Biochemical Control Aspects in Lignin Polymerization

Anders Holmgren

Doctoral Thesis

Royal Institute of Technology School of Chemical Sciences and Engineering Department of Fibre and Polymer Technology Division of Wood Chemistry and Pulp Technology

Stockholm 2008

Biochemical Control Aspects in Lignin Polymerization

Supervisors:

Professor Gunnar Henriksson

and

Dr Liming Zhang

AKADEMISK AVHANDLING

som med tillstånd av Kungliga Tekniska Högskolan i Stockholm framlägges till offentlig granskning för avläggande av teknologie doktorsexamen fredagen den 29 februari 2008, kl 10:00 i STFI-salen, STFI-Packforsk, Drottning Kristinas Väg 61. Avhandlingen försvaras på engelska.

Paper I Reproduced by permission of The Royal Society of Chemistry

TRITA-CHE-Report 2008:10 ISSN 1654-1081 ISBN 978-91-7178-864-1 Abstract

Lignins are produced by all vascular plants and they represent one of the most abundant groups of biopolymers in nature. Lignin chemistry research, which has been of great importance for the progress of pulping technologies, has been plagued by the difficulties of its isolation and characterization. The pioneering work of Karl Freudenberg in the 1950’s with synthetic models of lignin paved the way for a detailed structural characterization of many lignin substructures. His work with the so-called “synthetic lignins” or dehydrogenative polymers (DHP) also laid a foundation for understanding how different lignin substructures are formed, reinforcing the already existing theory of lignin polymerization. However, subsequent structural characterizations of DHPs and lignins have repeatedly put this theory to the test. In the past decade, even a new radically different hypothesis for lignin polymerization has emerged and is sustained by a few researchers in the field. In this work, DHPs were produced from phenolic monomers, mostly coniferyl alcohol, a common lignin monomer, in a variety of reaction conditions. This was done in order to establish how different chemical factors, potentially active in the plant cell wall during lignin polymerization, influence the polymer’s final properties. In the presence of nicotine amide adenine dinucleotide (NADH), a quinone methide model, which is an intermediate formed during lignin polymerization, was effectively reduced. An equivalent reduced structure was produced during DHP synthesis in the presence of NADH. These studies showed that reduction might take place during oxidative polymerization, possibly explaining how reduced lignin structures are formed in the plant cell wall. Another reductive agent, ascorbic acid, was also tested during synthesis of DHPs. It displayed a totally different effect than NADH, probably due to its anti-oxidant nature, by altering the final amounts of certain inter-unit substructures, in favour of β-O-4′ structures, which are so prominent in natural lignins. Furthermore, the new suggested model for lignin polymerization, stating that lignin itself possesses the ability for template replication, was tested by synthesizing DHPs in the presence of a simple β-β′ substructure model. The DHPs produced the same amounts of β-β′ substructures as a control synthesis without the model structure, indicating that no replication had occurred. Finally, the role of the monolignol γ-carbon oxidation state in lignin polymerization, was studied. Hypothetically, lignin- like polymers could be produced by the plant, using monolignol biosynthetic precursors which exhibit γ-carbonyl groups instead of an alcohol group, like the common lignin monomer. Synthetic lignins produced with ferulic acid, coniferaldehyde and the normal monolignol, coniferyl alcohol, displayed important differences in chemical and physical properties. Both the ferulic acid and coniferaldehyde polymers exhibited almost no saturated inter-unit substructures and very few cyclic structures, both of which are very common in coniferyl alcohol dehydrogenative polymers and natural lignins. This could have significant implications for the formation of an important type of lignin carbohydrate complexes (LCC). Also the hydrophobicity of the alcohol-type polymer was lower than the other two. The biological implications of all these findings are discussed and some suggestions are made to explain how all these factors might affect lignin polymerization and structure in nature. Sammanfattning

Lignin tillverkas av alla kärlväxter och är därmed en av de vanligaste naturligt förekommande polymererna på jorden. Forskningen kring ligninets kemi, som har varit viktig för utvecklingen av massatillverkningsprocesser, har länge lidit av svårigheter vad gäller dess isolering och karaktärisering. Karl Freudenbergs arbete med syntetiska ligninmodeller (DHP) på 1950-talet lade grunden för den kemiska strukturbestämningen av många av ligninets viktiga bindningstyper. Genom dessa upptäckter förstärktes den dåvarande, och idag allmänt accepterade, teorin för ligninpolymeriseringen. Efterföljande forskning kring DHP- och ligninstrukturen har vid upprepade tillfällen satt denna teori på prov. Några forskare har även utvecklat ytterligare en hypotes, som skiljer sig markant från den befintliga teorin. I föreliggande arbete har ligninmodeller (DHP) framställts av olika fenoliska monomerer och i olika reaktionsförhållanden med syftet att försöka bestämma hur och vilka kemiska faktorer, som kan vara aktiva i växternas cellväggar, påverkar ligninpolymeriseringen och därmed ligninets egenskaper. Studier med två biologiskt viktiga reduceringsmedel visade att sådana ämnen kan spela en roll i ligninpolymeriseringen och påverka dess struktur. För det första påvisades hur en kinonmetidmodell, som föreställer en vanlig ligninpolymeriseringsintermedjär, kunde reduceras av nikotinamid adenin dinukleotid (NADH) och bilda en så kallad reducerad ligninstruktur. Dessutom användes samma ämne, NADH, under polymerisering av DHP, vilket producerade bl a motsvarande reducerade strukturer. Dessa studier påvisade hur reduktion kan äga rum under oxidativ polymerisering och därmed ge en förklaring till hur reducerade ligninstrukturer bildas i cellväggen. För det andra visades att bildningen av vissa bindningstyper (β-O- 4′, den vanligaste bindningstypen i lignin) gynnades framför andra när DHP syntetiserades i närvaro av en antioxidant, askorbinsyra, som har rapporterats förekomma i cellväggen. En del av arbetet uppmärksammar den nya modellen som framhåller att ligninpolymeriseringen står under strikt biokemisk kontroll genom att påstå bl a, att lignin kopierar sig självt under polymeriseringen (eller replikerar sig) genom ”template” polymerisering. Ett experiment där DHP syntetiserades i närvaro av en enkel modell, en β-β′ struktur som är en vanligt förekommande bindningstruktur i lignin, visade att mängden β-β′ strukturer inte ökade överhuvudtaget i modellens närvaro jämfört med en motsvarande syntes utan modellen. Därigenom kunde den enligt hypotesens förväntade kopiering inte påvisas i detta fall. Slutligen har också den roll studerats, som monolignolernas (ligninmonomererna) kemiska struktur spelar i ligninpolymeriseringen med avseende på γ-kolets oxidationstal. I teorin skulle någon ligninlik polymer kunna bildas i cellväggen med vissa av monolignolernas biosyntetiska föregångare som har en aldehyd- eller karboxylsyragrupp i γ-kolet i stället för en alkoholgrupp, som de vanliga monolignolerna har. DHP har syntetiserats med ferulsyra, koniferylaldehyd och koniferylalkohol (en vanlig monolignol) var för sig och polymerernas kemiska och fysiska egenskaper har jämförts med varandra. De mer oxiderade monomererna, ferulsyra och koniferylaldehyd (båda med karbonylgrupper) bildade mera hydrofobiska polymerer och dessutom förekom i stort sett inga saturerade bindningstyper, vilka annars är vanliga i koniferylalkohol-DHP och i naturligt lignin. Betydelsen för bildningen av en viktig typ av lignin-kolhydrat komplex (LCC) behandlas därefter. Alla resultat diskuteras i ett biologiskt sammanhang och förslag lämnas på hur alla dessa faktorer påverkar ligninpolymeriseringen och därigenom också dess struktur.

If we knew what it was we were doing, it would not be called research, would it? Albert Einstein

List of Papers

This thesis is based on the following works which will be referred to by their numbers:

Paper I A. Holmgren, G. Brunow, G. Henriksson, L. Zhang and J. Ralph. “Non- Enzymatic Reduction of Quinone Methides during Oxidative Coupling of Monolignols: Implications for the Origin of Benzyl Structures in Lignins.” Organic & Biomolecular Chemistry, 2006, 4, 3456-3461.

Paper II A. Holmgren, G. Henriksson and L. Zhang. “Effects of a Biologically Relevant Anti-oxidant on the Dehydrogenative Polymerization of Coniferyl Alcohol.” Manuscript

Paper III A. Holmgren, L. Zhang and G. Henriksson. “In vitro Monolignol Dehydrogenative Polymerization in the Presence of a Candidate Dimer Template Model.” Manuscript

Paper IV A. Holmgren, M. Norgren, L. Zhang and G. Henriksson. “On the Role of the Monolignol γ-carbon in Lignin Biosynthesis.” Manuscript

The author’s contribution to each paper:

Papers I-III Principal author, performed all experimental work.

Paper IV Principal author, performed all experimental work except for the preparation of spin-coated surfaces and their characterization.

Related Material:

A. Holmgren, L. Zhang, G. Brunow and G. Henriksson. “Origin of Reduced Lignin Structures: Quinone Methides” 13th ISWFPC Proceedings, Auckland NZ, 2005, 3, 147-15 Table of contents

INTRODUCTION 1 OBJECTIVES 1

BACKGROUND 2 BIOLOGICAL PERSPECTIVES 2 LIGNINS STRUCTURE AND CHEMISTRY 5 LIGNIN BIOSYNTHESIS 9 LIGNIN AMONG OTHER POLYCYCLIC BIOPOLYMERS 15

METHODS 19

RESULTS AND DISCUSSION 22 REDUCTION DURING OXIDATIVE COUPLING (PAPER I) 22 EFFECTS OF AN ANTI-OXIDANT DURING LIGNIN POLYMERIZATION (PAPER II) 26 TESTING THE LIGNIN TEMPLATE POLYMERIZATION MODEL (PAPER III) 30 ON THE ROLE OF THE MONOLIGNOL γ-CARBON (PAPER IV) 33

CONCLUSIONS 42

ACKNOWLEDGEMENTS 44

REFERENCES 45

GLOSSARY 59

Introduction

In spite of more than 100 years of research in the field of wood chemistry, neither the structure of lignins* nor their polymerization is fully understood. Much of the research on lignins has been driven by the development of pulping technologies and it has therefore been focused on their degradation and chemical modification to improve pulp properties. So a considerable amount of fundamental knowledge has been obtained over the years making lignins the best known biopolymers of their kind. Nevertheless, in comparison with the knowledge collected on other common biopolymers such as cellulose, proteins and DNA, the understanding of lignins is vague. Merely comparing the available structural data on lignins with these other biopolymers makes them look like chaotic and complex compounds. Yet the plant, which deposits lignin into the walls of its cells, must somehow be able to control its structure and consequently its properties.

Lignins’ particular chemical and physical properties are believed to depend on the polymerization process, which leads to the lignification of the plant cell wall. The polymerization step in lignin biosynthesis has been a matter of discussion and speculation for a long time and recently a debate has arisen over the control mechanisms at work in the plant cell wall. This is the main subject of the present work.

This thesis starts with a background which describes lignins from two different but related contexts: biology and chemistry. The biosynthesis of lignins is also considered in the light of our knowledge to date and a special focus is placed on the polymerization step by presenting the currently accepted theory and its challenges. In addition, lignins and their biopolymerization are compared to other natural polycyclic polymers: melanins. Subsequently there is a general description of the experimental methods applied during the research that has led to the present thesis, followed by a summary of the results obtained and some discussion. The conclusions in summary form are presented at the end, and additional detailed information can be found in the appended articles.

Objectives It is the aim of this work to shed more light into how plants control lignin polymerization and structure. Understanding what roles the different biochemical factors play in building lignin is useful, e.g., when “engineering” a lignin structure that will improve pulping technologies. Based on existing models for lignin polymerization, experiments have been performed to gain understanding on the potential biochemical factors influencing the polymerization process. Factors such as reductive agents, possible template effects and the importance of the monomeric chemical structure were assessed for their potential influence upon the structure of the lignin polymer.

* In the present work, the term “lignins” is used to refer to the wide variety of naturally occurring lignin-type polymers that fulfil similar functions in the cell walls of plants.

