new regulators of xylem lignification in arabidopsis
TRANSCRIPT
New regulators of xylem
lignification in Arabidopsis
Bernadette Sztojka
Umeå Plant Science Centre
Department of Plant Physiology
Umeå 2020
This work is protected by the Swedish Copyright Legislation (Act 1960:729)
Dissertation for PhD
ISBN (printed version): 978-91-7855-428-7
ISBN (digital version): 978-91-7855-429-4
Cover design: Schematic representation of an Arabidopsis thaliana plant; Illustration of
transverse sections of the xylem in the base of the stem and in the hypocotyl of
Arabidopsis thaliana; Schematic view of xylem cells. (by Bernadette Sztojka)
Electronic version available at: http://umu.diva-portal.org/
Printed by: KBC Service Centre, Umeå University
Umeå, Sweden 2020
Szüleimnek.
To my parents.
i
Table of Contents
Table of Contents i
Abstract iii
Sammanfattning iv
Abbreviations v
List of publications ix
Introduction 1
1. The importance of wood 1
2. Xylem 2
2.1. Vascular cambium 2
Vascular cambium and secondary growth 2
Vascular cambium specification and maintenance 2
2.2. Arabidopsis as a model to study the basic molecular mechanisms of
secondary growth 6
2.3. The function of the xylem 7
2.4. Different cell types building up the xylem 8
Tracheids and vessel elements 8
Fibers 9
Parenchyma cells and ray cells 10
3. Secondary cell wall 11
3.1. Cellulose 11
3.2. Hemicellulose 13
3.3. Cell wall proteins 14
3.4. Lignin 15
The evolution of lignin 15
Major monolignols 15
Distribution of lignin in different species and cell types 16
The function of lignin 16
3.4.1. Biosynthesis of the lignin monomers 16
ii
3.4.1.a. The lignin biosynthetic pathway 16
3.4.1.b. Regulation of lignin biosynthesis 18
Transcriptional regulation 19
Chromatin-level regulation 22
Circadian regulation 22
Additional levels of regulation 23
3.4.2. Transport of the lignin monomers 23
3.4.3. Lignin polymerization 25
Laccases and peroxidases 25
Secondary substrates 27
Assembly of the polymer 28
3.4.4. Cell-autonomous vs. non-cell-autonomous lignification 29
3.4.5. Lignin as a resource for biotechnological applications 31
Research aims and objectives 34
Results and discussion 35
PIRIN2 is a non-cell-autonomous regulator of S-type lignin accumulation 35
HISTONE MONOUBIQUITINATION2 (HUB2) affects the lignin
composition of xylem vessels 37
PIB is a potential modulator of the diurnal timing of lignin biosynthesis 39
Conclusions and perspectives 41
Acknowledgements 43
References 45
iii
Abstract
The ability of land plants to grow upright, bear their own weight and withstand
adverse environmental conditions is largely dependent on the secondary
xylem tissues of the stem. The xylem cells acquire thick secondary cell walls
which are composed of cellulose, hemicellulose and lignin. The chemical
structure of lignin renders the secondary cell wall rigid and waterproof,
facilitating the transport of water and solutes through the vascular system.
Lignin is a polyphenolic polymer composed of three different types of lignin
units, guaiacyl (G), syringyl (S) and p-hydroxyphenyl (H), derived from the
coniferyl, sinapyl and p-coumaryl alcohol, respectively. Lignin biosynthesis,
monolignol transport and lignin polymerization (collectively called as
”lignification”) are controlled by numerous transcription factors and other
regulators.
This thesis work uncovers three novel regulators of lignification in the
secondary xylem tissues of Arabidopsis (Arabidopsis thaliana) stem and
hypocotyl. The cupin domain containing protein PIRIN2 (PRN2) suppresses
S-type lignin accumulation. PRN2 functions in a non-cell-autonomous
fashion: it is expressed in the cells next to the xylem vessel elements, but
affects the lignin composition of the vessel and fiber cell walls of the
neighbouring cells. Two protein interactors of PRN2 are characterized here in
connection to lignification. Opposite to the function of PRN2, the chromatin-
modifying protein HISTONE MONOUBIQUITINATION2 (HUB2)
promotes S-type lignin deposition. In line with this, PRN2 and HUB2
antagonistically regulate the expression of FERULATE-5-HYDROXYLASE1
which encodes the key S-type lignin-biosynthetic enzyme. Possibly, PRN2
antagonizes the S-lignin promoting function of HUB2 to ensure that the cell
walls of the vessel elements get enriched in G-type lignin. Finally,
identification of a potential diurnal modulator of lignin biosynthesis is
described in this work. The PRN2-interacting basic helix-loop-helix
transcription factor (PIB) does not influence the lignin content or composition
of the secondary cell walls. However, PIB affects the diurnal expression
pattern and promoter activity of some lignin-biosynthetic genes. Altogether,
PRN2, HUB2 and PIB highlight the importance of intercellular co-operation
in lignification, and uncover novel regulatory aspects of this process.
iv
Sammanfattning
Växternas förmåga att växa upprätt, bära sin egen vikt och tåla ogynnsamma
miljöförhållanden styrs till stor del av stammens vaskulär vävnad som består
av floem- och xylemceller. Xylemceller ackumulerar tjocka sekundära
cellväggar som är sammansatta av cellulosa, hemicellulosa och lignin. Den
kemiska strukturen hos lignin gör cellväggen styv och vattentät, vilket
möjliggör transport av vatten och näringsämnen genom det vaskulära
systemet. Lignin är en polyfenol som består av tre olika typer av lignin
enheter: guaiacyl (G), syringyl (S) och p-hydroxyfenol (H). Lignin biosyntes,
monolignol transport och lignin polymerisering (kollektivt kallad som
"lignifiering") styrs av ett flertal transkriptionsfaktorer och andra regulatorer.
Denna avhandling avslöjar tre nya regulatorer av lignifiering i xylemvävnader
av Arabidopsis (Arabidopsis thaliana) stam och hypokotyl. Genetiska och
kemiska analyser avslöjade ett nytt protein, PIRIN2 (PRN2), som dämpar S-
typ lignin ackumulering. PRN2 fungerar på ett icke-cell-autonomt sätt: det
uttrycks i cellerna bredvid xylemkärlselement, men påverkar
ligninsammansättningen hos de angränsande kärlens cellväggar. I
avhandlingsarbetet identifierades och karakteriserades också två andra
proteiner som interagerar med PRN2 i lignifiering. I motsats till funktionen
av PRN2, det kromatin-modifierande HISTONE
MONOUBIQUITINATION2 (HUB2) främjar S-typ lignin ackumulering. I
linje med detta reglerar PRN2 och HUB2 antagonistiskt uttrycket av
FERULATE-5-HYDROXYLASE1 som kontrollerar biosyntes av S-typ lignin.
Det verkar troligt att PRN2 motverkar S-lignin främjande funktion av HUB2
för att säkerställa anrikning av G-typ lignin i kärlelementens cellväggar.
Slutligen beskrivs här en helix-loop-helix transkriptionsfaktor (PIB) som
potentiellt modulerar lignin biosyntes enligt dygnsrytmen. PIB påverkar inte
lignin innehållet eller sammansättningen av de sekundära cellväggarna men
påverkar tajmningen av genuttrycket och promotoraktiviteten hos vissa
lignin-biosyntetiska gener. Sammantaget belyser PRN2, HUB2 och PIB
vikten av intercellulärt samarbete i lignifiering och avslöjar nya
regleringsaspekter av denna process.
