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Keywords:

  • cell wall;
  • lignin;
  • pathway discovery;
  • phenolic metabolism;
  • phenolic profiling;
  • synthetic biology

Abstract

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

Contents

 Summary978
I.Introduction979
II.Phenolic metabolism979
III.Lignin biosynthesis and structure983
IV.Alternative lignin monomers for biofuel applications985
V.Candidate alternative monolignols in biomimetic systems991
VI.From phenolic profiling to lignomics992
VII.Phenolic pathway engineering towards alternative monolignols993
 Acknowledgements994
 References994

Summary

Lignin, a phenolic polymer in the secondary wall, is the major cause of lignocellulosic biomass recalcitrance to efficient industrial processing. From an applications perspective, it is desirable that second-generation bioenergy crops have lignin that is readily degraded by chemical pretreatments but still fulfill its biological role in plants. Because plants can tolerate large variations in lignin composition, often without apparent adverse effects, substitution of some fraction of the traditional monolignols by alternative monomers through genetic engineering is a promising strategy to tailor lignin in bioenergy crops. However, successful engineering of lignin incorporating alternative monomers requires knowledge about phenolic metabolism in plants and about the coupling properties of these alternative monomers. Here, we review the current knowledge about lignin biosynthesis and the pathways towards the main phenolic classes. In addition, the minimal requirements are defined for molecules that, upon incorporation into the lignin polymer, make the latter more susceptible to biomass pretreatment. Numerous metabolites made by plants meet these requirements, and several have already been tested as monolignol substitutes in biomimetic systems. Finally, the status of detection and identification of compounds by phenolic profiling is discussed, as phenolic profiling serves in pathway elucidation and for the detection of incorporation of alternative lignin monomers.

I. Introduction

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

The development of lignin was a keystone event in the evolution of vascular land plants because lignin provided the necessary strength and hydrophobicity to fiber and vessel cell walls to allow plants to grow tall in a gravitropic environment and to transport water and nutrients in their vascular system (Rogers & Campbell, 2004; Weng & Chapple, 2010). Lignin is a complex aromatic polymer in which the cell wall polysaccharides (mainly cellulose and hemicelluloses) and cell wall glycoproteins are embedded. It is synthesized from the oxidative coupling of p-hydroxycinnamyl alcohol monomers and related compounds (Boerjan et al., 2003; Ralph et al., 2004b; Vanholme et al., 2010a). The main units in the polymer, p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) units, are derived from the monolignols p-coumaryl, coniferyl and sinapyl alcohol, monomers differing in the number of methoxyl substituents on the aromatic ring. The relative abundance of these main units varies among plant species, tissues, cell types, and developmental stages. Research on lignin biosynthesis has become a primary focus since it became clear that lignin is the major factor in lignocellulosic biomass recalcitrance to efficient processing. Lignin hinders the release of monosaccharides during enzymatic hydrolysis of cell wall polysaccharides – a process called saccharification – which is necessary for the production of second-generation biofuels and materials from biomass-derived sugars (Chen & Dixon, 2007; Yoshida et al., 2008). More specifically, lignin negatively affects the saccharification process by immobilizing cellulases (and associated enzymes) and blocking them from reaching their polysaccharide substrates (Chang & Holtzapple, 2000; Nakagame et al., 2011). In order to facilitate saccharification, mechanical, thermal and chemical pretreatments have been developed to disrupt cell wall structure, rendering the lignocellulosic material more accessible to the polysaccharidases (Carvalheiro et al., 2008; Kristensen et al., 2008; Hendriks & Zeeman, 2009; Chundawat et al., 2011). With the recognition that lignin is a problem, it has been appealing to consider reducing lignin contents via genetic engineering, a strategy that would reduce the input of chemicals and energy during the pretreatment reactions (Chen & Dixon, 2007). Drastic reductions in the biosynthesis of lignin, however, have negative effects on plant growth and development (Li et al., 2010; Gallego-Giraldo et al., 2011b; Voelker et al., 2011). By contrast, plants can tolerate large shifts in lignin composition, often without visible effects on plant development and morphology (Ralph et al., 1997; Meyer et al., 1998; Marita et al., 1999; Franke et al., 2000; Stewart et al., 2009; Vanholme et al., 2012). In fact, plants also show a remarkable ability to augment their polymer make-up by incorporating novel phenolic monomers, as is particularly evident when pathway gene down-regulations, limiting the flux to the traditional monolignols, lead to the build-up of products from pathway intermediates, for example, hydroxycinnamaldehydes, and 5-hydroxyconiferyl alcohol (Ralph et al., 2001b). Advances in analytical techniques have revealed that numerous phenolic metabolites act as natural lignin monomers in wild-type plants – examples include acylated hydroxycinnamyl alcohols, hydroxybenzaldehydes and dihydro-hydroxycinnamyl alcohols (Ralph et al., 1997, 2008a; Vanholme et al., 2008). The incorporation of atypical monomers that are rare, or even absent, in lignin of wild-type plants (hereafter called alternative lignin monomers) can be accomplished through genetic engineering (Jackson et al., 2008; Ralph et al., 2008b; Eudes et al., 2012). Therefore, research is now also focusing on the biosynthesis and incorporation of alternative monomers into lignin to alter the structure of the lignin polymer to facilitate lignin removal from lignocellulosic biomass by chemical pretreatments or to improve the penetration and action of hydrolytic enzymes (Simmons et al., 2010; Chundawat et al., 2011). Various modified lignin polymers might be envisioned to maintain the biological role of lignin in the plant while permitting more efficient conversion of lignocellulosic biomass for industrial saccharification.

II. Phenolic metabolism

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

The lignin polymer is the product of oxidative coupling of phenolic metabolites, normally p-hydroxycinnamyl alcohol monomers (the so-called monolignols). Steering the pathway to produce alternative monomers therefore requires a fundamental knowledge of phenolic metabolism, that is, the enzymes and metabolites involved in the pathways, and how these pathways are regulated. Although many phenolics are specific for one or a few plant species, several major classes of phenolics are found throughout the plant kingdom. The biosynthetic route towards these major classes has been studied via metabolic and genomic tools, mostly in Arabidopsis. In this section, the current knowledge of the main phenolic pathways in Arabidopsis and in other model species is briefly presented as a prelude to describing alternative monomers for lignification (Fig. 1).

image

Figure 1. Phenolic metabolism in plants. The phenolic metabolite classes are given (in gray frames), as well as pathways and metabolic sinks that use phenolic metabolites or shikimate pathway intermediates as substrates. Representative metabolites are given for phenolic classes. Not every phenolic metabolic class shown is present in every plant species. The major route towards the monolignols p-coumaryl, coniferyl and sinapyl alcohol is given in color; the shikimate pathway (yellow), phenylalanine biosynthesis (orange), general phenylpropanoid pathway (pink) and monolignol-specific pathway (purple). Arrows with dashed lines designate known routes that involve multiple enzymatic steps; for simplicity, the individual enzymatic steps are not shown. Arrows with dotted lines designate unknown or unauthenticated routes. Arrows with a question mark are routes that have been suggested in the literature. DHS, 3-deoxy-D-arabino-heptulosonate 7-phosphate synthase; DQS, 3-dehydroquinate synthase; DHQD, 3-dehydroquinate dehydratase; SD, shikimate dehydrogenase; SK, shikimate kinase; EPSPS, 5-enolpyruvylshikimate-3-phosphate synthase; CS, chorismate synthase; AT, amino transferase; TAL, tyrosine ammonia-lyase; PAL, phenylalanine ammonia-lyase; C4H, cinnamate 4-hydroxylase; 4CL, 4-coumarate: CoA ligase; HCT, hydroxycinnamoyl-CoA: shikimate/quinate hydroxycinnamoyltransferase; C3H, p-coumarate 3-hydroxylase; CCoAOMT, caffeoyl-CoA O-methyltransferase; CCR, cinnamoyl-CoA reductase; F5H, ferulate 5-hydroxylase; COMT, caffeic acid O-methyltransferase; CAD, cinnamyl alcohol dehydrogenase; UGT, UDP-glucosyltransferase; HCALDH, hydroxycinnamaldehyde dehydrogenase; BGLU, β-glucosidase.

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The shikimate pathway, which is present in bacteria, yeasts and plants, but not in animals, is the entry pathway towards a plethora of phenolic compounds. This plastid-localized pathway is highly transcriptionally and post-translationally controlled (Chen et al., 2006b; Tzin & Galili, 2010). In seven enzymatic steps, the glycolytic intermediate phosphoenol pyruvate and the pentose phosphate pathway intermediate erythrose-4-phosphate are metabolized into chorismate via 3-dehydroshikimate as an intermediate (Herrmann & Weaver, 1999). Although the biosynthetic route is not yet fully elucidated, 3-dehydroshikimate also serves as a precursor for gallic acid and, thus, gallotannin biosynthesis (Dewick & Haslam, 1969; Werner et al., 2004). Chorismate serves as the precursor for p-aminobenzoate (an intermediate in tetrahydrofolate biosynthesis; Basset et al., 2004) and the aromatic amino acids phenylalanine, tyrosine and tryptophan (Knaggs, 2003). Phenylalanine is produced from chorismate by the action of two enzymes, a dehydratase and an aminotransferase, the exact order of action being unknown (Cho et al., 2007; Yamada et al., 2008; Corea et al., 2012). Tryptophan is necessary for the production of auxin and secondary indolic metabolites, such as indolic glucosinolates and camalexin (Malitsky et al., 2008). In addition, chorismate is the main precursor for salicylic acid in Arabidopsis, although salicylic acid can also be produced from benzoic acid (Léon et al., 1995; Wildermuth et al., 2001; Métraux, 2002; Strawn et al., 2007).

The general phenylpropanoid pathway uses phenylalanine as an entry substrate and results, after seven steps, in feruloyl-CoA (Humphreys & Chapple, 2002; Boerjan et al., 2003). Following deamination of phenylalanine to cinnamate by phenylalanine ammonia-lyase (PAL), hydroxylation of the aromatic ring leads to p-coumarate, a reaction catalyzed by cinnamate 4-hydroxylase (C4H). In grasses, a PAL isozyme catalyzing the deamination of both phenylalanine (PAL activity) and tyrosine (tyrosine ammonia-lyase activity) in vitro might be implicated in the direct conversion of tyrosine to p-coumarate in vivo (Rösler et al., 1997). Activation of the acid to a thioester by 4-coumarate: CoA ligase (4CL) yields p-coumaroyl-CoA. The subsequent 3-hydroxylation of p-coumaroyl-CoA to caffeoyl-CoA involves three enzymatic steps, at least in dicots. First, p-coumaroyl-CoA is transesterified to its quinic or shikimic acid ester derivative by hydroxycinnamoyl-CoA: shikimate/quinate hydroxycinnamoyltransferase (HCT). p-Coumaroyl shikimate or quinate is then hydroxylated by p-coumarate 3-hydroxylase (C3H, named when it was assumed that p-coumarate was the direct substrate) and then transesterified again by HCT to caffeoyl-CoA. Recently, an alternative 3-hydroxylation route has been found; the poplar (Populus trichocarpa) heterodimeric C4H/C3H protein complex efficiently converts p-coumaric acid to caffeic acid (Chen et al., 2011), after which 4CL might convert caffeic acid into caffeoyl-CoA. Further methylation of the 3-hydroxyl group by caffeoyl-CoA O-methyltransferase (CCoAOMT) yields feruloyl-CoA. Various pathways branch off from the general phenylpropanoid pathway, including the monolignol-specific pathway and the pathways towards flavonoids, benzenoids, coumarins, and sinapate and ferulate esters.

The monolignol-specific pathway includes four well-studied enzymatic steps that convert feruloyl-CoA into the monolignols coniferyl alcohol and sinapyl alcohol (Humphreys & Chapple, 2002; Boerjan et al., 2003). First, feruloyl-CoA is reduced to coniferaldehyde by cinnamoyl-CoA reductase (CCR). Hydroxylation at the 5-position is catalyzed by ferulate 5-hydroxylase (F5H), which is also now often called coniferaldehyde 5-hydroxylase (CAld5H) to reflect its preferred substrate, to produce 5-hydroxyconiferaldehyde (Humphreys et al., 1999; Osakabe et al., 1999). The subsequent methylation of the newly introduced 5-hydroxyl group is catalyzed by caffeic acid O-methyltransferase (COMT), whose preferred substrate is also now known to be the aldehyde (Li et al., 2000; Parvathi et al., 2001), to provide sinapaldehyde. Further reduction to their corresponding alcohols, coniferyl alcohol and sinapyl alcohol, is catalyzed by cinnamyl alcohol dehydrogenase (CAD). A specific sinapyl alcohol dehydrogenase (SAD) involved in the reduction to sinapyl alcohol has been proposed in aspen (Populus tremuloides) based on in vitro enzymatic assays (Li et al., 2001), but an in vivo role of SAD in monolignol biosynthesis has never been convincingly demonstrated (Guo et al., 2010; Barakate et al., 2011). Although the depicted pathway is thought to occur in many species (certainly Arabidopsis, tobacco (Nicotiana tabacum) and poplar), an alternative sequence of reactions for sinapyl alcohol production probably occurs in Medicago (Lee et al., 2011) and the pathways in grasses are currently being evaluated (Withers et al., 2012). In Medicago, the flux towards coniferyl and sinapyl alcohol bifurcates after caffeoyl-CoA production (Lee et al., 2011). Caffeoyl-CoA destined for sinapyl alcohol synthesis is reduced by CCR to caffealdehyde, which is then converted to sinapaldehyde by the sequential actions of COMT, F5H and COMT.

The monolignols that result from the above-described monolignol-specific pathway are used for at least three different product classes: oligolignols/lignin, monolignol 4-O-hexosides and (neo)lignans. Oligolignols are racemic radical coupling products of monolignols that arise during lignin polymerization (Morreel et al., 2004a, 2010a,b; Ralph et al., 2004b). Lignans are formed by the initial stereospecific β-β coupling (see Fig. 2 for nomenclature) of two monolignol radicals (Umezawa, 2003). Secondary metabolites arising from two monolignol radicals that are stereospecifically β-O-4- or β-5-coupled are called neolignans (Umezawa, 2003). Stereospecific coupling reactions in (neo)lignan biosynthesis appear to be assisted by dirigent proteins (Davin et al., 1997; Umezawa, 2003; Beejmohun et al., 2007; Pickel et al., 2010). Because of their antioxidant properties, (neo)lignans are believed to be involved in defense responses (Davin et al., 1997). Some may also have a hormonal function; dehydrodiconiferyl alcohol glucoside (DCG) has been associated with cell division-promoting activities (Binns et al., 1987; Teutonico et al., 1991; Li et al., 2010). The third metabolic class derived from monolignols includes the 4-O-glucosylated monolignols (e.g. coniferin and syringin). Several glucosyl transferases involved in their biosynthesis have been described in Arabidopsis, as well as β-glucosidases that convert the monolignol 4-O-glucosides back to their respective aglycones (Lim et al., 2005; Escamilla-Treviño et al., 2006; Lanot et al., 2006). The biological role of monolignol 4-O-hexosides has not been unequivocally defined, but they could serve as storage forms for their aglycones (Lim et al., 2005; Vanholme et al., 2012). This hypothesis is supported by the finding that monolignol 4-O-glucosides are sequestered into the vacuoles of Arabidopsis, whereas monolignol aglycones are transported to the apoplast (Miao & Liu, 2010; Alejandro et al., 2012).

image

Figure 2. Oxidative radicalization (i.e. dehydrogenation) and coupling of the traditional lignin monomer, coniferyl alcohol. (a) The 4-O-localized radical formed upon dehydrogenation is stabilized via delocalization, as shown by the five resonance forms. By convention, α, β and γ, and 7, 8 and 9 are used to indicate the aliphatic carbon positions in lignin monomers/polymers and (neo)lignan/oligolignols, respectively. To ensure consistent nomenclature in the text, α, β and γ are used for both lignins and (neo)lignan/oligolignols. (b) β-O-4 coupling of two coniferyl alcohol monomers. An external nucleophile (Nu:) provides the pathway for re-aromatization of the quinone methide intermediate. In the (common) case where the nucleophile is water, this results in the hydroxylation at the Cα position. However, if the nucleophile is a hydroxyl function from a hemicellulosic sugar moiety, a covalent ether bond between lignin and the hemicellulose is formed (see Fig. 4). (c) β-5 coupling of two coniferyl alcohol monomers. Re-aromatization of the quinone methide intermediate is via internal trapping (via the formal 4-OH that can also be considered to result from keto-enol tautomerization of the intermediate shown), and a phenylcoumaran structure is formed. (d) β-β coupling of two coniferyl alcohol monomers. The γ-hydroxyl groups serve as nucleophiles for internal trapping of the quinone methide, resulting in two re-aromatization reactions (that are likely to be sequential, and not necessarily concerted as shown here), and the formation of a resinol structure. The bond formed during the radical coupling reaction is the one used to describe the various dehydrodimerization reactions and is shown in bold. Lignification proceeds mainly via endwise addition of new monolignols to the free-phenolic end, following renewed radicalization, at the 4-O- or 5-positions (and also, to a small extent, at the 1-position) shown by the dotted arrows.

