon wounding (D’Auria et al., 2007), and AT5G17540 is involved in the metabolism and homeostasis of brassinosteroids in Arabidopsis (Zhu et al., 2013b). In grape, GSVIVT01022762001 encodes an anthraniloyl-CoA:methanol acyltransferase that impacts flavour and aroma (Wang & De Luca, 2005). The alcohol acetyltransferases encoded by Solyc08g005770 and GSVIVT01024259001 in tomato and grape, respectively, accept a range of alcohol substrates (Maoz et al., 2018; Goulet et al., 2015). And finally, Potri.013G074500 and Potri.019G043600 encode BAHD enzymes that act as benzoyl-CoA: benzyl/salicyl alcohol O-benzoyltransferases in poplar (Chedgy et al., 2015). Clearly, sequence homology is a poor predictor of activity in BAHD enzymes, as has been noted previously (Luo et al., 2007). The observation that some of the closest homologues of HcPMT in eudicots have activity on various other acyl donors and acceptors suggests that HcPMT likely evolved independently of the monocot PMTs. Taxon-specific clustering in the phylogenetic analysis further indicates that HcPMT and the related eudicot BAHD sequences arose following the divergence of their common ancestors (i.e., long after the division between monocots and eudicots). Convergent evolution has been documented for BAHD enzymes previously (Luo et al., 2007; Peng et al., 2016). At present, it remains unclear how broadly pCA occurs within the eudicots beyond kenaf (order: Malvales) and mulberry (order: Rosales; Yamamoto et al., 2020). However, it is possible that PMT enzymes evolved repeatedly among the eudicots. Only further investigations on lignin acylations in diverse lineages will resolve this question. Of course, it is also possible that p-coumaroylation, or lignin acylation more generally, could be an ancestral trait that was subsequently lost from many extant genera. Given that a majority of plant taxa apparently lack lignin acylation, convergent evolution appears to offer a more plausible explanation. Independent evolution of plant cell wall components is hardly atypical. For example, mixed-linkage glucans evolved separately in horsetails (Equisetum spp.) and in the Poaceae monocots (Sørensen et al., 2008; Fry et al., 2008). Similarly, the biosynthesis of sinapyl alcohol, the monomer that yields S lignin units, developed independently in the angiosperms, the moss Selaginella moellendorffii and apparently in various basal gymnosperms (Weng et al., 2008; Espiñeira et al., 2011). It has also been postulated that the ability to acylate monolignols with ferulate has arisen at least twice since nonhomologous FMT sequences 123 occur in the eudicot Angelica sinensis and among the commelinid monocots (Wilkerson et al., 2014; Karlen et al., 2016). Indeed, the same logic may hold for other lignin acylations. For example, p-hydroxybenzoylation of lignin has been detected in the disparate angiosperm families Salicaceae and Araliaceae, as well as the monocot families Arecaceae and Posidoniaceae (Smith, 1955b; Pearl et al., 1959; Hibino et al., 1994; Rencoret et al., 2020). Clearly, the discovery of pCA in a eudicot species elicits new questions on the evolution of lignin. An equally important pursuit for future studies will be to investigate whether lignin acylations play any important biological role(s) in cell walls that could be adaptive for plants. In the meantime, the availability of HcPMT provides a new tool to modify lignin pendent groups and enable further progress in lignin engineering. 124 Chapter 6: Atypical lignification in eastern leatherwood (Dirca palustris) 6.1 Summary Lignin is a complex phenolic biopolymer found mainly in the secondary cell walls of vascular plants, where it contributes to mechanical strength, water conduction, and plant defence. We studied the lignin of eastern leatherwood (Dirca palustris) because this slow-growing woody shrub is known for its flexible stems. Various analytical techniques and microscopy methods were employed to examine the composition and distribution of lignin and structural polysaccharides in leatherwood xylem in comparison with trembling aspen (Populus tremuloides) and white spruce (Picea glauca). We found that leatherwood has low overall levels of lignin, a high syringyl lignin content, and a unique distribution of lignin. Most remarkably, the cell corners and middle lamellae remain unlignified in mature xylem. These findings help explain the flexibility of leatherwood and also call into question the classical model of lignification, which purports that lignin polymerisation begins in the cell corners and middle lamellae. This atypical lignification regime vividly illustrates the diversity in plant secondary cell wall formation that abounds in Nature and casts leatherwood as a new model for the study of lignin biogenesis. 6.2 Introduction Eastern leatherwood (Dirca palustris L.) is a slow-growing woody angiosperm found sporadically across eastern North America in rich, mesic soils of hardwood and mixed forests (Zasada et al., 2008). Leatherwood, a member of the family Thymelaeaceae, was traditionally used by indigenous peoples and early settlers for natural cordage as the stems are remarkably flexible (Thoreau, 1856; Figure A2-1; section A2.1.1). Much of the contemporary work has focused on this species’ horticultural potential and ecological context due to its attractive arboresque growth form and because the ephemeral flowers emerge in early spring when pollinator activity is limited (Williams, 2004; Peterson & Graves, 2011). The earliest recorded description of leatherwood was in 1700 by the Canadian physician and naturalist Michel Sarrazin (Mottiar, 2012). Nearly 200 yr later, the French botanist Van Tiegham first reported weakly lignified cell walls in various other members of the family Thymelaeaceae (Van Tieghem, 1893). 125 It has been suggested that poor lignification of the xylem is the cause of leatherwood’s flexibility (Nevling, 1962), but this has not been thoroughly examined. As lignin is both biologically and industrially important, a careful study of plants with atypical lignification could help address outstanding questions in lignin biology and could potentially inspire ongoing efforts to breed or engineer commercially relevant species for key industrial applications. Lignin is a phenolic polymer found mainly in the secondary walls of plant cells, where it contributes to structural integrity, xylem conductance, and plant defence (Weng & Chapple, 2010). Since lignin represents a significant investment of photosynthate derived carbon, its biosynthesis is tightly regulated in a spatiotemporal manner. Accordingly, lignin content and composition vary widely between cell types, throughout plant development, among plant taxa, and across different environmental conditions (Boerjan et al., 2003). Lignin biosynthesis occurs in the plastid and cytosol of plant cells via the shikimate and phenylpropanoid pathways, which give rise to p-coumaryl, coniferyl, and sinapyl alcohols, the primary lignin precursors. These three monolignols are subsequently exported to the apoplast, where enzyme-catalysed dehydrogenation yields radical monomers that undergo coupling and cross-coupling reactions to produce polymeric lignin with p-hydroxyphenyl, guaiacyl, and syringyl units, respectively (Ralph et al., 2004a). Perturbation in the supply of monolignols leads to lignin polymers with altered composition and structure, and noncanonical monomers can also be incorporated (see Chapter 2). The prevailing model of lignification recognises three phases: lignin initiation in the cell corners and middle lamellae, slow lignification during secondary cell wall formation, and rapid lignification following the deposition of cellulose and hemicellulose (Wardrop, 1957; Donaldson, 2001). As the cell corners and middle lamellae are the first to become lignified, it has been hypothesized that polymerisation begins at discrete nucleation sites located in these regions (Terashima, 1990; Donaldson, 1994). Although ferulate moieties bound to glucuronoarabinoxylan have been described as potential nucleation sites in the cell walls of grasses (Ralph et al., 1995), no universal initiation mechanism has yet been identified. Surveys of Nature’s diversity have helped demystify the molecular underpinnings of lignification, but much of the focus has been on model species, and reports of woody plants with unusual lignification 126 remain sparse. Herein, we describe the cell wall composition of leatherwood and reveal a unique distribution of lignin throughout the xylem and across the cell wall. Together, these results portray a remarkable example of atypical lignification that diverges from the classical model of lignin deposition and cell wall organisation. 6.3 Materials and Methods 6.3.1 Wood samples Woody stems were harvested from mature plants of D. palustris L. (eastern leatherwood) near Pembroke, Ontario (45° 49' N, 77° 60' W). Similarly, stems of Populus tremuloides Michx. (trembling aspen) and Picea glauca (Moench) Voss (white spruce) were collected as reference samples. Upright-growing stems were chosen to avoid any reaction wood, and all stems were devoid of any signs of damage or attack by pests or pathogens. Three biological replicates were harvested for each species from the same wood lot. All stems were between 1 and 3 cm in diameter, and samples used for microscopy and imaging were taken near the base. Additional samples of D. palustris, Dirca occidentalis, Dirca decipiens, and Dirca mexicana were generously provided by Z. Hudson (Iowa State University). 6.3.2 Analysis of lignin and structural polysaccharides Established protocols were followed for Fourier-transform infrared (FTIR) spectroscopy (potassium bromide pellets), lignin content (Klason lignin), structural polysaccharides composition (ultra-high performance liquid chromatography with diode array detection of benzocaine-derivatised acid hydrolysates), acetyl content (high-performance liquid chromatography with refractive index detection of alkaline hydrolysates), lignin composition (gas chromatography with flame ionisation detection of thioacidolysis reaction products), cellulose content (alpha cellulose), cell wall crystallinity (reflection mode X-ray diffraction), and cellulose microfibril angle (differential interference contrast microscopy). Detailed descriptions of the methodology are provided in section A2.2. 6.3.3 Microscopy Stem specimens for microscopy were sectioned using a sliding block microtome. A droplet of water was placed on the blade to maintain sample integrity while preparing 15 µm thick sections. Bright-field 127 microscopy was performed using a Zeiss Stemi 2000-C stereomicroscope (Carl Zeiss AG, Oberkochen, Germany) and a Leica DMR compound microscope (Leica Microsystems GmbH, Wetzlar, Germany) equipped with a Canon EOS Rebel T5 digital camera (Canon Inc., Tokyo, Japan). Confocal microscopy was performed to visualise lignin autofluorescence in unstained sections using an Olympus FV1000 MPE laser scanning microscope (Olympus Corp., Tokyo, Japan) equipped with a 405 nm excitation laser and emission filter sets spanning 425–525 nm. Images were cropped and scale bars were added using Adobe Illustrator (Adobe Systems Inc., San Jose, CA, U.S.A.), and pixel intensity plots were generated using ImageJ (National Institutes of Health, Bethesda, MD, U.S.A.). Double histochemical staining of lignified and nonlignified cell walls was achieved using safranin-O (1% dye in water) and astra blue (1% dye in 2% tartaric acid). After a 2 min incubation, sections were washed thoroughly with distilled water and examined by light microscopy. Staining of syringyl units in lignin was performed using the Mäule reaction. After a 5 min incubation in 0.5% potassium permanganate, sections were washed thoroughly with distilled water and incubated in 3.7% hydrochloric acid for an additional 2 min. Sections were then transferred to concentrated ammonium hydroxide and examined by light microscopy. Staining of lignin-associated aldehyde units was achieved using the Wiesner reaction with phloroglucinol–hydrochloric acid. After a 2 min incubation in freshly prepared stain (2 : 1 mixture of 3% aqueous phloroglucinol dye with concentrated hydrochloric acid), sections were examined by light microscopy. 6.3.4 Confocal Raman imaging A CM3050 S cryomicrotome (Leica) was used to prepare 16 μm thick sections for imaging with an Alpha300RA Raman instrument (WITec GmbH, Ulm, Germany) as described previously (Gierlinger et al., 2012). The samples were excited using a linear polarised VIS laser (λex = 532 nm, laser power 30 mW) via a 100× oil immersion objective lens (Zeiss, numerical aperture 1.4, coverslip correction 0.17 mm). Raman signals were backscattered through the same objective, directed through a 50 μm optical multifibre to a UHTS 300 spectrometer (WITec) fitted with a 600 g mm-1 grating, and finally to a DU401 BV CCD camera (Andor Technology Ltd, Belfast, UK). Control Four acquisition software (WITec) was 128 used for data collection with 0.3 μm steps at 0.07 s per spectrum, and the Project Four Plus software (WITec) was used for data analysis. After cosmic ray removal, Raman images were calculated based on band integration, and a hierarchical cluster analysis was performed as described previously (Gierlinger et al., 2012). 6.3.5 Time-of-flight secondary ion mass spectrometry imaging Stem specimens for time-of-flight secondary ion mass spectrometry (ToF-SIMS) analysis were extracted and solvent-exchange-dried with ethanol sequentially from 30% to 95% and then sectioned using a rotary microtome equipped with a diamond knife. Spectra were recorded by Surface Interface Ontario (University of Toronto) using a ToF-SIMS IV instrument (ION-TOF GmbH, Mnster, Germany) employing the scan parameters reported previously (Goacher et al., 2011). Since lignin contributes considerable charge background, low-energy electron flooding was used to facilitate the collection of positively charged secondary ions. Nominal mass resolution spectra were obtained using a pulsed current of c. 0.1 pA. Spectra were calibrated to the C+, CH3+, C2H3+, C2H5+ and C7H7+ ions using the Surface Lab 6.1 software (IONTOF). False-colour ion images were then assembled with lignin ions being those reported previously as ‘Group 1’ assignments by Goacher et al. (2011) as well as m/z 167 and 181 for syringyl lignin and 107 and 121 for p-hydroxyphenyl lignin moieties, and structural polysaccharides being those reported as ‘Group 2’ assignments with the addition of m/z 115 and 133 for xylan. 6.4 Results 6.4.1 Fourier transform infrared spectroscopy We initially used FTIR spectroscopy to survey the chemical composition of leatherwood xylem in comparison to samples of aspen (P. tremuloides) and spruce (P. glauca). These taxa were chosen for comparison because their cell wall composition and xylem histology have been studied extensively (Pettersen, 1984; Schweingruber et al., 2011). Principal component analysis was used to identify chemical features unique to leatherwood (Figure A2-2; section A2.1.2). To further investigate key spectral differences between the three species, curve fitting and a semi-quantitative analysis were then performed using spectra recorded on preparations of chlorite holocellulose and acid-insoluble lignin (Figure 6-1A, B). 129 Leatherwood holocellulose produced a stronger signal at 1740 cm-1 corresponding to the C=O stretching mode of esters and unconjugated ketones compared to aspen and spruce, pointing to higher levels of wall-bound acetate or uronic acids in leatherwood (Figure 6-1C; see Table A2-1 for annotations of FTIR wavenumbers). In addition, leatherwood gave a considerably weaker response at the 1420 cm-1 wavenumber, which has been ascribed to the CH2 bending mode of cellulose, suggesting that leatherwood may have lower cell wall crystallinity or less cellulose. Curve fitting of the lignin spectra revealed weaker signals for leatherwood at 1510 and 1270 cm-1 compared with spruce, and a stronger response at 1330 cm-1 compared with aspen (Figure 6-1D). The 1510 cm-1 wavenumber corresponds to aromatic skeletal vibrations in lignin, whereas the 1270 and 1330 cm-1 bands originate from the ring-breathing and C–O stretching modes of guaiacyl and syringyl lignin, respectively, suggesting that leatherwood xylem may have less lignin and a higher ratio of syringyl: guaiacyl lignin units. 