v
Abbreviations
ABC ATP-binding cassette
AGP Arabinogalactan-rich glycoproteins
AHP6 ARABIDOPSIS HISTIDINE PHOSPHOTRANSFER
PROTEIN6
ANT AINTEGUMENTA
Arabidopsis Arabidopsis thaliana
ARF Auxin response factor
ARR Arabidopsis response regulator
BES1 BRI1-EMS SUPPRESSOR1
BIL1 BRASSINOSTEROID-INSENSITIVE2-LIKE1
CAD Cinnamyl alcohol dehydrogenase
CCoAOMT Caffeoyl-CoA O-methyltransferase
CCR Cinnamoyl-CoA reductase
CLE CLAVATA3/EMBRYO SURROUNDING REGION-
related
COMT Caffeic acid O-methyltransferase
CSC Cellulose synthase protein complex
C3H P-coumarate 3-hydroxylase
C4H Cinnamate 4-hydroxylase
4CL 4-coumarate:CoA ligase
C-type lignin Caffeyl alcohol
ER Endoplasmic reticulum
ESB1/DIR10 ENHANCED SUBERIN1/DIRIGENT10
FAO Food and Agriculture Organization of the United Nations
FLA Fasciclin-like arabinogalactan-protein
F5H Ferulate 5-hydroxylase
GlcA Glucuronic acid
vi
Abbreviations (2/4)
GRP Glycine-rich glycoproteins
GSK3 Glycogen synthase kinase
GT43 Glycosyltransferase family 43
GUX Glucuronic acid substitution of xylan
GXMT Glucuronoxylan methyltransferase
G-type lignin Guaiacyl-type lignin
HCT Hydroxycinnamoyltransferase
HD-ZIP III Class III homeodomain-leucine zipper
HRGP Hydroxyproline-rich glycoproteins
H3K4me3 Histone H3 lysine K4 methylation
H3K27me3 Histone H3 lysine K27 methylation
H2O2 Hydrogen peroxide
H-type lignin P-hydroxyphenyl-type lignin
IRX Irregular xylem
IRX-L Irregular xylem-like
KFB Kelch-repeat F-box
KNOX Knotted-like homeobox
LAC Laccase
LHW LONESOME HIGHWAY
LOG LONELY GUY
MED Mediator
[Me]GlcA Methyl-glucuronic acid
MOL1 MORE LATERAL GROWTH1
MP MONOPTEROS
MYB Myeloblastosis
vii
Abbreviations (3/4)
NAC NAM, ATAF1/2, CUC2
NADPH oxidase Nicotineamide adenine dinucleotide phosphate hydrogen
oxidase
NST NAC SECONDARY WALL THICKENING
PROMOTING FACTOR
O2 Molecular oxygen
PAL Phenylalanine ammonia-lyase
PCW Primary cell wall
PEAR PEAR PHLOEM EARLY DOF1
PIN PIN-FORMED
PRP Proline-rich glycoproteins
PRX Peroxidase
PXY PHLOEM INTERCALATED WITH XYLEM
TDR TRACHEARY ELEMENT DIFFERENTIATION
INHIBITORY FACTOR RECEPTOR
RUL1 REDUCED IN LATERAL GROWTH1
RWA REDUCED WALL ACETYLATION
SCW Secondary cell wall
SERKs SOMATIC EMBRYOGENESIS RECEPTOR KINASEs
SMRE Secondary wall MYB-responsive element
SMXL5 SUPPRESSOR OF MAX2 1-LIKE5
SNBE Secondary wall NAC-binding element
SND SECONDARY WALL-ASSOCIATED NAC DOMAIN
SOD Superoxide dismutase
S-type lignin Syringyl-type lignin
TDIF TRACHEARY ELEMENT DIFFERENTIATION
INHIBITORY FACTOR
viii
Abbreviations (4/4)
TE Tracheary element
TED Tracheary element differentiation-related
TF Transcription factor
TMO TARGET OF MONOPTEROS
T5L1 TMO5-LIKE1
VND VASCULAR RELATED NAC DOMAIN
VNI2 VND-INTERACTING2
VNS VND, NST/SND, and SMB-related protein
WOX WUSCHEL-related homeobox
XND1 XYLEM NAC DOMAIN1
YUC4 YUCCA4
Other abbreviations are explained in the text, when they first appear.
ix
List of publications
Paper I
Zhang B*, Sztojka B*, Escamez S, Vanholme R, Hedenström M, Wang Y,
Turumtay H, Gorzsás A, Boerjan W, Tuominen H. 2020. PIRIN2 suppresses S-
type lignin accumulation in a noncell-autonomous manner in Arabidopsis xylem
elements. New Phytologist 225: 1923-1935.
* These authors contributed equally.
Paper II
Zhang B*, Sztojka B*, Seyfferth C, Escamez S, Miskolczi P, Chantreau M, Bakó
L, Delhomme N, Gorzsás A, Bhalerao RP, Tuominen H. 2020. The chromatin-
modifying protein HUB2 is involved in the regulation of lignin composition in
xylem vessels. Journal of Experimental Botany 71: 5484-5494.
* These authors contributed equally.
Paper III
Sztojka B, Escamez S, Seyfferth C, Wang Y, Zhang B, Gorzsás A, Demedts B,
Boerjan W, Tuominen H. (manuscript). PIB – a potential modulator of
lignification in Arabidopsis.
Publication not included in the PhD thesis
Escamez S, André D, Sztojka B, Bollhöner B, Hall H, Berthet B, Voß U, Lers A,
Maizel A, Andersson M, Bennett M, Tuominen H. 2020. Cell death in cells
overlying lateral root primordia facilitates organ growth in Arabidopsis. Current
Biology 30: 455-464.
x
Author’s contribution
Paper I: Obtained plant material and prepared samples for FT-IR and Raman
microspectroscopy, took part in the data analysis and interpretation. Performed
GUS staining of the Populus tremula x tremuloides proPttPRN2::GUS lines.
Obtained plant material for the PRN2 overexpression lines and prepared samples
for pyrolysis-GC/MS, performed the data analysis. Contributed to data
interpretation, the writing and formatting of the manuscript.
Paper II: Obtained plant material for the PRN2OE6 and hub2-1 sample set,
prepared samples for pyrolysis-GC/MS, performed the data analysis. Obtained
plant material and prepared samples for FT-IR and Raman microspectroscopy,
took part in the data analysis and interpretation. Performed the greenhouse
experiment, collected plant material, and prepared RNA for RNA-sequencing,
took part in the data analysis. Contributed to data interpretation, the writing and
formatting of the manuscript.
Paper III: Cloned the proPIB:GUS constructs, generated and selected transgenic
lines, and performed GUS staining. Cloned the proPIB::PIB:mCherry construct,
generated and selected transgenic lines. Performed DAPI staining and confocal
microscopic imaging of the proPIB::PIB:mCherry seedlings. Obtained plant
material, prepared samples for pyrolysis-GC/MS, performed the data analysis.
Performed the greenhouse experiment, collected plant material, and prepared
RNA for RNA-sequencing, took part in the data analysis. Performed the diurnal
experiment, analyzed the data. Assessed the gametophytic lethality of the pib-1
pib-like-1 double mutants. Obtained plant material and determined the acetyl
bromide soluble lignin content. Obtained plant material and prepared samples for
FT-IR microspectroscopy, took part in the data analysis. Prepared samples for the
monosaccharide analysis by acidic methanolysis and TMS derivatization,
performed the data analysis. Carried out the growth assay of inflorescence stems.
Performed two of the experiments to assay root elongation in response to different
sucrose concentrations. Wrote the manuscript with contribution from the co-
authors.
1
Introduction
1. The importance of wood
Wood is an essential, renewable bioresource with massive ecological and
economic importance. As of 2015, 30,6 % of the global land area is covered by
forest (UN, 2017). Wood is utilized in numerous ways, for example as raw
material for construction, furnishings, paper and packaging, toolmaking, and art
(Meents et al., 2018). According to data from the Food and Agriculture
Organization of the United Nations (FAO), one third of the world’s population,
more than 2.4 billion inhabitants, use wood as energy source in their daily lives
(FAO, 2014; 2017A). Indigent rural populations worldwide base their livelihood
on wood and non-wood forest products. Forest products constitute 40 % of the
global renewable energy, the same proportion as wind, solar and hydroelectric
power combined (FAO, 2017B). Furthermore, in the era of accelerating climate
change, the crucial function of forest as a carbon sink cannot be emphasized
enough. Trees act as a carbon reservoir, by absorbing about 2 billion tonnes of
carbon dioxide yearly (FAO, 2018).
While forests have beneficial role in binding greenhouse gases, deforestation
scores as the second prominent cause of climate change. To mitigate the harmful
effect of deforestation, sustainable forest management as well as biotechnological
developments for improved tree performance offer solutions. The combination of
natural variation, molecular and genetic tools can generate trees with beneficial
properties for the forest industry. Such properties may be enhanced growth,
climate resilience, wood characteristics which allow more efficient processing
and thus lessen the environmental repercussions. Hence, the speedy development
of basic and applied biology tools which address the abovementioned features is
crucial to facilitate forest improvement. The Paris Climate Agreement (2015)
established the shared responsibility in utilizing forests for the good of the
climate, and biotechnological innovations can help in addressing this.
