Download Deciphering the Enigma of Lignification: Precursor Transport

Molecular Plant
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Volume 5
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Number 2
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Pages 304–317
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March 2012
REVIEW ARTICLE
Deciphering the Enigma of Lignification:
Precursor Transport, Oxidation, and the
Topochemistry of Lignin Assembly
Chang-Jun Liu1
ABSTRACT Plant lignification is a tightly regulated complex cellular process that occurs via three sequential steps: the
synthesis of monolignols within the cytosol; the transport of monomeric precursors across plasma membrane; and the
oxidative polymerization of monolignols to form lignin macromolecules within the cell wall. Although we have a reasonable understanding of monolignol biosynthesis, many aspects of lignin assembly remain elusive. These include the precursors’ transport and oxidation, and the initiation of lignin polymerization. This review describes our current knowledge
of the molecular mechanisms underlying monolignol transport and oxidation, discusses the intriguing yet leastunderstood aspects of lignin assembly, and highlights the technologies potentially aiding in clarifying the enigma of plant
lignification.
Key words:
lignification; monolignol transport; ABC transporter; laccase; peroxidase.
INTRODUCTION
Lignin is complex phenypropanoid polymer derived primarily
from three cinnamyl alcohols: p-coumaryl, coniferyl, and
sinapyl alcohols (termed monolignols). It is a crucial structural
component preserving the integrity of plant cell wall, imparting stiffness and strength of vascular plants, enabling the
transport of water and solutes through the tracheary elements
in the vasculature system, and affording physical barriers
against invasions of phytopathogens, and other environmental stresses (Boerjan et al., 2003; Ralph et al., 2004a). However,
its presence contributes to the recalcitrance of cell wall to degradation, and thus is detrimental to using cellulosic fibers in
cattle feedstock, for pulping and paper making, and for producing liquid biofuels (Chen and Dixon, 2007; Li et al., 2008;
Weng et al., 2008).
Plant lignification is a cellular process generating lignin
polymer in the cell wall. In general, it occurs in three stages:
the biosynthesis of monolignols in the cytosol; the transport
of these monolignols to the cell wall; and their subsequent oxidative dehydrogenation and polymerization to form heterogeneous macromolecules. Over several decades, extensive
studies have centered on monolignol biosynthesis, particularly
on elucidating its pathways and the molecular regulation of
the biosynthetic processes. The progresses have been intensively reviewed correspondingly (Anterola and Lewis, 2002;
Boerjan et al., 2003; Ralph et al., 2004a; Ralph, 2007; Li
et al., 2008; Vanholme et al., 2008; Zhong and Ye, 2009; Umezawa, 2010; Vanholme et al., 2010; Zhao and Dixon, 2011).
Compared to the knowledge of monolignol biosynthesis,
our understanding of lignin assembly, namely the transport
of lignin precursors, their deposition, and subsequent activation and polymerization, is fragmentary. In this review, we outline the different speculations on the sequestration, transport,
and subsequent spatial deposition of the monolignols, and describe recent progresses in molecular genetics and biochemical
studies aimed at gaining an understanding of the underlying
molecular mechanisms for translocating monolignols across
cell membranes, advances in identifying the oxidative dehydrogenation enzymes, and progresses and issues in detailing
the topochemistry of lignification. Meanwhile, we highlight
the technological developments that might be applied productively to explore lignin assembly.
1
To whom correspondence should be addressed. E-mail [email protected],
tel. 631-344-2966, fax 631-344-3407
ª The Author 2012. Published by the Molecular Plant Shanghai Editorial
Office in association with Oxford University Press on behalf of CSPB and
IPPE, SIBS, CAS.
doi: 10.1093/mp/ssr121, Advance Access publication 3 February 2012
Received 29 September 2011; accepted 13 December 2011
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Biology Department, Brookhaven National Laboratory, Upton, NY 11973, USA
Liu
MONOLIGNOL BIOSYNTHESIS AND THE
POTENTIAL PHYSICAL ORGANIZATION
OF THE ENZYMES
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are membrane-anchored cytochrome P450 proteins that predictably associate with the outer surface of the endoplasmic reticulum (ER) by virtue of their N-terminal membrane anchor (Li
et al., 2008). Nevertheless, many other enzymes, such as phenylalanine ammonia lyase (PAL), 4-hydroxycinnamoyl CoA ligase
(4CL), caffeoyl CoA O-methyltransferase (CCoAOMT), hydroxycinnamoyl CoA reductase (CCR), caffeic acid/5-hydroxyferulic
acid 3/5-O-methyltransferase (COMT), and (hydroxy)cinnamyl
alcohol dehydrogenase (CAD) found in diverse species operationally are soluble proteins and likely to be located within
the cytosol (Takabe et al., 1985; Nakashima et al., 1997; Chen
et al., 2000). Apparently, the different compartmental propensity of the monolignol biosynthetic enzymes, together with the
exclusive plastidic localization of the shikimate pathway that
leads to the synthesis of aromatic amino acids for phenylpropanoids (Herrmann and Weaver, 1999; Rippert et al., 2009), implies
either the occurrence of multiple subcellular sequestration of
metabolic intermediates or the ideal physical organization of
Figure 1. The Scheme of the Simplified Shikimate–Phenylpropanoid–Lignin Biosynthetic Pathway, Illustrating Different Compartmentalization of the Biosynthesis.
