Preparation and Characterization of Lignin

The Pennsylvania State University

The Graduate School

College of Agricultural Sciences

PREPARATION AND CHARACTERIZATION OF LIGNIN-

PROTEIN COVALENT LINKAGES

A Dissertation in

Biorenewable Systems

Brett Galen Diehl

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

May 2014

The dissertation of Brett Galen Diehl was reviewed and approved* by the following:

Nicole R. Brown Associate Professor of Wood Chemistry Dissertation Adviser Chair of Committee

John E. Carlson Professor of Molecular Genetics

Jeffrey M. Catchmark Associate Professor of Agricultural and Biological Engineering

Emmanuel Hatzakis Director of NMR facility

John Ralph Special Member Professor of Biochemistry University of Wisconsin at Madison

Paul Smith Head of Biorenewable Systems department

*Signatures are on file in the Graduate School.

Abstract

Lignin is a natural aromatic polymer that is bio-synthesized in the cell walls of almost all land plants. Great strides have been made in understanding lignin’s biological origins and chemical and physical properties. However, many unanswered questions remain. For example, the extent to which lignin interacts with other cell wall components, such as proteins, is largely unknown. In order to help address this question, the preparation and characterization of lignin- protein covalent linkages is reported here for the first time. Chapter 1 provides a more detailed introduction, justification, and literature review.

Chapter 2 focuses on the preparation of low molecular weight lignin-protein model compounds. The compounds were not prepared under biomimetic conditions. Instead, the primary focus of this study was on the characterization of the model compounds, leading to the identification of diagnostic lignin-protein NMR chemical shifts.

Chapter 3 describes the characterization of lignin-protein linkages prepared under biomimetic conditions of lignin DHP formation. NMR showed that cysteine and tyrosine containing peptides covalently crosslink with lignin, while other amino acids do not. IR and EDS were useful for showing the general incorporation of protein into the lignin, but were incapable of distinguishing covalent and non-covalent interactions.

Chapter 4 describes the interaction between lignin and gelatin protein. It was found, using EDS and IR, that gelatin was incorporated into lignin DHP. However, a lack of diagnostic NMR signatures revealed that the crosslinking was likely dominated by non-covalent interactions such as physical entanglement. This seems likely, as gelatin is lacking in both cysteine and tyrosine residues, which were shown to be the only reactive amino acids towards lignin.

Chapter 5 details attempts at identifying lignin-protein linkages in wild type Arabidopsis. Arabidopsis was grown to maturity, then lignin was extracted from cell wall material using acidified dioxane. Elemental analysis was used to show that the lignin was contaminated with about 3.75% protein; however, NMR was not able to identify lignin-protein covalent linkages.

Chapter 6 details some future experiments that could be used to explore lignin-protein linkages, and it is hoped that this work will pave the way for such studies.

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TABLE OF CONTENTS

List of Figures…………………………………………………………………………………....vii List of Tables……………………………………………………………………………………viii Abbreviations……………………………………………………………………………………..ix Acknowledgements…………………………………………………………………………...... x

Chapter 1. Introduction ...………………………………………………………………………... 1 1.1. Problem statement ...………………………………………………………………… 1 1.2. Literature review ……………………………………………………………………. 1 1.2.1. Lignin biosynthesis ……………………………………………………….. 1 1.2.2. Plant cell wall structural proteins …………………………………………. 6 1.2.3. Evidence for lignin-protein linkages …………………………………….. 10 1.3. Methods for investigating lignin-protein linkages ………………………………… 12 1.3.1. Preparation of lignin-protein compounds ……………………………….. 12 1.3.1. Characterization of lignin-protein compounds ………………………….. 16 1.4. References …………………………………………………………………………. 21

Chapter 2. Towards lignin-protein crosslinking: Amino acid adducts of a lignin model quinone methide …………………………………………………………………………………………. 25 2.1. Abstract ……………………………………………………………………………. 25 2.2. Introduction ………………………………………………………………………... 25 2.3. Experimental ………………………………………………………………………. 28 2.3.1. Materials ………………………………………………………………… 28 2.3.2. Model compound preparations ………………………………………….. 28 2.3.3. Model compound properties …………………………………………….. 29 2.3.4. Nuclear magnetic resonance spectroscopy ……………………………… 42 2.3.5. Mass spectrometry ………………………………………………………. 42 2.3.6. Computational methods …………………………………………………. 43 2.4. Results and discussion …………………………………………………………….. 44 2.4.1. Preparation of quinone methide-amino acid adducts ……………………. 44 2.4.2. Solution-state NMR of compounds 3-9 and density functional theory calculations for compounds 10 and 11 …………………………………. 46 2.4.3. Adduct isomer determination ……………………………………………. 50 2.5. Conclusions ………………………………………………………………………... 50 2.6. Acknowledgements ………………………………………………………………... 51 2.7. References …………………………………………………………………………. 51

Chapter 3. Lignin crosslinks with peptides under biomimetic conditions ……………………... 55 3.1. Abstract ……………………………………………………………………………. 55 3.2. Introduction ………………………………………………………………………... 55 3.3. Experimental ………………………………………………………………………. 57

3.3.1. Materials ………………………………………………………………… 57 3.3.2. Synthesis of lignin DHP and lignin-peptide adducts ……………………. 57 3.3.3. Scanning electron microscopy and energy dispersive X-ray spectroscopy 57 3.3.4. Nuclear magnetic resonance spectroscopy ……………………………… 58 3.3.5. Fourier-transform infrared spectroscopy ………………………………... 58 3.4. Results and discussion …………………………………………………………….. 58 3.4.1. Preparation and yields of the lignin-peptide adducts ……………………. 58 3.4.2. Lignin-peptide morphology ……………………………………………... 59 3.4.3. Lignin-peptide linkage identification ……………………………………. 60 3.4.4. Supporting techniques for lignin-peptide characterization ……………… 64 3.5. Conclusions ………………………………………………………………………... 66 3.6. Acknowledgments …………………………………………………………………. 67 3.7. References …………………………………………………………………………. 67

Chapter 4. Preparation and characterization of lignin-gelatin complexes ……………………... 71 4.1. Abstract ……………………………………………………………………………. 71 4.2. Introduction ………………………………………………………………………... 71 4.3. Experimental ………………………………………………………………………. 73 4.3.1. Materials ………………………………………………………………… 73 4.3.2. DHP and DHP-Gel syntheses …………………………………………… 74 4.3.3. Fourier-transform infrared spectroscopy ………………………………... 74 4.3.4. X-ray photoelectron spectroscopy ………………………………………. 74 4.3.5. Scanning electron microscopy and energy dispersive X-ray spectroscopy 75 4.3.6. Nuclear magnetic resonance spectroscopy ……………………………… 75 4.4. Results and discussion …………………………………………………………….. 75 4.4.1. Preparation of DHP-Gel adducts ………………………………………... 75 4.4.2. Fourier-transform infrared spectroscopy of DHP-Gel adducts ………….. 76 4.4.3. Morphology and nitrogen content of DHP-Gel adducts ………………… 77 4.4.4. Nuclear magnetic resonance spectroscopy of DHP-Gel adducts ………... 80 4.5. Conclusions ………………………………………………………………………... 82 4.6. Acknowledgments …………………………………………………………………. 82 4.7. References …………………………………………………………………………. 83

Chapter 5. Searching for lignin-protein linkages in Arabidopsis ……………………………… 86 5.1. Abstract ……………………………………………………………………………. 86 5.2. Introduction ………………………………………………………………………... 86 5.3. Experimental ………………………………………………………………………. 87 5.3.1. Growth and lignin extraction from Arabidopsis ………………………… 87 5.3.2. Elemental analysis of Arabidopsis lignin ……………………………….. 88 5.3.3. Nuclear magnetic resonance spectroscopy of Arabidopsis lignin ………. 88 5.4. Results and discussion …………………………………………………………….. 88

5.4.1. Lignin extractions from Arabidopsis ……………………………………. 88 5.4.2. Protein content of Arabidopsis extracts …………………………………. 89 5.4.3. Nuclear magnetic resonance spectroscopy of Arabidopsis lignin ………. 90 5.5. Conclusions ………………………………………………………………………... 91 5.6. Acknowledgments …………………………………………………………………. 92 5.7. References …………………………………………………………………………. 92

Chapter 6. Conclusions ………………………………………………………………………… 93 6.1. Research summary ………………………………………………………………… 93 6.2. Future endeavors …………………………………………………………………... 94 6.3. References …………………………………………………………………………. 97

List of Figures

1.1. Three ‘common’ and three ‘uncommon’ monolignols …………………………………...….3 1.2. Resonance forms of monolignol radicals ………………………………………………….....3 1.3. Typical lignin inter-unit linkages …………………………………………………….……....4 1.4. Formation via radical coupling of β-ether QMs during lignin polymerization ……………...5 1.5. Nucleophilic amino acids that could potentially react with lignin QMs …………………….5 1.6. Tyrosine radicals and cross-coupled products ……………………………………………….7 1.7. Lignin-protein complex formed via lignin-carbohydrate linkage ....…………………………9 1.8. Preparation of a lignin β-ether model compound and its corresponding QM analog ...…….13 1.9. Preparation of lignin DHP ...………………………………………………………………..14 1.10. General structure of peptides added to lignin DHP preparations ...……………………….14 1.11. 1H NMR spectrum of lignin DHP ...……………………………………………………….17 1.12. HSQC spectrum of lignin DHP ...………………………………………………………….18 1.13. FT-IR ATR spectrum of lignin DHP ...……………………………………………………19 1.14. SEM image of lignin DHP ...………………………………………………………………20

2.1. Formation of β-ether QMs via radical coupling, and their rearomatization ...……………...26 2.2. Guaiacylglycerol-β-guaiacyl ether 1 and its derived quinone methide (QM) 2 ...………….27 2.3. QM-AA model compounds …………………………………………………………………45 2.4. Overlaid HMQC side chain regions of compounds 3 and 5 ...……………………………...48 2.5. HSQC NMR spectrum of lignin DHP with overlaid α- and β-correlation data of 3-11 ...….49

3.1. Lignin-peptide crosslinking mechanism ……………………………………………………56 3.2. SEM images of DHP and lignin-peptide adducts ...………………………………………...60 3.3. HSQC NMR of lignin-CGG adduct ...………………………………………………………61 3.4. HSQC NMR of lignin-YGG adduct ...……………………………………………………...62 3.5. HSQC NMR of lignin-HGG adduct ..………………………………………………………63 3.6. FT-IR spectra of DHP and lignin-peptide adducts …………………………………………65

4.1. FT-IR of neat DHP and DHP-Gel adducts …………………………………………………77 4.2. SEM of neat DHP and DHP-Gel adducts …………………………………………………..78 4.3. Morphology and nitrogen atomic percentages for DHP-Gel adducts ……………………...79

4.4. HSQC NMR spectrum of DHP-Gel1 ……………………………………………………….81

5.1. Optical microscopy of solvents extracted and ball milled Arabidopsis cell wall material …89 5.2. HSQC NMR spectrum of Arabidopsis lignin ………………………………………………81

6.1. Cell wall models ……………………………………………………………………………95 6.2. Synthetic route to α-13C coniferyl alcohol ………………………………………………….96 6.3. Standard lignin α-shifts and α-shifts of lignin-protein linkages ……………………………97

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List of Tables

2.1. 1H and 13C NMR chemical shifts for lignin-amino acid adducts …………………………...47 2.2. Observed and DFT calculated α-13C NMR chemical shifts for compound 3 ………………50

3.1. Yield data for the DHP and lignin-peptide adducts ………………………………………...59 3.2. Inter-unit linkage ratios of the DHP and lignin-peptide adducts …………………………...64 3.3. EDS elemental analysis data for DHP and the lignin-peptide adducts ……………………..66

4.1. Nucleophilic amino acid abundance (g/100 g dry, ash-free protein) in gelatin …………….72 4.2. Preparation and yields of DHP and DHP-Gel adducts ……………………………………..76 4.3. Inter-unit linkage ratios of DHP and DHP-Gel adducts ……………………………………81

5.1. Estimated protein content of Arabidopsis and extracted lignin …………………………….89 5.2. Inter-unit linkage ratios of Arabidopsis acidic dioxane lignin ……………………………...90

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Abbreviations

AA – amino acid AGP – arabinogalactan protein ATR – attenuated total reflectance DFT – density functional theory DHP – dehydrogenation polymer EDS – energy dispersive X-ray spectroscopy FT-IR – Fourier-transform infrared spectroscopy GRP – glycine-rich protein HRGP – hydroxyproline-rich glycoprotein NMR – nuclear magnetic resonance PRP – proline-rich protein QM – quinone methide SEM – scanning electron microscopy XPS – X-ray photoelectron spectroscopy

Acknowledgements

Here, at the end of my doctoral dissertation, I would like to take the opportunity to thank the people, without whose assistance this work would not have been possible. This list is not exhaustive, and I apologize greatly for any unintentional omissions.

First, I would like to thank my advisor, Dr. Nicole Brown, and my committee members, Drs. John Ralph, Jeff Catchmark, Emmanuel Hatzakis, and John Carlson. I would also like to thank Professor Emeritus Alan Benesi, who graciously served on my committee until his happy and healthy retirement. Without guidance and patience from these individuals, this work would not have been possible.

I would like to thank Dr. John Ralph’s entire research group, especially Matt Regner and Yuki Tobimatsu, who not only helped with my research but also made life immensely enjoyable when I visited John’s lab in May of 2012. I plan to revisit Madison as often as possible.

I would like to thank the seemingly countless number of individuals who have helped me with myriad technical matters throughout the course of my PhD. To the folks in the Materials Characterization Lab, Josh Stapleton, Trevor Clark, Vince Bojan, Tim Tighe, Julie Anderson, Melisa Yashinski, and Joe Stitt, for assistance in collecting and interpreting an endless tide of spectral data, my deepest thanks. To Wenbin Luo and the members of the Scott Showalter group, sincere thanks for assistance with all manner of NMR technical support.

My funding sources and the programs and research they fostered were instrumental toward the completion of this dissertation. I would like to acknowledge the USDA National Needs Fellowship, which provided tuition and research funding support for several years. A very special thanks is warranted to the DOE sponsored Center for Lignocellulose Structure and Formation (CLSF) and all of the members therein. The center provided not only funding and facilities to support this research, but also the breadth and depth of intellectual power necessary to inspire all its members to perform to their highest capabilities. A very special thanks is also warranted to the NSF CarbonEARTH fellowship program. This program provided me with financial support, but more importantly, opportunities and memories that will last a lifetime. Many thanks to all of my fellow CarbonEARTH’ers for advice, support, and fun times.

I would like to thank my lab mates, Paul Munson and Curtis Frantz, for companionship throughout the seemingly endless process of graduate school, and for research related insights.

Finally, I would like to thank my parents for their love and patience, and for instilling in me the skills I need to make it through life’s challenges. I am indebted to them in ways that can never be repaid.

Chapter 1

Introduction

1.1. Problem statement

The purpose of this research was to investigate the reactivities of amino acids and possibly proteins toward lignin, ultimately resulting in the formation of lignin-protein crosslinks. It has been suggested that proteins located in plant cell walls may interact with lignin in many ways. For example, enzymatic proteins such as peroxidases and laccases are necessary for lignin polymerization. Furthermore, structural proteins (non-enzymatic proteins that assist in cell wall scaffolding) may assist in the initial stages of lignin deposition, which occurs in the cell wall corner region. Several mechanisms could be envisioned, with the structural proteins playing a relatively passive role, or an active one in which they template lignin polymerization, perhaps influencing the inter-unit linkage sequence of the final lignin polymer. Lignin-protein linkages may also play a role in genetically engineered plant lines. Research has shown that plants with up-regulated cell wall protein expression sometimes exhibit altered physical and chemical properties, including enhanced lignin extractability, which may be due to increased levels of lignin-protein linkages.

In spite of the potential implications of lignin-protein crosslinks in both wild type and mutant plant lines, there have been few studies addressing the fundamental aspects of lignin- protein linkages and their formation. For example, prior to this work, it was largely unknown which amino acids (if any) were reactive toward lignin, how stable lignin-protein linkages were, and how the linkages could be identified using standard analytical tools. The goal of this work was to address and answer some of those questions, mostly through in vitro studies, while trying to keep in mind the future necessity of lignin-protein identification in vivo.

The remainder of this chapter provides a literature review, which focuses on lignin biosynthesis, plant cell wall structural proteins, evidence for lignin-protein linkages, and a section detailing the methods used in this study for lignin-protein linkage preparation and characterization. The second, third, and fourth chapters are presented in manuscript form and should be considered stand-alone publications. The final chapter presents a summary of pertinent findings, discusses limitations, and provides suggestions for future work.

1.2. Literature review

1.2.1. Lignin biosynthesis

Plant cell walls are composed of a network of interacting polymers, namely cellulose, hemicelluloses, pectins, lignin, and structural proteins (Cosgrove, 2005; McQueen-Mason and Cosgrove, 1994). Of these, lignin is the most abundant aromatic biopolymer, and the second most abundant biopolymer overall (Boerjan et al., 2003). The lignin polymer is unique among

1 the plant cell wall polymers in that it is composed of phenylpropanoid monomers known as the so-called monolignols, which undergo radical polymerization via a mechanism that is apparently not under biological control beyond the generation of the lignin radicals themselves. Lignin polymerization exhibits incredible plasticity among and within species. It is thought that lignin benefits the plant by providing strength and rigidity to the cell wall, enhanced water conductivity, and pathogen resistance. Pathogen resistance in particular is provided due to the recalcitrance of lignin towards degradation. Unfortunately, this recalcitrance negatively impacts human efforts to effectively use plant cell wall materials as biorenewable resources. Specifically, lignin recalcitrance affects the pulp and paper industry, the developing biofuels industry, the agricultural industries, and the chemical industries, all of whom seek higher value products from lignin (Chapple et al., 2007; Chen and Dixon, 2008; Jung and Allen, 1995; Jung, 1989; Li et al., 2008; Stewart et al., 2006). Thus, a greater understanding of lignin chemistry and biochemistry is desirable towards controlling and minimizing its recalcitrance, as well as engineering lignin- based products.

The process of lignification begins with the biosynthesis of the monolignols. Typically, monolignols are biosynthesized from phenylalanine via a series of enzymatic steps (Boerjan et al., 2003). There is evidence that the monolignols may be stored and/or transported to the cell wall as monolignol glucosides (i.e., the phenolic hydroxyl of the monolignol is blocked by 4-O- glycosylation); however, this may not always be the case. In addition, the mode of monoglignol transport into the cell wall is unknown (i.e., golgi-derived vesicles versus plasma membrane pumps) (Boerjan et al., 2003). As noted above, the process of lignification exhibits plasticity, and this is evidenced by the variability of monolignol biosynthesis and expression. The three most common lignin monomers are shown in Fig 1.1. The expression of these monolignols varies among plant taxa. For example, gymnosperm lignin is almost entirely composed of G-units (i.e., coniferyl alcohol based) with some traces of H-units (p-coumaryl alcohol based), dicotyledonous angiosperm lignin is mainly composed of G- and S-units (sinapyl alcohol based), and monocotyledonous angiosperm lignin is composed of all three units, as well as ferulates, sinapates, and p-coumarates. Other monolignols are biosynthesized and incorporated into the lignins of both wildtype and mutant plant lines. For example, lignin found in the seed coats of some wildtype vanilla orchids and cacti is almost completely composed of caffeyl alcohol (Chen et al., 2012). In caffeic acid/5-hydroxyconiferaldehyde O-methyltransferase (COMT) deficient mutants, 5-hydroxyconiferyl alcohol is incorporated into the lignin polymer (Li et al., 2000; Ralph et al., 2001). And in cinnamyl alcohol dehydrogenase (CAD) deficient mutants, coniferaldehyde and other aldehydes are incorporated into the lignin polymer (Ralph et al., 2001).

Fig 1.1. Three ‘common’ and three ‘uncommon’ monolignols. From left to right: p-coumaryl alcohol, coniferyl alcohol, sinapyl alcohol, caffeyl alcohol, 5-hydroxyconiferyl alcohol, and coniferaldehyde. The side chain carbons of the monolignols are often referred to as α, β, and γ- positions (see leftmost structure). This nomenclature will be used throughout the document.

Once the monolignols are shuttled to the cell wall, polymerization occurs via enzymatic dehydrogenation followed by radical recombination. Glycosyl hydrolases are implicated in the removal of the glucose residue from monolignol glucosides (Boerjan et al., 2003). Dehydrogenation is then catalyzed by peroxidases and/or laccases. The exact peroxidase and/or laccase isozymes responsible for monolignol oxidation have yet to be elucidated and may vary among species (Boerjan et al., 2003). Hydrogen peroxide is necessary for peroxidase catalyzed monolignol oxidation, and the source of this peroxide is uncertain, though NADPH oxidases may play a role. Again, further research in this area is necessary. The monolignol radical is stabilized by resonance (Fig 1.2), a direct consequence of which is the multiple lignin inter-unit linkage types that are observed (Fig 1.3).

Fig 1.2. Resonance forms of monolignol radicals. R typically represents H or OCH3.

There is currently no evidence for enzymatic control over the process of monolignol radical recombination (Ralph et al., 2008). Instead, the relative ratios of lignin inter-unit linkages can vary substantially and can be influenced by many factors, including but not limited to, monolignol composition (i.e., which monolignols are present), monolignol concentration, oxidant concentration (e.g., H2O2), catalyst/enzyme concentration, pH, the polymerization matrix (i.e., is the lignin polymerizing in a hemicellulose-rich, pectin-rich, or protein-rich environment, or bulk water, etc.), and other physical and chemical concerns (Boerjan et al., 2003; Cathala et al., 1998). In general though, the predominant inter-unit linkage type in native lignins is the so- called β-ether (β-O-4) linkage, with varying quantities of other linkages, including phenylcoumaran (β-5), resinol (β-β), dibenzodioxocin (5-5/β-O-4/α-O-4), spirodienone (β-1),

3 biphenyl ether (4-O-5), biphenyl (5-5), and β-ether/α-aryl ether (β-O-4/α-O-4) (Boerjan et al., 2003; Capanema et al., 2005; Vanholme et al., 2010).

Fig 1.3. Typical lignin inter-unit linkages. Linkage ratios vary and are influenced by many factors. Linkage ratios depicted here are not indicative of ratios observed in native lignins.

In the case of the predominant β-ether linkage, radical recombination results in the formation of an unstable quinone methide (QM) intermediate (Fig 1.4) that cannot be trapped intramolecularly, but instead must be trapped by an external nucleophile (in contrast to β-5 and β-β QMs, which can be trapped intramolecularly). The nucleophile is most often water, yielding the β-ether/α-OH structure. However, other cell wall nucleophiles are known to quench the QM. For example, lignin has long been understood to covalently crosslink with plant cell wall components such as hemicelluloses through nucleophilic reactions (via hydroxyl or carboxylic acid groups) with the α-carbon of lignin QMs (Balakshin et al., 2011; Leary, 1980; Miyagawa et al., 2012; Ralph et al., 2009; Toikka et al., 1998; Yuan et al., 2011).

Fig 1.4. Formation via radical coupling of β-ether QMs during lignin polymerization. L = lignin polymer, Nuc = nucleophile, R = H or OCH3.

The crosslinking of lignin with cell wall constituents other than hemicelluloses has not been well investigated. Cell wall structural proteins contain amino acid residues with nucleophilic side chains that could react with lignin QMs (Harrak et al., 1991; Jose and Puigdomenech, 1993; Ryser et al., 1997; Kieliszewski et al., 2011). Specifically, the amino acids cysteine (Cys), lysine (Lys), histidine (His), aspartic acid (Asp), glutamic acid (Glu), serine (Ser), threonine (Thr), tyrosine (Tyr) and hydroxyproline (Hyp) (Fig 1.5) all contain nucleophilic side chain groups. Cell wall proteins containing these amino acids vary in quantity among species and cell types, ranging from as low as 1-2% to 20% dry weight basis (Albersheim et al., 2010; Cassab et al., 1988). They have previously been postulated to crosslink with lignin, and it has been suggested that they may serve as nucleation sites or templates during lignification, but this has not been adequately tested (Boerjan et al., 2003; Cassab et al., 1988; Harrak et al., 1991; Albersheim et al., 2010; Beat et al., 1989). If true, this mechanism could provide spatial and temporal control over lignin deposition and architecture (Beat et al., 1989). The following section will discuss the various classes of cell wall structural proteins that contain nucleophilic amino acids and are likely to be in close spatial proximity to lignin within the cell wall.

Fig 1.5. Nucleophilic amino acids (nuc side chain groups are highlighted in green) that could potentially react with lignin QMs. In their free amino acid forms (shown here), the α-amine and α-acid groups could also be nucleophilic; therefore, these groups were blocked to prevent competing reactions in the studies described below. From left to right, starting at the top:

cysteine (Cys), lysine (Lys), histidine (His), aspartic acid (Asp), glutamic acid (Glu), serine (Ser), threonine (Thr), tyrosine (Tyr), hydroxyproline (Hyp). Amino acid stereochemistry is not shown; L-isomers dominate in nature.

1.2.2. Plant cell wall structural proteins

Plant cell wall structural proteins account for a relatively small percentage (dry weight basis) of the total cell wall material in mature tissues. Early studies showed that primary cell walls of dicots typically contain 5-10% protein and 2% hydroxyproline (Hyp), which originates primarily from extensins (Lamport, 1974; Talmadge et al., 1973). Once secondary walls are deposited, the relative protein content drops. The following section describes the classes of cell wall proteins that may potentially interact with lignin, as well as proposed interaction mechanisms. Proteomics of specific plant species of interest are discussed in a later section.

There are two broad classes of cell wall structural proteins that seem most likely to interact with lignin: glycine-rich proteins (GRPs), and the proline/hydroxyproline-rich glycoproteins, which are often further subdivided into the proline-rich proteins (PRPs), hydroxyproline-rich glycoproteins (HRGPs), and arabinogalactan-proteins (AGPs). These protein classes are evolutionarily related, resulting in structural and functional similarities. Some evidence has indicated that these proteins may interact with lignin, or even serve as nucleation sites for lignification in the cell corners and/or the general compound middle lamella. However, conclusive evidence for lignin-protein linkages has yet to be described.

