Download The cell biology of lignification in higher plants

Annals of Botany 115: 1053–1074, 2015
doi:10.1093/aob/mcv046, available online at www.aob.oxfordjournals.org
REVIEW
The cell biology of lignification in higher plants
Jaime Barrosy, Henrik Serk1, Irene Granlundz and Edouard Pesquet*
1
Umeå Plant Science Centre (UPSC), Department of Plant Physiology, Umeå University, 901 87 Umeå, Sweden
* For correspondence. E-mail [email protected]
†
Present address: Department of Biological Sciences, University of North Texas, 1155 Union Circle #305220, Denton, TX
76203, USA.
‡
Present address: ProTest Diagnostics, c/o Umeå Biotech Incubator AB, Tvistevägen 48c, 907 19 Umeå, Sweden.
Received: 5 January 2015 Returned for revision: 23 February 2015 Accepted: 10 March 2015 Published electronically: 15 April 2015
Background Lignin is a polyphenolic polymer that strengthens and waterproofs the cell wall of specialized plant
cell types. Lignification is part of the normal differentiation programme and functioning of specific cell types, but
can also be triggered as a response to various biotic and abiotic stresses in cells that would not otherwise be
lignifying.
Scope Cell wall lignification exhibits specific characteristics depending on the cell type being considered. These
characteristics include the timing of lignification during cell differentiation, the palette of associated enzymes and
substrates, the sub-cellular deposition sites, the monomeric composition and the cellular autonomy for lignin monomer production. This review provides an overview of the current understanding of lignin biosynthesis and polymerization at the cell biology level.
Conclusions The lignification process ranges from full autonomy to complete co-operation depending on the cell
type. The different roles of lignin for the function of each specific plant cell type are clearly illustrated by the multiple phenotypic defects exhibited by knock-out mutants in lignin synthesis, which may explain why no general
mechanism for lignification has yet been defined. The range of phenotypic effects observed include altered xylem
sap transport, loss of mechanical support, reduced seed protection and dispersion, and/or increased pest and disease
susceptibility.
Key words: Lignin, lignification, non-cell autonomous processes, plant cell wall, laccases, peroxidases, monolignols, Arabidopsis thaliana.
INTRODUCTION
Lignin (Latin lignum ‘wood’) is a polyphenolic polymer deposited directly in the cell wall of specialized cells. It is not only
restricted to plant woody tissues but represents an integral feature ensuring the proper cellular function of many other cell
types in different tissues/organs of the plant. The appearance of
lignin during plant evolution coincided with the emergence of
the vascular land plants, or tracheophytes, in the Devonian
(Weng and Chapple, 2010). Mechanically weaker than cellulose, lignin nevertheless adds a significant reinforcement to any
cell wall, providing an additional tensile strength of 25–75 MPa
and a Young’s modulus of 25–37 GPa (Gibson, 2012).
Despite the fact that lignin is the second most abundant terrestrial biopolymer after cellulose (Boerjan et al., 2003), our understanding of lignin formation remains fragmentary. In
contrast to cellulose, which presents a defined biochemical
structure independently of the type of plant cell, lignin formation is cell specific and exhibits both distinct sub-cellular localization and monomeric composition: a general lignification
mechanism cannot thus be drawn for all lignified cell types and
may explain why lignification is still only partly understood.
Our current biochemical understanding is that lignin forms in
the spaces between the cellulose microfibrils by the oxidative
coupling of free lignin monomers secreted directly into the
plant cell wall (Boerjan et al., 2003).
The canonical lignin monomers, called monolignols, are the
non-methoxylated p-coumaryl alcohol, the monomethoxylated
coniferyl alcohol and the dimethoxylated sinapyl alcohol which
respectively form H- (hydroxyphenyl), G- (guaicyl) and
S- (syringyl) units in the lignin polymer. Once these monomers
are activated in the cell wall by phenoloxidases, they can displace the radical charge through their conjugated unsaturation,
leading to various mesomeric resonance forms. The lignin polymer then forms by the end-wise addition of new activated monomers to its growing ends and branches (Davin and Lewis, 1992;
Boerjan et al., 2003), and the different linkages between subunits (ether and carbon–carbon bonds between the aliphatic propene and/or the aromatic moieties) depend on the mesomeric
form coupled. Our current lack of understanding of the biological processes behind lignin synthesis is due to the unknown
mechanisms enabling the formation of distinct lignin polymers
in specific cells – such as between wood fibres and vessels
which are neighbours but show specific lignin accumulation and
composition. Moreover, the fact that lignin cannot be removed
once deposited suggests that plants require specific regulatory
mechanisms to control lignin polymer biosynthesis and its subcellular localization at specific stages during the differentiation
of plant cells. This review focuses on the cell biology of lignin,
presenting the current knowledge of the different mechanisms
controlling cell wall lignification in specific cell types.
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Barros et al. — The cell biology of lignification in higher plants
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LIGNIFIED CELL TYPES IN PLANTS
Cell wall lignification occurs during the differentiation of distinct cell types, but also in response to specific environmental
changes. The appropriate timing and localization of lignin deposition in each distinct cell type is essential for the proper
function and adaptation of plants to their environment. Figure 1
represents a schematic view of the developmental and stressinduced lignin formation in the different cell types in intact
Arabidopsis thaliana plants. The cell types accumulating lignin
during their differentiation include the following.
sap-conducting cylinders (Fig. 1) and they are formed by undergoing cell suicide to remove their cell content (Fukuda, 1997)
and reinforcing their side walls with lateral lignified secondary
cell walls mainly composed of G-units (Terashima and
Fukushima, 1989; Higuchi, 1990). Genetic or pharmacological
reduction of TE lignification in whole plants results in collapsed TEs due to the inability of the cell to withstand the negative pressure associated with the rising of the sap (Smart and
Amrhein, 1985; Turner and Somerville, 1997; Jones et al.,
2001; Thévenin et al., 2011).
Tracheary elements (TEs)
Sclerenchyma cells
These specialized cells are an important component of the
xylem: the vascular tissue responsible for the hydro-mineral sap
distribution and the mechanical resistance of plants to gravity
(Tyree and Zimmermann, 2002). TEs act as the plant
These secondary cell wall-forming cells include fibres and
sclereids, found in many different plant tissues such as xylem,
phloem, epidermis and cortex in grasses and cereals, and in fruit
fleshy tissues (i.e. stone cells in pear fruit; Tao et al., 2009).
Development
Stress
Replum
Wounding
Pathogen attack
Drought
Temperature
Valves
Seed dispersion
Nutrient availability
CO2
Ozone
UV radiation
SILIQUE
Seed coat
Seed protection
Replum, valves and seed coat
LEAF
Sclerenchyma: mechanical support
Mesophyll
STEM
TEs: mechanical support
and nutrient transport
Xylem fibres and TEs
Parenchyma
Casparian
strip
ROOT
Unlignified cell wall
Lignified primary cell wall
Lignified secondary cell wall
Endodermis
Endodermis: transport barrier
FIG. 1. Lignified cell types in higher plants. The role of lignin: as a transport barrier; in water and nutrient transport; for mechanical support; for seed protection and
dispersion; and as a response to biotic and abiotic factors.
Barros et al. — The cell biology of lignification in higher plants
The function of these cells is to strengthen the central axis of
plant organs mechanically against gravity, mechanical disturbances and physical damage (Fig. 1). The sclerenchyma fibres
and sclereids have lignified secondary cell walls mainly composed of S-units (Higuchi, 1990). Genetic modification of
xylem fibre lignification by loss-of-function mutation or gene
silencing results in plants unable to withstand gravity (Jones
et al., 2001; Smith et al., 2013).
Endodermal cells
This cell type constitutes the root endodermis which delimits
the root cortex from its vascular system. Endodermal cells selectively allow passage of water and solutes to the root vascular
system by forming an apoplastic barrier against the extracellular diffusion of substances (Steudle, 2000; White and Broadley,
2001; Geldner, 2013): the Casparian strip. This longitudinal orientated strip, composed of lignin-like polymer combined with
suberin, tightly links the plasma membranes and the apoplastic
space between adjacent endodermal cells (Fig. 1). The
Casparian strip lignin-like polymer is composed of a mixture of
G- and S-units in monocolyledonous species, whereas dicotyledon plants exhibit more H- and G-units than S-units (Zeier and
Schreiber, 1997; Zeier et al., 1999; Naseer et al., 2012).
Genetic or pharmacological modification of Casparian strip
lignification in whole plants leads to a loss of the apoplastic
barrier function of the endodermis (Geldner, 2013).
Seed coat cells
During seed development, the ovule integuments differentiate into several cell layers composed of different specialized
cells which will form the protective seed coat or testa (Bewley,
1997; Debeaujon et al., 2007) (Fig. 1). Seed coat cells develop
heavily lignified secondary cell walls to reinforce the outer surface of the seed mechanically and to make it impermeable to
liquids and gasses (Kelly et al., 1992; Liang et al., 2006; Chen
et al., 2012, 2013). Seed coat lignins are different between angiosperm species (orchids and cactaceaes) and are composed of
a mixture of classic G- and S-units and/or non-canonical
C- (caffeyl) units which derive from an unusual monomer, caffeyl alcohol (para- and meta-hydroxylated but not methoxylated) (Tobimatsu et al., 2013). The importance of seed coat
lignin for the proper function of the seed is clearly illustrated
by the arabidopsis transparent testa 10 mutant, which accumulates less lignin in seeds and shows reduced germination rates
after vernalization (Liang et al., 2006). However, the physicochemical properties and potential effects of lignin on embryogenesis, seed fitness, dormancy and germination still remain
unknown.
Siliques cells
In A. thaliana and related species, the external envelope of
fruits consists of three main tissues: the valves or seedpod
walls; the replum or central ridge located between the valves;
and the valve margins which separate the valves from the
replum to disperse the seeds (Fig. 1). The release of seeds from
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their pods is enabled by a thin band of cells which form the dehiscence zone located between the replum and the valves. Cell
wall lignification occurs specifically in valve margin cells adjacent to the dehiscence zone as well as in an internal valve cell
layer (Fig. 1). During the shattering of siliques, the middle lamella between the dehiscence zone cells breaks and the separation of the cells allows the valve to separate from the replum to
release the seeds. The lignin polymer composition of silique
valve cells has not been reported to date.
