# Plant cell wall lignification and monolignol metabolism

Yin Wang, Maxime Chantreau, Maxime, Richard Sibout, Simon Hawkins

► To cite this version:

Yin Wang, Maxime Chantreau, Maxime, Richard Sibout, Simon Hawkins. Plant cell wall lignification and monolignol metabolism. Frontiers in Plant Science, 2013, 4, pp.1-14. 10.3389/fpls.2013.00220. hal-01204067

HAL Id: hal-01204067

https://hal.science/hal-01204067v1

Submitted on 29 May 2020

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>

Distributed under a Creative Commons CC BY 4.0 - Attribution - International License
<figure><img src="image_2.png" /></figure>

# Plant cell wall lignification and monolignol metabolism

Yin Wang<fnref n="1" />, Maxime Chantreau<fnref n="2" />,<fnref n="3" />, Richard Sibout<fnref n="1" />* and Simon Hawkins<fnref n="2" />,<fnref n="3" />

**Edited by:**

Samuel P. Hazen, University of
Massachusetts, USA

**Reviewed by:**

Dominique Loqué, Lawrence Berkeley
National Laboratory, USA
Curtis G. Wilkerson, Michigan State
University, USA

*Correspondence:

Richard Sibout, Unite Mixte
de Recherche 1318, Institut
Jean-Pierre Bourgin, Institut National
de la Recherche Agronomique, Saclay
Plant Sciences, F-78026 Versailles,
France
e-mail: richard.sibout@versailles.inra.fr

Plants are built of various specialized cell types that differ in their cell wall composition and structure. The cell walls of certain tissues (xylem, sclerenchyma) are characterized by the presence of the heterogeneous lignin polymer that plays an essential role in their physiology. This phenolic polymer is composed of different monomeric units – the monolignols – that are linked together by several covalent bonds. Numerous studies have shown that monolignol biosynthesis and polymerization to form lignin are tightly controlled in different cell types and tissues. However, our understanding of the genetic control of monolignol transport and polymerization remains incomplete, despite some recent promising results. This situation is made more complex since we know that monolignols or related compounds are sometimes produced in non-lignified tissues. In this review, we focus on some key steps of monolignol metabolism including polymerization, transport, and compartmentation. As well as being of fundamental interest, the quantity of lignin and its nature are also known to have a negative effect on the industrial processing of plant lignocellulose biomass. A more complete view of monolignol metabolism and the relationship that exists between lignin and other monolignol-derived compounds thereby appears essential if we wish to improve biomass quality.

**Keywords:** monolignol, lignin, lignan, metabolism, biomass

## INTRODUCTION

The majority of plant biomass consists of different cell wall poly-
mers produced by living plant cells. In most cases, these polymers
are energy-rich linked sugars that form the major structural com-
ponents in plant cell walls, particularly in the thick secondary cell
walls that characterize certain tissues. In addition to polysaccha-
rides, another major cell wall polymer – lignin – limits access
to cell wall sugars and negatively affects human utilization of
biomass (livestock feed, paper manufacturing, and lignocellulosic
biofuel production; Chapple et al., 2007). Because of its significant
economic impact and central role in higher plant development,
lignification is an important topic in plant biochemistry.

Despite a few exceptions (Martone et al., 2009), lignin is a
phenolic biopolymer that is only detected in vascular plants. It
is generated by radical coupling of hydroxycinnamyl alcohols
named monolignols: coniferyl alcohol, sinapyl alcohol, and *p*-coumaral alcohol. When introduced in the lignin polymer the
corresponding monolignols are named guaiacyl (G), sinapyl (S),
and hydroxyl-coumaroyl (H) units, respectively. The biosynthe-
sis of monolignols is initiated from the general phenylpropanoid
pathway (Dixon et al., 2001). Although tyrosine was proposed to
be the starting point of phenylpropanoid metabolism in some
plants such as grasses (Neish, 1961; Higuchi, 1990), it is gener-
ally recognized that monolignols are derived from phenylalanine
via a series of enzymatic reactions, catalyzed by the follow-
ing enzymes: phenylalanine ammonia lyase (PAL), cinnamate
4-hydroxylase (C4H), 4-coumarate coenzyme A ligase (4CL), fer-
ulate 5-hydroxylase (F5H), *p*-coumarate 3-hydroxylase (C3H),

*p*-hydroxycinnamoyl-CoA:quinate/shikimate hydroxycinnamoyl transferase (HCT), caffeoyl-CoA *O*-methyltransferase (CCOAO MT), cinnamoyl-CoA reductase (CCR), caffeic acid *O*-methyltransferase (COMT), and cinnamyl alcohol dehydrogenase (CAD; Vanholme et al., 2009).

The composition of lignin varies between plant species and tis-
sues. In general, lignins from gymnosperms and related species are
rich in G units and contains low amounts of H units, whereas dicot
lignins are mainly composed of G and S units (Weng and Chapple,
2010). It is important to note that differences in monolignol con-
tent are controlled by key enzymes that are often limiting (e.g., F5H
is limiting for S unit production; Weng and Chapple, 2010). Con-
sequently, certain natural genotypes/varieties may show unusual
proportions of monolignols as illustrated by the recent discovery
of a *Populus nigra* variety containing a truncated HCT enzyme
and producing substantial amounts of H units (Vanholme et al.,
2013) that are usually almost completely absent in poplar lignin.
Similarly, a loblolly pine naturally affected in cinnamyl alcohol
dehydrogenase activity and showing modified lignin structure was
also detected 20 years ago in a natural population (MacKay et al.,
1995; Ralph et al., 1997).

From a developmental point of view, lignification is generally
initiated during the formation of the secondary cell wall although
cell-/tissue-specific differences in the developmental pattern can
occur. For example, lignification is initiated in the compound pri-
mary cell wall of xylem elements (middle lamella/cell corners)
at the start of secondary cell wall formation (Donaldson, 2001;
Fromm et al., 2003). In contrast, lignin may be deposited at later

<footnote n="1">¹ Unite Mixte de Recherche 1318, Institut Jean-Pierre Bourgin, Institut National de la Recherche Agronomique, Saclay Plant Sciences, Versailles, France</footnote>

<footnote n="2">² Lille 1 UMR 1281, Université Lille Nord de France, Villeneuve d'Ascq, France</footnote>

<footnote n="3">³ Unite Mixte de Recherche 1281, Stress Abiotiques et Différenciation des Végétaux Cultivés, Institut National de la Recherche Agronomique, Villeneuve d'Ascq, France</footnote>
stages in other tissues (Boerjan et al., 2003; Baghdaddy et al., 2006; Harrington et al., 2012). For instance, parenchyma cells present in the secondary xylem of *Arabidopsis* hypocotyls are devoid of lignin during the vegetative stage (Sibout et al., 2008). This period may range over months according to different *Arabidopsis* accessions but ends as soon as the plant start flowering when parenchyma cells start to reinforce their secondary cell walls by producing lignin. Parenchyma therefore develops into sclerenchyma, a tissue dedicated to mechanical support. Biotic- and/or abiotic-stresses can also induce lignification in the walls of cells that do not normally lignify under non-stress conditions (Srivastava et al., 2007; Kim et al., 2008). Compression wood in coniferous trees also contains significant amounts of *p*-coumaryl alcohol (Sjöström, 1993). The observation that lignin biosynthetic genes are up-regulated (or activated) following mechanical injury suggests that stress lignification is a tightly controlled phenomenon. In contrast, the expression of monolignol biosynthesis genes is not always correlated with the presence of the lignin polymer. Although this is quite surprising at first glance, this situation can be explained by the fact that a wide range of other (non-lignin) products (hydroxycinnamic acids, phenols esters and phenol amides, or lignans; **Figure 1**) are also produced by the phenylpropanoid pathway.

In summary, lignification is a dynamic, flexible process reinforcing cell walls according to the different needs (water conduction, mechanical support, defense) of the plant during its life. Nevertheless, once produced, lignins (and the associated carbon) remain anchored in the cell wall since plants do not possess the enzymatic machinery necessary to recycle this polymer in contrast to other cell wall polysaccharides such as (1,3;1,4)-beta-D-glucans for instance (Fincher, 2009). The irreversibility of this situation also underlines the necessity for tight regulation of the lignification process.

Metabolic engineering can be used not only to reduce or modify natural lignin structure, but can also be employed to produce novel lignin structures in biomass crops with the goal of improving its structure for biomaterial or bioethanol production (Dixon et al., 1996; Vanholme et al., 2012a). Nevertheless, as underlined by a recent systems biology approach that investigated the consequences of lignin perturbations in *Arabidopsis* mutants, it is clear that our knowledge of this complex process is far from complete and that modifications in the lignification process can be accompanied by unexpected changes in gene expression and metabolism (Vanholme et al., 2012b). In this review, we focus on another aspect of lignin biology where our understanding is only partial – the different processes that occur after monolignol biosynthesis, including the acylation and glycosylation of monolignols in the cytoplasm, their transport through the membrane to the apoplast, and their deglycosylation, and polymerization into lignins. We also discuss lignans and related compounds produced by the same biosynthetic pathway. In this review we aim to point out some gaps in our knowledge of lignification that are potentially limiting for biomass use and engineering.

