Download Secondary Cell Walls: Biosynthesis, Patterned

Ruiqin Zhong and Zheng-Hua Ye*
Department of Plant Biology, University of Georgia, Athens, GA 30602, USA
*Corresponding author: E-mail, [email protected]; Fax, +1-706-542-1805.
(Received September 18, 2014; Accepted September 29, 2014)
Secondary walls are mainly composed of cellulose, hemicelluloses (xylan and glucomannan) and lignin, and are deposited in some specialized cells, such as tracheary
elements, fibers and other sclerenchymatous cells.
Secondary walls provide strength to these cells, which lend
mechanical support and protection to the plant body and, in
the case of tracheary elements, enable them to function as
conduits for transporting water. Formation of secondary
walls is a complex process that requires the co-ordinated
expression of secondary wall biosynthetic genes, biosynthesis
and targeted secretion of secondary wall components, and
patterned deposition and assembly of secondary walls. Here,
we provide a comprehensive review of genes involved in
secondary wall biosynthesis and deposition. Most of the
genes involved in the biosynthesis of secondary wall components, including cellulose, xylan, glucomannan and lignin,
have been identified and their co-ordinated activation has
been shown to be mediated by a transcriptional network
encompassing the secondary wall NAC and MYB master
switches and their downstream transcription factors. It has
been demonstrated that cortical microtubules and microtubule-associated proteins play important roles in the targeted
secretion of cellulose synthase complexes, the oriented deposition of cellulose microfibrils and the patterned deposition of secondary walls. Further investigation of many
secondary wall-associated genes with unknown functions
will provide new insights into the mechanisms controlling
the formation of secondary walls that constitute the bulk of
plant biomass.
Keywords: Cellulose Cortical microtubule Lignin Microtubule-associated protein Secondary wall Secondary wall NAC Xylan.
Abbreviations: Ara, arabinose; BC, brittle culm; CesA, cellulose synthase A; COBL, cobra-like; CSI, cellulose synthaseinteracting protein; CslA, cellulose synthase-like A; CTL, chitinase-like protein; DUF, domain of unknown function; ESK,
eskimo; FRA, fragile fiber; GalA, galacturonic acid; GlcA, glucuronic acid; GT, glycosyltransferase; GUX, glucuronic acid
substitution of xylan; GXM, glucuronoxylan methyltransferase; IRX, irregular xylem; KOR, korrigan; LAC, laccase; MAP,
microtubule-associated protein; MeGlcA, methylglucuronic
acid; MIDD, microtubule depletion domain; NST, NAC secondary wall thickening-promoting factor; PRX, peroxidase;
Rha, rhamnose; RWA, reduced wall acetylation; SMRE, secondary wall MYB-responsive element; SNBE, secondary wall
NAC-binding element; SND, secondary wall-associated NAC
domain protein; SWN, secondary wall NAC; VND, vascularrelated NAC-domain; XAT, xylan arabinosyltransferase; XAX,
xylosyl arabinosyl substitution of xylan; Xyl, xylose.
Introduction
Secondary walls are mainly found in tracheary elements (tracheids in seedless vascular plants and gymnosperms and vessels
in angiosperms) and fibers in the primary xylem and the secondary xylem (wood) (Fig. 1). They provide mechanical
strength to these cell types, which serve as mechanical tissues
to enable vascular plants to grow to a great height. Deposition
of lignified secondary walls in tracheary elements not only reinforces these water conduits to resist the negative pressure
generated during transpiration, but also renders them waterproof for efficient water transport. The ability of plants to deposit secondary walls in water-conducting elements was
evolved during the Silurian period (about 430 million years
ago), which is considered to be one of the pivotal steps for
vascular plans to conquer the terrestrial habitats (Raven et al.
1999). Secondary wall-containing cell types are also present in
tissues other than xylem for protection and structural support,
such as phloem fibers in stems of many dicot plants, extraxylary
fibers beneath the epidermis in grass stems, interfascicular
fibers between the vascular bundles in Arabidopsis stems
(Fig. 1A), and sclereids in pear fruits (also called stone cells)
and seed coats of jojoba (Mauseth 1988). In addition, secondary
wall deposition in anther endothecium is required for the generation of the tensile force necessary for stomium rupture to
release pollen grains (Yang et al. 2007). Similarly, secondary wall
deposition in extraxylary fibers of dehiscent seed pods is necessary for seed pod dehiscence leading to seed dispersal (Liljegren
et al. 2000).
The major components of secondary walls are cellulose,
hemicelluloses (xylan and glucomannan) and lignin (Fig. 1D).
Cellulose is the load-bearing unit, in which cellulose microfibrils
cross-link with hemicelluloses to form the framework of secondary walls. Hemicelluloses, such as xylan, are required for the
normal assembly and mechanical strength of secondary walls;
reduction in xylan content causes a severe decrease in cellulose
deposition, secondary wall thickening and mechanical strength
Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140, Advance Access publication on 7 October 2014,
available online at www.pcp.oxfordjournals.org
! The Author 2014. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
All rights reserved. For permissions, please email: [email protected]
Special Focus Issue – Review
Secondary Cell Walls: Biosynthesis, Patterned Deposition and
Transcriptional Regulation
R. Zhong and Z.-H. Ye | Secondary cell walls
Fig. 1 Deposition of secondary walls in xylem and fibers. (A) Cross-section of an Arabidopsis stem showing lignified secondary walls (red; stained
with phloroglucinol-HCl) in vessels (ve), xylary fibers (xf), interfascicular fibers (if) and phloem fibers (pf). co, cortex. (B) Transmission electron
micrograph showing three layers (S1, S2 and S3) of secondary walls in interfascicular fibers of an Arabidopsis stem. (C) Cellulose microfibrils in the
S2 layer of a developing interfascicular fiber cell of an Arabidopsis stem showing their nearly parallel orientations to the cell elongation axis
(double arrows). Scale bars in (A), (B) and (C) = 180, 1.3 and 0.28 mm, respectively. (D) Structural formula of secondary wall components,
including cellulose, hemicelluloses (glucuronoxylan, glucuronoarabinoxylan and glucomannan) and monolignols.
of vessels and fibers (Zhong et al. 2005b, Brown et al. 2007). The
essential role of xylan in secondary wall assembly is in sharp
contrast to primary wall xyloglucan, a complete loss of which
has no discernible effect on primary wall assembly (Cavalier
et al. 2008). Lignin, being hydrophobic and inert, impregnates
the cellulose and hemicellulose network to provide additional
mechanical strength, rigidity and hydrophobicity to secondary
walls. The composition of secondary walls, i.e. the proportion of
cellulose, hemicelluloses and lignin, may vary among different
plant species and even in different secondary wall-containing
cell types of the same species. For example, wood from a
gymnosperm species (Pinus strobus) consists of 41% cellulose,
9% arabinoglucuronoxylan, 18% galactoglucomannan and 29%
196
lignin, whereas wood from an angiosperm species (Populus
tremuloides) contains 48% cellulose, 24% glucuronoxylan, 3%
glucomannan and 21% lignin (Timell 1967). Secondary wall
composition may also change in response to different developmental and environmental stimuli. For example, compression
wood, which forms on the lower side of a leaning gymnosperm
stem, contains a higher lignin content and a lower cellulose
content compared with normal wood, whereas tension wood,
which forms on the upper side of a leaning angiosperm stem,
has an additional gelatinous layer in secondary walls that are
rich in cellulose but low in lignin content (Timell et al. 1967). In
addition, some specialized secondary walls may be composed
predominantly of one or two polymers. For example, secondary
Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140
walls of cotton fibers contain >90% cellulose with little xylan or
lignin (Haigler et al. 2012), and secondary walls of phloem fibers
from flax and hemp consist of cellulose and hemicelluloses
without lignin (Mauseth et al. 1988).
Because secondary walls in the form of wood and fibers are
important raw materials for our daily uses and a potential
source for biofuel production, there have been extensive studies
on the molecular mechanisms controlling secondary wall composition, biosynthesis and assembly. In this review, we provide
an update on the biosynthesis of secondary wall components,
patterned deposition of secondary walls, and transcriptional
regulation of secondary wall biosynthesis. Much of the current
knowledge on genes involved in secondary wall biosynthesis
stems from research in the model flowering plant,
Arabidopsis thaliana, and therefore we will mainly focus on
discussions of Arabidopsis secondary wall genes and only briefly
remark on their homologs in other species. Due to the evolutionary conservation of secondary wall biosynthesis, it is anticipated that knowledge gained from the study of Arabidopsis
secondary wall genes could be extrapolated into identifying
genes involved in secondary wall biosynthesis in other species.
