Download 1 Laccases direct lignification in the discrete secondary cell wall

Plant Physiology Preview. Published on August 25, 2014, as DOI:10.1104/pp.114.245597
1
Laccases direct lignification in the discrete secondary cell wall domains of
protoxylem.
Mathias Schuetz,1 Anika Benske,1,2 Rebecca A. Smith,1,2 Yoichiro Watanabe,1 Yuki
Tobimatsu,3,5 John Ralph,3 Taku Demura,4 Brian Ellis,2 and Lacey Samuels1
1
Department of Botany, University of British Columbia, Vancouver, BC, Canada
2
Michael Smith Laboratories, University of British Columbia, Vancouver, BC,
Canada
3
Department of Biochemistry and the US Department of Energy’s Great Lakes
Bioenergy Research Center (GLBRC), the Wisconsin Energy Institute, University of
Wisconsin-Madison, Madison, WI, USA
4
Nara Institute of Science and Technology, Nara, Japan
5
Graduate School of Agriculture, Kyoto University, Kyoto, Japan
Running title: Laccases in Protoxylem Lignin Deposition
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
Copyright 2014 by the American Society of Plant Biologists
2
Abstract
Plants precisely control lignin deposition in spiral or annular secondary cell wall
domains during protoxylem tracheary element (TE) development. Because
protoxylem TEs function to transport water within rapidly elongating tissues, it is
important that lignin deposition is restricted to the secondary cell walls, in order to
preserve the plasticity of adjacent primary wall domains. The Arabidopsis inducibleVASCULAR NAC DOMAIN7 (VND7) protoxylem TE differentiation system permits
the use of mutant backgrounds, fluorescent protein tagging, and high-resolution livecell imaging of xylem cells during secondary cell wall development. Enzymes
synthesizing monolignols, as well as putative monolignol transporters, showed a
uniform distribution during protoxylem TE differentiation. In contrast, the oxidative
enzymes, LACCASE4 and LACCASE17 were spatially localized to secondary cell
walls throughout protoxylem TE differentiation. These data support the hypothesis
that precise delivery of oxidative enzymes determines the pattern of cell wall
lignification. This view was supported by lac4lac17 mutant analysis demonstrating
that laccases are necessary for protoxylem TE lignification. Overexpression studies
showed that laccases are sufficient to catalyze ectopic lignin polymerization in
primary cell walls when exogenous monolignols are supplied. Our data support a
model of protoxylem TE lignification where monolignols are highly mobile once
exported to the cell wall, and in which precise targeting of laccases to secondary cell
wall domains directs lignification deposition.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
3
Introduction
During all stages of a land plant’s life cycle, lignified secondary cell walls provide
critical mechanical properties for water transport and the upright growth habit. Waterconduction in xylem tissue occurs in tracheary elements (TEs), whose secondary cell
walls are composed of cellulose, hemicelluloses, and the phenolic lignin polymer.
Lignification of the secondary cell wall imparts strength, rigidity and water
impermeability to the polysaccharide components. The deposition of lignin in
secondary cell walls is developmentally regulated, and different cell types generate
distinctive secondary cell wall patterns. Protoxylem TEs, for example, form annular
or helical secondary wall thickenings, whereas metaxylem TEs deposit secondary cell
walls in a reticulated or pitted pattern (Esau, 1965). Protoxylem TEs form in young
elongating plant tissues and therefore the restriction of lignin deposition to the annular
or helical secondary cell wall thickenings, as opposed to the intervening primary cell
walls, is crucial to allow continued axial elongation. The mechanisms restricting
lignin deposition specifically to secondary cell wall thickenings have not been
identified.
Monolignol (lignin monomer) biosynthesis occurs in the cytosol in close
proximity to the endoplasmic reticulum (ER), because the pathway includes both
cytosolic- and ER-localized enzymes (Bonawitz and Chapple, 2010). Multi-enzyme
complexes, potentially anchored at specialized sub-domains on the ER surface, have
been postulated to channel phenolic metabolite production during lignification (Chen
et al., 2011a; Bassard et al., 2012). Both ATP-binding cassette (ABC) transporters of
monolignols and proton-dependent transporters of monolignol glucosides have been
proposed as monolignol export mechanisms (Ehlting et al., 2005; Liu, C.J., 2012;
Miao and Liu, 2010; Tsuyama et al., 2013). However, genetic evidence does not
support a role for monolignol glucosides as direct precursors of lignin in Arabidopsis
(Chapelle et al., 2012). Monolignol transport activity has been reported for
heterologously expressed AtABCG29, which was capable of transporting p-coumaryl
alcohol in yeast cells and isolated PM vesicles (Alejandro et al., 2012). Localized
monolignol biosynthesis and/or export are hypothesized to be mechanisms for
concentrating monolignols in secondary, but not primary, cell wall domains in
protoxylem TEs.
An alternative mechanism for controlling the pattern of lignin deposition could be
the site-specific localization of the enzymes required for monolignol polymerization
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
4
within the cell wall. Monolignol polymerization occurs by oxidative combinatorial
coupling of monolignols, catalyzed by cell wall-localized laccases and/or peroxidases
(Vanholme et al., 2012). A peroxidase (ZPO-C), originally isolated from xylogenic
Zinnia elegans cell cultures, has been localized to secondary cell walls, which
demonstrates the spatial association of oxidative enzymes with lignin deposition (Sato
et al., 2006), but the function of this peroxidase in vivo has not been demonstrated.
Recent analysis of the Arabidopsis LACCASE4 (LAC4), LACCASE11 (LAC11) and
LACCASE17 (LAC17) genes showed that loss-of-function of these genes had a
dramatic effect on lignification of metaxylem and fiber cells in inflorescence stems
(Berthet et al., 2011; Zhao et al., 2013). However, the roles of either laccases or
peroxidases in controlling the spatial pattern of cell wall lignification in protoxylem
TEs have not been investigated.
Studying protoxylem TE development is challenging as they are located deep
within root or shoot tissues, but the identification of key transcriptional regulators that
activate differentiation of protoxylem TEs has led to the development of a genetically
tractable experimental system. In this system, the activity of VASCULAR NAC
DOMAIN 7 (VND7), a NAC domain transcription factor that regulates protoxylem TE
cell fate (Kubo et al., 2005), is induced so that nearly all cells in Arabidopsis
seedlings transdifferentiate into protoxylem TE-like cells (Yamaguchi et al., 2010).
This system allows protoxylem development to be studied in diverse mutant
backgrounds, and permits the expression and high-resolution imaging of proteins
tagged with green fluorescent protein (GFP) during xylogenesis.
The present study, examines the cellular mechanisms underlying the discrete
lignification of spiral or annular secondary cell walls in protoxylem TEs. With GFPtagging and live-cell monitoring in the VND7-VP16-GR lines, we tested if monolignol
biosynthetic enzymes, putative monolignol transporters, or laccases were specifically
localized in lignifying cell wall domains. Fluorescently tagged monolignols were used
to assay whether monolignols would polymerize in secondary cell wall domains in
laccase loss-of-function mutants, as well as in primary cell walls of overexpression
lines. While earlier studies identified which members of the laccase gene family were
active in lignification of metaxylem vessel and fiber cell types in the Arabidopsis
stem (Berthet et al., 2011; Zhao et al., 2013), this work directly addresses the role of
these oxidative enzymes in directing lignin polymerization in lignified secondary cell
wall domains adjacent to unlignified primary cell walls of the protoxylem TE.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
5
Results
Live-cell imaging of lignification of induced protoxylem TEs
Although the genes encoding lignin biosynthetic enzymes are up-regulated in plant
lines expressing the master transcription factor VND7, the presence of lignin in the
induced protoxylem TEs has not been established. To examine lignification in
induced protoxylem TEs, cell wall phenolic autofluorescence was profiled using twophoton excitation microscopy, with ultraviolet (UV) excitation (350-370 nm) and
emission (420-540 nm) (Fig. 1). In hypocotyls from uninduced control seedlings, the
only intrinsic fluorescence from lignified secondary cell walls came from endogenous
protoxylem TEs in the central vascular cylinder (Fig. 1A). However, within 48 hours
of VND7 induction, nearly 100% of hypocotyl epidermal cells of VND7-VP16-GR
seedlings transdifferentiated into protoxylem TEs with highly fluorescent secondary
cell walls (Fig. 1B,C).