2
2. Xylem
2.1. Vascular cambium
Vascular cambium and secondary growth
Secondary (lateral) growth is a developmental process characterized by the
activity of the post-embryonic meristems, vascular cambium and phellogen,
which generate cylindrical tissue layers in the plant body. These two meristems
produce tissues in a bidirectional manner, a feature which is strictly orchestrated
by molecular and hormonal factors (Ragni and Greb, 2018).
The vascular cambium is responsible for the secondary growth of stems and roots,
and the formation of the woody tissue in dicots and gymnosperms. Two types of
vascular stem cells exist in the vascular cambium: the long and narrow fusiform
initials, and the short and small ray initials. Through periclinal cell divisions
(parallel to the surface) fusiform initials give rise to the specialized cell pool for
xylem and phloem, the vital tissues for mechanical support and long-distance
transport. The ray initials are the precursors of ray parenchyma cells, which form
the storage and transport network between phloem and xylem (Fosket, 1994;
Fischer et al., 2019).
The phellogen, also called cork cambium, generates phelloderm inwards and cork
outwards. These tissues have protective role against environmental stresses, like
water loss, biotic and abiotic attacks (Campilho et al., 2020).
Vascular cambium specification and maintenance
Whether xylem and phloem precursors within the vascular cambium are derived
from one or more radial stem cell layers, has been long debated. The 150-year-
old uniseriate cambium organization theory (Sanio, 1873) was supported recently
by lineage-tracing experiments, revealing that the vascular cambium consists of
a single layer of bifacial stem cells (Bossinger and Spokevicius, 2018; Smetana
et al., 2019; Shi et al., 2019). Furthermore, the work of Smetana et al. (2019)
demonstrated that the cambial cell division and differentiation programs are
orchestrated by the meristem organizer cells that are located adjacent to the
cambial stem cells. Local auxin response maxima promote the xylem identity and
quiescence of the organizer cells by the upregulation of CLASS III
HOMEODOMAIN-LEUCINE ZIPPER (HD-ZIP III) transcription factors.
Positive feedback regulation between auxin, AUXIN RESPONSE FACTORS
(ARF) and HD-ZIP III transcription factors assures vascular cambium
maintenance. The stem cell organizer of the vascular cambium is dynamic; the
3
organizer differentiates into a xylem vessel and an adjacent cambial cell becomes
the new organizer. Members of the WUSCHEL RELATED HOMEOBOX
(WOX) transcription factor family mark all three stem cell organizers;
WUSCHEL in the shoot apical meristem, WOX5 in the root meristem and WOX4
in the vascular cambium (Mayer et al., 1998; Sarkar et al., 2007; Smetana et al,
2019), and they are all downstream of CLE peptide signaling (Schoof et al., 2000;
Stahl et al., 2009; Hirakawa et al., 2010).
Smetana et al. (2019) and Shi et al. (2019) dissected and defined the cambial
regions based on the activity of molecular markers. Bifacial stem cells are
characterized by the overlapping activity of WOX4, PHLOEM
INTERCALATED WITH XYLEM/TRACHEARY ELEMENT
DIFFERENTIATION INHIBITORY FACTOR RECEPTOR (PXY/ TDR),
AINTEGUMENTA (ANT) and SUPPRESSOR OF MAX2 1-LIKE5 (SMXL5).
The xylem cell precursors are marked by WOX4 and PXY, the stem cells by high
levels of ANT, while those destined for phloem development express SMXL5.
Furthermore, it was recently uncovered that a concentration gradient formed by
six mobile PEAR PHLOEM EARLY DOF1 (PEAR) transcription factors
demarks the procambial zone and promotes cell division. A negative feedback
loop exists between the cytokinin-inducible PEAR proteins and the xylem cell
fate-promoting HD-ZIP III transcription factors which are induced by auxin. This
regulatory network establishes the foundation of radial growth, restricting the
stem cell niche by transcriptional regulation and the control of protein movement
(Fig. 1) (Miyashima et al., 2019).
A fundamental mechanism regulating cambium activity and maintenance is the
receptor-ligand complex formed between PXY/TDR and the
CLAVATA3/EMBRYO SURROUNDING REGION-related 41
(CLE41)/CLE44 peptides (Hirakawa et al., 2008; Etchells and Turner, 2010).
CLE41 and CLE44 encode the 12-amino-acid-long peptide ligand called
TRACHEARY ELEMENT DIFFERENTIATION INHIBITORY FACTOR
(TDIF). The CLE peptides are expressed mainly in the phloem and they travel to
the dividing cambial cells and bind to PXY/TDR, a plasma membrane-bound
leucine-rich repeat receptor-like kinase (Fig. 1). SOMATIC EMBRYOGENESIS
RECEPTOR KINASEs (SERKs) are coreceptors in CLE41/TDIF-PXY/TDR
signaling (Zhang et al., 2016). Signaling pathways downstream of CLE41 and
PXY/TDR promote cambial cell division and cell fate maintenance, inhibit
differentiation of xylem cells and control vascular patterning (Hirakawa et al.,
4
2008; 2010; Whitford et al., 2008; Etchells and Turner, 2010; reviewed by
Fischer et al., 2019).
The CLE41-PXY/TDR interaction is crucial to promote procambial cell
proliferation, by enhancing WOX4/WOX14 expression (Hirakawa et al., 2010;
Etchells et al., 2013). Simultaneously, CLE41-PXY/TDR maintains homeostasis
between xylem and phloem production by the activation of the GLYCOGEN
SYNTHASE KINASE3 (GSK3) pathway and the brassinosteroid-dependent
signaling cascade (Kondo et al., 2014; Han et al., 2018). GSK3s negatively
regulate xylem differentiation by suppression of BRI1-EMS SUPPRESSOR1
(BES1) which itself promotes xylem cell fate (Kondo et al., 2014). One member
of the GSK3s, BRASSINOSTEROID-INSENSITIVE2-LIKE1 (BIL1), inhibits
cambial activity downstream of CLE41-PXY via ARF5/MONOPTEROS (MP)
(Han et al., 2018). ARF5/MP promotes the transition of stem cells to xylem cells
by suppressing cytokinin response through the negative regulators of cytokinin
signaling ARABIDOPSIS RESPONSE REGULATOR7 (ARR7)/ARR15, and
BIL1 enhances this by the phosphorylation of ARF5/MP. PXY blocks the BIL1-
ARF5-cytokinin pathway by inhibiting BIL1 and its negative regulation of
cambial activity (Han et al., 2018). Notably, BIL1 is key in cambium
maintenance since it connects peptide signaling with auxin and cytokinin
signaling. Despite the complex regulations encompassing vascular cambium
maintenance, Lebovka et al. (2020) demonstrated by computational modelling
that the CLE41-PXY complex is sufficient to define tissue organization in the
cambium.
Additionally, REDUCED IN LATERAL GROWTH1 (RUL1) and MORE
LATERAL GROWTH1 (MOL1) are two receptor-like kinases regulating
cambial cell proliferation independent of PXY, RUL1 promotes and MOL1
suppresses the process (Agustí et al., 2011; Gursanscky et al., 2016).
As mentioned above, cambium organization is regulated by hormonal cross talks.
Mutual inhibition takes place between auxin and cytokinin signaling, which is
essential to define the developing xylem and phloem regions (Bishopp et al.,
2011A; Mellor et al., 2017). High cytokinin levels promote auxin flow towards
the xylem via the PIN-FORMED (PIN) auxin efflux carriers (Bishopp et al.,
2011B). In contrast, in xylem auxin promotes the expression of the cytokinin
signaling inhibitor, ARABIDOPSIS HISTIDINE PHOSPHOTRANSFER
PROTEIN6 (AHP6) (Bishopp et al., 2011A). In general, high levels of auxin
promote xylem development, while elevated cytokinin signaling stimulates cell
proliferation (Mellor et al., 2017; Ruonala et al., 2017).
5
The auxin-dependent ARF3, ARF4 and ARF5/MP transcription factors have
distinct roles in cambium regulation. ARF3 and ARF4 stimulate cambium
activity from outside the stem cell region, while ARF5/MP promotes the
differentiation of stem cells to xylem cells by suppressing WOX4 expression and
activating xylem-related genes (Brackmann et al., 2018).
Figure 1. Regulatory pathways of the vascular cambium activity.