Lignin biosynthesis can also be triggered in responses to various biotic and abiotic stresses such as during wounding
(Delessert et al., 2004; Kim et al., 2006), pathogen infection
(Moerschbacher et al., 1990; Martı́n et al., 2007), drought (Fan
et al., 2006; Yoshimura et al., 2008), UV radiation (Rozema
et al., 1997; Hilal et al., 2004), low temperature (Hausman
et al., 2000; El Kayal et al., 2006), reduced nutrient availability
(Blodgett et al., 2005; Tahara et al., 2005) and CO2 or ozone
exposure (Davey et al., 2004; Cabané et al., 2004) (Fig. 1).
These stress-induced lignins generally impregnate the primary
cell wall of cells that are normally not lignified (i.e. leaf epidermal or stem pith parenchyma cells) and exhibit lower amounts
of S-units and higher amounts of G- and H-units as well as
more condensed C–C linkages between units (Lange et al.,
1995; Stange et al., 2001; Hawkins and Boudet 2003; Cabané
et al., 2004).
CELL BIOLOGY OF LIGNIN MONOMER
SYNTHESIS
Lignin deposition depends on the cell type, the developmental
stage and the species. This spatial distribution is characterized
by differences in time, amount, size and monomeric composition of the lignin polymer (Terashima et al., 2012). For example, TE lignin in gymnosperm wood is typically composed of
G-units with a minor contribution of H-units, while in angiosperm wood, TE secondary walls contain mainly G-units and
sclerenchyma fibres have G- and S-units. The overall H/G/S
proportions in plant TE-rich tissues vary from 0–5/95–100/0 in
gymnosperms, to 0–8/25–50/46–75 in dicot angiosperms and
5–33/33–80/20–54 in monocots/grasses (Ek et al., 2009). The
pathway synthesizing these different lignin monomers combines the reduction of the c-carbon terminal function from a
carboxylic acid to an alcohol and the meta and para substitution
of the aromatic ring with hydroxyl and methoxyl groups.
Lignin monomer precursor(s) derive from the aromatic
amino acid phenylalanine, synthesized in the plastid, which
is converted into 4-hydroxyphenylpropene alcohols. Monocotyledon plants also possess the capacity to use tyrosine as an additional precursor (Fig. 2). The enzymatic steps leading to the
synthesis of these lignin monomers have been extensively reviewed (Boerjan et al., 2003; Bonawitz and Chapple, 2010;
Vanholme et al., 2010), although new biosynthetic steps within
the pathway are still being discovered. The caffeoyl shikimate
esterase (CSE) (Vanholme et al., 2013), which catalyses the
conversion of caffeoyl shikimic/quinic acid into caffeic acid,
has been identified, as has the monocotyledon-specific p-coumaroyl-CoA:monolignol transferase (PMT) (Petrik et al., 2013)
which enables the formation of c-ester-bound monolignols acetylated with p-coumarate. In addition, the mobilization and
Barros et al. — The cell biology of lignification in higher plants
1056
COMT
CSE
PAL
phenylalanine
C4H/
REF3
4CL
4CL
HCT
ferulic acid
caffeic acid
tricin
tyrosine
TAL
C3H/
REF8
feruloyl hexose
CAldh/
REF1
CCoAOMT
HCT
feruloyl malate
4CL
SGT
cinnamic acid
p-coumaric acid
p-coumaroyl CoA
CCR
CCR1
p-coumaroyl
shikimic /
quinic aicd
caffeoyl shikimic
/quinic aicd
caffeoyl CoA
CCR
CCR1
feruloyl CoA
sinapic acid
sinapioyl glucose
CAldh/
REF1
CCR
CCR1
F5H/
FAH1
COMT
PMT
SMT
COMT
cinnamoyl hexose
p-coumaraldehyde
caffeyl aldehyde
CAD
CAD
coniferaldehyde
CAD
CAD
F5H/
FAH1
COMT
cinnamoyl malate
p-coumaroyl
alcohol
caffeyl alcohol
5-hydroxyconiferaldehyde
coniferyl alcohol
UGT
BGLU
sinapaldehyde
CAD
sinapoyl malate
COMT
5-hydroxyconiferyl
alcohol
sinapyl alcohol
UGT
BGLU
Cytoplasm
p-coumaroyl p-coumarate R=R’=H
coniferyl p-coumarate R=H, R’=OCH3
sinapyl p-coumarate R=R’=OCH3
Lignin monomers
coniferin
syringin
Plasma membrane
BGLU
Secondary cell wall
H units
H - H/G/S units
C units
G units
BGLU
S units
Primary cell wall
FIG. 2. General phenylpropanoid pathway showing lignin biosynthesis gene mutations in Arabidopsis thaliana (REF3, REF8, CCR1, REF1 and FAH1) and respective
side pathway reactions (coloured boxes) affected by the mutational change. Dotted boxes represent lignin monomers that are incorporated in the lignin polymer.
PAL, phenylalanine ammonia-lyase; TAL, tyrosine ammonia-lyase; C4H, cinnamate 4-hydroxylase; 4CL, 4-coumarate:CoA ligase; C3H, p-coumarate 3-hydroxylase; HCT, p-hydroxycinnamoyl-CoA:quinate/shikimate p-hydroxycinnamoyltransferase; CSE, caffeoyl shikimate esterase; CCoAOMT, caffeoyl-CoA O-methyltransferase; CCR, cinnamoyl-CoA reductase; CAD, cinnamyl alcohol dehydrogenase; COMT, caffeic acid O-methyltransferase; F5H, ferulate 5-hydroxylase; PMT,
p-coumaroyl-CoA:monolignol transferase; CAldh, coniferaldehyde dehydrogenase; UGT, UDP-glucosyltransferase; BGLU, b-glucosidase; SGT, sinapic acid:UDPglucosyl sinapoyltransferase; SMT, sinapoylglucose:malate sinapoyltransferase.
storage of monolignols is suggested to be regulated through
glycosylation/deglycosylation involving the enzymes UDPglucosyltransferase (UGT) and b-glucosidase (BGLU) (Liu,
2012). An overall scheme including these newly identified
genes/proteins is presented in Fig. 2.
The phenylpropanoid pathway forms a crossroad whose
branches lead to the synthesis of hormones, flavonoids, suberins
and many different phenolic compounds such as (neo)lignans
which derive from the same precursors as lignin (Vogt, 2010;
Weng and Chapple, 2010). This crossroad position is clearly
observed when key genes/enzymes of the pathway are genetically altered, leading to disturbances in both lignin and phenolic
compound accumulation. The disturbances include an increase
of cinnamoyl-malate in the At-REF3 mutant (loss of function
Barros et al. — The cell biology of lignification in higher plants
in cinnamate 4-hydroxylase, C4H; Fig. 2), an increase of
feruloyl-malate in the At-CCR1 mutant (loss of function in cinnamoyl-CoA reductase, CCR; Fig. 2) or a reduction of sinapoyl-malate in the At-REF1 mutant (loss of function in
coniferaldehyde dehydrogenase, CAldh; Fig. 2) as well as in
the At-REF8 mutant (loss of function in p-coumaroyl shikimate
3-hydroxylase, C3H; Fig. 2) and the At-FAH1 mutant (loss of
function in ferulate 5-hydroxylase, F5H; Fig. 2) (Nair et al.,
2002; Mir Derikvand et al., 2008; Bonawitz and Chapple,
2010).
Tissue/cell-specific expression of lignin monomer biosynthetic
genes
During plant development, lignin monomer biosynthetic
genes are expressed in all lignifying tissues (endodermis and
xylem) as well as in non-lignifying tissues such as the phloem
parenchyma and the epidermis. In lignifying tissues such as xylem, lignin monomer biosynthetic genes are also expressed in
the non-lignified cambium and in xylem parenchyma cells surrounding the lignifying TEs and fibres (Table 1; Fig. 3). During
normal development, it is difficult to decipher if both lignifying
and surrounding non-lignifying cells are producing lignin
monomers. The expression of the genes for phenylalanine ammonia-lyase (PAL), C4H, 4-coumarate:CoA ligase (4CL),
C3H, caffeoyl-CoA O-methyltransferase (CCoAOMT), cinnamyl alcohol dehydrogenase(CAD) and caffeic acid O-methyl
transferase (COMT) also depends on the circadian/diurnal cycle
(Rogers et al., 2005) and responds to biotic and abiotic stresses
(Hawkins and Boudet, 2003; El Kayal et al., 2006; Fan et al.,
2006; Kim et al., 2006; Hano et al., 2006; Yoshimura et al.,
2008; Moura et al., 2010). As for many other genes, lignin biosynthetic genes are part of small multigenic families which
sometimes show specificity for a distinct type of lignin. This is
the case of At-CCR in A. thaliana which has two isozymes
coded by different genes: At-CCR1 is preferentially expressed
during development, and its loss-of-function mutants have reduced lignification (Jones et al., 2001), while At-CCR2 is expressed at a low level during development but is strongly
induced during the defence reaction triggered by the pathogen
Xanthomonas ssp. and could participate in formation of stressinduced lignin (Lauvergeat et al., 2001). The differential expression of genes coding for isoforms of enzymes in lignin
monomer biosynthesis can represent one of the mechanisms
contributing to the specific type of lignin deposited in distinct
cell types: At-CCR1 during formation of developmental lignin
and At-CCR2 during formation of pathogen stress-induced
lignin
Sub-cellular localization of related lignin monomer synthesis
enzymes
While the aromatic amino acid precursor(s) initiating lignin
monomer synthesis derive from plastids, the enzymes implicated in lignin monomer synthesis are cytoplasmic and/or associate with the outer surface of the endoplasmic reticulum (ER)
(Table 2). All the different studies which have investigated the
sub-cellular localization of these enzymes are listed in Table 2
and summarized in Fig. 4. Based on these studies, the
1057
cytochrome P450 oxidoreductases responsible for the aromatic
ring hydroxylation (C4H, C3H and F5H) are on the outer surface of the ER, whereas all the other lignin monomer biosynthetic enzymes (PAL, 4CL, CCoAOMT, CCR, CAD and
COMT) are in the cytoplasm. However, a connection exists between the cytoplasm- and ER-localized enzymes as protein
complexes can form between cytoplasmic PAL and ER-localized C4H (Achnine et al., 2004), as well as between all the ERlocalized cytochrome P450 oxidoreductases: C4H, C3H and
F5H (Chen et al., 2011). Moreover, HCT (p-hydroxycinnamoyl-CoA:quinate/shikimate p-hydroxycinnamoyltransferase)
and 4CL were found to be partially associated with the ER
upon expression of C3H, suggesting an interaction between
these proteins (Bassard et al., 2012). The oxido-reductive state
of the different cytochrome P450 oxidoreductases, required for
the hydroxylation of the aromatic ring, is maintained by the cytochrome P450 reductase 2 (ATR2) (Sundin et al., 2014). Thus,
the synthesis of monolignols occurs in specialized sub-cellular
areas at the interface between the cytoplasm and the outer surface of the ER (Fig. 4B).