1 Background

Biological Perspectives

The Appearance and Roles of Lignins In plant history, the appearance of vascular tissues such as xylem and phloem implied a cellular specialization and a dedicated infrastructure for water and nutrient transport. The emergence of the first vascular plants, around 430 million years ago, is associated with the appearance of lignins [1- 3]. Lignins are therefore believed to have been crucial for the development of large land plants [4]. Today, there is a great variety of vascular plants including club mosses, horsetails, ferns and the seed-based gymnosperms and angiosperms. Woody plants, called softwoods and hardwoods, are part of the gymnosperm and angiosperm groups respectively. Vascular plants have the ability to grow taller than non-vascular plants, a factor which is important in the competition for sunlight. Their rigid structures have efficiently overcome the challenge of gravity; as is evident in certain redwoods and eucalypts which reach heights of over 100 meters. The ability to grow tall is also dependent on an effective system for the distribution of water to the whole plant. Lignins are believed to play a role which is essential for both rigidity and water transport.

Lignins can be found mostly in the cell walls and middle lamella of tissues involved in mechanical support and water transport like phloem fibres, schlerenchyma cells and mainly in the xylem tissue [1,5]. The xylem exhibits a remarkable morphological variation and specialization at the cellular level. In some cases the same cell type, tracheids, is responsible for both support and water conduction. Then there are parenchyma cells, which are for storage. Another type of diversification and specialization can be observed in the difference between earlywood and latewood tracheids in the xylem of softwoods. Earlywood tracheids, specialized for water transport, display a large lumen (Figure 1) and thin cell wall whereas the opposite relation is observed in latewood tracheids, where a thick cell wall is favoured to provide structural rigidity while the lumen is practically absent in many cases. Angiosperms have developed a particular cell type for water transport called vessel elements while the structural rigidity function is taken care of by other cell types such as libriform fibres. Figure 1 Softwood xylem tissue displaying latewood and earlywood tracheids (cross- section photomicrograph of spruce, bar: 100 µm).

2 As mentioned above, the first lignins appeared together with the development of the first vascular plants. Lignins are believed to be part of the product of the evolution of phenol metabolism and more specifically the phenylpropanoid pathway. This secondary metabolic pathway is thought to be important in the adaptation of plants to land life providing, among other things, protection against UV-light and pathogens [6-8]. Other phenolic compounds derived from this pathway are lignans, the phenolic portion of suberin, flavonoids, coumarins, styrylpyrones and stillbenes, some of which are present in bryophytes, which are non-vascular. Lignins have commonly been attributed not only the roles of increasing cell wall rigidity and hydrophobicity but also of improving the plant’s pathogenic defence [5,9-11]. Lignins fix cells to each other and harden the cell wall. A lignified cell wall is considered necessary for effective water conduction and the hydrophobic nature of lignins enhances cell wall impermeability. Furthermore, lignins’ chemical properties are believed to improve the cell wall’s efficiency as a barrier against microbial attack [12]. Looking at the enormous diversity of plants that produce lignins, it is interesting to notice their chemical variations as well. Since diversification can mean gain or loss of functions, to what extent does the lignin structure need to be conserved to retain the necessary properties? In other words, to what extent does the plant need to control its lignin structure and chemistry?

Figure 2 a. Wheat leaf schlerenchyma cells (cross-section TEM) showing middle lamella (ML), primary wall (PW) and secondary wall (SW). Bar: 1µm. From Brett et al. 1996 [5] with kind permission of Springer Science and Business Media. b. Model of wood cell wall displaying Middle lamella (ML), Primary wall (P) and Secondary wall (S1, S2 and S3). Adapted from Côté 1967 [13].With permission of The University of Washington Press.

The Plant Cell Wall and the Lignification Process The cell wall acts as a skeleton in plants, bringing shape and strength to the cell and rigidity to the whole plant. In woody plants, over 95% of the dry weight is attributed to the cell wall. It is a dense material, limiting the movement of large molecules, such as proteins or nucleic acids, in and out of the cell. Consequently, it also works as an effective barrier against pathogenic organisms, blocking even viruses. During cell growth, the cell wall is deposited in a series of layers (Figure 2). The first layer formed is the one between two neighbouring cells, the generally very thin middle lamella (0.1-1

3 µm). The primary cell wall then follows, varying between 0.1 and 0.3 µm in thickness. Certain specialized cells, e.g., tracheids, fibres and vessels deposit a secondary cell wall which is normally thicker than the primary cell wall (1-5 µm). For cells that do lignify their cell walls, lignification means a non- reversible process. In the xylem the completion and lignification of the secondary cell wall is followed by cell death. Accordingly, this process implies a terminal step for the cell and tissue, determining the final ultrastructure of the cell wall and its properties. The morphological distribution of lignin in cell walls of woody plant xylem has been studied with many methods but the best results have been obtained by UV and electron microscopy of woody plants [14,15]. It has generally been observed that the concentration of lignin is very high in the middle lamella and primary cell wall, ranging between 40 and 100% whereas the secondary wall displays a lignin concentration of about 20%.

The material of the cell wall is a composite of mainly three types of biopolymers: cellulose, hemicelluloses, and lignins. The cell wall is thus a highly heterogeneous material that can be divided into two phases: the microfibrillar phase composed of cellulose in a more or less homogenous phase with a high degree of crystallinity and a heterogeneous phase, the matrix, composed of different polysaccharides, proteins and phenolics (e.g., lignins) [5]. The cell wall synthesis, i.e., the deposition of the different components, generally follows an order. Thus lignins are never deposited in the absence of carbohydrates [1] and they are deposited from the outside and inwards [16,17]. Lignification follows three stages in xylem tissue. It is believed to start in the cell corner and middle lamella when the deposition of carbohydrates in the first layer (S1) of the secondary cell wall, begins. The second stage starts when the second layer (S2) of carbohydrates is deposited in the secondary cell wall, and the main lignification stage starts when cellulose fibrils are deposited in the third layer (S3) of the secondary wall (Figure 2b) [18,19]. Lignification proceeds, as described, towards the inside of the cell through the secondary cell wall, incrusting it by filling the pores of the polysaccharide matrix [20,21]. The carbohydrate contribution to the matrix varies considerably in composition within the cell wall. The middle lamella is mainly composed of pectins while the primary and secondary cell walls are made of different hemicelluloses. In addition, hemicelluloses vary between cell types and plant species. The physical and chemical conditions such as charge, hydrophobicity/hydrophilicity and pH are affected by the polysaccharides’ own properties. As a result, the local environment into which lignin is deposited varies. This raises questions about what influence, if any, the environment, and more specifically each type of carbohydrate, exerts on the deposition of lignins [22-25].

So far, the understanding of the organization of the different components in the plant cell wall and how these interact with each other is very limited. This is especially important when ascribing the general properties of the cell wall to the different components. The role(s) of each component should be regarded in the context of the cell wall as a whole [11]. Maybe the most studied type of interactions is the covalent linkages of lignin to carbohydrates, called lignin carbohydrate complexes (LCC) [26-30]. How LCCs are actually formed in the cell wall is a matter of debate, and it is further complicated by the fact that there are several types of LCCs. It is nonetheless generally accepted that LCCs exist naturally in the plant cell wall and they

4 may even cross-link the matrix polysaccharides to each other [30]. Non- covalent interactions between lignins and carbohydrates have been less investigated but they could involve, e.g., high interfacial affinity of lignin to hemicelluloses rather than cellulose [31] and to hydrophobic regions of the cell wall matrix [32]. Several tentative models describing possible organizations of the different components of the lignified cell wall have been proposed [33-37].

Lignins Structure and Chemistry

Description and Characterization Chemically, lignins can be described as aromatic polymers of methoxylated phenylpropanoid units, connected by both ether and carbon-carbon bonds. They are optically inactive [38] although they contain multiple chiral centres and are generally considered to be branched and to act as networks that cross-link cell wall components. No satisfactory exact definition is available [11,38]. Broad and simple definitions leave out important aspects of lignins and reflecting upon their obvious complexity, it is doubtful there will ever be one. Several molecular models of lignins have been proposed [39-42] giving an idea of the complexity involved (see Figure 3 for a softwood lignin model). These models are meant to consolidate the available experimental data and produce a qualitative representation of the native polymer, i.e., the unmodified polymer as it exists in situ. New models have updated previous ones as new features of lignins have been revealed. Nevertheless, no thorough 3-dimensional structural determination of any lignin has been achieved. The closest to such a determination has been achieved by a Raman spectroscopy study of spruce secondary cell wall. The study revealed how the aromatic rings in the lignin are positioned relatively parallel to each other and in the plane of the cell wall [43].

It is not possible to prepare a lignin sample that represents the whole lignin in its original state with any of the currently available methods. There are two reasons: lignin is heterogeneous within the structure itself and it is strongly associated with cell wall polysaccharides in different ways [16] making its complete isolation very difficult. Most structure determination studies have been made on isolated lignin samples, which traditionally involve such treatments that induce chemical changes [44-46]. Major advances in lignin analysis have been achieved by the development of acidolytic degradative methods, which selectively cleave certain bond-types in lignins. These methods, together with different chromatographic and mass spectroscopic tools, provide a good deal of structural information [47]. There are also so-called nondegradative methods, which basically involve common spectroscopic techniques like IR and UV. Among these, probably the most powerful method for structural characterization of lignins is solution-state nuclear magnetic resonance spectroscopy (NMR) [47-49]. However there are three generic analytical limitations despite the latest developments [11]:

Only fractions of lignin can be characterized. There is no method to completely depolymerize lignin and make it all available for analysis. Lignin isolation is plagued by contamination by other cell wall components, mostly carbohydrates. Rough total lignin content determinations are uncertain.

Figure 3 Softwood lignin model proposed by Brunow, et al 1998 [39] Reprinted with permission. Copyright (1998) American Chemical Society.

Since there is no isolation procedure available today to produce quantitatively and chemically unmodified lignin, its true molecular weight is unknown. Instead, molecular weight determinations are made on isolated lignins using a variety of methods such as Size exclusion chromatography (SEC) among others. Reports of isolated lignin molecular weights range in general from 102 to 106 g mol-1 [46]. A molecular weight of 106 g mol-1 corresponds roughly to a degree of polymerization (DP) of 5000. Molecular weight averages of approximately 20 000 g mol-1 (DP of ~100) have been

6 reported for spruce milled wood lignin [50]. For technical lignins, such as lignosulfonates, the molecular weights range from 100 to 10 000 g mol-1 [51] (DP from ~1 to ~50). Anyhow, since the isolation procedures most likely affect the size of the polymers, any molecular weight determination of such samples will represent rather low values of the actual polymers in nature.

Chemical Heterogeneity There are three monomers, called monolignols, from which most lignins are built: coniferyl alcohol, sinapyl alcohol and p-coumaryl alcohol. In the polymer, they are found as guaiacyl, syringyl and p-hydroxyphenyl residues respectively (Figure 4).

Figure 4 The common monolignols - and carbon number nomenclature - and their respective residues as found on the lignin polymer.

The evident variety of lignins between different plant species has led to the development of different classification systems. A general way, at first adopted by most researchers, was to follow plant taxonomy. According to this simple classification, gymnosperm lignin exhibits mostly guaiacyl residues, angiosperms, a mix of guaiacyl and syringyl residues and grasses, a mix of p- hydroxyphenyl, guaiacyl and syringyl residues. However this classification leaves out herbaceous angiosperm lignins, vascular cryptogam lignins (e.g., ferns) and overlooks exceptions such as compression wood of gymnosperms and the “exceptional” conifers which display lignin of a guaiacyl-syringyl type [52]. Thus a classification that is more consistent has been proposed [53], based on the lignins’ chemical characteristics rather than on the taxonomy of the plants from which the lignin comes. The basis for this classification is the predominance of each of the three lignin residues, and has been developed from two main lignin groups into 4 main groups [45]: Type-G lignins, Type G- S lignins, Type H-G-S lignins and Type H-G lignins (G, S and H stand for guaiacyl, syringyl and p-hydroxyphenyl respectively). The complexity of this classification system for lignins represents one aspect of their substantial chemical diversity.

7 Not only do lignins vary between plant species, they also vary within the same plant. Differences in monomer composition, from fibres compared to vessels, have been observed in a wide variety of hardwoods [54], and differences in lignins between cell types have also been observed in spruce [55]. Furthermore, physical and chemical differences have been noted within the cell wall, between lignins from the middle lamella as well as from the secondary wall [56,57].