The CLE41/44 peptides are expressed in the phloem and they move to the cambium,
where they bind to the PXY receptor. CLE41-PXY promotes cambial proliferation
through WOX4/WOX14. In addition, CLE41-PXY maintains xylem-phloem
homeostasis by activating GSK3, which suppresses xylem differentiation through
BES1. Downstream of CLE41-PXY, BIL inhibits cambial activity via ARF5/MP.
ARF5/MP suppresses cytokinin signaling through ARR7/ARR15. The auxin-
dependent ARF3 and ARF4 promote cambium activity from outside the stem cell
region. RUL1 promotes cambial cell proliferation and MOL1 suppresses the process.
The negative feedback loop between the cytokinin-inducible PEAR proteins and the
auxin-inducible HD-ZIP III transcription factors restricts the stem cell niche by
transcriptional regulation and protein movement control. High cytokinin levels
stimulate auxin flow towards the xylem via the PINs. ARF5/MP connects cytokinin
and auxin signaling by targeting TMO5. The TMO5-LHW heterodimer induces
AHP6 expression, which inhibits cytokinin signaling thus promoting xylem
formation. The TMO5-LHW complex can promote cytokinin biosynthesis through
the upregulation of LOG3 and LOG4, and auxin biosynthesis via YUC4, resulting in
a positive feedback loop.
6
MP is an important connection between auxin and cytokinin, MP targets
TARGET OF MONOPTEROS3 (TMO3)/TMO5/TMO7 which are upstream of
the cytokinin signaling cascade (Schlereth et al., 2010). TMO5/TMO5-LIKE1
(T5L1) form heterodimers with LONESOME HIGHWAY (LHW) (De Rybel et
al., 2013) and induce the expression of AHP6 (Ohashi-Ito et al., 2014), which in
turn promotes xylem formation by inhibiting cytokinin signaling (Mähönen et al.,
2006). Strikingly, the TMO5-LHW transcription factor complex can also
promote cytokinin production by the upregulation of the biosynthetic genes,
LONELY GUY (LOG3) and LOG4, which leads to enhanced cell proliferation
(Ohashi-Ito et al., 2014; Vera-Sirera et al., 2015). A key transcriptional hub for
cell proliferation downstream of TMO5-LHW is the transcription factor DOF2.1
which together with its close homologs control procambial cell divisions by
regulating a subset of cytokinin-dependent genes (Smet et al., 2019).
Furthermore, the TMO5-LHW dimer is able to upregulate auxin biosynthesis
through YUCCA4 (YUC4), leading to a positive feedback loop since elevated
auxin levels maintain the expression of TMO5, LHW and their downstream
targets (Ohashi-Ito et al., 2019). On the other hand, a negative feedback loop,
involving the polyamine signaling molecule thermospermine, confines the
TMO5-LHW levels (Katayama et al., 2015; Vera-Sirera et al., 2015).
Such intricate control mechanisms are essential to initiate vascular development
and modulate that based on the needs of the indefinitely growing plant body and
the changing environment, without compromising the integrity of the organism.
2.2. Arabidopsis as a model to study the basic molecular
mechanisms of secondary growth
Arabidopsis thaliana (from here on “Arabidopsis”) is a small, herbaceous annual
plant, a native species of Western Eurasia, which has colonized habitats across
the globe with vast climatic and environmental fluctuations. Arabidopsis not only
colonized our geographical world, but also the world of plant science, becoming
the central model plant and reference organism. The first study of this species
was published more than a century ago in 1907 by the German botanist, Friedrich
Laibach (Laibach, 1907; Krämer, 2015). Since then, research fields from cell
biology to ecology have adopted Arabidopsis as a model plant thanks to the small
size of the plant, its small genome and all the available toolkits. Aware of the
limitations of Arabidopsis, we shall acknowledge that this model plant has
enabled the molecular understanding of key biological concepts. One such
concept is secondary growth. Despite its herbaceous nature, the root, hypocotyl
7
and stem of Arabidopsis undergo secondary growth, making it a simple and handy
model. The root and the stem display a secondary growth gradient, with only
primary growth present close to the apical meristems and extensive secondary
growth in the proximity of the rosette. In contrast, the secondary growth of the
hypocotyl is longitudinally quite uniform, forming a cylinder of wood similar to
angiosperm trees (reviewed by Ragni and Greb, 2018), which makes it an
attractive model for secondary xylem development (Chaffey et al., 2002). During
secondary growth, a continuous cambial ring develops in the root and hypocotyl,
producing xylem on the inner and phloem on the outer side. Initially the
secondary xylem consists of vessel elements and parenchyma cells (phase I). A
developmental shift towards xylem expansion takes place when floral transition
occurs, and the cell type repertoire broadens with the emergence of fiber cells
(phase II) (Chaffey et al., 2002). In the inflorescence stem, uniform vascular
cambium is formed at the very base of the stem where the fascicular cambia of
the vascular bundles are linked together by the interfascicular cambia (Altamura
et al., 2001). A wide array of molecular genetics findings from Arabidopsis have
been corroborated in tree species (reviewed by Barra-Jiménez and Ragni, 2017).
Some of the features which make Arabidopsis a popular model system, can
occasionally become disadvantageous. For example, the small size of the plant
can make dissection of specific xylem cell types difficult. Due to its annual
nature, long-term research of the same individual is limited. Arabidopsis has a
poor biomass yield compared to trees, limiting biomass studies and the
characterization of certain traits important for the forest industry, such as the
mechanical properties of wood and sugar release upon enzymatic pretreatment.
2.3. The function of the xylem
Early in the evolution of vascular plants the development of xylem and phloem
enabled long-distance transport, leading to the gradual colonization of terrestrial
habitats. However, it should be noted that non-vascular plants also exist,
demonstrating that the presence of the vascular system is more of an evolutionary
advantage than a prerequisite for survival (Agustí and Blázquez, 2020).
Xylem has two key functions; transport and mechanical support. The transport of
xylem sap (water, mineral nutrients and hormones) from the root towards the
aboveground organs occurs through dead xylem cells which form hollow pipes.
Vascular plants invest resources into xylem development, which in turn allows
water transport at low energetic cost under negative pressure. Water transport is
driven by the transpiration rate of the leaves. In the negative pressure environment
8
of the xylem conduits water is metastable, thus susceptible for cavitation. Air-
blockage or embolism of the water-conducting cells may lead to the impediment
of water flow, thus compromising plant health. Environmental factors, such as
drought and frost, affect resistance to embolism (Lens et al., 2013; Brodersen et
al., 2019). To perform water transport efficiently and maintain integrity, plants
have developed strategies to prevent and withstand the formation of air bubbles
under negative pressure.
The biomechanical demands of up-right growth and endurance of the water flow-
derived tension are assured by specific morphological features of the xylem and
mechanical support of the secondary cell walls. The cellulose, hemicellulose and
lignin content of the secondary cell walls and the interaction of these polymers
determine the chemical and physical properties of the xylem. These properties
also affect the degree at which a species is able to sequester carbon from the
atmosphere (Myburg et al., 2013).
2.4. Different cell types building up the xylem
Environmental factors, hormone levels and cross talk among the different plant
hormones promote the wood anatomical diversity seen between and within
species (Carlquist, 2013). The morphology, distribution and characteristics of the
specialized cell types present in the xylem determine the properties of wood
(Myburg et al., 2013). The three main cell types are tracheids or vessel elements
(depending on the plant species), fibers and parenchyma cells.
Tracheids and vessel elements
Tracheids and vessel elements, collectively called as tracheary elements, are the
water-conducting cells in the xylem. Tracheary elements are able to withstand the
negative pressure associated with sap flow thanks to their patterned and heavily
lignified secondary cell walls (Turner et al., 2007). Tracheids are narrow (8-80
µm), elongated (0.5-10 mm) cells, essential for xylem sap transport and structural
support, predominantly present in the xylem of gymnosperms (Panshin and de
Zeeuw, 1970; Pittermann, 2010; Wiedenhoeft, 2013). Vessel elements differ in
morphology, being shorter (0.1-1.2 mm) and wider in diameter (typically 50-200
µm). Vessel elements are specialized on sap transport and they are the
characteristic of angiosperms (Tyree and Zimmermann, 2002; Wiedenhoeft,
2013). The final steps of differentiation for these cell types are programmed cell
death, clearance of cell content through autolysis and the formation of bordered
pits and perforation plates (Fukuda, 1996). Bordered pits are lined by
9
semipermeable membrane-like structure made of the primary wall and middle
lamella at the pits of neighbouring cells, permitting fluid transport but blocking
the passage of large air bubbles and pathogens. The tracheids in gymnosperms
are highly connected through bordered pits; every tracheid is connected to several
others via pits along the radial walls (Choat et al., 2008). As tracheids overlap
with their lower and upper neighbouring cells 20-30% of their length, water needs
to take a zigzag route to pass through the pits. Tracheids are relatively inefficient
conduits due to their small diameter and the presence of the pit membrane which
imposes resistance to the water flow (Wiedenhoeft, 2013). Furthermore, the level
of connectivity within the xylem affects sap flow efficiency and susceptibility to
embolism (Brodersen et al., 2019). During vessel element development,
secondary cell wall is not deposited at the apical and basal ends of the cells.