Cellulose Biosynthesis
Cellulose, being the load-bearing polymer in secondary walls,
constitutes about 40–50% of wood components. It is made of
linear chains of b-1,4-linked glucosyl residues with a degree of
polymerization of approximately 10,000 (Timell et al. 1967)
(Fig. 1D). It has long been held that 36 glucan chains are coalesced via hydrogen bonding into a highly insoluble crystalline
microfibril. However, recent studies using X-ray and nuclear
magnetic resonance techniques give an estimate of the diameter of a primary wall microfibril to be about 3 nm, which fits
well with a model of 18 glucan chains per microfibril (Newman
et al. 2013). The biosynthesis of cellulose is mediated by cellulose synthase complexes located at the plasma membrane
(McFarlane et al. 2014). The complexes are visualized via transmission electron microscopy as rosettes made of six particles
(McFarlane et al. 2014), and each particle is proposed to be an
aggregate of three cellulose synthase A (CesA) subunits in the
18-CesA model (Newman et al. 2013). The 18-CesA model predicts that the 18 CesA subunits in each rosette synthesize 18
glucan chains that aggregate into a microfibril, which is consistent with the structural analysis of bacterial CesA revealing
that each CesA polypeptide synthesizes and translocates one
glucan chain (Morgan et al. 2013).
The Arabidopsis genome harbors 10 genes encoding CesA
proteins, which belong to glycosyltransferase family GT2
(McFarlane et al. 2014). Biosynthesis of secondary wall cellulose
has been shown to involve three of them, CesA4/IRX5, CesA7/
IRX3 and CesA8/IRX1, which function non-redundantly (Taylor
et al. 1999, Taylor et al. 2000, Taylor et al. 2003) (Table 1).
Mutation of any of them causes a severe reduction in cellulose
content and secondary wall thickening, and concomitantly a
collapsed xylem phenotype. The three secondary wall CesAs
interact with each other within the same cellulose synthase
complex, and the presence of all of them is required for the
normal assembly and targeting of the complex (Taylor et al.
2004, Timmers et al. 2009). In the 18-CesA model, each particle
in the six particle rosette is proposed to contain one of each of
the three interacting secondary wall CesAs (Newman et al.
2013). Close homologs of CesA4, CesA7 and CesA8 are present
in other vascular plants and their disruption has been shown to
result in a reduction in cellulose content and secondary wall
thickening in rice, Brachypodium and Populus (Tanaka et al.
2003, Joshi et al. 2011, Handakumbura et al. 2013).
In addition to CesA genes, a number of other genes have
been implicated in the biosynthesis and assembly of secondary
wall cellulose (Table 1). Mutation of the Arabidopsis
KORRIGAN (KOR) gene, which encodes an endo-b-1,4-glucanase, causes a reduction in cellulose content in both primary
and secondary walls (Nicol et al. 1998, Szyjanowicz et al. 2004).
Although KOR does not associate with the secondary wall CesA
complex (Szyjanowicz et al. 2004), it has been shown to be an
integral part of the primary wall CesA complex (Vain et al.
2014). KOR is proposed to function as a cellulase in releasing
the growing microfibrils from the cellulose synthase complex
and/or in increasing the amount of non-crystalline cellulose
microfibrils (Szyjanowicz et al. 2004, Takahashi et al. 2009).
Arabidopsis COBRA-like 4 (COBL4), a glycosylphosphatidylinositol-anchored protein, and its rice ortholog, brittle culm 1
(BC1), are also implicated in cellulose biosynthesis since their
mutations result in a reduction in cellulose content and secondary wall thickening (Li et al. 2003, Brown et al. 2005, Sato
et al. 2010). BC1 has a carbohydrate-binding module that interacts with crystalline cellulose, and this interaction is proposed
to modulate cellulose assembly and cellulose microfibril crystallinity (Liu et al. 2013). The Arabidopsis chitinase-like protein
CTL2, whose expression is associated with secondary wall CesA
genes, is proposed to modulate cellulose crystallization and
cellulose interaction with hemicelluloses via binding to growing
cellulose microfibrils (Sanchez-Rodriguez et al. 2012). A role for
CTLs in modulating cellulose biosynthesis is consistent with the
reduced cellulose content and mechanical strength of the rice
brittle culm mutant bc15, which harbors a mutation in a CTL
gene (Wu et al. 2012). The xylem-associated TED6 protein has
been shown to interact with secondary wall CesA7, and RNA
interference of its expression results in a reduction in secondary
wall thickening, indicating its involvement in cellulose biosynthesis (Endo et al. 2009). Several additional genes are also implicated in secondary wall cellulose biosynthesis, including two
Arabidopsis genes encoding fasciclin-like arabinogalactan proteins, AtFLA11 and AtFLA12, whose mutations reduce cellulose
content and alter cellulose microfibril angles (MacMillan et al.
2010), a rice gene encoding a dynamin-related protein (BC3/
OsDRP2B), whose mutation causes a brittle culm phenotype
with a decreased cellulose content and an altered secondary
wall structure (Hirano et al. 2010, Xiong et al. 2010), and another rice gene encoding a DUF266 domain protein, whose
mutation results in a brittle culm phenotype with a reduced
cellulose content (Y. Zhou et al. 2009). In addition, a number of
proteins with unknown functions have been shown to copurify with cellulose synthase complexes in Populus xylem,
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R. Zhong and Z.-H. Ye | Secondary cell walls
Table 1 Arabidopsis genes that are involved in secondary wall biosynthesis, patterned deposition and transcriptional regulation
Gene
AGI code
Cellulose biosynthesis and assembly
CesA4/IRX5
At5g44030
CesA7/IRX3
At5g17420
Functions
References
Cellulose synthase catalytic subunits; functioning redundantly in secondary wall cellulose biosynthesis
Taylor et al. (1999, 2000, 2003)
CesA8/IRX1
At4g18780
KORRIGAN
At5g49720
Endo-b-1,4-glucanase; required for normal
cellulose microfibril deposition
Nicol et al. (1998), Szyjanowicz et al. (2004)
COBL4
At5g15630
A glycosylphosphatidylinositol-anchored protein; modulating cellulose microfibril
deposition
Brown et al. (2005)
CTL2
At3g16920
A chitinase-like protein; modulateing cellulose microfibril deposition
Sanchez-Rodriguez et al. (2012)
TED6
At1g43790
Interacting with CesA7
Endo et al. (2009)
AtFLA11
At5g03170
MacMillan et al. (2010)
AtFLA12
At5g60490
Fasciclin-like arabinogalactan proteins;
involved in cellulose microfibril deposition
Two functionally non-redundant groups of
GT43 proteins (IRX9/ I9H vs. IRX14/I14H)
required for xylan backbone elongation
Brown et al. (2007), Pena et al. (2007), Lee
et al. (2010), Wu et al. (2010)
Xylan xylosyltransferases (GT47); catalyzing
the elongation of xylan backbone
Brown et al. (2009), Wu et al. (2009), Jensen
et al. (2014), Urbanowicz et al. (2014)
Xylan glucuronyltransferases (GT8); catalyzing
the addition of GlcA side chains in xylan
Mortimer et al. (2100), Lee et al. (2012a)
Glucuronoxylan methylatransferases
(DUF579); catalyzing GlcA methylation of
xylan
Lee et al. (2012d), Urbanowicz et al. (2012)
DUF579 proteins required for normal xylan
biosynthesis
Brown et al. (2011), Jensen et al. (2011)
Involved in acetylation of xylan; homologs of
fungal CAS1 protein required for acetylation of glucuronoxylomannan in fungi
Lee et al. (2011)
Xylan biosynthesis
IRX9
At2g37090
I9H/IRX9L
At1g27600
IRX14
At4g36890
I14H/IRX14L
At5g67230
IRX10/GUT2
At1g27440
IRX10L/GUT1
At5g61840
GUX1
At3g18660
GUX2
At4g33330
GUX3
At1g77130
GXM1
At1g09610
GXM2
At4g09990
GXM3/GXMT1
At1g09610
IRX15
At3g50220
IRX15L
At5g67210
RWA1
At5g46340
RWA2
At3g06550
RWA3
At2g34410
RWA4
At1g29890
ESK1/TBL29
At3g55990
Xylan acetyltransferase catalyzing 2-O- and 3O-monoacetylation of xylan (DUF231)
Xiong et al. (2013), Yuan et al. (2013),
Urbanowicz et al. (2014)
FRA8
At2g28110
At5g22940
Functional paralogs (GT47); required for the
biosynthesis of xylan reducing end
sequence
Zhong et al. (2005b), Lee et al. (2009)
F8H
IRX8
At5g54690
A GT8 protein; required for the biosynthesis
of xylan reducing end sequence
Brown et al. (2007, Pena et al. (2007,
Persson et al. (2007
PARVUS
At1g19300
A GT8 protein; required for the biosynthesis
of xylan reducing end sequence
Brown et al. (2007), Lee et al. (2007b)
Glucomannan synthases; cellulose synthaselike A family
Liepman et al. (2005), Goubet et al. (2009)
Glucomannan biosynthesis
CslA2
At5g22740
CslA3
At1g23480
CslA9
At5g03760
(continued)
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Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140
Table 1 Continued
Gene
AGI code
Lignin biosynthesis and polymerization
PAL1
At2g37040
Functions
References
Phenylalanine ammonia lyases; highly expressed in stems
Raes et al. (2003)
PAL2
At3g53260
PAL4
At3g10340
C4H
At2g30490
Cinnamate 4-hydroxylase; highly expressed in
stems
Raes et al. (2003)
4CL1
At1g51680
At3g21240
4-Commarate CoA ligases; highly expressed
in stems
Raes et al. (2003)
4CL2
HCT
At5g48930
Hydroxycinnamoyl CoA:shikimate hydroxycinnamoyl transferase; highly expressed in
stems
Raes et al. (2003)
C3H1
At2g40890
p-Coumaroyl shikimate 30 -hydroxylase; highly
expressed in stems
Raes et al. (2003)
CSE
At1g52760
Caffeoyl shikimate esterase; converting caffeoyl shikimate into caffeate
Vanholme et al. (2013)
Caffeoyl CoA O-methyltransferases; highly expressed in stems
Raes et al. (2003
CCoAOMT1
At4g34050
CCoAOMT7
At4g26220
CCR1
At1g15950
Cinnamoyl CoA reductase; highly expressed
in stems
Raes et al. (2003)
F5H1
At4g36220
Ferulate 5-hydroxylase; highly expressed in
stems
Raes et al. (2003)
COMT
At5g54160
Caffeic acid O-methyltransferase; highly expressed in stems
Raes et al. (2003)
Cinnamyl alcohol dehydrogenases; highly expressed in stems
Raes et al. (2003)
Laccases; catalyzing the polymerization of
monolignols
Berthet et al. (2011), Zhao et al. (2013)
Peroxidases; catalyzing the polymerization of
monolignols
Herrero et al. (2013b), Shigeto et al. (2013,
2014)
CAD3
At4g34230
CAD6
At4g37970
LAC4
At2g38080
LAC11
At5g03260
LAC17
At5g60020
AtPRX2
At1g05250
AtPRX25
At2g41480
AtPRX71
At5g64120
AtPRX72
At5g66390
PDR1/AtABCG29
At3g16340
ATP-binding cassette-like transporter;
involved in the transport of p-coumaryl
alcohol
Alejandro et al. (2012)
MED5a/REF4
At2g48110
At3g23590
Transcriptional co-regulatory complex subunits; required for homeostatic repression
of phenylpropanoid biosynthesis
Bonawitz et al. (2014)
MED5b/RFR1
Interacting with cortical microtubules and
CesAs; functioning as a scaffold protein to
guide the movement of CesA complexes
along the track of cortical microtubules
Gu et al. (2010), Bringmann et al. (2012), S.