CM, chorismate mutase; TYRA, arogenage dehydrogenase; ADT, arogenate dehydratase; PAT, prephenate aminotransferase; PAL, phenylalanine ammonia lyase; TAL, tyrosine ammonia lyase; C4H, cinnamic acid 4-hydroxylase; 4CL, 4-hydroxycinnamoyl CoA ligase; HCT, hydroxycinnamoyl; CoA, shikimate/quinate hydroxycinnamoyltransferase; C3#H, p-coumaroylshikimate 3#-hydroxylase; CCoAOMT, caffeoyl CoA
O-methyltransferase; CCR, cinnamoyl CoA reductase; Cald5H/F5H, coniferaldehyde/ferulate 5-hydroxylase; COMT, caffeic acid/5-hydroxyferulic
acid O-methyltransferase; CAD, (hydroxy)cinnamyl alcohol dehydrogenase; POX, peroxidase; LAC, laccase.
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Originating from the shikimate pathway, wherein plant produces aromatic amino acids, the biosynthesis of monolignols
starts from the deamination of phenylalanine by the entry
point enzyme phenylalanine ammonia lyase (PAL), and then
undergoes subsequent aromatic-ring modifications via hydroxylation and methylation, and the transformation of the
carboxylic moiety of the propane tail through esterification
and reduction. This process yields primarily three hydroxycinnamoyl alcohols, p-coumaryl, coniferyl, and sinapyl alcohols
(i.e. monolignols) (Umezawa, 2010; Vanholme et al., 2010).
More than 10 enzymes sequentially catalyze this synthetic
pathway (Figure 1). Three of them, namely cinnamic acid
4-hydroxylase (C4H), p-coumaroylshikimate 3#-hydroxylase
(C3#H), and coniferaldehyde/ferulic acid 5-hydroxylase (F5H),
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physically organized and/or compartmentalized and how such
macromolecular organizations affect lignin deposition are intriguing aspects for future explorations.
TRANSPORT OF MONOLIGNOLS ACROSS
CELL MEMBRANE
After monolignols are synthesized in the cytosol, they must be
moved into the cell wall, where they are oxidized, and integrated into the matrix of the secondary cell wall. The exact
mechanism whereby this occurs remains unclear; at least three
models or pathways are envisaged, as illustrated in Figure 2: (1)
exocytosis via ER-Golgi-derived vesicles; (2) passive diffusion
via hydrophobic reactions through the plasma membrane;
and (3) active transport via plasma membrane-located transporters or the other facilitators. We have discussed these potential mechanisms in detail in a recent review (Liu et al., 2011).
To keep the integrity of this paper, we reiterate some related
contents here but focus more on the up-to-date progresses on
this subject.
Monolignol Glucosylation and Deglucosylation
Monolignols in cells occur both as free aglycones (non-sugar
compounds) and as water-soluble 4-O-b-D-glucosides. The latter,
namely p-coumaryl alcohol glucoside, coniferin, and syringin,
are primarily found in gymnosperms (Terazawa and Miyake,
1984; Whetten and Sederoff, 1995). A set of angiosperm species
also can generate those glucoconjugates in particular tissues; for
instance, coniferin and syringin accumulate in Arabidopsis roots
and their levels can be raised by light treatment (Hemm et al.,
2004). In conifers, the build-up of coniferin correlates spatially
and temporally with the beginning of secondary growth
(Savidge, 1988, 1989). Presumably, these glucoconjugates are
stored within the vacuoles of cambial cells (Leinhos and Savidge,
1993; Dharmawardhana et al., 1995). The possible presence of
the glucosides of monolignols (or of cinnamaldehydes) in
vacuoles of differentiating xylem cells led to the suggestion that
those glucoconjugates may serve the storage and transport
functions of monolignols at least in gymnosperms and some evolutionarily less-advanced angiosperms (Steeves et al., 2001; Tsuji
and Fukushima, 2004). Presumably, those glucosides are sequestrated first from the cytosol to vacuoles and then exported to the
cell wall (Dharmawardhana et al., 1995, 1999; Samuels et al.,
2002; Escamilla-Treviño et al., 2006).