Instead, membrane-free perforation plates are formed by partial digestion of the
primary walls (Turner et al., 2007). When differentiation is completed, multiple
vertically aligned vessel elements form hollow pipes called vessels, which
facilitate the free movement of water. Vessels are much longer than tracheids,
and may reach several meters in length (Brodersen et al., 2019). The individual
vessel elements are connected with each other through intervessel pits located on
their lateral walls, while half-bordered pits form between vessel elements and ray
cells (Wiedenhoeft, 2013). These properties make vessels far more effective in
water transport than tracheids (Lewis and Boose, 1995). Tracheids also occur
outside the gymnosperms, for example in the Fagaceae family, but due to their
smaller diameter the water transport performed by individual tracheids is
negligible compared to vessels. Primitive angiosperms, such as those in the genus
Amborella, lack vessels (Hacke et al., 2007; Sperry et al., 2007). Overall, the
general evolutionary trend within vascular plants was to increase xylem conduit
diameter and length, thus improving water conductance and capability to adapt
to a wide variety of environments (Lewis and Boose, 1995).
Fibers
Fibers provide mechanical support in the xylem. Generally, fibers are shorter than
tracheids (0.2-1.2 mm) but longer than the vessel elements of the same species
(Wiedenhoeft, 2013). Their thick lignified secondary cell walls are the main
determinants of the mechanical strength and density of wood. The model plant,
Arabidopsis, has long-living fibers (Bollhöner et al., 2012), in contrast to most
angiosperms where fibers, similarly to tracheary elements, undergo cell death.
However, this developmental program differs greatly between the two cell types;
DNA degradation in the fibers’ nuclei begins much prior to cell death, while in
10
tracheary elements DNA breakage seems to occur only shortly prior to the
vacuolar collapse. Upon vacuolar burst the cell content of xylem vessels is rapidly
degraded, in contrast to the gradual hydrolysis observed in fibers (Courtois-
Moreau et al., 2009; reviewed by Bollhöner et al., 2012).
Parenchyma cells and ray cells
Xylem parenchyma cells are the facilitators of a functional xylem by their
involvement in a multitude of processes. Tridimensional parenchyma cell
networks spam throughout the secondary xylem and phloem, ensuring high level
of interconnectivity. Xylem parenchyma cells often deposit secondary cell walls,
which have simpler structure than the cell walls of tracheary elements, resembling
thickened primary walls (Panshin and de Zeeuw, 1980). The bulk of the living
cells in the wood are the ray and axial parenchyma cells. Axial parenchyma cells
are derived from fusiform cambial initials, and as their name suggests, they are
axially elongated (Carlquist, 1988), while ray cells develop from ray initials. Ray
parenchyma cells are radially connected and they extend across the phloem,
cambium and xylem, enabling xylem-phloem exchange. A subset of parenchyma
cells is in direct contact with water-conducting cells through cell wall pits or
indirectly via the apoplast (reviewed by Spicer, 2014). The cells within the rays
are connected through plasmodesmata, enabling the continuous flow of resources
(Sauter and Kloth, 1986; Chaffey and Barlow, 2001).
Parenchyma cells have a wide range of functions from storage and transport of
non-structural carbohydrates, lipids and storage proteins (Höll, 2000; Plavcová
and Jansen, 2015) to water storage (Holbrook, 1995; Borchert and Pockman,
2005) and defense against biotic factors (Schmitt and Liese, 1993). Thanks to the
interconnection between tracheary elements and parenchyma cells, the secondary
cell walls of dead, functional TEs are structurally reinforced by the lignin
monomers coming from the parenchyma cells (Smith et al., 2013; Pesquet et al.,
2013; Zhang et al., 2020A). Furthermore, parenchyma cells are implicated in long
distance water transport, whereby they alter xylem hydraulic conductance and
sap flow rate through ion exchange into the transpiration stream (Jansen et al.,
2011; Nardini et al., 2011; reviewed by Ménard and Pesquet, 2015). Parenchyma
is also involved in restoring water transport in embolized tracheary elements by
the active export of sucrose, ions and H+ into the embolized cell, thus reducing
the water potential and triggering sap influx (Salleo et al., 2004; 2009; Brodersen
et al., 2010; Secchi and Zwieniecki, 2011; 2012). When a water-conducting cell
gets fatally damaged, the neighbouring xylem parenchyma cells will fill the
lumen of the dysfunctional cell with tyloses, pectin-rich gels and gums, thus
11
sealing the cell off from the conducting network (Kitin et al., 2010; Sun et al.,
2008).
The fraction, anatomy and spatial distribution of parenchyma cells are highly
variable in different species, and this variability has been exploited as a diagnostic
feature for wood identification (IAWA Committee, 1989; 2004). The quantitative
and qualitative features related to parenchyma cells are influenced by genetics
and environmental factors, such as temperature, precipitation and seasonality. In
general, coniferous gymnosperms contain very few parenchyma cells, while stem
succulents and lianas exhibit the highest proportions of xylem parenchyma within
the plant kingdom (Morris et al., 2016). In the xylem of the model plant
Arabidopsis, parenchyma cells are interspersed between fibers and vessel
elements (Smith et al., 2013). Under normal growth, Arabidopsis does not form
rays, but artificial weight treatment may induce this developmental process,
highlighting the plasticity of xylogenesis (Mazur and Kurczynska, 2012).
3. Secondary cell wall
Generally, secondary cell wall deposition takes place after plant cells have
reached their final size by expansion. By definition, a “secondary cell wall” has
a specific chemical composition, characterized by the presence of cellulose,
hemicellulose and lignin, with less contribution from cell wall proteins and
enzymes (Sjöström, 1993). The primary cell wall is mainly composed of pectin,
hemicellulose, cellulose and glycoproteins (Fry, 2004). The secondary cell wall
is deposited inside the primary cell wall, between the primary cell wall and the
plasma membrane. In order to begin secondary cell wall formation, the primary
cell wall biosynthetic machinery is replaced by new enzymes to meet the
biosynthetic and regulatory demands imposed by the secondary cell walls. First,
the cell wall polysaccharides, hemicellulose and cellulose, are deposited,
followed by lignification (Meents et al., 2018).
3.1. Cellulose
With a global annual yield of approx. 100 billion tonnes, cellulose is the most
abundant natural polymer on earth (de Souza Lima and Borsali, 2004). Cellulose
contributes to the load-bearing scaffold of the cell, accounting for 40-50% of the
SCW in land plants (Timell, 1967). Cellulose is composed of a linear chain of β-
1,4 linked glucose units, organized into microfibrils through intra- and
intermolecular bonds and Van der Waals forces (Kim et al., 2013). In contrast to
12
cellulose present in the primary cell walls, the cellulose of the secondary cell
walls displays higher degree of polymerization and crystallinity.
Cellulose microfibrils are synthesized at the plasma membrane by the large,
mobile, rosette-shaped cellulose synthase protein complex (CSC). The CSC is a
complex structure, built of multiple individual CesA subunits. The enzymes of
CSC specialized on secondary cell walls are CesA4/IRX5, CesA7/IRX3 and
CesA8/IRX1 (Turner and Sommerville, 1997; Taylor et al., 2003). Several recent
studies provided evidence that the CSC is likely to be a hexamer composed of
trimers or tetramers of 18-24 CesAs instead of the previously proposed structure
of the CSC composed of a minimum of 36 CesAs (reviewed by Meents et al.,
2018). The precise stoichiometry of the complex differs between species. In
Arabidopsis, both primary and secondary cell wall CesAs are present in
equimolar (1:1:1) amounts, likely composing a CSC with 18 subunits (Gonneau
et al., 2014; Hill et al., 2014). Using cytosolic UDP-glucose as a substrate, the 18
subunits simultaneously synthesize 18 glucan chains bound into a cellulose
microfibril with a diameter of approx. ~2.5 nm (reviewed by Polko and Kieber,
2019). Zhang et al. (2018) demonstrated that the gymnosperm Norway spruce
(Picea abies) has the same equimolar CesA stoichiometry as in Arabidopsis,
while the angiosperm aspen (Populus tremula) had a CesA ratio of 3:2:1 for
PtCesA8a/b:PtCesA4:PtCesA7a/b.