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The general phenylpropanoid- and monolignol-specific pathways also provide hydroxycinnamic acids, which include p-coumaric, caffeic, ferulic and sinapic acids. Hydroxycinnamic acids can be esterified or amidated by a variety of moieties such as malate, quinate, glucose, sucrose, choline, putrescine, spermidine, hydroxyanthranilate and tyramine; this can differ between plant species and plant tissues (Dimberg et al., 1993; Martin-Tanguy, 1997; Schmidt et al., 1999; Mahesh et al., 2007; Milkowski & Strack, 2010). In Arabidopsis, the largest portion of the ferulic and sinapic acid pool is made from coniferaldehyde and sinapaldehyde via hydroxycinnamaldehyde dehydrogenase (HCALDH) (Nair et al., 2004). The presence of ferulate and sinapate esters in hcaldh and ccr1 mutants, however, proves the existence of an alternative pathway probably involving feruloyl-CoA (Nair et al., 2004; Chen et al., 2006a; Mir Derikvand et al., 2008; Vanholme et al., 2012). Alternatively, ferulic acid might be made from p-coumaric acid via 3-hydroxylation and 3-O-methylation by C3H/C4H and COMT (Chen et al., 2011). Sinapate esters (e.g. sinapoyl glucose and sinapoyl malate) are putatively important as UV-protectants in Brassicaceae and the genes involved in their biosynthesis are well described in Arabidopsis (Lorenzen et al., 1996; Fraser et al., 2007; Sinlapadech et al., 2007). By contrast, the biological function and biosynthesis of ferulate esters are poorly understood. Ferulate esters are likely intermediates in the production of cell wall-bound ferulates (Rohde et al., 2004; Mir Derikvand et al., 2008) and of suberins (Bernards et al., 1995; Franke & Schreiber, 2007; Soler et al., 2007; Molina et al., 2009; Rautengarten et al., 2012). Ferulate-polysaccharide esters have well-established roles in polysaccharide–polysaccharide and lignin–polysaccharide cross-linking in grasses (Ralph et al., 1994a,b, 1998; Hatfield et al., 1999; Grabber et al., 2000; Ralph, 2010), and may even function as nucleation sites for lignification (Ralph et al., 1995; Grabber et al., 2002). Ferulate esters are also found in neolignan oligomers (e.g. G(β-O-4)feruloyl malate) for which the biological role is as yet unknown (Rohde et al., 2004; Böttcher et al., 2008; Meißner et al., 2008; Huis et al., 2012; Vanholme et al., 2012). Hydroxycinnamic acid polyamine conjugates are present in numerous plant species (Martin-Tanguy, 1997) and several of their biosynthetic genes have been characterized in Arabidopsis (Fellenberg et al., 2009; Grienenberger et al., 2009; Matsuno et al., 2009). Hydroxycinnamic acids are also 4-O-glucosylated, resulting in metabolites that share characteristics with 4-O-glucosylated monolignols and hydroxycinnamaldehydes (Lim et al., 2005). Finally, sinapyl and coniferyl p-coumarate esters serve as (nontraditional) monolignols of lignin (Lu & Ralph, 1999; Ralph, 2010).

Via the action of polyketide synthases, several phenolic pathways diverge from phenylpropanoid biosynthesis (Yu & Jez, 2008). For instance, the flavonoid biosynthetic pathway derives mainly from p-coumaroyl-CoA. The flavonoid biosynthetic pathway can be further divided into the general flavonoid, flavonol, isoflavonoid, anthocyanin and proanthocyanin (condensed tannin) pathways (Tanner et al., 2003; Lapčík, 2007; Yonekura-Sakakibara et al., 2008). Structural genes of the general flavonoid pathway are well characterized in Arabidopsis (Yonekura-Sakakibara et al., 2008), but enzymatic steps involved in flavonoid modification (e.g. glycosylation and acylation) are poorly understood. The committed step towards flavonoids, catalyzed by the polyketide synthase chalcone synthase (CHS), condenses p-coumaroyl-CoA with three acetyl-CoA molecules to produce naringenin chalcone. Although aromatic hydroxylations can occur within the flavonoid pathway itself, in some species CHS may accept multiple hydroxycinnamoyl-CoA substrates to create a diverse set of hydroxylated and methoxylated flavonoids after the first step in flavonoid biosynthesis (Dao et al., 2011). For example, sinapoyl, caffeoyl and p-coumaroyl-CoA are at least formal precursors of various flavones, such as tricin (found in grasses), luteolin and apigenin. Flavonoid biosynthesis and monolignol biosynthesis are even more intertwined; for example, Arabidopsis COMT also carries out its methoxylation reaction on flavonols to convert quercetin to isorhamnetin (Muzac et al., 2000; Goujon et al., 2003), and tricin has just been implicated as a monomer in grass lignins (del Río et al., 2012). In specific plant species, hydroxycinnamoyl-CoAs are the substrates for other polyketide reactions. For instance, stilbene synthase catalyzes the biosynthesis of stilbenes in pine (Pinus sylvestris and Pinus densiflora), grape (Vitis vinifera), peanut (Arachis hypogaea) and sorghum (Sorgum bicolor) (Hammerbacher et al., 2011).

Benzenoids are characterized by a C6-C1 skeleton. Some benzenoids, such as benzoic, p-hydroxybenzoic and vanillic acids, are made via chain-shortening of the C6-C3 skeleton of phenylpropanoids, whereas others, such as gallic acid, are synthesized from 3-dehydroshikimate. As described in the second paragraph of Section II, salicylic acid can be synthesized via chorismate or from benzoic acid. Two possible routes have been proposed for the biosynthesis of benzoic acid from cinnamic acid: the CoA-independent (non-β-oxidative) route via benzaldehyde and the CoA-dependent (β-oxidative) route via benzoyl-CoA (Hertweck et al., 2001; Boatright et al., 2004; Ibdah et al., 2009). Except for an aldehyde oxidase involved in the CoA-independent biosynthesis of benzoic acid in seeds of Arabidopsis (Ibdah et al., 2009) and a cinnamate:CoA ligase (CNL) and a 3-ketoacyl-CoA thiolase involved in the CoA-dependent biosynthesis of benzoic acid in petunia (Petunia hybrida BA) flowers (Van Moerkercke et al., 2009; Klempien et al., 2012), none of the genes involved in these pathways have been described. Because of similarities between fatty acid catabolism and the CoA-dependent chain-shortening of phenylpropanoids to benzenoids, the respective enzymes of these routes are expected to be (distant) homologs (Hertweck et al., 2001). p-Hydroxybenzoates acylate lignins in Populus, Salix and Palmae species (Ralph, 2010), and benzenoids such as methylsalicilate, methylbenzoate, benzylbenzoate and benzylacetate, are volatiles and part of the floral scent (Dudareva et al., 2004).

Another biosynthetic route that branches from the general phenylpropanoid biosynthetic pathway leads to coumarins. Following C2-hydroxylation of the hydroxycinnamoyl-CoA esters, lactonization of the side-chain produces the corresponding coumarins, which are often stored as glycosides. This yields umbelliferone, esculetin and scopoletin as the coumarin products of p-coumaric, caffeic and ferulic acids, respectively. The enzyme responsible for the first step, that is, the C6′ hydroxylation of feruloyl-CoA, has been described in Arabidopsis (Kai et al., 2008).

Although much information about phenolic metabolism has been gathered over recent decades, the low proportion of identified metabolites in phenolic profiling studies (Morreel et al., 2010a; Vanholme et al., 2010c) underscores the complexity of these pathways and the need for methods to speed up structural characterization. Certainly, knowledge gaps in metabolic products and pathways will hamper attempts to bioengineer new types of lignin using alternative monomers from the phenylpropanoid metabolism.

III. Lignin biosynthesis and structure

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

After their biosynthesis, monolignols are translocated to the apoplast via a largely unresolved mechanism probably involving ATP-binding cassette (ABC) transporters (Miao & Liu, 2010). Recently, an Arabidopsis ABC transporter (AtABCG29) has been identified that is capable of transporting p-coumaryl alcohol when expressed in yeast, whereas Arabidopsis mutated for this transporter contained less lignin and was more sensitive to p-coumaryl alcohol (Alejandro et al., 2012). Upon entering the cell wall matrix, monolignols are oxidized by peroxidases and/or laccases to monolignol radicals that eventually polymerize into the lignin macromolecule via combinatorial radical–radical coupling reactions (Fig. 2). Laccases and peroxidases are encoded by large gene families and the individual members have overlapping activities (Vanholme et al., 2010a). Two laccases involved in lignification have been identified (Berthet et al., 2011). Direct contact between the peroxidase/laccase and the substrate is not needed – radical-transfer reactions can also pass the radical from one molecule to another. All radical coupling reactions in lignification are termination events; thus, continued lignification of cell walls requires new hydrogen-abstraction of monomers and the growing lignin oligomer following each coupling reaction. It is assumed, but not proven, that a monolignol radical can transfer its single electron to the growing polymer, but other transfer agents may be involved. For instance, the p-coumarate moiety in sinapyl p-coumarate, or manganese oxalate, may act as radical shuttles (Takahama et al., 1996; Takahama & Oniki, 1997; Önnerud et al., 2002; Ralph et al., 2004a; Hatfield et al., 2008). Because radical coupling is a purely chemically driven process, independent of control by any proteinaceous agent, any phenolic molecule having the proper chemical kinetic and thermodynamic radical-generation and cross-coupling propensities can couple into the lignin polymer (Harkin, 1967; Lu & Ralph, 1999, 2002, 2008; Boerjan et al., 2003; Morreel et al., 2004a; Ralph et al., 2004b; Ralph, 2006; Vanholme et al., 2010a).

The monolignol radical is resonance-stabilized, having various sites of enhanced single-electron density in the molecule (Fig. 2a). Mutual coupling of monolignols (dimerization) and cross-coupling with the growing polymer lead not only to the characteristic H, G and S units, but also to various inter-unit linkage types (Fig. 2b–d). At least 60% of the inter-unit linkages in dicots are β-aryl ethers arising from β-O-4 coupling of a monolignol at its β-position to the 4-O-position of the growing oligomer. These β-aryl ether linkages can, unlike other prevalent interunit linkages, be cleaved by harsh alkaline or acidic pretreatment of the lignocellulosic biomass (Sarkanen & Ludwig, 1971). The two other major inter-unit linkages in lignin are phenylcoumarans and resinols formed by β-5 and β-β coupling, respectively. Both are carbon–carbon linkages (among the so-called ‘condensed linkages’) that can only be broken under extremely harsh conditions that would also degrade the polysaccharides. For the three major types of linkages, the incoming monolignol radical reacts exclusively at its β-position, enabling the resulting 4-O-phenolic function produced after re-aromatization of the quinone methide intermediate to enter another coupling reaction. Although the β-O-4, β-5, and β-β couplings yield a linear polymer, branching can occur whenever the 4-O- or 5-position of one lignin oligomer or polymer couples with the 5-position of another lignin oligomer or polymer, producing 5-5 and 4-O-5 linkages. The coupling reactions involved in lignification have been previously reviewed in detail (Ralph et al., 2004b).

Which primary units are present in the lignin polymer depends largely on the taxon of the plant. In general, gymnosperm lignins are rich in G units, with small amounts of H units, but no S units. Lignins from dicots are composed of both G and S units with only traces of H units, whereas lignin of monocot grasses also contains G and S units with modest levels (typically < 5%) of H units. It should be pointed out that H unit proportions in grasses are often overestimated because p-coumarate units acylating lignin are often mistakenly quantified as H units (Boerjan et al., 2003; del Río et al., 2012). Nevertheless, it should be stressed that the lignin unit composition is highly variable, not only between species, but also between tissue and cell types, and even within a single cell wall. In addition, any phenolic molecule entering the cell wall region may be oxidized and incorporated into the lignin polymer (Harkin, 1967; Lu & Ralph, 1999, 2002, 2008; Boerjan et al., 2003; Morreel et al., 2004a; Ralph et al., 2004b; Ralph, 2006; Vanholme et al., 2010a). Many alternative monomers are found in the lignin of wild-type plants. For example, traditional monolignols are often acylated at their γ-position with acetate, p-hydroxybenzoate or p-coumarate (Lu & Ralph, 1999, 2002, 2008; Morreel et al., 2004a). Such acylated units can even be highly abundant; for example, coniferyl and sinapyl acetate may constitute 50% or more of the units in lignin from kenaf (Hibiscus cannabinus) (Ralph, 1996; Del Río et al., 2007). Also, dihydro-hydroxycinnamyl alcohols, hydroxybenzaldehydes and hydroxycinnamic acids and products from an incomplete monolignol biosynthesis, such as hydroxycinnamaldehydes, are found in lignins of wild-type plants (Baucher et al., 1996; Ralph et al., 1997, 2008a; Sibout et al., 2002; Boerjan et al., 2003).

Lignin composition can be steered via genetic engineering. For example, F5H-deficient plants produce lignin composed almost entirely of G units rather than the normal complement of both S and G units (Meyer et al., 1998; Marita et al., 1999). Conversely, F5H up-regulation can lead to plants with extremely high proportions of S units (Meyer et al., 1998; Marita et al., 1999; Franke et al., 2000; Stewart et al., 2009). Plants deficient in COMT produce elevated amounts of 5-hydroxyconiferyl alcohol, an alternative monomer that has a high propensity to undergo β-O-4 coupling, producing novel benzodioxane structures within the lignin (Marita et al., 2001; Ralph et al., 2001a,b; Jouanin et al., 2004; Morreel et al., 2004b; Lu et al., 2010). These inter-unit linkages are below or close to the detection limit in wild-type plants (Atanassova et al., 1995; Morreel et al., 2004b; Lu et al., 2010; Huis et al., 2012). In an extreme case, an Arabidopsis comt mutant with concomitant F5H overexpression produced a lignin with over 90% of its units linked by benzodioxane structures (Vanholme et al., 2010b; Weng et al., 2010). The coupling of alternative monomers into lignins was also enhanced in plants with reduced CAD or CCR activity. Dicots that are deficient in CAD accumulate lignin units derived from hydroxycinnamaldehydes (Kim et al., 2000, 2002, 2003; Lapierre et al., 2004; Ralph et al., 2004b; Sibout et al., 2005; Leplé et al., 2007) and plants deficient in CCR are characterized by lignins containing small amounts of ferulic acid-derived units (Dauwe et al., 2007; Leplé et al., 2007; Mir Derikvand et al., 2008; Ralph et al., 2008b). The observation that plants readily incorporate alternative monomers to form lignins with altered physicochemical properties opens up the possibility of bioengineering various phenolic pathways to produce phenolic monomers that can be exported to the cell wall to create new types of lignins designed for efficient industrial processing of biomass.