6.4.1 Lignin content and composition To corroborate the FTIR results, we measured the Klason lignin content by two-stage acid hydrolysis and analysed the lignin composition using thioacidolysis (Table 6-1). In this analysis, leatherwood had less than half as much acid-insoluble lignin as spruce and 37% less than aspen. On the other hand, leatherwood produced more than 12 times as much acid-soluble lignin as spruce and 33% more than aspen. In addition, p-hydroxyphenyl units were absent in leatherwood, and the lignin was found to contain 80% syringyl units compared with 71% in aspen. 6.4.2 Structural polysaccharides On account of the comparatively lower lignin content of leatherwood xylem, we considered whether the structural polysaccharides might also be altered (Table 6-1). This analysis revealed that leatherwood has more xylose (39% more than aspen, 322% more than spruce) and contains more rhamnose (70% more than aspen, 329% more than spruce), but has less glucose (24% less than aspen, 18% less than spruce) and mannose (67% less than aspen, 93% less than spruce). Leatherwood also has more galactose and arabinose than aspen (68% and 305%, respectively) and less galactose than spruce (52%). Furthermore, alkaline 130 Figure 6-1. Semi-quantitative FTIR analysis of leatherwood, aspen and spruce. Peak fitting of spectra from (A) chlorite holocellulose and (B) acid-insoluble lignin derived from leatherwood (blue), aspen (green), and spruce (red) xylem with deconvoluted peaks shown only for leatherwood (black). Responses for several wavenumbers are plotted by normalised unit area for (C) chlorite holocellulose and (D) acid-insoluble lignin. Three replicates were analysed for each species and sample type, and the error bars show standard deviation. No fitted peak was obtained for spruce at 1330 cm-1. hydrolysis of cell-wall-bound acetate revealed that leatherwood xylem contains 25% more acetyl groups than aspen and four times more than spruce. Leatherwood has twice as much glucuronic acid as aspen and spruce, and about 1.5 times as much 4-O-methyl glucuronic acid (Table 6-1). In addition, leatherwood contains 2.6 times more galacturonic acid than aspen and 3.3 times more than spruce, pointing to higher levels of pectin in leatherwood xylem. It should be emphasised that these values are almost certainly underestimates, as uronic acids are not completely liberated during sulphuric acid hydrolysis (Willför et al., 2009). The alpha cellulose procedure was used to quantify the amount of cellulose, and X-ray diffraction and differential interference contrast microscopy were used to evaluate the physical properties of cellulose microfibrils (Figure A2-3; Table 6-1; section A2.1.3). The cellulose content of leatherwood was found to be 14% less than aspen and about 11% less than spruce, and the crystallinity index was 11% lower than 131 Table 6-1. Cell wall analysis of leatherwood, aspen and spruce. Cell wall properties Leatherwood Aspen Spruce Lignin content (% w/w) Acid-insoluble lignin 11.30 ± 0.19 17.96 ± 0.31 28.54 ± 1.30 Acid-soluble lignin 5.24 ± 0.23 3.92 ± 0.06 0.43 ± 0.02 Total lignin content 16.61 ± 0.25 21.91 ± 0.30 28.96 ± 1.32 Lignin composition (% w/w) p-Hydroxyphenyl Not detected 0.53 ± 0.11 2.38 ± 0.35 Guaiacyl 19.53 ± 1.11 28.65 ± 0.38 97.62 ± 0.35 Syringyl 80.47 ± 1.11 70.82 ± 0.27 Not detected Structural polysaccharides (% w/w) Glucose 35.00 ± 0.32 45.87 ± 0.36 42.84 ± 1.77 Xylose 23.19 ± 0.37 16.69 ± 0.42 7.19 ± 0.12 Mannose 0.66 ± 0.03 2.01 ± 0.08 10.15 ± 0.56 Galactose 1.14 ± 0.05 0.68 ± 0.04 2.37 ± 0.50 Arabinose 1.60 ± 0.10 0.52 ± 0.04 1.61 ± 0.15 Rhamnose 0.84 ± 0.04 0.49 ± 0.03 0.25 ± 0.05 Galacturonic acid 3.52 ± 0.13 1.37 ± 0.06 1.06 ± 0.19 Glucuronic acid 0.77 ± 0.11 0.38 ± 0.03 0.36 ± 0.07 4-O-Methyl glucuronic acid 2.59 ± 0.05 1.65 ± 0.04 1.36 ± 0.06 Other cell wall properties Cellulose content (% w/w) 31.44 ± 0.66 36.61 ± 2.42 35.49 ± 0.54 Ash content (% w/w) 1.20 ± 0.18 0.44 ± 0.07 0.19 ± 0.08 Wall-bound acetyl (% w/w) 5.74 ± 0.13 4.58 ± 0.04 1.29 ± 0.05 Crystallinity index 0.31 ± 0.01 0.34 ± 0.02 0.37 ± 0.02 Microfibril angle (°) 57.9 ± 1.9 32.1 ± 5.4 39.5 ± 1.9 Three biological replicates were analysed in triplicate and standard deviations are shown alongside mean values. aspen and 16% lower than spruce. On the other hand, leatherwood xylem showed substantially larger microfibril angles than the other species. 6.4.3 Cell wall histochemistry We next set out to dissect the histology of leatherwood xylem in light of its notable cell wall composition. In addition to the unlignified ray parenchyma cells, astra blue staining showed concentric rings of unlignified axial parenchyma laid down at the end of each growth cycle (Figure 6-2A, B). Safranin-O uncovered lignin-rich vessel elements in oblique strands amongst smaller thick-walled tracheids forming an unusual dendritic pattern within a field of libriform fibres. For comparison, staining of aspen showed small clusters of vessels arranged diffusely amidst fibres and ray parenchyma (Figure 6-2C), whereas tracheids and rays predominated in spruce xylem (Figure 6-2D). 132 Figure 6-2. Lignin histochemical analysis of leatherwood, aspen and spruce. Transverse cross-sections of leatherwood, aspen, and spruce xylem stained with (A–D) astra blue and safranin-O, (E–H) by the Mäule reaction, and (I–L) with phloroglucinol–hydrochloric acid. Images show vessel elements (V), fibres (F), ray parenchyma (R), axial parenchyma (A), and tracheids (T). Two magnifications are shown for leatherwood. Bars: (A, E, I) 200 µm; (B–D, F–H, J–L) 10 µm. To ascertain whether lignin composition varies in a cell-type specific manner in leatherwood, we examined cross-sections using the syringyl lignin-specific Mäule reaction. Axial and ray parenchyma remained unstained, whereas fibres, vessels, and vasicentric tracheids appeared to be stained equally (Figure 6-2E, F). Staining of vessels in aspen was weaker than fibres, indicating less syringyl lignin, as expected (Figure 6-2G), and spruce cell walls were not stained due to the absence of syringyl lignin in gymnosperms (Figure 6-2H). Staining with phloroglucinol was confined primarily to the strands of vessels and vasicentric tracheids 133 in leatherwood (Figure 6-2I, J), suggesting a difference in the composition of lignin associated aldehydes between the walls of fibres and those of vessels and vasicentric tracheids. No such patterns were observed in aspen or spruce, but the overall level of staining was more intense, reflecting the inherently higher lignin concentrations (Figure 6-2K, L). 6.4.4 Label-free chemical imaging To further resolve the cell-type-specific differences in cell wall composition, high spatial-resolution Raman imaging was performed on leatherwood xylem cross-sections. The signal at 1600 cm-1 corresponds to aromatic skeletal vibrations in lignin and can be used to examine the distribution of lignin (Gierlinger et al., 2012). Integrating this band revealed stronger signals in vessels and vasicentric tracheids, pointing to a difference in lignin content and/or composition (Figure 6-3A). An even more drastic difference was seen with the guaiacyl lignin-sensitive band centred around 1273 cm-1 (Agarwal et al., 2011), suggesting that vessels and vasicentric tracheids have more lignin and particularly more guaiacyl units (Figure 6-3B). The signal at 856 cm-1, corresponding to C6–C5–O5–C1–O1 backbone vibrations of pectin (Synytsya et al., 2003), was confined to cell corners and middle lamellae, and to the walls of ray and axial parenchyma (Figure 6-3C). Finally, the band at 1738 cm-1, ascribed to C=O stretching of ester and acetyl groups (Agarwal et al., 2011), was most prominent in the cell walls of vasicentric tracheids, but was also present in vessels and fibres (Figure 6-3D). In order to delve deeper into the spectral differences between cell types in leatherwood xylem, we performed a cluster analysis on the Raman hyperspectral dataset and identified three pixel clusters corresponding to the cell walls of vessels, fibres and vasicentric tracheids, as well as ray and axial parenchyma (Figure 6-3E). Examination of the corresponding spectra (Figure 6-3F) revealed that parenchyma differ broadly from vessels, tracheids and fibres having less of the acetyl (1735 cm-1) and lignin-specific signals (1600, 1335, 1273 cm-1), but more of the pectin-specific band (856 cm-1). This analysis also confirmed that vessels have more lignin than fibres and tracheids due to a much stronger response near 1600 cm-1. Comparing these spectra also showed that vessel cell walls have more guaiacyl lignin due to a greater response at 1273 cm-1 and, in accordance with the phloroglucinol staining results, 134 more aldehyde groups revealed by the shoulder at 1620 cm-1, which corresponds to the C=C stretching mode of aldehydes (Figure 6-3F inset; Agarwal et al., 2011). Figure 6-3. Raman imaging of leatherwood. Transverse cross-sections of leatherwood xylem imaged by confocal Raman microscopy. Chemical images are shown for specific integrated bands corresponding to (A) total lignin (1600 cm-1), (B) guaiacyl lignin (1273 cm-1), (C) pectin (856 cm-1), and (D) acetyl (1738 cm-1). (E) A false-colour image and (F) corresponding Raman spectra are shown from the cluster analysis to highlight the spectral differences between vessels and vasicentric tracheids (blue), fibres (red), and parenchyma (green). The inset in (F) shows zoomed-in spectra normalised at 2920 cm-1. Images show vessel elements (V), fibres (F), ray parenchyma (R), axial parenchyma (A), and tracheids (T). Bars, 50 µm. 135 6.4.5 Lignin-deficient middle lamellae Perhaps most striking from the Raman analysis was the absence of all lignin-related signals in the cell corners and middle lamellae (Figures 6-3A, B; Figure A2-4B). Debonding of cell walls was also frequently observed while sectioning leatherwood xylem (Figure 6-4A). To analyse this further, we examined the ultraviolet autofluorescence of lignin across adjacent cell walls and compound middle lamellae of neighbouring cells in leatherwood xylem and observed a drop in the lignin content within the middle lamella between vessels, fibres, and, somewhat less consistently, between vasicentric tracheids (Figure 6-4B). By contrast, lignin autofluorescence peaked within the middle lamella between vessels and fibres in aspen (Figure 6-4E, F) and between tracheids in spruce (Figure 6-4I, J). Similarly, a weaker response of the lignin-specific 1600 cm-1 Raman signal was observed in middle lamellae between leatherwood fibres, vessels, and tracheids (Figure 6-4C). As expected, aspen and spruce again showed the strongest lignin signal in the middle lamella (Figure 6-4G, K). ToF-SIMS imaging was also used to confirm that leatherwood possesses lignin-deficient middle lamellae. Although all three species showed characteristic signals for structural polysaccharides in the cell wall, only aspen and spruce exhibited lignin-related peaks in the middle lamella and cell corners (Figure 6-4D, H, L). Finally, examination of leatherwood samples collected from across the range of D. palustris as well as the three congener leatherwood species demonstrated that lignin-deficient middle lamellae is a trait that is conserved throughout the genus Dirca (Figure A2-5; section A2.1.5). 6.5 Discussion Although lignin biogenesis has been studied extensively and great strides have been made, gaps remain in our understanding of this complex biological process. Fundamental questions persist, including: What is the role of lignin in the middle lamella? Is there a functional significance of lignin structural diversity? What are the evolutionary origins of lignification? And, how is lignification initiated in the first place? To address these and other ongoing topics in lignin biology, new models are sorely needed (Neutelings, 2011). 136 Figure 6-4. Evidence of lignin-deficient middle lamellae in leatherwood. Transverse cross-sections of (A–D) leatherwood, (E–H) aspen, and (I–L) spruce xylem viewed by (A, E, I) ultraviolet (UV) microscopy, (C, G, K) confocal Raman imaging, and (D, H, L) time-of-flight secondary ion mass spectrometry (ToF-SIMS). Pointwise signal intensity of lignin autofluorescence is shown (B, F, J) for line traces (A, E, I) across adjacent cell walls of fibres (red), vessels (green), and tracheids (blue). The orange arrowheads (A) point to debonding of adjacent cells. Raman images show integration at the lignin-specific 1600 cm-1 wavenumber, and false-colour images for ToF-SIMS show the distribution of ions corresponding to lignin and structural polysaccharides. Bars, 10 µm. 137 6.5.1 Unusual cell wall composition and lignin distribution In this study, we have shown that leatherwood, a slow-growing woody shrub, contains conspicuously low levels of lignin and displays a unique distribution of lignin throughout the xylem and across the cell wall. Compared with other woody angiosperms reported in the literature, leatherwood is among the lowest in terms of lignin content (Pettersen, 1984). This is consistent with previous reports that senescent leatherwood leaves contain less lignin than all other shrubs and trees examined (Ferrari, 1999; Jo et al., 2016). Leatherwood is a member of the family Thymelaeaceae in the order Malvales, a diverse group of trees, shrubs, and lianas, but there is no evidence that poor lignification is a common feature of this family. For example, the Klason lignin contents of Aquilaria, Gonystylus, and Daphne spp. range from 23% to 31% (Syafii & Yoshimoto, 1991; Yamamoto & Hong, 1994; Shrestha & Budhathoki, 2012). Nevertheless, there are family resemblances in histology, including the presence of axial parenchyma and vasicentric tracheids in the xylem and thick bast fibres in the bark (Herber, 2003; Schweingruber et al., 2011; Figure A2-6). Leatherwood xylem contains strands of highly lignified thick-walled vessels and vasicentric tracheids set amongst thin-walled fibres that are only weakly lignified (Choquette, 1925). Using histochemical staining and Raman imaging, we found that leatherwood primarily lignifies only the water-conducting vessels and tracheids, and these cell walls also contain higher levels of guaiacyl lignin and more lignin-associated aldehyde groups compared to fibres. Such variability in lignin composition may be tailored to differences in cell function, as vessels are specialised for water conduction whereas fibres primarily confer mechanical support (Donaldson, 2001; Pereira et al., 2018). If so, leatherwood xylem may represent an extreme example of cell-type-specific lignin specialisation, and further investigations could help elucidate the functional significance of lignin structural diversity. We also found that leatherwood produces a syringyl-rich lignin and greater levels of cell-wall-bound acetyl, two important targets for the improvement of biomass feedstocks in the pulping and biofuel industries. Syringyl lignin content is positively correlated to wood pulpability and biomass pretreatability (Huntley et al., 2003; Mansfield et al., 2012), whereas acetyl levels affect carbohydrate recovery in 138 pretreatment (Johnson et al., 2017). Moreover, acetyl decorations impact xylan deposition and can influence interactions between lignin and other cell wall constituents (Grantham et al., 2017). In this way, acetylation in leatherwood could help provide structural compensation for the lower overall levels of lignin. Raman imaging showed that vasicentric tracheids contain the highest concentration of acetyl groups, perhaps relating to the strengthening role of acetylation in these cell walls (Pawar et al., 2013). 