While ejecting glucan chains which polymerize and crystallize to cellulose
microfibrils, the CSCs move along cortical microtubule rails in the plasma
membrane (Paredez et al., 2006). The linker protein CSI1/POM2 physically
connects CSCs to the microtubules (Li et al., 2012; Bringmann et al., 2012;
Derbyshire et al., 2015; Schneider et al., 2017). Recently, the microtubule-
independent movement of CSCs was demonstrated, where the CSCs progress on
trails left by previous complexes, likely by interacting with recently formed
cellulose microfibrils. The microtubule-guided movement of CSCs is more
common, and can override the microtubule-independent CSC movement (Chan
and Coen, 2020). The CSCs in the SCW are present in higher density than in
PCW (Watanabe et al., 2015). They form clusters which move on coordinated
trajectories, depositing aggregates of cellulose microfibrils characteristic of the
SCW (reviewed by Meents et al., 2018).
Beyond the core cellulose synthase machinery, several non-CesA proteins have
been linked to SCW cellulose synthesis although their mode of action remains to
be scrutinized. KORRIGAN, a membrane-bound endoglucanase, contributes to
proper cellulose biosynthesis, likely by regulating cellulose crystallinity
13
(Szyjanowicz et al., 2004; Maloney and Mansfield, 2010). The transmembrane
proteins Tracheary Element Differentiation-Related6 (TED6) and TED7 bind to
the SCW CSCs, suggesting that they may be accessory components of the
complex (Endo et al., 2009; Rejab et al., 2015). COBRA-LIKE4, a
phosphatidylinositol-anchored glycoprotein, binds to cellulose microfibrils and
affects cellulose crystallinity (Persson et al., 2005; Liu et al., 2013A).
3.2. Hemicellulose
10-40 % of the SCW is composed of hemicelluloses. Hemicelluloses contribute
to the load-bearing capacity of the SCW by their interaction with cellulose and
lignin. A variety of hemicelluloses occur in terrestrial plants, including
xyloglucans, xylans, mannans, glucomannans and β-(1-3, 1-4)-glucans. Xylans
are the predominant noncellulosic polysaccharide of dicot secondary cell walls.
Mannans, in the form of galactoglucomannans, are the major hemicellulose in the
SCW of gymnosperms (reviewed by Scheller and Ulvskov, 2010).
In contrast to cellulose, hemicelluloses are synthesized in the medial section of
the Golgi apparatus and subsequently transported to the plasma membrane. For
efficient hemicellulose biosynthesis the enzymes involved in the process are
likely to form complexes. In planta assays done in wheat (Triticum aestivum) and
asparagus (Asparagus officialis) revealed that the xylan biosynthetic enzymes
form homo- and heterodimers in the Golgi (Zeng et al., 2010; 2016). There are
three key enzymes which generate the xylan backbone; the Glycosyltransferase
Family 43 (GT43) members IRX9 and IRX14, and a GT47 member IRX10. They
all have functionally redundant paralogs; IRX9-L, IRX14-L and IRX10-L (Lee
et al., 2007; Brown et al., 2007; Wu et al., 2010). Since IRX10 does not have a
transmembrane domain, the interaction and complex formation with IRX9 and
IRX14 may be necessary for its proper Golgi localization and function (reviewed
by Meents et al., 2018). Functional characterization studies revealed that
IRX9/IRX9-L may have structural rather than enzymatic function,
IRX14/IRX14-L plays a role in substrate binding, while IRX10/IRX10L
presumably has xylosyl transferase activity (Ren et al., 2014; Urbanowicz et al.,
2014; Zeng et al., 2016)
The glucuronic acid (GlcA) substitutions on the xylan backbone are introduced
by two members of the GT8 family, Glucuronic Acid Substitution of Xylan
(GUX) 1 and GUX2 (Rennie et al., 2012). GUX1 adds the GlcA side chains
preferentially to evenly spaced xylose residues, always on the same side of the
backbone. GUX2 generates more tightly clustered decorations, substituting
14
xylose residues on either side of the backbone. The activity of GUX1 and GUX2
yields two different xylan domains which are likely to be part of the same
heterogenous xylan molecule. These distinct xylan domains may contribute to the
crosslinking capacity of the molecule with other cell wall polymers (Bromley et
al., 2013). Next, methyl groups may be added to the GlcA residues by members
of the glucuronoxylan methyltransferase (GXMT) family, which are DUF579
domain-containing proteins. GXMT1 methylates 75% of the GlcA residues in
Arabidopsis (Urbanowicz et al., 2012). IRX15 and IRX15L are putative
methyltransferases with no detected catalytic activity, likely playing a structural
role in the xylan synthase complex, similarly to IRX9/IRX9-L (Brown et al.,
2011; Jensen et al., 2011).
Initially, the DUF231 domain-containing acetyltransferase, ESK1/TBL29, was
shown to perform the monoacetylation of xylan at O-2 and O-3 position (Yuan et
al., 2013; Xiong et al., 2013). ESK1/TBL29 is required for evenly patterned xylan
acetylation, which then guides the activity of GUX1 for the evenly distributed
GlcA depositions (Grantham et al., 2017). Recently, several studies dissected the
role of other TBL family members in xylan acetylation. TBL3 and TBL31
specifically affect O-3 acetylation (Yuan et al., 2016A), while TBL32, TBL33,
TBL34 and TBL35 mediate the acetyl substitutions of xylosyl residues at O-2
and O-3 (Yuan et al., 2016B; 2016C). These TBL enzymes may also be essential
for the establishment of uniform xylan decorations. The four REDUCED WALL
ACETYLATION (RWA) genes in Arabidopsis also contribute to xylan
acetylation and proper secondary cell wall formation, but they have lower
substrate specificity than the TBL proteins since they can acetylate mannans and
xyloglucans as well. RWA and TBL proteins may form complexes where the
specificity of TBL drives the substrate preference. Alternatively, RWA might
perform xylan acetylation first, thus providing the substrate to TBL (Manabe et
al., 2013).
The even pattern of xylan substitution is a characteristic of vascular plants and
essential for the binding of xylan to the hydrophilic faces of cellulose microfibrils
to ensure normal secondary cell wall development (Grantham et al., 2017).
3.3. Cell wall proteins
Cell wall proteins account for a minor fraction of the SCW and their exact
function is poorly understood (Liu et al., 2013A). Cell wall proteins typically
belong to the following groups: glycine-rich (GRPs), proline-rich (PRPs),
arabinogalactan-rich (AGPs) and hydroxyproline-rich glycoproteins (HRGPs or
15
extensins) (Showalter, 1993). Fasciclin-like AGP (FLA) proteins are enriched in
the SCWs, and supposedly involved in cellulose deposition on the basis of the
presence of a GPI-anchor on several FLA and reverse genetic studies (reviewed
by Kumar et al., 2016).
3.4. Lignin
Lignin is the product of the phenylpropanoid pathway, an aromatic polymer,
which constitutes 15-36% of the woody cell wall biomass (Zobel and van
Buijtenen, 1989). Lignification, the final step of SCW biosynthesis, occurs prior
as well as after cells undergo programmed cell death (Smith et al., 2013; Pesquet
et al., 2013; Zhang et al., 2020A).
The evolution of lignin
Early in the evolution of the monolignol biosynthetic pathway its role might have
been to produce UV-protectant molecules (reviewed by Weng & Chapple, 2010).
Eight key enzymes required for the biosynthesis of the monolignols p-coumaryl
alcohol and coniferyl alcohol were most likely recruited from the primary
metabolism. The pathways within the primary metabolism are vital for plant
survival, and their products serve as precursors for the diverse reactions of the
secondary metabolism (reviewed by Pott et al., 2019). Homologs of the lignin-
biosynthetic enzymes can be identified within the primary metabolism. Besides
lignin, other hydrophobic polymers such as cutin, suberin, sporopollenin and
soluble metabolites such as flavonoids, tannins, stilbenes and lignans are the end
products of the phenylpropanoid pathway (reviewed by Weng & Chapple, 2010).