Monolignol glucosylation is catalyzed by coniferyl/sinapyl alcohol glucosyltransferase (Steeves et al., 2001). The activity of
this enzyme was detected in many gymnosperms and angiosperms; the woody angiosperms exhibited higher enzyme activities than did the herbaceous species (Ibrahim, 1977). On the
other hand, monolignol deglucosylation, catalyzed by specific
b-glucosidase, is thought to occur in the cell wall at the
site of lignin polymerization. The apoplastic location of
b-glucosidase was demonstrated by immunohistochemical
studies (Marcinowski et al., 1979; Burmeister and Hosel,
1981). The enzymes were identified in Norway spruce (Picea
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the pathways to eliminate the phytotoxicity of aromatic intermediates and to enhance the transformation efficiency of the
intermediate compounds. Such physical organization of metabolic pathways is variously termed ‘metabolic compartment’,
‘metabolic channeling’, or ‘metabolon’. The metabolic channels
may contain arrays of consecutive, physically associated enzymes
assembled on membranes or other physical structures, or interacted directly with each other. This association can channel biosynthetic intermediate from one enzyme to the next without
releasing them into the total cellular metabolic pool (Srere,
1987; Hrazdina and Jensen, 1992; Burbulis and Winkel-Shirley,
1999; Winkel-Shirley, 1999). Earlier immunolocalization studies
on monolignol enzymes in the lignifying mesophyll cells of Zinnia elegans and the differentiating xylem cells of Eucalyptus and
Populus revealed that the isoforms of PAL, CAD, OMT, and 4CL
could associate with ER-Golgi-derived vesicles, and then dispersed into the cytosol (Takabe et al., 2001; Takeuchi et al.,
2001). In contrast, the peroxidase involved in the dehydrogenative polymerization of monolignols was synthesized in the
rough ER and then transported to the lignifying cell wall (Takabe
et al., 1985; Nakashima et al., 1997; Takabe et al., 2001; Takeuchi
et al., 2001). Metabolic labeling experiments also revealed
that a PAL isoform, PAL1, in tobacco was in close physical
association with the membrane-bound P450 enzyme C4H on
the microsomes for the synthesis of the active intermediate,
p-coumaric acid (Czichi and Kindl, 1977; Hrazdina and Jensen,
1992; Deshpande et al., 1993; Rasmussen and Dixon, 1999; Dixon,
2000). Membrane fractionation and protein gel-blot analyses
corroborated this observation. Furthermore, the transient coexpression of the PAL1 or PAL2 fused with green fluorescence
protein in the cells expressing C4H demonstrated that two
PAL isoforms exhibited different affinities in associating with
the ER microsome and C4H likely organized the enzyme complex
containing PAL1 and/or PAL2 (Achnine et al., 2004). These data
suggest that monolignol biosynthetic enzymes are partially
organized within the lignifying cells. Consequently, Dixon
et al. (2001) speculated that there was a complicated metabolic
channel for producing syringyl lignin, wherein three cytochrome
P450 enzymes, C4H, C3#H, and F5H, anchor other operationally
soluble enzymes, including COMT and perhaps specific forms of
PAL, 4CL, and/or CCR, to the surface of ER. However, experimentally substantializing this appealing hypothesis proved challenging and technically demanding. Recently, many biochemical,
biophysical, molecular, and cellular approaches were developed
and applied for dissecting protein–protein interactions or protein complex organization (Lalonde et al., 2008; Miernyk and
Thelen, 2008). Among them, several may offer the possibility
to precisely evaluate the potential physical/spatial organization
of lignin biosynthesis at cellular and subcellular levels. Particularly promising are the split ubiquitin membrane yeast
two-hybrid screening (Obrdlik et al., 2004), surface resonance
spectroscopy, fluorescence-based technology, such as bimolecular fluorescence complementation (BiFC) and the fluorescence
energy resonance transfer (FRET), and three-dimensional
super-solution microscopy. How lignin biosynthetic enzymes
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Symplastic transport of monolignols may export them to the cell wall through active transport or by passive diffusion. Alternatively, they
may be sequestrated and stored as glucoconjugates into the vacuoles in gymnosperms, before their subsequent transport to the cell wall
and hydrolysis to free monolignols for polymerization. The deposited monolignols in the cell wall diffuse to initiation sites where the
polymerization process begins. The polymerization to form different bond-linkages of lignin is known to be a random chemical process.
However, the nature of initiating sites and the way in which the amount and type of lignin formation is controlled across the cell wall are
poorly understood.
abies) (Marcinowski and Grisebach, 1978), pinus species
(Leinhos et al., 1994; Dharmawardhana et al., 1995), and
A. thaliana (Escamilla-Treviño et al., 2006). The cDNAs encoding coniferin b-glucosidase were isolated from both lodgepole pine and Arabidopsis; their expression was proved
highly specific at the site of lignification (Dharmawardhana
et al., 1999; Escamilla-Treviño et al., 2006). Based on the discovery of UDP-glucose:coniferyl alcohol glucosyltransferase/
coniferin b-glucosidase, a monolignol glucosyltransferase and
b-glucosidase-mediated transport pathway was proposed,
where the two enzymes coordinately regulate the storage
and mobilization of monolignols for lignin biosynthesis
(Dharmawardhana et al., 1995, 1999; Samuels et al., 2002;
Escamilla-Treviño et al., 2006). However, recent experiments
failed to support this speculation. A small cluster of glucosyltransferase genes were functionally characterized from
Arabidopsis. The encoded enzymes exhibited ability to glucoconjugate monolignols in vitro (Lim et al., 2001). However, disrupting those genes’ expression did not perturb lignin
deposition (Vanholme et al., 2008), although a corresponding
reduction or accumulation of soluble monolignol glucosides
in the roots or leaves of transgenic plants was recorded (Lanot
et al., 2006). Moreover, feeding [3H]-Phe to the dissected xylem
of lodgepole pine (Pinus contorta) revealed that the radiolabels
were incorporated directly into the monolignols and the lignins
accumulated in the cell wall. The substantial amount of the
radiolabels was associated neither with monolignol glucoside
(coniferin) nor with the interior of the central vacuole, where,
expectedly, coniferin is stored (Kaneda et al., 2008). These data
argue against the proposed roles of monolignol glucosylation–
deglucosylation in exporting lignin precursors across plasma
membrane, and imply that monolignol aglycone is the chemical
form used for such transport.