Lignin monomer transport mechanisms
Although the synthesis of monolignols occurs within the
cell protoplast, lignin deposition is restricted to the cell walls.
Three types of transport mechanisms are suggested for the extracellular secretion of lignin monomers, namely passive diffusion (PD), vesicle-associated exocytosis and active ATPdependent transport using ABC transporters and/or proton
coupled antiporters (Fig. 5). The ATP-dependent transport
mechanism using ABC-G transporters has been identified for
p-coumaryl alcohol (H-unit) export (Miao and Liu, 2010;
Alejandro et al., 2012). However, the inhibition of ABC transporters in different xylem tissues (poplar, hybrid poplar,
Japanese cypress and pine) using vanadate did not affect
the transport of coniferin, whereas proton gradient erasers
markedly reduced the transport, suggesting an export via proton-coupled antiporters (Tsuyama et al., 2013). The passive
diffusion is supported by in vitro observations of the partitioning of lignin monomers by immobilized liposomes and lipid
bilayer discs (Boija and Johansson, 2006; Boija et al., 2007);
however, this mechanism is considered unlikely to play a major role in the mobilization of monolignols (Miao and Liu,
2010). Vesicle-associated secretion of lignin monomers was
initially supported by the labelling of ER- and Golgi-derived
vesicles in xylem TEs when feeding with tritium-labelled phenylalanine and subsequent autoradiographical imaging using
electron microscopy (Pickett-Heaps, 1968; Fujita and Harada,
1979; Takabe et al., 1985). Improved radiolabelling feeding
and detection techniques have since allowed the observation
that part of the label incorporation is due to proteins rather
than monolignols (Kaneda et al., 2008). Exocytosis of lignin
monomers by vesicles was pharmacologically tested in in vitro
TE differentiating cell cultures from Zinnia elegans using the
exocytosis inhibitor brefeldin A (Ito et al., 2004). These experiments revealed a reduction of secreted phenolics, although
the impact on lignin accumulation was not estimated. The
mechanisms ensuring lignin monomer biosynthesis and export
to the lignifying sites within the cell wall are still unclear.
Mostly in veins and midrib
Mostly in veins and midrib
Mostly veins and midrib
Mostly in veins and midrib
Not defined
Not defined
Not defined
Veins and midrib
Primary xylem of midrib
No expression
Not defined
Not defined
Xylem vessel
Not defined
Veins and midrib
Not defined
Not defined
Not defined
Not defined
Not defined
Veins and midrib
No expression
Whole leaf, veins and midrib
Not defined
Not defined
Nicotiana tabacum
Arabidopsis thaliana
Arabidopsis thaliana
Arabidopsis thaliana
Loblolly pine
Nicotiana tabacum
Medicago sativa
Arabidopsis thaliana
Populus tremula alba
Populus trichocarpa
Populus tremula alba
Populus tremuloides
Fragaria ananassa
Saccharum officinarum
Arabidopsis thaliana
Zea mays
Nicotiana tabacum
Saccharum officinarum
Populus sieboldii grandidentata
Medicago sativa
Zea mays
Arabidopsis thaliana
Arabidopsis thaliana
Populus trichocarpa
Populus tremuloides
Aecondary xylem
Vascular bundles
Cambium, ray parenchyma and XPs
Cambium, ray parenchyma and XPs
Vascular bundles
Not defined
Not defined
XPs
Phloem, xylem fibres and XPs
Vascular bundles and interfascicular
fibres
Diff. xylem cells; XPs
Xylem tissue
developping xylem
Xylem and phloem parenchyma
Not defined
Xylem and phloem parenchyma
Xylem vascular bundle and interfascicular fibres
Ray parenchyma and XPs
xylem and phloem parenchyma
XPs
Xylem fibres, young ray parenchyma, cambium and xylem
vessels
Xylem and phloem parenchyma
Vascular system
XPs
Columella root cap; central
cylinder
No expression
Vascular system
Not defined
Not defined
Not defined
XPs
Not defined
Not defined
Diff. xylem cells; XPs
No expression
Not defined
XPs
Xylem vessel
Not defined
Cambium
Vascular system
Vascular system in mature
xylem zone
Vascular system
Vascualr cylinder
Diff. xylem cells
Not defined
Not defined
Not defined
Xylem tissue
Low expression
Whole root
Vascular bundles
Not defined
Vascular system
Not defined
Vascular tissue and
epidermis
Entire root except meristem
and root tip
Entire root except meristem
Vascular system
Not defined
Not defined
Epidermis; primary xylem;
phloem fibres
Not defined
Not defined
Root
GUS
GUS
IL
IL
IL
GUS
IL
IL
IL
IL
GUS
qRT-PCR
ISH
IL
IL
IL
GUS
GUS
GFP/GUS
IL
IL
IL
GUS
GUS
GUS
GUS
IL
GUS
GUS and ISH
IL
GUS
GUS
GUS
ISH
GUS
Li et al. (1999)
Goujon et al. (2003)
Li et al. (2001)
Li et al. (2001)
Ruelland et al. (2003)
Quentin et al. (2009)
Li et al. (2001)
Ruelland et al. (2003)
Kersey et al. (1999)
Sato et al. (2009)
Chen et al. (2000)
Shi et al. (2010)
Lacombe et al. (1997)
Ruelland et al. (2003)
Blanco-Portales et al. (2002)
Ruelland et al. (2003)
Sibout et al. (2003)
Nair et al. (2002)
Vanholme et al. (2013)
Hoffmann et al. (2004)
Kersey et al. (1999)
Zhong et al. (2000)
Do et al. (2007)
Nair et al. (2002)
Capellades et al. (1996)
Hu et al. (1998)
Hoffmann et al. (2004)
Lee et al. (1995)
Gui et al. (2011)
Sato et al. (2009)
Pesquet et al. (2013); Serk et al. (2015)
Nair et al. (2002)
Franke et al. (2000)
Harding et al. (2002)
Bell-Lelong et al. (1997)
Ohl et al. (1990)
Hsieh et al. (2010)
Zhu et al. (1995)
Sato et al. (2009)
IL
GUS
IL
GUS
Gray-Mitsumune et al. (1999)
Osakabe et al. (2006)
Reference
GUS
GUS
Technique
XP, xylem parenchyma; GUS, b-glucuronidase; ISH, in situ hybridization; IL, immunolabelling; GFP, green fluorescent protein; qRT-PCR, quantitative real-time PCR.
F5H
COMT
CCR
CAD
CSE
CCoAOMT
HCT
C3H
Nicotiana tabacum
Populus tremuloides
4CL
Arabidopsis thaliana
Oryza sativa
Populus sieboldii grandidentata
Cambium, xylem parenchyma
Vascular bundles
Xylem vessel and XPs
Phloem fibres; rays; xylem;
cambium
Diff. xylem cells
Mostly phloem cells
Not defined
Epidermis and vascular bundles
Xylem fibres, young ray parenchyma, cambium and xylem
vessels
Vascular bundles
XPs; diff. xylem cells
Veins and midrib
Mostly veins and midrib
Not defined
Mostly in xylem of midrib
Xylem of midrib
No expression
Mostly in veins and midrib
Epidermis and vascular bundles
Not defined
Xylem and sclerified parenchyma
Mostly veins and midrib
Arabidopsis thaliana
Epidermis; primary xylem; phloem
fibres
Xylem fibres, young ray parenchyma, cambium and xylem
vessels
Vascular system
Vascular tissue and epidermis
Vascular tissue and epidermis
Xylem and phloem fibres
Stem
C4H
Whole leaf except veins and
midrib
epidermis; primary xylem;
phloem fibres
Not defined
Leaves
Mostly veins and midrib
Not defined
Not defined
Populus trichocarpa deltoides
Populus sieboldii grandidentata
Populus kitakamiensis
Species
Arabidopsis thaliana
Bambusa oldhamii
Oryza sativa
PAL
Protein
TABLE 1. Tissue-specific localization of lignin monomer biosynthesis transcripts/proteins in different species found in the literature
1058
Barros et al. — The cell biology of lignification in higher plants
Barros et al. — The cell biology of lignification in higher plants
1059
B
A
C
D
TEs
XPs
RPs
XFs
6 PAL
83
83
67
100
4 C4H
50
100
25
50
5 4CL
100
80
60
60
1 HCT
100
100
100
100
2 C3H
100
100
1 CSE
100
100
100
100
100
100
67
6 CCoAOMT 50
100
1 CCR
100
100
100
100
5 CAD
40
80
60
20
7 COMT
57
100
71
57
1 F5H
100
100
100
100
FIG. 3. Cell-specific expression of lignin monomer synthesis transcripts and proteins. Stem cross-sections of (A) arabidopsis, (B) Brachypodium and (C) Populus indicating the different cell types included in the studies in (D). Blue arrow, tracheary elements (TEs); orange arrow, xylem parenchyma (XPs); red arrow, ray parenchyma (RPs); purple arrow, xylary fibres (XFs). Scale bars ¼ 100 lm. (D) Percentage of studies supporting cell-specific expression of lignin monomer biosynthesis
genes in TEs, XPs, RPs or XFs. Numbers in the first column indicate the number of individual studies, respectively, for each gene.