Glycine-rich proteins (GRPs) are a diverse group of proteins that are often expressed in plant cell walls. As their moniker implies, they are glycine-rich and typically contain between 60% and 70% glycine, which is much higher than most other enzymatic or structural proteins found in plants or animals. They most commonly occur in tracheary elements of protoxylem and metaxylem tissues, and are involved in diverse cellular processes during plant development and adaptation to environmental change (Chen et al., 2007; Ringli et al., 2001). Their function varies among cell types, as does their structure, which is the basis for the most current GRP classification system. Class I GRPs may contain a signal peptide followed by a highly conserved

(GGX)n region, where X is often Ala, Ser, Val, His, Phe, Tyr or Glu. Class II GRPs may also contain a characteristic cysteine-rich C-terminal. Class III GRPs typically contain fewer glycine- rich regions compared to other GRPs. Class IV GRPs are RNA-binding and contain either an RNA-recognition motif or a cold-shock domain. And class V GRPs are glycine-rich with mixed glycine repeat patterns that are not typically observed in the other classes (Mangeon et al., 2010). For in-depth information regarding GRP tissue expression pattern, subcellular localization, structure, and function, three excellent reviews are Sachetto-Martins et al. (2000), Ringli et al. (2001), and Mangeon et al. (2010).

Based on the amino acid composition of GRPs, two modes of lignin-GRP crosslinking may be envisioned. The first mode of crosslinking is through QM-nucleophile reactions, the

6 chemistry of which was discussed in a preceding section. In GRPs, the amino acids most likely to react with lignin in this manner are His, Glu, Ser, and Tyr. Another potential lignin-GRP crosslinking mechanism is through oxidative coupling of lignin with amino acid moieties, specifically tyrosine. It has been shown that GRPs are often tyrosine-rich (up to 10% Tyr), and they crosslink in an intra- and inter-peptide manner via peroxidase mediated reactions (Ringli et al., 2001; Ryser et al., 2004). The tyrosine radical and experimentally observed tyrosine cross- coupled products are shown in Fig 1.6. Such intra- and inter-peptide linkages are also observed with PRPs and HRGPs, as discussed below. When lignin is in close proximity to GRPs, lignin- tyrosine crosslinking via this oxidative mechanism may result. Alternatively, lignin-tyrosine radical coupling may be discouraged if the oxidation potentials of the monolignols and tyrosine are quite different. This seems likely, given that monolignols exhibit radical delocalization over five resonance forms (Fig 1.2), while tyrosine only exhibits four (Fig 1.6) (Cong et al., 2013). The work described here mainly focuses on the preparation and characterization of lignin-peptide linkages formed through QM-nucleophile chemistry, but some attempts were made to identify putative lignin-tyrosine radical mediated linkages, as well. More work in this area is warranted.

Fig 1.6. Tyrosine radicals and cross-coupled products. Top row: tyrosine radical resonance forms. Middle row: isodityrosine, dityrosine, and pulcherosine. Bottom row: di-isodityrosine.

Of the GRPs, those filling cell wall structural functions may be in closest spatial proximity to lignin. Previous research has shown that GRPs may interact with lignin, though covalent linkage formation has not been clearly demonstrated. In 1989, Beat et al. noted that GRPs and lignin were localized to the same cell types within Phaseolus vulgaris (common bean), and it was hypothesized that the GRPs might provide nucleation sites for lignification via tyrosine residues. The benefits to the plant would include spatial and temporal control of various lignin properties including density and three-dimensional pattern (Beat et al., 1989). Similar results were obtained by Ye and Varner in 1991, this time with regards to soybean (Ye and Varner, 1991b). In 2004, Ryser et al. demonstrated that GRPs act as linkages between secondary cell wall thickenings, mainly composed of lignin, in protoxylem elements of seed plants as the cells passively expand following apoptosis (Ryser et al., 2004). Yet no attempt was made to determine how the GRPs anchor to the lignin-rich thickenings. Interestingly, in 2007, Chen et al. showed that an Arabidopsis GRP (AtGRP9) exhibits subcellular localization comparable with that of AtCAD5, a major Arabidopsis cinnamyl alcohol dehydrogenase localized to the cell wall. Yeast two-hybrid analysis also revealed that the two proteins interacted strongly, suggesting that GRPs may play a role in lignin monolignol synthesis, which occurs prior to lignin polymerization.

Proline-rich proteins (PRPs) display great heterogeneity in their amino acid sequences, but they all contain amino acids with nucleophilic side chains such as Lys, His, Glu, Ser, and Tyr (Jose and Puigdomenech, 1993), potentially allowing for QM-nucleophile crosslinking or lignin- tyrosine oxidative crosslinking. Ryser et al. (1997) stated, "localization of PRPs in lignified secondary walls and the secretion of the protein during lignification support the hypothesis of Ye et al. (1991a) that PRP localization is related to the pattern of lignification." They also made the bold claim that, "it may be speculated that PRPs function as a scaffold for lignin deposition via their tyrosine groups followed by oxidative cross-linking of lignin monomers" (Ryser et al., 1997). A similar conclusion was reached with regards to primary cell walls by Harrak et al. (1999), as it was found that a certain PRP located in wild tomato is down-regulated in response to drought, as is lignin production. The authors concluded that lignin and protein potentially interact with one another on the basis that they are up-regulated and down-regulated together and are located within the same cellular compartment (Harrak et al., 1999).

Hydroxyproline-rich glycoproteins (HRGPs) contribute to tissue integrity and tensile strength. The most abundant and well-studied HRGPs are the extensins, which are defined by

Ser-Hyp4 glycomodules. The proline hydroxyl groups and glycomodules (typically consisting of one through four arabinose residues) are post-translationally added and their placement and abundance is determined by the sequence of the peptide chain (Cannon et al., 2008; Kieliszewski et al., 2011). Extensins are generally tyrosine-rich, enabling them to crosslink via extensin peroxidase. These networks involve short motifs, where isodityrosine forms very short intramolecular crosslinks. This isodityrosine moiety may then react with a tyrosine residue to form pulcherosine, or react with another isodityrosine residue to form the tyrosine tetramer, di-

8 isodityrosine (Fig 1.6) (Kieliszewski et al., 2011). It is conceivable that lignin could crosslink with these tyrosine residues via the radical mechanism described previously. In addition, nucleophilic amino acids are abundant in HRGPs, and include Cys, Lys, His, Tyr, Thr, Asp, and Ser and Hyp residues that remain un-glycosylated, which may allow for QM-nucleophile crosslinking. It is also possible that the hydroxyproline-bound arabinose groups may crosslink with lignin, as the primary hydroxyl of arabinose has been shown to react with lignin QMs in vitro (Toikka et al., 1998). If this occurs in vivo, then lignin might be indirectly coupled to HRGPs via lignin-carbohydrate linkages (Fig 1.7). Observing this scenario may be difficult using standard lignin characterization techniques such as HSQC NMR, and warrants further investigation.

Fig 1.7. Hypothetical lignin-protein complex formed via lignin-carbohydrate linkage. Protein fragment sequence is Ser-Hyp-Hyp-Hyp, with varying degrees of arabinose glycosylation. Lignin-carbohydrate linkage forms through reaction of the arabinose primary hydroxyl (C6-OH) with the electrophilic α-carbon of the lignin QM.

The arabinogalactan-proteins (AGPs) are much more highly glycosylated than the PRPs and HRGPs, with type II arabino-3,6-galactans (5 – 25 kDa) accounting for 90% to 98% (w/w) of the AGP (Ellis et al., 2010). The miniscule protein component is often rich in Hyp, Pro, Ala, Ser, and Thr. Of these, Hyp, Ser, and Thr could potentially be reactive toward lignin QMs. However, it seems unlikely that lignin-AGP crosslinking would occur via addition of nucleophilic amino acids to the QM, both because the protein component is so insignificant and because the oligosaccharides likely encase the protein, shielding it from inter-polymer interactions. Crosslinking between lignin and AGPs would likely occur through the mechanism shown in Fig 1.7.

GRPs, PRPs, HRGPs, and AGPs are abundant plant cell wall structural proteins. Liyama et al. (1994) stated, "there is evidence that both HRGPs and Gly-rich proteins are associated with lignin and possibly act as foci for lignin polymerization. However, no information as to the

9 nature of possible covalent linkages or their biosynthetic route is available". There may be at least two mechanisms for lignin-protein crosslinking in plant cell walls. One mechanism involves radical crosslinking, perhaps via lignin and tyrosine moieties, while the second mechanism involves reactions of nucleophilic amino acid side chains with lignin QM intermediates. The latter mechanism is the primary focus of the research described here, but lignin-protein oxidative coupling will also be studied where possible.

1.2.3. Evidence for lignin-protein linkages

In the previous section it was shown that lignin and cell wall structural proteins are often co-localized within the plant cell wall, leading some researchers to speculate on the formation of lignin-protein complexes. These lignin-protein linkages have proven difficult to detect conclusively, especially in vivo. Nevertheless, evidence (which is largely anecdotal) suggests that lignin-protein linkages may occur. The prevailing theory of lignin biosynthesis supports this hypothesis. Under the prevailing theory, monolignol radicals couple to form lignin inter-unit linkages under conditions that are free of enzymatic control. This results in the formation of the predominant β-ether linkage and the subsequent QM intermediate (Fig 1.4), which reacts quickly with the most abundant and/or most chemically compatible nucleophile. Because the quenching of the QM is under chemical control, the QM could be expected to react with any nearby nucleophile, including nucleophiles located on proteins. Indeed, the quenching of QMs by nucleophiles that are often present on amino acids has been studied in non-lignin systems. For example, the thiol group of glutathione reacts with an o-QM generated from the flavonoid quercetin (Awad et al., 2000), the thiol group of cysteine reacts with the relatively unreactive p- QM, 2,6-di-tert-butyl-4-methylene-2,5-cyclohexadienone (Bolton et al., 1997), and thiols and thiolates react with QMs derived from anthracyclines (Ramakrishnan and Fisher, 1983). Similarly, amines (but not amino acids) have been shown to trap lignin QMs (Ralph and Young, 1983). A wide array of acid and hydroxyl-containing reagents react with p-QMs (Leary et al., 1977). And primary (and to a much lesser extent, secondary) hydroxyl groups of carbohydrates react with lignin QMs (Toikka et al., 1998). Thus, given the general ability of soft (and even relatively hard) nucleophiles to quench QMs, and given that lignin QM quenching is under simple chemical control, it seems plausible that similar reactions could occur in vivo between lignin QMs and nucleophilic amino acids.

In vitro experiments have provided some evidence for lignin-protein coupling. In 1978 and 1982, F. W. Whitmore published three articles regarding lignin-protein interactions (Whitmore, 1978a, 1978b, and 1982). Whitmore isolated cell walls of Pinus elliottii (slash pine) in such a way that native peroxidase enzymes were left intact and active. Lignin dehydrogenation polymer was then added to one group of cell walls (control), and coniferyl alcohol was added to another (experimental). Upon extraction, the experimental lignin contained significantly more protein than the control, providing evidence that proteins were incorporated into lignin during polymerization and not merely physically entangled in lignin following polymerization. Whitmore then determined that hydroxyproline interacted more strongly with the lignin than

10 other amino acids, perhaps by forming ether linkages. He hypothesized that extensin was most responsible for lignin-protein crosslinking (Whitmore, 1978, 1982). However, failure to directly observe the proposed lignin-protein linkage rendered the results inconclusive. With quantitative 1D and 2D NMR experiments now commonplace, it is perhaps time to revisit these experiments in order to more accurately ascertain the exact nature of the lignin-protein interactions.

More recently, evidence for lignin-protein interactions has been obtained through the use of dynamic mechanical analysis (DMA) and Fourier-transform infrared spectroscopy (FT-IR). Salmen and Petterson (1995) found that only one glass transition was observed for protein and lignin within the primary cell wall, indicating an association that is roughly homogenous in nature. Upon treatment with a protease, the glass transition temperature increased due to removal of protein and subsequent increase in the relative concentration of the thermally stable lignin polymer. Between 2006 and 2008, Stevanic and Salmen used DMA and FT-IR to study the primary cell walls of Norway spruce, resulting in three publications. The first article found that, "strong interactions were evident between lignin and protein, between cellulose and xyloglucan, and between cellulose and pectin" (Stevanic and Salmen, 2006). A similar conclusion was reached in the second publication, with the authors stating, "to a certain extent, all the polymers in the surface material...took part in the stress transfer...indicating an intimately linked network structure" (Stevanic and Salmen, 2008a). Finally, the third publication reported similar findings, namely that there appear to be lignin-protein and lignin-pectin interactions within the primary cell wall (Stevanic and Salmen, 2008b). DMA and dynamic FT-IR can indicate that polymer- polymer interactions exist, but the exact nature of these interactions cannot be determined using these methods, so further studies are warranted. It has been shown that horseradish peroxidase enzymes can crosslink, or at least strongly interact, with a growing lignin polymer. This may be why active peroxidases persist in lignified plant cells even after apoptosis (Evans and Himmelsbach, 1991). Kaewtip et al. (2010) showed an interaction between wheat gluten and lignin, and postulated that thiol groups on cysteine residues reacted with the double bonds of lignin to form lignin-protein linkages. Unfortunately, they were unable to conclusively confirm such linkages. It is interesting to note that, using FT-IR, blood plasma protein was observed to hydrogen bond to lignin (Polus-Ratajczak et al., 2003). It is important to keep in mind that in addition to covalent crosslinking, non-covalent interactions between lignin and protein could play an important role in the structure and function of plant cell walls.

In summary, previous work has shown that a variety of nucleophiles react with non-lignin QMs, indicating the possibility for lignin-protein linkages to form via QM-nucleophile chemistry. Furthermore, evidence has shown that lignin interacts with proteins under in vitro conditions as well as in native plant cell walls. Yet there has been no attempt to directly observe in vitro or in vivo lignin-protein linkages using modern techniques such as multidimensional NMR. Given the economic importance of lignin and its ubiquitous nature within the biosphere, increased knowledge of its structure and function should be a priority. The work described here

11 extends our fundamental understanding of lignin chemistry by characterizing lignin-protein covalent linkages as well as lignin-protein non-covalent interactions.

1.3. Methods for investigating lignin-protein linkages

1.3.1. Preparation of lignin-protein compounds

Lignin-protein model compounds were first prepared and characterized under relatively simple, in vitro conditions. The simplest lignin-protein model compounds (in terms of chemical structure and molecular weight) were prepared by reacting single nucleophilic amino acids with a lignin model quinone methide (QM). A nucleophile (meaning, “nucleus loving”) is broadly defined as a chemical group containing a partial negative charge that is relatively free to react with a complementary group of opposite charge called an electrophile (meaning, “electron loving”). As described above, some amino acids contain nucleophilic side chains (Fig 1.5), as well as nucleophilic α-amine and α-acid groups. In order to prevent side reactions, these α-amine and α-acid groups were chemically blocked, resulting in the side chain groups becoming the sole nucleophilic species in the amino acids. It was hypothesized that the amino acids would react with a lignin QM, which is an unstable electrophile that forms during lignin polymerization according to the mechanism shown in Fig 1.4. The model lignin QM used here (Fig 1.8) was chosen because it can be prepared cleanly, it is relatively small and simple (chemically speaking), and it is structurally representative of QMs that form in native guaiacyl-based lignins (Kawai et al., 1999; Landucci et al., 1981; Ralph and Young, 1983). Cross-coupling reactions were carried out in dichloromethane to obtain the desired lignin-protein model compounds and to prevent addition of nucleophilic solvent to the QM. Chapter 2 provides detailed descriptions of the preparation and characterization of these lignin-amino acid compounds.

Fig 1.8. Preparation of a lignin β-ether model compound and its corresponding QM analog. The amino acids shown in Fig 1.5 were then reacted with the QM to form lignin-protein model compounds via reaction of the amino acid nucleophilic side chain with the electrophilic α-carbon of the lignin QM.

In order to explore lignin-protein coupling under more biomimetic conditions, tripeptides were added to lignin dehydrogenation polymer (DHP) during the lignin polymerization process. Lignin DHP has been used for decades to approximate the natural lignification process. It is usually prepared by slowly combining lignin monomer, peroxidase enzymes, and hydrogen peroxide over the course of hours or days (Fig 1.9). This results in a synthetic lignin that is chemically similar to native lignin, though DHP typically exhibits increased resinol and phenylcoumaran structures and a corresponding reduction in β-ether structures compared to native lignins (Freudenberg, 1968; Terashima et al., 1995). For this study, DHP was prepared according to previously published methods using coniferyl alcohol as the sole lignin monomer, dilute hydrogen peroxide as initiator, and horseradish peroxidase as enzymatic catalyst (Terashima et al., 1995). The pH of the DHP solution was 6.5, which is standard for DHP preparations and only slightly higher than biologically relevant pH (4.5 - 6.0) (Cosgrove, 2005).

Fig 1.9. Preparation of lignin DHP. Over the course of several days, a peristaltic pump combines coniferyl alcohol and horseradish peroxidase (and in this case, peptides) with dilute hydrogen peroxide, forming lignin DHP (cream-colored solution in flask on left).

Peptides were added to the lignin polymerization reaction with the general formula of XGG (Fig 1.10), where X was any of the amino acids shown in Fig 1.5. The C-termini and N- termini of the peptides were blocked via amidation and esterification, respectively, to ensure that the amino acid of interest (i.e., residue X) was the only nucleophilic moiety. Glycine was chosen as the "place holder" residue due to its expected inertness towards lignin. Peptide length was limited to three residues in order to inhibit the formation of large lignin-peptide complexes that may have been insoluble and thus difficult to characterize (e.g., liquid state NMR may have become impractical). Peptides were added in 25% mol/mol ratio to the lignin monomer (coniferyl alcohol) because it was previously reported that lignin DHPs contain between 20 and 30% β-ether linkages (Tobimatsu, 2012). Thus, the ratio of nucleophilic residues to lignin β- ether QMs was expected to be approximately 1:1 over the course of the polymerization reaction. In summary, this experiment was designed to explore the ability of amino acids to outcompete water and other nucleophiles for addition to the QM under aqueous conditions. Chapter 3 provides detailed descriptions of the preparation and characterization of these lignin-peptide compounds.

Fig 1.10. General structure of peptides added to lignin DHP preparations. X represents the amino acid nucleophilic side chain.

Lignin DHP was prepared in the presence of gelatin protein under conditions similar to those described above. Though gelatin is of animal origin, the lignin-gelatin complex was expected to be informative for several reasons. First, gelatin is both glycine and hydroxyproline- rich, as are many plant cell wall structural proteins. Second, gelatin has a rather high molecular weight (20 kDa – 100 kDa depending on gelatin type), and is thus more similar in size to cell wall structural proteins compared to tripeptides. And third, gelatin was previously shown to interact with lignin, though the presence or absence of covalent linkages was not definitively determined (Whitmore, 1978b). Gelatin contains amino acids that could potentially be nucleophilic towards lignin (see Chapter 4); however, two key amino acids, namely cysteine and tyrosine, are almost entirely lacking. Chapter 4 provides a detailed description of the preparation and characterization of these lignin-gelatin complexes.

Finally, in an attempt to identify lignin-protein linkages formed under natural conditions of lignin biosynthesis, Arabidopsis (wild-type Columbia-0) plants were grown to maturity (8 weeks), then lignin was extracted from the inflorescence stems and characterized. The cell wall proteome of Arabidopsis has been studied more extensively than most other plant species, with 20 published papers and 500 proteins with predicted signal peptide identified (Albenne et al., 2013). Inconsistencies surrounding the Arabidopsis cell wall proteome remain, and much more work is needed. For example, the size of the cell wall proteome for five-day-old cell suspension cultures has been estimated at anywhere between 33 and 96 proteins (Chivasa et al., 2002; Feiz et al., 2006; Kwon et al., 2005; Robertson et al., 1997), while one study, which characterized three-day-old cell suspension cultures, estimated the proteome at 792 (Bayer et al., 2006)! It has been estimated that structural proteins account for 1.6% of the Arabidopsis cell wall proteome (Albenne et al., 2013). The quantity of structural protein in mature Arabidopsis cell wall, in terms of dry weight percentage, is unclear. In order to estimate the protein content of Arabidopsis and extracted Arabidopsis lignin, nitrogen analysis was performed on various Arabidopsis extracts. This allowed for protein estimation by multiplying the nitrogen percentage by a factor of 6.25, assuming that all nitrogen in the sample was due to protein (Chang et al., 2008; Fukushima and Hatfield, 2001).

Lignin was extracted from Arabidopsis following a previously described acidic dioxane method (Fukushima and Hatfield, 2004). In short, Arabidopsis inflorescence stem material was pre-ground in a Wiley mill, extracted (water, ethanol, chloroform, and acetone) in a Soxhlet apparatus, then ball milled in a cryomill. This cell wall material was then extracted by refluxing with 90:10 dioxane/2 M HCl, to obtain a crude lignin extract. The crude lignin extract was “purified” by precipitation in water followed by multiple washings with diethyl ether to yield ~30-35 mg lignin per g of Arabidopsis cell wall material. It has been postulated that this extraction method selectively cleaves α-ether linkages, which should raise concerns regarding the cleavage of putative lignin-protein linkages, as well. However, this method was deemed useful for several reasons. First, it was not possible to extract lignin using the typical milled wood lignin procedure of refluxing the sample in 96:4 dioxane/water. This method has been

15 employed for decades; however, during preliminary investigations with Arabidopsis, only ~2 mg of lignin was extracted per 1 g of Arabidopsis cell wall material, which is extremely inefficient and yields far too little lignin for effective characterization. Furthermore, lignin-protein linkages are expected to be low in quantity in wild type plants, so observing the putative linkages in cellulolytic enzyme lignins or whole cell walls seems unlikely due to very low signal to noise. Chapter 5 provides a detailed description of the extraction and characterization of the Arabidopsis lignin.

1.3.2. Characterization of lignin-protein compounds

The lignin-protein model compounds and Arabidopsis lignin extracts were characterized using a variety of complementary methods. Perhaps the single most useful of these, at least in terms of ability to directly detect lignin-protein covalent linkages, is nuclear magnetic resonance (NMR) spectroscopy. NMR relies on exploiting the quantum mechanical property of spin. When atomic nuclei with an odd number of protons and/or neutrons are placed in a magnetic field the magnetic nuclear spins align with the field. A radio frequency (RF) pulse is then applied to the sample and the nuclear spins align perpendicular to the magnetic field. The nuclear spins spontaneously relax, realigning with the magnetic field in a finite amount of time through a series of complex relaxation phenomena based on their local environment. In doing so, they re- emit radio frequencies at slightly different wavelengths than the original RF pulse, determined by the local chemical environment of each nucleus. This leads (following Fourier-transform) to the generation of the NMR spectrum, expressed in ppm.

As noted above, any atomic nucleus with an odd number of protons and/or neutrons is, in principle, NMR active, though in reality active isotopes exhibit varying degrees of sensitivity to the NMR technique, and the natural abundance of the varying isotopes is also of critical importance. For the study of lignin-protein linkages, the most useful atomic isotopes are proton (1H), carbon-13 (13C, because the most abundant isotope of carbon, 12C, is not NMR active), and potentially nitrogen-15 (15N, because 14N gives broad NMR peaks). Lignin and proteins also contain oxygen; however, the NMR active isotope of oxygen (17O) is extremely low in natural abundance and is quite insensitive to the NMR technique. Thus, 17O NMR is almost never employed.

There are many NMR techniques, based on the various active nuclei as well as various pulse programs. Furthermore, NMR data can be acquired as 1-dimensional (1D), 2-dimensional (2D), or higher dimensional spectra, and in either the solid or liquid state. For the study of lignin- protein linkages, 1D and 2D liquid state spectra are likely the most useful, but require solubility, which is sometimes limited. The simplest NMR experiments (in terms of pulse programs, not necessarily in terms of spectral interpretation) for the study of lignin are the 1D 1H and 13C experiments. 1H spectra can be collected within minutes, and provide information on functional groups within a range of ~0-12 ppm. This technique is very useful for the study of small, simple molecules. However, for complex molecules such as lignin, the relatively narrow ppm range

16 results in significant chemical shift degeneracy (Fig 1.11). The 13C NMR experiment exhibits a broad chemical shift range of ~0-200 ppm and is therefore more diagnostic for determining lignin chemical structure compared to the 1H experiment. However, the low sensitivity of the 13C experiment due to the low natural abundance and the low magnetogyric ratio of the 13C nucleus, means that relatively large sample quantities (tens of milligrams) are required. This, coupled with extremely long 13C T1 relaxation times, result in experimental times of many hours or even days to collect high resolution, quantitative spectra. Despite these disadvantages, the usefulness of quantitative 13C NMR in determining lignin structure has been well documented (Capanema et al., 2004; Capanema et al., 2005; Holtman and Kadla, 2004; Holtman et al., 2006).

Fig 1.11. 500 MHz 1H NMR spectrum of lignin DHP collected in DMSO-d6/pyridine-d5.

In addition to 1D NMR, 2D NMR has proven quite useful for elucidating lignin chemical structure. For example, the heteronuclear single quantum coherence (HSQC) technique (Fig 1.12) shows peaks that correspond to direct 1H-13C coupling, and it has the advantage of relatively high sensitivity while at the same time largely eliminating the chemical shift degeneracy that arises in 1D spectra. The HSQC technique has been used, sometimes in conjunction with quantitative 13C NMR, to identify novel lignin structures and/or interpolymer crosslinking, for example in the case of the so-called lignin-carbohydrate complexes that arise from lignin-polysaccharide coupling (Balakshin et al., 2007; Balakshin et al., 2011; Chen et al., 2012; Kim and Ralph, 2010; Mansfield et al., 2012). The following chapters show that this technique is also quite useful for the investigation of lignin-protein coupling. Other 2D NMR techniques useful for investigating lignin-protein linkages include heteronuclear multiple

17 quantum coherence (HMQC), which shows direct 1H-13C coupling but uses a different pulse program than HSQC, heteronuclear multiple bond correlation (HMBC), which shows long-range (typically 2 and 3-bond) through-bond 1H-13C coupling, correlation spectroscopy (COSY) or total correlation spectroscopy (TOCSY), which show long-range through-bond 1H-1H coupling, and nuclear Overhauser effect spectroscopy (NOESY), which shows 1H-1H through-space interactions.

Fig 1.12. 500 MHz 1H-13C HSQC spectrum of lignin DHP collected in DMSO-d6/pyridine-d5. Each shift is indicative of a unique inter-unit linkage type or other functional group, which collectively represent the lignin polymer.