Mechanisms for Transporting Monolignols across Cell
Membranes
Exocytosis via Vesicles Derived from Endoplasmic
Reticulum-Golgi Bodies
The non-cellulosic polysaccharides of plant cell wall, such as pectin and hemicellulose, are known to be synthesized within the
Golgi bodies and exported to the cell wall through an exocytosis
mechanism (Lerouxel et al., 2006; Sandhu et al., 2009). Early autoradiographic and immunochemical studies implied a similar
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Figure 2. The Scheme for Monolignol Deposition and the Subsequent Initiation of Lignin Polymerization within the Cell Wall.
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Passive Diffusion
Genetic engineering of lignin biosynthesis and chemical analysis of the resulting transgenic cell walls reveal that many nontraditional phenolics can be incorporated into the lignin
polymer. In transgenic plants and/or mutants deficient
in monolignol biosynthesis, some metabolite intermediates
or precursors, such as 5-hydroxyconiferyl alcohol (Van
Doorsselaere et al., 1995; Ralph et al., 2001), hydroxycinnamaldehydes (Baucher et al., 1996), and hydroxycinnamic acids
(Leple et al., 2007; Ralph et al., 2008), were integrated into lignin. In addition, in monocot and some dicot species, monolignols are often further acylated with p-hydroxybenzoate,
p-coumarate (Lu et al., 2004), and acetate (Lu and Ralph,
2002). These derivatives occur in the cell wall lignin (del Rio
et al., 2007, 2008). Such accommodation of alternative monomers in lignification implies a potential non-selective passive
diffusion of lignin monomers through plasma membrane
(Vanholme et al., 2008, 2010). This possibility was further evidenced from in vitro partitioning experiments using immobilized liposomes or lipid-bilayer discs as the model membranes.
Incubating monolignol-like compounds with artificial model
membranes caused them to freely partition into the membrane phase due to hydrophobic–hydrophilic interactions
(Boija et al., 2007, 2008). However, so far, we lack definitive
biochemical evidence for such transport in plant tissues.
Transporter-Mediated Active Sequestration and Export of
Monolignols
Global transcriptomic, proteomic studies, or functional genomics
in gymnosperms and angiosperms have frequently uncovered
highly expressed membrane transporters in lignifying or lignified tissues. For example, several ESTs encoding for ATP-binding
cassette (ABC) transporters were disclosed from gene expression
profiling during wood transformation (Allona et al., 1998;
Hertzberg et al., 2001; Kirst et al., 2003; Egertsdotter et al.,
2004). Transcriptional profiling based on the first-generation
loblolly pine microarray revealed the abundance of distinct
ABC transporter-like genes in the mature and compression
woods, wherein different types of lignins accumulated (Whetten
et al., 2001; Egertsdotter et al., 2004). In global transcript profiling of A. thaliana stems, seven ABC transporter genes exhibited
similar expression patterns with the functionally known monolignol biosynthetic genes (Ehlting et al., 2005). ABC transporter
genes were also recognized in the Eucalyptus xylem-subtractive
library (Paux et al., 2004) and in the cultures of lignifying Zinnia
tracheary elements (Pesquet et al., 2005). Also, the cell wall
macro-array analysis of maize (Zea mays) brown mid-rib (bm)
mutants indicated that the lower levels of guaiacyl lignin in
bm2 mutants might reflect the decreased transcription of one
ABC transporter gene (Guillaumie et al., 2007). Furthermore,
in a recent proteomics study on plasma membranes from poplar
leaf, xylem, and cambium/phloem, a set of ABC transporters
showed a rather ‘specific’ tissue distribution; some members
of the ABC transporter subfamily B and G were identified only
in the cambium/phloem (Nilsson et al., 2010). These data imply
that ABC transporters might play a significant role in cell wall
lignification, presumably in transporting monolignols to the wall
(Samuels et al., 2002; Douglas and Ehlting, 2005).
ABC transporters constitute a large, diverse family of proteins
that translocate a broad range of substances across cell membranes (Higgins, 1992; Sánchez-Fernández et al., 2001; Verrier
et al., 2008). The genomes of Arabidopsis and rice (Oryza sativa)
each contain more than 120 putative ABC transporters (SánchezFernández et al., 2001; Jasinski et al., 2003). Many of these proteins exhibited diverse transport activities to a variety of small
molecule metabolites, such as glutathione conjugates (Lu
et al., 1998; Tommasini et al., 1998), auxins (Geisler et al.,
2005; Strader and Bartel, 2009; Ruzicka et al., 2010), abscisic acid
(Kang et al., 2010; Kuromori et al., 2010), inorganic ions (Suh
et al., 2007), malate (Lee et al., 2008), wax and cutin precursors
(Pighin et al., 2004; McFarlane et al., 2010), defensive secondary
metabolites such as antifungal terpenoids (Jasinski et al.,
2001), tropane alkaloids (hyoscyamine and scopolamine)
(Goossens et al., 2003), benzylisoquinoline alkaloid (berberine)
(Shitan et al., 2003; Shitan and Yazaki, 2007), flavone glucuronides (Klein et al., 2000; Frangne et al., 2002), isoflavone genistein (Sugiyama et al., 2007), and many xenobiotics (Baerson
et al., 2005).