## TRANSPORT AND COMPARTMENTATION

Monolignols are synthesized in the cytoplasm and translocated to the cell wall for subsequent polymerization (Alejandro et al.,

2012). For many years, the mechanism of transport remained unknown and different hypotheses were suggested (passive diffusion, exocytosis, active transport, etc.; Liu et al., 2011; Liu, 2012). An elegant biochemical study has recently demonstrated that the glycosylation status determines monolignol transport and subcellular compartmentation (Miao and Liu, 2010). Plasma membrane-derived vesicles prepared from *Arabidopsis* and poplar cells transported coniferyl alcohol (the aglycone form), but not coniferin (the glycosylated form of coniferyl alcohol), in an ATP-dependant manner, whereas tonoplast-derived vesicles transported the glycosylated form but not the aglycone. Taken together these results would suggest that coniferyl alcohol is transported into the cell wall across the plasma membrane in an ATP-dependant process by an ABC-transporter and is subsequently polymerized by laccases (LACs) and/or peroxidases (PRXs). In contrast, the glycosylated form (e.g., coniferin) is transported into the vacuole for storage. Genetic confirmation of the involvement of ABC transporters in monolignol transport into the cell wall was recently reported (Alejandro et al., 2012). Co-expression studies in *Arabidopsis* identified an ABCG transporter gene (*AtABCG29*) co-regulated with phenylpropanoid gene expression and that was expressed in lignin-containing organs and tissues. Protein-fusion studies demonstrated that AtABCG29 was localized in the plasma membrane and transport studies in yeast suggested that the protein was involved in transporting *p*-coumaryl alcohol, but not coniferyl alcohol. A minor activity was also observed toward sinapyl alcohol. *Abcg29* mutant lines showed reduced root growth when grown on medium containing *p*-coumaroyl alcohol, but not coniferyl alcohol or sinapyl alcohol supporting the idea that AtABCG29 is involved in *p*-coumaryl alcohol transport. Lignin analyses of roots from two mutant lines revealed decreases in the amounts of H, G, and S subunits as compared to WT roots, an unexpected result since AtABCG29 is highly specific for *p*-coumaryl alcohol and it would be expected that only H units would be reduced. Interestingly, the mutant lines also showed metabolomic modifications in soluble phenolics, flavonoids and glucosinolates together with reductions in associated gene expression. These modifications are reminiscent of changes observed in different lignin mutants (Vanholme et al., 2012b) underlining the fact that phenylpropanoid metabolism can be perturbed not only at the biosynthesis level, but also by modifications of monolignol transport. The results also raise the possibility that the observed cell-specific variation in lignin composition (e.g., vessel element walls are rich in G lignin whereas fiber walls contain S–G lignin) may also be regulated via specific ABC transporters. Plant genomes contain large ABC transporter gene families (e.g., 130 genes in *Arabidopsis*) but ABC transporters for coniferyl alcohol and sinapyl alcohol remain to be identified and characterized. In addition, it is probable that monolignol glucosides from the vacuole participate to tracheary element lignification following cell death and membrane disruption (Pesquet et al., 2013). Additional information on the potential role(s) of ABC transporters in the lignification process is discussed by Sibout and Hofte (2012).

## MONOLIGNOL GLYCOSYLATION AND DEGLYCOSYLATION

Glycosylation of monolignols is catalyzed by UDP-glycosyltransferases (UGTs) belonging to the glycosyltransferase (GT) family 1.
<figure><img src="image_3.png" /></figure>

**FIGURE 1 | Schematic view of the monolignol biosynthetic pathway.** The synthesis of monolignols from phenylalanine and shikimate involves cytosolic (PAL, HCT, 4CI, CCR, CAD, CCoAOMT, and OMT) and ER membrane-anchored (the cytochrome P450 enzymes F5H, C3H, and C4H) enzymes. Monolignols and lignans may be conjugated by UGTs and then transported to the vacuole or directly transported to the cell wall for oxidative cross-linking by apoplastic peroxidases and laccases into lignins. In grasses, some hydroxycinnamic acids are conjugated to arabinose by BAHD enzyme for export via the Golgi apparatus and future incorporation into polysaccharides (pectins or arabinoxylans) whereas some are conjugated to monolignols (mainly sinapyl alcohol) in the cytosol and incorporated into lignin. Other phenolic compounds (soluble phenolic esters and amides, flavonoids) are shown and may indirectly interact with lignification. Their production implies common steps (enzymes) with the monolignol pathway. Most of these transport routes and storage forms still remain to be discovered and the localization of some enzymes (GH,

Perox, Lacc) confirmed. The dashed lines delineate putative pathways and full lines delineate known routes. Enzymes of the monolignol pathway are represented by circles. The brown color indicates involvement of the enzyme in the monolignol pathway; yellow indicates involvement in the flavonoid pathway; and pink indicates involvement in the sinapate ester pathway. PAL, phenylalanine ammonia lyase; C4H, cinnamic acid 4-hydroxylase; 4CL, 4-hydroxycinnamoyl-CoA ligase; HCT, hydroxycinnamoyl-CoA:shikimate hydroxycinnamoyl transferase; C3H, *p*-coumaroyl shikimate 3'-hydroxylase; CCoAOMT, caffeoyl-CoA *O*-methyltransferase; CCR, hydroxycinnamoyl-CoA reductase; F5H, ferulic acid 5-hydroxylase; COMT, caffeic acid/5-hydroxyferulic acid *O*-methyltransferase; CAD, cinnamyl alcohol dehydrogenase; UGT, UDP-glycosyltransferase; GH, beta-glucosidase; Perox, peroxidase; Lacc, laccase; DPs, dirigent proteins; BAHD, hydroxycinnamic acid transferase; ER, endoplasmic reticulum.
These enzymes play an important role in stabilization, enhance-
ment of water solubility and deactivation/detoxification of a wide
range of natural products including hormones and secondary
metabolites (Lim and Bowles, 2004). A recent phylogenetic recon-
struction (Caputi et al., 2012) of family 1 UGTs reveals a dramatic
evolutionary increase in the number of family 1 GTs from one in
the model green alga *Chlamydomonas* to 243 in poplar reflect-
ing the increased complexity of plant life on Earth. Less than
20% plant UGTs have been functionally characterized (Yonekura-
Sakakibara et al., 2012). A major problem is that plant GTs may
be redundant. Functional analyses in *Arabidopsis* (Lanot et al.,
2006) of the UGT72E clade suggested that the *UGT72E2* gene
product was responsible for monolignol glycosylation. Chemi-
cal analyses of soluble phenolics in light-grown roots from the
triple mutant showed a significant decrease in the quantity of
coniferyl alcohol- and sinapyl alcohol-glucosides as compared to
WT plants. Over-expression of the *UGT72E2* gene resulted in a
10-fold increase in coniferin levels and a lower increase in sinapyl
alcohol glucoside in roots suggesting that this is the principal
gene involved in monolignol glycosylation in *Arabidopsis*. Interest-
ingly, rosette leaves accumulated sinapoyl glucoside to higher levels
than coniferyl alcohol glucoside hinting at organ-specific varia-
tions in glycosylation mechanisms. However, no impact on lignin
content/composition was observed. Evidence suggesting a link
between lignification and monolignol glycosylation was recently
demonstrated (Wang et al., 2012) when the over-expression in
tobacco plants of a poplar UGT (*PtGT1*) homolog to the *Ara-
bidopsis* *UGT72E1-3* genes was shown to be associated with an
increased lignification as well as early flowering. Nevertheless, the
observation that recombinant PtGT1 showed no activity toward
monolignols and the fact that transformed plants did not contain
higher amounts of monolignol glycosides led the authors to sug-
gest that the observed impact of *PtGT1* over-expression in tobacco
was indirect. Further research is clearly needed to identify not
only the UGTs specifically involved in monolignol glycosylation,
but also to demonstrate a functional link between modifications
in *UGT* gene expression and lignification. The potential impor-
tance of monolignol glycobiology in regulating lignin biosynthesis
is also complicated by the fact that, in addition to glucose, plant
UGTs can also use different sugars (glucose, rhamnose, galactose,
glucuronic acid, etc.) to glycosylate monolignols (Caputi et al.,
2012).

If monolignol glucosides give rise to monolignols that are
then incorporated into the lignin polymer, then enzymes (beta-
glucosidases) should be co-localized with coniferin and syringin.
Beta-glucosidases capable of hydrolyzing monolignol glucosides
were first identified in cell wall fractions of *Picea abies* hypocotyls
and roots (Marcinowski and Grisebach, 1978). Subsequently,
a beta-glucosidase showing high affinity for coniferin was co-
localized with PRX activity to cell walls of young differentiating
xylem in *Pinus contorta* (Dharmawardhana et al., 1995, 1999). In
Angiosperms, beta-glucosidase activity was first located in the cell
walls of *Cicer arietinum* L. (Hosel et al., 1978; Burmeister and
Hosel, 1981). The co-localization of beta-glucosidase activity and
monolignol glucosides in differentiating xylem of conifers and
some angiosperms led to the hypothesis that monolignol gluco-
sides could represent lignin precursors. However, subsequent work

in *Ginkgo biloba* (Tsuji et al., 2005) showing that a beta-glucosidase demonstrated a 10 times greater affinity toward coniferaldehyde as compared to coniferin threw doubt on this initial idea. Subsequent work with radioactive precursors indicated that coniferyl alcohol derived from coniferin was only weakly incorporated into the lignin polymer and that greater amounts were converted into coniferaldehyde glucoside followed by subsequent deglycosylation and CAD-mediated conversion of the coniferaldehyde into coniferyl alcohol. These results suggested that coniferin represented the storage form of coniferyl alcohol and was not the direct precursor of coniferyl alcohol for lignin. In *Arabidopsis*, three orthologs (BGLU45/At1g61810; BGLU46/At1g61820; BGLU47/At4g21760) of the *Pinus contorta* beta-glucosidase gene are present and belong to the group 10 of the *Arabidopsis* GH1 family (Xu et al., 2004). Heterologous expressions of BGLU45 and BGLU46 showed that BGLU45 is highly specific for the three monolignol glucosides, whereas BGLU46 preferentially uses *p*-coumaryl alcohol glucoside as well as showing broad activity toward other phenolic glucosides (Xu et al., 2004; Escamilla-Trevino et al., 2006). A recent functional study of *Arabidopsis* *bglu45-1* and *bglu46-2* T-DNA mutants (Chapelle et al., 2012) showed only slight reductions (8.5 and 6.5%, respectively) in global beta-glucosidase activity and protein amounts as estimated by Western blots following Concanavalin-A column chromatography, presumably because of functional redundancy – at least four other glucosidases are expressed in *Arabidopsis* stems – and antibody cross-reactivity. Lignin analyses showed a slight but significant increase in Klason lignin for two BGLU45 mutants of the WS accession (*bglu45-1* and *bglu45-3*) under long-day conditions only but not in Col0 bglu45 mutants. No significant modification in lignin content was observed for *bglu46* mutants. Similarly, thioacidolysis revealed no changes in lignin subunit structure. Metabolomics (ultra performance liquid chromatography–mass spectrometry, UPLC–MS) identified around 100 different small glycosylated phenolic compounds, most of which were increased in the mutants. For the 10 most important differentials, five were identified/partially identified and all were derived from G units. Interestingly, an important ecotype effect was observed with, for example, an approximate 20-fold increase of coniferin in WS *bglu45-1* mutants, but only a 3.4-fold increase in Col10 mutants. While coniferyl alcohol, sinapyl alcohol, coniferaldehyde, sinapaldehyde, and syringin levels were not significantly modified, certain (neo)lignans increased [lariciresinol hexoside or isodihydrodehydrodiconiferyl alcohol (IDDDC) hexoside, dehydrodiconiferyl alcohol (DDC) hexoside]. Based on these observations, the authors proposed that monolignol glucosides are the storage form of monolignols and not the direct precursors. Nevertheless, it is interesting to note that *BGLU45* and *BGLU46* expression are deregulated under various biotic stresses suggesting that the monolignol storage pool might be used under pathogen attack in the fabrication of “defense lignin” and/or phytoalexins. Certainly, such a hypothesis allows a better understanding of the basic “spatial problem” associated with beta-glucosidase enzyme and substrate compartmentation (Figure 1). As indicated above, the *Pinus contorta* beta-glucosidase was localized to differentiating xylem walls and BGLU45/46 proteins are secreted to the cell wall (Chapelle et al., 2012), whereas glycosylated monolignols are stored in the
vacuole. Under such conditions it is perhaps not surprising that beta-glucosidase gene down-regulation has little/no effect on lignin levels under standard conditions and that the critical step occurs at the glycosylation step deciding whether monolignols are targeted to the cell wall without glycosylation for lignification, or glycosylated for storage in the vacuole. In contrast, cell and vacuole disruption following pathogen attack, insect/herbivore feeding and potentially severe abiotic stress (cell freezing, desiccation) or during programmed cell death (xylem vessel) would lead to contact between the monolignol glucosides and beta-glucosidases at the cell wall where PRXs and/or LACs could rapidly polymerize the aglycone forms. The role of beta-glucosidases in producing a wide range of bioactive defense molecules has been largely documented in plants (Morant et al., 2008).