Lignins also exhibit an important heterogeneity of substructures with six major inter-unit structures along with a number minor variations and additional features. The inter-unit bonds are generally very stable carbon- carbon and ether bonds (Figure 5 and Table 1). An additional variation aspect in inter-unit bond distribution is observed depending on the type of lignin, e.g., syringyl lignin displays no inter-unit bonds involving the 5-carbon. Apart from this, there are considerable variations in bond-type distribution within the plant cell wall. Lignins originating from the middle lamella of some gymnosperms contain more “condensed” lignin (i.e., lignin poor in β-O-4′ bonds) than those coming from the secondary cell walls [58,59]. A similar tendency has been noted in poplar [60]. Monomer distribution in the cell wall, which is probably associated with bond-type distribution, has been reported to follow a general pattern in both gymnosperms and angiosperms [36]. According to this, the cell corner and middle lamella contain mostly p- hydroxyphenyl and/or guaiacyl residues while the secondary cell wall contains mostly guaiacyl and/or syringyl residues. However, as far as softwoods are concerned, there is some disagreement in this matter [61].

Figure 5 The most common bonds in lignins. Some evidence exists that β-1′ and 5-5′ are actually derived from trimeric forms in lignins: spirodienone [62,63] and dibenzodioxocin [64] respectively as reviewed by Ralph et al. 2004 [65]. The most common form of β-O-4′ structures in lignins is the benzyl alcohol (R’=OH). Non-cyclic benzyl aryl ether (R′=Ar) are much less common. β-O-4′ may also display lignin carbohydrate bonds (R′=carbohydrate)

8 Table 1 Frequency of some lignin substructures in Spruce and Birch. Sources: Adler 1977 [66], Zhang et al 2006 [62] and Zhang and Gellerstedt 2007 [67]. One C9 unit corresponds to a phenylpropanoid unit in the polymer.

Intermonomeric bond- Frequency per 100 C9 unit Frequency per 100 C9 type in Spruce unit in Birch

β-O-4′ 35-45 60 β-5′ 9-12 6 β-β′ 3-4 3 β-1′/spirodienone* 2 / 1-2 1 / 2-3 5-5′/dibenzodioxocin* 9-11 / 4 ~5/ ~0 4-O-5′ ~4 ~7 *See Figure 5 for further details about these structures

Lignin Biosynthesis

Monolignol Biosynthesis The biosynthesis of lignin monolignols originates in the phenylpropanoid metabolic pathway, which is now relatively well established [8,68]. As previously mentioned, this pathway leads to a whole group of important plant phenolics but only a short description of the monolignol pathway will be presented here. The biosynthesis of the building units of lignin is thus a part of the secondary metabolism of plants, i.e., a dispensable metabolism for survival. Nonetheless, the plant spends important amounts of biochemical resources on lignification, making it a very energetically expensive process [69]. Also, substantial amounts of carbon are invested, as lignin is richer in carbon than polysaccharides [70]. All this is dedicated to a non-reversible process.

The general pathway starts with the amino acid phenylalanine. The specific pathway leading to the monolignols involves at least eight different enzymes and cofactors, biochemical energy (ATP) and reductive power (NADPH). Here, the pathway is summarized in three steps (Figure 6) - not necessarily in this order:

1. Removal of the amino group from phenylalanine and creation of a double bond on the side-chain. 2. A series of hydroxylations and methoxylations of specific aromatic carbons. 3. Reductions of the side-chain carbonyl group, from (a) carboxylic acid to aldehyde and (b) from aldehyde to alcohol.

It is worth noticing that lignin-like dehydrogenation polymers cannot only be formed with the alcohol monomers (the monolignols) but also with the different precursors like the acid or aldehyde [71,72]. However, plants apparently do “prefer” to build their lignin with the alcohol monomer although they could save energy by using any of the phenolic precursors instead.

Figure 6 Summarized phenylpropanoid pathway leading to monolignols. In terms of energy, step 2 requires one NADPH for each hydroxylation and step 3 requires one ATP and two NADPH to reduce each monomer to alcohol. Thus from phenylalanine to coniferyl alcohol one ATP and four NADPH are required, which is equivalent to a total of 13 ATP molecules [73]. For a more detailed description, see appended article IV.

“Unexpected” Plasticity of Lignin There is a considerable amount of observations describing alternative phenolics apparently being naturally incorporated to the lignin polymer (Figure 7a). Kenaf lignins contain over 50% γ-acetylated monomers [74] while grass lignins have approximately 10% p-coumaric acid [75]. The presence of γ-p-hydroxybenzoate substituted monomers in various plant lignins has been observed repeatedly [76-79]. Ferulates have been shown to be incorporated into lignin and to cross-link lignin to cell wall polysaccharides in grass [80]. In addition, other puzzling features in lignins are the reduced lignin structures in end-groups and dimeric structures (Figure 7b), which have been observed in softwood lignins [81-83].

Figure 7 a. Examples of phenolic monomers incorporated into lignins either naturally or in mutants/transgenics. b. Reduced lignin structures of undetermined origin.

Also, the study of plant transgenics and mutants has revealed even more chemical diversity of lignins. The genetic changes in all these specimens affect the monolignol biosynthetic pathway. Up- or down-regulations of specific enzymes, due to the genetic changes, lead to over- or under-

10 production of certain phenolics of this pathway. The plant’s lignin is thereby affected: the relative amounts of the different monolignols are changed [84,85] and in other cases, other phenolic species than the main monolignols are found in the lignin [86-88], resulting in remarkably different lignins deposited in the cell walls of these plants. The viability or growth is considerably affected in certain cases but interestingly, a lignin is nevertheless synthesized independently of the availability of the main three monolignols. Thus lignin possesses a rather unique ability among biopolymers of monomeric “plasticity” or flexibility, i.e., a capacity of accommodating monomers of moderately varied chemical structures.

Oxidative Polymerization Karl Freudenberg and co-workers are internationally recognized for their extensive work on lignin polymerization using model polymers made from the three common monolignols (Figure 4) [89]. The in vitro polymers formed from the oxidation of these monolignols by an oxidizing enzyme were called “dehydrogenation polymers” (DHP). The composition, the reaction products and the spectral properties of DHPs from coniferyl alcohol displayed a great resemblance with spruce lignin to the extent that they were originally described as identical [90]. Later on, however, DHPs were repeatedly shown to have noteworthy differences compared to lignin [91-94]. Although DHPs produce largely the same substructures as lignin, these are not found in the same proportions. DHPs are also difficult to produce in large molecular weights. Moreover, DHPs produced in different conditions differ as well, particularly by varying the rate of addition of the monolignols. Interestingly, Freudenberg himself noticed this difference and coined two general types of DHPs syntheses: Zulaufverfahren for one-time addition versus Zutropfverfahren for gradual addition. As a result, the validity of DHPs as lignin models has been discussed and questioned [95] but all the same they have been used and studied in connection with lignin biosynthesis because of two main reasons: DHPs produce all the major substructures of lignin and they can be produced without the typical impurities of isolated lignins, such as carbohydrates. As the differences were increasingly recognized numerous attempts have been made to make up for them by synthesizing DHPs in a great variety of conditions. These conditions represent different chemical and physical factors in the cell wall during lignin polymerization, which could potentially affect the structure of lignin. Apart from studying pH effects [96-98], the monolignol rate of addition [99,100], different oxidation agents [101-103] and other environmental conditions have also been tested. DHP syntheses in the presence of carbohydrates [23,104,105], trying to mimic the cell wall environment also proved to affect the structure of the polymer in different ways. Although no DHP has shown identical properties with isolated lignin, it has become evident that a DHP’s structural outcome is sensitive to its polymerization conditions [106] and thus its properties can be indirectly controlled. Which cell wall conditions actually influence lignin polymerization and to what degree they do so, is poorly understood.

From a theoretical point of view, a model for lignin polymerization had already been proposed by Erdtman in the 1930’s [107] in the light of a dehydrogenation experiment performed as early as 1908 by two French chemists [108]. This model described lignin as the product of the dehydrogenation of propenyl phenol derivatives. Freudenberg and co-workers

11 provided solid evidence for this model with their extensive experimental work on dehydrogenation models. The great value of Freudenberg’s work lies in the structural elucidation of more than forty products of DHP formation, from monomers to hexamers [89]. It became clear that the dehydrogenative polymerization process follows certain rules and the knowledge of these rules provided a basis for explaining the majority of the structures found in lignin. Today, this theoretical model is essentially the same and it can be summarized as follows:

Lignin polymerization is the result of radical coupling of resonance-stabilized monolignol radicals and phenolic radicals on the lignin polymer, followed by nucleophilic additions to the quinone methide intermediates (Figure 8).

Figure 8 β-O-4′ coupling of monolignol, coniferyl alcohol (CA), to guaiacyl polymer (PG) end-group. The final step constitutes a nucleophilic addition to the α-carbon of the quinone methide intermediate.

This model was supplemented by Sarkanen [95] with the idea of Bulk polymerization vs. End-wise polymerization. A bulk polymer is the result of predominant dimerization and subsequent coupling of dimer or oligomer radicals to each other, whereas an end-wise polymer is the product of monomer radicals’ coupling to a growing polymer chain. A bulk polymer will have more of a branched character whereas the end-wise polymer is more linear. According to Sarkanen [95], this then accounts for the differences observed in DHPs structures depending on the monolignol rate of addition as in Zulaufverfahren and Zutropfverfahren. The formation of certain bonds is particularly favoured depending on the type of polymerization. As a result, an end-wise polymer displays large amounts of β-O-4′ bonds while a bulk polymer contains large amounts of β-5′ and β-β′. The bulk polymer will also display a greater amount of α―β unsaturated side chains and phenolic end- groups. Since natural lignins contain β-O-4′ structures mostly, they are considered to be the product of end-wise polymerization [65,66,95].

Challenges

Polymerization Control The model offered by Erdtman and later supported by Freudenberg and others implies no strict control over the coupling step which ultimately defines much of the lignin structure. Thus according to this model, lignin is not expected to display any particular sequence of monolignols and of the bond-types created at each coupling event. As a result, lignin polymerization has subsequently been described as a random or near-random coupling of

12 monolignols [9,109], giving the impression that any monolignol might couple in any of the possible ways. Pure statistical randomness is not chemically possible in phenolic radical coupling [65] and Erdtman already supposed there would be a certain degree of order in lignins [110]. By looking at monolignol and bond distributions in lignins of different plant-types (Table 1), one would expect some kind of influence or “control” over the polymerization. The inability of DHPs to completely reproduce lignins in vitro, especially the large amounts of β-O-4′, makes clear the lack of understanding of the regulating mechanisms at work in the plant cell. Several suggestions have been made to explain how different factors might influence bond and monomeric distribution, representing possible ways of indirect regulation in the plant cell wall:

The rate of monolignol addition to the reaction site (alternatively the rate of radical generation) in the cell wall is very slow and very difficult if at all possible to reproduce in vitro. It has been shown that different addition rates will result in substantial differences in bond distributions [66,95,100,111]. Precursor/monolignol availability will define the monomer composition of lignin. The most obvious examples are mutants and transgenics which apparently incorporate available phenolic compounds as lignin monomers when lacking the common monolignols. The different monolignol availability will also influence the bonding pattern since depending on the monolignol structure some bonds are generally not formed (e.g., no 5-carbon bonds in syringyl lignin). The reaction environment, the heterogeneous polysaccharide matrix of the cell wall plays an important role in lignin polymerization and will ultimately influence bond formation [23,105,112,113]. As mentioned previously local pH and hydrophobicity vary within the cell wall. Purely chemical factors determine the outcome of the different coupling reactions in lignin polymerization: the oxidation potentials of the different phenolic species and their radical reactivities [110,114]. If lignin is mostly the product of the addition of monolignols to a growing lignin polymer, then cross-coupling (i.e., a monolignol radical coupling to a phenolic end-group radical) must be a main coupling event in lignin biopolymerization [65]. The regioselectivity of such reaction is not well understood but some experimental evidence suggests that the chemical effect, depending on the chemistry of the different phenolic species, over bond- formation can be quite strong. An interesting work by Syrjänen and Brunow showed how β-O-4′ bonds are strongly favoured when cross- coupling coniferyl alcohol to a phenolic end-group model [111].