To ascertain whether the transport of monolignol is an active process that involves ABC transporters, we recently
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mechanism for transporting lignin precursors. A vesicle trafficking between cytosol and plasmalemma in the differentiating
tracheids of wheat xylem was observed (Pickett-Heaps, 1968).
Feeding the developing xylem with isotopically labeled Phe,
Tyr, and cinnamic acid resulted in their incorporation into the
rough ER, the Golgi apparatus, and some vesicles that were associated with plasma membrane, or aggregated near the bands
of cell wall microtubules (Pickett-Heaps, 1968; Fujita and
Harada, 1979; Takabe et al., 1985). These findings suggested
that the ER-Golgi apparatus are likely to be involved in synthesizing and transporting monolignols to the cell wall. However,
since the applied radiotracers are assimilated efficiently into lignin and into proteins, it might complicate the interpretation of
these autoradiographic data. Indeed, in a recent experiment,
Kaneda et al. (2008) fed the [3H]-Phe to the cambium/developing xylem tissues of lodgepole pine that were isolated by cryofixation and freezing substitution before sectioning (those
treatments substantially minimize the damage to the cells
and thus prevent pseudo-autographic imaging). By selectively
inhibiting phenylpropanoid or protein biosynthesis, they discovered that the radioactivity presented in the ER-Golgi was
primarily derived from proteins rather than phenylpropanoids/monolignols. Further, they observed that the abundant
Golgi and Golgi-vesicle clusters in the developing xylem cells
did not load with phenylpropanoids (Kaneda et al., 2008). Their
study suggests that ER-Golgi vesicle-mediated exocytosis is
unlikely to play a significant role in transporting monolignols.
Liu
Issues in Identifying Particular Transporters Responsible
for Depositing Lignin
To identify the specific ABC transporters functioning in lignification, Kaneda et al. (2011) analyzed a subset of Arabidopsis
ABC transporters whose expressions have a good correlation
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with phenylpropanoid–lignin biosynthesis and are higher in
the vascular or interfascicular tissues of the primary stem
(Ehlting et al., 2005). However, mutant lines singly deficient
in those ABC transporter genes did not attenuate xylem lignification. Instead, two mutant lines respectively disrupting
ABCB14 and ABCB15 impaired auxin polar transport in the
stem, even though the former was recently characterized as
a malate importer in leaf guard cells (Lee et al., 2008). These
findings highlight the difficulty in characterizing the specific
monolignol transporters. Since lignification is a very complicated cellular process, orchestrating a large set of gene expression during cell differentiation and secondary cell wall
thickening, the ABC transporters found in vascular tissues
and correlated with their lignification may be involved in distinct biological processes. Furthermore, lignin deposition and
distribution are known to be tissue/cell-type specific, which
may require a set of distinct membrane transporters. Many
ABC transporter genes have overlapping tissue expression pattern and therefore possible functional redundancy. Analyzing
plants with a single gene deletion might fail to distinguish its
consequence on lignification, particularly when lignin content
is measured in large-sized, pooled cell wall samples. In addition, ABC transporters typically exhibit low substrate specificity (Verrier et al., 2008; Ruzicka et al., 2010); some members
may transport a set of unrelated molecules and fulfill disparate
functions, as implied by the researches on ABCB14 (Lee et al.,
2008; Kaneda et al., 2011). Besides creating multiple genedeficient lines for the related ABC transporters, we must
precisely measure the deficit of lignification in a tissue or
cell-type-specific manner. Laser capture microdissection of vascular bundles and interfascicular fibers, followed by microthioacidolysis or pyrolytic GC/MS analysis (Patten et al.,
2010) may offer a practical, higher-resolution strategy for
identifying the particular set of ABC transporters or other
membrane transporters involved in transporting monolignols
and depositing lignin.
ENZYMES RESPONSIBLE FOR OXIDATIVE
DEHYDROGENATION
After their deposition in the cell wall, monolignols undergo
dehydrogenation, producing resonance-stabilized radicals.
Then, the resulting phenoxy radicals couple each other or to
the growing lignin polymer, if the polymer was also oxidized,
to form different types of sub-linkages. Monolignol oxidation/
dehydrogenation is proposed to be catalyzed by peroxidases,
laccases, or other phenol oxidases. Those enzymes are widely
found in lignin-forming tissues and can oxidize a variety of
phenolics, including monolignols in vitro (Freudenberg,
1959; Ralph et al., 2004a; Davin et al., 2008).