In order to verify that the UV autofluorescence observed in VND7-induced TEs is
dependent on monolignol biosynthesis, the expression of two key monolignol
biosynthetic genes, CINNAMATE-4-HYDROXYLASE (C4H) or CINNAMOYL-CoA
REDUCTASE (CCR1) was suppressed through the use of targeted artificial
microRNAs (Schwab et al., 2006). Plants constitutively overexpressing the C4H or
CCR1 microRNA recapitulated the range of phenotypes previously described in c4h
and ccr1 mutants (Schilmiller et al., 2009; Jones et al., 2001; Thevenin et al., 2011),
including irregular xylem and a marked reduction in lignin deposition
(Pro35S:miRNA C4H lines, Supplemental Fig. 1; Pro35S:miRNA CCR lines, Smith et
al., 2013). Constitutive expression of the C4H or CCR1 microRNA during VND7induced TE formation did not inhibit secondary cell wall formation (Fig. 1D,E insets)
but two-photon microscopy revealed severely reduced UV autofluorescence in these
cell wall domains (Fig. 1D,E), compared to wild type (Fig. 1C). Together, these
results indicate that the autofluorescence in secondary cell walls of VND7-induced
protoxylem TEs is from lignin, as it is dependent not only on the function of the
general phenylpropanoid pathway enzyme C4H, but also on the function of the
monolignol-specific pathway enzyme CCR1. This establishes the VND7-inducible
protoxylem system as a valid model for analysis of cell wall lignification.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
6
Dynamic rearrangement of monolignol biosynthetic enzymes during protoxylem
TE differentiation
One hypothesis to explain specific deposition of lignin only in spiral cell wall
thickenings of protoxylem TEs is that monolignol biosynthetic enzymes in these cells
may be preferentially organized around sites of secondary cell wall deposition. To
directly image the subcellular distribution of C4H during protoxylem TE
differentiation, a C4H-GFP translational fusion was stably transformed into VND7VP16-GR plants. Similar C4H-GFP fusions have been shown to be functional in
plants and localized to the endoplasmic reticulum when transiently expressed in
tobacco leaf cells (Bassard et al., 2012; Ro et al., 2001). In epidermal cells of
uninduced C4H-GFP-transformed VND7-VP16-GR plants, C4H-GFP was clearly
localized to the reticulate network of tubular ER, as confirmed by colocalization with
an ER-localized RFP-HDEL marker (Supplemental Fig. 2). Induction of VND7mediated protoxylem TE differentiation had a profound impact on the pattern of C4HGFP distribution. Live-cell imaging was used to visualize the C4H-GFP-labeled ER
pattern, as it changed from an open tubular ER network early in development
(Supplemental Fig. 2A-C) to a denser lamellar pattern concentrated between
developing secondary cell wall domains (Fig. 2B; Supplemental Fig. 2D-G).
Continuous ER strands were parallel to, and generally excluded from, domains
directly below developing secondary cell wall thickenings, which were observed in
bright-field microscopy (Fig. 2A). Evidence of ER exclusion below secondary cell
wall domains was also observed in endogenous root protoxylem TEs using
transmission electron microscopy (TEM) of high-pressure frozen/freeze-substituted
samples (Fig. 2C). The changes of C4H-GFP distribution during VND7-induced TE
differentiation are therefore associated with overall changes in ER distribution and
morphology, but do not appear to be due to re-localization of C4H-GFP to specific
regions of the ER.
As CINNAMOYL-CoA REDUCTASE1 (CCR1) is specific to monolignol
biosynthesis, its subcellular distribution was tracked during VND7-mediated TE
lignification. First, the function of the CCR1-GFP fusion protein was tested by
complementing ccr1irx4 loss-of-function mutants, where CCR1-GFP rescued the
irregular xylem phenotype of ccr1irx4 mutants (Supplemental Fig. 3). CCR1 is
predicted to be a cytosolic enzyme and CCR1-GFP localization exactly matched
cytosolic-localized GFP (cytGFP) (Fig. 2D,E). During all stages of VND7-mediated
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
7
TE differentiation, both CCR1-GFP and cytGFP signals were observed throughout the
cytoplasm, with no bias towards secondary cell wall domains (Fig. 2F). Thus, spiral
secondary cell wall lignification is not correlated with the spatial distribution of the
C4H and CCR1 monolignol biosynthetic enzymes.
ATP-Binding Cassette (ABC) transporters localize to all plasma membrane
domains during protoxylem TE differentiation
ABCB11, ABCG29 and ABCG33 transporters have previously been identified as
candidate monolignol exporters (Ehlting et al., 2005; Kaneda et al., 2011; Alejandro
et al., 2012). To test the hypothesis that these putative monolignol transporters could
have a restricted PM localization adjacent to secondary cell wall domains, GFP
translational fusions of ABCB11, ABCG29 and ABCG33 were transformed into
VND7-VP16-GR plants. When VND7-mediated protoxylem TE differentiation was
induced, all fluorescently tagged ABC transporters were localized to plasma
membranes (Fig. 3; Supplemental Fig. 4). YFP-tagged ABCG11, which has
previously been shown to function as a plasma membrane localized cuticular wax and
cutin exporter (Bird et al., 2007), was included as a positive control for plasma
membrane localization (Supplemental Fig. 4). Secondary cell wall domains were
counterstained with the cellulose-specific Pontamine S4B dye (Fig. 3B,C; Anderson
et al., 2010), and when the GFP channel signal was merged with that of the cellulose
counter-stain, it was clear that transporters were equally distributed along both
secondary and adjacent primary wall regions (Fig. 3A,C). This uniform plasma
membrane distribution of candidate monolignol ABC transporters persisted until the
initiation of programmed cell death, when fragmentation of the GFP-labeled plasma
membranes was observed (Supplemental Movie 1 and Supplemental Fig. 5). These
results demonstrate that the candidate monolignol ABC transporters, including
ABCG29 (Fig. 3D), are not preferentially excluded from, nor localized to, the plasma
membrane domains facing developing secondary cell walls.
Laccase-dependent monolignol polymerization in secondary cell wall domains
The results of the preceding experiments suggest that monolignols are likely
produced and exported into the cell wall in a diffuse, rather than targeted, manner. If
this model is correct, then exogenously supplied monolignols are predicted to be
highly mobile throughout the cell wall matrix, and should only become polymerized
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
8
within the secondary cell wall domains. The tools to assess this prediction have
recently been developed, in the form of fluorescently tagged monolignol analogs,
such as γ-nitrobenzofuran-tagged coniferyl alcohol (NBD-CA), that become
polymerized into lignin via oxidase-mediated radical coupling (Tobimatsu et al.,
2011). When applied in planta, these NBD-tagged probes were incorporated into the
lignifying tissues in Arabidopsis and Pinus radiata plants (Tobimatsu et al., 2013).
The availability of NBD-tagged-monolignols together with the VND7-induced
protoxylem system provided the opportunity to document the lignification of discrete
cell wall domains in individual protoxylem TEs. NBD-CA and untagged coniferyl
alcohol (CA) were applied together to induced VND7-VP16-GR seedlings and
labeling of the differentiating TEs was monitored. To ensure that free monolignols
were not simply being trapped in the polymer matrix following treatment with
fluorescent monolignols, and that only the insoluble lignin polymer was being
imaged, treated seedlings were thoroughly washed in 49% methanol: 1% acetic acid
at 45 °C for up to 72 h to remove excess NBD-CA, prior to imaging.