Major monolignols
To date 35 different naturally occurring lignin monomers have been identified,
most of which are very species specific. The three most widely occurring
monomers, the p-hydroxyphenyl (H), guaiacyl (G) and syringyl (S) units, are
derived through the polymerization of p-coumaryl, coniferyl and sinapyl alcohol,
respectively. Branching and interconnections curtail the flexibility of the
polymer. The formation of many chain-starting monomers results in lignin with
low molecular weight (reviewed by Vanholme et al., 2019). The more rare
compounds have potentially been under selective pressure, therefore the three
major lignin units contribute to the bulk of the lignin polymer.
16
Distribution of lignin in different species and cell types
Lignin content, monomer composition and interunit linkage distribution vary
depending on the species, cell type, cell wall layer as well as upon environmental
stimuli (Campbell and Sederoff, 1996; Studer et al., 2011). Lignin has a very
wide pattern for deposition, being present in tracheary elements, sclerenchyma
cells, endodermal cells, seed coat and silique cells (Barros et al., 2015).
Gymnosperm species contain mostly G units with minor amounts of H units,
while angiosperm lignin is primarily formed of G and S units. The H unit content
is higher in monocots than in dicots (Boerjan et al., 2003). S-type lignin is specific
to flowering plants and the lycophyte Selaginella, while H- and G-type lignin are
elemental for all tracheophytes (reviewed by Weng and Chapple, 2010).
The water-conducting xylem vessels contain mostly G units and lignify first,
while the structural fibers containing both G and S units lignify later. H units are
incorporated into the middle lamella of vessel elements and fibers early on during
SCW formation (Fukushima and Terashima, 1990). H units are typically
deposited at the beginning of the lignification process in the cell corners and
middle lamella, G units are first deposited in the middle lamella and later in the
SCW layers, and finally S units are incorporated into the SCW (Terashima and
Fukushima, 1988; Terashima et al., 1988; Fukushima and Terashima, 1991;
reviewed by Donaldson, 2001). The relative abundance of each lignin unit affects
the overall structure and thus the physical properties of the polymer (reviewed by
Bonawitz and Chapple, 2010).
The function of lignin
The chemical structure of lignin confers rigidity and hydrophobicity to the cell
wall, which facilitates the transport of water and solutes through the vascular
system (Boerjan et al., 2003; Vanholme et al., 2010; Barros et al., 2015).
Moreover, lignin protects the plants against various environmental stresses, such
as pathogen attacks (reviewed by Miedes et al., 2014). Different developmental
and environmental factors, such as wounding, metabolic stress or perturbation in
the cell wall integrity, can trigger lignin biosynthesis (Cano-Delgado et al., 2003;
Tronchet et al., 2010).
3.4.1. Biosynthesis of the lignin monomers
3.4.1.a. The lignin biosynthetic pathway
Monolignols are synthesized most often from phenylalanine, in grasses also from
tyrosine (Barros et al., 2016), by sequential steps involving eleven enzymes of
17
the phenylpropanoid pathway. The H-, G- and S-lignin units are distinguished by
their degree of methoxylation.
Phenylalanine, the precursor of the phenylpropanoid pathway, is derived from the
shikimate pathway and converted in the first enzymatic step to cinnamic acid by
phenylalanine ammonia-lyase (PAL). Subsequently, the aromatic ring is p-
hydroxylated by the enzyme cinnamate 4-hydroxylase (C4H) and further
converted to p-coumaryl-CoA by the 4-coumarate:CoA ligase (4CL) enzyme.
After this, two possible directions can be taken; if H-lignin unit is being produced
then the p-coumaryl-CoA is converted to p-coumaryl alcohol by cinnamoyl-CoA
reductase (CCR) and cinnamyl alcohol dehydrogenase (CAD). If G or S-lignin
unit synthesis is taking place, p-coumaroyl-CoA is meta-hydroxylated by p-
hydroxycinnamoyl-CoA shikimate/quinate hydroxycinnamoyltransferase (HCT)
and p-coumarate 3-hydroxylase (C3H). The meta-hydroxyl residue is then
converted into methoxy residue by caffeoyl-CoA O-methyltransferase
(CCoAOMT), which is followed by the conversion of the CoA-conjugate of the
propene residue to an aldehyde by CCR. At this step CCR redirects the
phenylpropanoid pathway towards lignin monomer biosynthesis. The aldehyde
moiety of the coniferaldehyde is converted to an alcohol by CAD, which leads to
G unit production. The synthesis of S unit monomers requires the hydroxylation
of the coniferaldehyde by the enzyme ferulate 5-hydroxylase (F5H), which is
followed by a methoxylation by caffeic acid O-methyltransferase (COMT) (Fig.
2).
The hydroxylation enzymes, C4H, C3H and F5H are bound to the endoplasmic
reticulum (ER), while all other lignin biosynthetic enzymes are located in the
cytosol (Takabe et al., 2001; Ruelland et al., 2003). The interactions found
between the ER bound and cytosolic enzymes suggest that the site of lignin
monomer biosynthesis is in the cytosol, close to the ER (Achnine et al., 2004).
The enzymes 4CL, CCR, CAD, HCT, COMT and F5H have multiple substrates,
hence the pathway is composed of multiple parallel routes towards coniferyl and
sinapyl alcohol production (reviewed by Vanholme et al., 2019). Several
biosynthetic enzymes form protein complexes to increase activity and efficiency;
CAD1 and CCR2 form heterodimers (Yan et al., 2019), the enzymes performing
the 4- and 3-hydroxylation of cinnamic acid, C4H1, C4H2 and C3H, form
heterodimers and trimers (Chen et al., 2011), and 4CL3 and 4CL5 form
heterotetrameric complexes (Chen et al., 2014). Furthermore, the cytochrome
P450 enzymes, C4H, C3H and F5H, form clusters through interactions with
scaffold proteins (Gou et al., 2018).
18
Figure 2. Monolignol biosynthesis through the phenylpropanoid pathway.
Each arrow represents a reaction, the abbreviated names of the enzymes catalyzing
the reaction are shown in orange. The name of the intermediates of the biosynthetic
pathway are shown in bold black. The light orange background highlights the core
part of the pathway as it is known to this date. PAL, Phenylalanine ammonia-lyase;
C4H, Cinnamate 4-hydroxylase; 4CL, 4-coumarate:CoA ligase; HCT,
Hydroxycinnamoyltransferase; C3H, P-coumarate 3-hydroxylase; CSE, Caffeoyl
shikimate esterase; CCoAOMT, Caffeoyl-CoA O-methyltransferase; CCR,
Cinnamoyl-CoA reductase; CAD, Cinnamyl alcohol dehydrogenase; HCALDH,
hydroxycinnamaldehyde dehydrogenase; F5H, Ferulate 5-hydroxylase; COMT,
Caffeic acid O-methyltransferase.
3.4.1.b. Regulation of lignin biosynthesis
Secondary cell wall formation including lignin deposition is strictly controlled in
order to direct and restrict lignin deposition to the proper cell types and
developmental stages, and to avoid abnormal growth and development (Behr et
al., 2019). Multiple regulatory steps underlie the production of lignin polymer
biosynthetic enzymes, production of lignin polymer precursors, transport of
lignin polymer enzymes and precursors, and finally building of the highly
organized lignin structure (reviewed by Kumar et al., 2016).
19
Transcriptional regulation
More than 2000 genes in Arabidopsis are possibly related to cell wall biogenesis
(Carpita et al., 2001). The activity of the genes involved in cell wall synthesis and
modification is under transcriptional and post-transcriptional control. Taylor-
Teeples et al. (2015) characterized the complex gene regulatory network of
Arabidopsis transcription factors and secondary cell wall metabolic genes,
illustrating that xylem cell differentiation, including phenylpropanoid
biosynthesis, is controlled by a highly branched rather than a linear regulatory
pathway. Therefore, perturbation of a side pathway can lead to massive changes
in the whole network. This network analysis detected feed forward loops as a
very frequent form of regulation. E2Fc was identified as a key upstream regulator
of the master regulators as well as the structural genes. E2Fc, a negative regulator
of endoreduplication (Del Pozo et al., 2007), belongs to the E2F family of
transcription factors, and can act as a transcriptional activator as well as repressor
in a dose-dependent manner.