Lignin monomer plasticity
The identification of 4-hydroxyphenyl propene alcohols as
the typical lignin monomers essentially results from in vitro polymerization studies. Monolignols can be used in combination
with phenoloxidases to produce dehydrogenation polymers
(DHPs) which have similar characteristics to lignin (Brunow
and Lundquist, 1980). Several recent studies, however, revealed
that lignin synthesis is not only restricted to the three canonical
lignin monomers but also incorporates other unusual phenylpropene molecules exhibiting other substitutions such as monolignol precursors (i.e. hydroxycinnamic acid and aldehydes),
esters of monolignols (coniferyl and sinapyl acetate or coumarate) (Morreel et al., 2004; Grabber et al., 2010; Vanholme
et al., 2010; Wilkerson et al., 2014) but also other phenolic residues such as the flavone tricin (Del Rı́o et al., 2012; Lan et al.,
2015) (Fig. 2). Furthermore, the specific accumulation of dilignols and trilignols has been observed in methanol-extractable
fractions of the apoplast of lignifying xylem tissues (Morreel
et al., 2004) and from the extracellular medium of lignifying
Zinnia TE cell cultures (Tokunaga et al., 2005).
However, the question remains whether these compounds
represent intermediates of lignin synthesis, distinct apoplastic
precursors needed for lignin synthesis or unrelated phenolics.
Purified phenolic compounds accumulating in the extracellular medium of TE differentiating Zinnia cell cultures (which
are chemically different from monolignols) were able to restore fully the lignin reduction of TEs treated with inhibitors
of PAL, thus supporting the large diversity of potential/actual
lignin monomers (Tokunaga et al., 2005). Moreover, due to
the potential toxicity and low water solubility of monolignols
(Whetten and Sederoff, 1995; Boerjan et al., 2003), 4-O-glucosides represent one of the substitutions which could combine both reduced toxicity and higher water miscibility.
These monolignol glucosides are unable to form lignin without the intervention of a b-glucosidase in vitro (Chapelle
et al., 2012) but can be taken up by plants to be incorporated
into lignin (Tsuji and Fukushima, 2004) and are specifically
transported by ATP-dependent transporters in the endomembrane system of lignifying woody tissues (Tsuyama et al.,
2013). In addition, monolignol glucosides are highly accumulated in arabidopsis mutants which have extremely reduced
Barros et al. — The cell biology of lignification in higher plants
1060
TABLE 2. Sub-cellular localization of related lignin monomer biosynthesis proteins in different species found in the literature
Protein
PAL
C4H
4CL
HCT
C3H
CCoAOMT
CCR
CAD
COMT
F5H
Species
Phaseolus vulgaris
Primula kewensis
Zinnia elegans
Poplar kitakamiensis
Populus
sieboldii grandidentata
Phyllanthus tenellus
Poplar kitakamiensis
Nicotiana tabacum
Arabidopsis thaliana
Nicotiana tabacum
Populus
sieboldii grandidentata
Populus trichocarpa
Nicotiana benthamiana
Phaseolus vulgaris
Poplar kitakamiensis
Oryza sativa
Nicotiana benthamiana
Nicotiana benthamiana
Nicotiana benthamiana
Populus trichocarpa
Medicago sativa
Medicago sativa
Zinnia elegans
Zea mays
Oryza sativa
Zea mays
Zinnia elegans
Populus tremula alba
Poplar euramericana,
Eucalyptus globulus
Zea mays
Medicago sativa
Poplar euramericana,
Eucalyptus globulus
Medicago sativa
Medicago sativa
Populus trichocarpa
Tissue/cell type
Localization
Technique
Reference
Hypocotyl/XP, TE
Head cells of glandular trichomes
TE
XP, fibre and vessels
XP, fibre and vessels
Cytosol
ER
Vesicles, PM and SCW
Cytosol
cytosol, ER, Golgi
IL
IL
IL
IL
IL
Smith et al. (1994)
Schöpker et al. (1995)
Nakashima et al. (1997)
Takabe et al. (2001)
Sato et al. (2004)
Leaf mesophyl cell
XP, TE
Leaf epidermal cell
Epidermal cells of hypocotyl,
cotyledon and VND7-induced TEs
Leaf epidermal cells
XP, fibre and vessels
Cytosol, chloroplast
Plastid, cytosol
Cytosol (col. C4H)
ER
IL
IL
FRET
GFP fusion
ER (col. PAL)
Cytosol, ER, Golgi
FRET
IL
Santiago et al. (2000)
Osakabe et al. (2006)
Achnine et al. (2004)
Ro et al. (2001) and
Schuetz et al. (2014)
Achnine et al. (2004)
Sato et al. (2004)
Protoplast
Leaves
Hypocotyl/XP, TE
Xylem fibre
Protoplast
Leaves
Leaves
Leaves
Protoplast
Stem/XP, TE
Stem
Developing TE
Mesocotyl and root
Protoplast
Mesocotyl and root
TE
Stem
GFP fusion
FRAP
IL
IL
GFP fusion
FRAP
FRAP
FRAP
GFP fusion
IL
Western
IL
Western
GFP fusion
Western, IL
IL
IL
Chen et al. (2011)
Bassard et al. (2012)
Smith et al. (1994)
Takabe et al. (2001)
Rastogi et al. (2013)
Bassard et al. (2012)
Bassard et al. (2012)
Bassard et al. (2012)
Chen et al. (2011)
Kersey et al. (1999)
Guo et al. (2002)
Ye (1997)
Ruelland et al. (2003)
Kawasaki et al. (2006)
Ruelland et al. (2003)
Nakashima et al. (1997)
Šamaj et al. (1998)
Xylem fibre
ER (col. C4H)
ER (col. C3H)
ER
Cytosol
Cytosol
cytosol (ER)
Cytosol (ER)
ER (col. C4H)
ER (col. C3H)
Cytosol
Cytosol
Cytosol
Cytosol
Cytosol
Cytosol
Vesicles, PM and SCW
ER and Golgi-derived
vesicles
cytosol
IL
Takabe et al. (2001)
Mesocotyl and root
Stem/XP, TE
differentiating xylem cells
Cytosol
Cytosol
Cytosol
Western, IL
IL
IL
Ruelland et al. (2003)
Kersey et al. (1999)
Takeuchi et al. (2001)
Stem
Stem
Differentiating xylem protoplasts
Cytosol
ER
ER
Western
Western
GFP fusion
Guo et al. (2002)
Guo et al. (2002)
Wang et al. (2012)
XP, xylem parenchyma; IL, immunolabelling; GFP, green fluorescent protein; FRET, fluorescence resonance energy transfer; FRAP, fluorescence recovery
after photobleaching; col., co-localization with PAL/C4H/C3H; ER, endoplasmic reticulum; PM, plasma membrane; SCW, secondary cell wall.
amounts of lignin (Zhao et al., 2013). However, the genetic
loss-of-function mutants in UGT and BGLU, required for the
addition and removal of the glucoside moiety, do not affect
lignin accumulation or composition in arabidopsis plants
(Lanot et al., 2006; Chapelle et al., 2012). In contrast to any
other cell wall polymer, lignin appears to exhibit a
tremendously high plasticity for its monomers.
Lignin monomer production – autonomous or co-operative
process
The localization of the lignin monomer biosynthetic gene
expression and their corresponding protein is present in the
non-lignifying cells next to lignifying cells. For example, all
the lignin monomer biosynthetic genes are expressed in both
the actively lignifying interfascicular fibres of arabidopsis and
the non-lignified meristematic cells of the cambium and in the
associated parenchyma (xylem parenchyma during primary
growth and ray cell files in secondary wood; Table 1; Fig. 3).
This observation has led to the idea that cell wall lignification
could proceed through cell co-operation where non-lignifying
cells would provide the monomers and/or other substrates/enzymes to the cell wall of actively lignifying cells. This mechanism is referred to as ‘the good neighbour hypothesis’ or ‘the
non-cell autonomous process’ (Smith et al., 2013; Pesquet
et al., 2013). The lignification of distinct cell types can therefore be autonomous (a cell undergoing lignification produces
enzymes and substrates for its lignification) and/or
co-operative (neighbouring cells provide enzymes and/or substrates to lignifying cells). Co-operative lignin synthesis is
clearly observed between TEs and fibres in the xylem of poplar
trees as fibres directly neighbouring TEs present an intermediate lignin structure enriched in G-units like TEs, in contrast to
their S-unit enrichment when only surrounded by fibres
(Gorzsás et al., 2011). Although cell co-operation was hypothesized many years ago using the Zinnia TE differentiating cell
Barros et al. — The cell biology of lignification in higher plants
1061
A
Tyr
Phe
Nucleus
Vacuole
ER
Monomer
glucoside
Common pathway
H units
Cytosol
8 PAL
6 C4H
3 4CL
1 HCT
2 C3H
3 CCoAOMT
2 CCR
4 CAD
4 COMT
2 F5H
ER
Cunits
G units
Golgi
Plastids
40
Vesicles
20
13
Electrons
S units
Primary cell wall
Plasma membrane
Secondary cell wall
B
Monomer
7
PM
13
7
CW
7
75
7
13
75
25
100
100
100
100
25
13
13
25
13
13
100
100
FIG. 4. Sub-cellular localization of lignin monomer biosynthesis proteins. (A) Scheme illustrating the lignin monomer biosynthesis pathway and monomer transport
in respect to sub-cellular localization of the lignin monomer synthesis proteins. Tyr, tyrosine; Phe, phenylalanine. (B) Percentage of studies supporting specific
subcellular localization in: cytosol, endoplasmic reticulum (ER), Golgi, plastids, vesicles, plasma membrane (PM) or cell wall (CW). Numbers in the first column
indicate the number of individual studies, respectively, for each protein. ATR2, arabidopsis cytochrome P450 reductase 2.
culture system (Hosokawa et al., 2001), it cannot be generalized to all lignifying cell types. The Casparian strip of endodermal cells appears to depend on cell autonomous lignification
(Alejandro et al., 2012), whereas xylem TEs appear to rely on a
co-operative lignification with neighbouring xylem TE precursors and/or the surrounding parenchyma cells (Pesquet et al.,
2013; Smith et al., 2013).