Secondary cell wall domains, as identified by S4B staining for cellulose, were
strongly labeled by NBD-CA treatment, whereas adjacent primary cell walls were not
(arrowheads and arrows respectively in Fig. 4A-C). Similarly, in leaf tissue in which
only a single leaf mesophyll cell had transdifferentiated into a protoxylem TE,
incorporation of NBD-CA was restricted to the secondary cell walls of the
transdifferentiated cell (Fig. 4D-F). Collectively, these results demonstrate that
monolignols can diffuse freely throughout the apoplast and become polymerized into
lignin only in the secondary cell walls of differentiating protoxylem TEs.
If monolignols are exported to, and are mobile in, all cell wall domains, but
become polymerized only in discrete secondary cell wall thickenings, then the
presence of laccases and/or peroxidases in those spiral wall thickenings could
determine the sites of lignin polymerization. The LACCASE4 (LAC4) and
LACCASE17 (LAC17) genes are two of the most strongly differentially expressed
genes in plant cells responding to increased VND7 or VND6 activity (Ohashi-Ito et
al., 2010; Yamaguchi et al., 2010; Yamaguchi et al., 2011). In order to directly test
the contribution of these laccases to the lignification patterns of protoxylem TE cell
walls, the inducible VND7 construct was transformed into lac4 lac17 double mutants.
In contrast to strong NBD-CA incorporation in the secondary cell walls of induced
protoxylem in the wild-type VND7-VP16-GR seedlings (arrowheads in Fig. 5A), lac4
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
9
lac17 double mutants did not incorporate the fluorescent monolignols following
induction of VND7 (arrowheads in Fig. 5C). At the same time, VND7-mediated
secondary cell wall formation was not affected in lac4 lac17 mutants because
secondary wall thickenings in both wild-type (Fig. 5B) and lac4 lac17 double mutants
(Fig. 5D) were identical, as assessed by cellulose staining with Pontamine S4B, and
immunofluorescent labeling of xylan using LM10 antibodies. These results
demonstrate that although other secondary cell wall components such as
glucuronoxylan or dense deposits of cellulose were present in the lac4 lac17 mutants,
the polysaccharides were not sufficient to trap free monolignols in secondary cell
walls in the absence of the required laccases.
The loss-of-function mutant analysis demonstrated that LAC4 and LAC17 were
necessary for protoxylem lignification. In order to evaluate if these laccase proteins
are sufficient for lignin polymerization, we generated plant lines constitutively
overexpressing LAC4 (Pro35S:LAC4) or LAC17 (Pro35S:LAC17) and tested whether
primary cell walled cells in Arabidopsis seedlings would lignify. We predicted that
the presence of LAC4 or LAC17 in a cell wall, even a primary cell wall, would be
sufficient to enable cell wall lignification when monolignols were exogenously
applied. When Pro35S:LAC4 or Pro35S:LAC17 seedlings were treated with
exogenously-supplied, fluorescently
tagged monolignols, primary cell walls in
cotyledon epidermal cells had strong fluorescence, thus indicating cell wall
lignification (Fig. 5E,F). In contrast, wild-type cotyledons did not incorporate the
fluorescently tagged monolignols (Fig. 5G,H). Taken together, it is evident that
laccases are both necessary and capable of directing lignification in cell walls when
monolignols are available.
LACCASE4 and LACCASE17 localize to secondary cell walls in protoxylem TEs
If the presence of the enzymes responsible for oxidative coupling of monolignols is
the key factor determining where lignin is deposited, then the spatial organization of
these apoplastic proteins is predicted to be tightly controlled. To test this hypothesis,
LACCASE4 and LACCASE17 were tagged with mCherry (ProLAC4:LAC4-mCherry
and ProLAC17:LAC17-mCherry), a pH-stable monomeric red fluorescent protein that
can function in the acidic environment of the apoplast. The ‘irregular xylem’
phenotype of lac4lac17 double mutant inflorescence stems was complemented by
expression of the LAC4-mCherry construct, thus demonstrating that the LAC4-
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
10
mCherry protein is functional in planta (Supplemental Fig. 3). In primary roots from
lac4lac17 seedlings expressing ProLAC4:LAC4-mCherry, the fluorescent signal was
specifically detected in endogenous TEs and was localized to the helical and pitted
secondary cell wall thickenings of protoxylem and metaxylem TEs, respectively (Fig.
6A,B). The expression and subcellular localization of LAC17-mCherry was identical
to those observed for LAC4-mCherry, and is shown in Supplemental Fig. 6.
To visualize the LAC4-mCherry pattern at high-resolution, protoxylem was
induced on the plant surface using VND7-VP16-GR plants transformed with the
ProLAC4:LAC4-mCherry construct. The LAC4-mCherry signal was specifically
localized to the developing secondary cell wall domains during the early stages of
VND7-induced TE differentiation, as shown in a single optical section (Fig. 6C,D).
The concentration of LAC4-mCherry within developing secondary cell walls
continued throughout TE differentiation and persisted after programmed cell death
(maximum projection image, Fig. 6E). The pattern of LAC4-mCherry distribution
was identical to the patterns of LM10 immunofluorescence labeling of
glucuronoxylan in secondary cell walls (immunolabeled sections in Fig. 6F and inset
in 6G). The localization of LACCASE4 and LACCASE17 exclusively to secondary
cell wall domains, and loss of monolignol incorporation in the lac4 lac17 double
mutants, indicate that apoplastic targeting of these laccases plays a key role in the
precise pattern of lignin deposition in protoxylem TEs.
Discussion
The goal of this study was to identify mechanisms that allow protoxylem TEs to
specifically deposit lignin in spiral cell wall thickenings, while the intervening
primary cell walls remain lignin-free. This pattern of secondary cell wall lignification
is crucial for the ability of protoxylem TEs to facilitate water transport to young and
rapidly expanding plant organs. The combination of two new technologies,
fluorescently tagged monolignols and the VND7-inducible protoxylem system in
Arabidopsis, provided a platform to directly visualize the lignification process in an
experimental system with rich transcriptomic and molecular genetic resources. The
precise spatial association of fluorescent-protein-tagged LACCASE (LAC4-mCherry
and LAC17-mCherry) with the spiral wall domains demonstrated that these oxidative
enzymes are localized to secondary cell walls. The loss of lignification in lac4lac17
mutants, and ectopic lignification in overexpression lines of LAC4 and LAC17,
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
11
demonstrated that laccases are necessary and sufficient to direct lignin
polymerization. In contrast, localization of GFP-tagged monolignol biosynthetic
enzymes and putative monolignol transporter proteins in developing protoxylem TEs
demonstrated that the spatial distribution of these components was not correlated with
the lignified cell wall pattern.
The relevant laccases in this protoxylem system were predicted from
transcriptomic studies that identified LACCASE4 and LACCASE17 as genes that were
strongly differentially expressed in response to the master regulator transcription
factor VND7 (Yamaguchi et al., 2010; Yamaguchi et al., 2011). The dramatic
reduction of fluorescent lignin incorporation in protoxylem TEs from lac4 lac17
mutants, was more severe than the 40% reduction in total lignin reported earlier in
lac4 lac17 double mutant inflorescence stems (Berthet et al., 2011). This difference
may reflect the different types of lignified cells being examined in each study, i.e. the
abundance of fibers and metaxylem TEs in the Arabidopsis inflorescence stem versus
protoxylem TEs. While lac4 lac17 double mutant phenotypes in stems included
hypolignified fibers and irregular metaxylem vessels (Berthet et al., 2011), loss of
function of a third laccase, in the lac4 lac17 lac11 triple mutants, led to severe growth
defects and a striking loss of lignified cells in the stem (Zhao et al., 2013). These data
made the case that LAC11 was a third redundant component required for lignification
in the fiber-rich stem, and this is consistent with the preferential activation of LAC11
expression by the fiber-specific transcription factor SECONDARY WALLASSOCIATED NAC DOMAIN PROTEIN1 (Ohashi-Ito et al., 2010; Zhao et al.,
2013). Thus, analysis of gene expression and mutant phenotypes both indicate that,
while the combination of LAC4, LAC17, and LAC11 supports lignification in stems,
LAC4 and LAC17 are the major players in protoxylem TE lignification.