Peroxidase uses hydrogen peroxide as its electron receptor
to oxidize a variety of phenolics (Harkin and Obst, 1973). Different peroxidases have distinct predilection in oxidizing the
conventional monolignols. Most peroxidases readily oxidize
coniferyl alcohol, and the resulting radicals can transfer to,
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conducted the in vitro uptake assay to monolignols and/or
their glucosides by using plasma and vacuolar membrane
vesicles prepared from both Arabidopsis rosette leaves and
poplar roots (Miao and Liu, 2010). Our study reveals several
lines of evidence demonstrating the involvement of ABC-like
transporters in monolignol sequestration and transport: (1)
the transport of monolignols across either plasma or vacuolar
membranes largely depends on the presence of nucleotide
phosphates, particularly ATP; lacking it severely impairs the
transport activity of either type of vesicles, suggesting that
transport is an active process requiring energy; (2) a set of specific ABC transporter inhibitors largely reduce the uptake activity of both plasma and vacuolar membrane vesicles for
lignin precursors in the presence of ATP; (3) the uptake of
monolignols or their glucosides by membrane vesicles displays
typical protein–ligand binding kinetics, indicating that it is
a membrane protein-mediated biochemical process rather
than passive diffusion; and (4) the uptake to lignin precursors
by membrane vesicles shows obvious selectivity. In the presence of ATP, the vacuolar membrane vesicles actively sequester
monolingol glucoconjugates, whereas the preference of the
inverted plasma membrane vesicles is for the monolignol aglycones; this difference suggests that the glucosylation of monolignols is a prerequisite for vacuolar sequestration, while the
aglycones are the form for direct export into cell walls. Also,
the findings implicate the distinct classes of ABC transporters
as being involved in either partitioning the polarized ‘storage
form’ glucosides of monolignols into vacuoles or exporting the
hydrophobic aglycones across plasmalemma. Compared to the
vacuolar membrane vesicles, plasma membrane ones exhibit
a more relaxed substrate specificity, being able to convey
p-hydroxycinnamyl alcohols and other related cinnamaldehydes, thereby suggesting the basis for the observed plasticity
of lignin biosynthesis (Miao and Liu, 2010). Promiscuous
substrate-transport activity and/or the low intrinsic level of
plasmalemma diffusion may engender the deposition of
non-classic lignin precursors in the cell wall. In addition, we
found that the chemical character of the para-hydroxyl of
monolignol substantially influences the nature of transport.
While the 4-O-glucosylation of monolignols likely dominates
the vacuolar sequestration, the etherification of the parahydroxyl with a methyl moiety largely releases the dependence of monolignol transport on the ATP-energized
mechanism. When incubated with 4-O-methyl coniferyl
alcohol in the absence of ATP, the amount of this etherified
monolignol derivative accumulates in the inverted plasmamembrane vesicles is comparable to that of conventional
monolignols accumulated in the presence of ATP (Y. Miao
and C.-J. Liu, unpublished data).
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their expressions closely concur across a wide variety of tissues/
development stages (Turlapati et al., 2011). A few family members display unique tissue/cell-type expression patterns, suggesting their specific roles in either lignification or other
oxidative processes (Pourcel et al., 2005; Turlapati et al.,
2011). Determining the role of laccases in lignification yielded
ambiguous findings until a recent genetic study wherein the
functions of the Arabidopsis laccase genes LAC4 and LAC17
in xylem lignification were demonstrated conclusively (Berthet
et al., 2011). About a 30–40% reduction of lignin content in the
stem of mutant plants simultaneously disrupting LAC4 and
LAC17 was registered. A deficiency of LAC17 primarily affected
the deposition of guaiacyl lignin and resulted in a change of
the S/G ratio in mutant lines, implying the preference of LAC17
for oxidizing coniferyl alcohol. However, definitive specifications of the biochemical properties of those LACs are lacking
presently. It will be interesting to see whether the identified
LACs are able to oxidize sinapyl alcohol and other higher molecule oligomers/polymers.
Interestingly, down-regulation of LAC broadly changed the
phenylpropanoid biosynthetic pathway. Almost the entire
batch of monolignol biosynthetic genes was suppressed coordinately with the disruption of LAC4 and 17 (Berthet et al.,
2011). These data imply the existence of some form of feedback regulatory mechanisms at the plasma membrane/cell wall
interior that prevents the accumulation/transport of monolignols when oxidative capacity is diminished. It is interesting
to decipher whether the accumulated monolignols or their
derivatives serve as signaling molecules, and how the set of
biosynthetic genes in the pathway are organized as a particular
‘regulon’.
SPATIAL DISTRIBUTION OF LIGNIN AND
TOPOCHEMISTRY OF LIGNIFICATION
Lignin is known to be selectively deposited in the walls of different cell types. For example, the vessels of birch wood are
enriched in G units, while the fiber wall incorporates both
S and G units (Fergus and Goring, 1970). This feature is also
demonstrated in Arabidopsis stems, where the lignin of the
vascular bundle rich in vessel cells primarily contains G units,
while the interfascicular fibers are enriched in S units (Chapple
et al., 1994). In a more subtle way, lignin is differentially deposited in discrete regions of the cell walls. In spruce wood,
the lignin of the middle lamella embeds more p-coumaryl
alcohol-derived units than does the secondary wall lignin that
mainly is derived from coniferyl alcohol (Whiting and Goring,
1982). Feeding labeled monolignols into developing xylem
reveals the preferential deposition of radiolabeled p-coumaryl
alcohol in the middle lamella or cell corners, whereas coniferyl
alcohol is located mostly in the secondary wall (Terashima
et al., 1993). Moreover, during the development of secondary
cell wall to form S1-, S2-, and S3- layers, the three kinds of
monolignols exhibit sequential deposition, in the order of
p-coumaryl alcohol (for the H unit), coniferyl alcohol (G unit),
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and thus oxidize, sinapyl alcohol. Sinapyl alcohol itself is a poor
substrate for oxidation, and is catalyzed only by a very specific
peroxidase (Sasaki et al., 2006, 2008). These properties led to
a radical mediation model to explain lignin polymerization, in
which the coniferyl alcohol radicals generated through the interaction of the cell wall-bound peroxidases with monolignols
diffuses to the growing lignin polymer; thereafter, they either
transfer their higher oxidation state to a lignin molecule, if the
lignin polymer is at a base oxidative state, or undergo a coupling reaction to form a covalent bond with the polymer, if the
latter is at a higher oxidation state (Hatfield and Vermerris,
2001). Further, a complementary model was derived from this
radical-transfer hypothesis by suggesting the operation of
a diffusible redox shuttle-peroxidase system in which a membrane-bound peroxidase oxidizes manganese, the oxidized
ions diffuse through small pores in the cell wall to the lignification site, and then they oxidize both the monolignols and
growing lignin polymers. As the redox ions directly oxidize
the monolignols and polymers, the radicals as the primary oxidation products from both constitute a high concentration, thus
facilitating the coupling frequency of the monolignols with
growing polymers (Onnerud et al., 2002). However, the need
for having radical or redox-shuttle mediation was challenged
by the identification of a Populus peroxidase specific for sinapyl
alcohol that can directly and efficiently oxidize both sinapyl alcohol and the higher molecular polymers (Sasaki et al., 2004).