Taken together, these observations indicate that the glycosylation/deglycosylation of monolignols play key roles in determining their availability for subsequent lignin biosynthesis thereby representing potentially interesting engineering targets for plant biomass improvement. In addition, similar mechanisms are also likely to affect the availability of ferulic- and *p*-coumaric acids for lignin-hemicellulose/pectin and polysaccharide-polysaccharide covalent cross-links in the cell wall underlining the necessity for further research into the glycosylation/deglycosylation of monolignols and phenolic molecules.

## DIMERIZATION OF MONOLIGNOLS AND SYNTHESIS OF (NEO)LIGNANS

Monolignols can also give rise to more than 3,000 different lignans and associated structures such as neo-, sesqui-, and flavanolignans (Schmidt et al., 2010). Lignans are optically active phenylpropanoid dimers that result from the stereo-selective dirigent protein-mediated coupling of the 8 and 8' *C*-atoms of two hydroxycinnamyl alcohol (monolignol) moieties (Davin et al., 1997). (Note: two different nomenclatures are used to describe the propane carbon atoms involved in monolignol linkages in the lignin polymer and in oligolignols. In oligolignols, the numbers 7, 8, and 9 are used whereas in the lignin polymer *α*, *β*, and *γ* are used to describe the same carbon atoms. A *β*–*β* lignin linkage therefore corresponds to an 8–8 *oligolignol* linkage). Lignan formation has been widely studied since these molecules are natural anti-oxidants, possess anti-microbial activity and have numerous beneficial effects on human (Ford et al., 2001). Lignans are known to play a role in the durability, longevity, and resistance of the heartwood of many tree species against woodrotting fungi and are believed to function as phytoalexins in other (non-woody) plant species (Pohjamo et al., 2003). The formation and accumulation of lignans in flax (*Linum usitatissimum* L.) seeds has been particularly studied since they are one of the richest natural grain sources of certain lignans. The genus *Linum* contains approximately 200 species worldwide that can be divided into two main groups depending upon the major lignan types accumulated – either cyclolignans belonging to the aryltetralin series (AT lignans) or cyclolignans of the aryldihydronaphthalene (ADN)/arylnaphthalene (AN) group (Schmidt et al., 2010). Some plants in both groups can also accumulate dibenzylbutyrolactone (DBBL) lignans such as matairesinol and furofuran (FF) lignans such as pinoresinol. Both FF and DBBL lignans

are precursors of the AT/ADN/AN lignans. In the seeds of flax, dimerization of two E-coniferyl alcohol molecules leads to the formation of 8S,8'S-(-)-pinoresinol, which is sequentially converted into 8S,8'S-(-)-lariciresinol and 8S,8'S-(++)-secoisolariciresinol via the action of the enzyme pinoresinol lariciresinol reductase (PLR). (+)-Secoisolariciresinol is then glycosylated to secoisolariciresinol diglucoside (SDG) that accumulates to high levels in flax seeds (linseed; Ford et al., 2001). The complexity of lignan stereochemistry was underlined in a recent study (Hemmati et al., 2010) that isolated two PLRs (PLR-Lu1, PLR-Lu2) showing opposite enantiospecificity. PLR-Lu1 utilizes 8S,8'S-pinoresinol, whereas PLR-Lu2 uses 8R,8'R-pinoresinol. Interestingly, gene expression analyses have shown that the *LuPLR1* gene is expressed in different flax seed tissues, whereas the *LuPLR2* gene is expressed in flax stem and leaf tissues (Hano et al., 2006; Hemmati et al., 2010; Venglat et al., 2011). In agreement with this organ-specific gene expression, 8S,8'S-SDG accumulates in flax seeds whereas aerial green parts accumulate 8R,8'R-SDG. Further analyses (Schmidt et al., 2012) of lignans in 16 different *Linum* species suggested that lignan stereochemistry also depended upon the species analyzed and whereas R,R-SDG was mainly accumulated in the seeds from species containing relatively high amounts of ADN/AN cyclolignans, S,S-SDG was accumulated in seeds from species containing only trace amounts of cyclolignans.

A further insight into monolignol/lignan biology in flax was provided by a combined transcriptomics and metabolomics study (Huis et al., 2012) of flax inner- (xylem rich) and outer- (bast fiber rich) stem tissues (Figure 2). More than eighty different lignans and neolignans including pinoresinol, lariciresinol, secoisolariciresinol, DDC, and IDDDC, as well as their glycosylated forms were detected in both tissues. In agreement with the presence of lignans and neolignans in the flax stem, transcripts corresponding to PLR, as well as the enzyme phenylcoumaran benzylic ether reductase (PCBER) catalyzing the reduction of DDC into IDDDC and previously identified as the most abundant protein in poplar xylem (Gang et al., 1999; Vander Mijnsbrugge et al., 2000) were also highly abundant. Although PCBER transcripts were more highly abundant in inner stem tissues, IDDDC was not detected in either inner- or outer-stem tissues, despite the presence of DDC in both tissues. In contrast IDDDC hexosides were detected in both stem tissues suggesting that the neolignan is rapidly glycosylated. Overall, the glycosylated forms were more abundant in outer stem tissues in comparison with inner stem tissues. Further work in other species is necessary to know whether such differences in (neo)lignan glycobiology are widespread and also the potential biological role. Interestingly, recent reports showing that lignans are detected in the phloem tissues (Ramos and Kato, 2012) and honey-dew of the stink-bug and that glycosylated flax lignans but not the aglycone forms significantly increased aphid *Myzus persicae* insect mortality (Saguez et al., 2012) raise the possibility that monolignol-derived compounds might be glycosylated and transported in phloem cells as a defense mechanism against insect predators.

The abundance of glycosylated (neo)lignans in flax and other species suggests the existence of different UGTs capable of glycosylating lignans. Lignan UGT activity was first identified in sesame (Noguchi et al., 2008) and more recently in *Forsythia* (Ono
<figure><img src="image_4.png" /></figure>

<table><thead><tr><th>1) MONOLIGNOLS</th><th>Inner tissues</th><th>Outer tissues</th><th>Out/Inn</th></tr></thead><tbody><tr><td>p-coumaroyl alcohol (H)</td><td>-</td><td>-</td><td>-</td></tr><tr><td>p-coumaroyl hex</td><td>2.3 x 10<sup>6</sup></td><td>1.0 x 10<sup>7</sup></td><td>4.34</td></tr><tr><td>Coniferyl alcohol (G)</td><td>4.8 x 10<sup>3</sup></td><td>1.3 x 10<sup>4</sup></td><td>2.7</td></tr><tr><td>Coniferin (CA hex)</td><td>3.2 x 10<sup>6</sup></td><td>1.2 x 10<sup>7</sup></td><td>3.75</td></tr><tr><td>Sinapyl alcohol (S)</td><td>-</td><td>-</td><td>-</td></tr><tr><td>Syringin (SA hex)</td><td>5.0 x 10<sup>3</sup></td><td>1.8 x 10<sup>4</sup></td><td>3.6</td></tr></tbody></table>

<table><thead><tr><th>2) LIGNANS</th><th>Inner tissues</th><th>Outer tissues</th><th>Out/Inn</th></tr></thead><tbody><tr><td>Pinoresinol</td><td>2.8 x 10<sup>3</sup></td><td>3.1 x 10<sup>3</sup></td><td>2.7</td></tr><tr><td>Pinoresinol hex</td><td>6.6 x 10<sup>3</sup></td><td>1.1 x 10<sup>5</sup></td><td>16.8</td></tr><tr><td>Lariciresinol</td><td>1.6 x 10<sup>4</sup></td><td>1.2 x 10<sup>4</sup></td><td>0.75</td></tr><tr><td>Lariciresinol hex 1</td><td>1.3 x 10<sup>4</sup></td><td>2.8 x 10<sup>5</sup></td><td>21.5</td></tr><tr><td>Lariciresinol hex 2</td><td>8.3 x 10<sup>3</sup></td><td>1.1 x 10<sup>5</sup></td><td>13.4</td></tr><tr><td>Secoisolariciresinol</td><td>-</td><td>-</td><td>-</td></tr><tr><td>Secoisolariciresinol hex 1</td><td>2.6 x 10<sup>4</sup></td><td>-</td><td>Inner</td></tr><tr><td>Secoisolariciresinol hex 2</td><td>2.1 x 10<sup>4</sup></td><td>3.1 x 10<sup>5</sup></td><td>14.7</td></tr></tbody></table>

<table><thead><tr><th>3) NEOLIGNANS</th><th>Inner tissues</th><th>Outer tissues</th><th>Out/Inn</th></tr></thead><tbody><tr><td>DDC</td><td>9.5 x 10<sup>4</sup></td><td>3.0 x 10<sup>4</sup></td><td>0.3</td></tr><tr><td>DDC hex</td><td>-</td><td>-</td><td>-</td></tr><tr><td>IDDC</td><td>-</td><td>-</td><td>-</td></tr><tr><td>IDDDC hex 1</td><td>7.6 x 10<sup>2</sup></td><td>2.0 x 10<sup>4</sup></td><td>26.3</td></tr><tr><td>IDDDC hex 2</td><td>2.1 x 10<sup>4</sup></td><td>5.6 x 10<sup>5</sup></td><td>26.7</td></tr></tbody></table>