Although clarification of the functional members of the laccase gene family is
interesting, it is the precise localization of these laccases to the spiral secondary cell
walls of protoxylem TEs that is most relevant for the targeted lignification of these
secondary cell wall domains. During protoxylem development, there are many Golgi
bodies that pause at the microtubule-lined plasma membrane domain underlying the
secondary cell wall (Wightman and Turner, 2008), presumably to deliver xylans,
cellulose synthases, and cell wall proteins. LAC4-mCherry fluorescence was detected
in nascent secondary cell wall thickenings from the earliest stages of secondary cell
wall formation, suggesting a testable hypothesis that laccases are secreted
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
12
simultaneously with secondary cell wall matrix polysaccharides. The molecular
mechanisms underlying the specific targeting of secondary cell wall cellulose
synthases, xylans, and laccases to the plasma membrane domains below developing
secondary cell walls are still unknown. The high flux of vesicle delivery to the
microtubule-lined plasma membrane domains may represent a transient reorientation
of secretory vesicle flow during this stage of differentiation. A similar phenomenon
was hypothesized to account for the directed delivery of plasma membrane proteins to
the periarbuscular membrane during arbuscular mycorrhizal symbiosis in Medicago
(Pumplin et al., 2012). Although a transient reorientation of secretory vesicle traffic
is the simplest explanation, an alternative hypothesis of uniform secretion of laccases
coupled to rapid re-mobilization toward secondary cell walls, resulting in the
observed pattern of LAC4 and LAC17 localization, cannot be ruled out. The
development of LAC4-mCherry and LAC17-mCherry proteins provides important
tools to study these hypothesized targeting mechanisms.
The model that monolignols are exported out of the cell into all areas of the
apoplast, rather than being selectively exported into secondary versus primary cell
walls, is in agreement with the recent characterization of Casparian strip lignification
in root endodermal cells. Lee et al. (2013) demonstrated that, while plasmamembrane-bound CASP proteins function to organize the extracellular oxidative
peroxidases specifically to lignifying Casparian strips, other plasma membrane
proteins, including the monolignol transporter ABCG29, are excluded from Casparian
strip domains by the CASP complexes (Roppolo et al., 2011; Alejandro et al., 2012).
Given the exclusion of ABCG29 from the lignifying domain of the Casparian strip,
we hypothesized that a similar, or inverse, pattern might be observed in induced
protoxylem TE expressing a variety of GFP-tagged putative monolignol transporters
(Alejandro et al., 2012). However, the uniform localization of several different ABC
transporters during protoxylem differentiation was not consistent with the hypothesis
that localized monolignol export facilitates specific lignification patterns. A careful
examination of the transcriptome of the VND7-induced seedlings (Yamaguchi et al.,
2010; Yamaguchi et al., 2011) did not reveal increased expression of CASP, CASPlike proteins or peroxidases. Instead, which cell domains lignify appears to be
determined by the oxidative machinery and is independent of the source of
monolignols, an observation compatible with the ‘good neighbor’ hypothesis that
cells surrounding lignifying TEs may actively contribute to TE lignification
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
13
(Hosokawa et al., 2001; Tokunaga et al., 2005; Pesquet et al., 2013; Smith et al.,
2013). However, future research may identify other components that may be limiting
for the lignification process during protoxylem TE differentiation in addition to LAC4
and LAC17 .
In lac4lac17 mutants, the secondary cell wall polysaccharide environment
remained intact, but this was not sufficient to create the secondary cell wall
lignification pattern. These data indicate that differences in nano-scale cell wall
polysaccharide environments between the pectin/xyloglucan-rich primary cell wall
and the xylan-rich secondary cell wall do not contribute to trapping monolignols in
protoxylem secondary cell wall domains. Instead, the presence of laccases in
secondary cell wall domains appears to be a key feature, because LAC4/LAC17 were
both necessary for, and capable of, directing cell wall lignification.
Materials and Methods
Plant Growth
All Arabidopsis plants were grown under 16 h light/21 °C and 8 h dark/16 °C
conditions. Seeds were surface sterilized using chlorine gas generated in a closed
container using 100 ml of bleach and 3 ml of concentrated HCl for 3-6 h. Seeds were
subsequently plated on ½ MS (Murashige and Skoog) medium (Sigma), kept in the
dark at 4 °C for 2-3 days and subsequently moved to growth conditions. Pro35S:RFPHDEL plants and plasmid (Nelson et al., 2007) were obtained from ABRC (stock
#CD3 960). lac4-2 lac17 lines were obtained from Richard Sibout and Lise Jouanin,
and originally described in Berthet et al. (2011). Pro35S:VND7-VP16-GR seedlings
and plasmid were originally described in Yamaguchi et al. (2010). VND7 activity in
7-10-day-old seedlings was induced as described in Yamaguchi et al. (2010).
All transgenic plant lines were generated using Col-0 Arabidopsis plants,
Agrobacterium tumefaciens (strain GV3101) and the floral dip method. Primary
transformants were plated on ½ MS media and selected for survival on 50 μg/ml
kanamycin, 25 μg/ml hygromycin, 25 μg/ml Basta (DL-phosphinothricin: Bayer), or
by spraying 10-day-old seedlings grown on soil with 120 μg/ml Basta.
Cell Wall Labeling
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
14
Cellulose staining using Pontamine Fast Scarlet 4B (S4B; Sigma) was performed
by incubating seedlings in 10mg/ml S4B solution in ½ MS media for 10-20 min and
washing once in ½ MS prior to imaging. Labeling cell wall lignin using NBD-CA was
carried out as follows: a mixture of 0.5 mM NBD-CA (Tobimatsu et al., 2011;
Tobimatsu et al., 2013) and 5 mM CA (Sigma) was added to liquid ½ MS media to a
final concentration of 1 and 10 μM, respectively, together with 10 μM dexamethasone
(Sigma) for VND7 induction. VND7-VP16-GR seedlings were incubated in this
induction solution for 4 to 48 hours, gently shaking under regular growth conditions
(listed above). No differences in NBD-CA incorporation were observed when preinducing VND7 activity for 24 hours prior to adding the NBD-CA mix. Seedlings
were subsequently rinsed 2x for 15 min in ½ MS solution and fixed in 4% paraformaldehyde, 50 mM PIPES, 5 mM MgSO4, 5 mM EGTA solution overnight.
Seedlings were subsequently washed 2x in TBST (10
0.1%
Tween
20)
mM Tris pH 7, 0.25 M NaCl,
solution and then in 49% methanol 1% acetic acid solution heated
to 45 °C for 4 to 72 h with gentle shaking to wash away unincorporated NBD-CA.