Li et al. (2012)
Patterned deposition of secondary walls
CSI1
At2g22125
AtKTN1/FRA2
At1g80350
Katanin microtubule-severing protein; essential for cortical microtubule organization
and oriented deposition of cellulose
microfibrils
Burk et al. (2001, 2002)
FRA1
At5g47820
Kinesin motor protein; required for cellulose
microfibril deposition and secondary wall
strength
Zhong et al. (2002)
MAP65–8
At1g27920
Microtubule-associated protein whose zinnia
ortholog is involved in bundling of
microtubules
Mao et al. (2006)
(continued)
199
R. Zhong and Z.-H. Ye | Secondary cell walls
Table 1 Continued
Gene
AGI code
Functions
References
MAP70–1
At1g68060
Pesquet et al. (2010)
MAP70–5
At4g17220
Plant-specific microtubule-associated proteins;
regulating the organization of cortical
microtubule bundles
MIDD1
At3g53350
Interacting with microtubules; acting as a
scaffold protein to recruit Kinesin-13A for
microtubule depolymeirzation
Oda et al. (2010), Oda and Fukuda (2013a)
Kinesin-13A
At3g16630
Microtubule depolymerization protein; depolymerizing cortical microtubules underneath the pit-forming areas
Oda and Fukuda (2013a)
ROP11
At5g62880
GTPase; recruiting MIDD1/Kinesin-13A to the
microtubule-depleting plasma membrane
domains
Oda and Fukuda (2012)
ROPGAP3
At2g46710
ROP GTPase-activating protein; inactivating
ROP11
Oda and Fukuda (2012)
ROPGEF4
At2g45890
Guanine nucleotide exchange factor;
activating ROP11
Oda and Fukuda (2012)
Vessel-specific secondary wall NAC master
switches; functioning redundantly in regulating secondary wall biosynthesis in
vessels
Kubo et al. (2005), Yamaguchi et al. (2008),
Zhou et al. (2014)
Secondary NAC master switches regulating
secondary wall biosynthesis in fibers
(NST1, NST2 and SND1) and anther
endothecium (NST1 and NST2)
Mitsuda et al. (2005), Zhong et al. (2006,
2007b), Mitsuda et al. (2007, 2008), Zhong
and Ye (2014)
Secondary wall-associated NACs required for
normal secondary wall biosynthesis
Zhong et al. (2008)
Secondary wall MYB master switches activating secondary wall biosynthesis
Zhong et al. (2007a), McCarthy et al. (2009)
Secondary wall-associated MYBs induced by
SND1, NSTs and VNDs; involved in regulating secondary wall biosynthesis
Zhong et al. (2008)
Lignin-specific transcriptional activators
Zhong et al. (2008), J. Zhou et al. (2009)
Transcriptional regulation
VND1
At2g18060
200
VND2
At4g36160
VND3
At5g66300
VND4
At1g12260
VND5
At1g62700
VND6
At5g62380
VND7
At1g71930
NST1
At2g46770
NST2
At3g61910
SND1
At1g32770
SND2
At4g28500
SND3
At1g28470
MYB46
At5g12870
MYB83
At3g08500
MYB20
At1g66230
MYB42
At4g12350
MYB43
At5g16600
MYB52
At1g17950
MYB54
At1g73410
MYB69
At4g33450
MYB103
At1g63910
MYB58
At1g16490
MYB63
At1g79180
MYB85
At4g22680
KNAT7
At1g62990
A homeodomain transcription factor proposed to repress secondary wall
biosynthesis
E. Li et al. (2012)
LBD18
At2g45420
Soyano et al. (2008)
LBD30
At4g00220
LOB domain proteins involved in a positive
feedback loop to regulate VND7
expression
XND1
At5g64530
A NAC domain protein shown to repress
secondary wall biosynthesis and programmed cell death in vessels
Zhao et al. (2008)
MYB4
At4g38620
MYB7
At2g16720
Sonbol et al. (2009), Fornale et al. (2010),
Legay et al. (2010), Shen et al. (2012)
MYB32
At4g34990
Induced by MYB46 and their orthologs in
other species known to be transcriptional
repressors of lignin biosynthesis
(continued)
Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140
Table 1 Continued
Gene
AGI code
Functions
References
BLH6
At4g34610
A BEL1-like homeodomain transcription
factor involved in regulating lignin
biosynthesis
Cassan-Wang et al. (2013)
MYB26
At3g13890
A MYB specifically regulating secondary wall
biosynthesis in anther endothecium
Yang et al. (2007)
Others genes important for secondary wall deposition
IFL1/REV
At5g60690
A homeodomain leucine-zipper protein essential for fiber differentiation and secondary wall thickening
Zhong and Ye (1999)
WAT1
At1g75500
A vacuolar auxin transporter essential for
fiber differentiation and secondary wall
thickening
Ranocha et al. (2013)
EXO70A1
At5g03540
A subunit of exocyst complex proposed to
function in vesicle trafficking in vessels
Li et al. (2013)
FRA3
At1g65580
At1g22620
Phosphoinositide phosphatases essential for
actin organization and secondary wall
deposition
Zhong et al. (2004, 2005a)
AtSAC1/FRA7
RHD3
At3g13870
A GTPase essential for actin organization
and secondary wall deposition
Hu et al. (2003)
suggesting their potential roles in cellulose biosynthesis (Song
et al. 2010).
Xylan Biosynthesis
Xylan, a major hemicellulosic component in secondary walls of
angiosperm species, is a polymer consisting of a backbone of b1,4-linked xylosyl residues with the degree of polymerization
around 100 (Fig. 1D). The xylosyl residues are often substituted
with monosaccharide residues, including a-1,2-linked glucuronic acid (GlcA), a-1,2-linked 4-O-methylglucuronic acid
(MeGlcA), and a-1,2 and/or a-1,3-linked arabinose (Ara)
(Ebringerova and Heinze 2000). Although monosaccharide substituents are dominant, xylans from grasses may contain disaccharide side chains composed of a-1,3-linked Ara substituted at
O-2 with Ara or xylose (Xyl) (Ebringerova and Heinze 2000).
Xylans from Eucalyptus globulus and Arabidopsis also contain
disaccharide side chains composed of a-1,2-linked GlcA/
MeGlcA substituted at O-2 with galactose (Shatalov et al.
1999, Zhong et al. 2014). Based on the nature of their substitutions, xylans are named glucuronoxylan, arabinoxylan and glucuronoarabinoxylan. Glucuronoxylan is the dominant form of
xylan in dicot secondary walls, and glucuronoarabinoxylan is
the dominant form in the wood of gymnosperms and cell walls
of grass species. In addition to the sugar substitutions, xylans
may be acetylated at C-2 and/or C-3, with the degree of acetylation varying from 40% to 70% (Teleman et al. 2002, Evtuguin
et al. 2003, Goncalves et al. 2008). In grasses, Ara substituents of
xylan may be covalently cross-linked with lignin via ferulic acid
residues (Ebringerova and Heinze 2000). Xylans from gymnosperms and dicots contain a distinct tetrasaccharide sequence,
b-D-Xyl-(1!3)-a-L-Rha-(1!2)-a-D-GalA-(1!4)-D-Xyl, at its
reducing end (Shimizu et al. 1976, Johansson and Samuelson
1977, Andersson et al. 1983, Pena et al. 2007, Lee et al. 2009a).