The functional demonstration of TE non-cell-autonomous
lignification was provided recently by several complementary
studies including: (1) the pharmacological inhibition of
lignification in Z. elegans in vitro TE differentiating cell cultures, which could be reverted when washing the inhibitor
away once all TEs have committed cell death – which implies
that the monolignols are supplied by the surrounding living
cells – thereby demonstrating the cell co-operation during TE
post-mortem lignification (Tokunaga et al., 2005; Pesquet
et al., 2013); (2) arabidopsis plants knocked-out in xylem
parenchyma-expressed genes implicated in co-operative lignification (i.e. At-RCD1 or At-MYB13) exhibited quantitative and
qualitative changes in xylem lignin (Pesquet et al., 2013);
Barros et al. — The cell biology of lignification in higher plants
1062
Monolignols
O2
PD
Monomer radical
activation
Monomeric coupling
(with dirigent proteins)
End-wise
polymer
extension
LACCASE
Vesicle
DiP
ABC trans.
/ PCA
O2
NADPH
NADPH
O2
oxidase
.
O2 -
DiP
Redox Shuttle
SOD
Mn (II)
Mn (III)
Monomeric coupling
NADP+ H+
Monolignols
H2O2
PD
Vesicle
PEROXIDASE
ABC trans.
/ PCA
Monomer
Symplast
Plasma
membrane
Dimer
Secondary cell wall
Lignin polymer
Primary
cell wall
FIG. 5. General lignin polymerization. Lignin monomers are exported across the plasma membrane either by passive diffusion (PD), by exocytosis (vesicle) or
through ABC transporters and/or proton-coupled antiporter (PCA). Laccases and peroxidases activate the monomer radicals, resulting in the end-wise addition of
and/or cross-reaction of the radical oligo/monomers with the extending polymer(s). Classical production of H2O2 and O2 derives from a two-step enzymatic process:
NADPH oxidase and superoxide dismutase (SOD). Dirigent proteins (DiPs) stereospecifically restrict the radical coupling to one type of linkage. Manganese (Mn)
acts as a redox shuttle to mediate radical activation (Onnerud et al., 2002). This schematic representation does not support equal importance or intervention of
different possibilities used by specific cell types to form the lignin polymer.
(3) specific silencing of the lignin monomer biosynthetic gene
At-CCR1 in cells developing secondary cell walls using the
IRX3/CesA7 promoter (specific to cells that develop secondary
cell walls) did not completely abolish TE lignification, suggesting that thin-walled parenchyma contributes to TE lignification
(Smith et al., 2013); and lastly (4) complementation of the lignin monomer biosynthetic gene At-C4H knock-out mutant with
the promoter of master regulator VND6 (specific to metaxylem
TE precursor cells) driving native C4H, which restored TE lignification, suggesting that thin-walled TE precursor cells might
also contribute co-operatively to TE lignification (Yang et al.,
2013). Similarly, the partial cell-autonomous lignification of
xylem fibres was also demonstrated by specifically silencing
At-CCR1 using the IRX3/CesA7 promoter, which reduces fibre
lignin accumulation (Smith et al., 2013) and by complementing
the At-C4H knock-out mutant with a VND6 promoter-driven
C4H, which partially restores lignin accumulation in fibres
(Yang et al., 2013). This suggests that fibre lignification is partially co-operating with the cambium and/or metaxylem TE
cells. Altogether, cell-specific lignin deposition appears to result from distinct mechanisms depending on the cell type, from
full cell autonomy to completely co-operative. However, the
exact biological function and significance of these different levels of cell autonomy for lignification still needs to be fully
understood.
LIGNIN POLYMER FORMATION
Compared with other cell wall polymers such as pectins and
hemicelluloses, which are assembled remotely and exported to
the cell wall, the formation of the lignin polymer occurs directly
in the cell wall by the oxidative polymerization of secreted
lignin monomers. The activated monolignol radicals present a
relatively long half-life (approx. 45 s for phenoxy radicals;
Harkin, 1967), partly stabilized by resonance, and are capable
of diffusing through the wall to polymerize into lignin.
Depending on both thermodynamical reactivity and steric accessibility, the different radical mesomers will assemble to
form the different linkages of the polymer including both condensed C–C linkages such as 5–5’, b–5’, b–b and b–1’, and
non-condensed C–O–C ether linkages such as b-O-4’
(Terashima and Fukushima, 1988). The diversity of the most
prevalent intermonomeric linkages in different plant taxa is illustrated in Table 3. Both the polymer length and the proportion
of different linkages can be explained by the availability and reactivity of specific lignin monomer(s) radicals (Ralph et al.,
2008).
The impact of monomer availability on the polymer length is
illustrated during in vitro DHP formation, which increases in
size when supplied continuously with new monomers compared
with when supplied with an initial bulk (Tanahashi and
Barros et al. — The cell biology of lignification in higher plants
1063
TABLE 3. Percentages of the different interunit linkages in lignin of different plant taxa
Species
Sample
b–O-4
5–5
b–5
b–1
b–b
Reference
Spruce
Pine
Birch
Poplar
Arabidopsis
MMW
MMW
MMW
Wood
Stem
Alfalfa
Maize
Stem
Stem
Wheat
Straw
48
62
60
69
79
77
81
100
60
75
10
–
5
–
–
1
1
–
–
3
10
20
6
3
7
15
8
–
27
11
7
–
7
–
–
1
–
–
3
3
2
18
3
28
14
6
6
–
10
4
Adler (1977)
Mansfield et al. (2012)
Adler (1977)
Mansfield et al. (2012)
Mansfield et al. (2012)
Bonawitz et al. (2014)
Marita et al. (2003)
Mansfield et al. (2012)
Min et al. (2014)
Del Rı́o et al. (2012)
b–O-4, beta-aryl ether; 5–5, biphenyl and dibenzodioxocin; b–5, phenylcoumaran; b–1, 1,2-diaryl propane; b–b, resinol; MMW, mature milled wood.
Higuchi, 1981). The proportion of the different linkages in the
polymer depends on the available monomer type: for example
in the case of knock-out/down mutation of the COMT gene
(Fig. 2), more of the normally rare lignin sub-structure benzodioxane is detected in arabidopsis, Medicago and Populus (Lu
et al., 2010; Moinuddin et al., 2010) due to the increased presence of the unconventional 5-hydroxyconiferyl alcohol. Unlike
cellulose, in which the addition of new monomers to the extending polymer occurs directly by a polymerizing enzyme,
lignin end-wise addition to extend the polymer with radical
oligo/monomers occurs independently of processing enzymes
(Fig. 5). The mono/oligolignol radicals are produced enzymatically by two kinds of phenol-oxidoreductase enzymes, O2dependent laccases and H2O2-dependent peroxidases, which
have both been shown to enable monolignols to polymerize
into DHPs in vitro (Sterjiades et al., 1992, 1993).
Laccase-catalysed lignin polymer formation
Plant laccases (EC 1.10.3.2), or blue copper-containing oxidoreductases, are part of a medium size multigenic family (17
genes in arabidopsis and 39 genes in Populus trichocarpa;
Weng and Chapple, 2010) with an N-terminal peptide signal
guiding the protein through the secretory pathway. Laccases
can act as p-diphenol:O2 oxidoreductases by using O2 directly
to oxidize all types of monolignols to form DHPs (Higuchi and
Ito, 1958; Sterjiades et al., 1992; Bao et al., 1993; Ranocha
et al., 1999; Kärkönen et al., 2002; Liang et al., 2006).
Laccases do not appear to have specifically evolved to enable
plant lignin polymer formation and present a high degree of
structural conservation between bacteria, fungi and plants
(Dwivedi et al., 2011). The number of laccases has slightly
increased, ranging during plant evolution from three in
Chlamydomonas reinhardtii, to 12 in Physcomitrella patens,
ten in Selaginella moellendorffii and 17–39 in angiosperms
(Weng and Chapple, 2010). Interestingly, laccases in saprophytic fungi exhibit lignolytic activity and catalyse wood lignin
disassembly (Coy et al., 2010; Crestini et al., 2010). The diversity/redundancy of laccase function in plants is shown by multiple minor phenotypic defects in single loss-of-function mutants
in arabidopsis laccases: they show minor changes such as a
higher susceptibility to polyethylene glycol (PEG)-induced dehydration for laccase-2, early flowering for laccase-8 or pale
seed colour for laccase-15 (Cai et al., 2006).
The experimental evidence for the intervention of laccases
during lignification is supported by (1) their specific transcription profiles – laccases are co-regulated with lignin monomer
biosynthesis genes (Gavnholt and Larsen, 2002), with secondary cell wall-forming genes (Brown et al., 2005) and expressed
in lignifying tissues in arabidopsis (Berthet et al., 2011), and
(2) their enzymatic activity: laccase activities were detected
during xylem lignification in several species (Dean and
Eriksson, 1994; Liu et al., 1994; Richardson and McDougall,
1997; Dean et al., 1998; Ranocha et al., 1999; Kärkönen et al.,
2002; Ranocha et al., 2002; Caparrós-Ruiz et al., 2006;
Koutaniemi et al., 2015). Functional studies have now confirmed the role of specific laccase isoforms during the lignification of specific cell types (Tables 4 and 5).