Seedlings were subsequently washed and rehydrated in a graded methanol series,
mounted in ½ MS and imaged. For whole mount immunolabeling, VND7-induced
seedlings were fixed as described above and stored in TBST buffer at 4 °C until
hybridization. Fixed seedlings were incubated in 5% BSA (Sigma) in TBST for 1-2
hours at room temperature, and subsequently incubated with LM10 primary antibody
(McCartney et al., 2005) at 1/50 dilution overnight at 4 °C. Seedlings were rinsed 3x
in TBST solution for 5 min, and subsequently incubated in 1/100 dilution of
secondary anti-rat Alexa 488 antibody (Invitrogen) at room temperature for 1 h. After
washing 2x with TBST, samples were mounted onto slides and imaged. For
immunolocalization of thick sections, the above procedure was performed on
sectioned material from LR white resin infiltrated seedlings after high-pressure
freezing and freeze-substitution in glutaraldehyde and uranyl acetate. Sections were
generated using a Leica Ultracut UCT Ultramicrotome and mounted on Teflon-coated
glass slides (Electron Microscopy Sciences).
Microscopy and Image analysis
For live-cell imaging a Perkin-Elmer UltraView VoX spinning disk confocal
mounted on a Leica DMI6000 inverted microscope and a Hamamatsu 9100-02 CCD
camera was used with the following excitation/emission filters: GFP (488/525), YFP
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
15
(514/540), and RFP (561/595). Samples were mounted in water and imaged using a
Leica oil immersion 63X objective (Plan-Apo, NA 1.4). All images were processed
using Volocity image analysis software (Improvision). GFP settings were used for
imaging lignin with NBD-CA and RFP settings were used for imaging cellulose after
treatment with S4B.
For visualizing lignin autofluorescence, an Olympus FV1000 Multiphoton Laser
Scanning Microscope with a tunable MaiTai BB DeepSee (710-990 nm) laser
adjusted to 730 nm was used. Simultaneous absorption of two 730 nm photons results
in excitation of lignin at approximately 350-370 nm. During imaging, two emission
channels were simultaneous collected using 420-460 nm and 495-540 nm filter sets.
Samples were fixed in ethanol:acetic acid (6:1) solution for 24 h, washed twice with
95% ethanol and gradually rehydrated using 70% ethanol, 30% ethanol and water
solutions. Samples were subsequently mounted in chloral hydrate solution (chloral
hydrate: glycerol: water, 9:1:3) and imaged using an Olympus XL Plan N25X
objective. All images were processed using Volocity image analysis software
(Improvision) and Photoshop software (Adobe systems). For Supplemental Figs. 1
and 3, hand sections from mature Arabidopsis inflorescence stems were stained with
toluidine blue (Ted Pella Inc.) or freshly mounted and imaged for lignin
autofluorescence using a Leica DMR microscope equipped with a standard mercury
arc lamp with excitation (340-380 nm) and emission (450 nm) filter sets. For TEM
analysis of TEs, 7-day-old seedling roots were high-pressure frozen, freeze
substituted, sectioned and prepared for TEM as previously described (Smith et al.,
2013).
Molecular biology
All DNA sequences were cloned into pDONR221 or pDONRzeo using Gateway™
cloning methodology (Invitrogen) and subsequently transferred into specific
expression vectors as indicated. All PCR amplifications were carried out using
Phusion High fidelity DNA polymerase (New England Biolabs) and two-step adapter
PCR was used to incorporate full-length Gateway™ attb sequences into the PCR
products.
Genomic regions containing the open reading frames (ORF) for ABCB11,
ABCG29, ABCG33, C4H, CCR1 were subsequently cloned into the vectors driven by
the UBIQUITIN10 promoter (Grefen et al., 2010) and were transformed into VND7-
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
16
VP16-GR
plants;
C-terminal
GFP
fusions
(ProUBQ10:ABCB11-GFP,
ProUBQ10:C4H-GFP, ProUBQ10:CCR1-GFP) and N-terminal GFP fusions
(ProUBQ10:GFP-ABCG29 and ProUBQ10:GFP-ABCG33). Pro35S:YFP-ABCG11
plants (Bird et al., 2007) were crossed with Pro35:VND7-VP16-GR plants using
standard crossing techniques. A genomic LAC4 and LAC17 fragment containing 2 kb
of the 5΄ promoter and ORF with no stop codon was amplified and cloned into a
modified pMDC111 vector (Curtis and Grossniklaus, 2003), where the GFP coding
region was excised by digesting with the Asc1 and Sac1 restriction enzymes, and
replaced with the mCherry coding sequence. Pro35S:LAC4 and Pro35S:LAC17
constructs were generated by cloning the respective genomic ORFs into the pK2GW7
vector (Plant Systems biology, VIB, Ghent University) and subsequent transformation
into wild-type plants.
Artificial microRNA targeting C4H and CCR1 was designed using WMD2
(wmd2.weigelworld.org) and generated as described in Schwab et al. (2006), and
were subsequently cloned into the pK2GW7 vector (Plant Systems Biology, VIB,
Ghent University).
Accession numbers
LAC4, AT2G38080; LAC17, AT5G60020; CCR1, AT1G15950; C4H, AT2G30490;
VND7, AT1G71930; ABCG11, AT1G17840; ABCB11, AT1G02520; ABCG29,
AT3G16340; ABCG33, AT2G37280.
Acknowledgements
We thank the UBC Bioimaging Facility for technical support and Etienne
Grienenberger for technical advice and comments on the manuscript. This research
was funded by the Natural Sciences and Engineering Research Council of Canada
through Discovery and CREATE grants to BEE and ALS. TD was supported in part
by JSPS KAKENHI Grant Numbers 24114002 and 25291062 and by NAIST Global
Initiative Program. YT and JR acknowledge funding from US Department of Energy,
the Office of Science (DE-SC0006930), and by the DOE Great Lakes Bioenergy
Research Center (DOE Office of Science BER DE-FC02-07ER64494).
Author Contributions:
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
17
MS: designed and initiated the research, performed the experiments, and wrote the
manuscript; RS and AB: performed experiments and contributed to writing the
manuscript; YW: provided materials, contributed to experimental design and writing
the manuscript; YT, JR, BE, and TD: provided unpublished materials and contributed
to writing the manuscript; LS: designed and initiated the research, and wrote the
manuscript.
Figure Legends
Figure 1. Lignin autofluorescence in secondary cell walls of endogenous and
VND7-induced Arabidopsis seedling TEs.
(A) In a control, uninduced, 10-day-old Arabidopsis hypocotyl, only protoxylem TEs
(arrow) emit autofluorescence when excited by UV light. (B) In an Arabidopsis
hypocotyl carrying VND7-VP16-GR, after induction with DEX, epidermal cells
transdifferentiated into protoxylem TEs (arrows) and emit autofluorescence. (C-E)
UV autofluorescence of protoxylem TEs imaged under identical conditions in VND7VP16-GR alone, or in plants expressing both VND7-VP16-GR and artificial
microRNAs targeting monolignol biosynthetic genes (D) C4H or (E) CCR1.
Protoxylem TE differentiation and secondary cell wall (arrowheads) formation was
not inhibited by constitutive expression of artificial microRNAs [insets in D and E].
Maximum projection images of z-stacks are shown. Scale bars are 50 μm.
Figure 2. CINNAMATE-4-HYDROXYLASE (C4H-GFP) and CINNAMOYLCoA-REDUCTASE (CCR1-GFP) are localized between secondary cell walls
domains during VND7-induced protoxylem TE differentiation.
(A) Bright field image of a differentiating protoxylem TE induced in VND7-VP16-GR
plants, showing secondary cell wall thickenings (arrowheads). (B) Maximum
projection image of optical sections through the same cell show in A, where
CINNAMATE-4-HYDROXYLASE (C4H-GFP) localization is observed in the
reticulate ER network that is excluded below developing secondary cell walls
(arrowheads) and is adjacent to primary cell wall domains. (C) Transmission electron
micrograph showing laminar ER (ER), Golgi (G), and mitochondria (m) in relation to
secondary cell walls (scw) of root protoxylem TEs. (D) Maximum projection images
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
18
of optical sections showing cytoplasmic localization of CINNAMOYL-CoA
REDUCTASE (CCR1-GFP) and (E) cytosolic GFP (cytGFP) localization in control
hypocotyl epidermal cells of VND7-VP16-GR seedlings without DEX induction.