<figure><img src="image_5.png" /><figcaption>**FIGURE 2** | Monolignol metabolism in flax stems. Olignolignols were extracted from inner (IT) and outer (OT) tissues of flax stems (A) and analyzed by ultra-high performance liquid chromatography–Fourier transform–ion cyclotron resonance–mass spectrometry (UHPLC–FT–ICR–MS; Huis et al., 2012). Outer tissues are characterized by the presence of bast fibers with thick cellulose-rich secondary cell walls and extremely low lignin levels as indicated by limited red coloration with phloroglucinol–HCl staining (B). Inner tissues contain xylem tissue with heavily lignified cell walls (C). Aglycone and</figcaption></figure>

glycosylated forms of monolignols (1), lignans (2), and neolignans (3) were detected in both inner- and outer-stem samples. Glycosylated forms were generally more abundant in outer stems. DDC, dehydrodiconiferyl alcohol; IDDDC, isodihydrodehydrodiconiferyl alcohol; PLR, pinoresinol lariciresinol reductase; PCBER, phenylcoumaran benzylic ether reductase. Red heat blocks indicate corresponding unigene expression in inner- versus outer-stem tissues. This figure illustrates the extremely rich metabolism post-monolignol biosynthesis. (A) Courtesy of Anne-Sophie Blervacq, Université Lille 1.
et al., 2010). Subsequent screening of a *Forsythia* cDNA library and biochemical characterization of recombinant UGT71A17 and UGT71A18 proteins showed that both proteins were capable of glycosylating (+)-pinoresinol albeit UGT71A17 activity was much weaker than UGT71A18 activity. Further characterization of the latter showed that it was active against different FF and DBBL lignans. Expression analyses indicated that both genes were expressed in leaves, floral buds, and petals, but that only *UGT71A18* was expressed in cell suspension cultures. It would obviously be interesting to see whether *UGT* genes are also expressed in other organs/tissues. Monolignol dimerization is presumably catalyzed by class III PRXs that are localized in the cell wall and/or vacuole suggesting that lignans should also be localized in these compartments. Although one study (Attoumbre et al., 2010) localized lignans to the cell wall in flax seeds, a positive signal was also observed in cytoplasmic inclusions raising the possibility that lignans might also be present in this compartment. Plant UGTs, unlike their mammalian counterparts, do not possess peptide signals and are therefore probably localized in the cytoplasm. In this case, where does (neo)lignan glycosylation take place – do plants possess plasma membrane/tonoplast ABC transporters capable of transporting neolignans into/out of cells/tonoplasts? In conclusion, although we now have a relatively good understanding of monolignol biosynthesis (even if our knowledge of transcriptional and post-transcriptional regulation processes is far from complete), it is clear that we have much to learn about the mechanisms regulating the availability of synthesized monolignols in the cell wall for subsequent lignification. A better understanding of these mechanisms should lead to the elaboration of new targets for engineering more efficient plant biomass.

## POLYMERIZATION

Beyond dimerization formation of lignins requires a polymerization step. The complex structure of the polymers built up of different subunits (H, G, and S), the presence of different types of linkages between subunits and the absence of repeated motifs suggest a highly unpredictable polymerization mechanism probably affected by its microenvironment (pH, polysaccharides, etc.;

Wingender et al., 1999). In the developing cell wall, lignin polymerization occurs by radical coupling reactions (Freudenberg, 1959; **Figures 3 and 4**). Type III PRXs and LACs are probably both involved in monolignol oxidation and participate to lignification (**Figure 3**). PRXs involved in lignification are exported to the apoplasm in lignifying tissues where monolignol oxidation takes place (Koutaniemi et al., 2005; Vanholme et al., 2010). The oxidation reaction uses H₂O₂ as co-substrate which is probably provided by combined action of NADPH oxidase or germin-like proteins, even though firm genetic evidence for the involvement of these enzymes is currently lacking (Davidson et al., 2009). PRX exist as large multigene families (73 and 138 in *A. thaliana* and *Oryza sativa*, respectively; Welinder et al., 2002) making the clarification of the biological role of each member a considerable challenge. Clear characterization is also complicated by the fact that PRXs generally show low substrate specificities (Hiraga et al., 2001). Nevertheless recent studies suggesting that some PRX isoforms may be more specialized in the polymerization of guaiacyl or/and syringyl units could help to explain how plants are able to control lignin monomeric structure (Barcelo et al., 2007; Martinez-Cortes et al., 2012). Both the isoelectric point of the proteins and specific motifs in the PRX primary structure could support protein-substrate interaction with one monomer type over another and therefore influence lignin composition. Two examples of genetic manipulation illustrate this hypothesis. Antisense suppression of one PRX gene led to a global reduction of both G and S units in tobacco (Blee et al., 2003) whereas analyses of PRX down-regulated poplar (Li et al., 2003) revealed that lignin contained less G units but that S content was unchanged. More recently, Herrero et al. (2013a) used *Zinnia elegans* PRX protein sequence data to identify *Arabidopsis* orthologs for functional characterization. Histochemical analyses of the *atprx72* mutant suggest that it has reduced overall lignin levels and a low syringyl unit content as compared to WT plants (Herrero et al., 2013b). However, the mutant shows a strong developmental phenotype and it would be necessary to confirm this result on a second mutant allele. In parallel, Shigeto et al. (2013) used similar strategies to functionally characterize other PRX gene candidates in

<figure><img src="image_6.png" /><figcaption>**FIGURE 3** | Oxidative radicalization of monolignols by peroxidases and laccases. Peroxidases use peroxide produced by super oxide dismutase proteins (SOD) and NADPH oxidase as co-substrate to make oxidative</figcaption></figure>

radicalization of phenols (coniferyl alcohol in this case) whereas laccases use oxygen. Note that peroxidase can oxidize both monolignols and oligomers to produce oxidized phenols.
<figure><img src="image_7.png" /><figcaption>**FIGURE 4** | Mechanism of phenol oxidation and polymerization in the cell wall. Dotted lines show transfert of radicals between phenols (monolignol or oligomer). White arrows represent combinatorial radical coupling between phenols. Dark arrows show the product of radical-radical coupling and the oxidation of a phenol.</figcaption></figure>

*Arabidopsis*. Chemical analyses indicated a significant decrease in the total lignin content and modified lignin structure in *atprx2* and *atprx25* mutants and altered lignin structure in *atprx71* mutants.

The mechanism of oxidative radicalization of phenols followed by combinatorial radical coupling is well described (Vanholme et al., 2010; van Parijs et al., 2010). Nevertheless, the kinetics of monomer incorporation into the lignin polymer by PRX remains poorly understood. In a recent article, Demont-Caulet et al. (2010) propose a general scheme for S–G copolymerization coordinated by a PRX (PRX34) purified from *A. thaliana*. The purified enzyme shows a high substrate specificity and seems unable to directly oxidize S monomers in the absence of G units. The authors studied kinetics of the monomer conversion and subsequent formation of di-, tri-, and tetramers by using monomer feeding experiments. Their results suggest that endwise and bulk polymerization mechanisms are likely to co-exist within the same system, the bulk mechanisms occurring after total consumption of the monomers. During endwise polymerization, monomers are added to the polymer one-by-one whereas during bulk polymerization different

oligomers are linked together at the same time (Figure 4). Interestingly, Demont-Caulet et al. (2010) show that while monomer availability may influence the relative preponderance of the two mechanisms, it is the nature of the monomer that exerts the most influence. Indeed, under *in vitro* oxidation conditions, the presence of S monomer converted the PRX34-mediated polymerization of G monomers from a bulk to an endwise mechanism in agreement with data from Argyropoulos et al. (2002). These observations underline the central role of PRXs in controlling lignin biosynthesis. In a recent article, the group of Niko Geldner proposed a very nice model where PRXs are the last pieces of a puzzle that orchestrate lignification in Casparian strips in *Arabidopsis* roots (Lee et al., 2013). The authors show that the PRX enzymes colocalize with the very specific Casparian strip domain protein (CASP) and that their activity is indirectly under the control of an NADPH oxidase named respiratory burst oxidase homolog F protein (RBOHF) that provides H₂O₂ to PRXs via the action of an unknown superoxide dismutase (SOD). These fascinating results show for the first time that a *trans*-membrane protein
(CASP) is able to control peroxidase activities thereby representing a potential mechanism for localized spatial control of lignification.

Therefore, PRXs represent interesting targets for modifying not only lignin content but also lignin structure, even if high redundancy due to the existence of large gene families compli- cates genetic approaches. It is also important to emphasize that PRXs exported to the extracellular matrix of some tissues may not only be involved in lignification. For instance, they may par- ticipate to suberin polymerization, cell membrane protection or else, catalyze di-ferulate bonds between polysaccharides particu- larly in grasses where arabinoxylans are highly feruloylated (Fry, 2004; Arrieta-Baez and Stark, 2006). Tannins that contribute to the heartwood color in some trees (e.g., walnut), as well as lignans (see above) may also be produced by PRXs.

Laccases are a large group of copper-containing glycopro-
teins in plants, bacteria, and fungi. In contrast to PRXs, LACs
use O₂ instead of H₂O₂ to oxidize the monolignols **(Figure 3)**.
Although there is not enough evidence to confirm that plant
LACs are localized in the cell wall, the presence of signal pep-
tides in the N-terminal protein region suggests a subcellular
localization (Cesarino et al., 2013) whereas some PRXs could be
detected in other compartments. A number of *in vitro* experi-
ments have identified different putative LAC substrates including
flavonoids, 4-Hydroxyindole, coniferyl alcohol, *p*-coumaryl alco-
hol, and sinapyl alcohol (Mayer and Staples, 2002). While certain
studies confirm that tree LACs can polymerize monolignols *in vitro*
(Sterjiades et al., 1992; Ranocha et al., 2002), the broad substrate
specificity of plant LAC needs to be confirmed *in vivo*. LACs, like
PRXs, belong to (smaller) multigenic families in plants and differ-
ent members may have redundant or specific biological functions.
This is based upon two contrasting observations showing that
in single antisense poplar plants with reduced LAC transcripts,
neither lignin content nor composition is significantly altered
(Ranocha et al., 2002). In contrast, the transgenic poplars exhib-
ited an increase in total soluble phenolic content and perturbation
of xylem fiber integrity (Ranocha et al., 2002). In *Arabidopsis*, 17
LACs have been reported (McCaig et al., 2005) but until 2011,
only indirect evidence linked these enzymes to lignification in
*Arabidopsis*. *AtLAC2* mutants showed a reduced root elongation
under PEG treatment suggesting a possible impact on vascula-
ture (Cai et al., 2006) and *AtLAC4* mutants under continuous
light conditions were associated with a weak irregular xylem phe-
notype (Brown et al., 2005). Similarly, the *Arabidopsis* *AtLAC15*
mutant was identified during a transparent testa screen since it
possesses seeds with a pale brown seed coat due to altered flavonoid
condensation (Pourcel et al., 2005). Mutants also showed a 59%
increase of soluble proanthocyanidins when compared to wild-
type and the extractable lignin content in *tt10* seeds was reduced
by nearly 30% (Egertsdotter et al., 2004). This observation would
suggest that both monolignols and proanthocyanidins could serve
as substrates for AtLAC15 in *Arabidopsis* seed coats. Transcrip-
tomics (Ehling et al., 2005) indicate that four LACs (*AtLAC4*,
*AtLAC11*, *AtLAC12*, *AtLAC17*) show moderate or high expression
levels in *Arabidopsis* developing stems. These four genes are up-
regulated during the transition from young to intermediate stage
stems and were therefore considered as good targets for modifying
lignin.