Fourier-transform infrared (FT-IR) spectroscopy is another useful lignin characterization technique. It has the advantages of quick spectral acquisition (seconds to minutes) with just a few milligrams of sample, especially if an attenuated total reflectance (ATR) accessory is used. In attenuated total reflectance, the IR beam passes through a crystal (typically germanium, zinc selenide, silicon, or diamond) and total internal reflection occurs. An evanescent wave, which penetrates several microns into the sample, is established at the boundary of the crystal. The sample absorbs some wavelengths of IR radiation stronger than others, resulting in the IR spectrum. An FT-IR ATR spectrum of lignin DHP is shown in Fig 1.13, and spectral

18 assignments are shown where possible (Faix and Beinhoff, 1988). IR is useful for showing protein incorporation into lignin because proteins exhibit unique IR signatures. The most diagnostic of these occur near 1540 and 1658 cm-1, which are attributed to N-H deformation with C-N stretching, and C=O stretching, respectively (Socrates, 2001). In addition, the overall shape of the OH/NH region is altered upon protein incorporation, generally becoming sharper, and sometimes exhibiting an enhanced shoulder at 3200 cm-1, attributed to N-H stretching in amide functional groups (Socrates, 2001). Unfortunately, unlike NMR, direct detection of lignin-protein linkages may not be possible with IR. This is because IR shifts diagnostic of lignin-protein linkages are likely to be of very low intensity and located within the crowded fingerprint region. Thus, IR may be useful for showing protein incorporation into lignin, but not necessarily capable of elucidating the mechanism of lignin-protein interaction (i.e., covalent vs. non-covalent).

Fig 1.13. FT-IR ATR spectrum of lignin DHP.

Scanning electron microscopy (SEM) can be used to determine how protein incorporation affects the physical morphology of lignin. Transmission electron microscopy (TEM) can also be used, but SEM has the advantage of negligible sample preparation. Furthermore, advantages of TEM, such as the ability to collect diffraction spectra, are nullified by the amorphous nature of lignin. An SEM image of lignin DHP is shown in Fig 1.14. SEM has been used in the past to show that lignin morphology is altered by the presence of cellulose (Micic et al., 2003), and to investigate native lignin morphology within the cell wall (Terashima et al., 2004; Terashima and Yoshida, 2006).

Fig 1.14. SEM image of lignin DHP. Scale = 1 µm.

Elemental analysis, in various forms, is an important analytical tool for characterizing lignin. Due to the chemical structures of the monolignol constituents, neat lignin contains only the elements carbon, oxygen, and hydrogen. These three elements also compose the lignin- carbohydrate complexes, which often form in planta. However, in addition to these three elements, proteins also contain nitrogen. Thus, if a lignin contains nitrogen, then protein incorporation/contamination should be suspected. It is common to perform bulk elemental analyses on extracted lignins to determine protein content (N% is multiplied by a factor of 6.25 to obtain protein percentage, assuming all nitrogen in the sample is from protein) (Chang et al., 2008; Fukushima and Hatfield, 2001). In addition to purely bulk elemental analyses, energy dispersive X-ray spectroscopy (EDS) and X-ray photoelectron spectroscopy (XPS) can be used to obtain elemental data. In EDS, elemental composition is determined by bombarding the sample with electrons, then analyzing characteristic X-rays emitted from the sample. One of the advantages of EDS is that it can be collected in the SEM instrument while imaging the sample. This allows for comparison of sample morphology and elemental composition across the sample on the micron scale (the EDS spot size can be ~1 mm in diameter with an information depth of ~1-2 µm depending upon e- accelerating voltage). XPS is essentially the reverse process of EDS, as it determines elemental composition by bombarding the sample with X-rays then observing ejected electrons with characteristic energy levels. Because electrons have a far shorter mean free path than X-rays, the information depth of XPS is only about 10 nm. This allows for elemental analysis of the surface region. Comparison of the EDS and XPS data can then be used to show variations in elemental composition throughout the samples.

The following chapters will show that the experiments and characterization techniques described above are useful for investigating lignin-protein linkages, specifically under in vitro conditions. Future research should address the possibility of lignin-protein linkage formation in native plant systems.

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Chapter 2

Towards lignin-protein crosslinking: Amino acid adducts of a lignin model quinone methide

(Published in Cellulose, available here)

2.1. Abstract

The polyaromatic structure of lignin has long been recognized as a key contributor to the rigidity of plant vascular tissues. Although lignin structure was once conceptualized as a highly networked, heterogeneous, high molecular weight polymer, recent studies have suggested a very different configuration may exist in planta. These findings, coupled with the increasing attention and interest in efficiently utilizing lignocellulosic materials for green materials and energy applications, have renewed interest in lignin chemistry. Here we focus on quinone methides— key intermediates in lignin polymerization—that are quenched via reaction with cell-wall- available nucleophiles. Reactions with alcohol and uronic acid groups of hemicelluloses, for example, can lead to lignin-carbohydrate crosslinks. Our work is a first step toward exploring potential quinone methide (QM) reactions with nucleophilic groups in cell wall proteins. We conducted a model compound study wherein the lignin model compound guaiacylglycerol-β- guaiacyl ether 1, was converted to its QM 2, then reacted with amino acids bearing nucleophilic side-groups. Yields for the QM-amino acid adducts ranged from quantitative in the case of QM- lysine 3, to zero (no reaction) in the cases of QM-threonine 10 and QM-hydroxyproline 11. The structures of the QM-amino acid adducts were confirmed via 1D and 2D nuclear magnetic resonance (NMR) spectroscopy and density functional theory calculations, thereby extending the lignin NMR database to include amino acid crosslinks. Some of the QM-amino acid adducts formed both syn- and anti-isomers, whereas others favored only one isomer. Because the QM- threonine 10 and QM-hydroxyproline 11 compounds could not be experimentally prepared under conditions described here but could potentially form in vivo, we used density functional theory to calculate their NMR shifts. Characterization of these model adducts extends the lignin NMR database to aid in the identification of lignin-protein linkages in more complex in vitro and in vivo systems, and may allow for the identification of such linkages in planta.

2.2. Introduction

Plant cell walls are composed of a network of interacting polymers, namely cellulose, hemicelluloses, pectins, lignin, and structural proteins (Cosgrove 2005; McQueen-Mason and Cosgrove 1994). Of these, lignin is the major aromatic component, derived from monolignols— phenylpropanoid units whose biosynthesis exhibits incredible plasticity (Boerjan et al. 2003; Ralph et al. 2004; Vanholme et al. 2010). Lignin’s mode of polymerization is unique among the cell wall polymers. Resonance stabilized radicals are enzymatically generated from the monolignols, and as the radical-bearing structures couple combinatorially, a heterogeneous polymer containing many types of inter-unit linkages forms. The variety of the inter-unit

25 linkages contributes notable recalcitrance to the plant cell wall, stymying not only natural degradation, but also affecting the economics of many industrial sectors, including the pulp and paper industry, the developing biofuels industry, agricultural industries, and chemical industries, which all seek higher value products from lignin (Chapple et al. 2007; Chen and Dixon 2008; Jung 1989; Jung and Allen 1995; Li et al. 2008; Stewart et al. 2006).

Inter-unit linkages are not, however, the sole factor influencing lignin’s recalcitrance in planta. Lignin may be crosslinked with other polymers in the plant wall. Hydroxyl and uronic acid groups of polysaccharides bear mildly nucleophilic groups that can react with a key lignin intermediate—the α-carbon of quinone methides (QMs) (Balakshin et al. 2011; Leary 1980; Miyagawa et al. 2012; Ralph et al. 2009; Toikka et al. 1998; Yuan et al. 2011). These QMs form each time a monolignol radical couples at its β-position and, because β-coupling is prevalent, the importance of QMs in lignin structure cannot be understated. In certain cases, particularly β-5- and β-β-coupling, QM intermediates are quickly trapped intramolecularly, producing phenylcoumaran and resinol units (Leary 1980; Ralph et al. 2009). However, in the case of the predominant β-O-4-coupling, which produces β-ether linkages, the QM’s α-carbon becomes susceptible to external nucleophilic attack (Fig 2.1) (Leary 1980; Ralph et al. 2009). This reactivity of the QM is the focus of the current study.

Fig 2.1. Formation of β-ether QMs via radical coupling, and their rearomatization during lignin polymerization. L = lignin polymer, Nuc = nucleophile (e.g., H2O, and also here Cys, Lys, His, Asp, Glu, Tyr or Ser), R = H or OCH3

The crosslinking of lignin with cell wall constituents other than hemicelluloses has not been well investigated. Cell wall structural proteins, including glycine-rich proteins (GRPs), proline- rich proteins (PRPs), and hydroxyproline-rich glycoproteins (HRGPs), all contain amino acid residues with nucleophilic side-chains that could react with lignin QMs (Harrak et al. 1991; Jose and Puigdomenech 1993; Kieliszewski et al. 2011; Ryser et al. 1997). Cell wall proteins vary in quantity among species and cell types, ranging from as low as 1-2% to 20% on a dry weight basis in wild type plants (Albersheim et al. 2010; Cassab and Varner 1988). In 1978 and 1982, Whitmore showed evidence for the formation of lignin-protein linkages in isolated cell walls of slash pine. Further literature sources suggest that structural proteins may crosslink with lignin, or possibly even nucleate, or provide a template for, lignin structure, but these ideas have not been adequately tested (Albersheim et al. 2010; Beat et al. 1989; Boerjan et al. 2003; Cassab and

Varner 1988; Harrak et al. 1991). If true, this mechanism could provide spatial and temporal control over lignin deposition and architecture (Beat et al. 1989). Furthermore, it has recently been suggested that over-expression of cell wall proteins could result in increased lignin-protein linkage formation, which may affect cell wall physical and chemical properties, for example increased sugar extractability (Liang et al. 2008; Xu et al. 2013). However, identifying such linkages in planta would be difficult without first determining diagnostic lignin-protein spectroscopic signatures under simpler, more controlled conditions.

As a first step toward investigating potential lignin-protein linkages in planta, we conducted a model compound study to characterize products formed when the lignin model compound guaiacylglycerol-β-guaiacyl ether 1 was converted to its QM 2 (Fig 2.2), then reacted with amino acids bearing nucleophilic side-groups. Thiols, amines, acids and alcohols have been shown to quench QMs in a diverse array of systems. The thiol group of glutathione reacts with an o-QM generated from the flavonoid, quercetin (Awad et al. 2000); the thiol group of cysteine reacts with the relatively unreactive p-QM, 2,6-di-tert-butyl-4-methylene-2,5-cyclohexadienone (Bolton et al. 1997); and thiols and thiolates react with QMs derived from anthracyclines (Ramakrishnan and Fisher 1983). Similarly, amines have been shown to trap lignin QMs (Ralph and Young 1983). A wide array of acid- and hydroxyl-containing compounds react with p-QMs (Leary et al. 1977), and primary (and to a much lesser extent, secondary) hydroxyl groups of carbohydrates may react with QM 2 (Toikka et al. 1988). However, similar nucleophile-QM adducts have not been characterized in lignin-protein systems.

Fig 2.2. Guaiacylglycerol-β-guaiacyl ether 1 and its derived quinone methide (QM) 2

The nucleophilic amino acids investigated here—cysteine (Cys), lysine (Lys), histidine (His), aspartic acid (Asp), glutamic acid (Glu), tyrosine (Tyr), serine (Ser), threonine (Thr) and hydroxyproline (Hyp)—occur in plant cell wall structural proteins and may react to form lignin- protein crosslinks in vivo (Jose and Puigdomenech 1993; Kieliszewski et al. 2011). Because cell wall proteins are thought to exist in the wall prior to lignification, the α-amine and α-acid groups of the amino acids were protected to mimic their inclusion within a peptide. This allowed reactions of the nucleophilic side-chains to be determined without the complication of competing reactions from the terminal α-amine and α-acid groups. The QM-amino acid adducts (Fig 2.3) were characterized by nuclear magnetic resonance (NMR) spectroscopy, density functional

27 theory (DFT), mass spectrometry, and UV/Visible (UV/Vis) spectrophotometry. The characterization of these model adducts extends the lignin NMR database to aid in the identification of lignin-protein linkages in more complex in vitro and in vivo systems (Ralph et al. 2004).

2.3. Experimental

2.3.1 Materials

All chemicals used in the preparation of compounds 1 and 2, and lignin dehydrogenation polymer (DHP), were purchased from Sigma. All amino acids used in the preparation of compounds 3-9 were purchased from Sigma with the exception of Boc-L-histidine methyl ester, which was purchased from Indofine Chemical Company.

2.3.2. Model compound preparations

Compound 1 was prepared according to previous methods, as was its QM analog (2) (Kawai et al. 1999; Landucci et al. 1981; Ralph and Young 1983). Protected amino acids (1.05 eq) were added directly to the anhydrous solution of 2 in dichloromethane at room temperature.

In the case of Lys, which was obtained as Nα-acetyl-L-lysine methyl ester hydrochloride, triethylamine (~5 eq) was added in order to deprotonate the terminal amine and facilitate dissolution. NMR was used to show that triethylamine was not reactive towards the QM. A stir bar was added, the flask was stoppered, and the atmosphere was rendered inert by alternating between vacuum and dry nitrogen several times. The reaction was monitored visually; dissipation of the yellow hue indicated consumption of the QM. Intermittently, the reaction was also monitored by TLC (1:1 ethyl acetate/hexanes). Lys and His reacted with the QM within minutes. Other amino acids reacted more slowly with the QM and were allowed to stir overnight (Cys, Asp, Glu, Tyr) or for several days (Ser, Thr, Hyp), again, with intermittent monitoring by TLC. When TLC revealed that the reaction had reached equilibrium the mixture was evaporated to dryness. In the case of Lys, the reaction went to completion (complete consumption of the QM) and the crude products were evaporated to dryness and submitted to NMR without further purification. QM reactions with other amino acids did not go to completion. In the case of QM- Thr and QM-Hyp, TLC and NMR showed no reaction even over the course of several weeks. The products were purified via flash chromatography using silica gel and 1:1 ethyl acetate/hexanes as eluent. The purified products were then characterized using nuclear magnetic resonance (NMR) spectroscopy, mass spectrometry, and density functional theory (DFT). In the case of QM-His (5), the product could not be chromatographically separated (a range of eluent solvent systems were attempted) from α-O-aryl products formed presumably due to self- dimerization of the QM (2); however, mass spec and 2D NMR techniques were still able to confirm the identity of the QM-His product. In the case of QM-Ser (8), the product could not be fully separated from unreacted serine. The neat serine shifts as well as the shifts of compound 8 are labeled in the NMR spectra (see below).

Lignin guaiacyl-based dehydrogenation polymer (DHP) was synthesized according to a previously published method (Terashima et al. 1995). The DHP was characterized via HSQC NMR as described below and was found to contain shifts typical of native lignin and DHP (Capanema et al. 2004; Kim and Ralph 2010).

2.3.3. Model compound properties

QM-Cys, 3 (2-tert-Butoxycarbonylamino-3-[3-hydroxy-1-(4-hydroxy-3-methoxy-phenyl)-2-(2- methoxy-phenoxy)-propylsulfanyl]-propionic acid methyl ester). Pale white oil (yield: 73% after purification). Theoretical mass: 537.20 g/mol (+ H+: 538.21 g/mol). Observed m/z + H+: 538.21. Major isomer (86%): 1H NMR (400 MHz, acetone-d6): δ = 1.39 (9H, s, H7), 2.82 (2H, m, H1), 3.53 (1H, m, Hγ), 3.67 (3H, s, H4), 3.74 (1H, m, Hγ), 3.80 (3H, s, OMeB), 3.87 (3H, s, OMeA), 4.34 (1H, t, J = 5.78, Hα), 4.42 (1H, m, H2), 4.65 (1H, m, Hβ), 6.77 (1H, m, HA5), 6.85 (1H, m, HB6), 6.90 (1H, m, HB4), 6.94 (1H, m, HA6), 6.97 (1H, m, HB5), 7.05 (1H, m, HB3), 7.34 (1H, m, HA2). Minor isomer (14%): 1H NMR (400 MHz, acetone-d6): δ = 1.42 (9H, s, H7), 2.74 (2H, m, H1), 3.58 (1H, m, Hγ), 3.65 (1H, s, H4), 3.72 (1H, m, Hγ), 3.79 (3H, s, 13 OMeB), 3.86 (3H, s, OMeA), 4.55 (1H, m, Hβ). Major isomer (86%): C NMR (75.5 MHz, acetone-d6): δ = 28.50 (C7), 33.70 (C1), 51.41 (Cα), 52.40 (C4), 54.13 (C2), 56.11 (OMeA), 56.25 (OMeB), 62.01 (Cγ), 79.54 (C6), 83.87 (Cβ), 113.51 (CA2), 113.78 (CB5), 114.80 (CA5), 117.64 (CB3), 121.61 (CB6), 122.79 (CB4), 123.37 (CA6), 130.87 (CA1), 146.78 (CA4), 148.03 (CA3), 149.07 (CB1), 151.50 (CB2), 155.95 (C5), 172.20 (C3). Minor isomer (14%): 13C NMR (75.5 MHz, acetone-d6): δ = 113.63 (CA2), 115.41 (CA5), 117.88 (CB3), 121.79 (CB6), 131.00 (CA1), 156.33 (C5). Major isomer (86%): 1H NMR (400 MHz, DMSO-d6/pyridine- d5): δ = 1.34 (9H, s, H7), 2.66-2.86 (2H, m, H1), 3.43 (1H, m, Hγ), 3.56 (3H, s, H4), 3.58 (1H, m, Hγ), 3.70 (3H, s, OMeA), 3.78 (3H, s, OMeB), 4.18-4.27 (1H, m, H2), 4.33 (1H, m, Hα), 4.70 (1H, m, Hβ), 5.08 (1H, s, γ-OH), 6.76 (1H, m, HA5), 6.85 (1H, m, HB5), 6.88 (1H, m, HB4), 6.91 (1H, m, HA6), 6.92 (1H, m, HB6), 7.10 (1H, m, HB3), 7.34 (1H, m, HA1), 7.39 (1H, d, J = 8.07, NH), 9.19 (1H, s, A4-OH). 13C NMR (75.5 MHz, DMSO-d6/pyridine-d5): δ = 28.09 (C7), 32.23 (C1), 50.27 (Cα), 51.93 (C4), 53.47 (C2), 55.35 (OMeB), 55.74 (OMeA), 60.68 (Cγ), 81.78 (Cβ), 112.74 (CB6), 113.53 (CA2), 114.60 (CA5), 115.34 (CB3), 120.78 (CB5), 121.44 (CB4), 122.48 (CA6), 129.22 (CA1), 130.90 (C6), 146.05 (CA4), 147.43 (CA3), 148.01 (CB1), 149.87 (CB2), 155.37 (C5), 171.78 (C3). Minor isomer (14%): 1H NMR (400 MHz, DMSO- d6/pyridine-d5): δ = 4.35 (1H, m, Hα), 4.61 (1H, m, Hβ). 13C NMR (75.5 MHz, DMSO- d6/pyridine-d5): δ = 28.12 (C7), 32.42 (C1), 50.60 (Cα), 51.97 (C4), 53.69 (C2), 60.96 (Cγ), 81.65 (Cβ), 155.63 (C5), 171.74 (C3).

QM-Lys, 4 (2-Acetylamino-6-[3-hydroxy-1-(4-hydroxy-3-methoxy-phenyl)-2-(2-methoxy- phenoxy)-propylamino]hexanoic acid methyl ester). Pale white oil (yield: quantitative, no purification necessary). Theoretical mass: 504.25 g/mol (+ H+: 505.25 g/mol). Observed m/z + H+: 505.25. 1H NMR (300 MHz, acetone-d6): δ = 1.4 (4H, m, H2, H3), 1.52 (2H, m, H4), 1.90 (3H, s, H7), 2.41 (2H, m, H1), 3.50 (1H, m, Hγ), 3.63 (3H, s, H9), 3.69 (1H, m, Hγ), 3.79 (3H, s,

OMeA), 3.86 (3H, s, OMeB), 3.97 (1H, d, J = 6.82 Hz, Hα), 4.18 (1H, m, Hβ), 4.38 (1H, m, H5), 6.77 (1H, m, HB6), 6.85 (1H, m, HA6), 6.93 (1H, m, HA5), 6.96 (1H, m, HB5), 6.96 (1H, m, HB4), 7.09 (1H, m, HA2), 7.14 (1H, m, HB3), 7.39 (1H, d, J = 7.47, acetyl-NH). 13C NMR (75.5 MHz, acetone-d6): δ = 22.55 (C7), 24.09 (C3), 30.24 (C2), 32.34 (C4), 47.34 (C1), 52.03

(C9), 52.97 (C5), 56.09 (OMeA), 56.09 (OMeb), 62.19 (Cγ), 64.62 (Cα), 87.12 (Cβ), 111.96 (CA2), 113.16 (CB5), 115.35 (CB6), 118.98 (CB3), 121.75 (CA6), 121.75 (CB4), 122.99 (CA5), 133.12 (CA1), 146.70 (CA4), 148.33 (CA3), 149.45 (CB1), 151.63 (CB2), 170.11 (C6), 173.60 (C8). 1H NMR (300 MHz, DMSO-d6/pyridine-d5): δ = 1.21 (4H, m, H2, H3), 1.58 (2H, m, H4), 1.88 (3H, s, H7), 2.31 (2H, m, H1), 3.45 (1H, m, Hγ), 3.59 (3H, s, H9), 3.67 (1H, m, Hγ),

3.71 (3H, s, OMeA), 3.79 (3H, s, OMeB), 3.90 (1H, d, J = 6.82 Hz, Hα), 4.21 (1H, m, Hβ), 4.27 (1H, m, H5), 5.00 (1H, s, γ-OH), 6.76 (2H, m, HA5, HA6), 6.84 (1H, m, HB5), 6.90 (1H, m, HB4), 6.95 (1H, m, HB6), 6.98 (1H, m, HA2), 7.15 (1H, m, HB3), 8.30 (1H, d, J = 7.47, acetyl- NH), 9.12 (1H, s, A4-OH). 13C NMR (75.5 MHz, DMSO-d6/pyridine-d5): δ = 23.24 (C7),

23.26 (C3), 29.21 (C2), 30.94 (C4), 46.61 (C1), 51.62 (C9), 52.04 (C5), 55.41 (OMeA), 55.46 (OMeb), 60.82 (Cγ), 63.00 (Cα), 85.89 (Cβ), 111.66 (CA2), 112.29 (CB6), 115.15 (CA5), 116.99 (CB3), 120.71 (CA6), 120.78 (CB5), 121.57 (CB4), 131.4 (CA1), 145.83 (CA4), 147.58 (CA3), 148.60 (CB1), 149.93 (CB2), 169.56 (C6), 172.96 (C8).

QM-His, 5a (2-tert-Butoxycarbonylamino-3-{3-[3-hydroxy-1-(4-hydroxy-3-methoxy-phenyl)-2- (2-methoxy-phenoxy)-propyl]-3H-imidazol-4-yl}-propionic acid methyl ester) and QM-His, 5b (2-tert-Butoxycarbonylamino-3-{1-[3-hydroxy-1-(4-hydroxy-3-methoxy-phenyl)-2-(2-methoxy-

32 phenoxy)-propyl]-1H-imidazol-4-yl}-propionic acid methyl ester) (Note: Some of the NMR shift assignments for this compound were based on interpretation of the 2D HMQC and HMBC NMR spectra because, as noted above, the QM-His products could not be chromatographically separated from a lignin QM dimer, leading to some shift degeneracy in the 1H and 13C 1D spectra.) Pale white oil (yield: 45% by NMR). Theoretical mass: 571.25 g/mol (+ H+: 572.26 g/mol). Observed m/z + H+: 572.26. 1H NMR (400 MHz, acetone-d6): 1.35 (9H, s, H12), 2.97 (2H, m, H6), 3.44 (1H, m, Hγ), 3.54 (3H, s, H9), 3.56 (1H, m, Hγ), 3.63 (6H, s, OMe), 4.37 (1H, m, H7), 4.96 (1H, m, Hβ), 5.66 (1H, m, Hα), 6.74 (1H, m, HA5), 6.77 (1H, m, HB3), 6.86 (1H, m, HB4), 6.88 (1H, m, HA6), 6.89 (1H, m, HB5), 6.90 (1H, m, H5), 7.06 (1H, m, HB6), 7.11 (1H, m, HA2), 7.55 (1H, s, H5), 7.81 (1H, s, H2). 13C NMR (75.5 MHz, acetone-d6): 27.80 (C12), 29.40 (C6), 51.00 (C9), 54.06 (C7), 55.60 (OMe), 59.90 (Cγ), 60.97 (Cα), 78.40 (C11), 81.45 (Cβ), 111.40 (CB6), 111.60 (CA2), 112.60 (CB5), 115.00 (CA5), 116.56 (C5), 117.30 (C4), 120.20 (CA6), 120.80 (CB3), 122.40 (CB4), 129.80 (CA1), 135.20 (C5), 138.00 (C2), 147.50 (CA3), 147.60 (CB1), 147.80 (CB4), 150.80 (CB2), 155.40 (C10), 172.60 (C8). 1H NMR

(400 MHz, DMSO-d6/pyridine-d5): 1.33 (9H, m, H12), 2.89 (2H, m, H6), 3.46 (3H, s, OMeA), 3.47 (2H, m, Hγ), 3.47 (3H, s, H9), 3.67 (3H, s, OMeB), 4.37 (1H, m, H7), 5.02 (1H, m, Hβ), 5.70 (1H, m, Hα), 6.76 (1H, m, HA5), 6.76 (1H, m, H5), 6.82 (1H, m, HA6), 6.89 (1H, m, HB5), 7.05 (1H, m, HB3), 7.08 (1H, m, HB6), 7.13 (1H, m, HA2), 7.13 (1H, m, HB4), 7.62 (1H, s, H2), 7.81 (1H, d, J = 8.42 Hz, acetyl-0NH), 9.23 (1H, s, A4-OH). 13C NMR (75.5 MHz,

DMSO-d6/pyridine-d5): 28.04 (C12), 29.77 (C6), 51.20 (C9), 53.98 (C7), 55.45 (OMeA), 55.63 (OMeβ), 59.31 (Cγ), 60.20 (Cα), 78.29 (C11), 80.20 (Cβ), 111.62 (CB6), 111.81 (CA2), 112.47 (CB5), 114.80 (C5), 114.83 (CA5), 116.60 (CB3), 120.69 (CA6), 130.00 (CA1), 134.98 (C2), 136.44 (C4), 147.20 (CA4), 147.88 (CB1), 148.05 (CA3), 150.66 (CB2), 155.41 (C10), 172.63 (C8).