After VND7-VP16-GR protoxylem TE induction for 24 h, both (F) CCR1-GFP and
(G) cytGFP were predominately localized in the cytoplasm between the secondary
cell wall thickenings and as shown in a single median optical slice in (H) CCR1-GFP
and (I) cytGFP. n, nucleus; m, mitochondria; G, Golgi; er, endoplasmic reticulum;
scw, secondary cell wall. Scale bars are 12μm A,B, D-G. and 500 nm in C.
Figure 3. ABC transporters are evenly localized in plasma membranes during
protoxylem TE differentiation in VND7-VP16-GR seedlings.
(A) Maximum projection image of cotyledon epidermal cells undergoing protoxylem
TE differentiation, expressing GFP-ABCG33. (B) Cellulose stain, Pontamine S4B,
counter-stained image of cells shown in E, highlighting secondary cell wall domains.
(C) Median optical slice of induced protoxylem TE, showing invaginations of GFPABCG33 (green), counter-stained with Pontamine S4B (red) demonstrating secondary
cell wall domains, with inset image of the optical cross section of one secondary wall
thickening. (D) Induced protoxylem TE from plants expressing GFP-ABCG29, with
inset showing plasma membrane localization over primary and secondary wall
domains. Scale bars are 15 μm.
Figure 4. Mobility of monolignols in the apoplast: fluorescently tagged
monolignols are specifically incorporated into the secondary cell walls of VND7induced protoxylem TEs.
(A) Green fluorescence of incorporated nitrobenzofuran-tagged coniferyl alcohol
(NBD-CA) specifically in secondary cell walls (arrowhead) and not primary cell walls
(arrows) of epidermal cells induced to transdifferentiate into protoxylem TEs. (B)
Cellulose-specific Pontamine S4B counter-stained image of cells shown in A,
demonstrating secondary cell walls (arrowhead) and primary cell walls (arrow). (C)
Merged image of A and B showing colocalization of signal in secondary cell walls.
(D) In an area where only one leaf mesophyll cell differentiated into a protoxylem TE,
exogenously added NBD-CA was specifically incorporated in secondary wall
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
19
thickenings and not in the surrounding neighbors. (E) Pontamine S4B counter-stained
image of protoxylem TE shown in D, demonstrating background stain of primary cell
wall domains of surrounding leaf cells, and strong secondary cell wall domains in
lone protoxylem TE. (F) Merged image of D and E. Scale bars are 15 μm.
Figure 5. LACCASEs are necessary and sufficient to direct lignin polymerization
in Arabidopsis cell wall domains.
(A) Maximum projection image of hypocotyl epidermal cells induced to
transdifferentiate into protoxylem TEs in VND7-VP16-GR seedlings, showing NBDCA fluorescence in the secondary cell walls (arrowheads). (B) VND7-induced
protoxylem TE showing cellulose deposition (i;Pontamine S4B stain), whole mount
immunolocalization of xylan (ii; LM10 antibody), and (iii) merged images. (C)
Maximum projection image of epidermal cells induced to transdifferentiate into
protoxylem TE, in VND7-VP16-GR seedlings in lac4 lac17 double mutant
backgrounds showing that NBD-CA was not incorporated into the secondary cell
walls (arrowheads). (D) VND7-VP16-GR induced protoxylem TE in lac4 lac17
double mutants showing identical deposition patterns for cellulose (i; Pontamine S4B
stain), xylan (ii; LM10 immunolocalization), and (iii) merged images, as observed in
VND7-VP16-GR alone shown in B. Constitutive expression of (E) LAC4 or (F)
LAC17 catalyzes lignin polymerization of NBD-CA in primary cell walls of
cotyledon epidermal cells. (G) Control wild-type cotyledon epidermal cells do not
incorporate NBD-CA into their cell walls. (H) Corresponding bright-field image of
control wild-type cotyledons with one cell outlined. Scale bars are 25 μm.
Figure 6. LACCASE4 localizes specifically to protoxylem TE secondary cell
walls.
(A) LACCASE4 (LAC4-mCherry) localization in endogenous protoxylem and (B)
metaxylem TE from primary control roots. (C) Median optical slice of an induced
protoxylem TE, in hypocotyl epidermis, after 18 hours of VND7-VP16-GR induction
showing LAC4-mCherry localization, specifically in developing secondary cell walls.
(D) Higher magnification of developing secondary cell wall outlined in C. (E)
Maximum projection image showing that LAC4-mCherry localization persists in
secondary cell walls after extended exposure to DEX and programed cell death. (F)
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
20
Immunolocalization of xylan (LM10 antibody) on a sectioned differentiating
protoxylem TE showing identical deposition patterns as observed for LAC4-mCherry.
(G) Higher magnification of the developing secondary cell wall outlined in F. Scale
bars are 5 μm in A,B and 12 μm in C-F.
Supplemental Figure 1. Phenotypic characterization of Pro35S:miRNA C4H
plants.
(A,B) Cross sections of the inflorescence stem base in wild-type and (C,D)
Pro35S:miRNA C4H plants showing severely reduced UV autofluorescence of lignin
compared to wild type. Toluidine blue stained sections show collapsed xylem vessels
in Pro35S:miRNA C4H genotypes (arrows in C) compared to (B) wild type. (E,F)
Segregating population of Pro35S:miRNA C4H plants that are impaired in generating
pigmented phenylpropanoids (anthocyanins) similar to those observed in c4h mutants
(Ruegger and Chapple, 2001). Seedling lethal primary Pro35S:miRNA C4H
transformants are also observed but cannot be propagated (inset E). Abnormal
swelling of inflorescence nodes (arrow in G), decreased pathogen resistance
(arrowheads in G), and flower sterility (H,I) are all observed in previously described
c4h mutants (Schilmiller et al., 2009).
Supplemental Figure 2. CINNAMATE-4-HYDROXYLASE (C4H-GFP)
localization and ER structure during VND7-induced protoxylem TE
differentiation.
Maximum projection images of z-stacks showing C4H-GFP and RFP:HDEL
localization in cortical ER and nuclear envelope (n). (A) The ER marker RFP-HDEL
shows a reticulate pattern in uninduced VND7-VP16-GR hypocotyl epidermal cells,
which also contain labeled ER bodies (asterisk). (B) Localization of C4H-YFP in the
reticulate ER network and ER bodies in uninduced controls. (C) Colocalization of A
and B, indicating RFP-HDEL localization to the ER lumen (asterisk), and C4H-YFP
signal in the ER membrane (arrow). (D) C4H-GFP localization, 18 h and (E) 36 h
following induction of VND7-VP16-GR in hypocotyl epidermal cells
transdifferentiated into protoxylem TEs. (F) RFP:HDEL localization, 18 h and (G) 36
h following induction of VND7-VP16-GR. Arrowheads in D-G identify the location of
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
21
two adjacent secondary cell wall domains whereas arrows show transition to laminar
ER morphology. Scale bars are 12 μm.
Supplemental Figure 3. Complementation of irregular xylem phenotypes of
ccr1irx4 and lac4 lac17 mutants with GFP or mCherry fusion proteins.
Toluidine blue stained sections from Arabidopsis inflorescence stems show collapsed
xylem vessels (arrows) in ccr1irx4 and lac4 lac17 mutants as indicated. Open
morphology of xylem vessels (arrowheads) is restored in ccr1irx4 and lac4lac17
mutants by ProUBQ10:CCR1-GFP and ProLAC4:LAC4-mCherry respectively.