In arabidopsis, laccase-15 (At-LAC15) is responsible for seed
coat lignification, and the loss-of-function mutant (named transparent testa 10 because of reduced seed coat brownish colour)
exhibits a 30 % decrease in seed lignin content (Liang et al.,
2006). The At-LAC15 mutant has a significantly reduced capacity to oxidize coniferyl alcohol and an increased presence of
b–b/b–5 and b–O-4 linkages in seed cells compared with wildtype seeds (Liang et al., 2006). Arabidopsis laccases-4, -11 and
-17 are responsible for the lignification of both xylem TEs and
fibres (Berthet et al., 2011; Zhao et al., 2013). Single and double
loss-of-function mutants in these laccases show a subtle effect
on stem growth in normal conditions, but double mutants present a significant reduction in G-type lignin (Berthet et al.,
2011) (Table 5). The triple mutant exhibits an extremely reduced growth and a dramatic reduction in xylem lignin, suggesting that xylem lignification results from the combined activity
of these three laccases (Zhao et al., 2013) (Table 5).
Arabidopsis laccase-17 (At-LAC17) is localized in the secondary cell walls of xylem vessels and stem interfascicular fibres,
as well as xylem parenchyma, and vascular and interfascicular
cambium, while At-LAC4 is present in cambium, xylem parenchyma and interfascicular fibres, visible in both the compound
middle lamella and the secondary cell wall of xylem vessels and
fibres (Berthet et al., 2011, Schuetz et al., 2014) (Table 4).
Similarly to arabidopsis, poplar xylem lignification also depends
on multiple laccase isoforms. The downregulation of a single
wood-expressed laccase using constitutive antisense constructs
did not reduce lignin quantity or composition but interestingly
altered the cell wall structure of xylem fibres (Ranocha et al.,
2002) whereas the general silencing of poplar laccases using the
constitutive overexpression of microRNA397a – which targets
Barros et al. — The cell biology of lignification in higher plants
1064
TABLE 4. Tissue-/cell-specific expression and impact on growth and vessel morphology of mutations in different peroxidases and laccases associated with lignification in Arabidopsis thaliana plants
Protein
AT - number
Expressed in tissue/cell type
Modulation
Growth
Vessel
Lac4
At2g38080
Cambium, xylem parenchyma,
interf. fibres
KO
No effect
IRX
Lac11
At5g03260
KO
No effect
–
Lac15
At5g48100
KO
Shorter root
–
Lac17
At5g60020
KO
No effect
Slight IRX
Lac4/Lac17
Lac4/Lac11
Lac11/Lac17
Lac4/Lac11/ Lac17
Prx2
Prx4
Prx25
Prx37
Prx47
Prx52
Prx53
Prx64
–
–
–
–
At1g05250
At1g14540
At2g41480
At4g08770
At4g33420
At5g05340
At5g06720
At5g42180
KO
KO
KO
KO
KO
KO
KO
35S OE
–
KO
–
KD
No effect
No effect
No effect
Dwarf
No effect
No effect
No effect
Shorter
–
No effect
–
–
IRX
–
–
–
No effect
No effect
No effect
No effect
–
No effect
–
–
Prx66
At5g51890
Xylem parenchyma, vascular and
interf. cambium
Seed coat, xylem parenchyma,
vascular and interf. cambium
Xylem vessels and parenchyma,
interf. cambium and fibres,
vascular cambium
–
–
–
–
–
–
–
Vascular bundle
Endodermis, xylem parenchyma
–
Vascular bundle
Cortex, epidermis, endodermis,
xylem fibres
Xylem vessels
–
–
–
Prx71
Prx72
Prx2/Prx25
Prx2/Prx71
Prx25/Prx71
At5g64120
At5g66390
–
–
–
–
–
–
–
–
KO
KO
KO
KO
KO
No effect
Shorter
No effect
No effect
No effect
No effect
IRX
No effect
No effect
No effect
Reference
Berthet et al. (2011);
Turlapati et al. (2011);
Zhao et al. (2013)
Turlapati et al. (2011);
Zhao et al. (2013)
Liang et al. (2006);
Turlapati et al. (2011)
Berthet et al. (2011);
Turlapati et al. (2011)
Berthet et al. (2011)
Zhao et al. (2013)
Zhao et al. (2013)
Zhao et al. (2013)
Shigeto et al. (2013)
Fernández-Pérez et al. (2015)
Shigeto et al. (2013)
Pedreira et al. (2010)
Tokunaga et al. (2009)
Fernández-Pérez et al. (2014)
Østergaard et al. (2000)
Tokunaga et al. (2009);
Lee et al. (2013)
Sato et al. (2006);
Tokunaga et al. (2009)
Shigeto et al. (2013)
Herrero et al. (2013)
Shigeto et al. (2015)
Shigeto et al. (2015)
Shigeto et al. (2015)
KO, knock-out mutant; KD, knock-down; OE, overexpressor; interf., interfascicular; IRX, irregular xylem phenotype (collapsed vessels).
29 out of the 47 poplar laccases – induces significant reduction
of wood lignin in poplar (Lu et al., 2013).
Peroxidase-catalysed lignin polymer formation
Plant peroxidases, mostly represented by class III type peroxidases, are part of a large multigenic family (73 genes in arabidopsis) with a peptide signal that targets the proteins through
the secretory pathway (Hiraga et al., 2001; Valério et al.,
2004). In contrast to laccases, the number of peroxidase isoforms has increased tremendously during plant evolution, ranging from none in C. reinhardtii, to 43 in the non-lignified moss
P. patens, 79 in S. moellendorffii and 73–138 in angiosperms
(Weng and Chapple, 2010). Plant class III peroxidases (EC
1.11.1.7) have been shown to catalyse the formation of DHPs
efficiently in vitro using H2O2 and monolignols (Sterjiades
et al., 1993; Guerra et al., 2000). The enzymatic activity of peroxidases to produce DHPs appears to be more specific to coniferyl alcohol and peroxidases appear to be unable to use sinapyl
alcohol as a substrate (At-PRX53, Østergaard et al., 2000; AtPRX34, Demont-Caulet et al., 2010). However, some purified
peroxidases from other plant species (poplar and silver birch)
have shown affinity for sinapyl alcohol (Tsutsumi et al., 1998;
Fagerstedt et al., 2010).
Like laccases, peroxidases are expressed in lignifying vascular tissue (Table 4). In TE differentiating cell cultures of Zinnia,
the homologous protein to At-PRX66 is localized in TE
secondary cell walls and its recombinant protein showed a
strong preference for coniferyl alcohol (Sato et al., 2006). Like
laccases, the active role of peroxidases during lignin formation
was shown by reverse genetic approaches in multiple species:
the downregulation of PRX60 in tobacco plants (Blee et al.,
2003) or of PRX3 in aspen (Li et al., 2003) resulted in phenotypes which had reduced lignin accumulation and altered lignin
composition. In A. thaliana, endodermal Casparian strip lignification was significantly reduced when silencing At-PRX64 in
the root endodermis (Lee et al., 2013). Moreover, arabidopsis
single or double knock-out mutants in At-PRX2, At-PRX71 and
At-PRX25 exhibited reduced lignin accumulation without affecting stem height (Shigeto et al., 2013, 2015), and the mutation of At-PRX72 caused a reduction in lignin and stem height
(Herrero et al., 2013) (Tables 4 and 5).
Second substrate requirement – H2O2 and/or O2 – during lignin
polymer formation
Besides the lignin monomer, peroxidases and laccases require additional substrates – hydrogen peroxide (H2O2) and molecular oxygen (O2), respectively – to form monolignol radicals
(Fig. 5). Limiting concentrations of O2 are unlikely in normal
growth conditions as O2 is present in large proportion in the air
and is transported in the xylem sap. Gansert (2003) suggests
that O2 contributes to 70 % of xylem oxygenation in birch.
However, during anoxia/hypoxia, the triggered arrest of plant
Barros et al. — The cell biology of lignification in higher plants
1065
TABLE 5. Impact on total lignin content and H-, G-, and S-unit composition of mutations in different peroxidases and laccases
associated with lignification in Arabidopsis thaliana plants
Protein
Lac4
Lac11
Lac15
Lac17
Lac4/Lac17
Lac4/Lac11
Lac11/Lac17
Lac4/Lac11/Lac17
Prx2
Prx4
Prx25
Prx37
Prx52
Prx64
Prx66
Prx71
Prx72
Prx2/Prx25
Prx2/Prx71
Prx25/Prx71
Modulation
KO
KO
KO
KO
KO
KO
KO
KO
KO
KO
KO
35S OE
KO
KD
–
KO
KO
KO
KO
KO
Total lignin
H-units
G-units
S-units
S/G ratio
;
No effect
;
;
;
;
;
;
;
;
;
–
;
–
–
No effect
;
;
;
;
No effect
No effect
–
No effect
No effect
No effect
No effect
–
–
–
–
–
–
–
–
–
:
No effect
No effect
No effect
;
No effect
–
;
;
;
;
–
No effect
:
:
–
:
–
–
:
–
:
:
:
No effect
No effect
–
No effect
–
;
;
–
:
;
:
–
;
–
–
:
–
:
:
:
;vb :if
–
–
;vb :if
–
–
–
–
:
;
:
–
;
–
–
:
No effect
:
:
:
:, increase; ;, decrease in total lignin; KO, knock-out mutant; KD, knock-down; OE, overexpressor; vb, vascular bundle; if, interfascicular fibres. References
for each study are provided in Table 4.
growth is coupled to an increase of H2O2 (Blokhina et al.,
2001) as well as an increase of lignin in both the Casparian strip
and the sclerenchyma cells in rice (Kotula et al., 2009). The
pharmacological scavenging of H2O2, using KI or catalase,
clearly reduces the lignification of the Casparian strip (Lee
et al., 2013), the co-operative lignin formation in TEs (Pesquet
et al., 2013) and the extracellular lignin accumulation in Picea
abies cell cultures (Kärkönen et al., 2002). Similarly to other
reactive oxygen species (ROS), H2O2 molecules have extremely short half-lives ranging from 109 s to 1 ms
(D’Autréaux and Toledano, 2007) and are thus unable to accumulate, which suggests that they are constantly produced. The
production of apoplastic H2O2 derives from a two-step enzymatic process using nicotinamide adenine dinucleotide phosphate hydrogen oxidase (NADPH oxidase) and superoxide
dismutase (Ogawa et al., 1997).