6.5.2 Wood flexibility and lignin-deficient middle lamellae The source of leatherwood’s flexibility has remained an open question. Although poor lignification is thought to play a role (Nevling, 1962), others have pointed to the arrangement of vessels (McMinn & Forderhase, 1935, in D. occidentalis) and to the shorter fibres, thinner cell walls, and larger cell lumina (Potzger & Geisler, 1937). To this list we can add lower cell wall crystallinity and larger microfibril angles. The largest microfibril angles in leatherwood occur in the walls of vessels (Figure A2-4; section A2.1.4), possibly because these cells must bear the stresses of water conduction. Although all the aforementioned factors likely contribute to stem flexibility, the occurrence of lignin-deficient middle lamellae offers the most compelling hypothesis yet. We interrogated the lignin distribution across the compound middle lamella using three analytical techniques and consistently found lignin-rich middle lamellae in aspen and spruce, but not in leatherwood. Without the rigidity conferred by lignified middle lamellae, it could be that adjacent cells in leatherwood xylem remain free to slip past one another slightly, thereby providing some flexibility to the woody stems. It has long been known that the cell corners and middle lamellae of mature xylem normally contain the highest concentrations of lignin, accounting for 16–35% of all the lignin in wood (Ritter, 1925; Fergus et al., 1969; Saka & Goring, 1988). And although some heterogeneity in middle lamellar lignin has been reported (Hardell et al., 1980; Singh & Schmitt, 2000; Wi et al., 2005), leatherwood represents the first known example of such a widespread lignin deficiency in the middle lamella. 6.5.3 Challenging the model of lignification Reports from diverse plant species have shown that lignification begins in the cell corners and middle lamellae, and then spreads inward to the layers of the cell wall (Donaldson, 2001). Accordingly, it has 139 been hypothesised that the onset of lignin polymerisation occurs in these regions (Terashima, 1990; Donaldson, 1994). Putative nucleation sites for lignin initiation include cell-wall-bound hydroxycinnamates, pectin, and cell wall proteins (Ralph et al., 1995; Boerjan et al., 2003). Similarly, laccase or peroxidase enzymes that drive lignin polymerisation may be specifically tethered to the cell corners and middle lamellae (Boerjan et al., 2003; Chou et al., 2018; Tobimatsu & Schuetz, 2019), but irrefutable evidence for any one mechanism is still lacking. Moreover, previously observed differences in lignin composition between middle lamellae and adjacent cell walls (Whiting & Goring, 1982) are still unexplained. With cell corners and middle lamellae that remain unlignified in mature xylem, it could be that leatherwood represents an exception to the rule or it may be the case that debunks the model. Accordingly, the lignin distribution should be re-examined in diverse plant taxa to quell this uncertainty. Early in the development of xylem tissue, protoxylem cells are formed with bands of lignified secondary cell wall interspersed by regions of primary wall that remain unlignified. This arrangement, which provides axial flexibility for these rapidly growing cells while enabling water conduction, is guided by the deployment of oxidative enzymes specifically to the regions of secondary cell wall (Schuetz et al., 2014). Lateral flexibility in protoxylem is afforded by unlignified middle lamellae, while pectin and cell wall proteins provide cell adhesion (Ryser et al., 2004). In leatherwood, lignin-deficient middle lamellae may also confer flexibility, and perhaps this also results from a localised absence of laccase or peroxidase enzymes. Alternatively, the monolignols needed for lignification may not be efficiently delivered to the middle lamella, initiation sites may be lacking in leatherwood, or perhaps radical formation and polymer growth are inhibited by some other means. It is also notable that leatherwood lacks p-hydroxyphenyl moieties, since these lignin units typically predominate in the middle lamella (Whiting & Goring, 1982; Terashima et al., 2012). As the mechanisms undergirding lignin initiation and deposition remain obscure, further studies on the topochemistry of lignification in leatherwood may help elucidate these dynamics and could inform revisions to the classical model. 140 6.5.4 Implications of lignin-deficient middle lamellae In most plants, lignin-rich middle lamellae help adjacent xylem cells adhere to prevent slippage. By contrast, leatherwood xylem retains the pectin-rich middle lamellae and cell corners normally found between unlignified primary walls. While secondary cell wall lignin contributes to compressive strength by increasing the hardness properties of the wall (Niklas, 1992; Gindl et al., 2002), the mechanical role of lignin in the middle lamella is less clear. Compared with secondary walls, lignified middle lamellae have a lower Young’s modulus, a similar hardness, and a greater shear strength (Niklas, 1992; Qin et al., 2018), possibly due to the globular macromolecular structures of lignin formed in the absence of cellulose microfibrils (Terashima et al., 2012).Without heavily lignified middle lamellae, leatherwood could be susceptible to xylem fracture. In this light, the clustering of strongly lignified vessels and vasicentric tracheids into oblique strands may be a compensatory adaptation that imparts the requisite shear support with minimal investment in lignin. Similarly, the thick and flexible bark with its strong bast fibres could provide mechanical compensation for a weaker xylem (Holm, 1921). In any case, lignin-deficient middle lamellae could lead to greater debonding of xylem cells during mechanical shearing (Donaldson, 1995), a highly desirable trait in the processing of woody biomass. As poor lignification presents a challenge for defence against pest and pathogen, Dirca spp. invest in a wide range of defensive compounds, including diterpenes, lignans, coumarins, phenolic glycosides, and a novel class of sulphur-containing cytotoxins called dirchromones (Badawi et al., 1983; Ramsewak et al., 1999; St-Gelais et al., 2015). The chemical defence strategy is also used by Aquilaria and Gyrinops spp., other members of the family Thymelaeaceae whose axial parenchyma produce similar compounds when exposed to wood-decay microorganisms, resulting in a valuable aromatic xylem called agarwood (Rasool & Mohamed, 2016). A lignin-deficiency in the middle lamella could also impact the transport of water and dissolved nutrients through the xylem. Lignin and pectin are highly co-localised within the middle lamella (Wi et al., 2005; Kim et al., 2015), and this lignin has been negatively correlated to ion-mediated changes in xylem conductance (Boyce et al., 2004). In this way, lignification of middle lamellae represents a trade-off between mechanical support and reducing conductance by restricting the accessibility of pectin. 141 It has been reported that the bordered pits of leatherwood fibres have vestures, while those of vessels and vasicentric tracheids are rarely vestured (Jansen et al., 2000; Dute et al., 2001). As these lignin-rich protrusions may be involved in regulating water flow and safeguarding against embolisms (Choat et al., 2008), further studies on leatherwood xylem conductance and safety could help elucidate the role of lignin in these important processes. It remains unclear what selective pressures led leatherwood to develop lignin-deficient middle lamellae, and it is currently unknown whether this phenomenon occurs in related taxa beyond the genus Dirca (Figure A2-5). Raman imaging of Daphne odora did not uncover similarly unlignified middle lamellae (Zhang et al., 2012), suggesting that leatherwood may be unusual even among the Thymelaeaceae. As lignin represents a substantial investment of photosynthate, this atypical lignification regime undoubtedly imparts considerable energy savings. Although it is not directly useful as a biomass feedstock, leatherwood may soon be enlisted to address ongoing questions in lignin biology and inspire new strategies in lignin engineering. 142 Chapter 7: Conclusions 7.1 Summary of major research contributions An expansive volume of studies with diverse plant taxa have examined the effects on lignin content, composition, and structure resulting from perturbation of genes in the core monolignol biosynthetic pathway (Campbell & Sederoff, 1996; Boerjan et al., 2003; Bonawitz & Chapple, 2010; Vanholme et al., 2012; Wang et al., 2014; Barros et al., 2015; Liu et al., 2018). Although lignification is highly flexible (see Chapter 2) and minor changes are readily tolerated, significant decreases in lignin content often result in stunted or stressed plants with compromised water transport systems (Anterola & Lewis, 2002; Coleman et al., 2008a; Voelker et al., 2011; Bonawitz & Chapple, 2013). The accumulation of pathway intermediates may also contribute to these phenotypes since carbon flux which passes beyond phenylalanine ammonia lyase (PAL) is committed to the formation of phenolic secondary metabolites that could cause pleiotropic effects. In this work, I targeted chorismate, a key branchpoint intermediate upstream of PAL and phenylpropanoid biosynthesis, to elucidate the role of chorismate mutase (CM) in regulating carbon allocation towards the biosynthesis of lignin and phenylpropanoids (see Chapter 3). This revealed an apparent and unexpected functional redundancy in the CM gene family of poplar. It also pointed to an under-appreciated link between the biosynthesis of lignin and the formation of salicylic acid (and derivatives thereof), an important regulator of diverse metabolic and developmental processes in plants. The observation that chorismate is an important branchpoint metabolite upstream of lignin biosynthesis also helped inspire a new strategy in lignin engineering. I transferred the chorismate pyruvate lyase (CPL) gene from E. coli (Siebert et al., 1994) into transgenic poplar in order to divert carbon away from lignin biosynthesis and convert chorismate into p-hydroxybenzoate (see Chapter 4). Lignin from poplar and willow is naturally decorated with ester-linked p-hydroxybenzoate (pHB) groups due to the participation of acylated monolignols in lignification (Smith, 1955b; Pearl et al., 1957; Lu et al., 2004; Morreel et al., 2004). Heterologous expression of bacterial CPL in poplar lead to increased p-hydroxybenzoylation of lignin which was accompanied by a small decrease in lignin content. An equally large pool was retained 143 as soluble metabolites including glucosides of p-hydroxybenzoate as well p-hydroxybenzoate-containing analogues of well-known poplar metabolites. Furthermore, I also developed a new mass spectrometry-based imaging technique which was used to demonstrate that pHB groups are largely restricted to the cell walls of fibres, both in wild-type trees and also in CPL-expressing poplars (see Appendix 3 and Chapter 4). As pHB groups are ester-linked to lignin polymers, they can be cleaved off using mild alkaline hydrolysis and subsequently upgraded, inter alia, into high-value biochemicals and biomaterials. Accordingly, such lignin acylations are eminently worthy research targets and this work demonstrates the effectiveness of CPL in the metabolic engineering of lignin for this purpose. pHB groups are not the only lignin acylations that occur in Nature. For example, the lignin of kenaf (Hibiscus cannabinus) bast fibres is highly acetylated (Ralph, 1996; Lu & Ralph, 2002). Similarly, p-coumarate groups decorate the lignin of commelinid monocots (Harris & Hartley, 1980; Karlen et al., 2018). I showed that kenaf core tissues contain lignin with γ-esterified p-coumarate, thereby providing evidence that these groups are not phylogenetically restricted to monocot lineages (see Chapter 5). Moreover, heterologous expression of a BAHD acyltransferase from kenaf led to novel p-coumaroylation of lignin in poplar. This enzyme, HcPMT, shares less than 30% amino acid identity with the reported monocot PMTs. Finally, the introduction of p-coumarate groups into poplar lignin did not lead to improved rates of enzymatic saccharification of wood powder. This work offers yet another example of convergent evolution in plant cell wall biosynthesis, and provides a new molecular tool with proven utility in lignin engineering. Eastern leatherwood (Dirca palustris) is an uncommon woody shrub that has largely escaped the attention of plant biologists. Some excellent work has been done in recent years on the ecology, distribution, and horticultural potential of this species (Schrader & Graves, 2005; Williams, 2009; Peterson & Graves, 2009; Floden, 2011; Peterson & Graves, 2016), but the only studies that examined the woody stems of leatherwood were published in the 1920s and 30s and were largely descriptive in nature (Holm, 1921; Choquette, 1925; McMinn & Forderhase, 1935; Potzger & Geisler, 1937). Using various analytical techniques and several complementary imaging tools, I uncovered evidence that leatherwood has evolved an unusual pattern of lignification (see Chapter 6). Lignin in leatherwood stems is 144 predominately deposited in the walls of water-conducting cells whereas the xylem fibres that make up the bulk of wood are only weakly lignified. I also conducted a thorough analysis of the chemical composition of the wood, and found that leatherwood stems have a relatively low lignin content but a high proportion of S-lignin units. Finally, close examination of the cell walls revealed that leatherwood has pectin-rich middle lamellae that appear to be largely devoid of lignin. These findings may help explain the flexibility of leatherwood stems, but they also draw attention to several unresolved questions in lignin biology. 7.2 Diverting carbon flux away from lignin biosynthesis Based on the reported gene expression profiles of CM in poplar stems (Tsai et al., 2006), it was reasonable to predict that RNAi-mediated suppression of PtCM1 would result in decreased carbon flux towards monolignol biosynthesis. Accordingly, the observed increase in lignin content reported in Chapter 3 was both surprising and highly informative. These results showed that the other two CM genes can be upregulated in a compensatory manner, and in so doing revealed apparent gene redundancy in the poplar CM gene family. I have also generated RNA interference (RNAi) poplar lines downregulated in the other two CM genes; however, these have not yet been fully analysed. It will be very interesting to examine what, if any, are the effects on plant growth and metabolism, and whether compensatory expression can be reciprocal. Specifically, is PtCM1 upregulated in response to suppression of either PtCM2 or PtCM3? One possible future experiment could be to use tissue-specific rather than constitutive promoters to drive expression of the RNAi constructs in order to minimise confounding effects and to provide more fine-scale information on gene redundancy. Unfortunately, RNAi approaches can suffer from various limitations including diminishing expression of the RNAi hairpin construct due to transgene silencing (Small, 2007). Beyond analysing additional RNAi lines for PtCM2 and PtCM3, one approach would be to generate knock-out mutants of all three CM genes in poplar, both individually and in combination. This could conceivably be achieved using CRISPR/Cas9 gene editing techniques which have been successfully deployed in poplar (Zhou et al., 2015; Fan et al., 2015; Bruegmann et al., 2019). More broadly, the study of PtCM1 highlighted the importance of chorismate as a key branchpoint metabolite and showed that genes upstream of phenylpropanoid biosynthesis could be useful targets in 145 lignin engineering. Of course, this study was not the first to show that lignin content, composition, and structure can be affected by upstream perturbations. For example, heterologous expression of the bacterial 3-dehydroshikimate dehydratase in Arabidopsis led to changes in lignin content, composition, and structure (Eudes et al., 2015). As such, it is plausible that other metabolites in the shikimate and aromatic amino acid pathways could be useful future targets to perturb carbon flux destined for lignin biosynthesis. It may even be possible to target metabolic steps even further upstream. It has long been known that phenylpropanoid biosynthesis is tightly controlled by the supply of photosynthate. Specifically, erythrose 4-phosphate, one of the key substrates of the shikimate pathway, is produced from fructose 6-phosphate by transketolase in the Calvin cycle or from sedoheptulose 7-phosphate by transaldolase in the oxidative pentose phosphate pathway (Maeda & Dudareva, 2012). Antisense suppression of transketolase in tobacco resulted in a significant decrease in the production of aromatic amino acids, phenylpropanoids, and lignin (Henkes et al., 2001), and a transaldolase knockout mutant of Arabidopsis showed reduced lignin levels (Vanholme et al., 2012b). These enzymes represent an important albeit under-appreciated link between carbohydrate metabolism and the production of lignin and other phenylpropanoids (Pinard et al., 2019). Recent work points to the importance of regulation at the entry point of the shikimate pathway (Yokoyama et al., 2021), so this may soon prove to be an exceptionally useful target in lignin engineering. 