QM-Asp, 6 (2-tert-Butoxycaronylamino-succinic acid 1-benzyl ester 4-[3-hydroxy-1-(4- hydroxy-3-methoxy-phenyl)-2-(2-methoxy-phenoxy)-propyl] ester). Pale white oil (yield: 58% after purification). Theoretical mass: 625.25 g/mol (+ Na+: 648.24 g/mol). Observed m/z + Na+: 648.24. Major isomer (74%): 1H NMR (400 MHz, acetone-d6): δ = 1.37 (9H, s, H6), 2.92

(2H, m, H2), 3.6 (1H, m, H3), 3.65 (1H, m, Hγ), 3.75 (1H, m, Hγ), 3.80 (3H, s, OMeB), 3.84 (3H, s, OMeA), 4.61 (1H, m, Hβ), 5.10 (2H, s, H8), 6.06 (1H, d, J = 4.42, Hα), 6.82 (1H, m, HA5), 7.03 (1H, m, HB3), 6.84 (1H, m, HB5), 6.94 (1H, m, HA6), 6.94 (1H, m, HB4), 6.96 (1H, m, HB6), 7.17 (1H, m, HA2), 7.34 (5H, m, H10, H11, H12, H13, H14), 7.65 (1H, s, A4-OH). 13 C NMR (75.5 MHz, acetone-d6): δ = 28.42 (C6), 37.13 (C2), 51.23 (C3), 56.20 (OMeA),

56.16 (OMeB), 61.18 (Cγ), 67.37 (C8), 75.82 (Cα), 79.57 (C5), 83.60 (Cβ), 112.22 (CA2), 113.48 (CB6), 115.14 (CA5), 119.10 (CB3), 121.65 (CA6), 121.65 (CB5), 128.69 (CB4), 128.72 (CA1), 129.18 (C10-C14), 136.81 (C9), 147.32 (CA3), 147.97 (CA4), 151.75 (CB1), 151.80 (CB2), 156.11 (C4), 169.98 (C1), 171.71 (C7). Minor isomer (26%): 1H NMR (400 MHz, acetone-d6): δ = 4.51 (1H, m, Hβ), 6.14 (1H, m, Hα). 13C NMR (75.5 MHz, acetone-d6): δ = 76.41 (Cα), 84.62 (Cβ), 119.27 (CB3), 169.81 (C1), 171.67 (C7). Major isomer (74%): 1H NMR (400 MHz, DMSO-d6/pyridine-d5): δ = 1.33 (9H, s, H6), 2.82 (2H, m, H2), 3.57 (1H, m,

Hγ), 3.66 (1H, m, Hγ), 3.70 (3H, s, OMeB), 3.76 (3H, s, OMeA), 4.55 (1H, m, H3), 4.67 (1H, m, Hβ), 5.09 (2H, s, H8), 6.04 (1H, m, Hα), 6.81 (1H, m, HA5), 6.83 (1H, m, HB3), 6.86 (1H, m, HB5), 6.89 (1H, m, HA6), 6.91 (1H, m, HB6), 7.09 (1H, m, HA2), 7.11 (1H, m, HB4), 7.37 (5H, m, H10, H11, H12, H13, H14), 7.48 (1H, d, J = 3.42, NH), 9.36 (1H, s, A4-OH). 13C NMR (75.5

MHz, DMSO-d6/pyridine-d5): δ = 27.98 (C6), 35.60 (C2), 50.24 (C3), 55.44 (OMeA), 55.54

(OMeB), 59.63 (Cγ), 65.98 (C8), 74.75 (Cα), 78.45 (C5), 81.39 (Cβ), 111.96 (CA2), 112.76 (CB6), 114.84 (CA5), 116.77 (CB4), 120.69 (CA6), 120.69 (CB3), 120.69 (CB5), 127.19 (CA1), 127.53 (C12), 127.66 (C13), 127.78 (C11), 127.95 (C14), 128.29 (C10), 135.98 (C9), 144.32 (C4), 146.65 (CA4), 147.30 (CA3), 147.67 (CB1), 150.12 (CB2), 169.12 (C1), 171.79 (C7). Minor isomer (26%): 1H NMR (400 MHz, DMSO-d6/pyridine-d5): δ = 4.55 (1H, m, Hβ), 6.11 (1H, m, Hα). 13C NMR (75.5 MHz, DMSO-d6/pyridine-d5): δ = 75.08 (Cα), 82.70 (Cβ), 112.58 (CB6), 116.87 (CB4), 147.27 (CA3), 150.16 (CB2), 168.92 (C1).

QM-Glu, 7 (2-tert-Butoxycaronlyamino-pentanedioic acid 1-tert-butyl ester 5-[3-hydroxy-1-(4- hyroxy-3-methoxy-phenyl)-2-(2-methoxy-phenoxy)-propyl] ester). Pale white oil (yield: 47% after purification). Theoretical mass: 605.28 g/mol (+ Na+: 628.27 g/mol). Observed m/z + Na+: 628.25. Major isomer (74%): 1H NMR (400 MHz, acetone-d6): δ = 1.42 (9H, s, H10), 1.46 (9H, s, H7), 1.92, 2.08 (2H, m, H3), 2.47 (2H, t, J = 8.02, H2), 3.69, 3.79 (2H, m, Hγ), 3.84 (3H, s, OMeB), 3.88 (3H, s, OMeA), 4.08 (1H, m, H4), 4.62 (1H, m, Hβ), 5.78 (1H, s, γ-OH), 6.06 (1H, d, J = 5.11, Hα), 6.79 (1H, m, HA5), 6.85 (1H, m, HB5), 6.94 (1H, m, HA6), 6.94 (1H, m, HB4), 6.96 (1H, m, HB6), 7.02 (1H, m, HB3), 7.17 (1H, m, HA2). 13C NMR (75.5 MHz, acetone-d6): δ = 27.69 (C3), 28.07 (C10), 28.49 (C7), 31.19 (C2), 54.57 (C4), 56.21 (OMeA), 56.21 (OMeB), 61.27 (Cγ), 75.23 (Cα), 79.17 (C9), 81.46 (C6), 83.89 (Cβ), 112.29 (CA2), 113.55 (CB6), 115.15 (CA5), 119.02 (CB3), 121.68 (CB5), 121.68 (CA6), 123.31 (CB4), 129.32 (CA1), 147.32 (CA4), 147.96 (CA3), 148.97 (CB1), 151.78 (CB2), 156.40 (C5), 171.93 (C1), 174.01 (C8). Minor isomer (26%): 1H NMR (400 MHz, acetone-d6): δ = 4.53 (1H, m, Hβ), 6.13 (1H, m, Hα). 13C NMR (75.5 MHz, acetone-d6): δ = 75.93 (Cα), 84.85 (Cβ), 115.53

(CA5), 119.26 (CB3), 129.72 (CA1), 147.53 (CA4), 148.20 (CA3), 172.10 (C1). Major isomer (74%): 1H NMR (400 MHz, DMSO-d6/pyridine-d5): δ = 1.34 (9H, s, H10), 1.36 (9H, s, H7),

1.83 (2H, m, H3), 2.39 (2H, m, H2), 3.58, 3.66 (2H, m, Hγ), 3.71 (3H, s, OMeB), 3.76 (3H, s,

OMeA), 3.93 (1H, m, H4), 4.67 (1H, m, Hβ), 5.10 (1H, s, γ-OH), 6.02 (1H, m, Hα), 6.81 (1H, m, HA5), 6.82 (1H, m, HB3), 6.85 (1H, m, HB5), 6.89 (1H, m, HA6), 6.92 (1H, m, HB6), 7.08 (1H, m, HA1), 7.10 (1H, m, HB4), 7.28 (1H, d, J = 7.86, NH), 9.34 (1H, s, A4-OH). 13C NMR (75.5 MHz, DMSO-d6/pyridine-d5): δ = 26.04 (C3), 27.48 (C10), 28.05 (C7), 30.36 (C2), 53.60

(C4), 55.45 (OMeA), 55.57 (OMeB), 59.71 (Cγ), 74.23 (Cα), 78.09 (C6), 80.37 (C9), 81.58 (Cβ), 111.91 (CA2), 112.77 (CB6), 114.90 (CA5), 116.73 (CB4), 120.02 (CB3), 120.56 (CB5), 120.70 (CA6), 127.53 (CA1), 146.61 (CA4), 147.30 (CA3), 147.72 (CB1), 150.13 (CB2), 155.65 (C5), 171.20 (C8), 171.38 (C1). Minor isomer (26%): 1H NMR (400 MHz, DMSO-d6/pyridine-d5): δ = 1.97 (2H, m, H3), 4.56 (1H, m, Hβ), 6.08 (1H, m, Hα). 13C NMR (75.5 MHz, DMSO- d6/pyridine-d5): δ = 26.31 (C3), 30.13 (C2), 53.86 (C4), 74.68 (Cα), 78.00 (C6), 80.24 (C9), 82.64 (Cβ), 111.35 (CA2), 115.21 (CA5), 128.32 (CA1), 146.72 (CA4), 147.47 (CA3), 148.42 (CB1), 149.98 (CB2), 154.74 (C5), 171.65 (C1).

QM-Ser, 8 (2-Benzyloxycarbonylamino-3-[3-hydroxy-1-(4-hydroxy-3-methoxy-phenyl)-2-(2- methoxy-phenoxy)-propoxy]-propionic acid methyl ester). Pale white oil (39% after purification). Theoretical mass: 555.21 g/mol (+ Na+: 578.20 g/mol). Observed m/z + Na+: 578.20. Major isomer (68%): 1H NMR (400 MHz, acetone-d6): δ = 3.65 (2H, m, H1), 3.70

(3H, s, H4), 3.76 (3H, s, OMeB), 3.81 (3H, s, OMeA), 3.81 (2H, m, Hγ), 4.32 (1H, m, Hβ), 4.43 (1H, m, H2), 4.60 (1H, m, Hα), 5.09 (2H, m, H6), 6.80 (1H, m, HB6), 6.82 (1H, m, HB3), 6.88 (1H, m, HA5), 6.89 (1H, m, HA6), 6.89 (1H, m, HB4), 6.92 (1H, m, HB5), 7.00 (1H, m, HA2), 7.15, (1H, m, H10), 7.38 (5H, m, H8, H9, H11, H12). 13C NMR (75.5 MHz, acetone-d6): δ = 52.35 (C4), 55.59 (C2), 56.20 (OMe), 61.74 (Cγ), 66.87 (C6), 69.92 (C1), 82.25 (Cα), 85.28 (Cβ), 111.70 (C10), 112.00 (CA2), 113.49 (CB5), 115.23 (CB6), 119.27 (CA5), 121.73 (CA6), 122.03 (CB3), 123.09 (CB4), 128.67 (C8, C12), 129.22 (C9, C11), 130.62 (CA1), 138.05 (C7), 147.23 (CA4), 148.17 (CA3), 149.20 (CB1), 151.76 (CB2), 157.06 (C5), 171.87 (C3). Minor 1 isomer (32%): H NMR (400 MHz, acetone-d6): δ = 3.72 (2H, m, Hγ), 3.79 (3H, s, OMeB), 13 3.83 (3H, s, OMeA), 4.55 (1H, m, Hα). C NMR (75.5 MHz, acetone-d6): δ = 55.42 (C2), 61.33 (Cγ) 69.47 (C1), 82.40 (Cα), 85.68 (Cβ), 111.94 (CA2), 119.15 (CA5), 130.19 (CA1), 147.29 (CA4), 148.95 (CB1), 156.95 (C5). Major isomer (68%): 1H NMR (400 MHz, DMSO- d6/pyridine-d5): δ = 3.39 (1H, m, Hγ), 3.56-3.65 (2H, m, H1), 3.65 (3H, s, H4), 3.67 (1H, m,

Hγ), 3.65 (3H, s, OMeB), 3.72 (3H, s, OMeA), 4.41 (1H, m, H2), 4.46 (1H, m, Hβ), 4.59 (1H, m, Hα), 5.08 (2H, s, H6), 6.73-6.83 (1H, m, HA5), 6.74-6.80 (1H, m, HA6), 6.80-6.87 (1H, m HB4), 6.81 (1H, m, HB6), 6.82-6.93 (1H, m, HB5), 6.94-7.07 (1H, m, HA2), 6.97-7.06 (1H, m, HB3), 7.25-7.38 (5H, m, H7-H12), 7.81 (1H, d, J = 8.42, NH), 9.23 (1H, s, A4-OH). 13C NMR

(75.5 MHz, DMSO-d6/pyridine-d5): δ = 54.20 (C1), 55.29 (OMeA), 55.38 (OMeB), 51.81 (C4) 60.08 (Cγ), 65.80 (C6), 67.90 (C1), 80.91 (Cα), 82.50 (Cβ), 111.48 (CA2), 112.75 (CB5), 114.82 (CA5), 115.06 (CB6), 116.38 (CB4), 120.57 (CA6), 120.59 (CB3), 127.80-128.38 (C7-C12), 128.72 (CA1), 137.05 (C7), 146.36 (CA4), 147.84 (CB1), 149.55 (CB2), 149.93 (CA3), 156.30 (C5), 170.81 (C3). Minor isomer (32%): 1H NMR (400 MHz, DMSO-d6/pyridine-d5): δ =

4.42 (1H, m, Hβ), 4.55 (1H, m, Hα). 13C NMR (75.5 MHz, DMSO-d6/pyridine-d5): δ = 80.69 (Cα), 82.90 (Cβ), 111.84 (CA2), 115.90 (CB4), 128.54 (CA1).

QM-Tyr, 9 (2-tert-Butoxycarbonlyamino-3-{4-[3-hydroxy-1-(4-hydroxy-3-methoxy-phenyl)-2- (2-methoxy-phenoxy)-propoxy]-phenyl}-propionic acid methyl ester). Pale white oil (yield: 45% after purification). Theoretical mass: 597.26 g/mol (+ H+: 598.27 g/mol). Observed m/z + H+: 598.29. 1H NMR (300 MHz, acetone-d6): δ = 1.33 (9H, d, J = 3.28 Hz, H13), 2.88 (1H, m, H7),

3.00 (1H, m, H7), 3.62 (3H, d, J = 3.74, H10), 3.78 (3H, s, OMeB), 3.81 (3H, s, OMeA), 3.81 (1H, m, Hγ), 3.91 (1H, m, Hγ), 4.31 (1H, m, H8), 4.55 (1H, m, Hβ), 5.13 (1H, d, J = 5.13, Hα), 6.07 (1H, d, J = 7.66, γ-OH), 6.78 (1H, m, HA5), 6.81 (1H, m, HB5), 6.87 (2H, m, H3, H5), 6.93 (1H, m, HB4), 6.95 (2H, m, HB3, HB6), 6.97 (1H, m, HA6), 7.06 (2H, m, H2, H6), 7.17 (1H, m, HA2), 7.55 (1H, s, Ph-OH). 13C NMR (100 MHz, acetone-d6): δ = 28.42 (C13), 37.29 (C7), 52.07 (C10), 56.02 (C8), 56.17 (OMe), 61.19 (Cγ), 79.22 (C12), 79.42 (Cα), 85.40 (Cβ), 112.09 (CA2), 113.50 (CB6), 115.29 (CA5), 116.81 (C3, C5), 119.21 (CB5), 121.43 (CA6), 121.70 (CB3), 123.22 (CB4), 130.32 (CA1), 130.82 (C2, C4, C6), 147.12 (CA4), 148.13 (CA3), 149.10 (CB1), 151.82 (CB2), 156.10 (C11), 157.77 (C1), 173.22 (C9). 1H NMR (400 MHz, DMSO- d6/pyridine-d5): δ = 1.27 (9H, s, H13), 2.71-2.95 (2H, m, H7), 3.62 (3H, s, H4), 3.67 (3H, s,

OMeB), 3.71 (3H, s, OMeA), 3.73 (1H, m, Hγ), 3.55 (3H, s, H10), 3.79 (1H, m, Hγ), 4.18 (1H, m, H8), 4.66 (1H, m, Hβ), 5.08 (1H, s, γ-OH), 5.49 (1H, d, J = 4.20, Hα), 6.76 (1H, m, HA5), 6.81 (3H, m, HB3, H3, H5), 6.84 (1H, m, HB5), 6.89 (2H, m, HA6, HB6), 7.05 (2H, m, H2, H6), 7.06 (1H, m, HB4), 7.11 (1H, m, HA2), 9.21 (1H, s, A4-OH). 13C NMR (75.5 MHz, DMSO- d6/pyridine-d5): δ = 28.10 (C13), 35.50 (C7), 51.60 (C10), 55.47 (OMeB), 55.50 (C8), 55.60

(OMeA), 59.78 (Cγ), 78.26 (C12), 78.30 (Cα), 82.69 (Cβ), 112.02 (CA2), 112.81 (CB6) 115.01 (CA5), 115.79 (C3, C5), 116.43 (CB4), 120.73 (CA6), 120.75 (CB3), 121.57 (CB5), 128.42 (CA1), 129.65 (C2, C6), 129.96 (C4), 146.35 (CA4), 147.37 (CA3), 148.03 (CB1), 150.06 (CB2), 156.30 (C1), 172.56 (C9).

QM-Thr, 10 (2-tert-Butoxycarbonylamino-3-[3-hydroxy-1-(4-hydroxy-3-methoxy-phenyl)-2-(2- methoxy-phenoxy)-propoxy]-butyric acid methyl ester). Not experimentally observed. DFT- calculated 1H NMR (syn-isomer, DMSO force field): 0.8 (H1), 3.2 (Hβ), 3.3 (Hγ), 3.3 (H3),

3.5 (OMeB), 3.6 (Hγ), 3.6 (H2), 3.6 (OMeA), 4.4 (Hα), 6.1 (HB6), 6.7 (HA5), 6.7 (HB3), 6.7 (HB5), 6.9 (HA2), 6.9 (HA6), 7.0 (HB4). DFT-calculated 13C NMR (syn-isomer, DMSO force field): 15.2 (C1), 52.1 (OMeA), 52.1 (OMeB), 55.9 (Cγ), 60.7 (C3), 69.2 (C2), 70.7 (Cα), 88.5 (Cβ), 109.0 (CA2), 112.4 (CB3), 113.4 (CA5), 121.5 (CB5), 123.6 (CA6), 124.5 (CB6), 125.7 (CB4), 131.4 (CA1), 144.5 (CA4), 145.8 (CA3), 145.9 (CB1), 151.9 (CB2). DFT-calculated 1H

NMR (anti-isomer, DMSO force field): 0.9 (H1), 3.1 (Hγ), 3.2 (H3), 3.3 (H2), 3.3 (OMeA), 3.4 (Hβ), 3.5 (OMeB), 3.9 (Hγ), 4.3 (Hα), 4.8 (HB6), 6.4 (HB5), 6.6 (HB3), 6.8 (HA5), 6.8 (HA6), 6.8 (HB4), 7.2 (HA2). DFT-calculated 13C NMR (anti-isomer, DMSO force field): 51.9

(OMeB), 52.6 (OMeA), 73.7 (Cα), 60.9 (Cγ), 90.4 (Cβ), 112.1 (CB3), 113.9 (CA5), 114.3 (CA2), 121.2 (CA6), 121.5 (CB5), 122.4 (CB6), 123.8 (CB4), 130.1 (CA1), 144.8 (CA4), 146.1 (CA3), 149.3 (CB2), 149.3 (CB1).

QM-Hyp, 11 (4-[3-Hydroxy-1-(4-hydroxy-3-methoxy-phenyl)-2-(2-methoxy-phenoxy)- propoxy]-pyrrolidine-1,2-dicarboxylic acid 1-tert-butyl ester 2-methyl ester). Not experimentally 1 observed. DFT-calculated H NMR (syn-isomer, DMSO force field): 3.2 (Hβ), 3.5 (OMeB), 3.6 (OMeA), 4.7 (Hα), 3.4 (Hγ), 3.5 (Hγ), 6.0 (HB6), 6.1 (H2), 6.6 (HA5), 6.7 (HB3), 6.7 (HB5), 6.8 (HA6), 7.0 (HB4), 7.2 (HA2), 7.2 (H4). DFT-calculated 13C NMR (syn-isomer, DMSO force field): 52.0 (OMeB), 52.2 (OMeA), 56.3 (Cγ), 80.5 (Cα), 87.7 (Cβ), 105.1 (C3), 109.8 (CA2), 112.4 (CB3), 112.5 (C1), 113.0 (CA5), 119.1 (C4), 121.6 (CB5), 122.3 (CA6), 124.5 (CB6), 125.5 (CB4), 132.7 (CA1), 144.1 (CA4), 145.3 (CA3), 147.1 (CB1), 147.6 (C2), 151.2 1 (CB2). DFT-calculated H NMR (anti-isomer, DMSO force field): 3.4 (OMeA), 3.4 (Hγ), 3.5 (OMeB), 3.6 (Hβ), 3.7 (Hγ), 4.0 (H2), 4.6 (Hα), 4.8 (HB6), 6.1 (H4), 6.5 (HB5), 6.6 (HB3), 6.8 (HA5), 6.8 (HA6), 6.9 (HB4), 7.2 (HA2). DFT-calculated 13C NMR (anti-isomer, DMSO

41 force field): 51.9 (OMeB), 52.4 (OMeA), 60.9 (Cγ), 78.7 (Cα), 90.9 (Cβ), 95.7 (C3), 99.8 (C1), 112.2 (CB3), 112.3 (C4), 113.0 (CA2), 113.9 (CA5), 119.5 (CA6), 121.8 (CB5), 122.4 (CB6), 124.1 (CB4), 131.8 (CA1), 144.5 (CA4), 145.9 (CA3), 139.0 (C2), 149.1 (CB1), 149.2 (CB2).

2.3.4. Nuclear magnetic resonance spectroscopy

NMR spectra were collected in both acetone-d6 (spectra shown above) and DMSO- d6/pyridine-d5 (4:1 v/v, 500 ul). DMSO-d6/pyridine-d5 was chosen because it is a preferred solvent for NMR of lignin DHP, milled wood lignin (MWL), and whole cell walls; using the same solvent system allows for accurate shift comparisons (Kim and Ralph 2010). In general, negligible shift migration was observed between the two solvent systems. NMR spectra were acquired on Bruker DPX-300 (300 MHz 1H resonance freq.), DRX-400 (400 MHz 1H resonance freq.), AV-III-500 (500 MHz 1H resonance freq.) with a cryogenically-cooled probe and inverse probe geometry (i.e. proton coils closest to sample), AV-III-600 (500 MHz 1H resonance freq.) with a cryogenically-cooled probe, and AV-III-850 (850 MHz 1H resonance freq.) with a cryogenically-cooled probe. Spectral processing was performed in Bruker's Topspin 3.1 software. Standard Bruker pulse programs were employed: 1H (8-16 scans), 13C (5k-10k scans), HMQC (Bruker pulse program 'inv4gptp’, 64 scans), and HMBC (Bruker pulse program 'inv4gslplrnd’, 64 scans). Spectra were calibrated to the central solvent peaks (acetone: 2.05/29.8 ppm; dimethyl sulfoxide: 2.50/39.5 ppm). In the case of lignin DHP, NMR spectra were acquired on a Bruker Biospin (Billerica, MA, USA) AVANCE 500 (500 MHz 1H resonance freq.) spectrometer fitted with a cryogenically-cooled probe having inverse geometry, i.e., with the proton coils closest to the sample. Spectra were processed with Bruker’s Topspin 3.1 software, using the central solvent peak as internal reference (δH/δC: dimethyl sulfoxide (DMSO), 2.50/39.5 ppm). The synthetic lignin DHP (~50 mg) was placed in an NMR tube (ID: 4.1 mm), swollen homogeneously in DMSO-d6/pyridine-d5 (4:1 v/v, 500 ul) with the aid of ultrasonication (~3h), and then subjected to adiabatic 2D-HSQC (‘hsqcetgpsisp2.2’) experiments using the parameters described by Mansfield et al. (2012). Processing used typical matched Gaussian apodization in F2 (LB = -0.3, GB = 0.001), and squared cosine-bell and one level of linear prediction (32 coefficients) in F1 (Mansfield et al. 2012).

2.3.5. Mass spectrometry Exact masses for compounds 3-6 (see online resource) were calculated using ChemBioDraw Ultra 13.0. Mass spectrometric analysis was performed on a Waters LCT Premier time-of-flight (TOF) mass spectrometer (Waters Corporation (Micromass Ltd.), Manchester, UK), using MassLynx™ software Version 4.0. Samples were introduced using a Waters 2695 high performance liquid chromatograph. Sample analysis utilized flow injection analysis (FIA). The mobile phase used was 90% acetonitrile (LC-MS grade) and 10% aqueous ammonium acetate (10mM). The flow rate was 0.25 mL/min. The nitrogen drying gas temperature was set to 300 °C at a flow of 7 L/min. The capillary voltage was 2200 V. The mass spectrometer was set to scan from 100-1000 m/z in positive ion mode, using electrospray ionization (ESI).

2.3.6. Computational methods

Eight conformational isomers of QM-Cys, QM-Thr, and QM-Hyp, and sixteen conformational isomers of QM-His were built using Materials Studio 6.0 (Accelrys Inc., San

Diego, CA). Eight of the QM-His models exhibited a CαQM-N1His bond and eight models exhibited CαQM-N3His bond; these models allowed us to determine which CαQM-NHis bond was occurring and to determine if an observed chemical shift (α13C) at 78.8 ppm was due to a C-N bond. Each set of eight models (i.e., compound 3 (QM-Cys), compound 5a (QM-His(N3)), compound 5b (QM-His(N1)), compound 10 (QM-Thr), or compound 11 (QM-Hyp)) contained two of each of the stereoisomers (R,R), (SS), (R,S), and (S,R), where the former two stereoisomers are syn and the latter two stereoisomers are anti. These models were built to determine if the calculated NMR chemical shifts could differentiate the observed shifts for the syn and anti stereoisomers of QM-His and QM-Cys. Experimental NMR shifts for QM-Thr and QM-Hyp were not obtained because Thr and Hyp did not react with the QM; however, we reported the calculated shifts for these compounds (below) as potential references for other researchers to use.

Each model was energy minimized without symmetry or atomic constraints using the density functional theory (DFT) method M05-2X, coupled with the 6-311++G(2df,2p) basis set using the program Gaussian 09 (Curtiss et al. 2001; Frisch et al. 2009; Hohenberg and Kohn 1964; Kohn and Sham 1965; Zhao et al. 2006). Following the geometry optimization calculations, frequency calculations assured that each model attained a potential energy surface (PES) minimum, where no imaginary frequencies were present (Frisch et al. 2009).

Subsequent gauge-independent atomic orbital (GIAO) calculations using Gaussian 09 at the mPW1PW91/6-31G(d) theory level provided the NMR magnetic shielding tensors (α13C and α1H) for the energy-minimized structures (Adamo and Barone 1998; Buhl et al. 1999; Cheeseman et al. 1996; Karadakov 2008; Lodewyk et al. 2012; Schreckenbach and Ziegler 1995; Wolinski et al. 1990). Because our experiments were conducted in dimethylsulfoxide (DMSO), the GIAO calculations were also performed in a dielectric continuum of DMSO using a self- consistent reaction field (SCRF) and the integral equation formalism variant of the polarized continuum model (IEFPCM) (Cances et al. 1997; Gogonea 1998). Note that the structures were not energy minimized within the polarized continuum because prior work showed that doing so did not improve the precision of the calculations (Watts et al. 2011). A multi-standard NMR method using benzene for sp2-hybridized C- and H-atoms, and methanol for sp3-hybridized C- and H-atoms led to the α13C and α1H results (Sarotti and Pellegrinet 2009; Sarotti and Pellegrinet 2012; Watts et al. 2011). Benzene and methanol were energy minimized using M05-2X/6- 311++G(2df,2p) and underwent subsequent GIAO calculations using mPW1PW91/6-31G(d).