An alternative to the purely chemically “controlled” model has been proposed by several researchers [115-117]. This model suggests that the polymerization step in lignin biosynthesis is actually under strict control, as is the general case for other biopolymers. Consequently this control dictates both the sequence of the monomers and the bond-types formed. Such in vivo control system could then explain why in vitro syntheses of lignin models such as DHPs, have not been able to reproduce lignin correctly, particularly the amount of β-O-4′ structures. According to this model a primary lignooligomer is formed into arrays of dirigent sites in the cell wall, which completely control the radical-radical coupling of monolignol radicals. The

13 nature of the dirigent sites has been proposed to be cell wall proteins, such as hydroxyproline-rich gluco-proteins, which are present in the plant cell walls. Thereafter, the first lignin chain, product of the arrays of dirigent sites, works itself as a polymeric template which is replicated into a new lignin chain, copy of the first. The second chain will then serve as a template for a third and so on. Thus lignins are suggested to possess similar biochemical properties to enzymes’ catalytic capabilities. In addition, the replication mechanism of this template model produces a mirror image stereoisomerically speaking. This mirror effect is suggested to explain the observed optical inactivity of lignins. The proposal of this new model implying strict control, is controversial since it stands in contrast to the purely chemically controlled model and has generated much debate [38,65,115,117-122]. Although this new model is in need of more experimental evidence to gain wider acceptance [11,119], it has found its way into a text-book and is presented as the model for lignin polymerization awaiting “[…]full clarification at the biochemical level.” [123]

Another puzzle in lignification is the polymerization initiation sites. It is believed that in the xylem, lignification starts at the cell corners and proceeds towards the centre of the cell as previously described. Remarkably, in other plant tissues, there are reports of just the opposite happening, i.e., lignin deposition starting in the innermost cellulosic layer [124-126]. How does the common xylem cell prevent polymerization from happening in the inner part of cell wall as monolignols are transported into the cell wall’s outer layers? Is lignification specifically “directed” to start at specific sites/molecular structures in the cell wall matrix? Several suggestions in the literature attempt to answer these questions. Enzymes/proteins as well as cell wall polysaccharides are possible candidates as initiation sites [17]. Cross-linking in the cell wall by ferulates has led some to suggest them as starting points for lignification [127,128]. Ferulate dimers have been shown to cross-link polysaccharides by ester bonds and so a polysaccharide ester ferulate group in its phenolic state could be a starting point for polymerization. Such groups have been shown to couple monolignols through dehydrogenative coupling in vitro and in vivo [129,130]. The strict control model of lignin polymerization mentioned above proposes hydroxyproline-rich proteins as initiation sites. Interactions between lignin and these cell wall proteins and others (proline- and glycine-rich proteins) are believed to exist but neither the nature of the linkages nor the proteins’ functions have been established [11].

Dehydrogenation The oxidation step, creating a phenoxy radical on either a phenolic end- group of the polymer or on a monomer, has been a key subject of many studies. Generally, it is believed that hydrogen peroxide (H2O2) is the main oxidizing agent along with cell wall peroxidases (class III) as catalysts [131,132]. It is difficult to assign specific functions to different isozymes because of peroxidases’ wide substrate specificity but there are indications of monolignol-specific peroxidases [133-135]. Moreover, there is significant evidence for the involvement of other oxidases, such as laccase in lignification [136-138].

For the polymer to grow, the oxidation of phenolic end-groups on the polymer must be at least as effective as the oxidation of monomers. To

14 imagine cell wall enzymes being involved in the oxidation of diffusing monomers in the cell wall matrix might not be too difficult, but to picture them oxidizing the phenolic residues on the polymer as effectively as the monomers raises several questions. The lignifying cell wall is increasingly hydrophobic, turning it into a hostile environment for most enzymes. The oxidation of phenolic residues on the lignin polymer, which is expected to be more or less immobile as it is attached to a solid phase, will be dependent on the diffusion rate of the enzyme in an already crowded cell wall matrix. Additionally, enzymatic oxidation of a phenolic residue on the polymer will need to overcome a higher oxidation potential, compared to a monolignol [114], and steric hindrance by the polymer itself. Recently, however, a cell wall peroxidase was discovered in poplar callus, which effectively oxidizes polymeric lignin and plays a significant role in secondary cell wall lignification [139,140].

Other possibilities for the oxidation step have been envisaged, possibly solving these challenges. Phenolic radicals might act as oxidative carriers or shuttles that, once oxidized by an enzyme, can diffuse more freely than enzymes onto the growing polymer and oxidize phenolic residues [98,122,141]. Low molecular weight oxidants such as superoxide radical ●¯ 3+ anion, O2 , [142] and manganese, Mn [103], have been proposed to carry out the same function.

Lignin among Other Polycyclic Biopolymers There are other phenolic biopolymers that, although still poorly characterized, exhibit noteworthy similarities with lignins, i.e., suberin, polymeric tannins and melanin. In the literature, melanin has been compared to lignin but only superficially [143-145]. A short description of melanin function and biosynthesis is presented here to account for a number of similarities which might be of interest in elucidating these complex biopolymers.

Melanin(s) is a group of polycyclic biopolymers that are produced by a wide variety of organisms: bacteria, fungi, animals, protozoans and plants. They can be described as “generally black biological macromolecules composed of various types of phenolic or indolic monomers…” [146]. Interestingly, as with lignin, most structure determinations have been done on harshly isolated samples and so the structure of intact melanin is little understood [146]. In comparison, lignin research and structural characterization has come a long way. There are essentially three types of melanin [147]: eumelanin (nitrogen containing melanin derived from tyrosine and phenylalanine), phaeomelanin (like eumelanin but generally also containing sulphur), allomelanin (nitrogen-free melanin derived from naphthalene derivatives and catechol among others). A rough generalization would classify eumelanin and phaeomelanin as animal melanin and allomelanin as plant, fungi and bacterial melanin [144].

General properties usually attributed to melanin are: absorbance of a wide spectrum of light (UV and visible light); to participate in one- and two- electron redox reactions and to display cation chelating properties [148]. More specifically there are several interesting benefits which melanins convey

15 to the organisms that synthesize them. Under anaerobic conditions bacteria may use them as electron acceptors or carriers, making possible energy producing processes, similar to oxidative phosphorylation [147]. Melanin protects some fungi against UV-light [149]. They provide a virulence factor for certain pathogenic bacteria and fungi by acting as radical scavengers and thus defending them against their host’s own defence mechanisms [150,151]. Also in fungi, production of melanin is associated with cell wall modification, thus providing structural strength and protection it against extreme environments [146]. In insect cuticle formation, melanin is believed to provide mechanical strength [152] and act as a structural component in wound healing in arthropods and plants [153,154]. Apart from this, melanin in animals are usually pigments that provide camouflage and mimicry properties or ornamentation for sexual display [143]. In humans, melanin has been reported to offer protection to skin cells from damage caused by UV-light [155].

Melanin biosynthesis bears some interesting similarities with lignin biosynthesis. As with lignin, melanin synthesis is part of the secondary metabolism of the organism and thus non-essential for survival. Also, the final polymer is a product of oxidative polymerization of phenolic (and indolic) compounds or precursors [146,148]. In Figure 9, some of the known melanin precursors are presented to be compared with lignin monolignols and alternative phenolic monomers in Figures 4 and 7. Animal melanin precursors are mainly 3,4-dihrydroxy-phenylalanine (DOPA) and cysteinyl DOPA while fungi melanin precursors are diverse but mostly 1,8-dihydroxynaphtalene (DHN). Other known precursors are homogentisic acid in a few bacteria [156] and Glutaminyl-4-dihydroxybenzene (GDHB) in certain fungi [157].

In fungi, the polymerization process is believed to be achieved by a phenoloxidase. Tyrosinase, peroxidase and laccase are considered possible candidates [146,158]. Very little is known about the coupling step and about what the DHN polymerization products are or what types of bonds are formed. In the case of animal melanin it is known that DOPA polymerizes to melanin through oxidation by tyrosinase. The oxidation leads to formation of quinones which react with diphenolic precursors to form oligomers. In vitro experiments have shown that in the presence of tyrosinase and a pH of at least 6.8 DOPA forms melanin and at pH 8 or above DOPA polymerizes without the oxidase [159]. An intriguing fact is that melanosomes, the cell organelles where animal melanin is synthesized, have an acidic pH. A low pH is an unfavourable condition for polymerization, which requires deprotonation. It has thus been suggested that this is a way for the cell to control melanin polymerization [160]. An important difference between animal and fungi melanin is that in animals melanin is synthesized and kept inside the cell (in melanosomes) while fungi melanin occurs in the cell walls or extracellularly [146,161].

In insect cuticle, diphenolic precursors such as DOPA have been isolated and generally believed to build melanin contributing to the sclerotization process, which involves the darkening and hardening of cuticles [148,152,162]. However, only the diphenolic monomers N-acetyl-dopamine (NADA) and N-beta-alanyldopamine (NBAD) (Figure 8) are believed to play an important role, at least in the hardening process [163]. Although it is not known to which extent any of these present diphenolic precursors actually

16 polymerize in the cuticle, there is evidence for an uncharacterized polyphenolic polymeric fraction [164,165]. As for NADA and NBAD there is evidence supporting their participation in cross-linking and polymerizing proteins of the cuticle [163]. Through oxidation by phenoloxidases these precursors form quinones and quinone methides which react with nucleophilic groups in the cuticle from e.g., proteins [145,163]. Interestingly, dimers of these precursors (Figure 9) have been found in insect cuticles [166].

Figure 9 Melanin precursors from different organisms and proposed coupling mechanisms. a. Animal melanin precursor (DOPA: 3,4-dihrydroxy- phenylalanine) oxidation and polymerization. b. Fungi melanin precursors (GDBH:Glutaminyl-4-dihydroxybenzene and DHN:1,8-dihydroxynaphtalene) c. Bacterial melanin precursor (homogentisic acid) d. Sclerotization agent in insects (NADA: N-acetyl-dopamine)

Maybe the most relevant and consistent difference between lignin and melanin precursors is the diphenolic nature of melanin precursors. The polymerization process is also chemically different: the oxidation of melanin precursors is believed to lead to quinones, not stabilized radicals as in lignin, which then polymerize. Despite these differences, from a biosynthetic point of view, lignin and melanin monomers display the same chemical potential.

17 Upon activation by an oxidative agent they are able to spontaneously polymerize without any catalytic assistance, as opposed to other biopolymers. The monomers of nucleic acids, proteins, polysaccharides among others, do not polymerize by themselves after activation; instead they all depend on a carefully orchestrated enzymatic process for the coupling step [167]. Although it has not been determined whether melanins are polymerized in nature under a strict control system, the possibility exists that they are not. At least, in the case of lignin biopolymerization there is substantial evidence supporting the idea that lignin is deposited in the plant cell wall without such control. This does not mean, of course, that there is no careful control over the polymerization step, but as an alternative, the control mechanisms might be indirect.

In summary, the similarities between lignin and melanin include:

A role in hardening of extra-cellular layers or cell wall Cross-linking extracellular components A role in defence against pathogenic attacks or wound healing Being the product of polymerization of phenolic compounds through oxidative coupling Being recalcitrant due to the stability of the inter-unit bonds The coupling/polymerization step is probably purely chemically controlled.

It seems that in nature, common functions such as mechanical strength, cross-linking polymers and pathogenic protection are, in a wide variety of organisms, achieved by synthesizing polycyclic biopolymers such as lignins and melanins through oxidative polymerization of phenolic/diphenolic monomers.

18 Methods

Detailed and individual descriptions of materials and methods can be found in each paper appended to this thesis. A general description of the methods used in this work is presented here.

Coniferyl Alcohol Synthesis Coniferyl alcohol (Figure 4) was repeatedly synthesized for the majority of DHP experiments and was consistently used throughout all the studies. It was obtained from the reduction of commercially available coniferyl aldehyde with sodium borohydride, as described elsewhere [168].

Dehydrogenation Polymers – DHP The in vitro synthesis of dehydrogenation polymers in different conditions and the analyses of them using common organic chemistry analytic methods is the principal basis of all the experimental work presented in this thesis. Karl Freudenberg and co-workers, who originally introduced model dehydrogenation experiments and with them elucidated the formation and structure of many lignin substructures, already stated two general ways of synthesizing the polymers that would affect their final properties: Zulaufverfahren and Zutropfverfahren: a one-time addition versus a drop-wise (or gradual) addition of monomers, respectively. Although, no standardized method exists for them, these methods (or principles for synthesis) have been used by many groups and in each case adapted or modified depending on the desired effect: the one-time addition giving more bulk, branched polymers and the drop-wise giving more linear polymers. Since lignin is recognized as a generally linear polymer and one objective of this work was to synthesize polymers as similar to lignin as possible, the Zutropfverfahren or drop-wise principle was chosen in all DHP experiments. It is certain that DHPs, as lignin models, do not fully represent lignins and their polymerization. Nevertheless, the fact that they can form most of the bonds that exist in natural lignins and that the contamination problems of lignin isolation are avoided means they are useful experimental modelling systems to elucidate the formation and structure of lignins.