Supplemental Figure 4. ABC transporters are evenly localized in plasma
membranes during protoxylem TE differentiation in VND7-VP16-GR seedlings.
Maximum projection images of hypocotyl epidermal cells undergoing protoxylem
TE differentiation showing uniform plasma membrane localization of (A) YFPABCG11, (B) ABCB11-GFP, and (C) GFP-ABCG33 over primary and secondary
wall domains. Insets show a magnified single optical slice from each image panel.
Scale bars are 15 μm.
Supplemental Figure 5. C4H-YFP and YFP-ABCG11 localization during
protoxylem programmed cell death of transdifferentiated Arabidopsis epidermal
cells.
(A,B) Fragmentation of C4H-YFP signal to unknown structures and (C) implosion of
YFP-ABCG11 labeled plasma membranes, during programmed cell death of VND7induced TEs. Scale bars are 15 μm.
Supplemental Figure 6. LACCASE17 localizes specifically to protoxylem TE
secondary cell walls.
(A) LACCASE17 (LAC17-mCherry) localization in endogenous protoxylem TEs
from cotyledons. (B) Maximum projection image and (C) corresponding median
optical slice, showing LAC17-mCherry localization in secondary cell walls of
hypocotyl epidermal cells that have differentiation into protoxylem TEs in VND7VP16-GR seedlings. Scale bars are 12 μm.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
22
Supplemental Movie 1. Z-stack animation of YFP-ABCG11 localization in
numerous plasma membrane derived vesicles during protoxylem TE
programmed cell death.
References
Alejandro, S., Lee, Y., Tohge, T., Sudre, D., Osorio, S., Park, J., Bovet, L.,
Geldner, N., Fernie, A.R., and Martinoia, E. (2012). AtABCG29 is a
monolignol transporter involved in lignin biosynthesis. Current Biology 22,
1207-1212.
Anderson, C.T., Carroll, A., Akhmetova, L., and Somerville, C. (2010). Real-time
imaging of cellulose reorientation during cell wall expansion in Arabidopsis
roots. Plant Physiology 152, 787-796.
Bassard, J.E., Richert, L., Geerinck, J., Renault, H., Duval, F., Ullmann, P.,
Schmitt, M., Meyer, E., Mutterer, J., Boerjan, W., De Jaeger, G., Mely,
Y., Goossens, A., and Werck-Reichhart, D. (2012). Protein-protein and
protein-membrane associations in the lignin pathway. Plant Cell 24, 44654482.
Berthet, S., Demont-Caulet, N., Pollet, B., Bidzinski, P., Cezard, L., Le Bris, P.,
Borrega, N., Herve, J., Blondet, E., Balzergue, S., Lapierre, C., and
Jouanin, L. (2011). Disruption of LACCASE4 and 17 results in tissuespecific alterations to lignification of Arabidopsis thaliana stems. Plant Cell
23, 1124-1137.
Bird, D., Beisson, F., Brigham, A., Shin, J., Greer, S., Jetter, R., Kunst, L., Wu,
X., Yephremov, A., and Samuels, L. (2007). Characterization of Arabidopsis
ABCG11/WBC11, an ATP binding cassette (ABC) transporter that is required
for cuticular lipid secretion. Plant Journal 52, 485-498.
Bonawitz, N.D., and Chapple, C. (2010). The genetics of lignin biosynthesis:
connecting genotype to phenotype. Annu Rev Genet 44, 337-363.
Chapelle, A., Morreel, K., Vanholme, R., Le-Bris, P., Morin, H., Lapierre, C.,
Boerjan, W., Jouanin, L., and Demont-Caulet, N. (2012). Impact of the
absence of stem-specific beta-glucosidases on lignin and monolignols. Plant
Physiology 160, 1204-1217.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
23
Chen, H.C., Li, Q., Shuford, C.M., Liu, J., Muddiman, D.C., Sederoff, R.R., and
Chiang, V.L. (2011a). Membrane protein complexes catalyze both 4- and 3hydroxylation of cinnamic acid derivatives in monolignol biosynthesis. Proc
Natl Acad Sci USA, 108, 21253-21258.
Curtis, M.D., and Grossniklaus, U. (2003). A gateway cloning vector set for highthroughput functional analysis of genes in planta. Plant Physiology 133, 462469.
Ehlting, J., Mattheus, N., Aeschliman, D.S., Li, E., Hamberger, B., Cullis, I.F.,
Zhuang, J., Kaneda, M., Mansfield, S.D., Samuels, L., Ritland, K., Ellis,
B.E., Bohlmann, J., and Douglas, C.J. (2005). Global transcript profiling of
primary stems from Arabidopsis thaliana identifies candidate genes for
missing links in lignin biosynthesis and transcriptional regulators of fiber
differentiation. Plant Journal 42, 618-640.
Esau, K. (1965). Vascular differentiation in plants. (New York: Holt, Rinehart and
Winston).
Grefen, C., Donald, N., Hashimoto, K., Kudla, J., Schumacher, K., and Blatt,
M.R. (2010). A ubiquitin-10 promoter-based vector set for fluorescent protein
tagging facilitates temporal stability and native protein distribution in transient
and stable expression studies. Plant Journal 64, 355-365.
Hosokawa, M., Suzuki, S., Umezawa, T., and Sato, Y. (2001). Progress of
lignification mediated by intercellular transportation of monolignols during
tracheary element differentiation of isolated Zinnia mesophyll cells. Plant &
Cell Physiology 42, 959-968.
Jones, L., Ennos, A.R., and Turner, S.R. (2001). Cloning and characterization of
irregular xylem4 (irx4): a severely lignin-deficient mutant of Arabidopsis.
Plant Journal 26, 205-216.
Kaneda, M., Schuetz, M., Lin, B.S., Chanis, C., Hamberger, B., Western, T.L.,
Ehlting, J., and Samuels, A.L. (2011). ABC transporters coordinately
expressed during lignification of Arabidopsis stems include a set of ABCBs
associated with auxin transport. Journal of Experimental Botany 62, 20632077
Kubo, M., Udagawa, M., Nishikubo, N., Horiguchi, G., Yamaguchi, M., Ito, J.,
Mimura, T., Fukuda, H., and Demura, T. (2005). Transcription switches for
protoxylem and metaxylem vessel formation. Gene Dev 19, 1855-1860.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
24
Lee, Y., Rubio, M.C., Alassimone, J., and Geldner, N. (2013). A mechanism for
localized lignin deposition in the endodermis. Cell 153, 402-412.
Liu, C.J. (2012). Deciphering the enigma of lignification: precursor transport,
oxidation, and the topochemistry of lignin assembly. Molecular Plant 5, 304317.
McCartney, L., Marcus, S.E., and Knox, J.P. (2005). Monoclonal antibodies to
plant cell wall xylans and arabinoxylans. J Histochem Cytochem 53, 543-546.
Miao, Y.C., and Liu, C.J. (2010). ATP-binding cassette-like transporters are
involved in the transport of lignin precursors across plasma and vacuolar
membranes. Proc Natl Acad Sci USA, 107, 22728-22733.
Nelson, B.K., Cai, X., and Nebenfuhr, A. (2007). A multicolored set of in vivo
organelle markers for co-localization studies in Arabidopsis and other plants.
Plant Journal 51, 1126-1136.
Ohashi-Ito, K., Oda, Y., and Fukuda, H. (2010). Arabidopsis VASCULARRELATED NAC-DOMAIN6 directly regulates the genes that govern
programmed cell death and secondary wall formation during xylem
differentiation. Plant Cell 22, 3461-3473.
Pesquet, E., Zhang, B., Gorzsas, A., Puhakainen, T., Serk, H., Escamez, S.,
Barbier, O., Gerber, L., Courtois-Moreau, C., Alatalo, E., Paulin, L.,
Kangasjarvi, J., Sundberg, B., Goffner, D., and Tuominen, H. (2013).