Direct evidence showing that specific LACs are involved in
monolignol polymerization *in planta* was obtained by the charac-
terization of *Arabidopsis lac4lac17* double mutants (Berthet et al.,
2011). The stems of double mutants possess collapsed vessels
and show reduced G lignin deposition in fibers. Mutants also
show a 20–40% decrease in total lignin content and a 130% increase in saccharification yields when compared to the wild-type
control. Lignin composition (higher S/G ratio) is also modi-
fied and is mainly due to the decrease of G lignin in fibers.
This is not surprising since LAC and lignin genes are tightly
co-expressed (Berthet et al., 2011) and promoter-GUS analyses
showed that *LAC4* is expressed in vascular bundles and interfas-
cicular fibers of stems while *LAC17* is specifically expressed in
fibers (Berthet et al., 2011). Phylogenetic analyses of *Arabidop-
sis* LACs showed that *LAC4* and *LAC17* are the best homologs
(McCaig et al., 2005) suggesting redundant or at least similar
activity for both proteins in lignification. Recently, Cesarino
et al. (2013) identified a sugarcane LAC gene, *SofLAC*, putatively
involved in lignification. Its transcripts are localized in the inner-
and outer-lignified zones of stem sclerenchymatic bundle sheaths.
*SofLAC* was expressed preferentially in young internodes and the
corresponding transcript level decreased along with stem matu-
ryity. The *Arabidopsis lac17* mutant was partly complemented by
expressing *SofLAC* under the control of the AtLAC17 promoter.
*SofLAC* expression restored the lignin content but not the lignin
composition in complemented *lac17* mutant lines. These results
suggest that specific LACs may also play a role in lignification in
monocots.

Studies on mutants affected in different specific steps of
monolignol biosynthesis indicate that plants are able to incor-
porate substantial amounts of components (e.g., aldehydes,
5-hydroxyconiferyl alcohols) that are poorly incorporated in “nor-
mal” conditions. Currently, it is not known whether incorporation
is mediated by PRXs or LACs, or by both proteins. Some attempts
have also been made to engineer plants so as to incorporate
unusual compounds (e.g., rosmarinic acid or hydroxybenzalde-
hyde and hydroxybenzoate derivatives) into their lignins (Eudes
et al., 2012; Tobimatsu et al., 2012). The incorporation of such
specific compounds has been proposed as a strategy to reduce the
degree of lignin polymerization. The incorporation of novel com-
pounds together with the existence of large LAC and PRX gene
families showing broad substrate specificities raises exciting new
possibilities for the production of modified lignins and improved
biomass.

ACYLATION

Plant lignins often contain various acids (acetates, *p*-hydroxybenzoate, and *p*-coumarate) in addition to conventional monolignols. It is usually accepted that some of them (ferulate, *p*-coumarate) are by-products or intermediates of the monolignol pathway since there is no evidence that an independent pathway exists. Several observations suggest that the degree of acylation could be reasonably expected to impact on biomass quality. First, amounts of acylated monomers show high natural variations between different species. Kenaf and Agave possess up to 60 and 80% of acylated lignins, respectively when compared to maize and wheat that show 18 and 3% of acylated lignins, respectively
(reviewed by Withers et al., 2012). Second, it is possible that acylation may affect lignin structure during polymerization as well as subsequent degradation by chemical or biochemical processes such as saccharification and digestibility. In grasses, *p*-coumarates are linked to syringyl units and plants with low coumarate levels also generally show modifications in their S, or S and G content (Harrington et al., 2012). Importantly, coumaroylation of cell wall polysaccharides is most likely (or at least partly) catalyzed in the cytosol prior to export and polymerization in the cell wall strongly suggesting the existence of specific monolignol transferases and transporters (Withers et al., 2012; Molinari et al., 2013; **Figure 1**).

Until recently, the genetics underlying coumarylation of both
lignin and arabinoxylans remained unknown, mainly because
of the lack of suitable plants (specifically engineered to modify
acylation) for functional characterization. It is generally accepted

that the genes associated with such reactions would encode BAHD
acyl-CoA transferase proteins. Recent work showing that trans-
genics have higher or lower feruloylation levels has suggested
that some members of the BAHD acyl-CoA transferase protein
family may be responsible for catalyzing the addition of ester-
linked coumarate to cell wall arabinoxylans (Fernando et al., 2009;
Withers et al., 2012; Bartley et al., 2013; Molinari et al., 2013). A
member of the BAHD acyl transferase family was recently iden-
tified in the rice genome (Withers et al., 2012). Analyses of the
recombinant protein produced in *E. coli* showed that it had a
specific coumaroyl transferase activity. This enzyme OsPMT (*O.
sativa* *p*-coumaric acid monolignol transferase) catalyzes the acy-
lation of a monolignol with *p*-coumaric acid via *p*-coumaroyl
CoA and the corresponding transcript is co-expressed with the
monolignol biosynthetic 4CL gene. Based on catalytic efficiency

<figure><img src="image_8.png" /><figcaption>**FIGURE 5** | **Co-expression network of genes involved in lignin and lignan biosynthesis in Arabidopsis.** (A) Dendrogram representing the co-expression of genes involved in the biosynthesis and polymerization of monolignols. Two separate clusters (II and I, respectively) are obtained using the tool "expression tree" from Genecat (http://genecat.mpg.de). (B) Co-expressed gene network of genes present in cluster II. (C) Co-expressed gene network of genes present in cluster I. Nodes and edges represent genes and significant positive co-expression relationships between genes. Red nodes represent genes used as baits to produce gene co-expression</figcaption></figure>

networks. Blue nodes depict genes known to be involved in monolignol
biosynthesis. Green nodes represent specific genes involved uniquely in
cellulose synthesis of secondary cell walls. Orange nodes represent genes
putatively involved in polymerization and lignan formation. DIR6, dirigent
protein 6; LAC4, laccase 4; LAC17, laccase 17; CCR1, hydroxycinnamoyl-CoA
reductase 1; C4H, cinnamic acid 4-hydroxylase; 4Cl, 4-hydroxycinnamoyl-CoA
ligase 1; CCoAOMT1, caffeoyl-CoA O-methyltransferase; PLR,
pinoresinol-lariciresinol reductase; PER64, peroxidase 64; CESAA4, 7, 8,
cellulose synthase 4, 7, and 8. Other abbreviations are given in **Figure 1**.
and reaction rate, the authors show that OsPMT would pro-
duce more *p*-coumaryl *p*-coumarates in saturating conditions but
would favor the synthesis of sinapyl *p*-coumarate over coniferyl
*p*-coumarate *in planta*. Interestingly this gene is grass specific and
its enzymatic hallmark therefore perfectly correlates with the high
level of coumaroylated sinapyl alcohols in these types of plant.
In addition, the BAHD protein accepts coumarate, but not feru-
late as a substrate. The production of rice plants with disrupted
OsPMT expression may be difficult because of gene redundancy
since there are at least two paralogs in this species. Nevertheless, it
is interesting to note that there are less paralogs in *Brachypodium*
(Withers et al., 2012) and the recent article published (Molinari
et al., 2013) could help to identify suitable candidate genes for
further study. The authors established a correlation between candi-
date *p*-coumaric acid monolignol transferase transcript levels and
amounts of bound ferulate in different organs at different devel-
opmental stages. They propose different candidate genes similar to
OsPMT genes to be responsible for the lignin bound *p*-coumaric
acid.

CO-EXPRESSION DATA MAY HELP TO SCREEN FOR
PROTAGONIST INVOLVED IN THE LAST STEPS OF
LIGNIN BIOSYNTHESIS

The last 20 years have seen the elucidation of the complete mono-
lignol biosynthetic pathway and a number of limiting enzymes
have been identified. Importantly, the expression of monolignol
biosynthesis genes is not always correlated with the presence of
the lignin polymer. This observation can be attributed to the fact
that the monomers can also be used to generate a wide range of
monolignol-derived compounds located in intra- or extracellular
compartments and underlines the complexity of plant monolig-
nol metabolism and regulation (Figure 1). One illustration of
this phenomenon can be seen with the use of co-expression tools.
Recent years have seen the emergence of large transcriptomic
data sets and co-expression tools have been developed by differ-
ent research groups (Ruprecht and Persson, 2012). These tools
allow the identification of genes putatively involved in the same
biological process as, in general, the expression of related genes
should be transcriptionally coordinated (Usadel et al., 2009). Two
recent articles have used this approach to analyze lignin biosynthe-
sis in *Arabidopsis* and *Brachypodium* (Ruprecht and Persson, 2012).
When monolignol biosynthesis genes (*4CL*, *C4H*, *CCoAOMT*) and
monolignol polymerization/isomerization genes (*Dirigent protein*

REFERENCES

Alejandro, S., Lee, Y., Tohge, T., Sudre, D., Osorio, S., Park, J., et al. (2012). AtABCG29 is a monolignol transporter involved in lignin biosynthesis. *Curr. Biol.* 22, 1207–1212. doi: 10.1016/j.cub.2012.04.064

Argyropoulos, D. S., Jurasek, L., Kristo-
fova, L., Xia, Z., Sun, Y., and Palus, E.
(2002). Abundance and reactivity of
dibenzodioxocins in softwood lignin.
*J. Agric. Food Chem.* **50**, 658–666. doi:
**10.1021/jf010909g**

Arrieta-Baez, D., and Stark, R. E. (2006). Modeling suberization with peroxidase-catalyzed polymerization

of hydroxycinnamic acids: cross-
coupling and dimerization reactions.
*Phytochemistry* **67**, 743–753. doi:
**10.1016/j.phytochem.2006.01.026**

Attoumbre, J., Bienaime, C., Dubois,
F., Fliniaux, M. A., Chabbert, B.,
and Baltora-Rosset, S. (2010). Devel-
opment of antibodies against sec-
oisolariciresinol – application to the
immunolocalization of lignans in
*Linum usitatissimum* seeds. *Phy-
tochemistry* **71**, 1979–1987. doi:
**10.1016/j.phytochem.2010.09.002**

Baghdady, A., Blervacq, A. S., Jouanin,
L., Grima-Pettenati, J., Sivadon, P.,
and Hawkins, S. (2006). *Eucalyptus*

6, LAC4, LAC17) are used as baits for the co-expression analysis with genecat (http://genecat.mpg.de/), two distinct gene networks clearly appear (Figure 5). LAC genes are clearly co-expressed with the cellulose synthase genes CESA4, 7, and 8, that are specific to secondary-, but not primary-cell wall formation. This co-expression is even stronger than that observed with 4CL, C4H, and CCoAOMT that are all highly co-expressed with other monolignol biosynthetic genes such as PAL, COMT, CAD, CCR, C3H, and HCT (Figure 5). Since the monolignol biosynthetic pathway shares steps with the general phenylpropanoid pathway involved in the production of various non-lignin compounds (Goujon et al., 2003; Bassard et al., 2010), the co-expression data could suggest that LAC4 and LAC17 might represent a more efficient bait than monolignol biosynthesis genes when trying to identify other “lignin-related” genes including those encoding transporters and other polymerization enzymes.