Peroxidases in plant are encoded by a multigene family. In
Arabidopsis, 73 class III peroxidase genes were identified
(Tognolli et al., 2002; Fagerstedt et al., 2010). Despite their
high degree of sequence and function redundancy that complicate the identification of lignin-specific enzymes, the biological functions of class III plant peroxidases in lignification
were demonstrated in a few genetic studies with altered expression of the peroxidase genes. Down-regulation of the
gene coding for a cationic peroxidase NtPrx60 (TP60) in tobacco resulted in up to a 50% reduction in total acetyl bromide
lignin, and the corresponding reduction in both the G and S
units (Blee et al., 2003); down-regulation of an anionic peroxidase PkPrx03 (PrxA3a) in aspen resulted in a decline of up to
20% of total lignin content in transgenic lines (Li et al., 2003).
These data suggest that species-specific oxidative enzymes are
involved in plant lignification; on the other hand, different
types of peroxidases may have overlapping biological functions in lignin polymerization.
Laccases (p-diphenol:dioxygen oxidoreductase, EC 1.10.3.2)
are copper-containing extracellular glycoproteins that require
O2 as secondary substrate to oxidize various phenolic, inorganic, and/or aromatic amine substrates (Reinhammar and
Malmstroem, 1981). Similar to peroxidase, laccases oxidize
monolignols in vitro (Freudenberg, 1959; Sterjiades et al.,
1992; Bao et al., 1993; Takahama, 1995; Richardson et al.,
1997), and laccase-like activities were detected in the lignifying cell walls of differentiating xylem (Driouich et al., 1992;
Bao et al., 1993; Liu et al., 1994; Richardson et al., 2000). Arabidopsis contains 17 putative laccase homologous genes, and
Liu
Enigma of Plant Lignification
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ticularly for ferulate, these ‘wall-bound’ phenolics can dimerize or polymerize with each other, or with other cell wall
phenolics, forming ester-to-ether linkages to cross-link the adjacent polysaccharides, lignins, and/or structural proteins
(Ralph et al., 2004a). These ‘wall-bound’ phenolics purportedly
also act as nucleation sites for lignin polymerization (Ralph
et al., 2004a; Ralph, 2010). Peroxidase activity was noted, catalyzing the formation of ferulate-mediated ester to ether linkages between polysaccharide and lignin composition, or the
cross-links of ferulate residues in polysaccharides (Burr and
Fry, 2009a, 2009b). However, conclusive evidence to establish
the role of ‘wall-bound’ ferulate in initiating lignin polymerization is needed.
Since lignin assembly depends on the quantity and type of
monolignol radicals within the particular region of the cell
wall (Hatfield and Vermerris, 2001; van Parijs et al., 2010),
the potential sub-compartmental localization of the radicalproducing enzymes, namely peroxidase and laccase, within
the cell wall may affect the initiation of lignin polymerization
and control the spatial deposition of a particular type of lignin.
Both enzymes are extracellular proteins, and are known from
immunohistochemical analyses to locate in middle lamellae,
cell corner, and secondary cell wall of the lignifying cells
(Bao et al., 1993; Liu et al., 1994; Richardson et al., 2000; Sasaki
et al., 2006). However, there is little detail about their potential
sub-compartmentalization and spatial movement within the
cell wall during developmental programming and the lignification process. An early study found that the sycamore maple
laccase is far less active on phenolic substrates containing multiple aromatic rings than is peroxidase (Sterjiades et al., 1993).