IV. Alternative lignin monomers for biofuel applications

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

Alternative lignin monomers must meet certain criteria to confer increased susceptibility of biomass to pretreatments. As mentioned above, the monomer must meet the minimal requirement for a molecule to be radicalized and to couple into the lignin polymer. Given all available data about molecules that are polymerized into lignin, both in vivo in wild-type and genetically engineered plants and in vitro in synthetic lignins (dehydrogenation polymers (DHPs)), this minimal requirement is the presence of a phenolic function (Harkin, 1967; Lu & Ralph, 1999, 2002, 2008; Boerjan et al., 2003; Morreel et al., 2004a; Ralph et al., 2004b; Ralph, 2006; Vanholme et al., 2010a). As a consequence of incompatibilities in radical coupling reactions, p-hydroxyphenyl moieties fare less well than guaiacyl or syringyl moieties, at least when incorporating into guaiacyl-syringyl lignins, but other phenolics have not been well studied. Phenolic molecules with an accessible β-position (i.e. a side-chain conjugated to the phenol) allowing for so-called ‘endwise’ β-O-4 coupling are also considered ideal (Ralph, 2006). Although the ability to efficiently cross-couple with monolignols is a prerequisite, the ultimate utility of alternative monomers is determined by their abilities to lessen the inherent inhibitory effects of lignin on cell wall saccharification or to render lignin easier to remove by chemical pretreatments. For this purpose, we envision that five types of phenolics could prove useful as alternative monomers for lignification. These include (1) monomers that directly produce a readily cleavable functionality in the polymer, (2) hydrophilic monomers, (3) difunctional monomers and monomer conjugates linked via a readily cleavable functionality, (4) monomers that minimize lignin–polysaccharide cross-linking and (5) monomers that give rise to shorter lignin polymers. As described in the one but last paragraph of this Section (IV), alternative monomers can possess one or several of these characteristics, which may be of value for enhancing the conversion of biomass into fermentable sugars. In addition to the above-mentioned restrictions, molecules composed of only carbon, oxygen and hydrogen are most attractive as alternative monolignols for biofuel applications. This is because molecules containing other elements, such as nitrogen and phosphorous, although they might result in added-value degradation products from lignin, would probably increase the need to provide plants with increased nitrogen- and phosphorus-containing fertilizer for their biosynthesis. In some cases, greater fertilizer use might be justified, but as a general rule this would undesirably increase the financial and environmental cost of growing biomass for biofuel production.

Monomers that directly produce a readily cleavable functionality in the lignin polymer Ferulic acid (11G) is an example of a monomer that produces a cleavable functionality upon incorporation in the lignin polymer, in this case an acetal that can be readily cleaved under mildly acidic conditions (Ralph et al., 2008b). By β-O-4 coupling of ferulic acid with the growing polymer, decarboxylation to form a side-chain truncated unit and β-O-4 coupling again with a monomer or the polymer, an acetal is formed – the reaction mechanism is depicted in Fig. 3. Ferulic acid is incorporated into the lignin of CCR-down-regulated plants (Dauwe et al., 2007; Leplé et al., 2007; Mir Derikvand et al., 2008; Ralph et al., 2008b). Heavily CCR-down-regulated plants (with lower lignin contents) are compromised, usually having stunted growth and collapsed vessels, but it is unlikely that this is solely attributable to the incorporation of low amounts of ferulic acid into the lignins – it is probably a result of the reduced lignin contents and other metabolic and structural changes. Finding the means to incorporate ferulic acid or other related monomers might still prove to be fruitful. In principle, other monomers with carboxylic acid side-chains could form acetals too, but they must be capable of undergoing the double β-O-4 coupling reactions. An alternative strategy to introduce acetal-type inter-unit linkages in the lignin polymer is via alternative lignin monomers that bear a hydroxyl or ether function at the β-position (as in compounds 16, 24, 26, 32, 48, 5759 and 79). In this case, β-O-4 coupling will directly lead to the formation of an acetal.

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Figure 3. Formation of acetal functionalities in lignin via the incorporation of ferulic acid. Aromatization of the quinone methide intermediate of ferulic acid β-O-4 coupled to a phenolic end group (of a generic guaiacyl lignin unit here) is via decarboxylation of the Cγ carboxylic acid function. The newly generated α-β double bond allows the ferulic acid-derived unit to enter a second β-O-4 coupling reaction via its β-position, ultimately creating the acetal functionality after the usual re-aromatization of the quinone methide. The dotted arrows indicate the positions where lignification can proceed.

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Hydrophylic monomers Incorporation of hydrophilic monomers into lignin could enhance the penetration and, therefore, the hydrolysis of the lignocellulosic biomass by saccharifying enzymes, even without pretreatment. Lignin hydrophobicity could be modulated by the incorporation of phenolics with extensive side-chain or aromatic ring hydroxylation (e.g. monomers such as guaiacylglycerol (4G)) or substitution with hydrophilic groups (e.g. feruloyl quinate (28G), feruloyl glucose (35G) or isoconiferin (34G)). Hydrophilic groups attached by ester and glycosidic linkages would probably be cleaved under alkaline or acidic conditions; thus, the value of alternative monomers containing such groups would be diminished under many pretreatment conditions. However, hydroxyl or carboxylic acid groups remaining after pretreatment could enhance the extractability of lignin fragments into aqueous media, aiding delignification. The effect of hydrophilic lignin on plant growth and development is not obvious at this stage. Hydrophobic lignin may facilitate water transport in vessel elements but, while this has often been speculated to be the case, this requirement has never been demonstrated experimentally. If hydrophobic lignins are indeed required for water transport, hydrophilic monomers could be targeted toward fibers or other nonconducting tissues.

Difunctional monomers and monomer conjugates linked via a readily cleavable functionality Introducing monomers or conjugates with compatible phenolic groups at both ‘ends’ of the molecule allows lignification to proceed in both directions to incorporate the monomer. If coupling at the side-chain β-position is possible, such monomers can form important branch points in the polymer. More importantly, if these units contain bonds that are readily cleaved during anticipated processing, they introduce labile groups into the polymer backbone, allowing it to be readily ‘unzipped’; delignification can therefore be achieved under less stringent conditions, releasing the polysaccharides with lower energy requirements and higher yields for enzymatic saccharification or other uses. A wide array of phenolics meet these criteria, but the simplest examples include coniferyl ferulate (7GG) (Grabber et al., 2008; Ralph, 2010) linked by an alkali- and acid-labile ester bond, 3-methoxytyramine ferulate (8GG) linked by a somewhat acid-labile amide bond and compounds such as disinapoyl glucose (78SS) and diferuloyl sucrose (81GG), where two phenolic units are linked by labile ester bonds to a core ‘spacer’. All possess guaiacyl or syringyl type moieties compatible with oxidative coupling with monolignols and are therefore expected to become integral components of lignin. Monomers with one or more p-hydroxyphenyl moieties might also be utilized, but any molecule with a lower propensity to undergo radical coupling might hinder full incorporation into lignin. ortho-Diphenol (catechol) and 1,2,3-triphenol (pyrogallol) monomers are also prevalent phenolic metabolites and can profitably be considered. Unlike monomers that upon incorporation render lignin more hydrophilic, such difunctional monomers could maintain the hydrophobicity of lignin that may be required for water transport or plant defense responses. Their incorporation into hydrophobic polymers should also shield their ester, amide or glycosidic linkages from attack by hydrolytic enzymes produced by pathogenic fungi or bacteria. Thus, these monomer substitutes are a way of introducing ‘zips’ into lignin that can be readily cleaved during processing while maintaining the functional properties of lignin required by plants.

Monomers that minimize lignin–polysaccharide cross-linking Monomers that minimize lignin–polysaccharide cross-linking should enhance the inherent degradability of lignocellulosic biomass by saccharifying enzymes. The adverse effects of lignin–polysaccharide cross-linking on wall polysaccharide digestibility have been demonstrated in grasses (Grabber et al., 1998a,b, 2000, 2002; Ralph et al., 1998; Hatfield et al., 1999; Grabber, 2005), where cross-linking is mediated by ferulates on arabinoxylans (Fry, 1986; Fry & Miller, 1989; Yamamoto et al., 1989; Ralph et al., 1998, 2004a; Hatfield et al., 1999; Ralph, 2010). As a result of this finding, efforts are underway to attack this cross-linking mechanism by targeting the putative transferase that acylates arabinoxylans with ferulate (Yoshida-Shimokawa et al., 2001; Mitchell et al., 2007; Buanafina, 2009; Piston et al., 2010). In grasses and all other plants, lignin–polysaccharide cross-linking also apparently results from polysaccharides adding to the quinone methide intermediates produced during lignification and such cross-linkings also appear to limit fiber saccharification (Grabber et al., 2003; Grabber & Hatfield, 2005). During β-O-4 coupling, re-aromatization of the quinone methide intermediate occurs mainly via the proton-assisted nucleophilic attack of water at the α-position (Fig. 2). Actually, any nucleophile present in the neighborhood can participate in the reaction instead of water, an example being hemicellulosic alcohol (or acid) groups (Ralph et al., 2004b; Simmons et al., 2010). In the latter case, lignin becomes covalently linked to the hemicellulosic network, rendering its removal more difficult. Quantifying these benzyl ether and ester cross-links in cell walls is problematic and limited to a small fraction of lignin that can be extracted from cell walls (Balakshin et al., 2008, 2011). Nevertheless, judicious choice of lignin monomers can minimize/eliminate this cross-linking mechanism in plants. Alternative monomers with ortho-diphenol structures, such as caffeyl alcohol (1C), 5-hydroxyconiferyl alcohol (1F) or epicatechin (73C), and with 1,2,3-triphenol structures, such as ethyl gallate and epigallocatechin (73L), readily form benzodioxane structures; rapid internal trapping of the quinone methide, which is produced following a monolignol's β-O-4 coupling with such o-diphenols, precludes any possibility of benzyl ether and ester cross-linking of hemicellulosic alcohol or acid groups with those units (Fig. 4a). Lignins derived solely from caffeyl alcohol have been discovered in seed coats (Chen et al., 2012); lignins incorporating caffeyl alcohol have been observed in CCoAOMT-deficient gymnosperm cell cultures (Wagner et al., 2011). Lignins incorporating 5-hydroxyconiferyl alcohol derive from various COMT-deficient dicots and monocots (Van Doorsselaere et al., 1995; Lapierre et al., 1999; Marita et al., 2001; Ralph et al., 2001a,b; Morreel et al., 2004b; Lu et al., 2010; Vanholme et al., 2010b; Weng et al., 2010). Where they have been tested, the cell walls from these materials have enhanced digestibilities (Guo et al., 2001a,b; Fu et al., 2011). The effect of introducing rosmarinic acid (49CC) into lignins has recently been evaluated (see Section V) and found to be effective at improving saccharification, even without pretreatment (Tobimatsu et al., 2012). Other candidate alternative monolignols, such as guaiacyl butenol, provide a side-chain-based intramolecular pathway to trap lignin quinone methides that are formed from the monomer itself following β-O-4, β-β, or β-5 coupling (e.g. 25, 27, 4347). Trapping by the δ- or ε-hydroxyl group would result in tetrahydrofuran (oxolane) or tetrahydropyran (oxane) structures (Fig. 4b).

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Figure 4. An interesting way to avert lignin–polysaccharide cross-linking during β-O-4 coupling. (a) The quinone methide intermediate formed via β-O-4 coupling is trapped via an intra-molecular reaction of the novel C5-phenolic function, thereby precluding the possibility of inter-molecular nucleophilic attack by hemicellulose (indicated by a crossed arrow). (b) Similarly, the intra-molecular trapping via an alcoholic function on the δ (n = 1) or ε (n = 2) position of suitable alternate lignin monomers also avoids the inter-molecular nucleophilic attack by hemicellulose (crossed arrow). The dotted arrows indicate the positions where lignification can proceed.

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Monomers that give rise to shorter lignin polymers Decreasing the average length of the lignin polymers should also enhance the extractability of lignin. Alternative monomers that, upon polymerization, end up as an aliphatic or phenolic end group, rather than becoming an internal unit, might serve as polymerization initiation or termination monomers, respectively. Augmenting their availability at the lignification site might lead to a higher rate of lignin initiation or termination reactions, probably yielding a higher number of shorter lignin molecules, that is, lignins with lower degrees of polymerization. Monomers that initiate polymerization, for example dihydroconiferyl alcohol (2G) (Ralph et al., 1997) or benzenoids (9, 20), enter only into a single coupling reaction, that is, at the 4-O- or 5-position, thus consuming the phenolic function during initial coupling. A proof of principle is apparent in a study in which the bacterial hydroxycinnamoyl-CoA hydratase-lyase (HCHL) was expressed in Arabidopsis (Eudes et al., 2012). p-Hydroxybenzaldehyde (9H) and p-hydroxybenzoate (20H) were incorporated into the lignin of HCHL engineered plants, which resulted in lignin with a reduced molecular weight and an improved saccharification of pretreated stem cell walls (Eudes et al., 2012). Importantly, total lignin and biomass yield were not affected. Alternatively, co-polymerization with a monomer possessing a rather high oxidation potential enhances the termination of polymerization and reduces the average length of the lignin polymers. Among the traditional monolignols, p-coumaryl alcohol has the highest oxidation potential (Syrjänen & Brunow, 1998). Consequently, phenolic profiling (see Section VI) of flax (Linum usitatissimum) stem tissues showed that H units were preferentially phenolic end groups of oligolignols (Huis et al., 2012) and thioacidolysis shows that a high fraction of H-units are free-phenolic (Lapierre et al., 1988; Pitre et al., 2007; Lapierre, 2010). In part, this is thought to be attributable to radical transfer (to more stable units) in the radical-limited system – a similar phenomenon occurs with p-coumarate esters, which also remain uncoupled (Ralph et al., 1994a, 2004a; Hatfield et al., 2008). Furthermore, lignins from transgenic or mutant plants that contain a high level of H units are of lower molecular weight (Ziebell et al., 2010) and are enriched in β-5 and β-β linkages as compared with wild-type lignin (Ralph et al., 2006; Wagner et al., 2007). The relative decrease in β-O-4 coupling reactions, which are the main lignin polymer elongation reactions, suggests the presence of shorter lignins in these transgenic and mutant plants.

Finally, alternative monomers combining several of the above-mentioned mechanisms for altering lignin properties would be especially attractive as genetic engineering targets. For example, disinapoyl glucose (78SS) features two readily incorporated sinapate moieties attached to a hydrophilic moiety by readily cleavable ester linkages. Rosmarinic acid (49CC, noted above), epigallocatechin gallate (74LL) and dicaffeoyl quinate (72CC) all possess two cross-link-preventing o-diphenol functionalities connected by labile ester linkages, and in addition the latter molecule also contain a hydrophilic moiety. Finally, gallotannins (86) contain multiple pyrogallol units connected by labile ester linkages and the extensive hydroxylation of this molecule could make an altered lignin more hydrophilic.