CONCLUSION

It is clear that the metabolism and polymerization of both
monolignols and other more unusual monomers following their
biosynthesis are more complex than previously thought. A number
of key potential regulation points are involved in these processes
allowing plants to rapidly respond to different developmental
and environmental cues and that could also constitute interest-
ing novel engineering targets for optimizing lignin production
and composition. This will be achieved by answering a long list of
crucial questions. For instance, where are glucosidases located –
in the cell wall, in the vacuole or in the cytosol? In which
conditions (development, stress, etc.) are they active? How are
glycosylated monolignols released from the vacuole? What are
their roles? Where does the dimerization occur for lignan for-
mation? Are cytosolic PRXs involved? How do lignin polymers
grow in the cell wall? What are the specific roles of LACs ver-
sus PRXs? Are CASP-like proteins also present in other lignified
tissues (xylem and sclerenchyma for instance)? The regulation
of lignification has long been studied and modified by scien-
tists at the monolignol biosynthetic level only, often with the
sole goal of reducing lignin contents. It is now becoming clear
that many other important control points exist that will allow the
development of novel strategies for improving plant biomass for
both energy and material production. We can reasonably expect
that some of these key questions will be answered during the
next decade.

gunnii CCR and CAD2 promoters are active in lignifying cells during primary and secondary xylem formation in *Arabidopsis thaliana*. *Plant Physiol. Biochem.* **44**, 674–683. doi: **10.1016/j.plaphy.2006.10.027**

Barcelo, A. R., Ros, L. V., and Car-
rasco, A. E. (2007). Looking for
syringyl peroxidases. *Trends Plant Sci.*
**12**, 486–491. doi: **10.1016/j.tplants**
**2007.09.002**

Bartley, L. E., Peck, M. L., Kim, S. R., Ebert, B., Maniseri, C., Chiniquy, D., et al. (2013). Overexpression of a BAHD acyltransferase, OsAt10,

alters rice cell wall hydroxycinnamic
acid content and saccharification.
*Plant Physiol.* **161**, 1615–1633. doi:
**10.1104/pp.112.208694**

Bassard, J. E., Ullmann, P., Bernier,
F., and Werck-Reichhart, D. (2010).
Phenolamides: bridging polyamines
to the phenolic metabolism.
*Phytochemistry* **71**, 1808–1824. doi:
**10.1016/j.phytochem.2010.08.003**

Berthet, S., Demont-Caulet, N., Pol-
let, B., Bidzinski, P., Cezard, L., Le
Bris, P., et al. (2011). Disruption of
LACCASE4 and 17 results in tissue-
specific alterations to lignification of
*Arabidopsis thaliana* stems. *Plant Cell*
23, 1124-1137. doi: 10.1105/tpc.110.082792

Blee, K. A., Choi, J. W., O'Connell, A. P., Schuch, W., Lewis, N. G., and Bolwell, G. P. (2003). A lignin-specific peroxidase in tobacco whose antisense suppression leads to vascular tissue modification. *Phytochemistry* 64, 163-176. doi: 10.1016/S0031-9422(03)00212-7

Boerjan, W., Ralph, J., and Baucher, M. (2003). Lignin biosynthesis. *Annu. Rev. Plant Biol.* 54, 519-546. doi: 10.1146/annurev.arplant.54.031902.134938

Brown, D. M., Zeef, L. A., Ellis, J., Goodacre, R., and Turner, S. R. (2005). Identification of novel genes in *Arabidopsis* involved in secondary cell wall formation using expression profiling and reverse genetics. *Plant Cell* 17, 2281-2295. doi: 10.1105/tpc.105.031542

Burmeister, G., and Hosel, W. (1981). Immunofluorometrische Lokalisierung von beta-Glucosidase in *Cicer arietinum* L. Keimlingen. *Planta Med.* 42, 102. doi: 10.1055/s-2007-971553

Cai, X., Davis, E. J., Ballif, J., Liang, M., Bushman, E., Haroldsen, V., et al. (2006). Mutant identification and characterization of the laccase gene family in *Arabidopsis*. *J. Exp. Bot.* 57, 2563-2569. doi: 10.1093/jxb/erl022

Caputi, L., Malnoy, M., Goremykin, V., Nikiforova, S., and Martens, S. (2012). A genome-wide phylogenetic reconstruction of family 1 UDP-glycosyltransferases revealed the expansion of the family during the adaptation of plants to life on land. *Plant J.* 69, 1030-1042. doi: 10.1111/j.1365-313X.2011.04853.x

Cesarino, I., Araújo, P., Sampaio Mayer, J. L., Vicentini, R., Berthet, S., Demedts, B., et al. (2013). Expression of SofLAC, a new laccase in sugarcane, restores lignin content but not S:G ratio of *Arabidopsis* lac17 mutant. *J. Exp. Bot.* 64, 1769-1781.

Chapelle, A., Morreel, K., Vanholme, R., Le-Bris, P., Morin, H., Lapierre, C., et al. (2012). Impact of the absence of stem-specific beta-glucosidases on lignin and monolignols. *Plant Physiol.* 160, 1204-1217. doi: 10.1104/pp.112.203364

Chapple, C., Ladisch, M., and Meilan, R. (2007). Loosening lignin's grip on biofuel production. *Nat. Biotechnol.* 25, 746-748. doi: 10.1038/nbt0707-746

Davidson, R. M., Reeves, P. A., Manosalva, P. M., and Leach, J. E. (2009). Germins: a diverse protein

family important for crop improvement. *Plant Sci.* 177, 499-510. doi: 10.1016/j.plantsci.2009.08.012

Davin, L. B., Wang, H. B., Crowell, A. L., Bedgar, D. L., Martin, D. M., Sarkanen, S., et al. (1997). Stereoselective bimolecular phenoxy radical coupling by an auxiliary (dirigent) protein without an active center. *Science* 275, 362-366. doi: 10.1126/science.275.5298.362

Demont-Caulet, N., Lapierre, C., Jouanin, L., Baumberger, S., and Mechlin, V. (2010). *Arabidopsis* peroxidase-catalyzed copolymerization of coniferyl and sinapyl alcohols: kinetics of an endwise process. *Phytochemistry* 71, 1673-1683. doi: 10.1016/j.phytochem.2010.06.011

Dharmawardhana, D. P., Ellis, B. E., and Carlson, J. E. (1995). A beta-glucosidase from lodgepole pine xylem specific for the lignin precursor coniferin. *Plant Physiol.* 107, 331-339. doi: 10.1104/pp.107.2331

Dharmawardhana, D. P., Ellis, B. E., and Carlson, J. E. (1999). cDNA cloning and heterologous expression of coniferin beta-glucosidase. *Plant Mol. Biol.* 40, 365-372. doi: 10.1023/A:1006226931512

Dixon, R. A., Chen, F., Guo, D., and Parvathi, K. (2001). The biosynthesis of monolignols: a "metabolic grid", or independent pathways to guaiacyl and syringyl units? *Phytochemistry* 57, 1069-1084. doi: 10.1016/S0031-9422(01)00092-9

Dixon, R. A., Lamb, C. J., Masoud, S., Sewalt, V. J., and Paiva, N. L. (1996). Metabolic engineering: prospects for crop improvement through the genetic manipulation of phenylpropanoid biosynthesis and defense responses - a review. *Gene* 179, 61-71. doi: 10.1016/S0378-1119(96)00327-7

Donaldson, L. A. (2001). Lignification and lignin topochemistry - an ultrastructural view. *Phytochemistry* 57, 859-873. doi: 10.1016/S0031-9422(01)00049-8

Egertsdotter, U., Van Zyl, L. M., Mackay, J., Peter, G., Kirst, M., Clark, C., et al. (2004). Gene expression during formation of earlywood and latewood in loblolly pine: expression profiles of 350 genes. *Plant Bioi. (Stuttg.)* 6, 654-663. doi: 10.1055/s-2004-830383

Ehling, J., Mattheus, N., Aeschliman, D. S., Li, E., Hamberger, B., Cullis, I. F., et al. (2005). Global transcript profiling of primary stems from *Arabidopsis thaliana* identifies candidate genes for missing links in lignin biosynthesis and transcriptional regulators of fiber differentiation. *Plant

J. 42, 618-640. doi: 10.1111/j.1365-313X.2005.02403.x

Escamilla-Trevino, L. L., Chen, W., Card, M. L., Shih, M. C., Cheng, C. L., and Poulton, J. E. (2006). *Arabidopsis thaliana* beta-glucosidases BGLU45 and BGLU46 hydrolyse monolignol glucosides. *Phytochemistry* 67, 1651-1660. doi: 10.1016/j.phytochem.2006.05.022

Eudes, A., George, A., Mukerjee, P., Kim, J. S., Pollet, B., Benke, P. I., et al. (2012). Biosynthesis and incorporation of side-chain-truncated lignin monomers to reduce lignin polymerization and enhance saccharification. *Plant Biotechnol. J.* 10, 609-620. doi: 10.1111/j.1467-7652.2012.00692.x

Fernando, P., Cristobal, U., Lianhai, F., James, L., John, L., and Jorge, D. (2009). Down-regulation of four putative arabinoxylan feruloyl transferase genes from family PF02458 reduces ester-linked flucate content in rice cell walls. *Planta* 231, 677-691. doi: 10.1007/s00425-009-1077-1