The general procedure for DHP synthesis can be described as follows. Phenolic monomers (normally coniferyl alcohol) were dissolved in an organic solvent, miscible with water, and diluted in a larger volume of aqueous buffered solution, generally 3-50 mmol/l H2KPO4 with regulated pH, generally near-neutral or slightly acidic in some specific cases. A reaction mixture was prepared with an oxidant, generally hydrogen peroxide and a catalyst, horseradish peroxidase (HRP), which were dissolved separately from the monomers in a buffered solution of the same type. The monomers were dropped into the reaction slowly, using a peristaltic pump at flow-rates ranging between 6.5 and 2.5 ml/h depending on the experiment. After 24

19 alternatively 48h of reaction at room temperature and protected from light, the product was precipitated (if needed) by evaporation of the organic solvent initially used. The precipitate was washed with pure water by centrifugation. This step was taken to get rid of enzymes, salts and unreacted hydrogen peroxide. The product was then suspended in water and freeze-dried. The dried product, generally a powder was weighed, and in certain cases, washed in an appropriate organic solvent, e.g., ethyl acetate, to get rid of low- molecular weight products, mostly dimers but also unreacted monomers.

Derivatization Both the low-molecular weight and the polymeric fractions were derivatized for the different chromatographic methods used.

Acetylation To make DHP polymers more soluble in THF, which was the solvent in the SEC system (described below), acetylation was required. One to two mg of polymeric DHP were dissolved in 1-2 ml pyridine:acetic anhydride (1:1) and left to react overnight. The reaction was stopped with an excess of methanol, alternatively water:methanol (1:1) and pyridine co-evaporated with toluene to dryness.

Silylation For analysis with GC-FID and GC-MS, sylilation of low molecular weight compounds such as dimers and monomers was carried out under nitrogen for at least 1h. The product was dissolved in methylene chloride, alternatively acetone, ideally at 1mg/ml concentration and N, O-Bis(trimethylsilyl)acetamide (BSTFA) was added together with pyridine (10%) as catalyst.

Gas Chromatography - Flame Ionization Detector, GC-FID GC-FID was used to check purity of different synthetic models and to quantify dimer amounts using tetracosane as internal standard and calibrated with authentic models when available. The GC-FID instrument was a Hewlett-Packard 6890 with a RTX 5 column (30m, 0.25 µm I.D., 0.25 µm film thickness).

Gas Chromatography - Mass Spectroscopy, GC-MS GC-MS was used to confirm chemical structures of synthetic models and identify monomers and/or dimers in product mixes such as low molecular weight fractions from DHPs. The instrument was from Thermo Finnigan Trace GC-MS, 2000 series. The column used was RTX 5MS (30 m, 0.32 mm, 0.25 µm).

Size Exclusion Chromatography – SEC Samples of acetylated high molecular weight DHP products were analyzed by SEC to study their hydrodynamic volume and obtain an idea of the relative

20 molecular weight compared to elution times of monodisperse polystyrene standards. The analyses were performed on a system of three Cross-linked PS columns in series (Steerage, Waters HR4+HR2+HR0.5) using THF as effluent at 0.8 mL min-1. Detection was made with UV-light detector at 280 nm.

Nuclear Magnetic Resonance - NMR Samples of underivatized, usually only high molecular weight DHP fractions were dissolved in deuterated solvents for NMR analyses. Both 1-D (qualitative and quantitative 13C) and 2-D (HSQC and HSQC-TOCSY) NMR experiments were run to detect and quantify specific DHP substructures. All NMR experiments were run on a Bruker Avance 400 MHz instrument.

21 Results and Discussion

Reduction during Oxidative Coupling (Paper I) In native lignin there are some types of structures that cannot be explained solely by oxidative coupling. Although the Erdtman/Freudenberg model explains the large majority of lignin structures, a number of structures that cannot be explained by this chemistry alone have long been noted in lignins, including the reduced lignin structures [169,170]. The most common structure is dihydroconiferyl alcohol (Figure 10), found as an end-group and suggested to be incorporated as such by dehydrogenative coupling since the plant cell may synthesize it and subsequently use it as a monomer [87]. However, there is a more puzzling structure, reduced β-β′s (Figure 10), found in softwood lignins [38,65,170]. Since this is a dimeric reduced structure it is even more intriguing to consider its incorporation as such into the polymer, thus maybe originating from a specific lignan: secoisolaresinol. Another possibility is that this structure is formed subsequent to polymerization, e.g., by the reduction of a resinol structure, in the same way the lignan is formed from pinoresinol [171]. A third possibility is that it is formed during the dehydrogenative polymerization process, e.g., by the reduction of a quinone methide intermediate. Any of these possibilities convey immediate problems. Lignans are optically active, meaning that if the lignan secoisolaresinol has been incorporated then only one stereoisomer would be found in lignin, which is not the case [38]. A post-lignification reduction seems even less likely because a lignified cell wall is very compact and rules out the only probable candidates able to achieve such a reduction: enzymes [172]. Although an enzyme that generates the lignan secoisolariciresinol could be trapped in the cell wall during lignification the product would still be optically active. Therefore, the third possibility was considered the most likely. We reasoned there must exist a reductive agent able to achieve a reduction in the overall oxidative environment of lignin polymerization. This was the objective of our investigation: is there a naturally occurring reductive agent able to carry out the reduction of a quinone methide intermediate during dehydrogenative polymerization of monolignols?

Quinone Methide Model Reduction Four low molecular mass biological reducing agents were screened for their capability to reduce a model compound (Figure 11a) for the β-ether quinone methide intermediate in lignin biopolymerization. The reducing agents were nicotinamide adenine dinucleotide (NADH), reduced glutathione, both being common reducing agents produced in virtually all living cells [167], ascorbic acid known as a redox buffer agent in plant cell walls [173] and the citric acid intermediate α-ketoglutaric acid. The extracellular enzyme cellobiose dehydrogenase (CDH) was included in the screen as an example of a reducing enzyme [174]. Sodium borohydride was used as a positive control reducing agent. The β-ether quinone methide model was only successfully reduced by NADH or NaBH4. Gas chromatograms of the sodium borohydride- treated sample and the NADH-treated sample displayed an equivalent peak with identical MS spectra, corresponding to the silylated reduced model

22 compound (Figure 11b). Three NMR experiments (1H, 13C-DEPT-135 and 2D HSQC), run on the sodium borohydride-treated sample, confirmed the identity of the product.

Figure 10 Reduced lignin structures present in softwood lignin [170].

Reduced Structures in DHPs NADH was tested in synthetic dehydrogenation polymers (DHPs). The interest was to determine how well oxidation and reduction could proceed in the same reaction medium. The 2D NMR technique HSQC was employed to reveal the DHP structures formed. Figure 12 shows the presence of β-aryl ether A, phenylcoumaran B, resinol C, and α,β-diaryl ether A2 structures which are common in conventional DHPs [49,66]. Furthermore, reduced β- ether structures AR (Figure 12) of the same type as compound b (Figure 11) were observed from DHPs made in the presence of NADH. Disappointingly secoisolariciresinol structures CR (Figure 12) were not observed, nor were dihydroconiferyl alcohol X5 moieties found.

Figure 11 Reduction of a quinone methide intermediate from a β-O-4′ model compound. First step achieved according to Brunow, G. et al. [175]. See appended article I for details.

From these results it can be argued that the origin of reduced structures like secoisolariciresinol CR is an uncatalyzed, protein-independent reaction of a biologically significant reductant such as NADH with the quinone methide intermediate, formed during lignin polymerization. It is not known whether this is the actual compound responsible for such a reduction and the inability of CDH to reduce the quinone methide here does not imply that there are no other enzymes able to catalyze the reaction. The substantial amount of reduced β-aryl ether units AR detected (~48% of the β-ethers) indicates that

23 NADH was able to reduce the β-aryl ether quinone methide in an oxidative polymerization system. NADH was able to reduce a β-O-4′ quinone methide structure but not the β–β′ quinone methide; at least not in detectable amounts. The conditions and/or reductants needed for such a reduction remain uncertain. It would seem the reduction of a β-aryl ether (such as β-O- 4′) quinone methide competes with a nucleophilic attack by an external nucleophile (water or a phenol), whereas the β–β′- or β–5′-quinone methides have intramolecular hydroxyl groups out-competing the attack on the α- carbon. Since we observed resinol (β-β′) and phenylcoumaran (β-5′) structures in the DHPs, NADH was apparently not effective in competing against the fast intramolecular post-coupling reactions under these conditions.

Until these results were published, reduced structures had only been observed in end-groups X5 and in β-β′ CR units in natural lignins [87,169,170,176]. Recently, however, reduced β-O-4′ structures (AR) have been observed in lignin [177], analyzed by NMR spectroscopy with a special cryo-probe that increases the analysis sensibility. While it is exciting to confirm that these structures do occur naturally and are not only a laboratory product, it is also important to note that they apparently occur in lesser amounts than the reduced β-β′s. The question that immediately arises is: what process favours the formation of the structure which is seemingly more difficult to produce?

In this work we did not detect any reduced β-β′ structures CR, but succeeded in producing reduced β-O-4′ structures AR. Although it cannot be confirmed, this could be due to the low sensitivity of the NMR analyses but more probably to the difference in the structure and the polymerization environment between lignin and DHPs. It is a known fact that DHPs are sensitive to polymerization conditions [106]. If the latter explanation is the case, then there must be some kind of mechanism directing the reducing effect towards β-β′-units in lignin polymerization. There are of course other options: enzymatic catalysis, low molecular weight catalysts or unusual conditions during the reduction. Or, are β-β′ quinomethides more long-lived in the natural conditions of the biopolymerization of lignin?

Whatever the reduction agent consumed in such a reaction, it appears to be a waste of energy if reduced structures are just the products of side- reactions. Spending one molecule of NADH corresponds to 3 of ATP [73]. What could be the benefits of reducing the lignin? Such structures could be expected to be more hydrophobic than their “normal” counterparts. Also, they offer fewer possibilities for hydrogen bonding and for the formation of covalent bonds to polysaccharides. This could potentially mean less LCC formation through ester and ether bonds to the α-carbon in lignin monomers, and this might in turn have an effect on the properties of the cell wall. Furthermore, a higher rotational freedom and thus flexibility should be expected of a reduced β-β′ compared to the cyclic resinol structure in the normal β-β′. Although reduced lignin structures might involve a biochemical cost they could be a way for the plant to control the properties of its lignin.

Figure 12 2-D NMR spectra (HSQC, correlating a carbon nuclei to its bonded hydrogen’s nuclei) of the non-aromatic region of DHPs synthesized in the presence of NADH. Signals for X5 and CR are only the expected signals of these structures according to Zhang et al. [170]

Effects of an Anti-oxidant during Lignin Polymerization (Paper II) In our experiments with reduced lignin structures (Paper I), ascorbic acid could not reduce the quinone methide model and was discarded as a reducing agent able to generate the observed lignin reduced structures. Nevertheless, the presence of ascorbic acid in the cell wall has been reported [178,179] and it is suggested to play a role in scavenging hydrogen peroxide in the cell wall as it does in vacuoles [180,181]. Ascorbic acid is proposed to reduce hydrogen peroxide indirectly by reducing phenolic radicals which in turn have been generated by cell wall peroxidases and hydrogen peroxide. If there are reducing agents present during lignin polymerization, what other effects might they have upon the structure of lignin? The phenolic monomer radical concentration is an important factor affecting the structural outcome of the lignin polymer [95,100,111]. As a result, the presence of an anti-oxidant, such as ascorbic acid, during lignin polymerization should influence it. In the present work, we investigated the effect of the anti-oxidant ascorbic acid during the synthesis of coniferyl alcohol dehydrogenation polymers (DHPs).

DHPs in the Presence of Ascorbic Acid Dehydrogenation polymers were synthesized in the presence and in the absence of ascorbic acid at pH 5 by enzymatic oxidation of coniferyl alcohol using horseradish peroxidase and H2O2. The amounts of three common lignin inter-unit bond types (β-5′; β-β′ and β-O-4′) were quantified by 13C-NMR of the polymeric fraction and compared to a control synthesis, i.e., an analogous DHP synthesis without ascorbic acid. A portion of the spectra and signal assignments, according to Landucci [92], are presented in Figure 13 and Table 2. The quantification was done as follows. The amount of β-5′ structures was calculated by taking the mean of the integration values of the two isolated signals for the α- and γ-carbons of this substructure (Figure 3, signals 1 and 4 respectively). Then, this value was subtracted from the integrated value of the overlapping signals of the β-carbons of β-5′ and β-β′ (signal 8) in order to determine the amount of β-β′ structures. The value for β-β′ structures was subtracted from the signal 2 - i.e., the overlapping signals from β-carbons from the benzyl alcohol β-O-4′ and non-cyclic benzyl aryl ether β-O-4′ (α-O-4′) (see structures in Figure 5) and α-carbons from β-β′ - to obtain the amount of all β-O-4′ structures. The amount of the benzyl alcohol form of β-O-4′ alone was obtained from the integrated value of the overlapping α-carbon and γ- carbon signals from β-O-4′ and β-β′ respectively (signal 3), also by subtracting the β-β′ value obtained earlier. The amount of total β-O-4′ structures was generally no more than 3% higher than the benzyl alcohol β-O-4′ form alone in each case. Thus the contribution to the total amount of β-O-4′ by the non- cyclic benzyl aryl ether form was low, which is consistent with the observations that low pH favours the formation of the benzyl alcohol form of β-O-4′ [182]. Signal 6 (Figure 4) was also assessed to determine the total amount β-O-4′ structures but some contamination is suspected to have occurred since 20 to 50% higher quantification values were obtained from this signal compared to the β-O-4′ values obtained from signal 2 and 3.