To exploit the biochemical variation of potential alternative monolignols, we screened the SciFinder Scholar database (CAS; http://www.cas.org/products/scifindr/index.html) for plant metabolites that fulfill one of the above-mentioned criteria, that is, the metabolites must have a phenolic function, upon incorporation in the lignin polymer they must render it easier to degrade by chemical pretreatments, and they must be composed of carbon, oxygen and hydrogen only. Over 160 plant metabolites satisfying the above-mentioned criteria for promising alternative lignin monomers were identified (Fig. 5). Given the fact that only a small proportion of plant metabolites have been characterized thus far, the list of candidate alternative lignin monomers will grow further as targeted phenolic profiling studies are being carried out. Bacteria and fungi also contain attractive enzymatic activities for other types of phenolic compounds that might be exploited for modifying plant lignin (Masai et al., 2007; Merali et al., 2007; Eudes et al., 2012). In principle, such lignin modifications can be accomplished in lignocellulosic biomass crops by cloning and expressing alternative monomer biosynthetic pathways in conjunction with appropriate tissue-specific promoters. In vitro testing is necessary to cut down this large number of the ‘candidate’ alternative lignin monomers (Grabber et al., 2008, 2010; Tobimatsu et al., 2012). Such in vitro tests should at least include the co-polymerization of the proposed alternative monomers with traditional monolignols in DHPs or biomimetic in vitro lignified cell walls followed by analysis of the DHPs and lignified cell walls for the effects of pretreatment on delignification and by analysis of the lignified cell walls for the saccharification potential. In addition, DHPs and in vitro lignified cell walls should be analyzed for the incorporation of the alternative monomer by phenolic profiling (Morreel et al., 2004a) and nuclear magnetic resonance (NMR) (Grabber et al., 2010; Tobimatsu et al., 2012). These tests are to determine whether the proposed alternative lignin monomers do incorporate in sufficient amounts in the lignin polymer, and also to what extent the efficiency of incorporation of the monomers depends on the competing phenolics present in the cell wall matrix.

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Figure 5. Compounds found within the plant kingdom that (potentially) satisfy criteria for alternative lignin monomers. (a) Monomers that have already been authenticated or implicated in lignification. (b) Alternative monomers that, upon incorporation into the lignin polymer, potentially make the lignin more susceptible to biomass pretreatment. Aromatic ring units are all phenols, invariably p-hydroxy-aryl units here. Substituents are labeled R/R′ for the ‘first’ aromatic ring, X/X′ for the second, Y/Y′ for the third and Z/Z′ for the fourth. In all cases, the descriptor notation uses the compound number followed by the defined rings, in the order described as: H (p-hydroxyphenyl), G (guaiacyl), S (syringyl), C (caffeyl), F (5-hydroxyguaiacyl), or L (gallyl) with A being used for any or all (generic) units. The convention is illustrated with compounds 7 as follows: 7SG, sinapyl ferulate (R = R′ = OMe, X = OMe, X′ = H); 7AH, general hydroxycinnamyl p-coumarate (R,R′ = H/OH/OMe; X = X′ = H). Where necessary, other variable substituents are used (P, T) and a variable single or double bond in some structures is designated with one solid and one dashed bond line. The names of the compounds, the plant species in which the compounds are found and references to the literature are available in Supporting Information Notes S1.

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V. Candidate alternative monolignols in biomimetic systems

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

Plant genetic engineering studies will, of course, ultimately determine the feasibility and utility of modified lignins and their compatibility with plant growth and development. The engineering of plants will, however, be much more efficient if in vitro DHP studies and biomimetic cell wall lignification studies are first carried out to test the compatibility of the various monolignol substitutes with lignification, and to determine their potential effects on cell wall delignification and saccharification. In the latter case, for example, isolated maize (Zea mays) cell walls containing bound peroxidases are stirred in water or buffer solutions and artificially lignified by slowly adding separate solutions of lignin precursors and dilute hydrogen peroxide (Grabber et al., 1996b, 1998c). Candidate monolignol substitutes are typically added with normal monolignols to comprise 35–45% of the weight of the precursor mixture, potentially yielding a shift in lignin composition comparable to that observed in some mutant or transgenic plants with altered lignin biosynthesis. The effects of alternative monomer on lignin formation and the susceptibility of cell walls to chemical pretreatments and saccharification are compared with lignified controls prepared with normal monolignols (Grabber et al., 2008, 2010, 2012; Tobimatsu et al., 2012).

Thus far the work with the DHPs and lignified cell wall model systems has mainly focused on difunctional monomers or monomer conjugates linked via a readily cleaved functionality and monomers that minimize lignin–polysaccharide cross-linking. For instance, incorporation of coniferyl ferulate (7GG) facilitated lignin depolymerization and increased lignin extractability by up to twofold in aqueous NaOH, providing an avenue for producing fiber with less lignin contamination and delignifying at lower temperatures or lower chemical consumption (Grabber et al., 2008; Ralph, 2010).

As alluded to in Section IV, more recent model studies have demonstrated the utility of rosmarinic acid (49CC), an ester conjugate with two catechol moieties (Tobimatsu et al., 2012). In in vitro DHP experiments, rosmarinic acid readily underwent peroxidase-catalyzed copolymerization with monolignols to form polymers with benzodioxane inter-unit linkages, suggesting that fewer lignin–carbohydrate cross-links could be formed via lignin quinone methide intermediates. Incorporation of rosmarinic acid permitted extensive depolymerization of in vitro lignified cell walls by mild alkaline hydrolysis, via cleavage of ester linkages within the rosmarinic acid moiety (in the lignin) itself. Copolymerization of rosmarinic acid with monolignols modestly depressed lignification of cell walls and promoted subsequent cell wall saccharification by fungal enzymes after mild alkali pretreatment. Incorporating rosmarinic acid also improved cell wall saccharification by fungal enzymes and by rumen microflora even without alkaline pretreatments, possibly by modulating lignin hydrophobicity and/or limiting cell wall cross-linking.

In other studies (Grabber et al., 2010, 2012), epigallocatechin gallate (74LL), epicatechin gallate (74CL), epicatechin vanillate (74CG), epigallocatechin (73L), galloyl hyperin (77CL) and pentagalloyl glucose (86) formed wall-bound lignin at moderate to high concentrations and their incorporation increased in vitro ruminal fiber fermentability by 20 to 33% relative to lignified controls. By contrast, ethyl gallate and corilagin (87) severely depressed lignification and increased fermentability by c. 50%. Thus, addition of these units probably acted indirectly to improve fermentability through severely reducing lignin content. Regardless of the mechanism, ethyl gallate and corilagin would probably be of limited value as target monomers, because they severely disrupted cell wall lignification. Such reductions in lignin content, also already attained by down-regulating enzymes in the general phenylpropanoid and monolignol pathway, often reduce plant fitness (Gallego-Giraldo et al., 2011b; Voelker et al., 2011). Improvements in fermentability with flavan-3-ols were associated with increased hydroxylation, but this response was not necessarily caused by increased lignin hydrophilicity because flavonol glycosides and gallate esters with more extensive hydroxylation (e.g. hyperoside (76C, P=Gal), galloyl hyperin (77CL), and pentagalloyl glucose (86)) had less pronounced effects on cell wall fermentability. Among flavan-3-ols (73), gains in cell wall fermentability were related to the presence of gallate and pyrogalloyl units. Furthermore, the copolymerization of monolignols with epicatechin gallate (74CL), epigallocatechin gallate (74LL) and, to a lesser degree, pentagalloyl glucose (86) reduced the proportion of ferulates that underwent cross-linking with lignin (Grabber et al., 2010, 2012). These reductions in ferulate–lignin cross-linking should contribute to the improved cell wall fermentability (Grabber et al., 2009). From these experiments, epigallocatechin gallate (74LL) appeared a promising target for incorporation into lignin for improving the delignification and saccharification of biomass crops (Grabber et al., 2012).

In addition to studying the incorporation of the alternative monomers in lignin and the resulting cell wall properties, the biomimetic systems are also interesting systems with which to reveal peroxidase inactivation. For instance, partial substitution of coniferyl alcohol with coniferyl ferulate (7GG) tended to accelerate peroxidase inactivation and reduce cell wall lignification and cross-linking of feruloylated xylans to lignin (Grabber et al., 2008). Analogous effects were seen with sinapyl p-coumarate (7SH), which is, however, a monomer conjugate that is heavily implicated as being successfully incorporated into grass lignins (Ralph et al., 1994a; Grabber et al., 1996a; Lu & Ralph, 1999, 2008; Hatfield et al., 2009; Ralph, 2010; Withers et al., 2012). Notably, because non-bound apoplastic peroxidases were removed before artificial lignification in these experiments, peroxidase inactivation would be more markedly manifested in the model system than in plants. Nevertheless, such observations need to be tracked when genetically altered plants are developed.

VI. From phenolic profiling to lignomics

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

Phenolic profiling is a technique ideally suited for identifying new alternative lignin monomers and pathway intermediates, and in addition for verifying their incorporation into in vivo or in vitro lignins and for monitoring plant responses to phenolic pathway engineering. Basically, this technique includes the identification of phenolic metabolites by reversed-phase (ultra) high-pressure liquid chromatography coupled to mass spectrometry (UHPLC-MS). This technique permits the detection and quantification of essentially every small phenolic molecule with a molecular mass below 1500 Da at concentrations in the micromolar to millimolar range. An important bottleneck in the field of phenolic profiling is the structural characterization of detected compounds in the UHPLC-MS chromatogram. The identification of unknown compounds might be revealed via NMR upon compound purification (Nakabayashi et al., 2009). However, purification is often impossible because of co-eluting impurities, compound degradation or extremely low compound concentrations. In such cases, retention times and MS data (including high mass accuracy MS, from which the chemical formula can be determined, and MSn spectra) are often the only means for resolving the structure.

Concerning the identification of alternative monomers, the study and identification of metabolites that make up the lignome are of special interest. The lignome is defined as the ensemble of all phenolics for which the biosynthesis is co-regulated with lignin biosynthesis and includes the oligolignol (small lignin polymers) pool (Morreel et al., 2010a). The oligolignol pool in model species such as poplar and Arabidopsis has been well characterized (Morreel et al., 2004a, 2010b). Although the main lignome components have been elucidated in these species, the identity of many compounds, which are potentially useful as alternative monomers, remains unknown. Important progress in the structural elucidation of oligolignols was made when it was discovered that various lignin units (i.e. G and S) and linkage types (i.e. β-O-4, β-5 and β-β) show characteristic MS fragmentation patterns (Morreel et al., 2010a,b). Given the characteristic MS fragmentation, an algorithm was developed allowing the sequencing of oligolignols, that is, determining the order of the monolignols in the polymer, as well as the bonds connecting them (Morreel et al., 2010a). This approach greatly simplifies the identification of oligolignols and aids the discovery of alternative lignin units. The algorithm allowed the complete sequencing of 36 of the 134 oligolignols present in poplar xylem extracts; for the remaining compounds only partial sequences were obtained because of the presence of unidentified units (bonds or monomers) (Morreel et al., 2010a). These partially identified compounds are therefore prime targets for further structural elucidation, which can help in finding new monomers and elucidating their coupling propensities. For example, an MS fragmentation pattern associated with the presence of an arylglycerol unit was frequently encountered when sequencing small poplar lignin polymers (Morreel et al., 2010a). Four G(β-O-4)S(β-5)Gglycerol isomers were detected. Intriguingly, the arylglycerol was always β-5-, but never β-O-4-coupled, suggesting that the glycerol monomer has different coupling propensities than coniferyl alcohol. An algorithm for annotating oligolignols is particularly useful for studying the lignome of plants that are genetically modified to synthesize alternative monomers and for sequencing the oligolignol fraction of DHPs made with alternative monomers. Thus far, the lignome of many taxa such as grasses and gymnosperms remains largely unexplored and may contain new types of alternative lignin monomers that may prove to be useful for modifying lignin properties.

Our Scifinder-based search resulted in over 160 reported plant metabolites that fit the criteria to be used as alternative monomers (see Section IV). Nevertheless, as only a fraction of the secondary metabolites are currently known (Hadacek, 2002; Saito & Matsuda, 2010), many more alternative monomers are waiting to be identified together with their biosynthetic pathways. Structural elucidation of plant phenolics is thus of prime interest. Given the above-mentioned limitations for isolating and identifying phenolics, many researchers have invested heavily in annotating metabolites by MS2 spectral information, often leading to the construction of species-specific databases (Moco et al., 2006; Farag et al., 2007; Böttcher et al., 2008; Matsuda et al., 2009, 2010, 2011). In addition to the use of MS2 spectral libraries, information about other phenolic metabolites within the same plant extract can be used for structural elucidation. This is because the substrates and products of well-known enzymatic conversions, such as methylation or glycosylation, are often observed in the same chromatogram (Iijima et al., 2008). To search for these ‘candidate substrate product pairs’ (CSPPs), an algorithm was developed that uses the input of mass differences corresponding to particular enzymatic conversions (K. Morreel et al., unpublished). This tool is expected to significantly accelerate the identification of alternative monomers and their pathway intermediates.

VII. Phenolic pathway engineering towards alternative monolignols

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

According to the strategy outlined in this paper, pathway engineering in lignocellulosic biomass crops should entail the biosynthesis of the most promising alternative monomers and their incorporation in the lignin polymer. In some cases, the cloning of a biosynthetic pathway might need to be combined with the mutation of (an) endogenous gene(s) to re-direct the flux into the newly established pathway to reduce the competition for substrates at the level of biosynthesis of the monomers, their transport and their polymerization.

Pathway engineering of lignocellulosic biomass crops implies the cloning of biosynthetic pathways. Unfortunately, for most of the candidate alternative lignin monomers depicted in Fig. 5, the biosynthetic enzymes are unknown. Nevertheless, the occurrence of these candidate alternative lignin monomers in plant species opens up the possibility of elucidating their biosynthetic pathways. The few exceptions whose pathways have been well elucidated are sinapate esters (29S, 35S, 78SS) (Nair et al., 2004; Niggeweg et al., 2004; Fraser et al., 2007; Sinlapadech et al., 2007; Liu, 2010) and rosmarinic acid (49CC) (Ellis & Towers, 1970; Matsuno et al., 2002; Petersen et al., 2009; Liu, 2010). Recently, enzymes capable of synthesizing ferulate conjugates (7AG) have been obtained (Wilkerson et al., 2011), and the putative p-coumarate analog in grasses has been identified (Withers et al., 2012). In addition, an enzyme that is likely to catalyze the first committed step towards phenylbutanoids has been isolated from rhubarb (Rheum palmatum; Abe et al., 2001). This enzyme, benzalacetone synthase (BAS), belongs to the polyketide synthase family and makes the phenylbutanoid p-hydroxybenzalacetone from malonyl-CoA and p-coumaric acid (Shimokawa et al., 2010). For some other alternative lignin monomers that are found in plants, an enzymatic activity might be conjectured. For example, for the biosynthesis of monomer conjugates linked by ester or amide functionalities, at least one acylation reaction might be suggested to take place. Acyltransferases, currently known to be responsible for the transfer of hydroxycinnamic acids to recipient molecules, fall into three families. They all generally depend on CoA- or glucose-conjugated phenylpropanoids (Steffens, 2000; D'Auria, 2006; Kang et al., 2006; Liu, 2010) and they produce O- or N-linked products depending on the specific acyltransferase. Hydroxycinnamoyl-CoA-dependent BAHD (BEAT, AHCT, HCBT and DAT – after the first four members characterized) superfamily acyltransferases and the 1-O-acylglucose ester-dependent serine carboxypeptidase-like proteins (SCPLs) catalyze both O- and N-transacylation, while the general control non-depressible 5 (GCN5)-related N-acyltransferases (GNAT) catalyze only N-transacylation (Milkowski & Strack, 2004; Vetting et al., 2005; D'Auria, 2006). Searching BAHD, SCPL and GNAT-coding expressed sequence tags (ESTs) in the tissue where the alternative lignin monomer is also found might therefore result in candidate genes that can further be tested via feeding assays and other reverse genetic tools. In addition, phenolic profiling can also help in pathway elucidation, as it might identify molecules that are structurally related to alternative lignin monomers. Given a pool of structurally related molecules, enzymatic conversions can be proposed for their synthesis, which again enables the search for ESTs.