Fincher, G. B. (2009). Exploring the evolution of (1,3;1,4)-beta-D-glucans in plant cell walls: comparative genomics can help! *Curr. Opin. Plant Biol.* 12, 140-147. doi: 10.1016/j.pbi.2009.01.002

Ford, J. D., Huang, K. S., Wang, H. B., Davin, L. B., and Lewis, N. G. (2001). Biosynthetic pathway to the cancer chemopreventive secoisolariciresinol diglucoside-hydroxymethyl glutaryl ester-linked lignan oligomers in flax (*Linum usitatissimum*) seed. *J. Nat. Prod.* 64, 1388-1397. doi: 10.1021/np010367x

Freudenberg, K. (1959). Biosynthesis and constitution of lignin. *Nature* 183, 1152-1155. doi: 10.1038/1831152a0

Fromm, J., Rockel, B., Lautner, S., Windeisen, E., and Wanner, G. (2003). Lignin distribution in wood cell walls determined by TEM and backscattered SEM techniques. *J. Struct. Biol.* 143, 77-84. doi: 10.1016/S1047-8477(03)00119-9

Fry, S. C. (2004). Primary cell wall metabolism: tracking the careers of wall polymers in living plant cells. *New Phytol.* 161, 641-675. doi: 10.1111/j.1469-8137.2004.00980.x

Gang, D. R., Costa, M. A., Fujita, M., Dinkova-Kostova, A. T., Wang, H. B., Burlat, V., et al. (1999). Regiochemical control of monolignol radical coupling: a new paradigm for lignin and lignan biosynthesis. *Chem. Biol.* 6, 143-151. doi: 10.1016/S1074-5521(99)89006-1

Goujon, T., Sibout, R., Pollet, B., Maba, B., Nussaume, L., Bechtold, N., et al. (2003). A new *Arabidopsis thaliana* mutant deficient in the expression of O-methyltransferase impacts lignins and sinapoyl esters. *Plant Mol. Biol.* 51, 973-989. doi: 10.1023/A:1023022825098

Hano, C., Martin, I., Fliniaux, O., Legrand, B., Gutierrez, L., Arroo, R. R., et al. (2006). Pinoresinol-lariciresinol reductase gene expression and secoisolariciresinol diglucoside accumulation in developing flax (*Linum usitatissimum*) seeds. *Planta* 224, 1291-1301. doi: 10.1007/s00425-006-0308-y

Harrington, M. J., Mutwil, M., Barrière, Y., and Sibout, R. (2012). "Chapter 3 – Molecular biology of lignification in grasses," in *Advances in Botanical Research*, eds J. Lise and L. Catherine (Waltham: Academic Press), 77-112.

Hemmati, S., Von Heimendahl, C. B., Klaes, M., Alfermann, A. W., Schmidt, T. J., and Fuss, E. (2010). Pinoresinol-lariciresinol reductases with opposite enantiospecificity determine the enantiomeric composition of ligands in the different organs of *Linum usitatissimum* L. *Planta Med.* 76, 928-934. doi: 10.1055/s-0030-1250036

Herrero, J., Esteban-Carrasco, A., and Zapata, J. M. (2013a). Looking for *Arabidopsis thaliana* peroxidases involved in lignin biosynthesis. *Plant Physiol. Biochem.* 67C, 77-86. doi: 10.1016/j.plaphy.2013.02.019

Herrero, J., Fernandez-Perez, F., Yebra, T., Novo-Uzal, E., Pomar, F., Pedreno, M. A., et al. (2013b). Bioinformatic and functional characterization of the basic peroxidase 72 from *Arabidopsis thaliana* involved in lignin biosynthesis. *Planta* 237, 1599-1612. doi: 10.1007/s00425-013-1865-5

Higuchi, T. (1990). Lignin biochemistry: biosynthesis and biodegradation. *Wood Sci. Technol.* 24, 23-63. doi: 10.1007/BF00225306

Hiraga, S., Sasaki, K., Ito, H., Ohashi, Y., and Matsui, H. (2001). A large family of class III plant peroxidases. *Plant Cell Physiol.* 42, 462-468. doi: 10.1093/pcp/pce061

Hosel, W., Surholt, E., and Borgmann, E. (1978). Characterization of beta-glucosidase isoenzymes possibly involved in lignification from chick pea (*Cicer arietinum* L.) cell suspension cultures. *Eur. J. Biochem.* 84, 487-492. doi: 10.1111/j.1432-1033.1978.tb12190.x

Huis, R., Morreel, K., Fliniaux, O., Lucau-Danila, A., Fenart, S., Grec,
S., et al. (2012). Natural hypolignification is associated with extensive oligolignol accumulation in flax stems. *Plant Physiol.* 158, 1893-1915. doi: 10.1104/pp.111.192328

Kim, Y. H., Kim, C. Y., Song, W. K., Park, D. S., Kwon, S. Y., Lee, H. S., et al. (2008). Overexpression of sweetpotato swpa4 peroxidase results in increased hydrogen peroxide production and enhances stress tolerance in tobacco. *Planta* 227, 867-881. doi: 10.1007/s00425-007-0663-3

Koutaniemi, S., Toikka, M. M., Karkonen, A., Mustonen, M., Lundell, T., Simola, L. K., et al. (2005). Characterization of basic *p*-coumaryl and coniferyl alcohol oxidizing peroxidases from a lignin-forming *Picea abies* suspension culture. *Plant Mol. Biol.* 58, 141-157. doi: 10.1007/s11103-005-5345-6

Lanot, A., Hodge, D., Jackson, R. G., George, G. L., Elias, L., Lim, E. K., et al. (2006). The glucosyltransferase UGT72E2 is responsible for monolignol 4-O-glucoside production in *Arabidopsis thaliana*. *Plant J.* 48, 286-295. doi: 10.1111/j.1365-313X.2006.02872.x

Lee, Y. C., Kim, M. I., Woo, M. A., Park, H. G., and Han, J. I. (2013). Effective peroxidase-like activity of a water-solubilized Fe-aminoclay for use in immunoassay. *Biosens. Bioelectron.* 42, 373-378. doi: 10.1016/j.bios. 2012.10.092

Li, Y., Kajita, S., Kawai, S., Katayama, Y., and Morohoshi, N. (2003). Downregulation of an anionic peroxidase in transgenic aspen and its effect on lignin characteristics. *J. Plant Res.* 116, 175-182. doi: 10.1007/s10265-003-0087-5

Lim, E. K., and Bowles, D. J. (2004). A class of plant glycosyltransferases involved in cellular homeostasis. *EMBO J.* 23, 2915-2922. doi: 10.1038/sj.emboj. 7600295

Liu, C. J. (2012). Deciphering the enigma of lignification: precursor transport, oxidation, and the topochemistry of lignin assembly. *Mol. Plant* 5, 304-317. doi: 10.1093/ mp/ssr121

Liu, C. J., Miao, Y. C., and Zhang, K. W. (2011). Sequestration and transport of lignin monomeric precursors. *Molecules* 16, 710-727. doi: 10.3390/molecules16010710

MacKay, J. J., Liu, W., Whetten, R., Sederoff, R. R., and O'Malley, D. M. (1995). Genetic analysis of cinnamyl alcohol dehydrogenase in loblolly pine: single gene inheritance, molecular characterization and evolution.

Mol. Gen. Genet. 247, 537-545. doi: 10.1007/BF00290344

Marcinowski, S., and Grisebach, H. (1978). Enzymology of lignification. Cell-wall bound beta-glucosidase for coniferin from spruce (*Picea abies*) seedlings. *Eur. J. Biochem.* 87, 37-44. doi: 10.1111/j.1432-1033.1978.tb12349.x

Martinez-Cortes, T., Pomar, F., Espineira, J. M., Merino, F., and Novo-Uzal, E. (2012). Purification and kinetic characterization of two peroxidases of *Selaginella martensii* Spring. involved in lignification. *Plant Physiol. Biochem.* 52, 130-139. doi: 10.1016/j.plaphy. 2011.12.008

Martone, P. T., Estevez, J. M., Lu, F., Ruel, K., Denny, M. W., Somerville, C., et al. (2009). Discovery of lignin in seaweed reveals convergent evolution of cell-wall architecture. *Curr. Biol.* 19, 169-175. doi: 10.1016/j.cub.2008.12.031

Mayer, A. M., and Staples, R. C. (2002). Laccase: new functions for an old enzyme. *Phytochemistry* 60, 551-565. doi: 10.1016/S0031-9422(02)00171-1

McCaig, B. C., Meagher, R. B., and Dean, J. F. (2005). Gene structure and molecular analysis of the laccase-like multicopper oxidase (LMCO) gene family in *Arabidopsis thaliana*. *Planta* 221, 619-636. doi: 10.1007/s00425-004-1472-6

Miao, Y. C., and Liu, C. J. (2010). ATP-binding cassette-like transporters are involved in the transport of lignin precursors across plasma and vacuolar membranes. *Proc. Natl. Acad. Sci. U.S.A.* 107, 22728-22733. doi: 10.1073/pnas.1007747108

Molinari, H. B. C., Pellny, T. K., Freeman, J., Shewry, P. R., and Mitchell, R. A. (2013). Grass cell wall feruloylation: distribution of bound ferulate and candidate gene expression in *Brachypodium distachyon*. *Front. Plant Sci.* 4:50. doi: 10.3389/fpls.2013.00050

Morant, A. V., Jorgensen, K., Jorgensen, C., Paquette, S. M., Sanchez-Perez, R., Moller, B. L., et al. (2008). beta-Glucosidases as detonators of plant chemical defense. *Phytochemistry* 69, 1795-1813. doi: 10.1016/j.phytochem.2008.03.006

Neish, A. C. (1961). Configuration of beta-phenyllactic acids (2-hydroxy-3-phenyl-propionic acid). *Can. J. Biochem. Physiol.* 39, 1205-1206. doi: 10.1139/o61-124

Noguchi, A., Fukui, Y., Iuchi-Okada, A., Kakutani, S., Satake, H., Iwashita, T., et al. (2008). Sequential glucosylation of a furofuran lignan,

(+)-sesaminol, by *Sesamum indicum* UGT71A9 and UGT94DI glucosyltransferases. *Plant J.* 54, 415-427. doi: 10.1111/j.1365-313X.2008.03428.x

Ono, E., Kim, H. J., Murata, J., Morimoto, K., Okazawa, A., Kobayashi, A., et al. (2010). Molecular and functional characterization of novel furofuran-class lignan glucosyltransferases from *Forsythia*. *Plant Biotechnol.* 27, 317-324. doi: 10.5511/plantbiotechnology.27.317