Figure 13 13C-NMR partial spectra of the high molecular weight fraction of DHPs produced in the presence of 50% ascorbic acid (relative amount) – 1,3,5-trimethoxy benzene. Signals assignments in Table 2.

Table 2 Signal assignments from 13C NMR spectra in Figure 13. β-O-4′ represents the benzyl alcohol form and β-O-4′(α-O-4′) is the non-cyclic benzyl aryl ether form (see structures in Figure 5). Signal δ Assignment 1 87 α-carbons from β-5′ 2 83-85 β-carbons from β-O-4′, β-O-4′(α-O-4′) and α-carbons from β-β′ 3 71-72 α-carbon from β-O-4′ and γ-carbons from β-β′ 4 63 γ-carbons from β-5′ 5 62 γ-carbon from coniferyl alcohol end-group 6 60 γ-carbons from β-O-4′ and β-O-4′(α-O-4′) 7 56 Methoxyl carbons from DHPs 8 53-54 β-carbons from β-5′ and β-β′ IS-a 93 Unsubstituted aromatic carbon from internal standard IS-b 55 Methoxyl carbon from internal standard

Ascorbic acid clearly increased the formation of β-O-4′ structures and decreased the formation of β-β′, at a 25% amount, relative to the coniferyl alcohol, compared to the control experiment (Figure 14). Additional DHPs were produced with larger amounts of ascorbic acid (50 and 75%) to study any potential concentration effects. The effect of ascorbic acid was enhanced with its concentration up to 50% for β-O-4′ and slightly enhanced for β-β′ structures (Figure 14). However β-5′ structures were practically not affected. Since ascorbic acid was added into the reaction mix at the same rate as the other reactants it was expected to continuously slow the reaction down by reducing the radical concentration. Consequently, the larger the amount of ascorbic acid, the slower the polymerization would proceed, enhancing an end-wise polymerization [95]. The observed increase in β-O-4′ structures is in agreement with other reports where the reaction was slowed down by limiting the monomer diffusion rate alone [100,111]. Limiting the monomer concentration limits the monomer radical concentration which in turn limits

27 dimerization. Dimerization is a known problem in DHP syntheses because of generally high monomer concentrations [65]. As previously described, the regioselectivity in dimerization reactions is different compared to cross- coupling and for end-wise polymerization to occur, monomer-oligomer cross- couplings are necessary. The negative effect that ascorbic acid had on the formation of β-β′ structures might be due to a favoured end-wise polymerization assuming β-β′ structures are mainly a product of dimerization [183]. Furthermore, the anti-oxidant had apparently no effect on the formation of β-5′. This could be due to the fact that these structures can be both formed through dimerization and end-wise coupling. Interestingly, at 75% relative ascorbic acid concentration, the effect was reversed for β-O-4′ and β-β′ structures and the formation of β-5′ structures was affected negatively. In addition, the total DHP amount was monitored as well (Figure 14) since ascorbic acid was expected to consume H2O2, the reaction yield would consequently be affected. Accordingly, the yield started to decrease at 50 % ascorbic acid and was seriously affected by 75%. The effects observed at 75% ascorbic acid might suggest that the polymerization was continuously quenched until ascorbic acid was depleted. Then, the polymerization would be allowed to take place, producing DHPs with a different substructure composition because of the accumulation of coniferyl alcohol.

Figure 14 Substructure quantification by 13C NMR of DHPs synthesized at pH 5 in the presence of different amounts of ascorbic acid. The control experiment is represented as 0% ascorbic acid. Total amounts of β-O-4′ are presented here.

Although it is likely that ascorbic acid primarily slows down the polymerization rate by reducing phenolic radicals it is not possible to rule out any effects of the anti-oxidant with the enzyme, either as a substrate or as an inhibitor, generating a similar structural outcome. However, horseradish peroxidase apparently prefers phenolics as substrates rather than ascorbic acid [184,185]. Neither is it possible to rule out any direct effect of ascorbic acid upon the regioselectivity of the coupling events themselves. However, if such a “catalytic” effect were the case, one would not expect an inverted effect after a certain concentration, as observed here.

The molecular weight determinations by size exclusion chromatography showed that the DHPs displayed similar sizes independently of the presence of ascorbic acid and up to a 50% (Figure 15). The results imply that similar- sized polymers were produced with differences in inter-unit substructure amounts. At 75% ascorbic acid, the sizes of the DHPs appeared to have been somewhat affected (Figure 15). Based on the NMR inter-unit substructure quantification results, it is likely that the polymerization occurred under different conditions, as previously described, thus affecting the sizes of the polymers.

All DHPs synthesized displayed a common maximum size (Figure 15), as if the polymerization conditions did not “allow” them to grow larger. Generating large molecular weight DHPs by conventional ways is known to be a problem [112]. The conditions here used for the polymerization involved a mixture of dioxane:water (2:3) which allowed the reaction to stay in solution all along, compared to purely aqueous systems where DHPs tend to precipitate very early in the reaction. This system was chosen to make all polymeric species available for oxidation and coupling as much as possible. However, the solubility of this complex mixture of polymers is probably not complete after all. It is likely that in our system, solubility is still an issue which defines the maximum size of the polymers. Improving the solubility of DHPs is believed to affect their molecular weight positively [104].

Figure 15 Size exclusion chromatograph of DHPs synthesized at pH 5 in different ascorbic acid concentrations (0, 25, 50 and 75% ascorbic acid relative to coniferyl alcohol).

The most common inter-unit substructure in lignins is the β-O-4′. These results provide further support to the idea that a low monomer radical concentration - in this case provided by ascorbic acid - enhances the formation of β-O-4′ bonds by favouring end-wise over bulk polymers. Since ascorbic acid has been reported to be present in the plant cell wall, it seems plausible that this component or similar ones may play a role in the biopolymerization of lignin, as an additional component besides monolignols, peroxidases and hydrogen peroxide. The presence and concentration of a phenolic anti-oxidant may also be a way for the plant to control and regulate the lignin structure, possibly explaining the differences in lignin structure

29 between, e.g., cell wall layers, in addition to pH, polysaccharide composition and monolignol addition rate.

Table 3 Molecular weight values (in g mol-1 relative to polystyrene standards) of DHPs synthesized at pH5 in different ascorbic acid (Asc A) concentrations.

Mw Mn P MP

Control (0%) 2469 1751 1.4 1626 Asc A (25%) 2498 1753 1.4 1667 Asc A (50%) 2641 1813 1.5 1704 Asc A (75%) 2520 1781 1.4 1913 Mw = Average molecular weight. Mn = Weighed average molecular weight. MP = Peak molecular weight P = Polydispersity.

Testing the Lignin Template Polymerization Model (Paper III)

In this study an experiment was specifically designed in order to test the template hypothesis of lignin polymerization [117]. This model proposes strict control through replication of a lignin template. For the sake of simplicity we considered synthesizing dehydrogenation polymers in the presence of a lignin model, an isolated coniferyl alcohol lignan called pinoresinol, which would represent the common lignin substructure (β-β′), Based on this hypothesis this lignin was expected to act as a template and replicate itself during in vitro polymerization of coniferyl alcohol. The phenolic groups of the lignin model were protected through methylation to prevent it from coupling during polymerization (Figure 16).

Figure 16 a. Pinoresinol and b. Pinoresinol dimethyl ether.

DHPs in the Presence of a β-β′ Model Structure Dehydrogenative polymers (DHP) were prepared from coniferyl alcohol in the presence of pinoresinol methyl ether and a control polymerization was performed in the absence of this dimer. During the reaction no precipitation of the formed DHPs was observed due to the dioxane/water environment used. The final material was fractionated into two parts for separate analyses: a low molecular weight part, containing dimers produced from polymerization and the methylated pinoresinol model; and a polymeric part. The low

30 molecular weight fraction was analyzed by GC-FID for dimer quantification. The polymeric material was studied by quantitative 13C NMR and the substructure amounts were determined by peak integration in the same way as described in Paper II except that only peaks 1, 3, 4 and 8 were quantified.

NMR quantification results indicated no increase in the relative amount of β-β′ structures in the template experiment compared to our control (Table 4). The amounts of two other common substructures (β-O-4′ and β-5′) were determined as well in order to see if a template effect would quantitatively influence their formation, either positively or negatively. It was concluded that there was no specific structural influence of β-β′ model dimer as a template for replication on the polymerization of coniferyl alcohol. Rather the same pattern of bond distribution in the template experiment was reproduced in the control. Of the three inter-unit structures studied here, β-5′ was the predominant one followed by β-O-4′ and β-β′ last. This is in line with two known observations: in vitro dimerization of coniferyl alcohol produces these three structures with β-5′ as the major product [186], and dimerization is an over-represented reaction in DHPs [65,95]. Interestingly, the polymerization system used here provided relatively large amounts of β-O-4′ structures (benzyl alcohol form), compared to purely aqueous systems with enzymic oxidants at near-neutral pH [106,187]. The formation of β-O-4′ bonds is otherwise known to be favoured at lower pH [187,188]. It is not possible to determine if dimerization itself has been affected by the solvent polarity, possibly favouring the formation of β-O-4′ [189,190] or if the solubility effects of this water:dioxane system influences the final bond distribution. Dimerization and cross-coupling reactions – as in coupling between a monomer and a phenolic dimer/oligomer – do not display the same regioselectivity. β-O-4′ becomes the predominant bond-type in the cross- coupling of a phenolic end-group model and coniferyl alcohol [111]. The longer phenolic oligomeric species stay in solution, competing with monomers in coupling other monomers, the more they will affect the final bond distributions.

Table 4 13C NMR quantification of three specific substructures in DHPs produced in the presence and the absence of pinoresinol dimethyl ether.

Template DHP Control DHP Substructure -1 -1 (μmol g ) (μmol g )

β-β' 460 480

β-5' 960 940

β-O-4' 810 820

Two dimers were quantified from GC-FID analyses: the added model dimer, pinoresinol dimethyl ether, and β-β′ dimer products of polymerization (Table 5). The dimer quantification indicated no increase in the relative amount of β- β′ dimers and thus no replication effect. Other dimers, such as β-O-4′ and β- 5′, could not be properly quantified because of the lack of pure substances needed for calibration, but relative amounts are presented. The identity of the three dimers was determined by GC-MS of both silylated and acetylated samples (see appended article III for details).

31 Table 5 GC-FID quantification of specific dimers from the low molecular weight fractions. Proper quantification of β-5′ and β-O-4′ dimers could not be obtained; the area ratios (dimer:internal standard per mg sample) are presented here. Template DHP Template DHP Control dimers Control dimers Dimer dimers -1 dimers -1 (μmol g ) (Area ratios) (μmol g ) (Area ratios) β-β′ 4.7 ±0.5 4.4 ±0.2 0.07 ±0.01 0.07 ±0.003

β-5′ - - 0.76 ±0.03 1.00 ±0.1

β-O-4′ - - 0.15 ±0.003 0.19 ±0.01 Pinoresinol dimethyl 137 ±6 94 ±11 17 ±0.8 12 ±1.,4 ether

One might argue that the lignin model used, a dimer, is too short to fulfil a template function as described by Chen and Sarkanen [117]. Hypothetically, other bond-types and higher molecular weight lignin might have a stronger catalytic or template replication effect. However, since monolignol radicals can indeed couple and polymerize in the absence of any template, and the mechanism suggested by these scientists implies a strict control over the primary structure, i.e., bond-type and monolignol type, the template effect of the lignin on monolignol polymerization ought to be strong. Therefore, some replication effect, if not complete, should be expected from our template experiment and we believe that if the template mechanism proposed is valid, then any kind of lignin structure ought to have a template effect on the polymerization of monolignols, even at the dimer level.