The precision of the multi-standard method versus the single-standard method (e.g., tetramethylsilane as the standard) is illustrated when comparing single-standard results recently reported by Mostaghni et al. (2013) with the multi-standard results of Watts et al. (2011). Both

43 groups reported the δ13C for β-O-4 linkages in lignin model compounds; however, the mean unsigned errors, root-mean-squared errors, and maximum errors reported by Mostaghni et al. (2013) were approximately 10, 12, and 23 ppm, while those reported by Watts et al. (2011) were approximately 2, 3, and 8 ppm. Therefore, the multi-standard method produced results that were more precise than those produced by the single-standard method for lignin model compounds with β-O-4 linkages.

x For each C- or H-nucleus, we used δ calc= σref - σcalc + δref to calculate the chemical shift x of each H- and C-nucleus of interest (δ calc) in the GG-amino acid models (Sarotti and Pellegrinet

2009; Sarotti and Pellegrinet 2012). Here, σref is the calculated tensor of the C- or H- nucleus of the standard (i.e., methanol or benzene), σcalc is the calculated tensor of the nucleus of interest from the GG-amino acid model, and δref is the experimental chemical shift of the C- and H- nuclei in benzene or methanol dissolved in DMSO (Gottlieb et al. 1997). The chemical shifts for each C- and H-nucleus was thermodynamically weighted using the relative, calculated Gibbs free energy of each model to account for the thermodynamic abundance of each model (Barone et al. 2002). The calculated δ13C and δ1H results were then correlated with their respective NMR data.

2.4. Results and discussion

2.4.1. Preparation of quinone methide-amino acid adducts

A lignin β-ether QM 2 was prepared cleanly from guaiacylglycerol-β-guaiacyl ether 1, as previously described (Kawai et al. 1999; Landucci et al. 1981; Ralph and Young 1983). One of nine amino acids bearing a nucleophilic side-group was then added to the QM, with each reaction monitored by thin layer chromatography. It was observed that amino acids with amine- containing side-chains (Lys and His) reacted with the QM quickly (within minutes), whereas thiol-, acid-, and hydroxyl-containing amino acids reacted slowly (over hours or days). In the case of the secondary hydroxyl-containing amino acids (Thr and Hyp) no cross-coupling was observed (i.e., compounds 10 and 11 did not form), despite attempts to catalyze the cross- coupling reaction (refer to the electronic supplement for detailed reaction protocols). Products were purified via column chromatography and yields ranged from quantitative in the case of compound 3 (QM-Lys) to zero (no reaction) in the cases of compounds 10 and 11 (QM-Thr and QM-Hyp). Cross-coupling reactions were carried out in dichloromethane to produce the desired lignin-protein adducts.

Fig 2.3. QM-AA model compounds. Lignin-cysteine (QM-Cys) 3, lignin-lysine (QM-Lys) 4, lignin-histidine (QM-His) 5, lignin-aspartic acid (QM-Asp) 6, lignin-glutamic acid (QM-Glu) 7, lignin-serine (QM-Ser) 8, lignin-Tyrosine (QM-Tyr) 9, lignin-threonine (QM-Thr) 10, and lignin-hydroxyproline (QM-Hyp) 11 adducts derived from QM 2

2.4.2. Solution-state NMR of compounds 3-9 and density functional theory calculations for compounds 10 and 11

Reaction products were characterized using solution-state 1D 1H and 13C NMR, as well as 2D heteronuclear multiple quantum coherence (HMQC) and heteronuclear multiple-bond correlation (HMBC) experiments. Full spectral assignments for compounds 3-9 are given in the methods sections (sections 2.3.3 and 2.3.4). Interpretation of these results is consistent with structures 3-9 (Fig 2.3), indicating that Cys, Lys, His, Asp, Glu, Ser and Tyr all add to QM 2 in vitro. Density functional theory (DFT) was used to predict NMR shifts for compounds 10 (QM- Thr) and 11 (QM-Hyp), which did not form under the synthetic conditions employed here.

Table 2.1 shows the lignin α and β 1H and 13C shifts for compounds 3-11. The γ-shifts of these compounds are almost entirely degenerate and are therefore considered non-diagnostic. Because threonine and hydroxyproline are abundant in cell wall structural proteins (especially hyp, which can account for up to 33% of the amino acid profiles of some structural proteins), the authors perceived that estimations of the QM-Thr and QM-Hyp NMR chemical shifts could still be useful. Thus, NMR shifts for compounds 10 and 11 were calculated using DFT. As a control, DFT was also used to calculate NMR shifts for compounds 3 and 5 (Fig 2.4), showing comparison to experimental results. Calculated 13C shifts were generally in agreement with experimentally observed shifts. For example, calculated 13C α-shifts overestimated the observed shifts by only 0.8-3.1 ppm. Calculated 13C β-shifts overestimated the observed shifts by 5.4-9.8 ppm. Similar discrepancies in DFT calculated β-shifts of β-ether compounds have been previously reported, and further work is necessary to refine these calculations (Watts et al. 2011). Calculated 1H shifts consistently underestimated the experimentally observed shifts by about 0.5- 1 ppm. Thus, the calculated 1H shifts for compounds 10 and 11 are not reproducing the observed 1H shifts; however it could be possible with future work to develop a method to correlate the calculated and experimental 1H shifts, because of the consistent underestimation of the experimental 1H shifts by the calculated shifts. Lodewyk et al. (2012) described a method for using empirical scaling factors to obtain improved correlation between experimental and calculated 1H and 13C shifts; however, doing so is beyond the scope of the present work. In addition to the use of scaling factors, further research to develop multi-standard methods that are based on DFT results is necessary. This work could require the development and assessment of DFT methods, as well as basis sets to obtain methods to calculate 1H shifts more precisely.

Table 2.1. 1H and 13C NMR chemical shifts for lignin-amino acid adducts.

α-shifts β-shifts Experimental Calculated Experimental Calculated Compound 1H/13C 1H/13C 1H/13C 1H/13C 4.3/50.3 3.8/53.4 4.7/81.8 3.7/89.6 3 (QM-Cys)a 4.4/50.6 3.5/52.9 4.6/81.7 4.0/87.1 4 (QM-Lys) 3.9/63.0 4.2/85.9 5 (QM-His)b 5.7/60.2 5.0/61.0 5.0/80.2 3.9/90.0 6.0/74.8 4.7/81.4 6 (QM-Asp)a 6.1/75.1 4.6/82.7 6.0/74.2 4.7/81.6 7 (QM-Glu)a 6.1/74.7 4.6/82.6 4.6/80.9 4.4/82.5 8 (QM-Ser)a 4.6/80.7 4.4/82.9 9 (QM-Tyr) 5.5/78.3 4.7/82.7 4.4/70.7 3.2/88.5 10 (QM-Thr)c n/a n/a 4.3/73.7 3.4/90.4 4.7/80.5 3.2/87.7 11 (QM-Hyp)c n/a n/a 4.6/78.7 3.6/90.9

Key: a, products exhibited two stereoisomers, shifts for the major isomer are shown first; b, only the calculated shifts of anti-5b are shown, see the electronic supplement for calculated shifts of additional isomers of 5; c, syn-isomer shifts are shown first.

Fig 2.4. Overlaid HMQC side chain regions of compounds 3 and 5. The α- and β-shifts are labeled; methoxyl and γ-shifts are not labeled due to substantial shift degeneracy. Grey shifts are non-diagnostic. DFT calculated α-shifts (red squares) and β-shifts (blue squares) are shown for compounds 3, 5a and 5b (both threo and erythro stereoisomers are shown). Calculated 13C shifts correlate relatively well with experimentally observed 13C shifts, though not well enough to allow for assignment of stereochemistry in the experimentally observed product shifts. Calculated 1H shifts are underestimated by about 0.5-1.0 ppm. Further research is necessary to refine the predicative abilities of DFT for 1H shifts of lignin compounds.

Fig 2.5 highlights the location of diagnostic HMQC NMR peak contours of the lignin-amino acid adducts overlaid on the spectrum of a synthetic lignin (a so-called dehydrogenation polymer, or DHP). Differences in chemical shifts among the lignin-amino acid adducts are most salient for the α-positions and, as expected, less for those from the β-positions. Most of the lignin-amino acid shifts are readily distinguishable from correlations of native structures in lignin; however, the α-shifts of compound 4 (QM-Lys) are degenerate with phenylcoumaran γ- shifts. In this case, identifying a lignin-lysine crosslink may be possible by observing the lignin- lysine β-shifts. The α- and β-shifts of compound 9 (QM-Tyr) are degenerate with benzyl aryl ether linkages (so called α-O-aryl linkages) sometimes observed in synthetic and native lignin polymers. These lignin-lignin linkages form when QMs are quenched by phenolic moieties, and degeneracy is not surprising given the structural similarities among tyrosine and the lignin monomers, p-coumaryl, coniferyl, and sinapyl alcohols. This may make it difficult to distinguish lignin-tyrosine crosslinking from lignin-lignin α-O-aryl linkages in native lignins.

Though not depicted graphically, the lignin-peptide linkages described herein are largely free from overlap with previously described polysaccharide shifts in both angiosperms and

48 gymnosperms. However, a few of the lignin-amino acid shifts may overlap with signatures attributed to lignin-carbohydrate linkages. For example, the α-shifts of compounds 6 and 7 exhibit degeneracy with lignin-carbohydrate benzyl esters (α-shifts at 6.1/75.0 ppm) due to structural similarity (Balakshin et al. 2011; Toikka et al. 1998). Likewise, the α-shifts of 8 and 11 exhibit degeneracy with lignin-carbohydrate benzyl ethers (α-shifts located at 4.6/80.5 ppm) (Balakshin et al. 2011; Toikka et al. 1998). Thus, caution should be exercised when attempting to discern certain lignin-protein and lignin-carbohydrate linkages using 1D and 2D NMR techniques. The results of the current study indicate that NMR identification of lignin-protein linkages, especially linkages of the benzyl thioether and benzyl amine types, should be possible in whole cell walls or lignin extracts provided the linkages are adequately abundant (Kim and Ralph 2010; Mansfield et al. 2012).

Fig 2.5. HSQC NMR spectrum of a lignin DHP with overlaid α- and β-correlation data from compounds 3-11 represented by red (α) and blue squares (β)

2.4.3. Adduct isomer determination

Of purely fundamental interest, we attempted to resolve the stereochemistry of the products by the use of DFT, but these efforts were largely unsuccessful. For example, in the case of QM- Cys, 3, the root mean-squared error (RMSE) between experimental and calculated shifts was too large to reliably assign the isomers (Table 2.2). Although it may have been possible to improve the DFT results through the addition of conformational isomers, the added computational cost may not have reduced the calculated RMSE to experimental uncertainty levels. Hence additional attempts to resolve stereoisomers (compounds 6, 7, 8) were abandoned; likewise, DFT was not used to identify which stereoisomer was produced in 4 and 9 (only one product was observed in each case). Previously, addition of primary amines were shown (via diagnostic NMR of tetrahydro-1,3-oxazine derivatives) to strongly (>90%) favor formation of the syn-isomer (Ralph and Young 1983), so product 4 is likely syn.

Table 2.2. Observed and DFT calculated α-13C NMR chemical shifts for compound 3.

α-13C chemical shifts (ppm) Observed Calculated RMSE 50.27 52.90 - (3, syn) 2.7 50.60 53.40 - (3, anti) 2.8

In the case of 5 (QM-His), one α-shift was observed, occurring at 5.7/60.2 ppm. The His system is an interesting one to consider given the tautomerization in the His imidazole group and the potential for various regio-isomeric products (Nagy et al. 2005). In the HMQC and HMBC spectra (see methods section) the α-1H shows correlations to positions 2 and 5 of the imidazole ring, though α-1H correlations to position 5 are weak and partially degenerate with correlations to carbon A6. The NMR results suggest the formation of both compounds 5a and 5b, resulting from either N1 or N3 addition, but quantification of these compounds via NMR was rendered impossible due to the aforementioned shift degeneracy. The Gibbs free energy-based Boltzmann factors in the gas-phase suggested that compound 5b is thermodynamically prevalent relative to compound 5a (93.5% to 6.5%, respectively), and prior work by Watts et al. (2011) suggested that models with greater thermodynamic abundance generally provided α-13C results that were better correlated with experimental NMR data.

2.5. Conclusions

This study is the first to report on the synthesis of lignin-protein model compounds and contributes to the growing lignin NMR database. QM-amino acid adducts were synthesized and characterized. Namely, Cys, Lys, His, Asp, Glu, Ser, Tyr, Thr, and Hyp were reacted with a lignin model quinone methide—an important intermediate in lignification. The selected quinone methide 2 represents the structure and reactivity of QMs native to lignin. The amino acids were

50 selected because of their nucleophilic side-groups; furthermore, these amino acids are common in plant cell wall structural proteins and represent functional groups (amines, thiols, acids, and alcohols) that are known to react with quinone methides (Awad et al. 2000; Bolton et al. 1997; Ramakrishnan and Fisher 1983). The selected amino acids quenched the QM with varying efficiencies (in general, amine > thiol > acid > hydroxyl) under neutral organic solvent conditions. The secondary alcohols (Thr, Hyp) did not react under the selected conditions.

Using the results from these model compounds to identify any lignin-protein crosslinks in planta is our goal. Based on the results herein, lignin-protein NMR shifts should be well dispersed and, in most cases, distinct even within the complex NMR spectra of polymerized lignin (Fig 2.5). This suggests that the linkages may be detectable in planta if they exist in significant quantities.

Although density functional theory was used to predict NMR chemical shifts of lignin- protein crosslinks, the calculated chemical shifts did not display the level of accuracy required to distinguish stereoisomers. Future studies are needed to improve the correlation between these DFT calculations and experimentally observed shifts.

2.6. Acknowledgements

This research was supported as part of The Center for Lignocellulose Structure and Formation, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Award Number DE-SC0001090, and the DOE Great Lakes Bioenergy Research Center (DOE Office of Science BER DE-FC02- 07ER64494). The authors would like to thank and acknowledge the Center for Lignocellulose Structure and Formation (CLSF) and the members thereof. Student fellowships were provided by the USDA National Needs Program and the National Science Foundation. The authors would like to thank Dr. Alan Benesi and Dr. Wenbin Luo for assistance in acquiring NMR spectra of the lignin model compounds, Dr. James Miller for acquiring mass spec data, and Dr. Josh Stapleton for providing assistance with UV/Vis. The primary author would also like to acknowledge Paul Munson and Curtis Frantz for valuable discussion, and valuable interactions with Dan Gall and other members of the Wisconsin lab.

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Chapter 3 Lignin crosslinks with peptides under biomimetic conditions (Target journal for publication is Biomacromolecules)

3.1. Abstract

The work presented here investigates the crosslinking of various nucleophilic amino acids with lignin under aqueous conditions, thus providing insight as to which amino acids might crosslink with lignin in planta. Lignin dehydrogenation polymer (DHP) was prepared in aqueous solutions that contained peptides with the general structure XGG, where X represents an amino acid with a nucleophilic side chain. Fourier-transform infrared spectroscopy and energy dispersive X-ray spectroscopy showed that peptides containing cysteine and tyrosine were incorporated into the DHP, while peptides containing other nucleophilic amino acids were not. Scanning electron microscopy showed that the physical morphology of the DHP was altered by the presence of peptides, regardless of peptide incorporation. Nuclear magnetic resonance (NMR) spectroscopy showed that cysteine-containing peptide crosslinked with lignin at the lignin α-position, whereas in the case of the lignin-tyrosine adduct the exact crosslinking mechanism could not be determined. This is the first study to use NMR to confirm crosslinking between lignin and peptides under biomimetic conditions. The results of this study may indicate the potential for lignin-protein linkage formation in planta, particularly between lignin and cysteine and/or tyrosine-rich proteins.

3.2. Introduction

Lignin is an abundant, aromatic biopolymer that forms in the lignocellulosic matrices of plant cell walls. Its free radical polymerization mechanism and heterogeneous nature make it unique within the plant kingdom. Lignin is economically important to the pulp and paper industries, the agricultural industries, and the biofuels and biorenewables industries, all of whom are hampered by its recalcitrance against extraction and/or degradation (Boerjan, 2003; Stewart, 2006; Chen, 2008; Li, 2008; Chapple, 2007; Jung, 1989; Jung, 1995). Many aspects of lignification are still poorly understood, in spite of its abundance and economic relevance. For example, the extent to which lignin interacts with surrounding cell wall polymers, particularly proteins, is largely unknown. It is understood that lignin forms covalent crosslinks with plant cell wall components, particularly hemicelluloses (Balakshin, 2011; Miyagawa, 2012; Toikka, 1998; Yuan, 2011). One prevalent mechanism for lignin-carbohydrate linkage formation is through the reaction of a nucleophilic moiety (e.g., an hydroxyl or carboxylic acid group) with the electrophilic α-carbon of the lignin quinone methide (QM) intermediate (Leary, 1980; Ralph, 2009). The crosslinking of lignin with other cell wall components, such as proteins, has not been well investigated, despite the fact that lignin-protein linkages may play important roles in wild type and transgenic plant lines. In most wild type plant lines the pattern of lignin deposition indicates the presence of so-called nucleation sites within specific regions of the plant cell wall (e.g., the cell corners), but the nature of these nucleation sites remains unknown (Boerjan et al., 2003). It has been suggested that nucleation sites may be rich in structural proteins, perhaps leading to lignin-protein

55 crosslinking, but this hypothesis has not been adequately tested. Furthermore, lignin-protein linkages may affect the physical and chemical properties of transgenic plant lines. For example, a recently engineered line of Populus secretes a tyrosine-rich peptide into the cell wall. Increased sugar extractability was observed in these Populus lines upon protease digestion of the walls, and it was hypothesized that this was due to lignin-protein linkage formation. However, the putative lignin-protein linkages have yet to be identified (Liang et al., 2008; Xu et al., 2013). Diehl et al. (2014) recently showed that amino acids bearing nucleophilic side chains, namely Cys, Lys, His, Asp, Glu, Ser, and Tyr all react with a lignin model QM in dichloromethane. The study identified diagnostic NMR shifts of lignin-peptide compounds, but did not investigate the propensity for such linkages to form under biomimetic conditions (i.e., conditions of higher molecular weight lignin formation with peptides in aqueous media). In order to expand upon these results, the work described here investigates the propensities for various amino acids (in peptide chains) to crosslink with lignin dehydrogenation polymer, which is a biomimetic lignin model compound (Terashima et al., 1995). It is anticipated that this will assist in future studies to help elucidate the interactions between lignin and proteins in planta. In order to investigate the propensity for lignin-peptide crosslinking under biomimetic conditions, lignin dehydrogenation polymer (DHP) was prepared in aqueous solutions containing peptides. Each peptide had the general structure X-glycine-glycine (XGG), with X being cysteine (C), lysine (K), histidine (H), aspartic acid (D), glutamic acid (E), serine (S), tyrosine (Y), threonine (T), or hydroxyproline (Hyp). These amino acids were previously identified as being reactive (or potentially reactive in the case of T and Hyp) toward lignin QMs (Diehl et al., 2014). The general peptide structure and predicted mode of lignin-peptide crosslinking is shown in Fig 3.1. The C-termini and N-termini of the peptides were blocked via amidation and esterification, respectively, to ensure that the amino acid of interest (i.e., residue X) contained the only nucleophilic moiety. Glycine was chosen as the "place holder" residue due to its expected lack of reactivity toward lignin. The lengths of the peptides were limited to three residues because reaction of larger peptides with DHPs results in the formation of lignin-peptide complexes that are insoluble and thus difficult to characterize (e.g., liquid state NMR becomes impractical) (results not yet published). Peptides were added in 25% mol/mol ratio to the lignin monomer (coniferyl alcohol) because it was previously reported that lignin DHPs contain between 20 and 30% β-ether linkages (Tobimatsu, 2012). Thus, the ratio of nucleophilic amino acids to lignin β- ether QMs was expected to be approximately 1:1 over the course of the polymerization reaction.

Fig 3.1. Proposed lignin-peptide crosslinking mechanism. Lignin-peptide crosslinks form when nucleophilic side chains of amino acids react with quinone methides formed during lignin β-ether coupling. R = H or OMe, L = lignin. Fourier-transform infrared spectroscopy (FT-IR), scanning electron microscopy (SEM), energy dispersive X-ray spectroscopy (EDS), and nuclear magnetic resonance spectroscopy (NMR) were used to characterize the lignin-peptide adducts. FT-IR and, more recently, NMR, have become staples of lignin characterization (Capanema et al., 2004; Faix, 1988; Kim and Ralph, 2010). Multidimensional NMR techniques (e.g., heteronuclear single quantum coherence (HSQC)) are particularly useful because the shift degeneracies observed in 1D spectra are largely eliminated. Furthermore, diagnostic NMR shifts of lignin-peptide model compounds have previously been assigned (Diehl et al., 2014). SEM imaging of synthetic and native lignins has not garnered much research attention, but the technique was employed here in order to monitor morphological differences between neat DHP and the lignin-peptide adducts (Micic et al., 2003; Terashima et al., 2004). It was convenient to also collect EDS elemental analysis data while the lignin-peptide samples were in the SEM instrument, with the presence of nitrogen suggesting peptide incorporation because neat lignin contains only carbon, hydrogen, and oxygen. Through the use of these techniques, this study provides new insights into the propensities and mechanisms of lignin-peptide linkage formation. It is expected that this will be useful toward the continued study of lignin formation in both native and mutant plant lines.

3.3. Experimental 3.3.1. Materials All chemicals necessary for DHP preparation were purchased from Sigma with the exception of the peptides (>95% purity), which were purchased from Peptide 2.0 (www.peptide2.com). 3.3.2. Synthesis of lignin DHP and lignin-peptide adducts Guaiacyl-based DHP was synthesized according to a previously published method in sodium phosphate buffer (pH 6.5) using coniferyl alcohol as the sole lignin monomer (Terashima, 1995). The DHP crude product was centrifuged (10k g, 20 min, 4 °C) and the pellet washed four times with distilled water. The DHP product was then lyophilized to yield dry DHP (typical yields 60-70%), which was characterized via NMR as described below and was found to contain shifts typical of G-DHP (Capanema, 2004; Kim, 2010). Lignin-peptide adducts were prepared as above, with the exception that 25% peptide to coniferyl alcohol (mol/mol basis) was added to the flask containing coniferyl alcohol prior to the start of the reaction. The crude reaction products were centrifuged and lyophilized as described above to yield tan powders. These adducts were characterized using IR, SEM, EDS and NMR. 3.3.3. Scanning electron microscopy and energy dispersive X-ray spectroscopy Scanning electron microscopy images were collected on a field emission SEM (FESEM - FEI NanoSEM 630) at 2 or 3 kV under high vacuum (1.7 x 10-6 Torr). Samples were not sputter coated prior to imaging. Characteristic X-rays were collected with an X-Max silicon drift detector (Oxford Instruments) inside the FESEM at 10 kV under low vacuum conditions (0.6

Torr) in order to prevent sample charging. Elements were selected and quantified using Aztec Energy Analyser Software (Oxford Instruments).

3.3.4. Nuclear magnetic resonance spectroscopy

The neat peptides (25 mg) were dissolved in DMSO-d6/pyridine-d5 (4:1 v/v, 500 ul), and proton (16 scans), carbon (4k scans), HMQC (64 scans) and HMBC (32 scans) spectra were collected using standard Bruker pulse programs on a Bruker DRX-400 (400 MHz 1H resonance freq.) using the central solvent peak [δH/δC: dimethyl sulfoxide (DMSO), 2.50/39.50 ppm] as internal standard. In the case of DHP and the lignin-peptide adducts, NMR spectra were acquired on a Bruker Biospin (Billerica, MA, USA) AVANCE 500 (500 MHz 1H resonance freq.) spectrometer fitted with cryogenically-cooled gradient probes having inverse geometry, i.e., with the proton coils closest to the sample. Spectra were processed with Bruker’s Topspin 3.1 software, using the central solvent peak as internal reference [δH/δC: dimethyl sulfoxide (DMSO), 2.50/39.5 ppm]. The DHP or lignin-peptide adducts (~45 mg) were placed in an NMR tube (ID: 4.1 mm), dissolved in DMSO-d6/pyridine-d5 (4:1 v/v, 500 ul), and subjected to adiabatic HSQC (‘hsqcetgpsisp2.2’) experiments, and, in the case of DHP-YGG, also subjected to HMBC (‘hmbcgpndqf’), COSY (‘cosygpqf’), and NOESY (‘noesyesgpph’) experiments in an attempt to determine the lignin-tyrosine crosslinking mechanism. Processing used typical matched Gaussian apodization in F2 (LB = -0.3, GB = 0.001), and squared cosine-bell and one level of linear prediction (32 coefficients) in F1 (Mansfield, 2012). For an estimation of the various inter-unit linkage types in DHP and DHP-peptide adducts (Table 2; β-ether/α-OH, β- ether/α-O-aryl, β-ether/α-peptide, phenylcoumaran, pinoresinol, and dibenzodioxocin), the well resolved Cα-Hα contours were integrated; no correction factors were used. 3.3.5. Fourier-transform infrared spectroscopy Lignin DHP, neat peptides, and lignin-peptide adducts were analyzed using a Bruker Vertex V70 Spectrometer (Bruker Optics Billerica MA) equipped with an MVP-Pro diamond single reflection ATR accessory (Harrick Scientific Pleasantville NY), and 100 scans at 6 cm-1 resolution were averaged for each sample using a DTGS detector and scan frequency of 5 kHz. In all cases, the spectrum of the clean diamond crystal was used as the reference spectrum. All spectral manipulations were performed using OPUS 6.0 (Bruker Optics, Billerica MA).