The biosynthesis of xylan requires a number of enzymes,
including xylosyltransferases, glucuronyltransferases, methyltransferases, acetyltransferases and arabinosyltransferases, that
are responsible for xylan backbone elongation and substitutions (Table 1). Genetic and biochemical analysis in
Arabidopsis has uncovered the essential roles of glycosyltransferases from GT43 and GT47 families in xylan backbone elongation. There exist four GT43 genes in the Arabidopsis genome,
IRX9, I9H/IRX9-L, IRX14 and I14H/IRX14-L, all of which are expressed in secondary wall-forming cells, including vessels and
fibers, and their encoded proteins are located in the Golgi
(Brown et al. 2007, Pena et al. 2007, Lee et al. 2010, Wu et al.
2010). Mutation of IRX9 or IRX14 alone is sufficient to cause a
drastic reduction in xylan xylosyltransferase activity, xylan chain
length and xylan content (Brown et al. 2007, Lee et al. 2007a,
Pena et al. 2007). As a result, the secondary wall thickness in
fibers and vessels of the mutants is severely reduced and the
vessels are collapsed due to their inability to resist the negative
pressure generated during transpiration. Genetic complementation studies have revealed that the four GT43 genes form two
functionally non-redundant groups, IRX9 and I9H being in one
and IRX14 and I14H being in another, both of which are
required for xylan backbone elongation (Lee et al. 2010). Coexpression of IRX9 and IRX14 in tobacco BY2 cells leads to a
significant increase in xylosyltransferase activity that is able to
add xylosyl residues successively from the UDP-Xyl donor onto
the xylooligomer acceptor (Lee et al. 2012b). GT43 genes from
Populus and rice also form two functionally non-redundant
groups (Lee et al. 2012c, Lee et al. 2014), indicating that the
recruitment of two functionally non-redundant groups of GT43
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genes in xylan backbone elongation is conserved in vascular
plants. Mutations of two GT47 genes, IRX10 and IRX10-L,
cause similar phenotypes to the mutation of IRX9 or IRX14,
such as a reduction in xylan xylosyltransferase activity, xylan
chain length, xylan content and secondary wall thickness
(Brown et al. 2009, Wu et al. 2009). Activity assays of recombinant IRX10 and IRX10L proteins and their close
homologs from moss and psyllium revealed that they exhibit xylan xylosyltransferase activity that is capable of
adding xylosyl residues onto the xylooligomer acceptor
(Jensen et al. 2014, Urbanowicz et al. 2014). In the developing mucilaginous tissues of psyllium where xylan is predominant, IRX10 homologs are highly expressed but GT43
homologs are not, suggesting the possibility that IRX10 proteins may function without GT43 proteins in xylan backbone elongation in the mucilaginous tissues of psyllium
(Jensen et al. 2013). Although IRX9, IRX14 and IRX10 are
all required for the xylan xylosyltransferase activity, IRX9
and IRX14 are proposed to play a structural instead of a
catalytic role in xylan backbone biosynthesis (Ren et al.
2014).
The addition of GlcA side chains in Arabidopsis xylan is
mediated by three glucuronyltransferase genes (GUX1, GUX2
and GUX3) that belong to family GT8 (Mortimer et al. 2010,
Lee et al. 2012a). These GUX genes are expressed in secondary
wall-forming cells and their encoded proteins are targeted to
the Golgi. Mutation of a single GUX gene causes a partial reduction in the amount of GlcA, but no effects on plant growth
and secondary wall thickness. In contrast, simultaneous mutations of all three GUX genes lead to a complete loss of GlcA side
chains in xylan, a reduction in secondary wall thickness and a
deformation of vessels (Lee et al. 2012a). The finding that GUXs
exhibit xylan glucuronyltransferase activities when heterologously expressed in tobacco cells (Lee et al. 2012a, Rennie
et al. 2012) further supports that GUX1/2/3 are glucuronyltransferases responsible for GlcA substitution of xylan during
secondary wall biosynthesis.
About 60% of the GlcA side chains in Arabidopsis xylan are
methylated at O-4, and it has been shown that the GlcA methylation of xylan is catalyzed by three Golgi-localized glucuronoxylan methyltransferases (GXM1, GXM2 and GXM3/GXMT1)
that belong to a family of DUF579 domain-containing proteins
(Lee et al. 2012d, Urbanowicz et al. 2012). The three GXM
genes are expressed in secondary wall-forming cells and their
recombinant proteins exhibit methyltransferase activity that is
capable of transferring the methyl group from S-adenosylmethionine to GlcA-substituted xylooligomers. Although mutation of individual GXM genes causes a partial reduction in the
degree of GlcA methylation, simultaneous mutations of all
three GXM genes lead to a complete loss of GlcA methylation
in xylan (Yuan et al. 2014a). No reduction in xylan content and
secondary wall thickening was detected in the gxm1/2/3 triple
mutant, indicating that loss of GlcA methylation has no negative impact on xylan biosynthesis and secondary wall assembly.
The GlcA side chains in Populus wood xylan are completely
methylated, which has been shown to be mediated by four
wood-associated GXM homologs, PtrGXM1/2/3/4 (Yuan et al.
202
2014b). Recombinant proteins of PtrGXM3 and PtrGXM4 exhibit 10 times lower Km values than GXM3/GXMT1 that is
responsible for the bulk of GlcA methylation of Arabidopsis
xylan, indicating that PtrGXM3 and PtrGXM4 have a much
higher substrate affinity than Arabidopsis GXM, which may
enable a complete methylation of GlcA side chains in Populus
xylan during wood formation. In Arabidopsis, the GXM activity
could be rate limiting so that it may not be able to catch up
with the rate of xylan biosynthesis to methylate all the GlcA
side chains during secondary wall biosynthesis. This hypothesis
is supported by overexpression analysis showing that an increase in GXM activity is sufficient to methylate all GlcA side
chains in Arabidopsis xylan (Yuan et al. 2014a). Two additional
DUF579 domain-containing proteins, IRX15 and IRX15L, are
involved in xylan biosynthesis, although their exact functions
remain to be investigated (Brown et al. 2011, Jensen et al. 2011).
The addition of Ara side chains in grass xylan has been
shown to be mediated by glycosyltransferases from family
GT61 (Anders et al. 2012). RNA interference inhibition of a
wheat GT61 gene, TaXAT1, causes a specific decrease in
a-1,3-linked Ara substitution of xylan in wheat endosperm.
Heterologous expression of TaXAT2 and its rice close homologs, OsXAT2 and OsXAT3, in Arabidopsis results in a-1,3linked Ara substitution of xylan, which normally does not
occur in Arabidopsis. These findings provide genetic evidence
demonstrating that XATs are probably arabinosyltransferases
that catalyze the a-1,3-linked Ara substitution of grass xylan.
Another GT61 gene, XAX1, is proposed to mediate the transfer
of b-1,2-linked Xyl onto the Ara side chains that are attached to
the xylan backbone at O-3 (Chiniquy et al. 2012).
O-Acetylation of xylan involves the RWA genes and the
ESK1/TBL29 gene. The Arabidopsis genome contains four
RWA genes, all which are expressed in secondary wall-forming
cells. Simultaneous mutations of the four RWA genes result in a
40% reduction in xylan acetylation, reduced secondary wall
thickening and a collapsed vessel morphology, indicating
their roles in xylan acetylation (Lee et al. 2011). RWAs are
close homologs of fungal CAS1 that is involved in acetylation
of glucuronoxylomannan, a main capsule polysaccharide
(Janbon et al. 2001). Because RWAs contain only multiple transmembrane domains but lack the putative acetyltransferase
domain found in CAS1, they are proposed to be putative transporters for channeling the acetyl donor, acetyl-CoA, from the
cytoplasm to the Golgi lumen (Gille and Pauly 2012). ESK1
belongs to a large, plant-specific DUF231 domain protein
family, several other members of which have been shown to
be involved in O-acetylation of plant cell wall polysaccharides
(Gille and Pauly 2012). ESK1 is specifically expressed in secondary wall-forming cells and its encoded protein is located in the
Golgi (Yuan et al. 2013). The esk1 mutation causes a reduction
in secondary wall thickening and a deformation of vessel
morphology, but no decrease in xylan content. Analysis of
xylan acetylation and acetyltransferase activity revealed that
the esk1 mutant had a specific reduction in 2-O- and 3-Omonoacetylation of xylan and a drastic decrease in xylan acetyltransferase activity compared with the wild type, indicating
that ESK1 is a putative acetyltransferase mediating 2-O- and
Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140
3-O-monoactylation of xylan (Xiong et al. 2013, Yuan et al.