NADPH oxidase
NADPH oxidases, also called Respiratory Burst Oxidase
Homolog (RBOH) proteins, are plasma membrane proteins
with six conserved transmembrane domains and cytosolic
FAD- and NADPH-binding sites, which catalyse the production
of apoplastic superoxide (O2–) from cytoplasmic NADPH
(Sagi and Fluhr, 2006). In arabidopsis, NADPH oxidases are
part of a small multigenic family of ten isoforms labelled from
A to J, which are expressed in different organs and in response
to both plant development and defence (Torres et al., 2006;
Sagi and Fluhr, 2006; Lee et al., 2013). In rice, the synthesis of
stress-induced lignin by pathogen attack is mediated by the
Rac/Rop small GTPases RAC1 which binds to CCR1
(Kawasaki et al., 2006) and activates NAPDH oxidase
(Kawasaki et al., 1999). Similarly, xylem parenchyma cells
have been shown to express specifically Rac small GTPase
(Nakanomyo et al., 2002) and produce O2–/H2O2 in a polarized
way, only on the cell side in contact with TEs (Ros Barceló,
1998, 2005). The functional association of NADPH oxidase
with lignification was unravelled using both pharmacological
inhibition by diphenyleneiodonium (DPI) and genetic analysis
using loss-of-function mutants. DPI treatment prevents the lignification of the endodermal Casparian strip (Lee et al., 2013)
and reduces the co-operative lignification of Zinnia TE cell cultures (Pesquet et al., 2013). Mutations in NADPH oxidase
RBOH-F (AT1G64060) deplete the endodermal Casparian strip
of lignin without apparently affecting TE lignification (Lee
et al., 2013). Moreover, mutants in xylem parenchymaexpressed At-RCD1 (Radical-Induced Cell Death1; Overmyer
et al., 2000) present an elevated amount of ROS as well as an
increase of lignin in the xylem (Pesquet et al., 2013). The AtRCD1 mutant phenotype can be partially compensated when
mutating NADPH oxidase isoforms D and F (Zhu et al., 2013),
suggesting that these enzymes control the appropriate supply of
ROS for lignification. Similarly to the cell-specific expression
of different phenoloxidase isoforms, distinct NADPH oxidase
isoforms appear to be associated with the lignification of specific cell types.
High isoelectric point superoxide dismutase (SOD)
Superoxide dismutases (EC 1.15.1.1), divided into three different forms (CuZn-SOD, Fe-SOD and Mn-SOD), catalyse the
dismutation of toxic superoxide radicals produced by NADPH
oxidase into O2 and H2O2 substrates which can be used by laccases or peroxidases, respectively (Liochev and Fridovich,
1994). Superoxide dismutation by CuZn-SOD (Karpinska
et al., 2001) was confirmed pharmacologically to occur in lignifying xylem tissues of spinach (Ogawa et al., 1997) and showed
distinct subcellular localization in the secondary cell walls of
1066
Barros et al. — The cell biology of lignification in higher plants
xylem TEs and fibres, as well as in the middle lamella of surrounding xylem parenchyma cells in scots pine, poplar and
Zinnia TEs (Karpinska et al., 2001; Karlsson et al., 2005;
Srivastava et al., 2007). Pharmacological inhibition of SOD reduced Zinnia TE H2O2 production as well as TE lignin accumulation (Karlsson et al., 2005) and similarly reduced the
lignification of the endodermal Casparian strip (Lee et al.,
2013). Nevertheless, genetic modulation of CuZn-SOD using
antisense constructs in transgenic poplar reduced the growth
rate but only slightly reduced lignin accumulation (Srivastava
et al., 2007). Interestingly, constitutive overexpression of cytoplasmic SOD in arabidopsis had the opposite effect and improved performance of plant growth under salt stress due to an
increase in secondary cell wall synthesis and possibly lignin accumulation (Gill and Tuteja, 2010). Overall, these results suggest that SODs are, at least partially, involved in the
lignification of different tissues.
lignification (Zhao et al., 2013), suggesting that laccases and
peroxidases do not function redundantly. However, this cell
specificity for phenoloxidases is not so clear-cut as xylem TE
lignification requires both laccases and peroxidases (Table 5),
and collapsed TE phenotypes have been observed in lossof-function mutants of both At-LAC4 (Brown et al., 2005;
Berthet et al., 2011) and At-PRX72 (Herrero et al., 2013). This
led to the idea that laccases and peroxidases could act in a sequential order during cell lignification (Sterjades et al., 1993),
starting with laccases and followed by peroxidases, to accomplish full cell wall lignification. The late intervention of peroxidases was further supported as they can form rigid cross-links
between lignin, hemicelluloses and extensins in the secondary
cell wall (Mäder et al., 1977, 1980; Lamport, 1986; Lagrimini
et al., 1987; Passardi et al., 2004) once the initial lignin
polymers/oligomers are made by laccases. Three hypothetical
models could therefore explain this dual intervention of
laccases and peroxidases during cell wall lignification.
Intermonomeric linkage and dirigent proteins
Model of sequential intervention due to different substrate
specificity. In this model, the radical activation of the mono-
The mechanisms controlling the proportion of different intermonomeric linkages within the lignin polymer in the different
parts of specific cell types is still poorly understood. Each specific linkage has been considered to result from differences in
thermodynamic reactivity of the monomer radical resonance
forms (Watts et al., 2011) but have also been suggested to
depend on specific proteins, named dirigent proteins (DiPs)
(Fig. 5). DiPs govern the stereochemistry of the oxidative coupling of coniferyl alcohol catalysed by both peroxidases and
laccases, leading to the production of a less thermodynamically
favourable b–b linkage compared with b–O-4 [observed during
(neo)lignan(s) synthesis; Halls and Lewis, 2002]. DiP genes
have been found to be expressed during xylem lignification,
and their corresponding proteins are localized in secondary cell
walls of cells undergoing lignification (Kwon et al., 1999).
Functional evidence of the intervention of DiP in lignification
was provided by the loss-of-function mutant of the DiP
enhanced suberin 1 (ESB1, AT2G28670) which exhibited both
an increased lignin deposition and a misorganization of lignin
in the endodermal Casparian strip (Hosmani et al., 2013). The
biological control of intermonomeric linkages by DiPs could
explain the structural difference observed between DHPs and
native lignins (Davin and Lewis, 2000). The extent to which
random chemical coupling compared with DiP-dependent coupling contributes to the cell-specific lignin polymer formation
remains, however, unknown.
Lignin polymerization: laccase and peroxidase?
Although both laccases and peroxidases can catalyse the formation of monolignol radicals and form DHPs in vitro, these
enzymes appear to be differently implicated in the cell wall lignification of specific cell types. In A. thaliana, the Casparian
strip lignification depends on peroxidases (essentially AtPRX64; Lee et al., 2013) whereas xylem TE and fibre lignification are mostly dependent on laccases (essentially At-LAC4,
At-LAC11 and At-LAC17; Zhao et al., 2013). Interestingly, the
laccase triple knock-out mutant which has severely reduced
xylem lignification is not affected in its Casparian strip
mer(s) first requires laccases to produce the initiating oligolignols and/or the core lignin polymers which are then subjected
to peroxidases for further branching and/or extension. This
hypothesis relies on the capacity of peroxidases and laccases to
oxidize distinct substrates, other than monolignols, as shown by
histological detection of phenoloxidase activities in lignifying
tissue: 4-methoxy-a-naphthol reveals the presence of peroxidases but not laccases, whereas ABTS [2,2-azinobis-(3-ethylbenzothiazoline-6-sulphonate)] and coniferaldehyde show both
peroxidase and laccase activities in lignifying poplar (De Pinto
and Ros Barceló, 1997; Ranocha et al., 1999). Thus, ligninassociated laccases and peroxidases would catalyse the radical
activation of distinct molecules produced sequentially during
the end-wise polymerization of lignin.
Model of sequential intervention due to different spatio-temporal
expression. In this model phenoloxidase enzymes are differen-
tially localized/secreted within the TE primary and secondary
cell wall. The localization of laccase and peroxidase gene expression is found in both lignifying and non-lignifying xylem
cells, suggesting both cell-autonomous and co-operative processes (Table 5). Moreover, differences in the spatio-temporal
expression of specific phenoloxidase during tissue lignification
could explain the sequential intervention of laccases and
peroxidases.
Model of differential protein complex formation. In this model,
phenoloxidases are associated together with other proteins
(DiPs or others) forming protein complexes restricting substrate
specificity and/or localization in the cell wall. This model is
supported by the laccase/cellobiose dehydrogenase interaction
in Pycnoporus cinnabarinus fungus which specifically enables
the formation of the phenol-derived pigment cinnabarinic acid
(Temp and Eggert, 1999). Moreover, a multicomponent protein
complex containing manganese (II)-dependent peroxidase,
laccase and b-glucosidase was found in the fungus Lentinula
edodes which was able to transform pentachlorophenol and 2,5dichlorophenol (Makkar et al., 2001). Such protein complexes
could form at a specific time and/or specific sites to enable
specific steps required for lignification.
Barros et al. — The cell biology of lignification in higher plants
Together, these models aim to explain the mechanisms leading to differential lignin accumulation, size and composition in
distinct sub-domains of the cell wall (middle lamella vs. secondary cell wall) of specific cell types (xylem TEs vs. fibres),
although currently our understanding is still largely
fragmentary.
LIGNIN FORMATION DURING THE
DIFFERENTIATION OF SPECIFIC CELLS
Lignification is an integral part of the differentiation process
of several specialized cell types to fulfil their physiological
function. However for each distinct cell type, lignin deposition exhibits differences in (1) its timing during the cell differentiation process (before and/or after cell death), (2) its
dependency on specific enzymes and/or substrates, (3) its
sub-cellular deposition in the cell wall, (4) its monomeric
composition and (5) its autonomy for the production of the
required enzymes and/or substrates. This complexity is
illustrated by the multiple phenotypic defects observed in
lignin synthesis knock-out mutants in A. thaliana which have
altered lignification of seeds (Liang et al., 2006), reduced
height with a lower lignification of xylem fibres (Smith
et al., 2013), defects in xylem TE resistance during sap conduction (Jones et al., 2001) and/or increased susceptibility to
root pathogen infection (Cahill and Mc Comb, 1992; Wuyts
et al., 2007). It is therefore difficult to generalize lignification to one common cellular mechanism and, in order to
provide a better picture of the different deposition mechanisms, the lignification processes during the differentiation
of three distinct cell types are described below.