3.4. Results and discussion 3.4.1. Preparation and yields of the lignin-peptide adducts Lignin DHP was prepared in aqueous solutions containing tripeptides (25% peptide:coniferyl alcohol mol/mol basis). Each tripeptide contained one nucleophilic amino acid and blocked N- and C-termini in order to mimic inclusion within a larger protein and to prohibit potential side reactions. The lignin-peptide adducts were collected via centrifugation, washed, and characterized via SEM, NMR, EDS, and FT-IR. The results, detailed below, indicate covalent incorporation of CGG and YGG peptides into the lignin polymer, while other peptides did not show significant reactivity. Yields for the DHP and lignin-peptide adducts are shown in Table 3.1. Yield A was determined by dividing the mass of recovered solids by the total starting mass (i.e., combined mass of lignin monomer and peptide), while yield B was determined by dividing the mass of

58 recovered solids by the starting mass of lignin monomer only. Thus, yield B is only valid for lignin-peptide reactions in which peptide incorporation into the lignin was negligible. Notably, the yields were very high when DHP was prepared in the presence of non-covalently reactive peptides (i.e., all peptides other than CGG and YGG). The reason for this was unclear. In the cases of the considerably reactive peptides (i.e., CGG and YGG) the yields of recoverable DHP were depressed. For DHP-CGG, a likely explanation is that the thiol group of the CGG peptide inhibited the catalytic ability of horseradish peroxidase, thus hampering polymerization (Tobimatsu et al, 2009; Veitch, 2004). It is not known why the yield was depressed in the case of DHP-YGG. The authors perceived that in the cases of DHP-CGG and DHP-YGG a portion of the lignin-peptide adducts may have been aqueous soluble and held in solution during the centrifugation process. However, extraction of the aqueous supernatants with ethyl acetate and chloroform followed by NMR analyses did not show evidence for lignin- protein complexes. Drying down the aqueous supernatant, re-suspending the solids in DMSO- d6/pyridine-d5, and analyzing the products via NMR similarly failed to provide evidence for lignin-protein crosslinking. This confirmed the depression of DHP yields in the cases of DHP- CGG and DHP-YGG. Table 3.1. Yield data for the DHP and lignin-peptide adducts

CA (mg) pep (mg) yield (mg) Yield A (%) Yield B (%) DHP 200.0 0.0 130.0 65.0 65.0 DHP-CGG 200.0 88.4 100.0 34.7 - DHP-KGG 200.0 83.6 174.5 61.5 87.3 DHP-HGG 200.0 86.1 176.0 61.5 88.0 DHP-DGG 200.0 80.0 180.0 64.3 90.0 DHP-EGG 200.0 83.9 197.9 69.7 99.0 DHP-SGG 200.0 72.2 172.8 63.5 86.4 DHP-YGG 200.0 93.3 114.2 38.9 - DHP-TGG 200.0 76.1 179.1 64.9 89.6 DHP-HypGG 200.0 79.4 177.4 63.5 88.7

Yield A was determined by dividing the mass of recovered solids by the total starting mass (i.e., combined mass of lignin monomer and peptide). Yield B was determined by dividing the mass of recovered solids by the starting mass of lignin monomer only. CA = coniferyl alcohol, pep = peptide. 3.4.2. Lignin-peptide morphology Scanning electron microscopy was used to compare the morphologies of DHP and the lignin-peptide adducts (Fig 3.2). The DHP particles clumped together to form nearly perfect spheres, as reported previously (Micic et al, 2003; Micic et al, 2004). Comparatively, spheres of lignin-peptide adducts tended to form large, amorphous domains. This alteration of morphology was observed regardless of whether the peptide in question was incorporated into the lignin. This change in morphology was unexpected but was most likely due to non-covalent interactions

59 occurring between the peptides and the growing lignin chain. Further research is necessary to determine the influence of non-covalent inter-polymer interactions during lignin polymerization.

Fig 3.2. SEM images of DHP (top), then, proceeding from left to right and top to bottom, DHP- CGG, DHP-YGG, DHP-HypGG, DHP-DGG, DHP-EGG, DHP-KGG, DHP-HGG, DHP-SGG, and DHP-TGG. Scale bar: 2 µm.

3.4.3. Lignin-peptide linkage identification Fig 3.3 shows the heteronuclear single quantum coherence (HSQC) spectrum of DHP- CGG. This 2D NMR technique is particularly useful for lignin analysis because the shift degeneracy observed in 1D NMR spectra is largely avoided. Novel shifts are shown in green, red, and blue, while standard lignin shifts are shown in black (Capanema, 2004; Kim, 2010). Reference shifts of neat CGG (purple) were added to the spectrum during processing; these shifts were not observed in the DHP-CGG spectrum. Some peptide shifts migrated as a result of DHP- 13 1 CGG crosslinking. For example, the cys- C/ Hα shift (originally at 4.5/56.0 ppm in neat CGG) 13 1 migrated to 4.6/52.8 ppm, and the cys- C/ Hβ shift (originally at 2.8/26.6 ppm in neat CGG) migrated to 2.8/32.9 ppm in the DHP-CGG adduct. These shifts migrated upon lignin-peptide crosslinking due to their proximity to the thiol group, which is the reactive center of the CGG

60 peptide. Shifts of proton and carbon atoms located far from the reactive thiol were largely unaffected by crosslinking (e.g., shifts at 3.9/42.8 and 3.8/42.5 ppm).

Two novel lignin shifts, found at 4.4/50.1 ppm (Fig 3.3, red peak) and 4.8/81.3 ppm (Fig 3.3, blue peak), confirmed covalent crosslinking of DHP with CGG. Similar lignin α- shifts (4.3/50.4 ppm) and β-shifts (4.7/81.7 ppm) were previously reported when a single cysteine residue was reacted with a lignin model quinone methide to yield a structure similar to that shown in Fig 3.3 (Diehl et al., 2014). The minute differences in shift locations can be attributed to changes in chemical environment between a small lignin model compound and a high molecular weight lignin. Volume integration of the HSQC contours showed that approximately 33% of the β-ether linkages in DHP-CGG exhibited cysteine functionality at the α-carbon, while the remaining β-ether linkages exhibited typical α-hydroxyl functionality and a minor fraction of α-aryl ether (α-O-aryl) moieties. This indicated that cysteine was an efficient trapper of lignin QMs under biomimetic conditions.

Fig 3.3. Side-chain and aromatic regions (inset) of the HSQC NMR spectrum of DHP-CGG. Black shifts are typical of G-DHPs, green shifts correspond to peptide α- and β-signals, and red and blue shifts correspond to lignin α- and β-signals in β-ether/α-cysteine structures (top left). Purple shifts were added during processing to indicate shifts of neat CGG peptide. Fig 3.4 shows the HSQC spectrum of DHP-YGG. As with the DHP-CGG adduct, incorporation of YGG peptide into lignin was evidenced by the appearance of diagnostic chemical shifts (Fig 3.4, green and orange contours). Reference shifts of neat YGG (purple, solid yellow, and solid orange contours) were added during processing. The authors perceived that given the similarity of tyrosine and coniferyl alcohol, crosslinking of YGG with DHP may have occurred via two mechanisms.

The first potential mechanism involves oxidation of the phenolic hydroxyl of tyrosine by horseradish peroxidase, followed by recombination of the tyrosine radical with a radical on the lignin polymer. This mechanism may be unfavorable because radicals generated on tyrosine could be shuttled to coniferyl alcohol, which exhibits an additional resonance structure compared to tyrosine, presumably making it more stable (Cong et al., 2013). In addition to HSQC NMR, we submitted the DHP-YGG adduct to heteronuclear multiple bond correlation (HMBC), correlation spectroscopy (COSY), and nuclear Overhauser effect spectroscopy (NOESY) techniques (spectra not shown), but were unable to conclusively assign NMR shifts of lignin- tyrosine linkages formed in this manner. This may indicate that the mechanism is not valid under our experimental conditions, and/or may illustrate the inadequacy of NMR to resolve shift degeneracy between lignin-tyrosine linkages and typical lignin shifts. A second crosslinking mechanism is possible when the phenolic hydroxyl of tyrosine quenches the lignin quinone methide to form the α-aryl ether structure shown in Fig 3.4. Again, shift degeneracy may complicate the investigation of this mechanism, as a lignin-tyrosine model compound exhibited similar NMR shifts (α-1H/13C: 5.5/78.3 ppm in DMSO/pyridine) to α-aryl ether linkages known to occur in neat DHPs (α-13C: 79.01 ppm in DMSO) (Diehl et al., 2014; Ralph, Ralph and Landucci, 2004). In an attempt to overcome this issue, the well-resolved HSQC α-signals of neat DHP and DHP-YGG adduct were integrated. It was observed that α-aryl ether shifts comprised approximately 4.2% of the total α-signal in DHP-YGG but only 1.9% in neat DHP synthesized under similar conditions. This increase could be due to imprecision in the HSQC volume integration or random variation among DHP syntheses (other lignin-peptide adducts displayed similarly high α-aryl ether signals), making it unclear if the structure shown in Fig 3.4 formed in the DHP-YGG adduct. In summary, the NMR, EDS, and IR data (shown below) strongly suggest that the YGG peptide crosslinked with lignin DHP; however, the mechanism of lignin-tyrosine crosslinking is still uncertain.

Fig 3.4. Side-chain and aromatic regions (inset) of the HSQC NMR spectrum of DHP-YGG. Black shifts are typical of G-DHPs, green shifts correspond to peptide α- and β-signals, and red and blue shifts correspond to lignin α- and β-signals in β-ether/α-tyrosine structures (top left) and/or lignin-lignin α-O-aryl structures. Purple shifts were added during processing to indicate shifts of neat YGG peptide. Within the aromatic region, solid yellow and orange shifts (added during processing) were assigned to the aromatic ring of tyrosine in neat YGG. It can be seen that the contours (orange) representative of tyrosine ring positions 3 and 5 shift downfield as a result of lignin-tyrosine crosslinking. The specific lignin-tyrosine crosslinking mechanism could not be determined by NMR, and the structure shown is one of several possibilities. It was notable that in the case of lignin-peptide adducts other than DHP-CGG and DHP- YGG, peptide peaks could always be observed when viewing the HSQC contours quite low (i.e., near the signal to noise limit). Fig 3.5 shows the HSQC spectrum of DHP-HGG. This sample showed the highest concentration of peptide after DHP-CGG and DHP-YGG. A putative lignin- α-histidine crosslink was observed at 5.7/60.4 ppm, in good agreement with the α-shift of a lignin-histidine model compound (5.7/60.2 ppm) (Diehl et al., 2014). Volume integration showed that the abundance of the lignin-α-histidine shift only accounted for ~0.1% of the total lignin α- signal. It is noteworthy that this low abundance of peptide was detected by HSQC NMR but not readily detected by IR or EDS, thus illustrating the sensitivity of multidimensional NMR toward investigating lignin-protein linkages. Other lignin-peptide adducts exhibited less abundant NMR peptide shifts than DHP- HGG. This indicated that negligible lignin-peptide crosslinking had occurred, in concurrence with IR and EDS data (below).

Fig 3.5. Side-chain region of the HSQC NMR spectrum of DHP-HGG. Black shifts are typical of G-DHPs, green shifts correspond to peptide α- and β-signals, and red and blue shifts correspond to lignin α- and β-signals in β-ether/α-histidine structures (top left). Purple shifts were added during processing to indicate shifts of neat HGG peptide. Volume integration of the HSQC contour signals allowed for comparison of the various lignin inter-unit linkages among DHP and lignin-peptide adducts (Table 3.2). The DHP contained linkage ratios typical of DHPs (Terashima et al., 1995 and 2009; Tobimatsu et al., 2012). Linkage ratios varied among the lignin-peptide adducts, but decreased β-ether content with increased pinoresinol (β-β) content was generally observed. This occurred regardless of covalent reactivity towards the lignin DHP, demonstrating the ability of a matrix material (in this case peptides) to influence lignin structure during polymerization. Table 3.2. Inter-unit linkage ratios of the DHP and lignin-peptide adducts

HSQC signal ratios β-ether/α-OH β-ether/α-O-aryl β-ether/α-pep β-5 β-β Dibenz. DHP 27.3 1.9 - 50.3 19.2 1.2 DHP-CGG 8.6 1.5 5.1 50.8 32.4 1.6 DHP-DGG 10.1 5.5 0.1 54.1 30.2 tr DHP-EGG 13.1 4.6 0.1 54.1 27.2 0.9 DHP-KGG 4.7 3.1 tr 62.3 29.9 tr DHP-HGG 20.4 0.9 0.1 57.7 20.3 0.6 DHP-SGG 11.1 2.3 tr 52.5 34.1 tr DHP-YGG 11.5 4.2 51.7 32.5 tr DHP-TGG 21.8 0.9 tr 54.1 22.2 1.0 DHP-HypGG 17.7 2.7 tr 53.2 26.2 0.2 Lignin inter-unit linkage ratios (as percentage of total α-signal) for DHP and lignin-peptide adducts. In the case of DHP-YGG the DHP-α-peptide shift was degenerate with standard lignin α-O-aryl shifts, thus the β-ether/α-O-aryl and β-ether/α-pep quantities were combined. tr, trace (<0.1%).

3.4.4. Supporting techniques for characterization of lignin-peptide entanglement In addition to NMR, which can provide direct evidence of covalent crosslinking, other techniques can be used to show peptide incorporation into lignin. Fig 3.6 shows FT-IR spectra of neat DHP and the lignin-peptide adducts. The neat DHP IR spectrum exhibited bands typical of lignin DHPs (Faix, 1988). The DHP-CGG and DHP-YGG spectra exhibited three peaks indicative of peptide incorporation into the lignin. The shoulder near 3200 cm-1 was attributed to N-H stretching in amide functional groups, the peak at 1658 cm-1 increased dramatically and was attributed to increased C=O stretching due to the incorporation of amide functional groups, and the shoulder at 1540 cm-1 was attributed to N-H deformation with C-N stretching, again indicating incorporation of amide functionalities (Socrates, 2001). It is notable that these shifts displayed greater intensity in the DHP-CGG adduct compared to the DHP-YGG adduct, suggesting greater incorporation of CGG peptide. These peaks were not observed in the IR spectra of other lignin-peptide adducts, suggesting a lack of peptide incorporation. In the case of DHP-CGG and DHP-YGG, incorporation of peptide into the lignin polymer caused an increase

64 in the band at 1505 cm-1, which was previously assigned to aromatic skeletal vibrations (Faix, 1988). This increase in peak height was not observed in other lignin-peptide adducts and the origin of this increased peak intensity is unclear. Though the IR results suggested incorporation of CGG and YGG peptides into lignin DHP, peaks directly attributable to lignin-peptide linkages were not identified. These results demonstrate that IR is a quick and reliable technique for indicating lignin-peptide interactions in general, but may be insufficient for determining the presence or absence of covalent crosslinks.

Fig 3.6. FT-IR spectra of DHP and lignin-peptide adducts.

Because neat lignin contains only carbon, oxygen, and hydrogen, elemental analysis techniques can be used to show incorporation of proteins into lignin when nitrogen is present (assuming no inorganic nitrogen contamination). In this case, energy dispersive X-ray spectroscopy (EDS) was used to determine the elemental compositions of DHP and lignin- peptide adducts because EDS spectra are readily attainable in the SEM instrument, and can therefore be carried out in conjunction with morphological studies. The DHP and most lignin- peptide adducts contained 0% nitrogen (Table 3), which indicated no detectable incorporation of peptide into the lignin. In comparison, DHP-CGG and DHP-YGG contained 3.5% and 2.0% nitrogen, respectively. This provided compelling evidence for incorporation of cysteine and tyrosine-containing peptides into the lignin DHP. It was not possible to determine quantitative ratios (for example, lignin-to-peptide wt/wt ratio) using this technique; however, it is possible to roughly compare peptide quantities among samples when necessary. Thus, the EDS data suggested that cysteine reacted more readily with lignin compared to tyrosine, which is in agreement with the IR data.

Table 3.3. EDS elemental analysis data for DHP and the lignin-peptide adducts.

Average atomic % (std. dev.) carbon oxygen nitrogen sulfur DHP 78.8 (2.8) 21.2 (2.9) 0.0 (0.0) 0.0 (0.0) DHP-CGG 76.5 (2.0) 18.9 (1.2) 3.5 (0.7) 1.2 (0.3) DHP-KGG 86.8 (2.5) 13.2 (2.5) 0.0 (0.0) 0.0 (0.0) DHP-HGG 90.5 (2.1) 9.5 (2.5) 0.0 (0.0) 0.0 (0.1) DHP-DGG 82.8 (3.8) 17.2 (3.8) 0.0 (0.0) 0.0 (0.0) DHP-EGG 83.7 (2.0) 15.4 (1.1) 0.0 (0.0) 0.6 (0.8) DHP-SGG 86.3 (2.9) 13.7 (2.9) 0.0 (0.0) 0.0 (0.0) DHP-YGG 84.0 (2.7) 14.0 (1.1) 2.0 (1.7) 0.0 (0.0) DHP-TGG 85.8 (1.1) 14.2 (1.1) 0.0 (0.0) 0.0 (0.0) DHP-HypGG 84.4 (0.2) 15.6 (0.2) 0.0 (0.0) 0.0 (0.0)

Atomic percentages are reported as averages of three sampling locations. Standard deviations are shown in parentheses. Trace levels of calcium account for the balance in the case of DHP-EGG. 3.5. Conclusions Amino acid residues with nucleophilic side chains were previously shown to react with a lignin model quinone methide in dichloromethane, yielding lignin-α-peptide structures (Diehl et al., 2014). In the present study, we extended this work by characterizing DHP-peptide covalent crosslinks and non-covalent effects of peptides on DHP formation, under biomimetic conditions. Lignin DHP was prepared using coniferyl alcohol as the sole lignin monomer and peptides were added having the general structure XGG, in which X was an amino acid residue with a nucleophilic side chain (i.e., C, K, H, D, E, S, Y, T, and Hyp). The lignin was precipitated via centrifugation to yield DHP-peptide adducts, and analysis using IR, EDS, and NMR showed that CGG and YGG were significantly reactive toward lignin while other peptides were not. In the case of DHP-CGG, HSQC NMR showed that crosslinking occurred at the lignin α-position

(Fig 3.4). The crosslinking mechanism of DHP with YGG could not be conclusively elucidated. SEM imaging showed that DHP-peptide adducts exhibited a unique morphology compared to neat DHP, regardless of peptide incorporation into the lignin. With regards to lignin inter-unit lignin ratios, the quantity of β-ether linkages was typically depressed in DHPs synthesized in the presence of peptides, while the quantity of pinoresinol structures increased. The yields of DHP- CGG and DHP-YGG were depressed compared to neat DHP yields; however, curiously, the yields increased when DHP was prepared in the presence of other peptides. We have shown that cysteine and tyrosine crosslink with lignin under biomimetic conditions. This suggests that similar crosslinking may occur in the cell walls of both native and transgenic plant lines. Further research is needed to investigate whether this crosslinking does occur, and to discover how the plant might control and benefit from such crosslinking (i.e., does it help stiffen the wall, assist water conduction, provide protection from pathogens, etc). In addition, a better understanding of lignin-protein linkages could lead to genetic manipulation (up-regulation or down-regulation of the linkages), as already suggested by Liang et al. (2008) and Xu et al. (2013). This could reduce lignin recalcitrance, which is currently a barrier to using lignocellulosic materials in developing industries such as biofuels. It is anticipated that this work will lead to future studies of lignin-protein linkages in planta, and a more thorough understanding of how such linkages could be tailored and modified.

3.6. Acknowledgements This material is based upon work supported as part of The Center for Lignocellulose Structure and Formation, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Award Number DE- SC0001090. Student fellowships were provided by the USDA National Needs Program and the National Science Foundation via the CarbonEARTH program. Many thanks to Julie Anderson and Melisa Yashinski (PSU MRI) for acquisition of SEM/EDS data and for valuable discussions. Thanks also to Dr. John Ralph, Yuki Tobimatsu, and Matt Regner for valuable discussions regarding multidimensional NMR of lignin. 3.7. References

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Chapple, C.; Ladisch, M.; Meilan, R. Loosening lignin's grip on biofuel production. Nat. Biotechnol. 2007, 25, 746-748. Chen, F.; Dixon, R. A. Genetic manipulation of lignin biosynthesis to improve biomass characteristics for agro-industrial processes. In Vitro Cell. Dev. Biol. - Animal. 2008, 44, S28- S29. Cong, F.; Diehl, B.G.; Hill, J.L.; Brown, N.R.; Tien, M. Covalent bond formation between amino acids and lignin: Cross-coupling between proteins and lignin. Phytochem. 2013, 96, 449-456. Diehl, B.G.; Watts, H.D.; Kubicki, J.D.; Regner, M.R.; Ralph, J.; Brown, N.R. Towards lignin- protein crosslinking: Amino acid adducts of a lignin model quinone methide. Cellulose 2014, available online. Faix O, Beinhoff O. 1988. Journal of Wood Chemistry and Technology, 8 (4): 505-522. "FTIR spectra of milled wood lignins and lignin polymer models with enhanced resolution obtained by deconvolution." Fisher, J.; Abdella, B. R. J.; McLane, K. E. Anthracycline antibiotic reduction by spinach ferredoxin-NADP+ reductase and ferredoxin. Biochemistry 1985, 24, 3562-3571. Jung, H. G.; Allen, M. S. Characteristics of plant cell walls affecting intake and digestibility of forages by ruminants. J. Animal Sci. 1995, 73, 2774-2790. Jung, H. G. Forage lignins and their effects on fiber digestibility. Agron. J. 1989, 81, 33-38. Kim, H.; Ralph, J. Solution-state 2D NMR of ball-milled plant cell wall gels in DMSO- d6/pyridine-d5. Org. Biomol. Chem. 2010, 8, 576-591. Leary, G. J. Quinone methides and the structure of lignin. Wood Sci. and Tech. 1980, 14, 21-34. Liang, H.; Frost, C. J.; Wei, X.; Brown, N. R.; Carlson, J. E.; Tien, M. Improved sugar release from lignocellulosic material by introducing a tyrosine-rich cell wall peptide gene in poplar. Clean 2008, 36(8), 662-668. Li, X.; Weng, J. K.; Chapple, C. Improvement of biomass through lignin modification. Plant J. 2008, 54, 569-581.

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Chapter 4 Preparation and characterization of lignin-gelatin complexes (Target journal for publication is Journal of Applied Polymer Science)

4.1. Abstract

Lignin dehydrogenation polymer (DHP) was prepared in the presence of gelatin protein to yield “DHP-Gel adducts.” The DHP-Gel adducts were characterized using Fourier-transform infrared spectroscopy (FT-IR), scanning electron microscopy (SEM), energy dispersive X-ray spectroscopy (EDS), X-ray photoelectron spectroscopy (XPS) and heteronuclear single quantum coherence nuclear magnetic resonance spectroscopy (HSQC NMR). FT-IR, EDS and XPS showed that gelatin was incorporated into the lignin even when added in small quantities. In addition, EDS and XPS showed that gelatin was distributed throughout the DHP when added during lignin polymerization, but adsorbed to the surface of DHP when added following polymerization. A lack of diagnostic lignin-protein crosslink signatures in the HSQC NMR spectra suggested that the lignin-gelatin interaction was largely non-covalent in nature. This may have implications toward lignification in planta, in which lignin is biosynthesized in a pre- deposited matrix of polysaccharides and proteins. Cell wall structural proteins, which are common in the cell corner region of the middle lamella (where lignification begins), may help “nucleate” lignification without necessarily covalently crosslinking with lignin. Key words: lignin, gelatin, crosslinking, non-covalent, scanning electron microscopy, energy dispersive X-ray spectroscopy, X-ray photoelectron spectroscopy, nuclear magnetic resonance spectroscopy.

4.2. Introduction

Lignin is the most abundant aromatic biopolymer on earth (Boerjan et al., 2003). It is most commonly biosynthesized in the cell walls of land plants from three monolignols (p- coumaryl, coniferyl, and sinapyl alcohols) that vary in their degree of aromatic ring methoxylation. It is commonly believed that lignin imparts three main evolutionary advantages to the plant: structural rigidity, water conductivity, and pathogen resistance. Lignin is commercially important to the pulp and paper industry, agricultural industries concerned with forage digestibility, and the developing biofuels industry, in which it is known to foul cellulose to ethanol conversion processes (Stewart, 2006; Chen, 2008; Li, 2008; Chapple, 2007; Jung, 1989; Jung, 1995). Lignin is also a potential source of renewable carbon for plastics, carbon fibers, solvents, and low and high value chemicals, to name a few (Gellerstedt, et al., 2010, Chen and Sarkanen, 2006, Dorrestijn, et al., 2000, Clark, et al., 2009).

Despite decades of research, some details of lignification are still poorly understood. For example, it has long been known that within most plants, lignin deposition begins in the cell corner region of the middle lamella. However, the mechanism by which the plant controls this pattern of lignin deposition is unknown. Lignification initiation sites (sometimes referred to as “nucleation sites”) have been postulated, with two commonly hypothesized initiation sites being

71 calcium-pectate complexes (which may bind anionic peroxidases necessary for lignin polymerization) and cell wall structural proteins (especially extensins, which are abundant in the cell corners) (Albersheim et al., 2010; Boerjan et al., 2003). Neither of these hypotheses has been adequately investigated in vitro or in vivo. Several studies have shown that proteins have an affinity to bind lignin; however, the nature of the lignin-protein binding (i.e., covalent vs. non- covalent) was not elucidated (Whitmore, 1978a; Whitmore, 1978b; Whitmore, 1982). With regard to lignin-protein covalent crosslinking, it was recently shown that Cys, Lys, His, Asp, Glu, Ser, and Tyr crosslink with lignin in non-polar solvents, and that Cys and Tyr crosslink with lignin even under aqueous, biomimetic conditions (Diehl et al., 2014; Diehl and Brown, in review). Here, we report the preparation and characterization of lignin DHP in the presence of gelatin, a glycine and hydroxyproline-rich animal protein.

Though gelatin protein does not originate from plants, the lignin-gelatin complex is interesting and potentially informative for several reasons. Gelatin is both glycine and hydroxyproline-rich, as are many plant cell wall structural proteins. Gelatin has previously been shown to interact with lignin, though the presence or absence of covalent crosslinks was not definitely determined (Whitmore, 1978b). Several potentially nucleophilic amino acid residues are found in gelatin; however, cysteine and tyrosine residues are almost entirely lacking (Table 4.1) (Eastoe, 1955). While nucleophilic amino acids other than cysteine and tyrosine have been shown to react with lignin under ideal conditions in non-polar solvents, only cysteine and tyrosine have been shown to covalently crosslink with lignin under biomimetic conditions of DHP preparation (Diehl et al., 2014; Diehl and Brown, in review). The affinity for gelatin to interact with lignin under aqueous conditions is therefore interesting, as it seems most likely to arise from physical entanglement and/or non-covalent interactions, or from lignin-protein crosslinkage types that have not previously been observed under conditions of DHP preparation. Understanding the interactions occurring within the lignin-gelatin complex may provide insights into lignin-protein interactions in planta, where lignin and cell wall structural proteins could be expected to be in close spatial proximity.