2013). Activity assay of recombinant ESK1 confirmed that it
exhibits acetyltransferase activity catalyzing 2-O- and 3-Omonoacetylation of xylan (Urbanowicz et al. 2014).
A number of genes involved in the biosynthesis of xylan
reducing end tetrasaccharide sequence have been identified
(Zhong et al. 2005b, Brown et al. 2007, Lee et al. 2007b, Pena
et al. 2007, Persson et al. 2007, Lee et al. 2009b). These genes
include FRA8, F8H, IRX8 and PARVUS, all of which are expressed
in secondary wall-forming cells, and their encoded proteins are
located in the Golgi, except for PARVUS that is predominantly
located in the endoplasmic reticulum. FRA8 and it close homolog, F8H belong to family GT47, and IRX8 and PARVUS belong
to family GT8. Mutation of FRA8, IRX8 or PARVUS alone causes
a reduction in xylan content and secondary wall thickness, and
a loss of the xylan reducing end tetrasaccharide sequence, b-DXyl-(1!3)-a-L-Rha-(1!2)-a-D-GalA-(1!4)-D-Xyl
(Brown
et al. 2007, Lee et al. 2007b, Pena et al. 2007, Lee et al. 2009b).
F8H is a functional paralog of FRA8, and simultaneous mutations of FRA8 and F8H result in a severe deformation of vessels
and an extreme retardation in plant growth (Lee et al. 2009b).
No reduction in xylan xylosyltransferase or glucuronyltransferase activities occurs in these mutants (Lee et al. 2007a). The
available evidence indicates that these glycosyltransferases mediate the biosynthesis of the xylan reducing end sequence.
Based on their catalytic mechanisms, it was proposed that
FRA8 might catalyze the transfer of Xyl to Rha or of Rha to
GalA, and IRX8 might mediate the transfer of GalA to the
reducing Xyl residue at the reducing end of xylan (Pena et al.
2007). PARVUS might be involved in the initiation of the reducing end sequence biosynthesis by transferring the reducing
end Xyl residue to an unknown acceptor (Lee et al. 2007b).
Glucomannan Biosynthesis
Glucomannan, a minor hemicellulosic component of dicot
wood, consists of a linear chain of b-1,4-linked mannosyl residues interspersed with glucosyl residues with a ratio of 2 : 1
(Timell 1967) (Fig. 1D). Galactoglucomannan, a major hemicellulosic component of gymnosperm wood, is composed of
the glucomannan backbone substituted with a-1,6-linked
galactosyl residues. Glucomannan from aspen and birch
wood has been shown to be O-acetylated at C-2 or C-3 of
mannosyl residues, with a degree of acetylation of about 0.3
(Teleman et al. 2003). The biosynthesis of glucomannan is catalyzed by members of the cellulose synthase-like A (CslA)
family, which belong to family GT2 (Dhugga et al. 2004,
Liepman et al. 2005). Recombinant Arabidopsis CslA proteins
are able to catalyze the transfer of both mannosyl and glucosyl
residues from GDP-mannose and GDP-glucose, respectively, to
produce b-linked glucomannan polymers, demonstrating that
CslA family members are glucomannan synthases. There are
nine CslA genes in the Arabidopsis genome, and three of
them, CslA2, CslA3 and CslA9, are responsible for the synthesis
of glucomannan in secondary wall-containing cells of stems
(Goubet et al. 2009) (Table 1). CslA genes are present in all
taxa of land plants, and those from tree species, including pine
and Populus, have been demonstrated to exhibit glucomannan
synthase activities (Suzuki et al. 2006, Liepman et al. 2007). The
a-1,6-linked galactosyl substitution in galactoglucomannan is
catalyzed by UDP-galactose-dependent a-1,6-D-glucomannan
galactosyltransferase, whose corresponding gene was identified
in fenugreek (Edwards et al. 1999).
Lignin Biosynthesis
Lignin is a complex three-dimensional polyphenolic polymer
that is generated through oxidative polymerization in muro
of three monolignols, p-coumaryl, coniferyl and sinapyl alcohols
(Fig. 1D). The lignin polymer is divided into three types,
p-hydroxyphenyl (H) lignin that is polymerized from p-coumaryl alcohol, guaiacyl (G) lignin polymerized from coniferyl
alcohol, and syringyl (S) lignin polymerized from sinapyl alcohol. Lignin composition, i.e. the proportion of each of three
polymerized monolignols, may differ significantly among different plant species and even among different cell types in the
same species. For example, secondary walls in gymnosperms
contain guaiacyl lignin but lack syringyl lignin, whereas those
from dicot species are typically rich in both guaiacyl lignin and
syringyl lignin. In secondary walls of grass species, a significant
proportion of p-hydroxyphenyl lignin is present in addition to
guaiacyl lignin and syringyl lignin (Bonawitz and Chapple 2010).
Monolignol biosynthesis is carried out through the common
phenylpropanoid pathway starting from phenylalanine to produce hydroxycinnamoyl-CoA esters. These esters are then
routed into the lignin branch pathway to generate monolignols.
At least 10 enzymes are involved in catalyzing monolignol biosynthesis; they are phenylalanine ammonia lyase (PAL), cinnamate 4-hydroxylase (C4H), 4-coumarate CoA ligase (4CL),
hydroxycinnamoyl CoA:shikimate hydroxycinnamoyl transferase (HCT), p-coumaroyl shikimate 30 -hydroxylase (C3H), caffeoyl CoA O-methyltransferase (CCoAOMT), ferulate 5hydroxylase (F5H), caffeic acid O-methyltransferase (COMT),
cinnamoyl CoA reductase (CCR) and cinnamyl alcohol dehydrogenase (CAD) (Bonawitz and Chapple 2010). It has recently been demonstrated that caffeoyl shikimate esterase
(CSE) that catalyzes the conversion of caffeoyl shikimate into
caffeic acid is another enzyme in the lignin biosynthetic pathway (Vanholme et al. 2013). Genes encoding these monolignol
biosynthetic enzymes have been identified and functionally
characterized in a number of species (Bonawitz and Chapple
2010). In Arabidopsis, 35 monolignol biosynthetic genes have
been identified, and the expression of 16 of them, i.e. PAL1,
PAL2, PAL4, C4H, 4CL1, 4CL2, HCT, C3H1, CSE, CCoAOMT1,
CCoAOMT7, CCR1, F5H, COMT, CAD3 and CAD6, is closely
associated with the development of lignified xylem and interfascicular fibers in stems, indicating that they are the core
monolignol biosynthetic genes responsible for lignin synthesis
during secondary wall formation in stems (Raes et al. 2003,
Vanholme et al. 2013) (Table 1). Similarly, expression analysis
identified 18 monolignol biosynthetic genes that are abundant
and specific in differentiating xylem in Populus and are likely to
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be responsible for lignin synthesis during wood formation (Shi
et al. 2010). Mutants with mutation of each of the monolignol
biosynthetic genes in Arabidopsis have been generated and
analyzed by transcriptomics and metabolomics. Although not
all of the mutants have reduced lignin content and altered
lignin composition, the expression of many genes and the
level of various metabolites are altered in these mutants
(Vanholme et al. 2012).
Monolignols are synthesized in the cytosol and then transported into cell walls, where they are polymerized into lignin via
oxidative reactions catalyzed by oxidases, including laccases and
peroxidases (Bonawitz and Chapple 2010). The transport of
monolignols into cell walls is proposed to be mediated by
plasma membrane-localized transporters (Kaneda et al. 2008,
Liu et al. 2011). Using isolated Arabidopsis plasma membrane
vesicles, it has been demonstrated that plasma membranes
contain ATP-binding cassette-like transporter activities that
are able to transport monolignols (Miao and Liu. 2010).
Heterologous expression of one of the Arabidopsis ATP-binding
cassette-like transporter genes, AtABCG29, in yeast cells results
in an increased activity of p-coumaryl alcohol transport, suggesting that AtABCG29 is involved in the transport of p-coumaryl alcohol (Alejandro et al. 2012). Transporters responsible
for the transport of coniferyl and sinapyl alcohols remain to be
identified. Several genes encoding laccases and peroxidases
have been shown to be involved in lignin polymerization
during secondary wall biosynthesis in Arabidopsis and
Populus (Table 1). Simultaneous mutation of Laccase 4
(LAC4) and LAC17 results in a reduction in lignin content in
Arabidopsis stems (Berthet et al. 2011) and simultaneous mutation of LAC4 and LAC17 together with LAC11 leads to plant
growth arrest and a loss of lignification in root vessels (Zhao
et al. 2013). Down-regulation of laccase genes in Populus by
overexpression of a microRNA that targets a number of laccase
genes leads to a reduction in laccase activity and lignin content
in transgenic wood (Lu et al. 2013). Several Arabidopsis peroxidase genes have been implicated in lignin polymerization
(Herrero et al., 2013a); mutation of AtPRX72 causes a reduction
in lignin content (Herrero et al. 2013b) and mutations of
AtPRX2, AtPRX25 and AtPRX71 result in reduced lignin content
and altered lignin structure in secondary walls (Shigeto et al.