This leads to the assumption that laccases may polymerize
monolignols into oligo-lignols during the early stages of lignification, whereas the cell wall peroxidases may function when
H2O2 is generated during the later stages of xylem cell development or in response to environmental stresses (Sterjiades
et al., 1993). Recently, intriguing findings from several pieces
of cell wall immunohistochemical studies revealed the existence of diverse microenvironments within cell walls that
are likely to influence the access of proteins or enzymes to specific targets (Lee et al., 2011). Some small-molecular-weight
carbohydrate binding modules or immuno-epitopes can be effectively blocked or restricted to access to their ligands with
the presence of pectic homogalacturonan (Herve et al.,
2010; Marcus et al., 2010). It is possible that the distribution
and localization of oxidative enzymes within the cell wall
are affected with the similar ‘polymer masking’ effects. Along
with characterizing lignin-specific laccases and peroxidases, it
now might be able to evaluate this appealing hypothesis. Detailed biochemical analyses of the identified laccases and peroxidases should clarify whether different types of oxidative
enzymes exhibit substrate specificity, particularly towards oxidizing high-molecule oligomers or polymers. Moreover, advanced microscopic techniques now enable us to dissect the
details of single protein behaviors in a particular cell compartment, as recently demonstrated in a series of researches
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and sinapyl alcohol (S unit) (Terashima et al., 1986a, 1986b,
1993). Disturbing lignin biosynthesis by down-regulating particular monolignol biosynthetic genes, such as CCR, selectively
affects the spatial deposition of lignin in the fiber and xylem
cells, and even within the different sub-layers of secondary cell
wall (Chabannes et al., 2001; Ruel et al., 2009; Thevenin et al.,
2011). These data strongly suggest that lignification is tightly
regulated during developmental programs and that lignin deposition within the wall is a highly organized process. Although the exact molecular mechanism in controlling lignin
deposition remains elusive, the lignin composition of each individual cell appears to be controlled by a complex array of
gene expression in the differentiating cells or in adjacent cells.
This complexity would determine the number and type of lignin precursors deposited at lignification sites, which were predicted as the key factors controlling lignin structure (van Parijs
et al., 2010). Accordingly, the spatial and temporal expression
of the monolignol transporters and their substrate specificity
may greatly contribute to the tissue and cell-type-specific lignification.
Once monolignols are laid down in the cell wall, they are
proposed to diffuse to the initiation site for polymerization.
Lignification, in general, begins at the cell corner of the middle
lamella and the S1 region of the secondary cell wall before
spreading across the secondary wall towards the lumen
(Donaldson, 2001; Moller et al., 2006). Lignification of primary
cell wall typically begins after the start of the formation of the
secondary cell wall, while lignification of secondary cell wall
usually starts when secondary cell wall formation is completed.
Currently, it is unclear yet how the precise topochemistry of
lignification occurs and is controlled. A type of non-enzymatic
dirigent protein was characterized functioning as a template
guiding the formation of the bond linkage of two monolignol
molecules, and defining the stereochemistry of the resulted
optically active compound lignan (Davin et al., 1997). This proteinaceous control mechanism was envisaged for lignin polymerization, where proteins bearing dirigent sites guide the
phenoxy radical coupling within the cell wall (Davin and Lewis,
2000, 2005; Davin et al., 2008). The dirigent sites were then predicted to be the lignin initiation sites (Donaldson, 2001). However, lignin is structurally known as an optical inactive, racemic
polymer containing many different types of bonds connecting
monolignol residues in a somewhat random pattern
(Vanholme et al., 2010). These facts apparently challenge
the involvement of dirigent proteins in polymerization. In
addition, although a set of dirigent homolog genes were
found in plants, so far, none was definitively defined in lignin
polymerization, either biochemically or genetically.
Because lignification starts at the region furthest from the
protoplast, it was suggested that the initiation sites may have
been bound to specific regions of the existing cell wall
(Donaldson, 1994). The cell walls of monocot grasses and some
dicot species, including Arabidopsis thaliana, contain significant quantities of ester-bound hydroxycinnamates, primarily
p-coumarate and ferulate (Ishii, 1997; Ralph et al., 2004b). Par-
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CONCLUDING REMARKS
Lignification is an extremely complicated but tightly controlled biological process, even though the bond formation
in lignin polymerization is a random chemical process. Lignin
deposition and assembly are spatially and temporally programmed in differentiating cell walls. Two important events
therein are the transport of precursors across the protoplast
membrane and the subsequent oxidative dehydrogenation
in the cell walls. In concert with a complex array of transcriptional regulation (reviewed by Zhong and Ye, 2009; Umezawa,
2010; Zhao and Dixon, 2011) and potential metabolic organization of monolignol synthesis, spatially and temporally controlled monolignol transport and oxidation in distinct cell
types and/or the particular sub-compartments of the cell wall
are envisioned, which may play a critical role in controlling the
elaborate processes of lignin deposition. Moving the monolignols across cell membranes takes an energy-requiring, selec-
tive transport process that involves ABC transporters. Despite
the complexity of this process, research is underway towards
clarifying and identifying particular monolignol transporters.
Aided by modern microscopic technologies, together with the
identification of the monolignol transporters and the specific
oxidative enzymes for generating phenox radicals, the enigma
of lignification in plants will be better elucidated.
FUNDING
This work was supported by the Division of Chemical Sciences, Geosciences, and Biosciences, Office of Basic Energy Sciences of the US
Department of Energy through Grant DEAC0298CH10886; the National Science Foundation through grant MCB-1051675; the Laboratory Directed Research and Development Program of Brookhaven
National Laboratory (No 11–007); the CAS/SAFEA International Partnership Program for Creative Research Teams in Plant Metabolisms,
and the National Science Foundation of China for Oversea Distinguished Young Scholars (31028003). No conflict of interest declared.
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