7.3 Lignin acylations, monolignol transferases, and designer lignins It has long been recognised that pHB groups occur naturally as γ-linked lignin acylations in poplar and willow (Smith, 1955b; Pearl et al., 1957). Despite the potential industrial significance of these ester-linked phenolics, no attempts to engineer the levels of pHB groups have previously been reported. However, ongoing work has sought to identify BAHD acyltransferase(s) involved in the synthesis of p-hydroxybenzoylated monolignol conjugates. In Chapter 4, I reported on a novel strategy to engineer pHB levels and simultaneously divert carbon flux away from the phenylpropanoid pathway. Although this was successful, a considerable pool of p-hydroxybenzoate (pHBA) accumulated as soluble metabolites in the xylem. Several candidate pHBA-CoA:monolignol transferases have recently been identified, and gene stacking experiments with CPL are ongoing. It will be very exciting to see whether pHB levels are 146 significantly improved in these forthcoming transgenic poplar lines. At present, a CesA promoter has been used to drive expression of the CPL transgene, but it may also be prudent to experiment with other promoters, including those from lignin biosynthetic genes, in order to maximise pHB levels. Various other complementary strategies should also be contemplated. Greater production of pHBA could perhaps be achieved through enhanced expression of the transgene. Most previous reports on heterologous expression of CPL in planta used plastid localisation signals; however, substantially greater activity has been achieved with biolistics-mediated plastid transformation (Viitanen et al., 2004). It could also be that pHBA is not efficiently delivered from the plastid to the cytosol where monolignol conjugates are presumably produced. One approach to remedy this would be the introduction of pHBA transporters such as those previously identified in Pseudomonas and Acinetobacter spp. (Harwood et al., 1994; Pernstich et al., 2014). And since CPL may be feedback inhibited (Siebert et al., 1994), improved export of the product from plastids might also enhance conversion rates. Another potential rate-limiting step could be the activation of pHBA into the CoA thioester. This reaction is likely catalysed by one of the endogenous 4-coumarate:CoA ligase (4CL) or 4CL-like enzymes that are known to accept diverse phenolic substrates (Cukovica et al., 2001), but this activity has not yet been reported in poplar. Introduction of a CoA ligase that is selective for pHBA, such as those reported in Rhodopseudomonas palustris (Merkel et al., 1989) and more recently in hornwort (Wohl & Petersen, 2020), could help drive the production of monolignol–pHB conjugates. In Chapter 5, I reported on the characterisation of a novel BAHD acyltransferase from kenaf which showed activity as a p-coumaroyl-CoA:monolignol transferase (PMT). Initially, the goal was to find an acetyl-CoA:monolignol transferase since kenaf is well-known for having extensively acetylated lignin (Ralph, 1996). Although HcPMT does accept acetyl-CoA as a substrate in vitro (Peers, 2012), acetylated monolignols are evidently not produced in planta. The discovery that this new BAHD acyltransferase functions as a PMT enzyme was particularly surprising since p-coumaroylated lignin has traditionally only been considered a feature of the cell walls of commelinid monocots (Karlen et al., 2018). From a phylogenetic analysis of related BAHD sequences, it appears that HcPMT arose independently from monocot PMTs. 147 Although this new acyltransferase could prove to be very useful in future lignin engineering work, there is still value in searching for acetyltransferases. Kenaf remains one of the best candidates for this work; however, other species with acetylated lignin such as sisal, abaca, and hornbeam may also be useful (del Río et al., 2007). It could also be worthwhile characterising additional dicot PMTs and searching for p-coumaroylation of lignin in these taxa as this would help validate the hypothesis that convergent evolution explains the occurrence of p-coumaroylated lignin among the dicots. Several BAHD enzymes with activity towards monolignols have been identified and characterised in recent years (Withers et al., 2012; Petrik et al., 2014; Marita et al., 2014; Wilkerson et al., 2014; Karlen et al., 2018). It would now be very exciting to employ protein engineering techniques to ‘evolve’ monolignol acyltransferases with novel activities. In this way, new acylations not observed in Nature could be introduced into lignins. Although random mutagenesis and high-throughput screening approaches could be useful for this, the availability of crystal structural models for monolignol acyltransferases would enable rational design strategies and accelerate work towards these goals. Of course, pHB and p-coumarate are not the only possible ester-linked pendent groups. Monolignol conjugates of ferulate, for example, occur naturally in many plant species and poplars engineered with increased levels exhibited improved saccharification (Wilkerson et al., 2014; Kim et al., 2017; Zhuo et al., 2017; Bhalla et al., 2018). Recently, benzoate has also been identified in the lignin of poplars deficient in cinnamate 4-hydroxylase and p-coumaroyl-CoA 3'-hydroxylase due to a shift in phenylpropanoid metabolism and the action of an endogenous monolignol transferase (Kim et al., 2020). It is conceivable that further hydroxylations of the phenolic ring will be compatible with lignification such that 3,4-dihydroxybenzoate (protocatechuate) and 3,4,5-trihydroxybenzoate (gallate) could be worthwhile future targets in lignin engineering. These compounds could be produced through hydroxylations, for example using a pHBA hydroxylase such as those from Pseudomonas spp. (Hosokawa & Stanier, 1966). Alternatively, the bacterial 3-dehydroshikimate dehydratase enzyme has previously been used to divert carbon flux from the shikimate pathway and make 3,4-dihydroxybenzoate directly (Eudes et al., 2015). In recent years, it has become apparent that the plasticity of lignification extends far beyond the core monolignols, acylated monolignol conjugates, and even beyond the phenylpropanoid pathway itself (see 148 Chapter 2). For example, the flavonoid tricin, the hydroxystilbenes resveratrol, isorhapontigenin, and piceatannol, and the phenolic amide diferuloylputrescine have all been identified as bone fide naturally-occurring lignin monomers in plants (Lan et al., 2015; del Río et al., 2017; del Río et al., 2018). Various research groups are currently working to transfer such lignin chemistry into biomass feedstock species and maximise the level of these value-added designer lignins. Without a doubt, additional non-canonical lignin monomers will be discovered in the coming years and researchers may even soon succeed at introducing monomers that are not naturally occurring in lignin, such as the recently reported attempts to incorporate the diphenylheptanoid curcumin into poplar lignin (Oyarce et al., 2019). 7.4 Unresolved questions in lignin biology Despite ample evidence that lignification is a highly dynamic and flexible process, it remains entirely unknown what, if any, biological consequences result from different lignin compositions and structures. For example, lignin deposited in response to stress is rich in H-lignin units (Cesarino, 2019). Also, it has long been known that the cell walls of poplar vessel elements are rich in G-lignin relative to xylem fibres (Musha & Goring, 1975). Similarly, in Appendix 3, I reported compelling evidence that p-hydroxybenzoate pendent groups occur predominantly, if not exclusively, in the xylem fibres of poplar and willow. And in Chapter 6, I described differences in S-lignin as well as lignin-associated aldehydes across the different xylem cell types in eastern leatherwood (Dirca palustris). One of the big challenges in lignin reserch, and particularly for those interested in exploiting lignin plasticity for engineering goals, will be to tease apart the biological function(s) of different lignin polymers. It could be that the resulting changes in the physicochemical properties can engender improvements in strength parameters, greater hydrophobicity, or enhanced resistance to chemical or biological degradation, for example. Alternatively, it could be that lignin variations are entirely inconsequential and merely a reflection of differences in metabolism. As such, further investigations on the mechanical, chemical, and biological properties of different lignin polymers are sorely needed. In Chapter 6, I reported on the atypical pattern of lignification that occurs in the stems of leatherwood. Although this discovery was quite interesting and unexpected, in many ways this work posed more 149 questions than it answered. Chiefly among these, why has leatherwood evolved such an unusual lignification pattern? To address this question, it could be helpful to determine when this trait appeared and whether leatherwood is simply one example among many others. A very similar distribution of lignin was apparent when I examined stems from other members of the genus, namely western leatherwood, Mexican leatherwood, and deceptive leatherwood (see Appendix 2). It would be worthwhile expanding upon this analysis to include other genera in the family Thymelaeaceae. In fact, I have already analysed representative stem samples from a dozen taxa and, thus far, none have shown signs of lignin-deficient middle lamellae. However, since most of these were African and South-east Asian species, other American genera should be examined next. Leatherwood also provokes questions on the evolution and biological functions of lignin. Archeological evidence suggests that lignin first evolved to fulfil a structural role in plants, and that it was only later recruited to support long-distance water transport and plant defence (Boyce et al., 2003; Sperry, 2003). In the case of leatherwood, water conduction was evidently prioritised as the cell walls of xylem fibre cells are poorly lignified. This allocation may be best understood in terms of the overall carbon budget since lignin represents a significant investment of photosynthetic carbon in plants (Amthor, 2003). Leatherwood maximises the return on this investment by predominantly lignifying the cell walls of water-conducting vessel elements and adjacent vasicentric tracheids. However, this is presumably only possible because leatherwood has evolved various compensatory adaptations that support plant defence and biomechanics. For example, leatherwood invests in a wide range of xylem metabolites which may be involved in defence, perhaps to compensate for the limited structural fortifications. Also, the thick bark features strong bast fibres that may help with strengthening. Even still, leatherwood stems are relatively weak and can be easily broken by heavy winter snowfalls. Interestingly, the cell walls of bast fibres in leatherwood bark are apparently devoid of lignin (Hudson, 2019). Leatherwood is not the only plant species that has divested in lignin. For example, the mechanical role of lignin has been largely replaced by silica in Equisetum spp. (Yamanaka et al., 2012). Recent work has shown that specifically altering lignin biosynthesis in fibre cells may be a promising engineering strategy (Liang et al., 2019; Gui et al., 2020), mirroring the natural lignification pattern 150 apparent in leatherwood. But what other lessons can be drawn from this species? It would be interesting to examine whether leatherwood bark contributes to stem strength as compensation for the weakly lignified xylem fibres. Similarly, leatherwood metabolites such as phenolic glycosides and dirchromones (Ramsewak et al., 1999; St-Gelais et al., 2015) have never before been studied in the context of plant defence. It could also be informative to investigate what, if any, are the impacts of this lignification patterns in terms of water transport and consequently on the overall physiology of leatherwood. Such work would go a long way toward unravelling the implications of atypical lignification in leatherwood. Although the genetic basis of monolignol biosynthesis has been largely mapped, there persist many unanswered questions surrounding monolignol export and lignin polymerisation (Perkins et al., 2019; Tobimatsu & Schuetz, 2019). Among these, the enigma of lignin initiation in the middle lamella remains particularly unresolved. As an apparent exception to the norm, leatherwood could be extremely useful in future studies on these topics. For example, one hypothesis to explain the lignin-deficient middle lamellae in leatherwood is that monolignol export or radical formation could be impaired. In preliminary experiments, I attempted to assess this by visualising the in situ formation of lignin following the exogenous application of fluorescently labelled monolignols. While this work yielded inconclusive results, further experiments in this area could help explain why leatherwood middle lamellae remain unlignified and, in so doing, could help demystify the dynamics of lignification in plants more broadly. Until now, all work on leatherwood has been conducted using responsibly harvested wild plants. Unfortunately, leatherwood is notoriously difficult to grow from seeds (Del Tredici, 1984; Schrader & Graves, 2005). Propagation from cuttings has not yet been reported and nor has the establishment of explants in tissue culture. As such, optimised procedures for the propagation of leatherwood are urgently needed. Furthermore, the pace of leatherwood research could be greatly accelerated by the availability of genomic and/or transcriptomic resources. Measurements of nuclear DNA content point to a rather large genome, roughly four times that of poplar (Fridley & Craddock, 2015). And so, it would be prudent to be selective in the choice of genotype before embarking on genome sequencing and assembly projects. Phytogeographic studies have identified a number of leatherwood populations that exhibit relatively low heterozygosity which could be good candidates for further study (Peterson et al., 2016). 151 Finally, any lignin treatise would be wholly incomplete without briefly mentioning the enigmatic lignin found among the recondite Rhopalocera. Although detailed reports remain scant at the time of this writing, it is apparent that lignins from arborescent butterflies stand apart from all other occurrences in Nature, both in terms of their unique composition and structure but also for the important epistemological questions which they elicit on the evolution and biological roles of lignin (Leisola et al., 2012). 