2013, Shigeto et al. 2014).
Mutants with a reduction in lignin content often lead to
impaired secondary wall deposition, deformed vessels and
reduced plant growth (Zhong et al. 1998, Jones et al. 2001,
Ruel et al. 2009), which may hinder the effort of generating
low-lignin crops for biofuel production. The recent finding
that the reduced growth and abnormal secondary wall deposition seen in the ref8 mutant, which is due to a mutation in the
C3H1 gene, could be partially rescued by mutations of two
genes encoding transcriptional co-regulatory complex subunits
MED5a and MED5b may provide a new tool to overcome the
plant growth defect caused by a reduced lignin content
(Bonawitz et al. 2014). An alteration in lignin structure without
a change in lignin content and plant growth has been achieved
by introduction into Populus of a monolignol ferulate transferase that generates monolignol ferulate conjugates (Wilkerson
204
et al. 2014). Lignin in transgenic Populus wood with incorporation of ester-linked monolignol ferulates is easier to remove,
which represents a novel approach to engineer biofuel crops
with easy removal of lignin.
Patterned Deposition of Secondary Walls
Secondary walls in tracheary elements and fibers of vascular
plants are deposited in elaborate patterns. Tracheary elements
(protoxylem) in elongating organs typically have secondary
walls deposited in a helical or annular pattern, which provides
the necessary mechanical support to the xylem conduits and at
the same time allows organs to continue to elongate. Tracheary
elements (metaxylem) in non-elongating organs typically have
secondary walls organized in a reticulated or pitted pattern
(Fig. 2A), which provides maximum mechanical support to
the xylem conduits and the plant body. Fiber cells generally
have secondary walls deposited uniformly between the primary
wall and the plasma membrane, except for the pit areas that are
used for solute transport (Mauseth, 1988) (Fig. 2B). Secondary
walls typically contain three distinct layers, called S1, S2 and S3,
when observed using transmission electron microscopy
(Fig. 1B). The layered deposition of secondary walls is caused
by changes in the orientations of cellulose microfibrils. Both the
S1 layer and the S3 layer are typically thin, and the cellulose
microfibrils in these two layers are oriented in a crossed organization and a flat helix relative to the cell elongation axis, respectively. The S2 layer is the thickest, with cellulose microfibrils
oriented almost parallel to the cell elongation axis (Fig. 1C), and
the angles of these microfibrils largely determine the mechanical strength of fibers in wood (Timell 1967). The number of
layers in secondary walls may change in response to environmental stimuli, and secondary walls in some plant species may
have more than three layers. For example, secondary walls in
compression wood lack the S3 layer but contain an extra layer
of lignin located between S1 and S2, whereas those in tension
wood may lack one or two of the three typical layers but often
have an additional cellulose-rich gelatinous layer next to the
lumen (Timell et al. 1967). In addition, up to six distinct layers
are observed in the secondary walls of bamboo fibers (Gritsch
and Murphy 2005).
It has long been known that cortical microtubules play a
crucial role in both the oriented deposition of cellulose microfibrils and the patterned deposition of secondary walls. At the
subcellular level, the orientation of cortical microtubules coaligns with that of cellulose microfibrils, and disruption of the
cortical microtubule organization by pharmacological drugs results in an altered orientation of cellulose microfibrils, which led
to the alignment hypothesis that cortical microtubules guide
the oriented deposition of cellulose microfibrils (Baskin 2001).
This hypothesis is supported by the study of fluorescent protein-tagged CesAs showing that CesA complexes move along
the track of cortical microtubules located underneath the
plasma membrane, and disruption of the cortical microtubule
organization alters the movement of CesA complexes (Paradez
et al. 2006). Primary wall CesAs have been found to interact
Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140
Fig. 2 Patterned deposition of secondary walls. (A) Ectopic deposition of secondary walls in a reticulated pattern in Arabidopsis
leaf epidermal cells overexpressing SND1. Note the gap areas between thickened walls. (B) Transmission electron micrograph
showing a pair of bordered pits in secondary walls of interfascicular fibers from an Arabidopsis stem. Scale bar = 2.6 mm. (C)
Diagram of regulation of patterned deposition of secondary
walls. Cortical microtubules are bundled beneath the plasma
membrane where secondary wall thickening occurs. Bundling of
cortical microtubules and delimitation of the borders of bundled
cortical microtubules are facilitated by microtubule-associated
proteins (MAP65 and MAP70). Cortical microtubules, actin cytoskeleton and the exocyst complex (including a subunit EXO70A1)
are involved in targeted transport of CesA complexes and vesicles
carrying secondary wall components. Formation of the pit area
with no secondary wall thickening is controlled by the ROP11/
MIDD1/Kinesin-13A complex that mediates local depolymerization
of cortical microtubules.
with cortical microtubules via cellulose synthase-interacting
protein 1 (CSI1) (Gu et al. 2010) (Table 1). CSI1 directly interacts with microtubules and the cytoplasmic domain of CesAs,
and csi1 mutation affects the distribution and the movement of
CesA complexes. CSI1 is suggested to function as a scaffold
protein to guide the movement of CesA complexes along the
track of cortical microtubules (Bringmann et al. 2012, S. Li et al.
2012). Although the functional roles of CSI1 in secondary wall
cellulose biosynthesis have not been investigated, it is likely that
CSI1 or CSI homologs may also guide the movement of secondary wall CesA complexes along cortical microtubules, and
thereby control the oriented deposition of cellulose microfibrils
in secondary walls.
Since the orientation of cortical microtubules is correlated
with the direction of CesA movement and the subsequent
orientation of cellulose microfibrils, it is likely that the different
orientations of cellulose microfibrils seen in the S1, S2 and S3
layers of secondary walls reflect the dynamic re-organization of
cortical microtubules during the transition of formation of different secondary wall layers. An important role for cortical
microtubules in controlling the oriented deposition of cellulose
microfibrils during secondary wall biosynthesis is exemplified by
mutation of the AtKTN1 gene in the fra2 mutant (Burk et al.
2007) (Table 1). AtKTN1 encodes a katanin microtubule-severing protein, and its mutation in the fra2 mutant causes a disorganization of cortical microtubules, an aberrant orientation
of cellulose microfibrils and a loss of layers in the secondary
walls of fibers. These findings provide genetic evidence supporting a role for cortical microtubules in directing the formation of
layered secondary walls (Burk et al. 2001, Burk and Ye. 2002). It
has been shown that the kinesin motor proteins, Arabidopsis
FRA1 and its rice ortholog BC12, also play a significant role in
the oriented deposition of cellulose microfibrils and the mechanical strength of secondary walls (Zhong et al. 2002, Zhang
et al. 2010) (Table 1). Mutation of FRA1 or BC12 has no effect
on cortical microtubule organization and cellulose content, but
it drastically reduces the mechanical strength of fibers, probably
due to the disorganization of cellulose microfibrils (Zhong et al.
2002, Zhang et al. 2010). FRA1 overexpression results in a reduction in cellulose content and secondary wall thickness in
fibers, a collapsed vessel phenotype and an increase in the
number of layers in secondary walls (Zhou et al. 2007). FRA1
and BC12 belong to the kinesin-4 subfamily, and kinesin-4 proteins in animal cells are implicated in regulating the movement
of multiple intracellular components, such as vesicle transport
(Martinez et al. 2008). It is likely that FRA1 and BC12 are
involved in the transport of vesicles carrying cell wall materials
along microtubules and thereby influence cellulose microfibril
deposition and secondary wall assembly.
It has been demonstrated that cortical microtubules in
tracheary elements form bundles beneath specific plasma
membrane domains where secondary wall thickening occurs
(Oda et al. 2005). Disruption of the cortical microtubule organization by pharmacological drugs alters the patterned deposition of secondary walls in tracheary elements, suggesting that
cortical microtubules determine the patterned deposition of
secondary walls. It has been shown that CesA complexes
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responsible for cellulose microfibril deposition and laccases
involved in monolignol polymerization are concentrated as
bands at the plasma membrane domains where secondary
wall thickening occurs, and the underlying cortical microtubule
bundles are essential for CesA localization (Wightman and
Turner 2008, Schuetz et al. 2014). It is evident that the cortical
microtubule bundles not only control the trajectory of CesA
movement in the plasma membrane but also direct the targeted transport of vesicles carrying the CesA complexes, hemicelluloses and monolignol polymerization enzymes to specific
plasma membrane domains, and thereby control the patterned
deposition of secondary walls. The bundling of cortical microtubules is mediated by MAP65 microtubule-associated proteins. ZeMAP65-1 has been shown to co-localize with cortical
microtubule bundles in the Zinnia tracheary elements and,
when overexpressed in Arabidopsis suspension cells, it induces
the normally evenly distributed cortical microtubules to form
thick helical or transverse bundles that resemble cortical microtubule bundles seen in the tracheary elements (Mao et al. 2006).
In addition, two plant-specific MAPs, the Arabidopsis MAP70-5
and MAP70-1, are proposed to play an essential role in defining
the locations of secondary wall thickening through regulating
the organization of cortical microtubule bundles (Pesquet et al.