Lignification of xylem tracheary elements
The vascular and mechanical support functions of TEs is established by the removal of the cell cytoplasm by programmed
cell death and the reinforcement of the TE cell side walls with a
lignified secondary cell wall (Pesquet and Lloyd, 2011). TE differentiation from the elongated cells of the meristematic cambium progresses sequentially with (1) a microtubule-guided
cellulose and xylan secondary cell wall deposition (Oda et al.,
2010; Pesquet et al., 2010), which is followed by (2) the cellautonomous programmed cell death, and concluded by (3) the
post-mortem lignification of the TE secondary cell wall and the
autolysis of the cell content (Fig. 6A). In primary tissues, TE
lignin deposition is restricted to the secondary cell wall thickenings and it is completely absent from the modified TE primary
cell walls in TEs of Coleus (Hepler et al., 1970), Zinnia (Taylor
et al., 1992) and arabidopsis (Schuetz et al., 2014; Serk et al.,
2015). On the other hand, in secondary tissues, TE lignin deposition is present in both secondary and primary cell walls
(Fromm et al., 2003; Gierlinger, 2014; Herbette et al., 2015;
Serk et al., 2015). Interestingly, the lignification of TEs in secondary tissues initiates at the cell corner of the middle lamella
and then gradually progresses through the secondary cell wall
layers (Terashima and Fukushima, 1988, 1989; Donaldsson
et al., 1999).
The initiation of TE lignification before or after TE cell death
is still debated, although all agree that TE lignin deposition will
1067
continue once TE cells have died (Pesquet et al., 2013; Smith
et al., 2013). This post-mortem lignification of TEs operates by
the co-operative supply of lignin monomers and ROS by both
the surrounding precursor cells and xylem parenchyma (Ros
Barceló et al., 2005; Pesquet et al., 2013). The main enzymes
associated with TE lignin oxidative polymerization are AtLAC4 (Brown et al., 2005) and At-PRX72 (Herrero et al.,
2013), as both single loss-of-function mutants have collapsed
TE cell walls. This defect is due to altered secondary cell wall
structures observed also when reducing lignin biosynthesis in
plants either pharmacologically or genetically such as when AtCCR1 is mutated in IRX4 (Jones et al., 2001). Based on the expression of other phenoloxidases in cells surrounding TEs
(Table 4), At-LAC4 and At-PRX72 probably act with other
phenoloxidases produced in both TEs and xylem parenchyma
(Berthet et al., 2011; Zhao et al., 2013).
The biological significance of TE post-mortem lignification
has not been yet unravelled, but one hypothesis is that TEspecific non-cell-autonomous lignification is a mechanism to
enable optimal hydro-mineral sap flow without renewing the
entire vascular system (Ménard and Pesquet, 2015). As the
plant increases in height and width, the tension pressure that is
necessary to enable sap rising as well as the pressure within the
stem tissues increases. Post-mortem lignification of TEs might
have evolved to withstand the increasing pressure in the stem
by prolonged lignification of the TEs along with the increase in
height and girth of the stem.
Lignification of xylem fibres
Sclerenchyma fibres are believed to derive from a primitive
ancestral xylem cell which evolved into TEs mainly for the sap
conduction and fibres for mechanical support (Lucas et al.,
2013). In contrast to TEs, fibres do not require cell death to
achieve their structural support function. Sclerenchyma fibre
cells derive from the xylem meristem and differentiate as they
elongate by tip growth with a microtubule-dependent deposition of xylan and cellulose in the secondary cell wall (Chaffey
et al., 2002) which is then impregnated with lignin, starting at
the cell corner of the compound middle lamella and progressing
gradually inwards towards the innermost layer of the secondary
cell wall (Donaldson, 2001) (Fig. 6B). Although both TEs and
fibres develop thick lignified secondary cell walls, xylem fibres
present a lignin enriched in S-units. Unlike TEs, xylem sclerenchyma fibres in arabidopsis are mostly lignified by cellautonomous processes. Reduced lignin accumulation in fibres
is observed in both the loss-of-function mutants in the lignin
monomer biosynthesis gene At-CCR1 (Jones et al., 2001) and
in plants silencing At-CCR1 using artificial microRNA driven
by the secondary cellulose synthase promoter IRX3 (Smith
et al., 2013). However partial co-operative lignification of xylem fibres is also observed with the neighbouring cells: poplar
xylem fibres next to TEs exhibit a lignin composition intermediate between that of TEs and fibres with a polar enrichment in
G-units (Gorzsás et al., 2011). Lignin oxidative polymerization
in sclerenchyma fibres depends on multiple laccases (Berthet
et al., 2011; Lu et al., 2013; Zhao et al., 2013) as well as peroxidases (Blee et al., 2003; Herrero et al., 2013; Shigeto et al.,
2013) (Tables 4 and 5).
Barros et al. — The cell biology of lignification in higher plants
1068
A
Co-operative lignification (tracheary elements)
Tracheary element (alive)
- hemi-/cellulose deposition -
Cambial cell
Neighbouring xylem/ray
parenchyma cell
Tracheary element (dead)
- lignin deposition -
Monolignols
Monolignols
PCD
Differentiation
H2O2
LAC/PRX
Lac17/Prx 72
LAC/PRX
Lac4/Prx72
.
O
O2
B
O2
2
Partial co-operative lignification (xylem fibres)
Xylem fibre
- Lignin deposition -
Xylem fibre
- hemi-/cellulose deposition -
Cambial cell
Monolignols
Differentiation
Neighbouring xylem/ray
parenchyma cell
Monolignols
Monolignols
Lignification
LAC/PRX
ROS
LAC/PRX
ROS
C
Autonomous lignification (casparian strip)
CASP domain
formation
Localization of NADPH
oxidase and SOD
Localization
of ESB1
Export of monolignolsand
peroxidases for lignification
Monolignols
O2
.
O2-
.O
2
PRX
Prx64
.O
2
H2O2
H2O2
PRX
Prx64
Monolignols
NADPH oxidase
Plasma membrane
CASP
Primary cell wall
Peroxidase
SOD
ESB1
Non-lignified secondary cell wall
ABCG transporter
Lignified secondary cell wall/
apoplastic space
FIG. 6. Different levels of co-operative and autonomous lignification depending on the cell types studied: xylem tracheary elements, xylem fibres and endodermal
cells. (A) Different stages of TE formation showing post-mortem TE lignification through neighbouring parenchyma cells that provide monolignols, laccases/
peroxidases (LAC/PRX) and O2/H2O2 produced by NADPH oxidase and superoxide dismutase (SOD). Living neighbouring TEs may also provide monolignols and
incorporate LAC/PRX in the secondary cell wall before programmed cell death (PCD). (B) Differentiation and partial co-operative lignification of xylem fibres
showing that monolignols, LAC/PRX and reactive oxygen species (ROS) are produced by both xylem fibres and neighbouring parenchyma cells. Living unlignified
fibres may also provide monolignols. (C) Autonomous lignification in endodermal cells during Casparian strip formation, showing the formation of the Casparian
strip domain by CASP proteins (Casparian strip membrane domain proteins) and the localization of ESB1 (enhanced suberin 1) in the Casparian strip zone. Later on,
H2O2 (produced by NADPH oxidase and SOD), monolignols (exported by ABCG transporters) and PRX are supplied to the Casparian strip zone.
Barros et al. — The cell biology of lignification in higher plants
Lignification of root endodermal cells
Endodermal cells constitute a tightly bound root cell layer
isolating and regulating the flux of compounds/nutrients from
the root cortex in contact with the rhizosphere to the root
medullar vascular system (Geldner, 2013). This barrier property
is directly due to the apoplastic polarized deposition of a lignified and suberized Casparian strip which frames and radially interconnects each endodermal cell (Fig. 1). Lignin deposition in
this cell type is achieved independently of cell death and is necessary to establish the apoplastic barrier (Naseer et al., 2012).
The differentiation of these cells derives from the root tip and,
once cells have fully elongated, the Casparian strip is formed
(Fig. 6C) by: (1) establishing specific polarized domains,
named CASPs (Casparian strip membrane domain proteins),
which will delimit the future sites of the Casparian strips both
at the plasma membrane (localization of At-CASP1; Roppolo
et al., 2011) and in the neighbouring apoplastic space (localization of ESB1; Hosmani et al., 2013); (2) NADPH oxidase
RBOH-F is targeted to the CASP domain and At-PRX64 is
exported into the CASP- delimited apoplastic space (Lee et al.,
2013); (3) lignin monomers are secreted in the apoplast,
some by an unpolarized ABC transporter such as ABCG29
(Alejandro et al., 2012); finally, (4) lignin is polymerized in
the apoplastic space delimited by the CASP domain using peroxidases, the apoplastic monomers and H2O2 (from O2– generated by RBOH-F and converted by an unidentified apoplastic
SOD).
CONCLUSIONS
Lignin is associated with many different specialized cells to fulfil specific physiological functions, but exhibits distinct properties for each cell type, which may explain why no general
mechanism for lignification has been yet defined. Thus depending on the desired lignin properties for the cell function, specific substrates and enzymes will be produced in distinct cell
types although a high plasticity for both substrates and enzymes
will still be retained. Depending on the different cell types, lignification will progress autonomously and/or co-operate with
the surrounding cells to ensure full lignification and to adapt to
environmental changes.
ACKNOWLEDGEMENTS
The authors would like to acknowledge the memory of Barek
Tamasloukht. The authors thank Professor Leszek A.
Kleczkowski, Drs Emiko Muruzoka and Raphael Decou, Mrs
Delphine Ménard and Mr Charilaos Dimotakis for critical
comments. This research was supported by Vetenskapsrådet
(VR) research grant 2010-4620 (to E.P.), the Gunnar Öquist
Fellowship from the Kempe Foundation (to E.P.) and the Carl
Trygger Foundation (to E.P.).
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