7.5 Outroduction Lignin research has progressed steadily from the earliest efforts which were largely descriptive, through to more mechanistic studies on lignin biochemistry, and has finally reached a point where discoveries in lignin biology can be readily applied in the engineering of biomass feedstock species for industrial gains (Erdtman, 1933; Freudenberg & Neish, 1968; Adler, 1977; Donaldson, 2001; Ralph et al., 2004a; also see Chapter 2). 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Mottiar Y, Chafe PDJ, Ribbens E. Imperfect flowers of Opuntia fragilis in Kaladar, Ontario. Haseltonia 2015, 20: 22–25. 2. Cass CL, Peraldi A, Dowd PF, Mottiar Y, Santoro N, Karlen SD, Bukhman YV, Foster CE, Thrower N, Bruno LC, Moskvin OV, Johnson ET, Willhoit ME, Phutane M, Ralph J, Mansfield SD, Nicholson P, Sedbrook JC. Effects of PHENYLALANINE AMMONIA LYASE (PAL) knockdown on cell wall composition, biomass digestibility, and biotic and abiotic stress responses in Brachypodium. J. Exp. Bot. 2015, 66(14): 4317–4345. 3. Kuglerová L, García L, Pardo I, Mottiar Y, Richardson JS. Does leaf litter from invasive plants contribute the same support of a stream ecosystem function as native vegetation? EcoSphere 2017, 8(4): e01779. 4. Murphy EK, Mottiar Y, Soolanayakanahally RY, Mansfield SD. Variations in cell wall traits impact saccharification potential of Salix famelica and Salix eriocephala. Biomass & Bioenergy 2021, 148: 106051. 5. Kim KH*, Mottiar Y*, Jeong K, Tran PHN, Tran NT, Zhuang J, Kim CS, Lee H, Gong G, Ko JK, Lee S-M, Kim SY, Shin JY, Song HK, Yoo CG, Kim N-K, Mansfield SD. One-pot conversion of engineered poplar into biochemicals and bioproducts using biocompatible deep eutectic solvents. Forthcoming publication. *Equally contributing first-authors. 6. Unda F, Mottiar Y, Mahon EL, Karlen SD, Kim KH, Loqué D, Eudes A, Ralph J, Mansfield SD. 3,4-Dihydroxybenzoate is compatible with lignification in poplar. Forthcoming publication. 188 Appendix 2. Supplementary Information Pertaining to Chapter 6 A2.1 Supplementary Notes A2.1.1 The Dirca genus The Dirca genus comprises four species of slow-growing sciophilous angiosperms: D. occidentalis Gray, which is found only in the San Francisco bay region of California; D. mexicana Nesom and Mayfield, which is endemic to a single valley in Mexico’s Sierra Madre Oriental Mountains; D. decipiens Floden, which is limited to Arkansas and Kansas; and D. palustris L., which is widespread but uncommon throughout eastern Canada and the United States (Figure A2-1A; Schrader & Graves, 2004; Floden et al., 2009). Traditionally, leatherwood was used for naturopathic medicines and for natural cordage. Leatherwood stems are so flexible that one can easily tie a woody twig into a knot without it snapping (Figure A2-1B; Anderson, 1933). A2.1.2 FTIR analysis FTIR was initially used to screen for unique chemical features in leatherwood xylem in comparison to aspen and spruce. Since FTIR spectra of powdered wood are notoriously complex due to overlapping peaks from lignin and structural polysaccharides, spectra of chlorite holocellulose and acid-insoluble lignin were also collected (Figure A2-2A–C). Principal component analysis (PCA) of the ensuing datasets was used to identify noteworthy chemical features in leatherwood (Figure A2-2D–F). The first principal component of the PCA model generated for powdered wood primarily differentiated aspen from spruce and accounted for over 78 % of the total variance (Figure A2-2G). Compared to spruce, aspen was rich in the regions around 1735 and 1240 cm-1. On the other hand, the spruce spectra were more prominent in the signals at 1510, 1385, 1090 and 1020 cm-1. The second principal component captured 21 % of the cumulative variance while distinguishing spruce and aspen from leatherwood (Figure A2-2J). Spruce and aspen contained more of the bonds characterised by 1725, 1510, 1270, 1165, 1120, 1060 and 1030 cm-1 whereas leatherwood featured comparatively more of the signals at 1760, 1625 and 975 cm-1. Of particular interest is the deficiency in the 1510 cm-1 signal corresponding to generic aromatic skeletal vibrations in lignin (Faix, 1991) which points to lower levels of lignin in leatherwood compared to the spruce and aspen 189 samples. To fully realise the potential of FTIR, it was prudent to consider lignin and holocellulose separately. The first principal component of the PCA model generated for holocellulose captured 94 % of the total variance and primarily distinguished leatherwood from spruce (Figure A2-2H). Leatherwood contained more of the bonds associated with 1740, 1245 and a wide region near 1045 cm-1 whereas spruce again featured more of the signal at 1510 cm-1. The second principal component, which captured less than 6 % of the total variance, separated aspen from leatherwood and spruce (Figure A2-2K). Aspen holocellulose was rich in the bonds associated with 1335, 1165, 1110 and 980 cm-1 while leatherwood and spruce produced stronger signals at 1760, 1610, 1510 and 1270 cm-1. Since several of these signals are lignin-specific bands (e.g., 1510, 1270 cm-1), small amounts of undigested lignin likely remained in the holocellulose preparations as is often the case with acid chlorite treatment (Browning, 1967). Even still, the overrepresentation of the signals at 1740 and 1245 cm-1 in leatherwood point to an unusual hemicellulose composition. This was explored further by curve fitting (Figure 6-1A, C). The first component of the PCA model generated for acid-insoluble lignin separated aspen from spruce and leatherwood, and accounted for about 65 % of the total variance (Figure A2-2I). It revealed that lignin from aspen contained more of the signal at 1165 and a wide band at 1110 cm-1 whereas lignin from spruce and leatherwood contained more of peaks at 1510 and 1270 cm-1. The second principal component captured about 34 % of the total variance and primarily distinguished spruce from leatherwood (Figure A2-2L). Lignin from spruce was rich in the peaks at 1510, 1270, 1140 and 1030 cm-1 whereas leatherwood contained more of the bonds related to 1720, 1670, 1545 and 1110 cm-1. The weaker signal at 1510 cm-1 again points to lower lignin levels in leatherwood compared to aspen and spruce. In addition, the lower abundance of the band near 1270 cm-1 points to a noteworthy lignin composition in leatherwood since this wavenumber is associated with guaiacyl-type lignin (Faix, 1991). This was explored further by curve fitting (Figure 6-1B, D). A2.1.3 Microfibril angle Differential interference contrast microscopy was used to measure the microfibril angle of xylem cells 190 following salt impregnation and sonication (Figure A2-3). For each species, 100 measurements were taken for fibres (leatherwood and aspen) and tracheids (spruce). The microfibril angle in leatherwood xylem was consistently higher than aspen and spruce, suggesting that leatherwood cell walls could have greater tensile strength. This comes despite the leatherwood xylem samples being older (it is a slow-growing species) than aspen or spruce. In addition, leatherwood xylem cells are short compared to other woody species (Potzger & Geisler, 1937). The extent to which high microfibril angles in leatherwood xylem contribute to stem flexibility warrants future investigation. A2.1.4 Additional Raman imaging Closer examination of additional Raman bands can be informative. For example, the signal at 2920 cm-1 corresponds to the C–H stretching mode of organic compounds. Integrating this band shows a slightly higher density of structural components in the cell walls of vessels and vasicentric tracheids compared to fibres (Figure A2-4A). Poor signals in the middle lamella and cell corners are also apparent. The syringyl lignin-sensitive signal at 1335 cm-1 was apparently evenly distributed (Figure A2-4B). Although this could indicate a uniform syringyl lignin distribution, interpretation of this signal must be done cautiously due to interference from the nearby cellulose-specific band at 1380 cm-1. The total lignin and guaiacyl lignin signals are much more informative (Figure 6-3A, B). The even signal at 380 cm-1 shows that similar levels of cellulose were present in vessels, tracheids and fibres (Figure A2-4C). Finally, the band at 1090 cm-1 can be used to probe cellulose microfibril organisation (Gierlinger et al., 2010) and the stronger response in vessels points to higher microfibril angles (Figure A2-4D). A2.1.5 Lignin distribution in other Dirca species To ascertain whether lignin-deficient middle lamellae is a trait that is common throughout the genus Dirca, additional leatherwood stem samples were examined by confocal microscopy. Firstly, samples of D. palustris collected from Maine, Massachusetts and North Dakota were sectioned and examined. All three showed a deficiency of lignin in the middle lamella (Figure A2-5A–C). Next, samples of D. occidentalis and D. decipiens were collected from the natural ranges in California and Kansas. Since the natural range of D. mexicana in Mexico is difficult to access, samples were obtained from cultivated 191 specimens. Samples of all three species were sectioned and examined, and all exhibited lignin-deficient middle lamellae (Figure A2-5D–F) suggesting that this trait is conserved across the genus Dirca. A2.2 Supplementary Materials and Methods A2.2.1 Sample processing Xylem samples used for the analysis of lignin, cellulose, acetyl and structural polysaccharides were debarked, air-dried and ground in a Wiley mill to pass a 40-mesh sieve (420 μm). Similarly, samples for FTIR and X-ray diffraction were comminuted to pass a 270-mesh sieve (53 μm). A2.2.2 Fourier-transform infrared spectroscopy Samples of wood flour, acid-insoluble lignin and acid chlorite-derived holocellulose (2 mg) were homogenised with FTIR-grade potassium bromide (200 mg) in a mortar and pestle and dried overnight at 105 ºC prior to being formed into compressed discs using a hydraulic press and a die mold. Chlorite holocellulose was prepared for each species according to the procedure described by Browning (1967) using sodium chlorite and acetic acid, and acid-insoluble lignin was retained from the lignin quantification procedure described below. Spectra were recorded using a Tensor 27 FTIR spectrometer (Bruker Optics, Ettlingen, Germany) at a resolution of 4 cm-1 with 150 scans from 4000 to 400 cm-1. The spectra were corrected for atmospheric contributions, cropped to the fingerprint region (1800 to 800 cm-1) and baseline-adjusted with the rubberband algorithm and 64 baseline points using the OPUS version 5.1 software package (Bruker Optics). Triplicate samples were processed for each species and each sample type. PCA analysis was performed for all three sample types using the PLS Toolbox Version 5.8 for Matlab (Eigenvector Research Incorporated, Wenatchee, WA, U.S.A.). For each of the three sample types, three spectra from each species were normalised by unit area prior to analysis. For the lignin and holocellulose spectra, peaks were identified using the second derivative scanning mode of OPUS version 5.1 with 9 smoothing points. Curve fitting was then performed using the Levenberg-Maquardt algorithm with variable peak intensity and width, with the peak positions fixed and with the peaks modelled by a pseudo-Voigt shape (50 % Gaussian, 50 % Lorentzian). 192 A2.2.3 Lignin content The lignin content was quantified using a standardised acid hydrolysis procedure for Klason lignin (Coleman et al., 2008b). Briefly, 40-mesh wood powder was Soxhlet-extracted using acetone for 24 hours. Extractive-free wood powder (200 mg) was oven-dried at 105 ºC and weighed prior to treatment with 72 % sulphuric acid (3 mL) for two hours at 20 ºC with stirring every 10 minutes followed by acid hydrolysis in 4 % sulphuric acid (dilution with 112 mL deionied water) for one hour in an autoclave at 121 ºC and 101 kPa. Acid-insoluble lignin was determined gravimetrically as the weight of residue remaining upon medium coarseness sintered-glass crucibles after filtering and drying in an oven at 105 ºC. Acid-soluble lignin was estimated by UV-spectroscopy of the acid hydrolysis filtrates using an extinction coefficient of 110 L g-1 cm-1 at 205 nm (Swan, 1965). The oven-dried acid-insoluble powder was subsequently used for FTIR analysis, while the acid hydrolysate was used to quantify the structural polysaccharides. Acid-insoluble lignin was corrected for the ash content which was determined as the weight of extractive-free wood powder remaining in porcelain crucibles following four hours at 575 ºC in a muffle furnace. Three biological replicates were analysed for each species using technical triplicates. A2.2.4 Structural polysaccharides composition The composition of structural polysaccharides was assessed by UHPLC-DAD of acid hydrolysates following reductive amination with benzocaine as described previously (Sakamoto et al., 2015), but with slight modifications. Derivatising reagent (80 μL) containing 100 mg mL-1 benzocaine, 117 μL mL-1 glacial acetic acid and 10 mg mL-1 sodium cyanoborohydride in methanol was added to an aliquot of acid hydrolysate (20 μL) with 3-O-methylglucose as an internal standard (5 μL, 1 mg mL-1). The reaction was incubated at 60 °C for 60 minutes and then cooled prior to analysis. Good separation of six neutral sugars and three uronic acids was obtained using an AQUITY BEH C18 column (100 mm × 2.1 mm, 1.7 μm particle size, Waters Inc., Milford, MA, U.S.A.) maintained at 50 °C and an Agilent 1290 Infinity II UHPLC apparatus equipped with a diode array detector (Agilent Technologies Inc., Santa Clara, CA, U.S.A.). An injection volume of 1 μL was used in non-isocratic separation at 0.5 mL min-1 with a gradient of 5 to 12 % acetonitrile in 200 mM borate buffer (pH 8.9) over 10 minutes. Benzocaine-derivatised 193 sugars and uronic acids were detected at the UV maximum of 308 nm, and a column clean-up was performed between samples using 90% acetonitrile in 200 mM borate buffer (pH 8.9). Calibration standards of known concentration enabled quantification and correction for mass loss upon hydrolysis. Three biological replicates were analysed for each species using technical triplicates. A2.2.5 Acetyl content The cell-wall-bound acetyl content was evaluated using HPLC-RI of ester-linked organic acids liberated by alkaline hydrolysis. Extractive-free 40-mesh wood powder (25 mg) was weighed into 2-mL screw-cap vials and 2 M sodium hydroxide (1 mL) was added prior to incubation for 24 hours at 30 ºC and 500 rpm in a ThermoMixer incubator (Eppendorf, Hamburg, Germany). Following incubation, sulphuric acid (100 μL, 72 % w/w) was added and the tubes were mixed by vortexing and placed on ice to terminate the saponification reactions. Finally, acetic acid was quantified by HPLC using a Shodex RI-101 detector and an Aminex HPX-87H column (BioRad Laboratories, Hercules, California) maintained at 60 ºC with a Summit HPLC apparatus (Dionex Corporation, Sunnyvale, CA, U.S.A.). An injection volume of 25 μL was employed and good separation of peaks was achieved with an isocratic eluent of 5 mM sulphuric acid at a flow rate of 0.6 mL min-1. Three biological replicates were analysed for each species using technical triplicates, and butyric acid was used as an internal standard. A2.2.6 Lignin composition The lignin composition was measured by thioacidolysis using the standard methodology coupled with gas chromatography (Robinson & Mansfield, 2009). Thioacidolysis was performed using 10-mg samples of 40-mesh extractive-free wood powder in reactions with 1 mL of 2.5 % boron trifluoride etherate and 10 % ethanethiol in freshly distilled dioxane for four hours at 100 ºC with periodic mixing. After stopping the reactions at -20 ºC and adding 400 mM sodium bicarbonate (0.3 mL) to adjust the pH, the reaction products were extracted using dichloromethane (1 mL) and deionised water (2 mL). The organic phase was