2010). It has been shown that MAP70-5 lines the borders of
each cortical microtubule bundle. MAP70-5 expression is
induced during tracheary element differentiation in
Arabidopsis suspension cells, and manipulating the level of
MAP70-5 or MAP70-1 by their down-regulation or overexpression leads to a loss of patterned secondary wall deposition or an
increase in the population of tracheary elements with the spiral
secondary wall pattern, respectively (Pesquet et al. 2010). These
findings indicate that MAP65 and MAP70 proteins play
crucial roles in bundling cortical microtubules and defining
the borders of cortical microtubule bundles, respectively, and
thereby regulate the pattered secondary wall deposition
(Table 1; Fig. 2C).
Cortical microtubule bundles are separated by distinctive
gaps that are low in microtubules. It was proposed that the
maintenance of low levels of microtubules in the gap areas is
mediated by the ROP11/MIDD1/Kinesin-13A complexes, which
are located underneath the microtubule-depleting plasma
membrane domains that coincide with regions where no secondary wall thickening occurs (Oda and Fukuda 2013b)
(Table 1; Fig. 2C). In these complexes, the microtubule depletion domain 1 protein, MIDD1, directly interacts with microtubules (Oda et al. 2010) and acts as a scaffold protein to recruit
the microtubule depolymerization protein, Kinesin-13A, to depolymerize cortical microtubules locally, leading to formation
of microtubule-deleting plasma membrane domains (Oda and
Fukuda 2013a). RNA interference inhibition of MIDD1 results in
a failure of localized depolymerization of cortical microtubules
and concomitantly a uniform deposition of secondary walls
(Oda et al. 2010). Similarly, RNA interference inhibition of
Kinesin-13A causes formation of secondary walls with smaller
pit areas. In contrast, overexpression of Kinesin-13A leads to
increased depolymerization of cortical microtubules and formation of secondary walls with larger pit areas (Oda and
206
Fukuda 2013a). The recruitment of MIDD1/Kinesin-13A to
the microtubule-depleting plasma membrane domains is
mediated by the active form of ROP11 GTPase. The inactivation
and activation of ROP11 are controlled by a ROP GTPase-activating protein (ROPGAP3) and a ROP guanine nucleotide exchange factor (ROPGEF4), respectively, both of which have
been shown to be localized at the microtubule-depleting
plasma membrane domains (Oda and Fukuda 2012). These
findings suggest that the spacing between cortical microtubule
bundles is delimited by the microtubule-depleting plasma
membrane domains containing the ROP11/MIDD1/Kinesin13A complexes, which control the patterns of cortical microtubule bundles and the subsequent patterns of secondary wall
deposition (Fig. 2C).
Transcriptional Regulation of Secondary Wall
Biosynthesis and Deposition
Because secondary walls are deposited after cessation of cell
elongation in secondary wall-forming cells, genes encoding
the suite of catalytic enzymes required to produce secondary
wall polymers need to be co-ordinately turned on temporaly
and spatially. This co-ordinate activation of secondary wall biosynthetic genes is controlled by a transcriptional network encompassing secondary wall NAC and MYB master switches and
their downstream transcription factors (Zhong et al. 2010a)
(Table 1; Fig. 3). In this transcriptional network, secondary
wall NAC master switches (SWNs) are the top-level master
switches that are capable of turning on the entire secondary
wall biosynthetic program. In Arabidopsis, 10 SWNs (VND1–
VND7, SND1, NST1 and NST2), which belong to the same subgroup of the NAC family, regulate secondary wall biosynthesis
in various secondary wall-forming cell types. VND1–VND7 are
specifically expressed in vessels, and their dominant repression
causes a loss of secondary wall thickening in vessels (Kubo et al.
2005, Yamaguchi et al. 2008, Zhou et al. 2014). SND1, NST1 and
NST2 are expressed specifically in xylary fibers and extraxylary
fibers in stems, roots and siliques. Simultaneous mutations of
SND1, NST1 and NST2 cause a complete loss of secondary wall
thickening in fibers (Zhong et al. 2006, Mitsuda et al. 2007,
Zhong et al. 2007b, Mitsuda and Ohme-Takagi 2008, Zhong
and Ye 2014). NST1 and NST2 are also expressed in anther
endothecium cells, and simultaneous mutations of NST1 and
NST2 led to a loss of secondary wall thickening in endothecium
cells and an anther indehiscence phenotype (Mitsuda et al.
2005). Overexpression of any of the SWNs induces the expression of the biosynthetic genes for cellulose, xylan and lignin, and
concomitantly results in ectopic deposition of secondary walls
in normally parenchymatous cells. Furthermore, expression of
any of the SWNs driven by the SND1 promoter in the snd1 nst1
double mutant rescues the secondary wall defect in fibers
(Zhong et al. 2010c, Yamaguchi et al. 2011, Zhou et al. 2014).
SWN homologs exist in all taxa of vascular plants, ranging
from seedless vascular plants to gymnosperms and angiosperms (Zhong et al. 2010a). Functional characterization
of SWN homologs from Populus, Eucalyptus, maize, rice,
Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140
Brachypodium, Medicago and spruce has revealed that similarly
to Arabidopsis SWNs, they are capable of activating secondary
wall biosynthetic genes leading to ectopic secondary wall deposition when overexpressed, causing a reduction in secondary
wall thickening when dominantly repressed, and/or restoring
secondary wall thickening in fibers of the snd1 nst1 double
mutant when expressed under the SND1 promoter (Zhong
et al. 2010a, Zhong et al. 2010b, Zhao et al. 2010, Zhong et al.
2011a, Zhong et al. 2011b, Valdivia et al. 2013, Yoshida et al.
2013). In Populus and rice, some of the SWN homologs have
alternative splicing forms, resulting in truncated SWNs without
the carboxyl activation domain that may serve as negative regulators of SWN functions (Q. Li et al. 2012, Zhao et al. 2014).
These findings indicate that vascular plants have evolved to
employ SWNs as master switches to regulate secondary wall
biosynthesis (Fig. 3).
SWNs have been shown to activate a battery of downstream
transcription factors, including SND2, SND3, MYB20, MYB42,
MYB43, MYB46, MYB52, MYB54, MYB58, MYB63, MYB69,
MYB83, MYB85, MYB103 and KNAT7 (Zhong et al. 2008)
(Table 1; Fig. 3). Among them, MYB46 and its close homolog,
MYB83, act as the second-level master switches regulating secondary wall biosynthesis (Zhong et al. 2007a, McCarthy et al.
2009). MYB46 and MYB83 are expressed in both vessels and
fibers, and their dominant repression or RNA interference inhibition results in a severe reduction in secondary wall thickening in fibers and vessels and a concomitant collapsed vessel
phenotype. Simultaneous mutations of MYB46 and MYB83
lead to a loss of secondary wall thickening in vessels and an
arrest of plant growth at the seedling stage (McCarthy et al.
2009). Similar to SWNs, MYB46 and MYB83 are also able to
induce the expression of the biosynthetic genes for cellulose,
xylan and lignin, thus causing ectopic deposition of secondary
walls (Zhong et al. 2007a, Ko et al., 2009, McCarthy et al. 2009).
MYB46/83 close homologs from gymnosperms and angiosperms have been demonstrated to be functional orthologs
of Arabidopsis MYB46/83 (Patzlaff et al. 2003, Goicoechea
et al. 2005, McCarthy et al. 2010, Zhong et al. 2010a, Zhong
et al. 2013), indicating that the recruitment of MYB46/83-like
transcription factors as second-level wall MYB master switches
is also evolutionarily conserved at least in gymnosperms and
angiosperms. Another MYB transcription factor, MYB26, may
function as a master switch regulating secondary wall biosynthesis in anther endothecium because its mutation causes a
loss of secondary wall thickening in anther endothecium
and an anther indehiscence phenotype, whereas its overexpression results in ectopic deposition of secondary walls. It is
suggested that MYB26 acts upstream of NST1 and NST2 in
regulating secondary wall biosynthesis in endothecium (Yang
et al. 2007).
A number of SWN- and MYB46/83-regulated downstream
transcription factors have been shown to play important roles
in regulating secondary wall biosynthesis (Table 1; Fig. 3).
Arabidopsis MYB58, MYB63, MYB85, BLH6 and their close
homologs in other species are specific activators of the lignin
biosynthetic pathway (Zhong et al. 2008, J. Zhou et al. 2009,
Cassan-Wang et al. 2013, Hirano et al. 2013b), LBD18 and
Fig. 3 Diagram of transcriptional regulation of the secondary wall
biosynthesis program in Arabidopsis. Secondary wall NAC and MYB
master switches bind to and activate the SNBE and SMRE sites, respectively, in the promoters of their targets, and, together with their
downstream transcription factors, activate the expression of genes
involved in secondary wall biosynthesis and deposition, leading to
secondary wall formation. VNDs also control the programmed cell
death in vessels.