Non-cell-autonomous postmortem lignification of tracheary elements in
Zinnia elegans. Plant Cell 25, 1314-1328.
Ro, D.K., Mah, N., Ellis, B.E., and Douglas, C.J. (2001). Functional
characterization and subcellular localization of poplar (Populus trichocarpa x
Populus deltoides) cinnamate 4-hydroxylase. Plant Physiology 126, 317-329.
Roppolo, D., De Rybel, B., Tendon, V.D., Pfister, A., Alassimone, J., Vermeer,
J.E., Yamazaki, M., Stierhof, Y.D., Beeckman, T., and Geldner, N. (2011).
A novel protein family mediates Casparian strip formation in the endodermis.
Nature 473, 380-383.
Ruegger, M., and Chapple, C. (2001). Mutations that reduce sinapoylmalate
accumulation in Arabidopsis thaliana define loci with diverse roles in
phenylpropanoid metabolism. Genetics 159, 1741-1749.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
25
Pumplin, N.1., Zhang, X., Noar, R.D., and Harrison, M.J. (2012). Polar
localization of a symbiosis-specific phosphate transporter is mediated by a
transient reorientation of secretion. Proc Natl Acad Sci USA, 109 665-672.
Sato, Y., Demura, T., Yamawaki, K., Inoue, Y., Sato, S., Sugiyama, M., and
Fukuda, H. (2006). Isolation and characterization of a novel peroxidase gene
ZPO-C whose expression and function are closely associated with lignification
during tracheary element differentiation. Plant & Cell Physiology 47, 493-503.
Schilmiller, A.L., Stout, J., Weng, J.K., Humphreys, J., Ruegger, M.O., and
Chapple, C. (2009). Mutations in the cinnamate 4-hydroxylase gene impact
metabolism, growth and development in Arabidopsis. Plant Journal 60, 771782.
Schwab, R., Ossowski, S., Riester, M., Warthmann, N., and Weigel, D. (2006).
Highly specific gene silencing by artificial microRNAs in Arabidopsis. Plant
Cell 18, 1121-1133.
Smith R.A., Schuetz M., Roach M., Mansfield S.D., Ellis B., and Samuels A.L.
(2013). Neighboring parenchyma cells can contribute to Arabidopsis xylem
lignification, while lignification of interfascicular fibers is cell autonomous.
Plant Cell 25, 3988-99.
Thevenin, J., Pollet, B., Letarnec, B., Saulnier, L., Gissot, L., Maia-Grondard,
A., Lapierre, C., and Jouanin, L. (2011). The simultaneous repression of
CCR and CAD, two enzymes of the lignin biosynthetic pathway, results in
sterility and dwarfism in Arabidopsis thaliana. Molecular Plant 4, 70-82.
Tobimatsu, Y., Davidson, C.L., Grabber, J.H., and Ralph, J. (2011).
Fluorescence-tagged monolignols: synthesis, and application to studying in
vitro lignification. Biomacromolecules 12, 1752-1761.
Tobimatsu, Y., Wagner, A., Donaldson, L., Mitra, P., Niculaes, C., Dima, O.,
Kim, J.I., Anderson, N., Loque, D., Boerjan, W., Chapple, C., and Ralph,
J. (2013). Visualization of plant cell wall lignification using fluorescencetagged monolignols. Plant Journal 76, 357-366.
Tokunaga, N., Sakakibara, N., Umezawa, T., Ito, Y., Fukuda, H., and Sato, Y.
(2005). Involvement of extracellular dilignols in lignification during tracheary
element differentiation of isolated Zinnia mesophyll cells. Plant & Cell
Physiology 46, 224-232.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
26
Tsuyama, T., Kawai, R., Shitan, N., Matoh, T., Sugiyama, J., Yoshinaga, A.,
Takabe, K., Fujita, M., and Yazaki, K. (2013). Proton-dependent coniferin
transport, a common major transport event in differentiating xylem tissue of
woody plants. Plant Physiology 162, 918-926.
Vanholme, R., Morreel, K., Darrah, C., Oyarce, P., Grabber, J.H., Ralph, J., and
Boerjan, W. (2012). Metabolic engineering of novel lignin in biomass crops.
New Phytologist 196, 978-1000.
Wightman, R., and Turner, S.R. (2008). The roles of the cytoskeleton during
cellulose deposition at the secondary cell wall. Plant Journal 54, 794-805.
Yamaguchi, M., Mitsuda, N., Ohtani, M., Ohme-Takagi, M., Kato, K., and
Demura, T. (2011). VASCULAR-RELATED NAC-DOMAIN7 directly
regulates the expression of a broad range of genes for xylem vessel formation.
The Plant Journal 66, 579-590.
Yamaguchi, M., Goue, N., Igarashi, H., Ohtani, M., Nakano, Y., Mortimer, J.C.,
Nishikubo, N., Kubo, M., Katayama, Y., Kakegawa, K., Dupree, P., and
Demura, T. (2010). VASCULAR-RELATED NAC-DOMAIN6 and
VASCULAR-RELATED NAC-DOMAIN7 effectively induce
transdifferentiation into xylem vessel elements under control of an induction
system. Plant Physiology 153, 906-914.
Zhao Q., Nakashima, J., Chen, F., Yin, Y., Fu, C., Yun, J., Shao, H., Wang, X.,
Wang, Z.Y., Dixon, R.A. (2013). Laccase is necessary and nonredundant
with peroxidase for lignin polymerization during vascular development in
Arabidopsis. Plant Cell 25, 3976-3987.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
A Wild type C B D VND7-­‐GR VND7-­‐GR + DEX E VND7-­‐GR; miRNA C4H VND7-­‐GR; miRNA CCR1 Figure 1. Lignin autofluorescence in secondary cell walls of endogenous and VND7induced Arabidopsis seedling TEs.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
A B C4H-­‐GFP C scw er n n G scw m er m D CCR1-­‐GFP E cytGFP F CCR1-­‐GFP G cytGFP H CCR1-­‐GFP I cytGFP Figure 2. CINNAMATE-4-HYDROXYLASE (C4H-GFP) and CINNAMOYL-CoAREDUCTASE (CCR1-GFP) are localized between secondary cell walls domains during
VND7-induced protoxylem TE differentiation.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
A GFP-­‐ABCG33 B Cellulose S4B stain C GFP-­‐ABCG33 D GFP-­‐ABCG29 Figure 3. ABC transporters are evenly localized in plasma membranes during
protoxylem TE differentiation in VND7-VP16-GR seedlings.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
A NBD – CA D NBD – CA B C merged Cellulose S4B stain E Cellulose S4B stain F merged Figure 4. Mobility of monolignols in the apoplast: fluorescently tagged monolignols are
specifically incorporated into the secondary cell walls of VND7-induced protoxylem
TEs.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
A VND7-­‐GR B VND7-­‐GR Figure 5. LACCASEs are necessary
and sufficient to direct lignin
polymerization in Arabidopsis cell
wall domains.
i ii iii NBD – CA C lac4 lac17; VND7-­‐GR D i ii iii NBD – CA E Pro35S:LAC4 NBD – CA G lac4 lac17; VND7-­‐GR F Pro35S:LAC17 NBD – CA Wild type H Wild type NBD – CA Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.
A LAC4-­‐ mCHERRY B E LAC4-­‐mCHERRY LAC4-­‐ C mCHERRY F LAC4-­‐mCHERRY D LM10-­‐xylan G Figure 6. LACCASE4 localizes specifically to protoxylem TE secondary cell walls.
Downloaded from on June 17, 2017 - Published by www.plantphysiol.org
Copyright © 2014 American Society of Plant Biologists. All rights reserved.