The exact subcellular location of the biosynthesis and storage of most candidate alternative monomers is unknown. Several monomers might be recognized in planta as ‘true’ monomers, because of their structural similarity to them, and therefore transported by default to the cell wall. That this is indeed the case is proven by the increase of ferulic acid, 5-hydroxyconiferyl alcohol and cinnamaldehydes in the lignin of CCR-, COMT- and CAD-deficient plants, respectively (Atanassova et al., 1995; Ralph et al., 2001b; Kim et al., 2003; Morreel et al., 2004b; Sibout et al., 2005; Dauwe et al., 2007; Mir Derikvand et al., 2008), and by the incorporation of hydroxybenzaldehyde and hydroxybenzoate in HCHL engineered plants (Eudes et al., 2012). However, alternative lignin monomers that bear a glucose, malate or quinate moiety are more prone to be stored in the vacuole, both in their endogenous species and when produced in lignocellulosic biomass crops (Wink, 1997; Bartholomew et al., 2002; Dean et al., 2003). This subcellular localization has been proven via vacuolar isolation followed by phenolic profiling for sinapoyl glucose (35S) and sinapoyl malate (29S) in Raphanus sativus and for chlorogenic acid (28C) in Catharanthus roseus (Sharma & Strack, 1985; Ferreres et al., 2011). For these monomers, rerouting to the cell wall is needed.

An important issue with genetic engineering of the lignin pathway is that plants with perturbed lignification often show unwanted pleiotropic effects (Chen & Dixon, 2007; Li & Chapple, 2010; Li et al., 2010; Vanholme et al., 2010b; Gallego-Giraldo et al., 2011b). Although the exact cause(s) of these effects is not fully known, they have been attributed to impaired water transport (Jones et al., 2001; Franke et al., 2002), altered levels of dehydrodiconiferyl alcohol glucoside (DCG) that might influence cell proliferation and expansion (Abdulrazzak et al., 2006; Li & Chapple, 2010), the lack of cell wall integrity or the release of elicitors that trigger responses at the level of gene expression (Li & Chapple, 2010; Seifert & Blaukopf, 2010), and the accumulation of phenylpropanoid pathway intermediates or products (Vanholme et al., 2010b; Gallego-Giraldo et al., 2011a). In this respect, the recent finding that reduced growth in Arabidopsis plants with impaired lignin biosynthesis is correlated with increased concentrations of salicylic acid is of particular interest (Gallego-Giraldo et al., 2011a; Lee et al., 2011). While HCT-down-regulated Arabidopsis plants were severely affected in growth, reducing salicylic acid biosynthesis by crossing in a mutation in the isochorismate synthase (ICS) gene could partially restore plant growth (Gallego-Giraldo et al., 2011a). This is an interesting observation because it shows that growth defects are (partly) caused by the unintended accumulation of phenolics, and that these defects can be (partly) complemented by redirecting the flux. How the phenolic steady state will react to pathway engineering will differ case by case and is difficult to predict. In the ideal case, the pathway engineering would only alter lignin biosynthesis and not plant growth and performance, but if it does, possible solutions are to engineer the new pathway in specific cell types only; for example, the use of fiber-specific promoters might restrict the altered lignin to be deposited in supportive but not conductive tissues. Alternatively, suppressor screens might be carried out to identify the molecular causes of the pleiotropic effects and to identify the genes to mitigate the unwanted phenotypes (Halpin, 2010).

It has been noted that the main building blocks of lignin seem relatively conserved over different taxonomic clades (Weng & Chapple, 2010). However, this observation may result from the fact that only few plant species have been investigated with current analytical (including powerful NMR) methods able to detect and identify ‘novel units’ and often the novelty is missed by failing to recognize that the lignins may have derived from, for example, acylated monolignols. There are numerous examples of lignins, in both ‘natural’ and transgenic plants, being partially to substantially derived from monomers that are not the three classical monolignols; these include the conjugates: monolignol acetates in many plants, monolignol p-coumarates in all grasses, monolignol p-hydroxybenzoates in Salix, Populus, and Palmae; but also catechol-type monomers such as caffeyl and 5-hydroxyconiferyl alcohol, double-bond-reduced monomers such as dihydroconiferyl alcohol, the hydroxycinnamaldehydes, etc. – as all noted in Sections I and III (Ralph et al., 1997, 2008a; Lu & Ralph, 1999; Lu et al., 2004; Morreel et al., 2004a; Vanholme et al., 2008; Stewart et al., 2009; Ralph, 2010; Chen et al., 2012; del Río et al., 2012). It is currently not clear whether these nontraditional lignins confer different properties to the plants that have made evolution to select these lignins over the classical HGS-type lignins, but they clearly have their niches. It also will remain to be tested how plants respond to the incorporation of high amounts of the proposed alternative monomers in terms of growth and development, biotic and abiotic stresses and mechanical properties. In cases such as introducing readily cleavable bonds into the backbone of the lignin polymer by utilizing various ester conjugates as monomer replacements, however, it may be sufficient to introduce these at reasonably low levels that will not greatly alter the structural properties but will render the polymer dramatically easier to cleave into smaller fragments during pretreatments. Evidence suggests that modifications such as some of the ones proposed here could lead to significantly improved plant materials (from the point of view of biomass conversion) and that some of the suggested modifications are likely to prove game-changing for plant cell wall utilization. Interesting times are ahead as researchers strive to introduce some of these new traits into important biomass crops.

Acknowledgements

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

The authors thank Annick Bleys for help in preparing the manuscript. This work was supported by grants from the Multidisciplinary Research Project ‘Biotechnology for a sustainable economy’ of Ghent University and from Stanford University's Global Climate and Energy Project (‘Towards New Degradable Lignin Types’ and ‘Efficient Biomass Conversion: Delineating the Best Lignin Monomer-substitutes’). R.V. is a postdoctoral fellow of the Research Foundation-Flanders. P.O. is a predoctoral fellow of the Agency for Innovation by Science and Technology of Chili.