The proposal of the proteinaceous dirigent array/template polymerization model, implying complete synthetic control over lignin polymerization, stems from scientists who question the “traditional” theory, which only confers certain chemical “control”. Although the traditional model does not comprehensively explain all known lignin features, the experimental evidence supporting this newer model is very limited. It consists mainly of two studies: the effect on the molecular weight of synthetic lignins (DHP), when macromolecular kraft lignin components are present during DHP polymerization – the kraft lignin being suggested to serve as template [191]; and the determination of the monomer sequence of a lignin hexamer fragment from Eucalyptus globulus, claimed to be a repeating fragment and used as evidence that lignin has a well defined primary structure [192]. A recent analysis of the latter [119] indicates however, that small amounts of this structure can be expected also by uncatalyzed radical polymerization, i.e., the Erdtman/Freudenberg-model. Apart from this, there are also studies localizing so-called dirigent arrays of proteins in the cell wall [115]. However, no actual “catalytic” involvement of such proteinaceous elements with lignin polymerization has been shown. From a more philosophical point of view, there is a potential problem in one of the premises, repeatedly used as an argument by the proponents of this new model [116,120,123,193-195]. The authors assume that lignin cannot be an exception compared to how biopolymers are usually synthesized in nature as expressed by the following statement: “[…]lignin formation is under full biochemical control, in harmony with the formation of all other known biopolymers.” [195]. However, no reasons are given to explain why lignin could not be an exception. It is argued that lignin polymerization could not be left to chance [116]; but no other alternatives than complete control, are envisaged. No convincing reasons are given to establish why an indirect or partial control cannot be enough. Furthermore, few lignin chemists, if any, would argue today that lignin polymerization is a completely random and uncontrolled process. On the

32 contrary, all the available reports on the different pH, hydrophobicity, monolignol radical concentration effects on the polymerization of DHPs are attempts at determining which factors influence and might “control” lignin polymerization in situ.

Part of the critique aimed at the traditional model of lignin polymerization is actually directed against the limitations of lignin analysis [121]. The critique is justified to some extent because most knowledge of lignin structure is based only on partial samples. It is assumed that the small samples are representative of the rest of the polymer. If lignin analysis has never been representative then no structure characterization should be used as conclusive evidence for (or against) the conventional theory of lignin polymerization. But then again, this criticism implies that most of the knowledge on lignin structure, developed through hundreds of studies, is seriously mistaken.

A probably relevant criticism lies in the choice of oxidase when modelling with dehydrogenation polymers [193]. Originally, oxidases from mushrooms, which do not synthesize lignin in vivo, were used by Freudenberg [196] and later also peroxidases and laccases [90]. In the last decades, horseradish peroxidase has probably been the most used enzyme in dehydrogenation polymer experiments. Although peroxidases have a wide substrate specificity and they certainly produce dehydrogenation polymers from monolignols, they do not necessarily exhibit the same activity with different substrates [98,140]. The structure of the lignin polymer should be expected to vary depending on the activity of the peroxidase used [197], especially when considering that lignin polymerization depends on an effective oxidation of monomers as well as polymer end-groups. One of the reasons for the general failure of DHPs in reproducing lignin structure more accurately might very well be the choice of oxidase.

On the Role of the Monolignol γ-carbon (Paper IV)

The biosynthesis of the monolignols consumes important amounts of energy. The normal monolignols used for lignin biosynthesis carry alcohol groups at the γ-carbon. In order to produce this alcohol the plant needs to reduce it from carboxylic acid to aldehyde and from aldehyde to alcohol (Figure 6). The γ-carbon is not actually involved in the coupling of other monolignols through radical polymerization so it is logical to ask why the plant needs to reduce this group. In addition, ferulic acid and coniferaldehyde (Figure 17) can form lignin-like polymers through dehydrogenation [71,72,198]. Is there a reason for the plant to invest energy in producing the alcohol for lignin? What effects would a lignin based on the γ-aldehyde/carboxylic acid have on the cell wall? The aim of this study was to chemically and physically characterize DHPs synthesized from three different monomers that are produced by plants: coniferyl alcohol, coniferyl aldehyde and ferulic acid. In addition, the biological significance of the γ-carbon reduction is also discussed.

Figure 17 Monomers used in different dehydrogenation synthesis of lignin-like polymers.

Characterization of DHPs from Different Phenolic Monomers Dehydrogenation polymers (DHP) from coniferyl alcohol, coniferyl aldehyde and ferulic acid (Figure 17) were synthesized by oxidation by horseradish peroxidase and H2O2. The weight yields of the DHPs were 35, 47 and 58% respectively, after extraction of low molecular weight compounds. In the conditions used here, coniferyl alcohol had a propensity to precipitate at the beginning of the reaction which could explain the relatively low DHP yield. Interestingly, the acid-based DHPs did not precipitate during the reaction in contrast to the other two DHPs.

Size exclusion chromatography analysis (SEC) (Figure 18) revealed that the three DHPs contained dispersed polymeric material. The weight average molecular weight (Mw) and the number average molecular weight (Mn) (Table 5) of the alcohol and the acid DHPs were similar, but the molecular weight at the peak (MP) of the latter was almost twice the MP of the other DHPs indicating that ferulic acid polymerization might have produced some important populations of larger polymers compared to the alcohol and aldehyde. However, caution should be exercised when comparing sizes of polymers containing different functional groups, as determined by SEC. Larger polymers might be expected of the ferulic acid polymerization because it was the only reaction that did not precipitate and solubility is important for the availability of the different oligomers for further oxidation and coupling.

Figure 18 Size Exclusion Chromatogram of acetylated DHP materials.

34 Table 5 Molecular weight values (in g mol-1 relative to polystyrene standards) from SEC analyses of the three acetylated DHP materials.

Mw Mn P MP Acid DHP 4489 3395 1,3 5017

Aldehyde DHP 2709 1898 1,4 2222 Alcohol DHP 4464 3206 1,4 2725

Mw: Weight average molecular weight; Mn: Number average molecular weight; P: polydispersity; MP: Peak molecular weight.

All three DHPs were analyzed by 13C NMR and the spectra can be viewed in Figure 19. Signal assignments were suggested (see appended article IV) based on reported values for synthesized model components but also NMR studies of isolated lignins [92,199-201]. The different substructures found are shown in Figure 20 and the following observations were made from the comparison of the three spectra:

The different DHPs could generally form the same type of bonding structures: β-O-4′, β-5′, β-β′ and 5-5′. Obvious structural differences were observed in the δ 60-90 ppm region of the three spectra. The alcohol DHP displayed a number of signals in this region corresponding to saturated α, β and γ carbons while the acid and aldehyde DHPs practically lacked any such signals. Saturated inter-unit substructures are formed when coniferyl alcohol radical coupling occurs through at least one β- carbon, i.e., β-O-4′, β-5′, β-β′ and β-1′. Although some of these bond-types occurred in the DHPs from ferulic acid and coniferyl aldehyde the structures formed were predominantly unsaturated and their signals appear in the 100-140 ppm region. There was a general lack of cyclic structures (e.g., β-5′ and β-β′) in both the acid and aldehyde DHPs compared to the alcohol DHP. The acid DHP did produce some signals, although weak, for cyclic β-5′ and β-β′ structures which contain α and β single bond structures. The acid and aldehyde DHPs lacked benzyl alcohol groups, i.e., α- hydroxyl groups, and benzyl aryl-ether structures that are typical of alcohol DHPs [66].

Figure 19 13C NMR Spectra of a. Ferulic acid DHP b. Coniferaldehyde DHP and c. Coniferyl alcohol DHP

Figure 20 Inter-unit substructures in the three different DHP materials

36 AFM images of very smooth spin-coated DHP model surfaces are presented in Figure 21. The contact angle of water was determined on the DHP model surfaces as a relative measurement of the hydrophobicity of the different polymeric materials. The results (Table 6) indicate that the coniferyl alcohol DHP was less hydrophobic than the coniferaldehyde DHP which was less hydrophobic than the ferulic acid DHP. All three DHPs indicated much higher hydrophobicity than has been previously determined for kraft lignin [202], but this was expected since the latter is a lignin derivative where both free phenolic groups and carboxyl groups are present. There are reasons to believe that the hydrophobicity of the aldehyde DHP was somewhat underestimated, since this material was incompletely dissolved in pyridine (the solvent used for making the surfaces). Therefore, the fraction of the DHP that was dissolved by the pyridine and used for making the surfaces, most likely represented a somewhat lower molecular weight DHP that was less hydrophobic.

The DHPs were also characterized with a solvent panel (Table 7). Generally, the aldehyde DHP displayed the lowest solubility; it was in some cases troublesome to dissolve completely. The carboxylic acid and alcohol DHPs showed rather similar patterns, although the ferulic acid DHP was more soluble in the more polar solvents than the alcohol (Table 7).

Figure 21 AFM tapping mode height images (1 × 1 μm) and rms roughness determinations of the different spin-coated DHP model surfaces that were used in the measurements of the contact angle of water. a) DHP from coniferyl alcohol, rms roughness 0.349 nm. b) DHP from coniferyl aldehyde, rms roughness 0.411 nm. c) DHP from ferulic acid, rms roughness 0.404 nm.

Table 6 Initial equilibrium contact angles of water on DHP model surfaces. Sample Contact angle (º)

DHP from ferulic acid 63

DHP from coniferaldehyde 60

DHP from coniferyl alcohol 58

Kraft lignin (spruce) 46* *from Norgren, et al. 2006 [202]

37 Table 7 Ocular inspection of the solubility of the different unmodified DHP materials in organic and aqueous solvents of increasing dielectric constants (except Dioxane:Water and Acetone:Water, which are mixes of solvents at 1:1 ratio).

Solvent Acid DHP Aldehyde DHP Alcohol DHP

Dioxane no no partly Ethyl acetate no no no THF yes no no

CH2Cl2 no no no Pyridine yes partly yes Acetone partly no no DMF yes partly yes DMSO yes yes yes Water no no no Dioxane: Water no no yes Acetone:Water yes No yes

In summary, our investigation has demonstrated that also monolignols with unreduced γ-carbons can form polymers with generally the same types of inter-monolignol bonds as in natural lignin. Thus, there is no reason to believe that reduction of γ-carbon is necessary to obtain a polymer. Based on the contact angle measurements and the solubility panel, we reasoned that coniferaldehyde generated the most hydrophobic polymeric material and the coniferyl alcohol the less hydrophobic. Therefore, the reduction of the γ- carbon could be a way to make the lignin slightly more hydrophilic. This is surprising for two reasons:

One of the biological roles of lignin is to make the cell wall hydrophobic and waterproof, which is needed in the development of water-conducting tissues in plants. However, a lignin structure that is too hydrophobic may interact poorly with cellulose and hemicelluloses and thereby not fulfill its biological functions efficiently.

The carboxylic acid in ferulic acid is a charged structure at physiological pH as opposed to the primary alcohol of coniferyl alcohol. It was therefore expected that the DHP made of ferulic acid would be considerably more hydrophilic than the DHP made from coniferyl alcohol, and not the opposite as in our contact angle results.

The characterization by 13C NMR demonstrated that in addition to the expected differences in the γ-carbon functional group, there was one main difference between the coniferyl alcohol DHP on one side and ferulic acid /coniferyl aldehyde DHPs on the other. The latter DHPs displayed almost only α―β unsaturated inter-unit substructures whereas the coniferyl alcohol DHP displayed both unsaturated and saturated α―β bonds. In alcohol DHPs unsaturated α―β structures commonly exist in end-groups whereas the unsaturated structures are formed through β-coupling. In lignins, α―β unsaturated structures are much less common than in DHPs. In addition, benzyl alcohols, a frequent product of β-coupling in coniferyl alcohol DHPs

38 and especially in lignins (β-O-4′ see Figure 6), may contribute considerably to the overall hydrophilicity of the polymer, offering an explanation for its lower hydrophobicity in our results. However, the relative hydrophobicity of the DHP made of ferulic acid was nonetheless surprising. Part of the explanation might lie in a loss of charge through loss of carboxylic acid groups. Apparently there is a tendency for decarboxylation of certain substructures during the oxidative polymerization of ferulic acid [72,199,203]. If and how much decarboxylation has occurred in our polymers is difficult to assert from our NMR studies, since signals from such structures were difficult to distinguish from the non-decarboxylated ones.