Table 4.1. Nucleophilic amino acid abundance (g/100 g dry, ash-free protein) in gelatin.

Residue Porcine Gelatin Bovine Gelatin Cys 0.00 0.05 Lys 4.14 5.20 His 1.01 0.63 Glu 11.30 12.10 Asp 6.70 6.90 Tyr 0.60 0.14 Ser 4.13 2.90 Thr 2.19 2.20 Hyp 13.50 14.40 Total 43.57 44.52

In order to elucidate the lignin-gelatin interaction, five lignin-gelatin complexes (DHP- Gel adducts) were prepared (see Table 4.2 for DHP-Gel preparations). The DHP was prepared by slowly combining monolignol (e.g., coniferyl alcohol) and hydrogen peroxide solutions to a horseradish peroxidase solution, as previously described (Terashima et al., 1995). In addition, various quantities of porcine (high molecular weight) and bovine (low molecular weight) gelatin were added to the DHP preparations. DHP-Gel1 and DHP-Gel5 contained equivalent quantities (wt/wt basis) of monolignol and porcine gelatin, with the only difference being that gelatin was dissolved in the same flask as the coniferyl alcohol in the case of DHP-Gel1, and was thus added slowly and continuously to the DHP preparation during polymerization, whereas in the case of

DHP-Gel5 gelatin was added to the DHP preparation following completion of polymerization. It was perceived that covalent crosslinking, if it occurred, might be more likely in the case of DHP-

Gel1 because the gelatin would potentially be in intimate contact with reactive lignin intermediates (i.e., quinone methides) over the course of the polymerization reaction. It was also perceived that addition of gelatin at different times might influence the morphology of the DHP-

Gel adducts. In the cases of DHP-Gel2, DHP-Gel3, and DHP-Gel4, decreasing quantities of bovine gelatin (2.5:1, 5:1, and 10:1 monolignol to gelatin wt/wt basis) were added to the DHP, again during the polymerization process. These DHP-Gel adducts allowed for investigation of lignin interactions with small quantities of low molecular weight gelatin.

The DHP-Gel adducts were characterized using several techniques. Fourier-transform infrared spectroscopy (FT-IR) was employed because it can show incorporation of protein into lignin, although it is admittedly deficient at identifying lignin-protein covalent crosslinks (Diehl et al., in review). Scanning electron microscopy (SEM) was used to determine the physical morphology of the DHP-Gel adducts. Energy dispersive X-ray spectroscopy (EDS) and X-ray photoelectron spectroscopy (XPS) were used to both confirm the incorporation of protein into the lignin (via quantification of nitrogen) as well as to elucidate morphological details. Finally, heteronuclear single quantum coherence (HSQC) nuclear magnetic resonance (NMR) spectroscopy was used to investigate potential lignin-gelatin covalent crosslinks. Investigation of the lignin-gelatin complexes reported here may lead to a better understanding of lignin-protein interactions in native plant systems.

4.3. Experimental

4.3.1. Materials

Coniferyl alcohol was prepared from coniferaldehyde (Sigma Aldrich) as previously described (Ludley and Ralph, 1996). Horseradish peroxidase (type I), hydrogen peroxide, sodium phosphate and porcine and bovine gelatin were purchased from Sigma Aldrich. The bovine gelatin (Sigma #G6650) had a bloom number of 75, with an estimated molecular weight of 20 to 25 KDa. The porcine gelatin (Sigma #G2500) had an estimated molecular weight of 100

KDa. The peristaltic pump used in the DHP synthesis was a Cole-Parmer Masterflex, model number 77120-52.

4.3.2. DHP and DHP-Gel preparations

Lignin DHP was synthesized according to a published method with a few modifications (Terashima et al., 1995). Coniferyl alcohol (200 mg) was added to 200 ml warm sodium phosphate (0.01 M, pH 6.5) buffer. Horseradish peroxidase (HRP) (4 mg) was added to this flask after the buffer temperature dropped below 40° C. In a second flask, hydrogen peroxide was added to 200 ml of buffer to a final concentration of 0.025%. A peristaltic pump was used to combine the contents of the flasks into a single 500 ml flask that initially contained 2 ml of buffer and 1 mg of HRP. Addition of reactants was performed at a rate of approximately 6 ml/min and the contents of the collection flask were allowed to stir for an additional 24 hours upon completion of reactant addition.

DHP-Gel adducts were prepared as above, but porcine or bovine gelatin (quantities shown in Table 4.2) were added to the flask containing coniferyl alcohol prior to the start of the reaction. In the case of DHP-Gel5, gelatin was added following DHP polymerization (i.e., gelatin was added approximately 24 hours after the complete addition of coniferyl alcohol and hydrogen peroxide).

Neat DHP and DHP-Gel adducts were centrifuged at 10k g for 20 min at 4° C. The supernatants were discarded and the samples were re-suspended in DI water and centrifuged again. This was repeated for a total of 5 washings. DHP-Gel adducts were then dried under vacuum at room temperature to obtain yields shown in Table 2. Solutions of neat gelatin and gelatin that had been subjected to the oxidative conditions of DHP preparation were centrifuged as described above and were not found to precipitate.

4.3.3. Fourier-transform infrared spectroscopy

Lignin DHP and DHP-Gel adducts were analyzed using a Bruker Vertex V70 Spectrometer (Bruker Optics Billerica MA) equipped with an MVP-Pro diamond single reflection attenuated total reflectance (ATR) accessory (Harrick Scientific Pleasantville NY), and 100 scans at 6 cm-1 resolution were averaged for each sample using a DTGS detector and scan frequency of 5 kHz. In all cases, the spectrum of the clean diamond crystal was used as the reference spectrum. All spectral manipulations were performed using OPUS 6.0 (Bruker Optics, Billerica MA).

4.3.4. X-ray photoelectron spectroscopy

The spectra were acquired with a Kratos Axis Ultra, using monochromatic Al-Kα X-rays. Analysis chamber pressures were in the mid-10-8 torr range during measurements. Samples were mounted on a 7mm x 7mm piece of Scotch Brand 3M double-sided tape (cat #137). The

74 materials covered the tape well enough to prevent exposure of the glue, and the tape was secured to a piece of OFHC copper which was slightly larger than the tape. All spectra were acquired with the analyzer set in hybrid mode, with the charge neutralizer on. The Pass Energy was set at 80 eV for surveys and 20 eV for high-resolution scans. Step sizes were 0.5 eV and 0.1 eV for survey and high-res scans, respectively. The survey scan dwell time was set at 150 ms, while values for high-resolutions scans varied from 600-2000 ms depending on the peak intensity.

4.3.5. Scanning electron microscopy and energy dispersive X-ray spectroscopy

Scanning electron microscopy (SEM) images were collected on a field emission SEM (FESEM - FEI NanoSEM 630) at 2 or 3 kV under high vacuum (1.7 x 10-6 Torr). Samples were sputter coated with iridium prior to imaging. Characteristic X-rays were collected with an X-Max silicon drift detector (Oxford Instruments) inside the FESEM at 10 kV under low vacuum conditions (0.6 Torr) in order to prevent sample charging. Samples were not sputter coated prior to EDS analysis. Elements were selected and quantified using Aztec Energy Analyser Software (Oxford Instruments).

4.3.6. Nuclear magnetic resonance spectroscopy

NMR spectra were acquired on a Bruker Biospin (Billerica, MA, USA) AVANCE 500 (500 MHz 1H resonance freq.) spectrometer fitted with a cryogenically-cooled gradient probe having inverse geometry, i.e., with the proton coils closest to the sample. Spectra were processed with Bruker’s Topspin 3.1 software, using the central solvent peak as internal reference [δH/δC: dimethyl sulfoxide (DMSO), 2.50/39.5 ppm]. The DHP or DHP-Gel adducts (~50 mg) were placed in an NMR tube (ID: 4.1 mm), swelled in DMSO-d6/pyridine-d5 (4:1 v/v, 500 ul), and subjected to adiabatic 2D-HSQC (‘hsqcetgpsisp2.2’) experiments. Processing used typical matched Gaussian apodization in F2 (LB = -0.3, GB = 0.001), and squared cosine-bell and one level of linear prediction (32 coefficients) in F1 (Mansfield, 2012). For an estimation of the various inter-unit linkage types in DHP and DHP-Gel adducts (Table 3; β-ether/α-OH, β-ether/α-

O-aryl, phenylcoumaran, pinoresinol, and dibenzodioxocin), the well resolved Cα-Hα contours were integrated; no correction factors were used.

4.4. Results

4.4.1. Preparation of DHP-Gel adducts

DHP-Gel adducts were prepared by adding various quantities of porcine or bovine gelatin to lignin DHP as it polymerized in the cases of DHP-Gel1, DHP-Gel2, DHP-Gel3, and DHP-Gel4, or following lignin polymerization in the case of DHP-Gel5. Adduct yields following centrifugation are shown in Table 4.2. The adducts were characterized using IR, EDS, XPS, SEM, and NMR, as detailed below.

Table 4.2. Preparation and yields of DHP and DHP-Gel adducts.

sample CA (mg) PG (mg) BG (mg) yield (mg) yield (%) DHP 200 0 0 124 62 DHP-Gel1 200 200 0 154 39 DHP-Gel2 200 0 80 139 50 DHP-Gel3 200 0 40 159 66 DHP-Gel4 200 0 20 138 63 DHP-Gel5 200 200 0 112 28

CA: coniferyl alcohol, PG: porcine gelatin, BG: bovine gelatin. Yields were calculated by dividing the mass of recovered product by the total mass of reactants (i.e., CA + PG or BG). In the case of DHP-Gel5, porcine gelatin was added following DHP polymerization.

4.4.2. Fourier-transform infrared spectroscopy of DHP-Gel adducts

Fig 4.1 shows FT-IR spectra of neat gelatin, neat DHP, and DHP-Gel adducts. The DHP FT-IR spectrum exhibited bands typical of lignin DHPs (Faix, 1988). The DHP-Gel adducts exhibited three peaks indicative of protein incorporation into the lignin. The peaks were located at approximately 3200 cm-1, 1658 cm-1, and 1540 cm-1, and were previously observed in lignin- protein adducts prepared by reacting DHPs with tripeptides (Diehl et al., in review). These peaks were indicative of protein incorporation but were not specifically diagnostic toward covalent versus non-covalent lignin-protein bonding. Qualitatively, FT-IR spectra showed that DHP-Gel1 apparently contained the most protein, with protein content decreasing through DHP-Gel2, DHP-

Gel3, and DHP-Gel4. Protein content of DHP-Gel5 (gelatin added following lignin polymerization) appeared to lie between that of DHP-Gel3 and DHP-Gel4.

Fig 4.1. FT-IR of neat DHP and DHP-Gel adducts. Y-scaling is arbitrary.

4.4.3. Morphology and nitrogen content of DHP-Gel adducts

Fig 4.2 shows SEM images of neat DHP and DHP-Gel adducts. The neat DHP was composed of smooth spheres of varying sizes, as previously observed (Micic et al., 2003). In the cases of DHP-Gel1, DHP-Gel2, DHP-Gel3, and DHP-Gel4, the samples exhibited spherical morphology, but the sizes and shapes of the spheres varied among samples and within samples.

DHP-Gel4 (lowest concentration of gelatin) exhibited the most variation in particle shape and size—an affect that was reproducible. The particles within these DHP-Gel adducts exhibited bumpy surfaces, which can be seen especially well in Fig. 2, DHP-Gel3, inset. Spherical particles of DHP-Gel5 (gelatin added following polymerization) exhibited smooth surfaces, similar to neat DHP.

Fig 4.2. SEM images of DHP-Gel1 (top left), DHP-Gel2 (top right), DHP-Gel3 (middle left), DHP-Gel4 (middle right), DHP-Gel5 (bottom left) and neat DHP (bottom right). Bar: 2 µm (DHP-Gel3 inset bar: 500 nm, DHP-Gel5 inset bar: 1 µm).

It was perceived that gelatin may not have been homogeneously dispersed throughout the DHP-Gel particles (see Fig 4.3, models A and B). In order to test this hypothesis, elemental analysis data was obtained using both XPS and EDS. XPS is a surface sensitive technique with an approximate information depth of only 10 nm. In contrast, EDS has an information depth of >1 µm when employing an accelerating voltage of 10 kV to a “light” substrate (i.e., an organic substance such as the lignin-gelatin complex). Because the DHP-Gel particles were generally several tens or hundreds of nanometers in diameter (Fig 4.2), XPS analyses were expected to reveal nitrogen content near the surface (i.e., shell) of the particles, while EDS analyses were expected to reveal nitrogen content throughout multiple particles.

Fig 4.3 shows the atomic nitrogen percentages of the DHP-Gel adducts as determined by XPS and EDS (averages were determined by sampling three locations per DHP-Gel complex).

Two sample t-tests (using unequal variances and α = 0.05) showed that DHP-Gel1, DHP-Gel2, DHP-Gel3, and DHP-Gel4 contained no significant differences in nitrogen as determined by XPS and EDS (p-values: DHP-Gel1: 0.215, DHP-Gel2: 0.783, DHP-Gel3: 0.659, DHP-Gel4: 0.212), suggesting a morphology similar to Fig 4.3, model A. In the case of DHP-Gel5 (gelatin added following polymerization), the nitrogen content determined by XPS was significantly higher (p = 0.005) than the nitrogen content determined by EDS. This suggested that gelatin preferentially bound to the lignin surface when added following lignin polymerization, suggesting morphology similar to Fig 4.3, model B.

Fig 4.3. Morphology and nitrogen atomic percentages for DHP-Gel adducts as determined by XPS and EDS (averages were determined by sampling three locations per DHP-Gel complex; error bars are one positive and one negative standard deviation). For DHP-Gel morphological models (A and B), black circles represent individual lignin modules (thin and thick lines represent hydrophilic and hydrophobic domains, respectively), which may aggregate to form macromolecules (Micic et al., 2004). Green lines represent gelatin. XPS and EDS showed that model A best represented DHP-Gel1, 2, 3, and 4, while model B best represented DHP-Gel5.

Within the plant cell wall, lignification of a given region generally occurs following the deposition of all other structural wall polymers, including structural proteins (Donaldson, 2001; Boerjan et al., 2003). In addition, cell wall lignin (especially lignin from the cell corner region of the middle lamella) adopts a similar morphology to the DHP and DHP-Gel adducts observed here (Donaldson, 1994; Hafren et al., 2000; Terashima et al., 2004). This suggests that lignin may surround and/or entangle cell wall proteins during lignification in planta, resulting in

79 morphology similar to Fig 4.3, model A. This physical entanglement, with the potential also for lignin-protein covalent crosslink formation, may explain why protein is often found as a contaminant in lignin extractions (Hatfield et al., 1994; Fukushima and Hatfield, 2001).

4.4.3. Nuclear magnetic resonance spectroscopy of DHP-Gel adducts

Fig 4.4 shows the HSQC NMR spectrum of DHP-Gel1, which was representative of the DHP-Gel adducts. Shifts in orange were added during processing and are representative of gelatin shifts. Gelatin shifts were visible when viewing the HSQC spectrum at a lower contour level than shown in Fig 4.4. There are several reasons why the gelatin shifts were less intense than the lignin shifts. First, the gelatin was the limiting reagent in most cases (except in the cases of DHP-Gel1 and DHP-Gel5). Second, the gelatin was not completely incorporated into the lignin. And third, the DHP-Gel adducts were not fully soluble in the NMR solvent system (DMSO-d6/pyridine-d5). The gelatin shifts could be expected to be depressed in intensity compared to the lignin shifts if the lignin component is more soluble than the gelatin component.

Diehl et al. (2014) previously identified diagnostic NMR shifts for lignin-protein crosslinks. None of these shifts were observed in NMR spectra of DHP-Gel adducts, which suggests that lignin and gelatin did not covalently crosslink. This may not be surprising, as Diehl et al. (in review) also showed that cysteine and tyrosine were the only amino acid residues to substantially crosslink with lignin under similar conditions of DHP preparation, and these residues are almost entirely absent from gelatin (Table 1.1). The apparent lack of DHP-Gel crosslinking suggested that non-covalent interactions were largely responsible for the observed

DHP-Gel interaction. In the case of DHP-Gel1 and DHP-Gel5, equal quantities of gelatin were added to the reactions, with the only difference being whether the gelatin was added during lignin polymerization or after. The DHP-Gel1 (gelatin added during lignin polymerization) showed greater gelatin incorporation compared to DHP-Gel5 (gelatin added after lignin polymerization) (Fig 4.1 and Fig 4.3). This increased gelatin incorporation was probably due to physical entanglement of the gelatin within the lignin, as evidenced by XPS and EDS analyses (Fig 4.3). Though DHP-Gel covalent crosslinks were not readily identified by NMR it may be inappropriate to completely rule out the possibility of crosslink formation, albeit in minor amounts. Isotopically labeled proteins and/or monolignols may be useful toward identifying trace lignin-protein crosslinkages in further in vitro and in vivo experiments.

Fig 4.4. Side chain and aromatic (inset) regions of the HSQC NMR spectrum of DHP-Gel1. Shifts in orange were added during processing. These shifts are indicative of gelatin and can be observed when the HSQC spectrum is viewed at a lower contour level than shown here.

Table 4.3 shows estimates of the lignin inter-unit linkage ratios for the DHP and DHP- Gel adducts as determined by volume integration of the well-resolved HSQC α-shifts. Neat DHP contained linkage ratios typical of DHPs (Terashima et al., 1995; Terashima et al., 2009; Tobimatsu et al., 2012). The variation in inter-unit linkage ratios exhibited no clear trend with regards to the quantity of gelatin added, suggesting that gelatin has no significant effect on the mechanism of lignin polymerization. The observed variation of inter-unit linkage ratios is most likely attributable to the fact that DHP syntheses are inherently difficult to reproduce (Cathala et al., 1998).

Table 4.3. Inter-unit linkage ratios of DHP and DHP-Gel adducts.

HSQC signal ratio (as % of total α-signal)

β-ether/α- β-ether/α- OH aryl β-5 β-β Dibenz. DHP 27.3 1.9 50.3 19.2 1.3 DHP-Gel1 23.5 0.5 53.4 18.1 4.6 DHP-Gel2 23.2 0.4 55.6 20.3 0.4 DHP-Gel3 13.5 0.7 58.7 25.4 1.7 DHP-Gel4 20.8 4.0 49.2 22.0 4.0 DHP-Gel5 32.8 1.3 45.3 16.4 4.1

4.5. Conclusions

DHP-Gel adducts were prepared under biomimetic conditions of lignin polymerization. A variety of methods was used to characterize the DHP-Gel adducts. FT-IR showed incorporation of gelatin into lignin DHP, but was unable to definitively show either the presence or absence of lignin-gelatin covalent linkages. SEM showed that the DHP-Gel adducts generally consisted of spherical particles ranging from tens to hundreds of nanometers in diameter, with morphological details varying among samples. XPS and EDS were used in combination to show that gelatin was relatively evenly dispersed throughout DHP-Gel particles when added during lignin polymerization (Fig 4.3, model A), but aggregated mostly at the surface of DHP-Gel particles when added following lignin polymerization (Fig 4.3, model B). It was interesting to note that covalent crosslinking was not observed by HSQC NMR. This may not be surprising, as Diehl et al. (in review) previously showed that cysteine and tyrosine were the only amino acid residues to substantially crosslink with lignin under similar conditions of DHP preparation, and gelatin is almost entirely lacking in these residues. The observation that gelatin adsorbed to the lignin surface when added to lignin post-polymerization provided further evidence that covalent crosslinking was not necessary to account for the lignin-gelatin interaction.

The lignin-gelatin interaction appears to be essentially non-covalent, and gelatin peptide was dispersed throughout the DHP-Gel particles when it was added over the course of lignin polymerization. In planta, lignification occurs in a pre-deposited polysaccharide and protein matrix. Based on the results of this study it seems plausible that cell wall structural proteins may become surrounded or physically entangled by lignin without necessarily forming covalent crosslinks. It may be possible for structural proteins to serve as initiation sites of lignification without lignin-protein covalent bond formation. Based on previous studies, covalent crosslinking may also occur if cysteine and tyrosine residues are present (Diehl et al., in review). Alternatively, lignin-protein crosslinking may occur at saccharide residues that are found on plant cell wall structural proteins (particularly extensins and arabinogalactan proteins) but not on gelatin (Lamport et al., 2011; Toikka et al., 1998; Wilson and Fry, 1986); the study reported here did not test such a hypothesis. The intimate physical entanglement of the lignin-protein complex may make covalent crosslinking favorable by bringing reactive lignin and protein species into close proximity. The formation of such complexes may not only serve as lignin nucleation sites, but may also help to rigidify and/or waterproof the cell wall. Further research is needed to determine the role of covalent and non-covalent lignin-protein complexes in plant cell walls.

4.6. Acknowledgements

This material is based upon work supported as part of The Center for LignoCellulose Structure and Formation, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, under Award Number DE-SC0001090. Student fellowships were provided by the USDA National Needs Program and the National Science Foundation via the CarbonEARTH program. Many thanks to Julie Anderson, Melisa Yashinski, and Vince Bojan

(Penn State Materials Research Institute) for acquisition of SEM, EDS and XPS data and for valuable discussions, Jenna Ferraraccio for assistance with figure preparation, and John Ralph, Yuki Tobimatsu, and Matt Regner for acquisition of NMR spectra and valuable discussions regarding multidimensional NMR of lignin.

4.7. References

Albersheim, P.; Darvill, A.; Roberts, K.; Sederoff, R.; Staehelin, A. Principles of Cell Wall Architecture and Assembly. In Plant Cell Walls; Garland Science, 2010; pp 227-27 Boerjan, W.; Ralph, J.; Baucher, M. Lignin Biosynthesis. Annu. Re. Plant Biol. 2003, 54, 519- 546. Cathala, B.; Saake, B.; Faix, O.; Monties, B. 1998. Evaluation of the reproducibility of the synthesis of dehydrogenation polymer models of lignin. Polymer Degradation and Stability 59:65-69.

Chapple, C.; Ladisch, M.; Meilan, R. Loosening lignin's grip on biofuel production. Nat. Biotechnol. 2007, 25, 746-748.

Chen, Y. and S. Sarkanen. 2006. Cellulose Chemistry and Technology, 40(3-4): 149-163. "From the macromolecular behavior of lignin components to the mechanical properties of lignin-based plastics." Chen, F.; Dixon, R. A. Genetic manipulation of lignin biosynthesis to improve biomass characteristics for agro-industrial processes. In Vitro Cell. Dev. Biol. - Animal. 2008, 44, S28- S29.

Clark, J. H., Deswarte, F. E. I. and T. J. Farmer. 2009. Biofuels, bioproducts and biorefining, 3: 72-90. "The integration of green chemistry into future biorefineries." Donaldson, L.A. 1994. Mechanical constraints on lignin deposition during lignification. Wood Sci. and Technol. 27:111-118. Donaldson, L.A. 2001. Lignification and lignin topochemistry – an ultrastructural view. Phytochemistry 57: 859-873. Dorrestijn, E., Laarhoven, L. J. J., Arends, I. W. C. E. and P. Mulder. 2000. Journal of analytical and applied pyrolysis, 54: 153-192. "The occurrence and reactivity of phenoxyl linkages in lignin and low rank coal." Eastoe, J.E. 1955. The amino acid composition of mammalian collagen and gelatin. Biochemical Journal 61(4):589-600. Faix O, Beinhoff O. 1988. Journal of Wood Chemistry and Technology, 8 (4): 505-522. "FTIR spectra of milled wood lignins and lignin polymer models with enhanced resolution obtained by deconvolution."

Fukushima, R.S.; Hatfield, R.D. 2001. Extraction and isolation of lignin for utilization as a standard to determine lignin concentration using the acetyl bromide spectrophotometric method. Journal of Agricultural and Food Chemistry 49(7):3133-3139. Gellerstedt, G., Sjoholm, E. and I. Brodin. 2010. The Open Agriculture Journal, 3: 119-124. "The wood-based biorefinery: A source of carbon fiber?"

Hafren, J., Funino, T., Itoh, T., Westermark, U., Terashima, N. 2000. Ultrastructural changes in the compound middle lamella of Pinus thunbergii during lignification and lignin removal. Holzforschung 54:234-240.

Hatfield, R.D.; Jung, H.G.; Ralph, J.; Buxton, D.R.; Weimer, P.J. 1994. A comparison of the insoluble residues produced by the Klason lignin and acid detergent lignin procedures. J. Sci. Food Agric. 65:51-58.

Jung, H. G.; Allen, M. S. Characteristics of plant cell walls affecting intake and digestibility of forages by ruminants. J. Animal Sci. 1995, 73, 2774-2790.

Jung, H.G. Forage lignins and their effects on fiber digestibility. Agron. J. 1989, 81, 33-38.

Lamport, D.T.A.; Kieliszewski, M.J.; Chen, Y.; Cannon, M.C. Plant Phys 2011, 156, 11-19.

Li, X.; Weng, J. K.; Chapple, C. Improvement of biomass through lignin modification. Plant J. 2008, 54, 569-581.

Ludley, F. H. and J. Ralph. 1996. Journal of Agricultural and Food Chemistry, 44: 2942-2943. "Improved preparation of coniferyl and sinapyl alcohols."

Mansfield, S. D.; Kim, H.; Lu, F.; Ralph, J. Whole plant cell wall characterization using solution-state 2D NMR. Nature Protocols. 2012, 7(9), 1579-1589.

Micic, M.; Radotic, K.; Jeremic, M.; Leblanc, R. M. Study of the self-assembly of the lignin model compound on cellulose model substrate. Macromol. Biosci. 2003, 3 (2), 100-106.

Micic, M.; Radotic, K.; Jeremic, M.; Djikanovic, D.; Kammer, S.B. 2004. Study of the lignin model compound supramolecular structure by combination of near-field scanning optical microscopy and atomic force microscopy. Colloids and Surfaces B: Biointerfaces 34:33-40.

Terashima, N., Atalla, R.H., Ralph, S.A., Landucci L.L., Lapierre, C., Monties, B. 1995. Holzforschung, 49: 521-527. "New preparations of lignin polymer models under conditions that approximate cell wall lignification."

Terashima, N., Awano, T., Takabe, K., Yoshida, M. 2004. Formation of macromolecular lignin in ginkgo xylem cell walls as observed by field emission scanning electron microscopy. C. R. Biologies 327: 903-910.

Toikka, M.; Sipila, J.; Teleman, A.; Brunow, Gosta. 1998. Lignin-carbohydrate model compounds. Formation of lignin-methyl arabinoside and lignin-methyl galactoside benzyl ethers via quinone methide intermediates. J. Chem. Soc., Perkin Trans. 1 3813-3818.

Whitmore, F.W. 1978a. Lignin-carbohydrate complex formed in isolated cell walls of callus. Phytochemistry 17:421-425.