References

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information
  • Abdulrazzak N, Pollet B, Ehlting J, Larsen K, Asnaghi C, Ronseau S, Proux C, Erhardt M, Seltzer V, Renou J-P et al. 2006. A coumaroyl-ester-3-hydroxylase insertion mutant reveals the existence of nonredundant meta-hydroxylation pathways and essential roles for phenolic precursors in cell expansion and plant growth. Plant Physiology 140: 3048.
  • Abe I, Takahashi Y, Morita H, Noguchi H. 2001. Benzalacetone synthase - a novel polyketide synthase that plays a crucial role in the biosynthesis of phenylbutanones in Rheum palmatum. European Journal of Biochemistry 268: 33543359.
  • Alejandro S, Lee Y, Tohge T, Sudre D, Osorio S, Park J, Bovet L, Lee Y, Geldner N, Fernie AR et al. 2012. AtABCG29 is a monolignol transporter involved in lignin biosynthesis. Curent Biology 22: 12071212.
  • Atanassova R, Favet N, Martz F, Chabbert B, Tollier M-T, Monties B, Fritig B, Legrand M. 1995. Altered lignin composition in transgenic tobacco expressing O-methyltransferase sequences in sense and antisense orientation. Plant Journal 8: 465477.
  • Balakshin M, Capanema E, Gracz H, H-m Chang, Jameel H. 2011. Quantification of lignin-carbohydrate linkages with high-resolution NMR spectroscopy. Planta 233: 10971110.
  • Balakshin MY, Capanema EA, Chang H-M. 2008 Recent advances in the isolation and analysis of lignins and lignin–carbohydrate complexes. In: Hu TQ, ed. Characterization of lignocellulosic materials. Oxford, UK: Blackwell Publishing, 148170.
  • Barakate A, Stephens J, Goldie A, Hunter WN, Marshall D, Hancock RD, Lapierre C, Morreel K, Boerjan W, Halpin C. 2011. Syringyl lignin is unaltered by severe sinapyl alcohol dehydrogenase suppression in tobacco. Plant Cell 23: 44924506.
  • Bartholomew DM, Van Dyk DE, Lau S-MC, O'Keefe DP, Rea PA, Viitanen PV. 2002. Alternate energy-dependent pathways for the vacuolar uptake of glucose and glutathione conjugates. Plant Physiology 130: 15621572.
  • Basset GJC, Quinlivan EP, Ravanel S, Rébeillé F, Nichols BP, Shinozaki K, Seki M, Adams-Phillips LC, Giovannoni JJ, Gregory JF III et al. 2004. Foliate synthesis in plants: the p-aminobenzoate branch is initiated by a bifunctional PabA-PabB protein that is targeted to plastids. Proceedings of the National Academy of Sciences, USA 101: 14961501.
  • Baucher M, Chabbert B, Pilate G, Van Doorsselaere J, Tollier MT, Petit-Conil M, Cornu D, Monties B, Van Montagu M, Inzé D et al. 1996. Red xylem and higher lignin extractability by down-regulating a cinnamyl alcohol dehydrogenase in poplar. Plant Physiology 112: 14791490.
  • Beejmohun V, Fliniaux O, Hano C, Pilard S, Grand E, Lesur D, Cailleu D, Lamblin F, Laine E, Kovensky J et al. 2007. Coniferin dimerisation in lignan biosynthesis in flax cells. Phytochemistry 68: 27442752.
  • Bernards MA, Lopez ML, Zajicek J, Lewis NG. 1995. Hydroxycinnamic acid-derived polymers constitute the polyaromatic domain of suberin. Journal of Biological Chemistry 270: 73827386.
  • Berthet S, Demont-Caulet N, Pollet B, Bidzinski P, Cézard L, Le Bris P, Borrega N, Hervé J, Blondet E, Balzergue S et al. 2011. Disruption of LACCASE4 and 17 results in tissue-specific alterations to lignification of Arabidopsis thaliana stems. Plant Cell 23: 11241137.
  • Binns AN, Chen RH, Wood HN, Lynn DG. 1987. Cell division promoting activity of naturally occurring dehydrodiconiferyl glucosides: do cell wall components control cell division? Proceedings of the National Academy of Sciences, USA 84: 980984.
  • Boatright J, Negre F, Chen X, Kish CM, Wood B, Peel G, Orlova I, Gang D, Rhodes D, Dudareva N. 2004. Understanding in vivo benzenoid metabolism in petunia petal tissue. Plant Physiology 135: 19932011.
  • Boerjan W, Ralph J, Baucher M. 2003. Lignin biosynthesis. Annual Review of Plant Biology 54: 519546.
  • Böttcher C, von Roepenack-Lahaye E, Schmidt J, Schmotz C, Neumann S, Scheel D, Clemens S. 2008. Metabolome analysis of biosynthetic mutants reveals a diversity of metabolic changes and allows identification of a large number of new compounds in Arabidopsis. Plant Physiology 147: 21072120.
  • Buanafina MMdO. 2009. Feruloylation in grasses: current and future perspectives. Molecular Plant 2: 861872.
  • Carvalheiro F, Duarte LC, Gírio FM. 2008. Hemicellulose biorefineries: a review on biomass pretreatments. Journal of Scientific & Industrial Research 67: 849864.
  • Chang VS, Holtzapple MT. 2000. Fundamental factors affecting biomass enzymatic reactivity. Applied Biochemistry and Biotechnology 84–86: 537.
  • Chen F, Dixon RA. 2007. Lignin modification improves fermentable sugar yields for biofuel production. Nature Biotechnology 25: 759761.
  • Chen F, Srinivasa Reddy MS, Temple S, Jackson L, Shadle G, Dixon RA. 2006a. Multi-site genetic modulation of monolignol biosynthesis suggests new routes for formation of syringyl lignin and wall-bound ferulic acid in alfalfa (Medicago sativa L.). Plant Journal 48: 113124.
  • Chen F, Tobimatsu Y, Havkin-Frenkel D, Dixon RA, Ralph J. 2012. A polymer of caffeyl alcohol in plant seeds. Proceedings of the National Academy of Sciences, USA 109: 17721777.
  • Chen Y, Zhang X, Wu W, Chen Z, Gu H, Qu L-J. 2006b. Overexpression of the wounding-responsive gene AtMYB15 activates the shikimate pathway in Arabidopsis. Journal of Integrative Plant Biology 48: 10841095.
  • Chen H-C, Li Q, Shuford CM, Liu J, Muddiman DC, Sederoff RR, Chiang VL. 2011. Membrane protein complexes catalyze both 4-and 3-hydroxylation of cinnamic acid derivatives in monolignol biosynthesis. Proceedings of the National Academy of Sciences, USA 108: 2125321258.
  • Cho M-H, Corea ORA, Yang H, Bedgar DL, Laskar DD, Anterola AM, Moog-Anterola FA, Hood RL, Kohalmi SE, Bernards MA et al. 2007. Phenylalanine biosynthesis in Arabidopsis thaliana – identification and characterization of arogenate dehydratases. Journal of Biological Chemistry 282: 3082730835.
  • Chundawat SPS, Donohoe BS, da Costa SousaL, Elder T, Agarwal UP, Lu F, Ralph J, Himmel ME, Balan V, Dale BE. 2011. Multi-scale visualization and characterization of lignocellulosic plant cell wall deconstruction during thermochemical pretreatment. Energy and Environmental Science 4: 973984.
  • Corea ORA, Ki C, Cardenas CL, Kim S-J, Brewer SE, Patten AM, Davin LB, Lewis NG. 2012. Arogenate dehydratase isoenzymes profoundly and differentially modulate carbon flux into lignins. Journal of Biological Chemistry 287: 1144611459.
  • Dao TTH, Linthorst HJM, Verpoorte R. 2011. Chalcone synthase and its functions in plant resistance. Phytochemistry Reviews 10: 397412.
  • D'Auria JC. 2006. Acyltransferases in plants: a good time to be BAHD. Current Opinion in Plant Biology 9: 331340.
  • Dauwe R, Morreel K, Goeminne G, Gielen B, Rohde A, Van Beeumen J, Ralph J, Boudet A-M, Kopka J, Rochange SF et al. 2007. Molecular phenotyping of lignin-modified tobacco reveals associated changes in cell-wall metabolism, primary metabolism, stress metabolism and photorespiration. Plant Journal 52: 263285.
  • Davin LB, Wang H-B, Crowell AL, Bedgar DL, Martin DM, Sarkanen S, Lewis NG. 1997. Stereoselective bimolecular phenoxy radical coupling by an auxiliary (dirigent) protein without an active center. Science 275: 362366.
  • Dean JV, Shah RP, Mohammed LA. 2003. Formation and vacuolar localization of salicylic acid glucose conjugates in soybean cell suspension cultures. Physiologia Plantarum 118: 328336.
  • Del Río JC, Marques G, Rencoret J, Martínez ÁT, Gutiérrez A. 2007. Occurrence of naturally acetylated lignin units. Journal of Agricultural and Food Chemistry 55: 54615468.
  • Dewick PM, Haslam E. 1969. Phenol biosynthesis in higher plants – gallic acid. Biochemical Journal 113: 537542.
  • Dimberg LH, Theander O, Lingnert H. 1993. Avenanthramides – a group of phenolic antioxidants in oats. Cereal Chemistry 70: 637641.
  • Dudareva N, Pichersky E, Gershenzon J. 2004. Biochemistry of plant volatiles. Plant Physiology 135: 18931902.
  • Ellis BE, Towers GHN. 1970. Biogenesis of rosmarinic acid in Mentha. Biochemical Journal 118: 291297.
  • Escamilla-Treviño LL, Chen W, Card ML, Shih M-C, Cheng C-L, Poulton JE. 2006. Arabidopsis thaliana β-Glucosidases BGLU45 and BGLU46 hydrolyse monolignol glucosides. Phytochemistry 67: 16511660.
  • Eudes A, George A, Mukerjee P, Kim JS, Pollet B, Benke PI, Yang F, Mitra P, Sun L, Çetinkol ÖP et al. 2012. Biosynthesis and incorporation of side-chain-truncated lignin monomers to reduce lignin polymerization and enhance saccharification. Plant Biotechnology Journal 10: 609620.
  • Farag MA, Huhman DV, Lei Z, Sumner LW. 2007. Metabolic profiling and systematic identification of flavonoids and isoflavonoids in roots and cell suspension cultures of Medicago truncatula using HPLC-UV-ESI-MS and GC-MS. Phytochemistry 68: 342354.
  • Fellenberg C, Böttcher C, Vogt T. 2009. Phenylpropanoid polyamine conjugate biosynthesis in Arabidopsis thaliana flower buds. Phytochemistry 70: 13921400.
  • Ferreres F, Figueiredo R, Bettencourt S, Carqueijeiro I, Oliveira J, Gil-Izquierdo A, Pereira DM, Valentão P, Andrade PB, Duarte P et al. 2011. Identification of phenolic compounds in isolated vacuoles of the medicinal plant Catharanthus roseus and their interaction with vacuolar class III peroxidase: an H2O2 affair? Journal of Experimental Botany 62: 28412854.
  • Franke R, Hemm MR, Denault JW, Ruegger MO, Humphreys JM, Chapple C. 2002. Changes in secondary metabolism and deposition of an unusual lignin in the ref8 mutant of Arabidopsis. Plant Journal 30: 4759.
  • Franke R, McMichael CM, Meyer K, Shirley AM, Cusumano JC, Chapple C. 2000. Modified lignin in tobacco and poplar plants over-expressing the Arabidopsis gene encoding ferulate 5-hydroxylase. Plant Journal 22: 223234.
  • Franke R, Schreiber L. 2007. Suberin – a biopolyester forming apoplastic plant interfaces. Current Opinion in Plant Biology 10: 252259.
  • Fraser CM, Thompson MG, Shirley AM, Ralph J, Schoenherr JA, Sinlapadech T, Hall MC, Chapple C. 2007. Related Arabidopsis serine carboxypeptidase-like sinapoylglucose acyltransferases display distinct but overlapping substrate specificities. Plant Physiology 144: 19861999.
  • Fry SC. 1986. Cross-linking of matrix polymers in the growing cell walls of angiosperms. Annual Review of Plant Physiology 37: 165186.
  • Fry SC, Miller JC. 1989 Toward a working model of the growing plant cell wall. Phenolic cross-linking reactions in the primary cell walls of dicotyledons. In: Lewis NG, Paice MG, eds. Plant cell wall polymers. Washington, DC, USA: American Chemical Society, 3346.
  • Fu C, Mielenz JR, Xiao X, Ge Y, Hamilton CY, Rodriguez M Jr, Chen F, Foston M, Ragauskas A, Bouton J et al. 2011. Genetic manipulation of lignin reduces recalcitrance and improves ethanol production from switchgrass. Proceedings of the National Academy of Sciences, USA 108: 38033808.
  • Gallego-Giraldo L, Escamilla-Trevino L, Jackson LA, Dixon RA. 2011a. Salicylic acid mediates the reduced growth of lignin down-regulated plants. Proceedings of the National Academy of Sciences, USA 108: 2081420819.
  • Gallego-Giraldo L, Jikumaru Y, Kamiya Y, Tang Y, Dixon RA. 2011b. Selective lignin downregulation leads to constitutive defense response expression in alfalfa (Medicago sativa L.). New Phytologist 190: 627639.
  • Goujon T, Sibout R, Pollet B, Maba B, Nussaume L, Bechtold N, Lu F, Ralph J, Mila I, Barrière Y et al. 2003. A new Arabidopsis thaliana mutant deficient in the expression of O-methyltransferase impacts lignins and sinapoyl esters. Plant Molecular Biology 51: 973989.
  • Grabber JH. 2005. How do lignin composition, structure, and cross-linking affect degradability? A review of cell wall model studies. Crop Science 45: 820831.
  • Grabber JH, Hatfield RD. 2005. Methyl esterification divergently affects the degradability of pectic uronosyls in nonlignified and lignified maize cell walls. Journal of Agricultural and Food Chemistry 53: 15461549.
  • Grabber JH, Hatfield RD, Lu F, Ralph J. 2008. Coniferyl ferulate incorporation into lignin enhances the alkaline delignification and enzymatic degradation of cell walls. Biomacromolecules 9: 25102516.
  • Grabber JH, Hatfield RD, Ralph J. 1998a. Diferulate cross-links impede the enzymatic degradation of non-lignified maize walls. Journal of the Science of Food and Agriculture 77: 193200.
  • Grabber JH, Hatfield RD, Ralph J. 2003. Apoplastic pH and monolignol addition rate effects on lignin formation and cell wall degradability in maize. Journal of Agricultural and Food Chemistry 51: 49844989.
  • Grabber JH, Mertens DR, Kim H, Funk C, Lu F, Ralph J. 2009. Cell wall fermentation kinetics are impacted more by lignin content and ferulate cross-linking than by lignin composition. Journal of the Science of Food and Agriculture 89: 122129.
  • Grabber JH, Quideau S, Ralph J. 1996a. p-Coumaroylated syringyl units in maize lignin: implications for β-ether cleavage by thioacidolysis. Phytochemistry 43: 11891194.
  • Grabber JH, Ralph J, Hatfield RD. 1998a. Ferulate cross-links limit the enzymatic degradation of synthetically lignified primary walls of maize. Journal of Agricultural and Food Chemistry 46: 26092614.
  • Grabber JH, Ralph J, Hatfield RD. 1998b Modeling lignification in grasses with monolignol dehydropolymerisate-cell wall complexes. In: Lewis NG, Sarkanen S, eds. Lignin and lignan biosynthesis. Washington, DC, USA: American Chemical Society, 163171.
  • Grabber JH, Ralph J, Hatfield RD. 2000. Cross-linking of maize walls by ferulate dimerization and incorporation into lignin. Journal of Agricultural and Food Chemistry 48: 61066113.
  • Grabber JH, Ralph J, Hatfield RD. 2002. Model studies of ferulate-coniferyl alcohol cross-product formation in primary maize walls: implications for lignification in grasses. Journal of Agricultural and Food Chemistry 50: 60086016.
  • Grabber JH, Ralph J, Hatfield RD, Quideau S, Kuster T, Pell AN. 1996b. Dehydrogenation polymer-cell wall complexes as a model for lignified grass walls. Journal of Agricultural and Food Chemistry 44: 14531459.
  • Grabber JH, Ress D, Ralph J. 2012. Identifying new lignin bioengineering targets: impact of epicatechin, quercetin glycoside, and gallate derivatives on the lignification and fermentation of maize cell walls. Journal of Agricultural and Food Chemistry 60: 51525160.
  • Grabber JH, Schatz PF, Kim H, Lu F, Ralph J. 2010. Identifying new lignin bioengineering targets: 1. Monolignol-substitute impacts on lignin formation and cell wall fermentability. BMC Plant Biology 10: 114.
  • Grienenberger E, Besseau S, Geoffroy P, Debayle D, Heintz D, Lapierre C, Pollet B, Heitz T, Legrand M. 2009. A BAHD acyltransferase is expressed in the tapetum of Arabidopsis anthers and is involved in the synthesis of hydroxycinnamoyl spermidines. Plant Journal 58: 246259.
  • Guo D, Chen F, Inoue K, Blount JW, Dixon RA. 2001a. Downregulation of caffeic acid 3-O-methyltransferase and caffeoyl CoA 3-O-methyltransferase in transgenic alfalfa: impacts on lignin structure and implications for the biosynthesis of G and S lignin. Plant Cell 13: 7388.
  • Guo D, Chen F, Wheeler J, Winder J, Selman S, Peterson M, Dixon RA. 2001b. Improvement of in-rumen digestibility of alfalfa forage by genetic manipulation of lignin O-methyltransferases. Transgenic Research 10: 457464.
  • Guo D-M, Ran J-H, Wang X-Q. 2010. Evolution of the cinnamyl/sinapyl alcohol dehydrogenase (CAD/SAD) gene family: the emergence of real lignin is associated with the origin of bona fide CAD. Journal of Molecular Evolution 71: 202218.
  • Hadacek F. 2002. Secondary metabolites as plant traits: current assessment and future perspectives. Critical Reviews in Plant Sciences 21: 273322.
  • Halpin C. 2010. Novel mutants optimized for lignin, growth and biofuel production. GCEP Research Symposium 2010. Stanford University, USA.
  • Hammerbacher A, Ralph SG, Bohlmann J, Fenning TM, Gershenzon J, Schmidt A. 2011. Biosynthesis of the major tetrahydroxystilbenes in spruce, astringin and isorhapontin, proceeds via resveratrol and is enhanced by fungal infection. Plant Physiology 157: 876890.