The structural differences between these DHPs are due to different rearomatization pathways of the quinone methide intermediates which are formed after oxidative coupling. A quinone methide intermediate is formed when a coupling occurs through at least one β-carbon. In the case of coniferyl alcohol, rearomatization always occurs through an intra- or intermolecular nucleophilic attack on the α-carbon. Due to the carbonyl group at the γ- position on both ferulic acid and coniferaldehyde, upon coupling through the β-carbon, the β-proton is acidic enough to be released for the rearomatization of the quinone methide intermediate. For ferulic acid, decarboxylation is an additional way of rearomatizing a quinone methide. These additional rearomatization pathways in the cases of the ferulic acid and coniferyl aldehyde dehydrogenative polymerization apparently limit any intermolecular nucleophilic attacks by water or a phenol compared to coniferyl alcohol polymerization. Because of the carbonyl groups at the Cγ, the recreation of a Cα―Cβ double bond during coupling is predominant for both ferulic acid and coniferaldehyde polymerization. In Figure 22, the hypothetical mechanisms are summarized for the creation of the most common intermonomeric bond in lignin, the β-O-4′ bond for coniferyl alcohol, conifer aldehyde and ferulic acid DHPs. Rearomatizing the quinone methide intermediate by an intramolecular nucleophilic attack leads to a cyclic structure. Although the possibility of forming cyclic structures, i.e., β-5′and β-β′ for ferulic acid and in the case of β-5′ for coniferaldehyde has been shown elsewhere [72,200], only weak signals from possible cyclic β-5′ and β-β′ structures were observed for the ferulic acid DHPs here. Other reports of coniferyl aldehyde dehydrogenative polymerization support the lack of cyclic β-5′ structure [71,204]. However, the cyclic β-5′ might be formed first but as it is not thermodynamically stable, it is subsequently transformed into the non-cyclic form [200]. In the case of the acid, a suggestion has been made that cyclic and non-cyclic dimers of β-5′ and β-β′ structures could be both present and in some sort of equilibrium [72,205].

How do these structural differences explain the large investment by plants in reducing the γ-carbon? Apart from an increased hydrophobicity, a coniferaldehyde or ferulic acid lignin might be expected to form less covalent bonds with hemicelluloses and other wood cell wall polysaccharides. Such covalent bonds, forming lignin carbohydrate complexes (LCC) [26,30,206], may be created by nucleophilic attacks on the α-carbon of the quinone methide intermediate of alcohols or carboxylic acids on the polysaccharide [207,208], i.e., these hydroxyl groups replace water or a phenol in the mechanism of Figure 22a. As the suggested mechanisms describe, and the results of this study show, the possibilities for extramolecular nucleophilic attacks appear to be limited and consequently the formation of this type of LCC might be limited as well. It has recently been demonstrated that LCC form entire networks in wood, where lignin crosslinks polysaccharides of different kinds. These networks have been suggested to play an important role for the mechanical properties of the wood [28].

Figure 22 Mechanism suggestions for the rearomatization of the β-O-4′ bond during coniferyl alcohol, coniferaldehyde and ferulic acid oxidative coupling. Quinone methide rearomatization in the case of (c) ferulic acid can occur as described for (b) coniferyl aldehyde too.

Thus, reducing the γ-carbon of the monolignols might convey at least one very tangible benefit, which is improving the formation of lignin- polysaccharide covalent networks. Interestingly, a mutant loblolly pine was discovered with a deficient expression of the enzyme cinnamyl alcohol dehydrogenase [87] which affects the phenylpropanoid pathway (see appended article IV for details). The viable plant exhibited decreased lignin content, but also abnormally large amounts of aldehyde groups from coniferyl aldehyde end-groups, among others. So, coniferyl aldehyde has the ability to co- polymerize with normal lignin monomers to some degree. However, this does not indicate that the reduction is unnecessary for obtaining a functional lignin, since the lignin of this mutant still contains mostly alcohol monomers which can more easily form α-carbon LCCs. Lignin has a unique ability among biopolymers for “metabolic plasticity”, i.e., that non-conventional monomers can be incorporated into the polymer [87,209]. However, this should not be understood as if any potential monomer can form a highly

40 functional lignin. The standard monomers are very well designed by nature for lignins to fulfill their biological functions, as discussed here for the reduction of the γ-carbon.

41 Conclusions

The present work adds one more factor - reducing agents - to the growing list of variables described in the literature as having influence over the structure of lignin: carbohydrate composition of the cell wall matrix, pH, oxidase activity and monolignol rate of addition. Biological reducing agents were shown to be able to affect the structure of DHPs in two ways: in one case, by producing reduced lignin structures during oxidative coupling and in another case by altering the frequency of inter-unit bonds, especially by favouring the formation of the most common lignin substructure, the β-O-4′. In addition, one part of this work has focused on a different aspect which also affects the structure of the polymer: the chemical structure of the monolignol itself. The expensive reduction of the lignin monomers’ γ-carbon all the way to an alcohol has considerable implications for the properties of the final polymer in that it allows the formation of saturated inter-unit substructures and possibly improves the formation of lignin carbohydrate complexes, important for the cell wall properties as a whole. Some attention has been given to the relatively newly suggested model of lignin polymerization implying a strict control of the lignin primary structure. Although this hypothesis is difficult to test, it is nevertheless interesting to consider whether lignin has any catalytic capabilities (or template replication capabilities). Our results here indicate no support for a replication effect as such. Nevertheless, any effect of the lignin macromolecule itself (and other cell wall macromolecular components for that matter) upon the polymerization cannot be excluded, but remains to be clarified.

The data in this thesis suggests that it is very unlikely that plants use a catalytic system, such as the one suggested by the proteinaceous dirigent array/lignin template replication hypothesis, to control the structure of lignin in great detail. Instead, it shows that plants can influence lignin polymerization and structure by ensuring the presence of reducing agents (probably of low molecular weight) during lignification. Furthermore, even though lignins are reportedly flexible in the choice of monomers, investing in the monomer γ-carbon reduction is indeed a way for the plant to generate a lignin that fulfils its biological functions fully in the plant cell wall.

In several ways, our understanding of lignin polymerization chemistry is only tentative. The control mechanisms at work defining lignin structure, especially during polymerization, are today still unclear. Although there is a suggestion for a strict catalytical system controlling it in the cell wall, there is no convincing evidence supporting it. It is therefore all the more exciting to elucidate an alternative system in nature of apparently rather subtle control: one that is not enzymatically based. For that reason, establishing how and what factors in the plant cell wall, chemical and physical, influence the final structure of the lignin polymer is necessary; this might be useful, e.g., when attempting to alter the structure of lignin in plants for the purpose of improving pulping results.

Future research with DHPs as models of lignin polymerization would benefit from some highly desired improvements. For example, the choice of peroxidase should be considered more carefully in light of the findings that have revealed differing activities of enzymes oxidizing monomers compared to polymer end-groups. The kinetics of radical couplings between the different phenolic species (monomer-monomer and monomer-oligomer) should be

42 determined to permit the better understanding of how different radical concentrations favour the formation of either bulk or end-wise polymers. Finally, the structure of DHPs is dependent on the solubility of the different oligomeric species, but this is probably not the case in the cell wall where growing lignin oligomers are more likely to be fixed to a “solid phase”, and thus also less likely to react with each other. These oligomer-oligomer reactions cannot be avoided in conventional “solution” DHPs. Thus, the development of an effective system where oligomers are fixed to a solid phase might improve end-wise polymerization and make the model polymers even more similar to natural lignins.

43 Acknowledgements

I would like to express a few words of gratitude to the many persons who have supported my work. Without them it would not have been possible.

First and foremost my two supervisors:

Professor Gunnar Henriksson, for always being available for discussions and always encouraging, as well as being a constant idea-generator in this project; and

Dr. Liming Zhang, for all your experimental guidance and ideas, and especially for all the practical assistance with the NMR.

And also:

Professor Gösta Brunow, for our many rewarding discussions and your vast knowledge of the relevant literature as well as for your cooperation in Paper I. Your practical and theoretical knowledge in the field has been invaluable.

Professor John Ralph, for our many valuable discussions and your genuine interest in our research project as well as for your cooperation in Paper I.

Dr. Magnus Norgren, for your valuable cooperation in Paper II.

Inga Persson, for taking care of all the formalities related to the project and especially for going the extra mile in many things and making the whole group feel more at home.

I would also like to thank the Swedish Research Council for Environment, Agricultural Sciences and Spatial Planning (FORMAS), contract no 229-2004- 1240, for financial support.

Many thanks to some special colleagues, Martin, Maria, Andrea, Lotta, Viviana, Sverker, Stefan, Jiebing, David, Juanjo and Dimitri, for many discussions and help, and for making my work more fun; and many thanks to all colleagues from the Fibre and Pulp technology groups for contributing to a great working atmosphere.

Last but not least, I want to thank my family which has doubled in size since I started my Ph.D. studies. You are a daily encouragement and I cannot imagine life without you. Many thanks also to my parents for all the support and valuable suggestions while writing the thesis, and …to God for helping me keep perspective on life.

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58 Glossary

Angiosperm: Vascular plant in which Libriform fibre: Angiosperm cell with a the ovule is fertilized and develops mechanical support function. into a seed in an enclosed hollow ovary. Lignans: Heterocyclic oligomeric molecules synthesized from the ATP: Adenosine triphosphate. Energy monolignols and involved in plant rich molecule and enzyme co-factor defence. involved in many important metabolic processes. Lignosulfonates: Water soluble polyelectrolyte lignin-based Club mosses: A small creeping vascular polymers recovered as a by-product plant which reproduce sexually from the sulfite pulp-making through spores. process.

Coumarins: Heterocyclic organic Lumen: The cell space left when molecules with characteristic emptied from all organelles and odours, involved in plant defence. cytosolic compounds. Lumen occurs in for example dead xylem Ferns: The most diversified group of cells. vascular plants which display true leaves and reproduce through Melanosome: Organelle of animal cells spores. (usually called melanocytes) which synthesize and contain melanin. Flavonoids: Heterocyclic organic molecules functioning as pigments, Monolignol: Specific name for the three for example in flower petals. common lignin monomers: p- coumaryl alcohol, coniferyl alcohol Gymnosperm: Vascular plant that and sinapyl alcohol. reproduces by means of an exposed seed, or ovule. NAD(P)H: Nicotinamide adenine dinucleotide (phosphate) reduced Hardwood: An angiosperm woody form. Electron rich molecules and plant, generally more dense and enzyme co-factors involved in with a more complex structure than important metabolic processes. softwood. Nucleic acids: Biopolymers such as Hemicellulose: Complex heteropolymer DNA or RNA, built of nucleotides of different sugar monomer usually carrying genetic composition found in the cell wall information. of plants. Number average molecular Weight (Mn): Horsetails: Small vascular plant It is the common average weight of growing a single hollow jointed any individual polymer in a mix of stem, which reproduce through varied-sized polymers. spores. Oxidative phosphorylation: The process Kenaf: Herbaceous annual (or biennial) in which biochemical energy (ATP) plant with a woody base growing is formed through an electron several meters high and transfer mechanism involving the reproducing trough seeds. reduction of molecular oxygen.

Laccase: Copper containing enzymes Parenchyma: Living plant cells with capable of carrying out one- thin cell walls dedicated to storage, electron oxidation of phenols by found in both xylem and phloem reducing molecular oxygen to tissues. water.

59 Pectins: Heterogenous polysaccharides Styrylpyrones: Heterocyclic compounds common in the cell walls of plants. involved in plant defence, structurally close to stillbenes. Phloem: Vascular plant tissue that carries organic nutrients. Suberin: Complex hydrophobic plant substance in plants preventing Polydispersity: A way of describing the free water diffusion. It is found in distribution of molecular weights in the roots and bark. a polymer sample. Tracheids: Xylem cells developed for Protozoans: Single cell eukaryotic water conduction and structural organisms generally very mobile. support. Some are parasitic like the amoebas. TEM: Transmission electron microscopy. Raman spectroscopy: Spectroscopic technique used to study, e.g., Vascular cryptogram: Collective term vibrational energy in molecular for all vascular plants which do not systems, complementary to IR reproduce through seeds. spectroscopy. Vessel elements: Angiosperm xylem Schlerenchyma: Plant cells with heavily cells specifically developed for thickened and lignified walls water transportation dedicated to mechanical support. Weight average molecular weight (Mw): Sclerotization: The process of It is an average molecular weight hardening and darkening of insect that has been weighed against the cuticles. size of the different polymer populations. Softwood: A Gymnosperm woody plant with a generally less dense and Xylem: Vascular plants tissue that with a simpler structure than carries water and dissolved hardwood. minerals.

Stillbenes: Polyphenolic compounds involved in plant defense, particularly in hardwood protection.