Whitmore, F.W. 1978b. Lignin-protein complex catalyzed by peroxidase. Plant Science Letters 13:241-245.

Whitmore, F.W. 1982. Lignin-protein complex in cell walls of Pinus elliottii: Amino acid constituents. Phytochemistry 21(2):315-318.

Wilson, L.G.; Fry, J.C. 1986. Extensin—A major cell wall glycloprotein. Plant, Cell and Environment 9:239-260.

Chapter 5

Searching for lignin-protein linkages in Arabidopsis

5.1. Abstract

In order to explore lignin-protein linkages in planta, Arabidopsis thaliana was grown to maturity and lignin was extracted from the inflorescence stems. Elemental analysis was used to estimate the protein content of the crude Arabidopsis, the Arabidopsis following solvents extraction, and the Arabidopsis lignin extracts. Nuclear magnetic resonance techniques were then used to search for putative lignin-protein covalent linkages. No apparent linkages were identified, but further work is needed in other wild type and mutant plant species, and isotopically labeled monolignols may be useful toward investigating lignin-protein linkages in both wild type and mutant plants.

5.2. Introduction

The previous chapters have shown that lignin crosslinks with several amino acids under ideal conditions (i.e., in neutral, non-polar solvents), and that cysteine and tyrosine crosslink with lignin under biomimetic conditions of lignin polymerization. Finally, in an attempt to identify lignin-protein linkages formed under natural conditions of lignin biosynthesis, Arabidopsis thaliana (wild-type Columbia-0) plants were grown to maturity (8 weeks), then lignin was extracted from the inflorescence stems and characterized.

Arabidopsis lignin is composed mainly of guaiacyl and syringyl type lignin moieties. The lignin content of mature Arabidopsis inflorescence stems has been estimated at around 14% using the Klason method (Chang et al., 2008), and 14-16% using the acetyl bromide method (Yong Bum Park, unpublished data). The quantity of structural protein in mature Arabidopsis cell wall, in terms of dry weight percentage, is unclear. However, Chang et al. (2008) previously showed that extracted Arabidopsis lignin (dioxane and Klason lignin) contained about 3.7% protein contamination. In order to estimate the quantity of proteins in Arabidopsis, nitrogen content was determined for mature Arabidopsis inflorescence stems at three levels of sample preparation: crude ball-milled Arabidopsis, Arabidopsis that was ball-milled and solvents extracted to yield cell wall material, and extracted Arabidopsis lignin. Nitrogen content can then be used to determine protein percentage by multiplying by 6.25 (assuming all nitrogen is due to protein) (Chang et al., 2008; Fukushima and Hatfield, 2001), and in this way the protein content can be monitored throughout the various steps of Arabidopsis lignin extraction.

Lignin was extracted from Arabidopsis following a previously described acidic dioxane (ADL) method (Fukushima and Hatfield, 2004). It has been postulated that this extraction method selectively cleaves α-ether linkages, which should raise concerns regarding the cleavage of putative lignin-protein linkages, as well. However, this method was deemed useful for several reasons. First, it was not possible to extract lignin using the typical milled wood lignin procedure of refluxing the sample in 96:4 dioxane/water. This method has been employed for decades;

86 however, during preliminary investigations with Arabidopsis, only ~2 mg of lignin was extracted per 1 g of Arabidopsis cell wall material, which is extremely inefficient and yields far too little lignin for effective characterization. Furthermore, lignin-protein linkages are expected to be low in quantity in wild type plants, so observing the putative linkages in cellulolytic enzyme lignins or whole cell walls seems unlikely due to very low signal to noise.

Following lignin extraction and protein content estimation, the Arabidopsis lignin was characterized using 2D nuclear magnetic resonance (NMR) spectroscopy. The two most important techniques were heteronuclear single quantum coherence (HSQC) and heteronuclear multiple bond correlation (HMBC) experiments, which were previously shown to be quite useful toward elucidating lignin-protein linkages.

5.3. Experimental

5.3.1. Growth and lignin extraction from Arabidopsis

Arabidopsis samples were prepared as follows. After 4 days of cold treatment at 4°C, Arabidopsis thaliana wild-type (Colombia (Col-0) ecotype) seedlings were grown on 1× MS medium (Murashige and Skoog, 1962) containing 1% sucrose for 1 week, and 500–600 seedlings were transferred onto soil and grown for 7–8 more weeks under 70 µmol m-2s-1 light intensity (day/night: 16/8 h, temperature: 22/16°C). The matured Arabidopsis inflorescence stems were collected and frozen at -80° C. Prior to lignin extraction, Arabidopsis was prepared by grinding in a blender, followed by freeze-drying. Samples (typically 2-3 g) were then Soxhlet extracted with water, ethanol, chloroform, and acetone, for eight hours per solvent. Solvents extracted Arabidopsis was then Wiley milled to pass a 60 mesh screen, then ball milled in a Retsch cryomill (1.5 g Wiley milled Arabidopsis for 2 hr at 10 Hz). Lignin was extracted following the previously described acidic dioxane lignin (ADL) method (Fukushima and Hatfield, 2004). Briefly, 1 g of solvents extracted Arabidopsis was refluxed for 45 min in 20 ml of 90:10 dioxane/2 M HCl (aq). The solubilized lignin was then filtered through a WhatmanTM glass microfiber filter, and the cell wall residue was rinsed with 96:4 dioxane/water. The dioxane filtrates were combined and neutralized with sodium bicarbonate, filtered through a 0.45 µm nylon membrane filter, then concentrated under reduced pressure. The concentrated lignin solution (~1-2 ml) was added dropwise to ~40 ml DI water in a centrifuge tube, then centrifuged at 10k g at 5° C for 20 min. The aqueous supernatant was poured off and saved, then the lignin pellet was resolubilized using a minimum volume of dioxane. Ether was added to the centrifuge tube and the sample was centrifuged at 10k g at 5° C for 15 min, and this solubilization followed by ether washing and centrifugation was repeated for a total of two cycles. The ether supernatants were then discarded and the lignin was freeze dried and stored at 4° C for future characterization. Typically, lignin yields were 30-35 mg per 1 g of solvents extracted Arabidopsis cell wall material.

5.3.2. Elemental analysis of Arabidopsis lignin

Nitrogen weight percentages were determined using a CE Instruments EA 1110 CHNS-O elemental analyzer. Approximately 3 mg of sample were massed to the nearest ten-thousandth of a milligram and analyzed according to the manufacturer’s instruction. Protein content was determined by multiplying nitrogen percentage by 6.25, as described previously (Chang et al., 2008; Fukushima and Hatfield, 2001).

5.3.3. Nuclear magnetic resonance spectroscopy of Arabidopsis lignin

NMR spectra were collected in DMSO-d6/pyridine-d5 (4:1 v/v, 500 ul). DMSO- d6/pyridine-d5 was chosen because it is a preferred solvent for NMR of lignin DHP, milled wood lignin (MWL), and whole cell walls (Kim and Ralph, 2010). NMR spectra were acquired on a Bruker Biospin (Billerica, MA, USA) AVANCE 500 (500 MHz 1H resonance freq.) spectrometer fitted with a cryogenically-cooled gradient probe having inverse geometry, i.e., with the proton coils closest to the sample. Spectra were processed with Bruker’s Topspin 3.1 software, using the central solvent peak as internal reference (δH/δC: dimethyl sulfoxide (DMSO), 2.50/39.5 ppm). Lignin (~20 mg) was placed in an NMR tube (ID: 4.1 mm), swollen homogeneously in DMSO-d6/pyridine-d5 (4:1 v/v, 500 ul), and then subjected to adiabatic 2D- HSQC (‘hsqcetgpsisp2.2’) experiments using the parameters described by Mansfield et al. (2012). Processing used typical matched Gaussian apodization in F2 (LB = -0.3, GB = 0.001), and squared cosine-bell and one level of linear prediction (32 coefficients) in F1 (Mansfield et al., 2012).

5.4. Results and discussion

5.4.1. Lignin extractions from Arabidopsis

Arabidopsis was grown to maturity, then inflorescence stems were harvested and prepared for lignin extraction by solvents extracting and cryomill grinding. The grinding time of approximately 2 hr per 1.5 g of Arabidopsis resulted in highly variable particle sizes (Fig 5.1). Longer grinding times may be necessary, although increased sample alteration following increased grinding times is always cause for concern. The grinding times employed here allowed for the extraction of ~30-35 mg of acidic dioxane lignin (ADL) per g of Arabidopsis cell wall material. This was similar to previously reported ADL yields for grassy plants such as alfalfa and red clover (Fukushima and Hatfield, 2001, 2004). This material was then subjected to protein content estimation and nuclear magnetic resonance spectroscopy, as described below.

Fig 5.1. Optical microscopy of solvents extracted and ball milled Arabidopsis cell wall material. Particle size distribution is highly variable. Scale bar = 50 µm.

5.4.2. Protein content of Arabidopsis extracts

Protein contents of crude Arabidopsis, solvents extracted Arabidopsis, and Arabidopsis lignin were estimated by multiplying the measured nitrogen atomic percentages by a factor of 6.25, as previously described (Chang et al., 2008; Fukushima and Hatfield, 2001). It was found that crude Arabidopsis inflorescence stems contained approximately 5.31% protein, Arabidopsis that had been solvents extracted contained 4.94% protein, and Arabidopsis ADL contained approximately 3.75% protein. This was very similar to the protein content determined by Chang et al. (2008) for dioxane and Klason Arabidopsis lignins, but was considerably higher than that of Loblolly ADL, which was typically <1%. This may be expected due to the prominent secondary cell walls in Loblolly and other plants with a strong tree habit, and the general lack of protein in secondary cell walls. Because of the increased protein content in Arabidopsis (and presumably grasses and other non-grasses exhibiting a grass-like growth habit), wild type and mutant Arabidopsis lines may be useful for future investigations of lignin-protein linkages.

Table 5.1. Nitrogen content and estimated protein content of Arabidopsis extracts.

N % Protein % Crude Arabidopsis 0.85 5.31 Solvents extracted Arabidopsis 0.79 4.94 Arabidopsis acidic dioxane lignin 0.60 3.75

5.4.3. Nuclear magnetic resonance spectroscopy of Arabidopsis lignin

Acidic dioxane lignin isolated from Arabidopsis inflorescence stems was analyzed using HSQC and HMBC NMR techniques in DMSO-d6/pyridine-d5. Table 5.2 shows estimates of the lignin inter-unit linkage ratios as determined by volume integration of the well-resolved HSQC α-shifts, and the ratios were typical of dicotyledonous G/S lignins (Capanema et al, 2004). An HSQC spectrum of Arabidopsis ADL is shown in Fig 5.2; however, the peaks discussed in further detail below were generally too weak to be observed at the contour levels shown.

Table 5.2. Inter-unit linkage ratios of Arabidopsis ADL.

HSQC signal ratio (as % of total α-signal) β-ether/α-OH β-ether/α-aryl β-5 β-β Dibenz. 70.6 tr 17.7 9.7 2.1 Estimates of the lignin inter-unit linkage ratios as determined by volume integration of the well- resolved HSQC α-shifts are shown. tr = trace.

A very weak shift was observed in the HSQC spectrum at 4.4/50.8 ppm. This is close to the observed α-shift of a lignin-cysteine linkage (4.4/50.6) identified in Chapter 2. However, the corresponding HMBC spectrum did not identify this shift as an α-shift, and thus the likelihood of this shift being attributable to a lignin-cysteine linkage seems low. This demonstrates the importance of using multiple NMR techniques when investigating putative lignin-protein crosslinks. No other putative lignin-protein shifts were identified in the HSQC and HMBC spectra. Furthermore, essentially no lignin α-ester and only very minor quantities of α-ether linkages were observed in the HSQC spectrum. This suggests that even lignin-hemicellulose linkages were essentially absent from the Arabidopsis ADL. This may have been due to the harshness of the extraction procedure, and less harsh lignin extraction techniques may be necessary in order to optimize the chances of lignin-protein linkage identification.

Fig 5.2. HSQC NMR of Arabidopsis acidic dioxane lignin. Typical lignin shifts and residual dioxane and pyridine solvent shifts are labeled. Cinnamyl end groups and dibenzodioxocin structures were not abundant enough to be seen at the contour levels shown. Lignin-protein linkages were not apparent even at low contour levels.

5.5. Conclusions

Wild type Arabidopsis was grown to maturity and inflorescence stems were harvested and lignin extracted using the acidic dioxane (ADL) method. It was found that the ADL contained a significant protein content (3.75%), so the lignin was characterized using HSQC and HMBC NMR techniques. No lignin-protein linkages were identified. Furthermore, the lignin appeared to be essentially free of lignin-carbohydrate linkages. The acidic dioxane extraction method may have cleaved such linkages if they were in fact present in the wild type Arabidopsis, and it may be beneficial to use a milder lignin extraction procedure. Alternatively, lignin-protein linkages may be very low in abundance (or non-existent), and thus below the NMR signal to

91 noise limit. Exploration of lignin-protein linkages in mutant plant lines and/or using isotopically labeled monolignols may help probe this question.

5.6. Acknowledgements

This material is based upon work supported as part of The Center for LignoCellulose Structure and Formation, an Energy Frontier Research Center funded by the U.S. Department of Energy, Office of Science, Office of Basic Energy Sciences under Award Number DE- SC0001090. Student fellowships were provided by the USDA National Needs Program and the National Science Foundation via the CarbonEARTH program. Many thanks to Ephraim Govere for assistance in acquiring elemental analysis data.

5.7. References

Capanema, E.A.; Balakshin, M. Y.; Kadla, J.F. J. Agric. Food Chem. 2004, 52, 1850-1860. Chang, X.F.; Chandra, R.; Berleth, T.; Beatson, R.P. J. Agric. Food Chem. 2008, 56, 6825-6834.

Fukushima, R.S.; Hatfield, R.D. J. Agric. Food Chem. 2001, 49, 3133-3139.

Fukushima, R.S.; Hatfield, R.D. J. Agric. Food Chem. 2004, 52, 3713-3720.

Kim, H.; Ralph, J. Org. Biomol. Chem. 2010, 8, 576-591.

Mansfield, S.D.; Kim, H.; Lu, F.; Ralph, J. Nat. Protoc. 2012, 7(9), 1579-1589. Murashige, T.; Skoog, F. Physiol. Plant 1962, 15, 473-497.

Chapter 6

Conclusions

6.1. Research summary

Lignin is a heterogeneous, aromatic polymer that is biosynthesized in the cell walls of almost all lands plants. It is economically relevant to several industries, especially pulp and paper, agricultural, and biofuels industries, where lignin’s natural recalcitrance to extraction and degradation pose problems. Additionally, lignin could potentially be used as a feedstock for many renewable products, including plastics, activated carbons, carbon fibers, solid and liquid fuels, and other specialty chemicals, but many of these lignin utilization schemes are beyond the reach of currently available technology. To this end, it is important that we continue to focus on both fundamental and applied advances in lignin and its related fields.

The plant cell wall is a complex matrix composed of structural polymers such as cellulose, hemicelluloses, pectins, lignin, and proteins, as well as lower molecular weight organic compounds. Lignin polymerization occurs within the pre-deposited polysaccharide and protein matrix, causing lignin to interact with its local chemical environment. This results not only in non-covalent interactions, but also in covalent crosslinking with hemicelluloses, and perhaps other matrix components, such as proteins. It has been hypothesized that lignin-protein linkages may be important in both wild type and mutant plant lines, yet lignin-protein linkages have not been previously described. This work helps fill that gap in the literature by describing the preparation and characterization of lignin-protein linkages.

In the first study, a low molecular weight quinone methide, representative of native lignin quinone methide structures, was reacted with single amino acids. It was found that under these ideal conditions, a diverse array of amino acids, including those bearing thiol, amine, carboxylic acid, and hydroxyl functional groups on their side chains, reacted with lignin quinone methides to form lignin-protein model compounds. Characterization of these compounds with NMR resulted in the identification of diagnostic lignin-protein crosslink signatures, which were helpful in identifying lignin-protein linkages in more complex model systems, and should also be helpful toward identifying lignin-protein linkages in native lignins.

The second study expanded upon the first by exploring the reactivities of amino acids toward lignin quinone methide intermediates under biomimetic conditions. Specifically, the amino acids were incorporated into short peptide chains and exposed to lignin dehydrogenation polymer as it polymerized in aqueous media. Using NMR, it was possible to show that cysteine and tyrosine-containing peptides were covalently incorporated into the synthetic lignin polymer, while other amino acids were almost entirely inert under such biomimetic conditions. This suggests that cysteine and/or tyrosine-rich proteins may be the most likely to covalently crosslink with lignin in planta. In this study it was also shown that Fourier-transform infrared

93 spectroscopy and elemental analysis (specifically energy dispersive X-ray spectroscopy in this case) were useful toward showing lignin-protein interactions.

A third study investigated the interactions between lignin DHP and gelatin protein. Gelatin was chosen because it shares similar characteristics with plant cell wall structural proteins, but is much more readily available. In addition, gelatin lacks cysteine and tyrosine, which were the only two amino acids shown in this work to covalently crosslink with lignin in aqueous media. Thus, the observed interaction between lignin and gelatin was considered worthy of investigation. SEM, EDS, and FT-IR showed that gelatin was incorporated into lignin DHP, but a lack of characteristic lignin-protein NMR signatures suggested that the interaction was largely (or entirely) non-covalent. This indicates that lignin-protein non-covalent interactions may be important in planta, and further work is necessary.

Finally, in a fourth study, an attempt was made to identify lignin-protein linkages in the wild type dicot plant, Arabidopsis. Unfortunately, the cell wall protein content was shown to be very low in this wild type plant, and NMR was unable to reveal lignin-protein linkages in acidic dioxane lignin extracts. This does not necessarily rule out the formation of these linkages, but may merely suggest that linkage abundance is below the detection limit of NMR. Furthermore, this study did not investigate the abundance or importance of linkages in mutant plant lines, and additional studies are needed.

Though the work described here has allowed for more efficient and reliable characterization of lignin-protein linkages in model systems, more work is necessary, especially regarding native plant systems. The final section of this document will outline some ways in which future investigations could be carried out.

6.2. Future endeavors

To further investigate lignin-protein linkages in more realistic model systems, it may be useful to lignify model cell walls, or cell walls that have been isolated from native plants, then characterize these walls using NMR and other techniques. Cybulska et al. (2010) and Dammstrom et al. (2005) previously described the preparation of model cell walls using either pure cellulose, or cellulose with the incorporation of hemicelluloses and/or pectins. In addition, Uraki et al. (2011) demonstrated the preparation of cellulose-based, honey-comb shaped model cell walls (Fig 6.1).

Fig 6.1. Cell wall models. Left: cellulose, pectin, and xyloglucan model (Dammstrom et al., 2005). Right: honey-comb cellulose model (Uraki et al., 2011). Incorporation of proteins into these models, followed by lignin DHP polymerization, may allow for investigations into lignin- protein crosslinking.

I propose the preparation of similar cell walls, but with the added inclusion of proteins of interest. Peroxidases could be incorporated into the cell wall models during assembly, or flowed over the models after assembly. Lignin monomers and hydrogen peroxide would then be flowed over the cell wall models, causing lignin polymerization to take place. The entire cell wall model would then be characterized without disturbing the micro and macro structures that formed, as this might provide the most insight into the lignin polymerization mechanisms. Techniques such as NMR could be used to explore putative lignin-protein covalent linkages, while other techniques could be used to explore the topology and distribution of the lignin DHP. The exploration of potential lignin nucleation sites would be particularly interesting (for example, lignin may nucleate at sites rich in hemicelluloses, pectins/pectates, peroxidases, structural proteins, or show no particular nucleation pattern at all). Visualization techniques such as SEM and AFM could be useful in probing this question. Also, the use of fluorescent monolignols (and perhaps additional labeling of other components) could be beneficial (Tobimatsu et al., 2011). Monitoring lignin distribution and topology at various time points of polymerization might also be helpful towards understanding the lignin polymerization mechanism.

In addition to more advanced model compound studies, studies using mutant plant lines are also warranted. While lignin-protein linkages may exist in wild type plant lines, and studies involving wild type plants should not be dismissed, mutant plant lines may show the most promise toward identifying and characterizing lignin-protein linkages. These mutant plant lines should be engineered to secrete cysteine and/or tyrosine-rich peptides into the plant cell walls, as these amino acids have been shown to be most reactive toward lignin in model studies. A mutant plant line meeting these characteristics has already been prepared, although lignin-protein covalent linkages have yet to be identified at the time of document preparation (Liang et al., 2008; Xu et al., 2013). It is hoped that lignin-protein linkages will be further explored in mutant

95 plant lines, as practical implications toward lignin extractability have been demonstrated in pilot studies.

It seems appropriate to briefly address a method that could be useful toward enhancing the identification of putative lignin-protein linkages via NMR. It has previously been shown that NMR is a powerful technique for characterizing plant cell walls and their constituent components, and this work has shown that the technique can also be applied to the identification of lignin-protein linkages. However, when exploring lignin-protein linkages in complex chemical systems, such as those that exist in planta, achieving adequate signal to noise is expected to become a problem. This is because the ratio of lignin-protein linkages to all other chemical structures in the sample is expected to be exceedingly low. Isotopic labeling experiments may help address this problem. Previously, α-13C coniferyl alcohol glucoside (coniferin) was prepared and fed to live ginkgo (Xie and Terashima, 1991), then the lignin was characterized via NMR and the α-shifts were shown to exhibit increased signal to noise. A possible route to α-13C coniferyl alcohol is shown in Fig 6.2 (slightly modified compared to Xie and Terashima’s original route to α-13C coniferin). This labeled coniferyl alcohol could be useful for identifying lignin-protein linkages in complex in vitro or in vivo systems, because NMR α-shifts of lignin- protein linkages (as well as standard lignin α-shifts) would exhibit greater signal to noise compared to spectra of unlabeled lignins (Fig 6.3).

Fig 6.2. Proposed synthetic route to α-13C coniferyl alcohol.

Fig 6.3. Standard lignin α-shifts (red) and α-shifts of lignin-protein linkages (red squares). These shifts would exhibit greater signal to noise compared to standard lignin shifts (black) in spectra of α-13C-enriched lignin, perhaps allowing for easier identification of putative lignin-protein linkages.

Efficient extraction and utilization of lignin is of great economic importance. In spite of this, there is still much about the lignin polymer that remains unknown. This work has illuminated fundamental aspects of lignin-protein linkage formation in an attempt to better understand how lignin interacts with the protein component of plant cell walls. It is hoped that this research will translate into a practical means of reducing lignin’s recalcitrance toward degradation and extraction.

6.3. References

Cybulska, J.; Vanstreels, E.; Tri Ho, Q.; Courtin, C.M.; Craeyveld, V.V.; Nicolai, B.; Zdunek, A.; Konstankiewicz, K. Mechanical characteristics of artificial cell walls. J. of Food Eng. 2010, 96, 287-294.

Dammstrom, S.; Salmen, L.; Gatenholm, P. The effect of moisture on the dynamical mechanical properties of bacterial cellulose/glucuronoxylan nanocomposites. Polymer 2005, 46, 10364- 10371.

Liang, H.; Frost, C.J.; Wei, X.; Brown, N.R.; Carlson, J.E.; Tien, M. Improved sugar release from lignocellulosic material by inducing a tyrosine-rich cell wall peptide gene in poplar. Clean 2008, 36(8), 662-668.

Tobimatsu, Y.; Davidson, C.L.; Grabber, J.H.; Ralph, J. Fluorescence-tagged monolignols: Synthesis, and application to studying in vitro lignification. Biomac. 2011, 12, 1752-1761.

Uraki, Y.; Tamai, Y,; Hirai, T.; Koda, K.; Yabu, H.; Shimomura, M. Fabrication of honeycomb- patterned cellulose material that mimics wood cell wall formation processes. Mat. Sci. and Eng. C 2011, 31, 1201-1208.

Xie, Y.; Terashima, N. Selective carbon 13-enrichment of side chain carbons of ginkgo lignin traced by carbon 13 nuclear magnetic resonance. Mokuzai Gakkaishi 1991, 37(10), 935-941.

Xu, Y.; Chen, C.; Thomas, T.P.; Azadi, P.; Diehl, B.; Tsai, C.; Brown, N.; Carlson, J.E.; Tien, M.; Liang, H. Wood chemistry analysis and expression profiling of a poplar clone expressing a tyrosine-rich peptide. Plant Cell Rep. 2013, 32, 1827-1841.

Vita Brett Galen Diehl

Education May, 2014 Ph.D. Biorenewable Systems Department of Agricultural and Biological Engineering The Pennsylvania State University, University Park, PA

May, 2009 B.S. Wood Products, Processing and Manufacturing Option School of Forest Resources The Pennsylvania State University, University Park, PA

Research 2009-2014 Graduate Research, Penn State, University Park, PA • Illuminated fundamental aspects of lignin-protein linkages, which may influence the physical and chemical properties of plant cell walls impacting industries such as agriculture, pulp and paper, and biofuels.

2008 NSF Research Experience for Undergraduates, Penn State, University Park, PA • Researched cellulose-producing Acetobacter xylinum and methods of generating biofuels from cellulosic materials.

2007 Paid Internship, Armstrong World Industries, Lancaster, PA • Studied exotic hardwood species using scanning electron microscopy.

Publications B.G. Diehl, H.D. Watts, J.D. Kubicki, M.R. Regner, J. Ralph, N.R. Brown. Towards lignin- protein crosslinking: Nucleophilic amino acid adducts of a lignin model quinone methide. Cellulose, accepted January, 2014, not yet published.

B.G. Diehl, N.R. Brown, C.W. Frantz, M.R. Lumadue, F. Cannon. Effects of pyrolysis temperature on the chemical composition of refined softwood and hardwood lignins. Carbon (2013), 60: 531-537.

Y. Xu, C. Chen, T.P. Thomas, P. Azadi, B.G. Diehl, C. Tsai, N. Brown, J.E. Carlson, M. Tien, H. Liang. Wood chemistry analysis and expression profiling of a poplar clone expressing a tyrosine-rich peptide. Plant Cell Reports (2013), 32(12): 1827-1841.

F. Cong, B.G. Diehl, J.L. Hill, N.R. Brown, M. Tien. Covalent bond formation between amino acids and lignin: Cross-coupling between proteins and lignin. Phytochemistry (2013), 96: 449- 456.

All data collected, preparing for publication, target journal is Biomacromolecules: B.G. Diehl, N.R. Brown. Lignin crosslinks with peptides under biomimetic conditions.

All data collected, preparing for publication, target journal is Journal of Applied Polymer Science: B.G. Diehl, N.R. Brown. Characterization of lignin-gelatin complexes.