  • Harkin JM. 1967. Lignin – a natural polymeric product of phenol oxidation. In: Taylor WI, Battersby AR, eds. Oxidative coupling of phenols. New York, NY, USA: Marcel Dekker, 243321.
  • Hatfield R, Ralph J, Grabber JH. 2008. A potential role for sinapyl p-coumarate as a radical transfer mechanism in grass lignin formation. Planta 228: 919928.
  • Hatfield RD, Marita JM, Frost K, Grabber J, Ralph J, Lu F, Kim H. 2009. Grass lignin acylation: p-coumaroyl transferase activity and cell wall characteristics of C3 and C4 grasses. Planta 229: 12531267.
  • Hatfield RD, Ralph J, Grabber JH. 1999. Cell wall cross-linking by ferulates and diferulates in grasses. Journal of the Science of Food and Agriculture 79: 403407.
  • Hendriks ATWM, Zeeman G. 2009. Pretreatments to enhance the digestibility of lignocellulosic biomass. Bioresource Technology 100: 1018.
  • Herrmann KM, Weaver LM. 1999. The shikimate pathway. Annual Review of Plant Physiology and Plant Molecular Biology 50: 473503.
  • Hertweck C, Jarvis AP, Xiang L, Moore BS, Oldham NJ. 2001. A mechanism of benzoic acid biosynthesis in plants and bacteria that mirrors fatty acid β-oxidation. ChemBioChem 2: 784786.
  • Huis R, Morreel K, Fliniaux O, Lucau-Danila A, Fénart S, Grec S, Neutelings G, Chabbert B, Mesnard F, Boerjan W et al. 2012. Natural hypolignification is associated with extensive oligolignol accumulation in flax stems. Plant Physiology 158: 18931915.
  • Humphreys JM, Chapple C. 2002. Rewriting the lignin roadmap. Current Opinion in Plant Biology 5: 224229.
  • Humphreys JM, Hemm MR, Chapple C. 1999. New routes for lignin biosynthesis defined by biochemical characterization of recombinant ferulate 5-hydroxylase, a multifunctional cytochrome P450-dependent monooxygenase. Proceedings of the National Academy of Sciences, USA 96: 1004510050.
  • Ibdah M, Chen Y-T, Wilkerson CG, Pichersky E. 2009. An aldehyde oxidase in developing seeds of Arabidopsis converts benzaldehyde to benzoic acid. Plant Physiology 150: 416423.
  • Iijima Y, Nakamura Y, Ogata Y, Tanaka K, Sakurai N, Suda K, Suzuki T, Suzuki H, Okazaki K, Kitayama M et al. 2008. Metabolite annotations based on the integration of mass spectral information. Plant Journal 54: 949962.
  • Jackson LA, Shadle GL, Zhou R, Nakashima J, Chen F, Dixon RA. 2008. Improving saccharification efficiency of alfalfa stems through modification of the terminal stages of monolignol biosynthesis. BioEnergy Research 1: 180192.
  • Jones L, Ennos AR, Turner SR. 2001. Cloning and characterization of irregular xylem4 (irx4): a severely lignin-deficient mutant of Arabidopsis. Plant Journal 26: 205216.
  • Jouanin L, Goujon T, Sibout R, Pollet B, Mila I, Leplé JC, Pilate G, Petit-Conil M, Ralph J, Lapierre C. 2004. Comparison of the consequences on lignin content and structure of COMT and CAD downregulation in poplar and Arabidopsis thaliana. In: Walter C, Carson M, eds. Plantation forest biotechnology in the 21st century. Kerala, India: Research Signpost, 219229.
  • Kai K, Mizutani M, Kawamura N, Yamamoto R, Tamai M, Yamaguchi H, Sakata K, Shimizu B. 2008. Scopoletin is biosynthesized via ortho-hydroxylation of feruloyl CoA by a 2-oxoglutarate-dependent dioxygenase in Arabidopsis thaliana. Plant Journal 55: 989999.
  • Kang S, Kang K, Chung GC, Choi D, Ishihara A, Lee D-S, Back K. 2006. Functional analysis of the amine substrate specificity domain of pepper tyramine and serotonin N-hydroxycinnamoyltransferases. Plant Physiology 140: 704715.
  • Kim H, Ralph J, Lu F, Pilate G, Leplé J-C, Pollet B, Lapierre C. 2002. Identification of the structure and origin of thioacidolysis marker compounds for cinnamyl alcohol dehydrogenase deficiency in angiosperms. Journal of Biological Chemistry 277: 4741247419.
  • Kim H, Ralph J, Lu F, Ralph SA, Boudet A-M, MacKay JJ, Sederoff RR, Ito T, Kawai S, Ohashi H et al. 2003. NMR analysis of lignins in CAD-deficient plants. Part 1. Incorporation of hydroxycinnamaldehydes and hydroxybenzaldehydes into lignins. Organic & Biomolecular Chemistry 1: 268281.
  • Kim H, Ralph J, Yahiaoui N, Pean M, Boudet A-M. 2000. Cross-coupling of hydroxycinnamyl aldehydes into lignins. Organic Letters 2: 21972200.
  • Klempien A, Kaminaga Y, Qualley A, Nagegowda DA, Widhalm JR, Orlova I, Shasany AK, Taguchi G, Kish CM, Cooper BR et al. 2012. Contribution of CoA ligases to benzenoid biosynthesis in petunia flowers. Plant Cell 24: 20152030.
  • Knaggs AR. 2003. The biosynthesis of shikimate metabolites. Natural Product Reports 20: 119136.
  • Kristensen JB, Thygesen LG, Felby C, Jørgensen H, Elder T. 2008. Cell-wall structural changes in wheat straw pretreated for bioethanol production. Biotechnology for Biofuels 1: 5.
  • Lanot A, Hodge D, Jackson RG, George GL, Elias L, Lim E-K, Vaistij FE, Bowles DJ. 2006. The glucosyltransferase UGT72E2 is responsible for monolignol 4-O-glucoside production in Arabidopsis thaliana. Plant Journal 48: 286295.
  • Lapčík O. 2007. Isoflavonoids in non-leguminous taxa: a rarity or a rule? Phytochemistry 68: 29092916.
  • Lapierre C. 2010. Determing lignin structure by chemical degradations. In: Heitner C, Dimmel DR, Schmidt JA, eds. Lignin and lignans: CRC Press, Boca Raton, USA: 1148.
  • Lapierre C, Monties B, Rolando C. 1988. Thioacidolysis of diazomethane-methylated pine compression wood and wheat straw in situ lignins. Holzforschung 42: 409411.
  • Lapierre C, Pilate G, Pollet B, Mila I, Leplé J-C, Jouanin L, Kim H, Ralph J. 2004. Signatures of cinnamyl alcohol dehydrogenase deficiency in poplar lignins. Phytochemistry 65: 313321.
  • Lapierre C, Pollet B, Petit-Conil M, Toval G, Romero J, Pilate G, Leplé J-C, Boerjan W, Ferret V, De Nadai V et al. 1999. Structural alterations of lignins in transgenic poplars with depressed cinnamyl alcohol dehydrogenase or caffeic acidO-methyltransferase activity have an opposite impact on the efficiency of industrial kraft pulping. Plant Physiology 119: 153164.
  • Lee Y, Chen F, Gallego-Giraldo L, Dixon RA, Voit EO. 2011. Integrative analysis of transgenic alfalfa (Medicago sativa L.) suggests new metabolic control mechanisms for monolignol biosynthesis. PLoS Computational Biology 7: e1002047.
  • Léon J, Shulaev V, Yalpani N, Lawton MA, Raskin I. 1995. Benzoic-acid 2-hydroxylase, a soluble oxygenase from tobacco, catalyzes salicylic acid biosynthesis. Proceedings of the National Academy of Sciences, USA 92: 1041310417.
  • Leplé J-C, Dauwe R, Morreel K, Storme V, Lapierre C, Pollet B, Naumann A, Kang K-Y, Kim H, Ruel K et al. 2007. Downregulation of cinnamoyl-coenzyme A reductase in poplar: multiple-level phenotyping reveals effects on cell wall polymer metabolism and structure. Plant Cell 19: 36693691.
  • Li L, Cheng XF, Leshkevich J, Umezawa T, Harding SA, Chiang VL. 2001. The last step of syringyl monolignol biosynthesis in angiosperms is regulated by a novel gene encoding sinapyl alcohol dehydrogenase. Plant Cell 13: 15671585.
  • Li L, Popko JL, Umezawa T, Chiang VL. 2000. 5-Hydroxyconiferyl aldehyde modulates enzymatic methylation for syringyl monolignol formation, a new view of monolignol biosynthesis in angiosperms. Journal of Biological Chemistry 275: 65376545.
  • Li X, Bonawitz ND, Weng J-K, Chapple C. 2010. The growth reduction associated with repressed lignin biosynthesis in Arabidopsis thaliana is independent of flavonoids. Plant Cell 22: 16201632.
  • Li X, Chapple C. 2010. Understanding lignification: challenges beyond monolignol biosynthesis. Plant Physiology 154: 449452.
  • Lim E-K, Jackson RG, Bowles DJ. 2005. Identification and characterisation of Arabidopsis glycosyltransferases capable of glucosylating coniferyl aldehyde and sinapyl aldehyde. FEBS Letters 579: 28022806.
  • Liu C-J. 2010. Biosynthesis of hydroxycinnamate conjugates: implications for sustainable biomass and biofuel production. Biofuels 1: 745761.
  • Lorenzen M, Racicot V, Strack D, Chapple C. 1996. Sinapic acid ester metabolism in wild type and a sinapoylglucose-accumulating mutant of Arabidopsis. Plant Physiology 112: 16251630.
  • Lu F, Marita JM, Lapierre C, Jouanin L, Morreel K, Boerjan W, Ralph J. 2010. Sequencing around 5-hydroxyconiferyl alcohol-derived units in caffeic acid O-methyltransferase-deficient poplar lignins. Plant Physiology 153: 569579.
  • Lu F, Ralph J. 1999. Detection and determination of p-coumaroylated units in lignins. Journal of Agricultural and Food Chemistry 47: 19881992.
  • Lu F, Ralph J. 2002. Preliminary evidence for sinapyl acetate as a lignin monomer in kenaf. Chemical Communications 1: 9091.
  • Lu F, Ralph J. 2008. Novel tetrahydrofuran structures derived from β-β-coupling reactions involving sinapyl acetate in kenaf lignins. Organic & Biomolecular Chemistry 6: 36813694.
  • Lu F, Ralph J, Morreel K, Messens E, Boerjan W. 2004. Preparation and relevance of a cross-coupling product between sinapyl alcohol and sinapyl p-hydroxybenzoate. Organic & Biomolecular Chemistry 2: 28882890.
  • Mahesh V, Million-Rousseau R, Ullmann P, Chabrillange N, Bustamante J, Mondolot L, Morant M, Noirot M, Hamon S, de Kochko A et al. 2007. Functional characterization of two p-coumaroyl ester 3 ‘-hydroxylase genes from coffee tree: evidence of a candidate for chlorogenic acid biosynthesis. Plant Molecular Biology 64: 145159.
  • Malitsky S, Blum E, Less H, Venger I, Elbaz M, Morin S, Eshed Y, Aharoni A. 2008. The transcript and metabolite networks affected by the two clades of Arabidopsis glucosinolate biosynthesis regulators. Plant Physiology 148: 20212049.
  • Marita JM, Ralph J, Hatfield RD, Chapple C. 1999. NMR characterization of lignins in Arabidopsis altered in the activity of ferulate 5-hydroxylase. Proceedings of the National Academy of Sciences, USA 96: 1232812332.
  • Marita JM, Ralph J, Lapierre C, Jouanin L, Boerjan W. 2001. NMR characterization of lignins from transgenic poplars with suppressed caffeic acid O-methyltransferase activity. Journal of the Chemical Society, Perkin Transactions 1: 29392945.
  • Martin-Tanguy J. 1997. Conjugated polyamines and reproductive development: biochemical, molecular and physiological approaches. Physiologia Plantarum 100: 675688.
  • Masai E, Katayama Y, Fukuda M. 2007. Genetic and biochemical investigations on bacterial catabolic pathways for lignin-derived aromatic compounds. Bioscience, Biotechnology, and Biochemistry 71: 115.
  • Matsuda F, Hirai MY, Sasaki E, Akiyama K, Yonekura-Sakakibara K, Provart NJ, Sakurai T, Shimada Y, Saito K. 2010. AtMetExpress development: a phytochemical atlas of Arabidopsis development. Plant Physiology 152: 566578.
  • Matsuda F, Nakabayashi R, Sawada Y, Suzuki M, Hirai MY, Kanaya S, Saito K. 2011. Mass spectra-based framework for automated structural elucidation of metabolome data to explore phytochemical diversity. Frontiers in Plant Physiology 2: 40.
  • Matsuda F, Yonekura-Sakakibara K, Niida R, Kuromori T, Shinozaki K, Saito K. 2009. MS/MS spectral tag-based annotation of non-targeted profile of plant secondary metabolites. Plant Journal 57: 555577.
  • Matsuno M, Compagnon V, Schoch GA, Schmitt M, Debayle D, Bassard J-E, Pollet B, Hehn A, Heintz D, Ullmann P et al. 2009. Evolution of a novel phenolic pathway for pollen development. Science 325: 16881692.
  • Matsuno M, Nagatsu A, Ogihara Y, Ellis BE, Mizukami H. 2002. CYP98A6 from Lithospermum erythrorhizon encodes 4-coumaroyl-4′-hydroxyphenyllactic acid 3-hydroxylase involved in rosmarinic acid biosynthesis. FEBS Letters 514: 219224.
  • Meißner D, Albert A, Böttcher C, Strack D, Milkowski C. 2008. The role of UDP-glucose: hydroxycinnamate glucosyltransferases in phenylpropanoid metabolism and the response to UV-B radiation in Arabidopsis thaliana. Planta 228: 663674.
  • Merali Z, Mayer MJ, Parker ML, Michael AJ, Smith AC, Waldron KW. 2007. Metabolic diversion of the phenylpropanoid pathway causes cell wall and morphological changes in transgenic tobacco stems. Planta 225: 11651178.
  • Métraux J-P. 2002. Recent breakthroughs in the study of salicylic acid biosynthesis. Trends in Plant Science 7: 332334.
  • Meyer K, Shirley AM, Cusumano JC, Bell-Lelong DA, Chapple C. 1998. Lignin monomer composition is determined by the expression of a cytochrome P450-dependent monooxygenase in Arabidopsis. Proceedings of the National Academy of Sciences, USA 95: 66196623.
  • Miao Y-C, Liu C-J. 2010. ATP-binding cassette-like transporters are involved in the transport of lignin precursors across plasma and vacuolar membranes. Proceedings of the National Academy of Sciences, USA 107: 2272822733.
  • Milkowski C, Strack D. 2004. Serine carboxypeptidase-like acyltransferases. Phytochemistry 65: 517524.
  • Milkowski C, Strack D. 2010. Sinapate esters in brassicaceous plants: biochemistry, molecular biology, evolution and metabolic engineering. Planta 232: 1935.
  • Mir Derikvand M, Berrio Sierra J, Ruel K, Pollet B, Do C-T, Thévenin J, Buffard D, Jouanin L, Lapierre C. 2008. Redirection of the phenylpropanoid pathway to feruloyl malate in Arabidopsis mutants deficient for cinnamoyl-CoA reductase 1. Planta 227: 943956.
  • Mitchell RAC, Dupree P, Shewry PR. 2007. A novel bioinformatics approach identifies candidate genes for the synthesis and feruloylation of arabinoxylan. Plant Physiology 144: 4353.
  • Moco S, Bino RJ, Vorst O, Verhoeven HA, de Groot J, van Beek TA, Vervoort J, de Vos CHR. 2006. A liquid chromatography-mass spectrometry-based metabolome database for tomato. Plant Physiology 141: 12051218.
  • Molina I, Li-Beisson Y, Beisson F, Ohlrogge JB, Pollard M. 2009. Identification of an Arabidopsis feruloyl-coenzyme A transferase required for suberin synthesis. Plant Physiology 151: 13171328.
  • Morreel K, Dima O, Kim H, Lu F, Niculaes C, Vanholme R, Dauwe R, Goeminne G, Inzé D, Messens E et al. 2010a. Mass spectrometry-based sequencing of lignin oligomers. Plant Physiology 153: 14641478.
  • Morreel K, Kim H, Lu F, Dima O, Akiyama T, Vanholme R, Niculaes C, Goeminne G, Inzé D, Messens E et al. 2010b. Mass spectrometry-based fragmentation as an identification tool in lignomics. Analytical Chemistry 82: 80958105.
  • Morreel K, Ralph J, Kim H, Lu F, Goeminne G, Ralph S, Messens E, Boerjan W. 2004a. Profiling of oligolignols reveals monolignol coupling conditions in lignifying poplar xylem. Plant Physiology 136: 35373549.
  • Morreel K, Ralph J, Lu F, Goeminne G, Busson R, Herdewijn P, Goeman JL, Van der Eycken J, Boerjan W, Messens E. 2004b. Phenolic profiling of caffeic acid O-methyltransferase-deficient poplar reveals novel benzodioxane oligolignols. Plant Physiology 136: 40234036.
  • Muzac I, Wang J, Auzellotti D, Zhang H, Ibrahim RK. 2000. Functional expression of an Arabidopsis cDNA clone encoding a flavonol 3 ‘-O-methyltransferase and characterization of the gene product. Archives of Biochemistry and Biophysics 375: 385388.
  • Nair RB, Bastress KL, Ruegger MO, Denault JW, Chapple C. 2004. The Arabidopsis thaliana REDUCED EPIDERMAL FLUORESCENCE1 gene encodes an aldehyde dehydrogenase involved in ferulic acid and sinapic acid biosynthesis. Plant Cell 16: 544554.
  • Nakabayashi R, Kusano M, Kobayashi M, Tohge T, Yonekura-Sakakibara K, Kogure N, Yamazaki M, Kitajima M, Saito K, Takayama H. 2009. Metabolomics-oriented isolation and structure elucidation of 37 compounds including two anthocyanins from Arabidopsis thaliana. Phytochemistry 70: 10171029.
  • Nakagame S, Chandra RP, Kadla JF, Saddler JN. 2011. Enhancing the enzymatic hydrolysis of lignocellulosic biomass by increasing the carboxylic acid content of the associated lignin. Biotechnology and Bioengineering 108: 538548.
  • Niggeweg R, Michael AJ, Martin C. 2004. Engineering plants with increased levels of the antioxidant chlorogenic acid. Nature Biotechnology 22: 746754.
  • Önnerud H, Zhang L, Gellerstedt G, Henriksson G. 2002. Polymerization of monolignols by redox shuttle–mediated enzymatic oxidation: a new model in lignin biosynthesis I. Plant Cell 14: 19531962.