LBD30 are involved in a positive feedback loop to regulate
VND7 expression (Soyano et al. 2008), XND1 is probably a
negative regulator of secondary wall biosynthesis and
programmed cell death (Zhao et al. 2008), KNAT7 is proposed
to be a negative regulator of secondary wall biosynthesis
(E. Li et al. 2012), and MYB4, MYB7, MYB32 and their close
homologs in other species are probably negative regulators of
lignin biosynthesis (Sonbol et al. 2009, Fornale et al. 2010,
Legay et al. 2010, Shen et al. 2012). Other transcription
factors, such as SND2, SND3, MYB52, MYB54, MYB69 and
MYB103, might be involved in fine-tuning the transcriptional
regulation of secondary wall biosynthesis because their
dominant repression causes a reduction in secondary wall
thickening but their overexpression does not result in
ectopic deposition of secondary wall components (Zhong
et al., 2008).
SWNs bind to a 19 bp imperfect palindromic consensus sequence, (T/A)NN(C/T)(T/C/G)TNNNNNNNA(A/C)GN(A/C/
T)(A/T), which is named secondary wall NAC-binding element
(SNBE), and thereby activate their target gene expression
(Zhong et al. 2010c). MYB46/MYB83 bind to a 7 bp secondary
wall MYB-responsive element (SMRE) that has a consensus
207
R. Zhong and Z.-H. Ye | Secondary cell walls
sequence of ACC(A/T)A(A/C)(T/C) (Zhong and Ye 2012).
The SMRE consensus sequence has eight variants, and three
of them are identical to the AC elements, which is congruent
with the early findings that pine and Eucalyptus MYB46 orthologs bind to the AC elements (Patzlaff et al. 2003, Goicoechea
et al. 2005). Another study showed that the MYB46-binding
element (namely M46R) is an 8 bp core motif, (T/C)ACC
(A/T)A(A/C)(T/C) (Kim et al. 2012a), which is identical to
SMRE except for having an additional nucleotide at the 50
end. SWN orthologs and MYB46/MYB83 orthologs from
other plant species also bind to SNBE and SMRE, respectively
(Zhong et al. 2011a, Zhong et al. 2011b, Valdivia et al. 2013,
Zhong et al. 2013, Yoshida et al. 2013), further demonstrating
the evolutionary conservation of SWNs and MYB46/MYB83like secondary wall MYBs in regulating secondary wall biosynthesis in vascular plants.
Direct target analyses have revealed that SWNs and
MYB46/MYB83 are able to activate directly the expression
of not only a suite of downstream transcription factors but
also a number of secondary wall biosynthetic genes (OhashiIto et al. 2010, Yamaguchi et al. 2011, Zhong and Ye 2012,
Zhong et al. 2010c, Kim et al. 2012b, Kim et al. 2014). In
addition, SWNs directly activate many genes involved in cell
wall modification and programmed cell death. These findings
provide important insights into the underlying mechanism of
the transcriptional network regulating secondary wall biosynthesis, suggesting that it does not utilize a simple linear
regulatory cascade; instead it employs a feed-forward loop
regulatory structure in which one transcription factor regulates another and they together regulate downstream targets
(Fig. 3). In this network, SWNs directly activate MYB46/
MYB83 and they together directly activate other transcription factors, such as KNAT7, and some biosynthetic genes for
xylan and lignin. Similarly, MYB46/MYB83 directly activate
MYB58/MYB63 and they together directly regulate the expression of lignin biosynthetic genes. This multileveled
feed-forward loop regulatory structure is adopted by plants
probably to ensure the efficient activation and regulation of
secondary wall biosynthetic genes during the development of
secondary wall-forming cells.
The ectopically deposited secondary walls induced by
overexpression of SWNs and secondary wall MYB master
switches form specific patterns similar to those of tracheary
elements (Kubo et al. 2005, Mitsuda et al. 2005, Zhong et al.
2006, Zhong et al. 2007a) (Fig. 2A), indicating that they also
regulate patterned deposition of secondary walls. Indeed, several genes known to be involved in cortical microtubule organization, oriented deposition of cellulose microfibrils and
patterned deposition of secondary walls, including MAP65-8
(ZeMAP65-1 ortholog), FRA1, MIDD1 and ROP11, are
induced by SWNs (Zhong et al. 2010c, Yamaguchi et al.
2011, Oda and Fukuda 2012). ROP11 is also a direct target
of MYB46 (Zhong and Ye 2012). It is apparent that SWNs
and secondary wall MYB master switches activate genes
involved in both biosynthesis and patterned deposition of
secondary walls. Furthermore, VNDs also regulate the expression of genes involved in programmed cell death in tracheary
208
elements (Ohashi-Ito et al. 2010, Zhong et al. 2010c,
Yamaguchi et al. 2011).
Concluding Remarks
Secondary wall formation is a complex process involving signaling that triggers the secondary wall biosynthetic program,
transcriptional activation of secondary wall biosynthetic genes,
biosynthesis of secondary wall components, targeted transport
of enzymes and secondary wall materials, and patterned deposition and assembly of secondary walls. Despite the tremendous
progress made toward identification and functional characterization of genes involved in secondary wall biosynthesis, a
number of aspects regarding secondary wall formation
remain to be investigated and a few of them are highlighted
below. First, secondary wall deposition is initiated in secondary
wall-forming cells after cessation of elongation, but signals that
trigger the secondary wall biosynthetic program have not been
well characterized. Identifying such signals will be of significance
in dissecting the molecular mechanisms underlying wood formation and have important implications in tree biotechnology.
Auxin, an important signal for initiation of differentiation of
secondary wall-forming cells, may serve as a signal for initiation
and/or maintenance of the secondary wall biosynthetic program since a reduction in auxin transport results in a defect
in secondary wall thickening in fibers (Zhong and Ye 1999,
Zhong and Ye 2001, Ranocha et al. 2013). Secondly, cortical
microtubules and the actin cytoskeleton are important for
trafficking of vesicles carrying CesA complexes (Wightman
and Turner 2008) and hemicelluloses to the plasma membrane
regions where secondary wall thickening occurs, but proteins
involved in vesicle trafficking for secondary wall formation are
largely unknown. The EXO70A1 gene, which encodes a subunit
of the exocyst complex, is proposed to function in vesicle trafficking during vessel differentiation (Li et al. 2013). The exo70a1
mutant has irregular secondary wall thickening, incomplete
perforation plates and large vesicular compartments in vessels.
Phosphoinositides, which are important for vesicle trafficking
and actin cytoskeleton organization in yeast and animals
(Takenawa and Itoh 2001), may also play crucial regulatory
roles in vesicle trafficking of secondary wall materials.
Mutations of two phosphoinositide phosphatase genes, FRA3
and AtSAC1 (FRA7), result in altered actin organization and
aberrant deposition of secondary walls (Zhong et al. 2004,
Zhong et al. 2005a). RHD3, a GTPase required for tubular endoplasmic reticulum generation and Golgi distribution (Chen
et al. 2011), is essential for normal actin organization and secondary wall deposition, probably also through regulating vesicle
trafficking of secondary wall materials (Hu et al. 2003). Further
deciphering the exocytosis machineries involved in the targeted
transport of secondary wall materials will help dissect the
mechanisms controlling the patterned deposition of secondary
walls. Thirdly, secondary wall composition varies among different plant species, in different cell types within the same species
and may change in response to developmental and environmental stimuli. With the increased understanding of genes
Plant Cell Physiol. 56(2): 195–214 (2015) doi:10.1093/pcp/pcu140
involved in the biosynthesis of secondary wall components and
its transcriptional regulation, tools are now available to investigate the molecular mechanisms controlling secondary wall
heterogeneity. Fourthly, secondary walls are deposited in specific patterns in xylem cells during different developmental
stages, e.g. an annular or helical pattern in protoxylem and a
reticulated or pitted pattern in metaxylem. Signals that trigger
the developmental changes of the patterned secondary wall
deposition and of the underlying cortical microtubule organization are yet to be identified. Similarly, little is known about
the molecular switches directing the oriented deposition of
cellulose microfibrils in the S1, S2 and S3 layers of secondary
walls. Finally, many additional secondary wall-associated genes
have been identified through genomic and co-expression analyses (Ko et al. 2009, Wilkins et al. 2009, Ohashi-Ito et al. 2010,
Yamaguchi et al. 2011, Zhong et al. 2010c, Zhong and Ye 2012,
Hirano et al. 2013a) and the functions of most of them are
unknown. Functional characterization of these genes may provide new insights into the molecular mechanisms of secondary
wall formation. Identification and functional characterization of
all players participating in the formation of secondary walls, the
major constituents of plant biomass, is not only important for
basic plant biology but also provides a knowledge foundation
for better utilizing plant biomass for various human needs.
Funding
The work in the authors’ lab was funded by the Division of
Chemical Sciences, Geosciences, and Biosciences, Office of
Basic Energy Sciences of the US.Department of Energy [DEFG02-03ER15415]; the US Department of Agriculture
National Institute of Food and Agriculture [AFRI Plant
Biology program (#2010-65116-20468)]; the National Science
Foundation [ISO-1051900].
Acknowledgments
We regret that many original articles on secondary wall biosynthesis could not be cited due to space limitation.
Disclosures
The authors have no conflicts of interest to declare.
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