dried by passing through a loosely packed column of anhydrous sodium sulphate and then evaporated to dryness using a vacuum centrifuge. Samples were resuspended in dichloromethane (0.7 mL) and lignin monomers were derivatised by adding pyridine (20 μL) and N,O-bis(trimethylsilyl) acetamide (100 μL) to 194 sample aliquots (20 μL). Finally, derivatised monomers were analysed using a Trace 1310 gas chromatography apparatus (Thermo Scientific, Waltham, MA, U.S.A.) equipped with a TraceGOLD 5MS column using helium as a carrier gas at a flow rate of 1 mL min-1, and a flame ionisation detector fed by hydrogen (35 mL min-1) and compressed air (350 mL min-1). A 1-μL aliquot was injected by splitless injection at 250 ºC and the oven conditions were as follows: hold at 130 ºC for 3 min, then ramp at 6.5 ºC min-1 to 230 ºC and hold for 6 min, and finally ramp at 2.5 ºC min-1 to 250 ºC and hold for 10 min. Three biological replicates were analysed for each species using technical triplicates, and an internal standard of tetracosane was added prior to phase separation. Peaks corresponding to derivatised products of p-hydroxyphenyl, syringyl and guaiacyl monomers were integrated using the Chromeleon 7 software package (Thermo). A2.2.7 Cellulose content The cellulose content was determined using the standard methodology for alpha cellulose following delignification (Browning, 1967). Extractive-free wood powder (100 mg) was reacted at 50 ºC for 16 hours with 20 % sodium chlorite (1.5 mL) in sodium acetate buffer (3.5 mL containing 60 mL L-1 glacial acetic acid and 1.3 g L-1 sodium hydroxide). This was repeated for another 16 hours using fresh reaction mixture. Delignified wood powder was then recovered on a medium coarseness sintered-glass crucible after washing with 1 % acetic acid (100 mL) and acetone (10 mL). After drying at 50 ºC, a portion of the recovered holocellulose (30 mg) was incubated at room temperature for 30 minutes with 17.5 % sodium hydroxide (2.5 mL), and then diluted to 8.75 % by the addition of deionised water (2.5 mL) and incubated for another 30 minutes. After filtering through a medium coarseness sintered-glass crucible and rinsing with deionised water (100 mL), the powder was soaked in 1 M acetic acid for 5 minutes and then rinsed again with water (100 mL). The alpha cellulose content was then determined gravimetrically after drying at 105 ºC. A2.2.8 Crystallinity X-ray diffraction of wood powder was performed in reflection mode with a Philips powder X-ray diffractometer comprised of a PW1830 HT generator, a PW1050 goniometer and a PW3710 control unit 195 alongside X'Pert Quantify data capture software (PANalytical B.V., Almelo, The Netherlands). The CuKα radiation generated at 40 kV and 40 mA was primarily monochromatic upon passing through a 15-μm PW1385/00 nickel filter. Scans were obtained from 5 to 55 degrees 2θ with steps of 0.02 degrees every 2.5 seconds. An amorphous profile was evaluated using stable amorphous cellulose prepared by dissolving Avicel PH-101 (50 μm particle size, Sigma-Aldrich, St. Louis, MO, U.S.A.) in sulphur dioxide-diethylamine-dimethylsulphoxide and regenerating in water before freeze-drying (Isogai & Atalla, 1991). Triplicate samples were processed for each species. The data were baseline-corrected and the amorphous cellulose curve was scaled to reach each experimental spectrum at only one point (Thygesen et al., 2005). The crystallinity index was then evaluated as the fraction of total area above the amorphous profile which is attributable to crystallinity. A2.2.9 Microfibril angle The microfibril angle was evaluated using differential interference contrast microscopy (Wang et al., 2001). Radial longitudinal xylem sections with a thickness of 15 μm were obtained using a sliding block microtome. Sections were incubated in a 5 % (w/v) cobalt chloride solution at 80 °C for 2 hours and then transferred to a 40 kHz sonicating water bath with the same solution for an additional 2 hours. Sections were then rinsed with water, mounted onto glass slides, allowed to dry overnight, and visualised using an Axioskop 2 microscope (Carl Zeiss AG, Oberkochen, Germany) equipped with a DFC 450 C camera (Leica Microsystems GmbH, Wetzlar, Germany). The angle of fractured lamellae of cellulose microfibril bundles was measured relative to the long axis of xylem fibres (leatherwood and aspen) and tracheids (spruce) using ImageJ (National Institutes of Health, Bethesda, MD, U.S.A.), and 100 measurements were averaged for each species. 196 A2.3 Supplementary Tables and Figures Table A2-1. Annotations of FTIR bands. Wavenumbers (cm-1) Band Assignments Origin Notes References 1760 – 1725 C=O stretching of acetyl or carboxyl groups Holocellulose Related to the content of acetyl and uronic acids 1, 2 1720 – 1710 C=O stretching of unconjugated ketone and carboxyl groups Lignin 3 1675 – 1660 C=O stretching of para-substituted aryl ketone groups Lignin 3 1640 – 1625 C=O stretching of conjugated groups and absorbed O–H Holocellulose 4, 5 1605 – 1595 Aromatic skeletal vibrations and C=O stretching Lignin 3, 6 1560 – 1530 Possibly N-H stretching from small amounts of contaminating proteins Protein 9 1515 – 1505 Aromatic skeletal vibrations Lignin Related to the total lignin content 3, 6 1470 – 1425 Aromatic skeletal vibrations and asymmetrical C–H deformations Lignin 3 1430 – 1420 Symmetrical CH2 deformations Holocellulose Related to the cellulose crystallinity 7, 8 1385 – 1375 Symmetrical C–H deformations Holocellulose and Lignin 3, 4, 5 1335 – 1325 Syringyl ring breathing and C–O stretching Lignin Related to the guaiacyl lignin content 3 1335 – 1315 In-plane O–H deformations and CH2 wagging Holocellulose 4,5 1275 – 1265 Guaiacyl ring breathing and C–O stretching Lignin Related to the syringyl lignin content 3 1250 – 1230 C–O stretching of acetyl groups Holocellulose 1, 2 1170 – 1160 C–O stretching of conjugated ester groups Lignin 6 1165 – 1125 Asymmetrical vibrations of C–O–C groups Holocellulose 4, 10 1145 – 1135 Guaiacyl in-plane C–H deformations Lignin 3 1130 – 1100 Syringyl in-plane C–H deformations Lignin 3, 5 1115 – 1105 Asymmetrical ring stretching Holocellulose 4 1090 – 1080 C–O deformation of secondary alcohols and aliphatic ethers Lignin 3 1060 – 1015 C–O stretching related to cellulose and hemicellulose Holocellulose 4, 5, 10 1035 – 1030 Guaiacyl in-plane C–H deformations and C–O deformations of primary alcohols Lignin 3 985 – 970 Out-of-plane trans =CH deformations Lignin 3 915 – 815 Aromatic C–H out-of-plane deformations Lignin 3 1 Marchessault (1962) 5 Pandey (1999) 8 Ciolacu et al. (2011) 2 Stefke et al. (2008) 6 Faix (1991) 9 Stuart (2004) 3 Hergert (1971) 7 O’Connor et al. (1958) 10 Kačuráková et al. (2000) 4 Fengel & Ludwig (1991) 197 Figure A2-1. The family Thymelaeaceae and the distribution of leatherwood. A partial phylogeny showing eastern leatherwood (Dirca palustris L.) within the family Thymelaeaceae and the distribution of the genus Dirca in North America sensu Floden et al., 2009 and Beaumont et al., 2009 (A, note that Gnidia remains polyphyletic as shown). A flowering branch of leatherwood that was tied in a knot but continues to grow unabated (B). 198 Figure A2-2. FTIR analysis of leatherwood, aspen and spruce. Principal component analysis for FTIR of powdered wood, chlorite holocellulose, and acid-insoluble lignin. Representative spectra (A–C) are shown for leatherwood (blue), aspen (green) and spruce (red). Models (D–F) captured over 99 % of the cumulative variance in two principal components and are based on triplicate scans for each species and sample type. Loadings corresponding to the first component of the models (PC1) are shown in panels G–I while loadings for the second component (PC2) are shown in panels J–L. Loadings values exceeding ± 0.05 have been labelled with the corresponding wavenumbers. 199 Figure A2-3. Microfibril angle measurements of leatherwood, aspen and spruce. Longitudinal xylem cross-sections of leatherwood (A), aspen (B) and spruce (C) imaged by differential interference contrast microscopy showing how microfibril angles can be measured from fractured lamellae of cellulose microfibril bundles which become visible after salt impregnation and sonication. The average angle for 100 measurements is shown alongside standard deviation. Scale bars represent 10 μm. 200 Figure A2-4. Additional Raman imaging of leatherwood. Transverse cross-sections of leatherwood xylem imaged by confocal Raman microscopy. Chemical images are shown for specific integrated bands corresponding to total organics (A, 2920 cm-1), syringyl lignin (B, 1335 cm-1), cellulose (C, 380 cm-1) and cellulose orientation (D, 1090 cm-1). Images show vessel elements (V), fibres (F), ray parenchyma (R), axial parenchyma (A), and tracheids (T). Scale bars represent 50 μm. 201 Figure A2-5. Lignin-deficient middle lamellae across the genus Dirca. Lignin autofluorescence imaged by UV microscopy for transverse xylem cross-sections of Dirca palustris from North Dakota (A), Massachusetts (B), and Maine (C), as well as Dirca occidentalis from California (D), Dirca decipiens from Kansas (E) and Dirca mexicana originally from Tamaulipas (F). Scale bars represent 10 μm. Figure A2-6. Photos of leatherwood. Photos showing the arboresque growth form (A), the thick bast fibres in the bark (B), the structure of the branches (C), and the leaves and developing drupes (D). 202 Appendix 3. ToF-SIMS imaging reveals that p-hydroxybenzoate groups specifically decorate the lignin of fibres in the xylem of poplar and willow A3.1 Summary Esterified phenolics occur as γ-linked lignin acylations in many plant taxa but little is known about the distribution or function of such groups. In the family Salicaceae, p-hydroxybenzoate groups are present in the lignin of poplars (Populus spp.) and willows (Salix spp.). In this study, the distribution of cell-wall-bound p-hydroxybenzoate was examined in different tissue and cell types of poplar and willow trees. This analysis showed that p-hydroxybenzoate groups were most prevalent in juvenile stems, and were elevated in tension wood and following nitrogen fertilisation. Closer examination of stem cross-sections using time-of-flight secondary-ion mass spectrometry (ToF-SIMS) directly showed that p-hydroxybenzoate groups occur predominantly in the cell walls of fibres and are largely absent from vessel elements. These results point to a role for ester-linked groups in modifying the syringyl-rich lignin of fibres, perhaps to strengthen the cell walls of these structural elements or to increase lignification rates. A3.2 Introduction Lignin is a primary component of the secondary cell walls of vascular plants. It contributes greatly to the mechanical strength of stems and branches, and it plays important roles in water conduction and plant defence (Novo-Uzal et al., 2012). Although lignin is indispensable for plant vitality, it poses a major challenge for the industrial processing of plant biomass. As such, lignin chemistry and biosynthesis have received considerable research attention, and great strides have been made in modifying lignin structures through plant biotechnology (see Chapter 2). Lignin is a complex phenolic polymer that is largely resistant to chemical degradation (Figure A3-1A). While the recalcitrance of lignin presents a challenge for efficient biomass deconstruction, it also provides opportunities for the development of biochemicals and biomaterials (Glasser, 2019). Traditionally, lignin was seen as a low-value byproduct of industrial processing and was typically burned for energy. Today, new processes are being developed to preserve phenolic structures during lignocellulose deconstruction in 203 Figure A3-1. Diagram of p-hydroxybenzoylated lignin. (A) Model of lignin showing ester-linked p-hydroxybenzoate groups (–pHB) in orange. (B) Acylated monolignol conjugates of coniferyl– p-hydroxybenzoate and sinapyl–p-hydroxybenzoate. (C) Fragmentation ions of p-hydroxybenzoyl (–pHB) and p-hydroxyphenyl (H unit) moieties in lignin (following Saito et al., 2006). a strategy termed ‘lignin-first biorefining’ (Schutyser et al., 2018). Various biomaterials are envisioned from recovered polymers and oligomers, and lignin-derived phenolics could be fed to microbes for fermentation into diverse secondary products just as cellulose and hemicellulose can be converted into biofuels (Beckham et al., 2016; Kai et al., 2016). Lignin is assembled primarily from three cinnamic acid derivatives known as monolignols: p-coumaryl, coniferyl and sinapyl alcohol (Boerjan et al., 2003). Once incorporated into a lignin polymer, these precursors give rise to p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) lignin units. The monolignols are produced in the cytosol and then exported from the cell to the apoplast where peroxidase and laccase enzymes generate phenoxy radicals that undergo coupling and cross-coupling reactions (Ralph et al., 2004a). As the resulting polymers arise through combinatorial assembly, perturbations in the supply of monomers lead to differences in lignin composition and structure. The plasticity of lignification extends beyond the three primary monolignols, and various non-canonical lignin monomers are found in diverse plant taxa. For example, hydroxycinnamaldehydes and hydroxybenzaldehydes hyper-accumulate in the lignin of pine and tobacco plants deficient in the 204 biosynthetic enzyme cinnamyl alcohol dehydrogenase (Kim et al., 2003). A linear lignin polymer built entirely from caffeyl alcohol has been identified in the seed coats of cactus and vanilla beans (Chen et al., 2012). And, lignin monomers can even be recruited from other biosynthetic pathways, such as the stilbenoids resveratrol and piceatannol, which have been found in the lignin of palm fruits (Del Río et al., 2017). Some non-canonical lignins naturally arise from the incorporation of acylated (ester-linked) monomers. In commelinid grasses, for example, p-coumarate groups are linked to the lignin backbone via the γ position (Karlen et al., 2018; Ralph et al., 1994; Shimada et al., 1971). It has been shown that members of the BAHD family of acyltransferases are responsible for this biochemistry by producing ester-linked conjugates of coniferyl–p-coumarate and sinapyl–p-coumarate which can, via the monolignol moieties, participate in the radical-coupling reactions that assemble lignin polymers (Hatfield et al., 2009; Withers et al., 2012). Analogously to the better-studied p-coumarate, ester-linked p-hydroxybenzoate groups (henceforth denoted as –pHB to emphasise the pendent nature) occur in poplars, willows, palms, and at particularly high levels in the seagrass Posidonia oceanica (Lu et al., 2015; Pearl et al., 1957; Rencoret et al., 2020; Smith, 1955b). These –pHB groups are also linked via the γ position and arise through the incorporation of acylated monolignol conjugates (Figure A3-1B; Lu et al., 2004; Lu et al., 2015; Morreel et al., 2004; Nakamura & Higuchi, 1978). The p-hydroxybenzoyl moieties of these conjugates prefer radical transfer over radical coupling, and therefore remain as pendent groups and are not integrated into lignin polymer backbones (Ralph, 2006). As they are ester-linked, mild alkaline hydrolysis (saponification) releases the phenolic pendent groups as a ‘clip-off’ which can be separated for use in fermentation or directly as a platform chemical (Rinaldi et al., 2016). Furthermore, the presence of ester-linked pendent phenolics may improve deconstruction and saccharification rates (Grabber et al., 2004; Sibout et al., 2016). Accordingly, cell-wall-bound phenolics have become an important target in lignin engineering although little is known about their biological function or distribution. Previous work with UV microscopy indicated that –pHB groups may occur primarily in the xylem fibres of poplar wood (Musha & Goring, 1975). Analysis of plant tissues by time-of-flight secondary-ion 205 mass spectrometry (ToF-SIMS) offers many benefits over classical imaging techniques including high surface sensitivity (top 1–2 nm), low detection limits (ppb–ppm), high spatial resolution (∼50–300 nm), and rich spectral data with high mass resolution (Goacher et al., 2014). No tagging or labelling is required for ToF-SIMS imaging, and microtomed plant sections can be analysed directly without applied matrices. Moreover, characteristic ToF-SIMS secondary ions unique to lignin and polysaccharides have been reported previously (Goacher et al., 2011; Goacher et al., 2012; Goacher et al., 2013). In this study, the occurrence of –pHB groups was surveyed in various tissues of poplar and willow by chemical analysis, and ToF-SIMS was then used to examine the distribution of –pHB in wood. To our knowledge, this report represents the first use of ToF-SIMS to examine the distribution of non-canonical lignin monomers. A3.3 Materials and Methods A3.3.1 Plant samples Samples for chemical analysis were harvested from trees grown in a glasshouse. Dormant cuttings of Populus nigra cv. ‘Italica’ (Lombardy poplar), Populus trichocarpa cv. ‘Nisqually-1’ (black cottonwood) and Salix eriocephala DUK-E5 (heart-leaf willow) were harvested from mature trees, dipped in rooting powder (0.8% indole-3-butyric acid), planted in two-gallon pots with peat-based potting soil and irrigated as needed with water containing Hoagland’s solution (Hoagland & Arnon, 1938). After six months of growth, the main stems were divided into 50 cm sections (i.e., young, juvenile, and mature) while the roots were separated into young and mature roots. Young and mature leaves were also collected, as were branches and bark which was peeled from the main stems. Stem samples were also harvested from glasshouse-grown Populus tremula × alba INRA 717-IB4 (hybrid poplar) fed with low nitrogen (0.1 mM ammonium nitrate) and high nitrogen (10 mM ammonium nitrate) fertilisers. Samples of tension wood, opposite wood, and normal wood from the same genotype were described previously (Gerttula et al., 2015). All samples were comminuted using a Wiley mill to pass a 40-mesh screen and then Soxhlet-extracted for 24 h with hot acetone prior to analysis. Three biological replicates were analysed for each genotype. 