Pesquet, E., Zhang, B., Gorzsas, A., Puhakainen, T., Serk, H., Escamez, S., et al. (2013). Non-cell-autonomous postmortem lignification of tracheary elements in *Zinnia elegans*. *Plant Cell* 25, 1314-1328. doi: 10.1105/tpc.113.110593

Pohjamo, S. P., Hemming, J. E., Willfor, S. M., Reunanen, M. H., and Holmbom, B. R. (2003). Phenolic extractives in *Salix caprea* wood and knots. *Phytochemistry* 63, 165-169. doi: 10.1016/S0031-9422(03)00050-5

Pourcel, L., Routaboul, J. M., Kerhoas, L., Caboche, M., Lepiniec, L., and Debeaujon, I. (2005). TRANSPARENT TESTA10 encodes a laccase-like enzyme involved in oxidative polymerization of flavonoids in *Arabidopsis* seed coat. *Plant Cell* 17, 2966-2980. doi: 10.1105/tpc.105.03154

Ralph, J., MacKay, J. J., Hatfield, R. D., O'Malley, D. M., Whetten, R. W., and Sederoff, R. R. (1997). Abnormal lignin in a loblolly pine mutant. *Science* 277, 235-239. doi: 10.1126/science.277.5323.235

Ramos, C. S., and Kato, M. J. (2012). Secondary metabolites from the phloem of *Piper solmsianum* (Piperaceae) in the honeydew of *Edessa meditabunda*. *Phytochem. Anal.* 23, 604-606. doi: 10.1002/pca.2361

Ranocha, P., Chabannes, M., Chamayou, S., Danoun, S., Jauneau, A., Boudet, A. M., et al. (2002). Laccase down-regulation causes alterations in phenolic metabolism and cell wall structure in poplar. *Plant Physiol.* 129, 145-155. doi: 10.1104/pp.010988

Ruprecht, C., and Persson, S. (2012). Co-expression of cell-wall related genes: new tools and insights. *Front. Plant Sci.* 3:83. doi: 10.3389/fpls.2012.00083

Saguez, J., Attoumbré, J., Giordanengo, P., and Baltora-Rosset, S. (2012). Biological activities of lignans and neolignans on the aphid *Myzus persicae* (Sulzer). *Arthropod Plant Interact.* 7, 225-233. doi: 10.1007/s11829-012-9236-x

Schmidt, T. J., Hemmati, S., Klaes, M., Konuklugil, B., Mohagheghzadeh, A., Ionkova, I., et al. (2010). Lignans in flowering aerial parts of *Linum* species - chemodiversity in the light of systematics and phylogeny. *Phytochemistry* 71, 1714-1728. doi: 10.1016/j.phytochem. 2010.06.015

Schmidt, T. J., Klaes, M., and Sendker, J. (2012). Lignans in seeds of *Linum* species. *Phytochemistry* 82, 89-99. doi: 10.1016/j.phytochem.2012.07.004

Shigeto, J., Kiyonaga, Y., Fujita, K., Kondo, R., and Tsutsumi, Y. (2013). Putative cationic cell-wall-bound peroxidase homologues in *Arabidopsis*, AtPrx2, AtPrx25, and AtPrx71, are involved in lignification. *J. Agric. Food Chem.* [Epub ahead of print]. doi: 10.1021/jf400426g

Sibout, R., and Hofte, H. (2012). Plant cell biology: the ABC of monolignol transport. *Curr. Biol.* 22, R533-R535. doi: 10.1016/j.cub.2012.05.005

Sibout, R., Plantegenet, S., and Hardtke, C. S. (2008). Flowering as a condition for xylem expansion in *Arabidopsis* hypocotyl and root. *Curr. Biol.* 18, 458-463. doi: 10.1016/j.cub.2008.02.070

Sjöström, E. (1993). *Wood Chemistry: Fundamentals and Applications*. San Diego: Academic Press.

Srivastava, V., Schinkel, H., Witzell, J., Hertzberg, M., Torp, M., Srivastava, M. K., et al. (2007). Downregulation of high-isoelectric-point extracellular superoxide dismutase mediates alterations in the metabolism of reactive oxygen species and developmental disturbances in hybrid aspen. *Plant J.* 49, 135-148. doi: 10.1111/j.1365-313X.2006.02943.x

Sterjiades, R., Dean, J. F., and Eriksson, K. E. (1992). Laccase from sycamore maple (*Acer pseudoplatanus*) polymerizes monolignols. *Plant Physiol.* 99, 1162-1168. doi: 10.1104/pp.99.3.1162

Tobimatsu, Y., Elumalai, S., Grabber, J. H., Davidson, C. L., Pan, X., and Ralph, J. (2012). Hydroxycinnamate conjugates as potential monolignol replacements: in vitro lignification and cell wall studies with rosmarinic acid. *ChemSusChem* 5, 676-686. doi: 10.1002/cscs.201100573

Tsuji, Y., Chen, F., Yasuda, S., and Fukushima, K. (2005). Unexpected behavior of coniferin in lignin biosynthesis of *Ginkgo biloba*. *Planta* 222, 58-69. doi: 10.1007/s00425-005-1517-5

Usadel, B., Poree, F., Nagel, A., Lohse, M., Czedik-Eysenberg, A., and Stitt,
M. (2009). A guide to using MapMan to visualize and compare Omics data in plants: a case study in the crop species, Maize. *Plant Cell Environ.* 32, 1211–1229. doi: 10.1111/j.1365-3040.2009.01978.x

Vander Mijnsbrugge, K., Beeckman, H., De Rycke, R., Van Montagu, M., Engler, G., and Boerjan, W. (2000). Phenylcoumaran benzylic ether reductase, a prominent poplar xylem protein, is strongly associated with phenylpropanoid biosynthesis in lignifying cells. *Planta* 211, 502–509. doi: 10.1007/s004250000326

Vanholme, B., Cesarino, I., Goeminne, G., Kim, H., Marroni, F., Van Acker, R., et al. (2013). Breeding with rare defective alleles (BRDA): a natural *Populus nigra* HCT mutant with modified lignin as a case study. *New Phytol.* 198, 765–776. doi: 10.1111/nph.12179

Vanholme, B., Kast, P., Haegeman, A., Jacob, J., Grunewald, W., and Gheysen, G. (2009). Structural and functional investigation of a secreted chorismate mutase from the plant-parasitic nematode *Heterodera schachtii* in the context of related enzymes from diverse origins. *Mol. Plant Pathol.* 10, 189–200. doi: 10.1111/j.1364-3703.2008.00521.x

Vanholme, R., Demedts, B., Morreel, K., Ralph, J., and Boerjan, W. (2010). Lignin biosynthesis

and structure. *Plant Physiol.* 153, 895–905. doi: 10.1104/pp.110.155119

Vanholme, R., Morreel, K., Darrah, C., Oyarce, P., Grabber, J. H., Ralph, J., et al. (2012a). Metabolic engineering of novel lignin in biomass crops. *New Phytol.* 196, 978–1000. doi: 10.1111/j.1469-8137.2012.04337.x

Vanholme, R., Storme, V., Vanholme, B., Sundin, L., Christensen, J. H., Goeminne, G., et al. (2012b). A systems biology view of responses to lignin biosynthesis perturbations in *Arabidopsis*. *Plant Cell* 24, 3506–3529. doi: 10.1105/tpc.112.102574

van Parijs, F. R., Morreel, K., Ralph, J., Boerjan, W., and Merks, R. M. (2010). Modeling lignin polymerization. *I. Simulation model of dehydrogenation polymers. Plant Physiol.* 153, 1332–1344. doi: 10.1104/pp.110.154468

Venglat, P., Xiang, D., Qiu, S., Stone, S. L., Tibiche, C., Cram, D., et al. (2011). Gene expression analysis of flax seed development. *BMC Plant Biol.* 11:74. doi: 10.1186/1471-2229-11-74

Wang, Y. W., Wang, W. C., Jin, S. H., Wang, J., Wang, B., and Hou, B. K. (2012). Over-expression of a putative poplar glycosyltransferase gene, PtGT1, in tobacco increases lignin content and causes early flowering.

J. Exp. Bot. 63, 2799–2808. doi: 10.1093/jxb/ers001

Welinder, K. G., Justesen, A. F., Kjaersgard, I. V., Jensen, R. B., Rasmussen, S. K., Jespersen, H. M., et al. (2002). Structural diversity and transcription of class III peroxidases from *Arabidopsis thaliana*. *Eur. J. Biochem.* 269, 6063–6081. doi: 10.1046/j.1432-1033.2002.03311.x

Weng, J. K., and Chapple, C. (2010). The origin and evolution of lignin biosynthesis. *New Phytol.* 187, 273–285. doi: 10.1111/j.1469-8137.2010.03327.x

Wingender, J., Neu, T. R., and Flemming, H. C. (1999). *Microbial Extracellular Polymeric Substances: Characterization, Structure and Function*. Berlin: Springer doi: 10.1007/978-3-642-60147-7

Withers, S., Lu, F., Kim, H., Zhu, Y., Ralph, J., and Wilkerson, C. G. (2012). Identification of grass-specific enzyme that acylates monolignols with *p*-coumarate. *J. Biol. Chem.* 287, 8347–8355. doi: 10.1074/jbc.M111.284497

Xu, Z., Escamilla-Trevino, L., Zeng, L., Lalgondar, M., Bevan, D., Winkel, B., et al. (2004). Functional genomic analysis of *Arabidopsis thaliana* glycoside hydrolase family 1. *Plant Mol. Biol.* 55, 343–367. doi: 10.1007/s11103-004-0790-1

Yonekura-Sakakibara, K., Fukushima, A., Nakabayashi, R., Hanada,

K., Matsuda, F., Sugawara, S., et al. (2012). Two glycosyltransferases involved in anthocyanin modification delineated by transcriptome independent component analysis in *Arabidopsis thaliana*. *Plant J.* 69, 154–167. doi: 10.1111/j.1365-313X.2011.04779.x

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Received: 28 March 2013; accepted: 06 June 2013; published online: 09 July 2013.

Citation: Wang Y, Chantreau M, Sibout R and Hawkins S (2013) Plant cell wall lignification and monolignol metabolism. *Front. Plant Sci.* 4:220. doi: 10.3389/fpls.2013.00220

This article was submitted to *Frontiers in Plant Biotechnology*, a specialty of *Frontiers in Plant Science*.

Copyright © 2013 Wang, Chantreau, Sibout and Hawkins. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits use, distribution and reproduction in other forums, provided the original authors and source are credited and subject to any copyright notices concerning any third-party graphics etc.