  • Osakabe K, Tsao CC, Li L, Popko JL, Umezawa T, Carraway DT, Smeltzer RH, Joshi CP, Chiang VL. 1999. Coniferyl aldehyde 5-hydroxylation and methylation direct syringyl lignin biosynthesis in angiosperms. Proceedings of the National Academy of Sciences, USA 96: 89558960.
  • Parvathi K, Chen F, Guo D, Blount JW, Dixon RA. 2001. Substrate preferences of O-methyltransferases in alfalfa suggest new pathways for 3-O-methylation of monolignols. Plant Journal 25: 193202.
  • Petersen M, Abdullah Y, Benner J, Eberle D, Gehlen K, Hücherig S, Janiak V, Kim KH, Sander M, Weitzel C et al. 2009. Evolution of rosmarinic acid biosynthesis. Phytochemistry 70: 16631679.
  • Pickel B, Constantin M-A, Pfannstiel J, Conrad J, Beifuss U, Schaller A. 2010. An enantiocomplementary dirigent protein for the enantioselective laccase-catalyzed oxidative coupling of phenols. Angewandte Chemie-International Edition 49: 202204.
  • Piston F, Uauy C, Fu L, Langston J, Labavitch J, Dubcovsky J. 2010. Down-regulation of four putative arabinoxylan feruloyl transferase genes from family PF02458 reduces ester-linked ferulate content in rice cell walls. Planta 231: 677691.
  • Pitre FE, Pollet B, Lafarguette F, Cooke JEK, MacKay JJ, Lapierre C. 2007. Effects of increased nitrogen supply on the lignification of poplar wood. Journal of Agricultural and Food Chemistry 55: 1030610314.
  • Ralph J. 1996. An unusual lignin from kenaf. Journal of Natural Products 59: 341342.
  • Ralph J 2006. What makes a good monolignol substitute? In: Hayashi T, ed. The science and lore of the plant cell wall biosynthesis, structure and function. Boca Raton, FL, USA: Universal Publishers (Brown Walker Press), 285293.
  • Ralph J. 2010. Hydroxycinnamates in lignification. Phytochemistry Reviews 9: 6583.
  • Ralph J, Akiyama T, Kim H, Lu F, Schatz PF, Marita JM, Ralph SA, Reddy MSS, Chen F, Dixon RA. 2006. Effects of coumarate 3-hydroxylase down-regulation on lignin structure. Journal of Biological Chemistry 281: 88438853.
  • Ralph J, Brunow G, Harris PJ, Dixon RA, Schatz PF, Boerjan W. 2008a. Lignification: are lignins biosynthesized via simple combinatorial chemistry or via proteinaceous control and template replication? In: Daayf F, Lattanzio V, eds. Recent advances in polyphenol research. Oxford, UK: Blackwell Publishing, 3666.
  • Ralph J, Bunzel M, Marita JM, Hatfield RD, Lu F, Kim H, Schatz PF, Grabber JH, Steinhart H. 2004a. Peroxidase-dependent cross-linking reactions of p-hydroxycinnamates in plant cell walls. Phytochemistry Reviews 3: 7996.
  • Ralph J, Grabber JH, Hatfield RD. 1995. Lignin-ferulate crosslinks in grasses: active incorporation of ferulate polysaccharide esters into ryegrass lignins. Carbohydrate Research 275: 167178.
  • Ralph J, Hatfield RD, Grabber JH, Jung H-JG, Quideau S, Helm RF. 1998 Cell wall cross-linking in grasses by ferulates and diferulates. In: Lewis NG, Sarkanen S, eds. Lignin and lignan biosynthesis. Washington, DC, USA: American Chemical Society, 209236.
  • Ralph J, Hatfield RD, Quideau S, Helm RF, Grabber JH, Jung H-JG. 1994a. Pathway of p-coumaric acid incorporation into maize lignin as revealed by NMR. Journal of the American Chemical Society 116: 94489456.
  • Ralph J, Kim H, Lu F, Grabber JH, Leplé J-C, Berrio-Sierra J, Mir Derikvand M, Jouanin L, Boerjan W, Lapierre C. 2008b. Identification of the structure and origin of a thioacidolysis marker compound for ferulic acid incorporation into angiosperm lignins (and an indicator for cinnamoyl CoA reductase deficiency). Plant Journal 53: 368379.
  • Ralph J, Lapierre C, Lu F, Marita JM, Pilate G, Van Doorsselaere J, Boerjan W, Jouanin L. 2001a. NMR evidence for benzodioxane structures resulting from incorporation of 5-hydroxyconiferyl alcohol into lignins of O-methyltransferase-deficient poplars. Journal of Agricultural and Food Chemistry 49: 8691.
  • Ralph J, Lapierre C, Marita JM, Kim H, Lu F, Hatfield RD, Ralph S, Chapple C, Franke R, Hemm MR et al. 2001b. Elucidation of new structures in lignins of CAD- and COMT-deficient plants by NMR. Phytochemistry 57: 9931003.
  • Ralph J, Lundquist K, Brunow G, Lu F, Kim H, Schatz PF, Marita JM, Hatfield RD, Ralph SA, Christensen JH et al. 2004b. Lignins: natural polymers from oxidative coupling of 4-hydroxyphenylpropanoids. Phytochemistry Reviews 3: 2960.
  • Ralph J, MacKay JJ, Hatfield RD, O'Malley DM, Whetten RW, Sederoff RR. 1997. Abnormal lignin in a loblolly pine mutant. Science 277: 235239.
  • Ralph J, Quideau S, Grabber JH, Hatfield RD. 1994b. Identification and synthesis of new ferulic acid dehydrodimers present in grass cell walls. Journal of the Chemical Society, Perkin Transactions 1: 34853498.
  • Rautengarten C, Ebert B, Ouellet M, Nafisi M, Baidoo EEK, Benke P, Stranne M, Mukhopadhyay A, Keasling JD, Sakuragi Y et al. 2012. Arabidopsis Deficient in Cutin Ferulate encodes a transferase required for feruloylation of ω-hydroxy fatty acids in cutin polyester. Plant Physiology 158: 654665.
  • del Río JC, Prinsen P, Rencoret J, Nieto L, Jiménez-Barbero J, Ralph J, Martínez AT, Gutiérrez A. 2012. Structural characterization of the lignin in the cortex and pith of elephant grass (Pennisetum purpureum) stems. Journal of Agricultural and Food Chemistry 60: 36193634.
  • Rogers LA, Campbell MM. 2004. The genetic control of lignin deposition during plant growth and development. New Phytologist 164: 1730.
  • Rohde A, Morreel K, Ralph J, Goeminne G, Hostyn V, De Rycke R, Kushnir S, Van Doorsselaere J, Joseleau J-P, Vuylsteke M et al. 2004. Molecular phenotyping of the pal1 and pal2 mutants of Arabidopsis thaliana reveals far-reaching consequences on phenylpropanoid, amino acid, and carbohydrate metabolism. Plant Cell 16: 27492771.
  • Rösler J, Krekel F, Amrhein N, Schmid J. 1997. Maize phenylalanine ammonia-lyase has tyrosine ammonia-lyase activity. Plant Physiology 113: 175179.
  • Saito K, Matsuda F. 2010. Metabolomics for functional genomics, systems biology, and biotechnology. Annual Review of Plant Biology 61: 463489.
  • Sarkanen KV, Ludwig CH. 1971. Lignins, occurrence, formation, structure and reactions. New York, NY, USA: Wiley-Interscience.
  • Schmidt A, Grimm R, Schmidt J, Scheel D, Strack D, Rosahl S. 1999. Cloning and expression of a potato cDNA encoding hydroxycinnamoyl-CoA: tyramineN-(hydroxycinnamoyl)transferase. Journal of Biological Chemistry 274: 42734280.
  • Seifert GJ, Blaukopf C. 2010. Irritable walls: the plant extracellular matrix and signaling. Plant Physiology 153: 467478.
  • Sharma V, Strack D. 1985. Vacuolar localization of 1-sinapoylglucose: L-malate sinapoyltransferase in protoplasts from cotyledons of Raphanus sativus. Planta 163: 563568.
  • Shimokawa Y, Morita H, Abe I. 2010. Structure-based engineering of benzalacetone synthase. Bioorganic & Medicinal Chemistry Letters 20: 50995103.
  • Sibout R, Baucher M, Gatineau M, Van Doorsselaere J, Mila I, Pollet B, Maba B, Pilate G, Lapierre C, Boerjan W et al. 2002. Expression of a poplar cDNA encoding a ferulate-5-hydroxylase/coniferaldehyde 5-hydroxylase increases S lignin deposition in Arabidopsis thaliana. Plant Physiology and Biochemistry 40: 10871096.
  • Sibout R, Eudes A, Mouille G, Pollet B, Lapierre C, Jouanin L, Séguin A. 2005. CINNAMYL ALCOHOL DEHYDROGENASE-C and -D are the primary genes involved in lignin biosynthesis in the floral stem of Arabidopsis. Plant Cell 17: 20592076.
  • Simmons BA, Loqué D, Ralph J. 2010. Advances in modifying lignin for enhanced biofuel production. Current Opinion in Plant Biology 13: 313320.
  • Sinlapadech T, Stout J, Ruegger MO, Deak M, Chapple C. 2007. The hyper-fluorescent trichome phenotype of the brt1 mutant of Arabidopsis is the result of a defect in a sinapic acid: UDPG glucosyltransferase. Plant Journal 49: 655668.
  • Soler M, Serra O, Molinas M, Huguet G, Fluch S, Figueras M. 2007. A genomic approach to suberin biosynthesis and cork differentiation. Plant Physiology 144: 419431.
  • Steffens JC. 2000. Acyltransferases in protease's clothing. Plant Cell 12: 12531256.
  • Stewart JJ, Akiyama T, Chapple C, Ralph J, Mansfield SD. 2009. The effects on lignin structure of overexpression of ferulate 5-hydroxylase in hybrid poplar. Plant Physiology 150: 621635.
  • Strawn MA, Marr SK, Inoue K, Inada N, Zubieta C, Wildermuth MC. 2007. Arabidopsis isochorismate synthase functional in pathogen-induced salicylate biosynthesis exhibits properties consistent with a role in diverse stress responses. Journal of Biological Chemistry 282: 59195933.
  • Syrjänen K, Brunow G. 1998. Oxidative cross coupling of p-hydroxycinnamic alcohols with dimeric arylglycerol β-aryl ether lignin model compounds. The effect of oxidation potentials. Journal of the Chemical Society, Perkin Transactions 1: 34253429.
  • Takahama U, Oniki T. 1997. Enhancement of peroxidase-dependent oxidation of sinapyl alcohol by an apoplastic component, 4-coumaric acid ester isolated from epicotyls of Vigna angularis L. Plant and Cell Physiology 38: 456462.
  • Takahama U, Oniki T, Shimokawa H. 1996. A possible mechanism for the oxidation of sinapyl alcohol by peroxidase-dependent reactions in the apoplast: enhancement of the oxidation by hydroxycinnamic acids and components of the apoplast. Plant and Cell Physiology 37: 499504.
  • Tanner GJ, Francki KT, Abrahams S, Watson JM, Larkin PJ, Ashton AR. 2003. Proanthocyanidin biosynthesis in plants - Purification of legume leucoanthocyanidin reductase and molecular cloning of its cDNA. Journal of Biological Chemistry 278: 3164731656.
  • Teutonico RA, Dudley MW, Orr JD, Lynn DG, Binns AN. 1991. Activity and accumulation of cell division-promoting phenolics in tobacco tissue cultures. Plant Physiology 97: 288297.
  • Tobimatsu Y, Elumalai S, Grabber JH, Davidson CL, Pan X, Ralph J. 2012. Hydroxycinnamate conjugates as potential monolignol replacements: in vitro lignification and cell wall studies with rosmarinic acid. ChemSusChem 2012: 676686.
  • Tzin V, Galili G. 2010. New insights into the shikimate and aromatic amino acids biosynthesis pathways in plants. Molecular Plant 3: 956972.
  • Umezawa T. 2003. Diversity in lignan biosynthesis. Phytochemistry Reviews 2: 371390.
  • Van Doorsselaere J, Baucher M, Chognot E, Chabbert B, Tollier M-T, Petit-Conil M, Leplé J-C, Pilate G, Cornu D, Monties B et al. 1995. A novel lignin in poplar trees with a reduced caffeic acid 5-hydroxyferulic acid O-methyltransferase activity. Plant Journal 8: 855864.
    Direct Link:
  • Van Moerkercke A, Schauvinhold I, Pichersky E, Haring MA, Schuurink RC. 2009. A plant thiolase involved in benzoic acid biosynthesis and volatile benzenoid production. Plant Journal 60: 292302.
  • Vanholme R, Van Acker R, Boerjan W. 2010c. Potential of Arabidopsis systems biology to advance the biofuel field. Trends in Biotechnology 28: 543547.
  • Vanholme R, Demedts B, Morreel K, Ralph J, Boerjan W. 2010a. Lignin biosynthesis and structure. Plant Physiology 153: 895905.
  • Vanholme R, Storme V, Vanholme B, Sundin S, Christensen JH, Goemine G, Halpin C, Rohde A, Morreel K, Boerjan W 2012. A systems biology view of responses to lignin biosynthesis perturbations in Arabidopsis. Plant Cell, doi:10.1105/tpc.112.102574.
  • Vanholme R, Morreel K, Ralph J, Boerjan W. 2008. Lignin engineering. Current Opinion in Plant Biology 11: 278285.
  • Vanholme R, Ralph J, Akiyama T, Lu F, Pazo JR, Kim H, Christensen JH, Van Reusel B, Storme V, De Rycke R et al. 2010b. Engineering traditional monolignols out of lignin by concomitant up-regulation of F5H1 and down-regulation of COMT in Arabidopsis. Plant Journal 64: 885897.
  • Vetting MW, de Carvalho LPS, Yu M, Hegde SS, Magnet S, Roderick SL, Blanchard JS. 2005. Structure and functions of the GNAT superfamily of acetyltransferases. Archives of Biochemistry and Biophysics 433: 212226.
  • Voelker SL, Lachenbruch B, Meinzer FC, Strauss SH. 2011. Reduced wood stiffness and strength, and altered stem form, in young antisense 4CL transgenic poplars with reduced lignin contents. New Phytologist 189: 10961109.
  • Wagner A, Ralph J, Akiyama T, Flint H, Phillips L, Torr K, Nanayakkara B, Kiri LT. 2007. Exploring lignification in conifers by silencing hydroxycinnamoyl-CoA: shikimate hydroxycinnamoyltransferase in Pinus radiata. Proceedings of the National Academy of Sciences, USA 104: 1185611861.
  • Wagner A, Tobimatsu Y, Phillips L, Flint H, Torr K, Donaldson L, Pears L, Ralph J. 2011. CCoAOMT suppression modifies lignin composition in Pinus radiata. Plant Journal 67: 119129.
  • Weng J-K, Chapple C. 2010. The origin and evolution of lignin biosynthesis. New Phytologist 187: 273285.
  • Weng J-K, Mo H, Chapple C. 2010. Over-expression of F5H in COMT-deficient Arabidopsis leads to enrichment of an unusual lignin and disruption of pollen wall formation. Plant Journal 64: 898911.
  • Werner RA, Rossmann A, Schwarz C, Bacher A, Schmidt H-L, Eisenreich W. 2004. Biosynthesis of gallic acid in Rhus typhina: discrimination between alternative pathways from natural oxygen isotope abundance. Phytochemistry 65: 28092813.
  • Wildermuth MC, Dewdney J, Wu G, Ausubel FM. 2001. Isochorismate synthase is required to synthesize salicylic acid for plant defence. Nature 414: 562565.
  • Wilkerson C, Ralph J, Withers S. 2011. BAHD acyltransferase that synthesizes coniferyl ferulate. Patent pending.
  • Wink M. 1997. Compartmentation of secondary metabolites and xenobiotics in plant vacuoles. Advances in Botanical Research 25: 141169.
  • Withers S, Lu F, Kim H, Zhu Y, Ralph J, Wilkerson CG. 2012. Identification of grass-specific enzyme that acylates monolignols with p-coumarate. Journal of Biological Chemistry 287: 83478355.
  • Yamada T, Matsuda F, Kasai K, Fukuoka S, Kitamura K, Tozawa Y, Miyagawa H, Wakasa K. 2008. Mutation of a rice gene encoding a phenylalanine biosynthetic enzyme results in accumulation of phenylalanine and tryptophan. Plant Cell 20: 13161329.
  • Yamamoto E, Bokelman GH, Lewis NG. 1989 Phenylpropanoid metabolism in cell walls. An overview. In: Lewis NG, Paice MG, eds. Plant cell wall polymers. Washington, DC, USA: American Chemical Society, 6888.
  • Yonekura-Sakakibara K, Tohge T, Matsuda F, Nakabayashi R, Takayama H, Niida R, Watanabe-Takahashi A, Inoue E, Saito K. 2008. Comprehensive flavonol profiling and transcriptome coexpression analysis leading to decoding gene-metabolite correlations in Arabidopsis. Plant Cell 20: 21602176.
  • Yoshida M, Liu Y, Uchida S, Kawarada K, Ukagami Y, Ichinose H, Kaneko S, Fukuda K. 2008. Effects of cellulose crystallinity, hemicellulose, and lignin on the enzymatic hydrolysis of Miscanthus sinensis to monosaccharides. Bioscience, Biotechnology, and Biochemistry 72: 805810.
  • Yoshida-Shimokawa T, Yoshida S, Kakegawa K, Ishii T. 2001. Enzymic feruloylation of arabinoxylan-trisaccharide by feruloyl-CoA: arabinoxylan-trisaccharide O-hydroxycinnamoyl transferase from Oryza sativa. Planta 212: 470474.
  • Yu O, Jez JM. 2008. Nature's assembly line: biosynthesis of simple phenylpropanoids and polyketides. Plant Journal 54: 750762.
  • Ziebell A, Gracom K, Katahira R, Chen F, Pu Y, Ragauskas A, Dixon RA, Davis M. 2010. Increase in 4-coumaryl alcohol units during lignification in alfalfa (Medicago sativa) alters the extractability and molecular weight of lignin. Journal of Biological Chemistry 285: 3896138968.

Supporting Information

  1. Top of page
  2. Abstract
  3. I. Introduction
  4. II. Phenolic metabolism
  5. III. Lignin biosynthesis and structure
  6. IV. Alternative lignin monomers for biofuel applications
  7. V. Candidate alternative monolignols in biomimetic systems
  8. VI. From phenolic profiling to lignomics
  9. VII. Phenolic pathway engineering towards alternative monolignols
  10. Acknowledgements
  11. References
  12. Supporting Information

Please note: Wiley-Blackwell are not responsible for the content or functionality of any supporting information supplied by the authors. Any queries (other than missing material) should be directed to the New Phytologist Central Office.

FilenameFormatSizeDescription
nph4337-sup-0001-NotesS1.docWord document653KNotes S1 The compounds, the plant species in which the compounds are found, and references used in the compilation of Fig. 5.