206 Wood samples for ToF-SIMS were harvested from the same Lombardy poplar and heart-leaf willow trees described above. Transversal xylem cross-sections were cut from the main stems to a thickness of roughly 100 μm using a sliding-block microtome and then allowed to air dry. Prior to ToF-SIMS analysis, several cross-sections were solvent-extracted to remove small-molecule extractives (Goacher et al., 2013). Sections were placed in cellulose thimbles and Soxhlet-extracted for 8 h with 95% ethanol, 8 h with 70% ethanol/30% toluene, and 4 h with distilled water, following ASTM standard D1105–96 (2007) but with a doubled ethanol extraction time. Subsequently, some of the untreated and extracted microtomed sections underwent mild alkaline hydrolysis to cleave ester-linked –pHB by soaking in 2M sodium hydroxide for 24 h at 30 °C with shaking at 60 rpm (Goldstein, 1984). After extraction and/or alkaline hydrolysis, the cross-sections were rinsed with 20 mL aliquots: once with distilled water, four times with 1 M acetic acid, and finally 10 times with distilled water (Braham & Goacher, 2015). Samples of wood powder from Betula alleghaniensis (yellow birch), an angiosperm genus which is devoid of –pHB groups (Bardet et al., 1985a), were used as a negative control for peak identification purposes. A3.3.2 Chemical analysis of –pHB groups and lignin The amount of ester-linked cell-wall-bound –pHB groups was determined by mild alkaline hydrolysis of extractive-free powdered samples. Duplicate 20 mg samples of oven-dried powder were weighed into 2 mL screw-cap vials and 1 mL of 2 M sodium hydroxide was added along with 100 μL of 1 mg mL−1 o-anisic acid as an internal standard. Tubes were incubated at 30 °C for 24 h. The saponification reactions were terminated by the addition of 100 μL of 72% sulphuric acid. After incubation on ice for 5 min, the supernatants were collected by centrifugation and filtered through 0.45 μm nylon syringe filters prior to analysis using reversed phase HPLC. Using a Dionex Summit HPLC apparatus equipped with a diode array detector (Thermo Scientific, Waltham, MA, U.S.A.), 20-μL samples were injected onto a Symmetry C-18 column (4.6 × 250 mm, 5 μm particle size, Waters Inc., Milford, MA, U.S.A.) maintained at 35 °C. Good peak resolution was achieved using non-isocratic separation with a gradient over 30 min from 5 to 45% of eluent A (0.1% trifluoroacetic acid in 70:30 acetonitrile:methanol) in eluent B (0.1% trifluoroacetic acid in water) at a 207 flow rate of 0.7 mL min−1. Integration of p-hydroxybenzoic acid and o-anisic acid peaks was performed at the UV maxima of 255 and 296 nm respectively, and a seven-point calibration curve from 1 to 100 ng mL−1 prepared with analytical-grade standards (at least 99% pure, Sigma-Aldrich Corp., St. Louis, MO, U.S.A.) was used for quantification. The amount and composition of lignin was determined using the Klason lignin method and thioacidolysis procedure followed by derivatisation and gas chromatography as described previously (Robinson & Mansfield, 2009; also see section A2.2.6). Heat maps of –pHB content (normalised per g lignin) were generated using Adobe Illustrator (Adobe Systems Inc., Mountain View, CA, U.S.A.). A3.3.3 ToF-SIMS ToF-SIMS data was acquired using two ToF-SIMS 5 instruments equipped with reflectron-type mass analysers (ION-TOF GmbH, Münster, Germany). Analysis was performed using Bi3++ primary ions at a 45° incident angle with 20 eV electron flooding for charge compensation. The untreated, extracted, and alkaline-treated sections were analysed using high-current conditions for fast spectral acquisition (50 keV Bi3++, 0.7 pA pulsed current, 65 μs cycle time, 256 × 256 pixels rastered randomly over either 100 × 100 or 500 × 500 μm2 areas). High resolution images were obtained of the extracted sections using burst alignment settings with 0.11 pA current, 50 keV Bi3++ at a 65 μs cycle time, and 256 × 256 pixels rastered randomly over a 200 × 200 μm2 area; or with 0.06 pA current, 60 keV Bi3++ at a 100 μs cycle time, and 512 × 512 pixels rastered randomly over a 200 × 200 μm2 area. The pressure during analysis was less than 3 × 10−7 mbar and delayed extraction was used to minimise the effects of surface topography. SurfaceLab 6.3 and 6.6 (IONTOF GmbH, Münster, Germany) software versions were used to calibrate the ToF-SIMS mass spectra to C3H5+ (m/z 41.03), C4H7+ (m/z 55.05), C7H7+ (m/z 91.05) and C5H5O2+ (m/z 97.03), and to generate images of characteristic ions. Mass interval lists were prepared with high mass resolution to export data to Matlab R2016b (MathWorks, Natick, MA, U.S.A.) for multivariate statistical analysis using the PLS Toolbox 7.0 and MIA Toolbox 3.0 (Eigenvector Research Inc., Manson, WA, U.S.A.). Pre-processing of spectra for principal component analysis (PCA) included Poisson (square root mean) scaling, normalisation, and mean centring. Curve-smoothing was performed for the spectra 208 presented in the m/z 121 peak envelope using Microsoft Excel (Microsoft Corp., Redmond, WA, U.S.A.) with a damping factor of 0.9. Images were pre-processed using Poisson scaling and mean centring for PCA, while only Poisson scaling was applied for image analysis via multivariate curve resolution (MCR). Figures were assembled and scale bars were added using Adobe Photoshop and Adobe Illustrator (Adobe Systems Inc.). A3.4 Results and Discussion A3.4.1 Occurrence of –pHB groups in poplar and willow To begin, the distribution of –pHB was surveyed throughout the vegetative tissues of poplar and willow. As –pHB moieties are ester-linked to the lignin backbone, mild alkaline hydrolysis was used to liberate these groups prior to analysis by HPLC. This survey revealed substantial variability between tissues, but also among the different taxa more broadly (Figure A3-2). In agreement with previous reports, Lombardy poplar had the highest –pHB levels, while heart-leaf willow had the lowest (Pearl et al., 1957; Venverloo, 1971). Figure A3-2. Occurrence of ester-linked p-hydroxybenzoate groups in poplar tissues. Levels of cell-wall-bound p-hydroxybenzoate groups (mg –pHB/g lignin) in vegetative tissues of (A) Lombardy poplar, (B) black cottonwood, and (C) heart-leaf willow, as well as in (D) stems of hybrid poplar grown with low and high levels of nitrogen fertilisation, and in tension wood, opposite wood and normal wood. Tissue types depicted in a–c include stems (in 50 cm increments), branches, roots (young and mature), leaves (young and mature), and bark. Gradient scales show the lignin-normalised amount of –pHB with yellow representing more –pHB and red representing less. All measurements were performed in duplicate with extractive-free whole cell wall material using three biological replicates. 209 In all three genotypes examined, the stems contained the greatest amounts of –pHB with levels ranging from 5 to 29 mg –pHB/g lignin, or in other words, 0.5–2.9% of the lignin (Figure A3-2A–C). By examining a developmental gradient, it became clear that the juvenile stems had the most –pHB (i.e., not the youngest stems, nor the oldest). This may help explain the discord in the literature where some reported higher levels in younger stems (Chen et al., 2017), while others found more in mature stems (Venverloo, 1969; Wang et al., 2020b) and at least one group reported no differences whatsoever (Nakano et al., 1961). In the present study, more –pHB was observed in mature roots compared to young roots, particularly in Lombardy poplar where –pHB represented 2.2% of the total lignin in the mature roots. Bark contained much less –pHB (at most 0.2% of the lignin), with the leaves having only trace levels. As –pHB was most prevalent in the stems, the effect of perturbations in stem development were considered next. Nitrogen fertilisation of trees has been shown to lead to changes in development and resource allocation, generally resulting in fast-growing stems with wider vessels and reduced total lignin content (Li et al., 2012). In the hybrid poplar samples analysed in the present study, –pHB groups were more abundant in the xylem of trees fertilised with more nitrogen, in agreement with previous observations (Figure A3-2D; Pitre et al., 2007). As nitrogen fertilisation is known to reduce lignin content in glasshouse-grown poplar (Novaes et al., 2009), it should be emphasised that this represents a decrease in –pHB per g of lignin, not merely per g of biomass. Reaction wood, formed as a gravitropic response in leaning stems, is another example of wood with “natural” perturbations in xylem morphology and cell wall composition. In poplar, tension wood has fibres with a thick G-layer comprising more cellulose and less lignin, as well as vessels that are narrower and fewer in number (Jourez et al., 2001). In the hybrid poplar samples analysed in the present study, tension wood had 2.3 times more –pHB than opposite wood, and 3.5 times more than normal wood (Figure A3-2D). This observation corroborates previous NMR studies on the lignin of reaction wood (Foston et al., 2011; Hedenström et al., 2009). As lignin levels are lower following nitrogen fertilisation and in tension wood, it is conceivable that the metabolic shift away from lignin leads to greater production of p-hydroxybenzoate, the key precursor of –pHB groups. Although the biosynthesis of p-hydroxybenzoate has not yet been fully elucidated in plants, 210 radiotracer studies have shown a link to lignin biosynthesis (Terashima et al., 1975). Alternatively, variations in –pHB could be due to the presence/absence of –pHB in different cell types, as both reaction wood and nitrogen fertilisation result in altered proportions of normal fibre and vessel cell walls in wood. A3.4.2 ToF-SIMS detection of –pHB groups In order to better understand the occurrence of –pHB in wood, ToF-SIMS imaging was employed to examine the distribution of these groups in the juvenile stems of poplar and willow. Principal component analysis (PCA) was initially used to identify characteristic secondary ions (see section A3.6.1; Figure A3-6). This work uncovered a putative signature for –pHB at m/z 121.03. It was also observed that solvent extraction was important in the sample preparation, as has been reported previously (Goacher et al., 2011; Goacher et al., 2013). Examination of high mass resolution scans showed several peaks in the vicinity of m/z 121, namely m/z 121.03, 121.07 and 121.10. As a quality control measure to test whether these were spurious peaks caused by sample roughness or topography, the ToF-SIMS extractor bias was altered from 0 to +50 V to reject any stray ions originating from cell sidewalls. None of the peaks at m/z 121 were influenced by the extractor voltage and were therefore all deemed representative of real chemistry and not instrumental artefacts. Several sample treatments were devised to examine the nature of these three ions. First, solvent extraction resulted in decreased intensity of m/z 121.10 in Lombardy poplar (Figure A3-3A), consistent with the PCA analysis. The reduction of m/z 121.10 was also observed with solvent extraction of heart-leaf willow samples (data not shown). Therefore, the peak at m/z 121.10 likely corresponds to a small-molecule aliphatic extractive, and its mass is consistent with a peak assignment of C9H13+. The aromatic ring structure of –pHB is identical to H-lignin units (Figure A3-1C). As such, it was important to also consider peaks arising from H-lignin in this study. Previously reported ToF-SIMS analysis of a synthetic 8-O-4' H-type polymer resulted in the assignment of three characteristic ions for H-lignin: m/z 107.05 for C7H7O+, m/z 121.03 for C7H5O2+, and m/z 121.07 for C8H9O+ (Saito et al., 2006). Of these, –pHB would most likely fragment as m/z 121.03 due to cleavage of the C–O bond (Figure A3-1C). 211 Figure A3-3. Identification of p-hydroxybenzoate-specific ToF-SIMS fragment. Positive-ion mass spectra of the m/z 121 peak envelope for (A) various treatments of Lombardy poplar wood, and for (B) solvent-extracted samples of three different species where all spectra were normalised to total ion intensity. The ions at m/z 107.05 and 121.07 are not expected for –pHB as they would require a rearrangement of –pHB and its associated lignin monomer unit to form. As –pHB groups are ester-linked (but H-lignin units are not), mild alkaline hydrolysis should lead to a reduction in any –pHB-related peaks. Indeed, treatment with sodium hydroxide was observed to reduce the intensity of the peak at m/z 121.03 for samples of Lombardy poplar (Figure A3-3A), confirming this ion as characteristic of –pHB. The residual signal observed at m/z 121.03 after treatment with sodium hydroxide could be due to remaining, uncleaved –pHB groups or could arise from H-lignin. Notably, the intensity of the peak at 121.07 was not altered after solvent extraction or sodium hydroxide treatment. Such insensitivity to solvents and alkaline hydrolysis would be expected for peaks corresponding to H-lignin units. Chemical analysis showed that juvenile stems of Lombardy poplar had seven times as much –pHB as 212 heart-leaf willow but a similar amount of H-lignin (Table A3-1), thus a comparison of the peak envelopes for these samples could also be informative. The intensity of the peak at m/z 121.03 was only about four times greater in Lombardy poplar than in heart-leaf willow (Figure A3-3B). And, while m/z 121.03 had greater relative intensity than m/z 121.07 in both poplar and willow, the opposite was true for yellow birch, which lacks –pHB but does have H-lignin units. Moreover, the contribution of H-lignin to m/z 121.03 can be estimated since yellow birch lacks –pHB but does have H-lignin units. Following this approach, the –pHB component of the m/z 121.03 peak was estimated to be 7.5 times greater in poplar than willow, closer in line with the chemical analysis. These observations provide further support for the assignment of m/z 121.03 as originating from –pHB (with some contribution from H-lignin), m/z 121.07 as originating entirely from H-lignin units, and m/z 121.10 as corresponding to an aliphatic extractive compound. A3.4.3 Distribution of –pHB groups in wood Chemical images of the m/z 121.03 peak originating from –pHB were prepared using false-colour overlays where the –pHB signal intensity is plotted in magenta and the intensity of two aromatic ions at m/z 77.04 (phenyl cation) and m/z 91.05 (tropylium cation) representing the overall signal from lignin are summed and plotted in green (Figure A3-4). The scales represent relative, not absolute, concentrations. The ToF-SIMS chemical images for Lombardy poplar indicate that –pHB is enriched in the cell walls of fibres and is largely absent in areas surrounding vessel elements, as well as in some regions adjacent to ray parenchyma (Figure A3-4A). Although this contrast is more evident in poplar where there is a higher concentration of –pHB, the same pattern can also be observed in willow (Figure A3-4B). Additional false-colour ion images are provided (Figure A3-7) to support the interpretation that –pHB is largely absent from vessels. However, it should be emphasised that some portion of the m/z 121.03 intensity in these ion images may originate from H-lignin depending on its relative abundance. Accordingly, the ToF-SIMS images were further examined using MCR analysis. This chemometric technique deconstructs complex multivariate datasets into model components to help elucidate the contributions of minor chemical constituents (Graham & Castner, 2012). MCR analysis for the ion images 213 Figure A3-4. ToF-SIMS imaging of poplar and willow. False-colour ion images with –pHB shown in magenta (m/z 121.03) and lignin shown in green (m/z 77.04 and 91.05) for extractive-free transversal xylem cross-sections of juvenile (A) Lombardy poplar (B) and heart-leaf willow stems. Gradient scales show the ion intensities for relative quantification. Scale bars represent 20 μm. of Lombardy poplar (Figure A3-5A) and heart-leaf willow (Figure A3-5B) uncovered two model components of interest. The first of these, plotted in blue in these false-colour images, corresponds to several lignin-related peaks including G-lignin peaks at m/z 137.05 and 151.03 (Figure A3-5C, D). Not surprisingly, this model component was evenly dispersed throughout the cell walls of both fibres and vessels in the xylem of poplar and willow. The second component of interest in the MCR modelling, plotted in yellow, was comprised primarily of the peak at m/z 121.03 related to –pHB as well as m/z 167.03 and 181.05 peaks which originate from S-lignin (Figure A3-5E, F). This component was found exclusively in the cell walls of fibres. The deficiency of S-lignin signatures in vessels is well known (Gorzsás et al., 2011; Musha & Goring, 1975; Zhou et al., 2011), and this observation helps validate the methodology. While the distribution of m/z 121.03 in the ion images could not be attributed entirely to –pHB as discussed earlier, the absence 214 Figure A3-5. Multivariate curve resolution modelling of ToF-SIMS spectral images. Modelling of spectra from extractive-free transversal xylem cross-sections of juvenile (A, C, E) Lombardy poplar and (B, D, F) heart-leaf willow stems identified model components (C, D) comprising peaks related mostly to lignin (shown in blue) dispersed throughout the xylem, whereas separate components (E, F) comprising peaks related to –pHB and S-lignin (shown in yellow) were confined to the cell walls of fibres. Gradient scales show the scores intensity of each model component for relative quantification. Scale bars represent 20 μm. here of other peaks related to H-lignin (i.e., m/z 107.05 and 121.07) indicates that H-lignin is not a contributor to this model component. In other words, the response at m/z 121.03 in this model component likely originates entirely from –pHB. Thus, MCR analysis of the ToF-SIMS spectral images provides confirmation that –pHB occurs primarily, if not exclusively, in the cell walls of fibres in poplar and willow. Musha & Goring (1975) used UV microscopy to examine the distribution of –pHB groups in poplar wood. Since lignin monomers have UV maxima around 280 nm while p-hydroxybenzoate absorbs strongly near 255 nm, they interpreted differences in the UV spectra of vessels and fibres as evidence that –pHB was present only in fibres. Interestingly, another study employed the same methodology to describe 215 a decrease in the amount of –pHB in poplar xylem following wounding (Frankenstein et al., 2006). Using a spectral unmixing approach, Donaldson (2013) identified a signature in the autofluorescence of poplar and willow fibres consistent with –pHB. Building on these studies, the results presented herein provide the first direct evidence using mass spectrometry that –pHB groups occur predominantly in the cell walls of fibres. This observation may help explain the abundance of –pHB in different plant tissues. Compared to mature wood, juvenile stems have more fibres and wider vessels (Zobel & Sprague, 1998). On the other hand, very young stems have more vessels as well as fibres with thinner cell walls. In this way, juvenile stems have the highest overall proportion of fibre cell walls per gram, thus accounting for the higher concentrations of –pHB. While the elevated levels in tension wood and following nitrogen fertilisation may also relate to changes in the amount of normal fibre cells walls, reduced lignin levels offer another explanation. It could be that more p-hydroxybenzoate precursors become available, or perhaps there is greater expression of the BAHD acyltransferase responsible for generating acylated monolignol conjugates. One clue is provided by radiotracer experiments which showed that exogenous p-hydroxybenzoate incorporates into lignin more readily in earlier stages of cell wall development (Terashima et al., 1979). Alternatively, perhaps p-hydroxybenzoate is deployed to the wall to use up any remaining phenolics prior to programmed cell death or as a way of promoting plant defence. In any case, it remains unclear whether acylations are used by plants to intentionally alter lignin structure, or merely as a consequence of differences in phenylpropanoid metabolism between cell types. A3.4.4 The importance of –pHB as lignin decorations Although lignin acylations have been the focus of considerable efforts in plant biotechnology (Petrik et al., 2014; Sibout et al., 2016; Smith et al., 2015), little is known about any potential biological role for these pendent phenolics. As p-hydroxybenzoyl moieties readily transfer radicals to monolignols rather than undergoing radical coupling reactions directly, these conjugates could serve to increase polymerisation rates (Ralph, 2006). For instance, sinapyl alcohol is not efficiently oxidised by many plant peroxidases (Marjamaa et al., 2009), so conjugation with p-hydroxybenzoate could be a particularly useful 216 strategy to enhance lignification for S-rich lignins such as those in the fibres of poplar and willow. To this point, there is evidence that –pHB occurs almost exclusively on S-lignin units in poplar (Regner et al., 2018; Stewart et al., 2009), so perhaps the BAHD acyltransferase responsible for conjugating p-hydroxybenzoate prefers sinapyl alcohol over coniferyl alcohol. However, one recent study reported a negative correlation between S-lignin and –pHB in a collection of natural poplar variants (Yoo et al., 2018). The co-localisation of –pHB with S-lignin in fibres is supported by the MCR analysis herein (Figure 5), although these results cannot distinguish conjugation of –pHB to S-lignin units specifically. As –pHB groups are linked via the γ position, these acylations lead to structural changes in lignin. For example, during β–β coupling of monolignols, the γ hydroxyl is normally used to trap the quinone methide intermediate leading to the formation of resinol structures in lignin (Ralph et al., 2004a). Since such structures are not possible when –pHB is present, alternative linkages such as β–O–4 prevail in highly γ-acylated S-rich lignin polymers (Martínez et al., 2008). Greater abundance of pendent groups might also lead to reductions in polymer length. For example, lignification with –pHB and p-hydroxybenzaldehyde in transgenic Arabidopsis led to lignin polymers with lower average molecular weight (Eudes et al., 2012). Similarly, the degree of polymerisation was reduced in synthetic dehydrogenative lignin polymers due to the incorporation of p-coumarate (Djikanović et al., 2012). However, the impacts of acylated monolignols on lignin molecular weight have not yet been conclusively demonstrated. Although what benefits these lignin alterations may offer to fibres remains an open question, the industrial significance of –pHB is much clearer. Lignin polymers with abundant β–O–4 ether bonds are more amenable to deconstruction (Stewart et al., 2006; also see Chapter 2). Interestingly, it has also been suggested that liberation of –pHB promotes acid-catalysed reactions and can act as a blocking agent to prevent condensation reactions that counteract lignin depolymerisation (Bardet et al., 1985b; Chua & Wayman, 1979). However, the most attractive aspect is that these ester-linked phenolics can be readily ‘clipped-off’ by mild alkaline hydrolysis. As a platform chemical, p-hydroxybenzoate can be used to make diverse valuable commodity chemicals that would otherwise be derived from fossil fuels (Wang et al., 2018b). Perhaps most notably, parabens (a class of preservatives widely used in cosmetics and pharmaceuticals) are produced via 217 p-hydroxybenzoate by esterification with alcohols such as methanol, butanol or benzyl alcohol (Yang et al., 2018). Terephthalate, the key precursor for PET plastics, is traditionally synthesised from petroleum-derived p-xylene but could also be produced via carboxylation of p-hydroxybenzoate (Bai et al., 2016). Novel processes are also being devised to synthesise high-value compounds from p-hydroxybenzoate such as paracetamol (acetaminophen) (Ralph et al., 2019b). Furthermore, engineered microorganisms have been developed that can metabolically upgrade p-hydroxybenzoate into sundry high-value products (Becker & Whittmann, 2019). The potential of clip-off phenolics will undoubtedly encourage future plant engineering efforts to increase the levels of –pHB groups in industrially useful biomass feedstock species, and the ToF-SIMS methodology developed here will be useful in evaluating the effectiveness of this work. A3.5 Conclusions While the biological role of cell-wall-bound phenolics such as –pHB remains obscured, there is considerable industrial potential for such clip-offs in lignin valorisation. Accordingly, ToF-SIMS imaging was used to examine the distribution of –pHB groups in the xylem of poplar and willow stems. The results presented here corroborate historical UV-microscopy studies and show that –pHB is found predominantly, if not exclusively, in the cell walls of fibres. This observation helps explain the occurrence of –pHB in different plant tissues and perhaps also in reaction wood and following nitrogen fertilisation. An improved understanding of –pHB could help elucidate the biological role of lignin decorations and may inform future efforts in plant biotechnology and lignin engineering. A3.6 Supplementary Information A3.6.1 Principal component analysis Principal component analysis (PCA) was used as an unbiased tool to pinpoint which peak(s) in the ToF-SIMS mass spectra could be attributed to –pHB. As these moieties are ester-linked to lignin, any peaks related to –pHB should decrease in intensity following alkaline hydrolysis with sodium hydroxide. The ToF-SIMS spectra of untreated, solvent-extracted, and sodium hydroxide-treated samples from two stems of Lombardy poplar and one stem of heart-leaf willow were modelled using PCA (Figure A3-6). 218 Figure A3-6. Principal component analysis (PCA) of poplar and willow ToF-SIMS spectra. A: PCA model of the ToF-SIMS spectra from m/z 100–200 for various treatments of Lombardy poplar and heart-leaf willow stems as labelled, where B: PC 1 describes differences due to solvent extraction, and C: PC 2 describes differences due to mild alkaline hydrolysis. 219 The first principal component (PC 1) of this model captured 54% of the variance due to the separation of several unextracted samples with positive scores from the remaining samples, which had negative scores (Figure A3-6A). This reveals that the largest difference between samples was due to solvent extraction. Peaks that loaded positively on PC 1 were present in higher relative intensity prior to solvent extraction and are therefore due to extractive compounds (Figure A3-6B). At a given nominal (unit) mass, the extractive-related peaks tend to have a slightly higher mass. For example, at nominal mass m/z 121 there is a lignocellulose-related peak at m/z 121.03 and an extractive-related peak at m/z 121.10. This is consistent with an increased number of hydrogen atoms in extractives due to a higher concentration of saturated bonds in extractives versus unsaturated bonds in aromatic lignin (Goacher, 2013). The PCA model of the ToF-SIMS spectra further described 18% of the variance on the second principal component (PC 2), where the scores plot (Figure A3-6A) separated samples which were sodium hydroxide-treated (negative scores) from those that were not (positive scores). The ToF-SIMS peaks that loaded positively on PC 2 (Figure A3-6C) therefore correspond with samples that did not undergo mild alkaline hydrolysis and thus retain –pHB groups. The corresponding peaks in the positive loadings included several with even masses that were likely nitrogen-containing proteins (e.g., m/z 120.09), inorganic peaks (e.g., salts) below the nominal mass (e.g., m/z 112.92), as well as the peak at m/z 121.03, which was flagged for further consideration as a potential signature of –pHB. It should be emphasised that the H-lignin-related ions at m/z 107.05 and 121.07 identified by Saito et al. (2006) loaded neutrally on PC 2, demonstrating that these peaks do not distinguish between wood before and after alkaline hydrolysis, and therefore likely do not arise from –pHB. The peak at m/z 149.02 is a well-known peak attributed to phthalate esters, which could be a carboxylation adduct with –pHB caused by rearrangement in the reactive sputter plume during SIMS analysis, or could arise from phthalate contamination of these samples from the sodium hydroxide reagent. 220 Table A3-1. Lignin analysis of poplar and willow samples analysed by ToF-SIMS. Lignin content and composition of the juvenile stems of Lombardy poplar and heart-leaf willow, as well as the sample of yellow birch. All measurements were performed in triplicate, and mean values are shown alongside the standard deviation. Klason lignin (% w/w) H units G units S units –pHB groups (% of lignin) Lombardy poplar 25.57 ± 0.15 1.25 ± 0.01 34.16 ± 0.33 61.82 ± 0.33 2.76 ± 0.02 Heart-leaf willow 22.02 ± 0.31 1.40 ± 0.04 31.17 ± 0.19 66.97 ± 0.21 0.46 ± 0.02 Yellow birch 22.47 ± 0.21 0.90 ± 0.06 31.86 ± 0.42 67.24 ± 0.48 Not detected Note: On a per gram basis, Lombardy poplar stems contained 0.2557 × 2.76 ∕ 0.2202 × 0.46 = 6.97 times more –pHB than heart-leaf willow. Figure A3-7. Additional ToF-SIMS imaging of poplar and willow. Responses for –pHB are shown in magenta (m/z 121.03) and responses for lignin are shown in green (m/z 77.04 and 91.05) for extractive-free transversal xylem cross-sections of (A, B) Lombardy poplar and (C, D) heart-leaf willow stems. Gradient scales show the ion intensities for relative quantification. Scale bars represent 20 µm.
Thesis/Dissertation
eng
Forestry
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Mansfield, Shawn D., 1969-
Exploring the plasticity of lignification
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