The majority of the transcription factors controlling secondary cell wall formation
belong to the NAC (NAM, ATAF1/2, CUC2) domain or to the MYB
(Myeloblastosis) transcription factor family, and they act in hierarchical order
(Fig. 3). The first level of transcription factors are the master regulators of
secondary cell wall formation. There are five master regulator NAC domain
transcription factors that belong to the VNS subfamily (VND, NST/SND, and
SMB-related protein): NST1 (NAC SECONDARY WALL THICKENING
PROMOTING FACTOR1), SND1 (SECONDARY WALL-ASSOCIATED
NAC DOMAIN1), NST2, VND6 (VASCULAR RELATED NAC DOMAIN6)
and VND7. The VNS proteins act as molecular master switches of xylem cell
differentiation by regulating genes encoding for cell wall biosynthetic enzymes
and genes related to programmed cell death (Ohtani and Demura, 2019). NST1 is
expressed in interfascicular fibers and differentiating vessels (Mitsuda et al.,
2005; 2007), and SND1 is expressed in interfascicular and xylary fibers (Zhong
et al., 2006). NST1 and SND1 regulate secondary cell wall formation in fibers
(Mitsuda et al., 2005; 2007; Zhong et al., 2006). In contrast, the expression and
regulatory activity of VND6 and VND7 are xylem vessel specific (Kubo et al.,
2005; Yamaguchi et al., 2008). Several molecular factors have been reported to
act as negative regulators of the master switches. VND7 interacts with VND-
INTERACTING2 (VNI2) which leads to the repression of vessel-specific genes
regulated by VND7, making VNI2 a transcriptional repressor of xylem cell fate
(Yamaguchi et al., 2010). XYLEM NAC DOMAIN1 (XND1) inhibits tracheary
element differentiation and premature SCW formation (Zhao et al., 2008; 2017).
20
XND1 interacts with NST1 thereby modulating SCW deposition (Zhang et al.,
2020B). WRKY12 directly binds to the NST2 promoter, thus suppressing the
SCW activator role of NST2 (Wang et al., 2010). Several of these so-called first
level transcription factors bind to a conserved cis-acting element, the secondary
wall NAC-binding element (SNBE), present in the promoter of all their targets
(Zhong et al., 2010).
Figure 3. Transcriptional regulation of secondary cell wall formation. Transcription
factors coordinate the transcriptional regulation, and E2Fc is the key upstream
regulator. The first and second layer master switches can regulate the transcription
factors below them or the secondary cell wall biosynthetic genes directly. The
downstream regulators directly bind to the secondary cell wall biosynthetic genes.
γMYB2 represses some of the second layer master switches and downstream
regulators. Subunits of the transcriptional co-regulator Mediator complex regulate
lignification through homeostatic repression.
MYB (MYB46, MYB83 and MYB55) and NAC (SND3 and XND1)
transcription factors are among the second layer of transcriptional regulators.
MYB46 and MYB83 were shown to bind to the secondary wall MYB-responsive
element (SMRE), and thus directly control an array of not only transcription
factors (MYB58, MYB63), but secondary cell wall biosynthetic genes as well
(Zhong and Ye, 2012). The third level comprises transcription factors which are
downstream targets of the master regulators and the members of the second layer
transcription factors. They control specifically the expression of cell wall
biosynthetic genes by binding to their promoters. MYB20, MYB69, MYB79,
21
MYB85, MYB58, MYB63, MYB103 and KNAT1 control the expression of the
lignin-biosynthetic genes (reviewed by Kumar et al., 2016; Öhman et al., 2013).
MYB58 and MYB63 transcriptionally activate all lignin-biosynthetic genes
(except F5H) as well as LAC4, through binding to the AC cis-regulatory elements
of the promoters. C4H and COMT were also directly activated, even though their
promoter may only contain deteriorated AC elements. Similar AC element
dependent activation of lignin-biosynthetic genes was shown for MYB85 (Zhou
et al., 2009). MYB103 affects S-type lignin content and S/G lignin ratio by
altering the expression of F5H through an unknown mechanism (Öhman et al.,
2013). The knotted-like homeobox (KNOX) protein, KNAT1, directly regulates
several lignin-biosynthetic genes by binding to their promoters (Mele et al.,
2003).
Furthermore, MYB4 and its two close homologs MYB7 and MYB32 are negative
regulators of the lignin-biosynthetic pathway (reviewed by Zhang et al., 2018;
Behr et al., 2019). Recently, MYB20, MYB42, MYB43 and MYB85 were shown
to act redundantly in the activation of phenylalanine and lignin biosynthesis. This
transcription factor quartet also suppressed flavonoid biosynthesis through
MYB4, thus ensuring sufficient phenylalanine supply for the lignin-biosynthetic
pathway (Geng et al., 2020). MYB52 is another negative regulator of
lignification. It is co-expressed with several SCW related TFs and biosynthetic
genes, proposed to have a role in maintaining the balance between the production
of different SCW components (Cassan-Wang et al., 2013). MYB75 promotes
anthocyanin biosynthesis and, together with KNAT7, suppresses SCW formation
(Bhargava et al., 2010; 2013). A member of the MYB coiled-coil family, γMYB2,
is also involved in SCW establishment by repressing SND3, MYB46, MYB63 and
MYB103 (Nguyen et al., 2019).
Besides the evolutionarily highly conserved NAC-MYB gene regulatory network
(Xu et al., 2014; Bowman et al., 2017), there are likely to be additional molecular
regulators and modulators involved in the control of lignification. These
molecular factors might be less conserved, but more species specific (Ohtani and
Demura, 2019).
In addition to the transcription factors, the transcription of lignin-biosynthetic
genes is also controlled by the Mediator complex (Fig. 3). The Mediator complex
is a transcriptional co-regulator in eukaryotes containing around 30 subunits,
accounting for the core head, middle and tail modules, and the dissociable kinase
module (Asturias et al., 1999; Tsai et al., 2014). MED5a/5b, two paralogs located
in the tail module of Mediator, are homeostatic repressors of the phenylpropanoid
22
metabolism (Bonawitz et al., 2012; 2014). MED5a/5b genetically interact with
the other subunits of the tail module MED2, MED16 and MED23 (Dolan et al.,
2017), as well as the subunits of the kinase module CDK8 and MED12 (Mao et
al., 2019A), thereby regulating lignin biosynthesis and plant growth (Mao et al.,
2019B). The role of Mediator in the regulation of lignification may be to integrate
various signals, ensuring appropriate developmental transitions.
Chromatin-level regulation
To fine-tune plant developmental processes, the transcriptional activity of genes
is commonly regulated at the chromatin level. The state of the chromatin affects
its accessibility for the transcription factor machinery. While a plethora of
transcription factors have been linked to lignification, knowledge on its
chromatin level regulation is sporadic. Studies done in the xylem of the
Eucalyptus tree model found correlations between the abundance of the
transcriptionally activating (H3K4me3) and repressive (H3K27me3) chromatin
marks and the expression level of some lignin-biosynthetic genes (Hussey et al.,
2015; 2017). The H3K27me3 enrichment of lignin and phenylpropanoid
biosynthetic genes during early xylem development corresponds to the lack of
lignification at that point (Hussey et al., 2017). Plants likely utilize the toolbox
of chromatin modifications to coordinate the developmental transitions within the
xylem. Genes involved in secondary cell wall development might be repressed in
the cell expansion zone and activated later on, however this aspect of lignification
needs to be elucidated in future studies.
Circadian regulation
The circadian clock is present in all living organisms. In plants the biological
timing coordinates the metabolism, growth, molecular and cellular processes with
environmental changes. Secondary growth is also likely to be under circadian
regulation.
It was demonstrated in Populus tremula x tremuloides trees that the circadian
clock differentially influences the auxin and cytokinin flow, which in turn affects
cell elongation and xylem formation in a diurnal fashion (Edwards et al., 2018).
Genes encoding for 23 enzymes from the phenylpropanoid pathway exhibit a
synchronized expression peak before dawn (Harmer et al., 2000). Furthermore,
starch turnover and carbohydrate availability influence the magnitude of
circadian oscillations of lignin biosynthetic gene expression. Independent of the
circadian clock these same biosynthetic genes may be impacted by light
perception as well (Rogers et al., 2005). 13C labelling experiments of the
23
developing wood of Populus tremula x tremuloides trees showed that the
incorporation of labelled carbon to lignin was highest at the end of the night,
supporting the early findings (Harmer et al., 2000) that lignin biosynthesis takes
place before dawn (Mahboubi et al., 2015).
Lignin is a major carbon sink and carbon availability is cyclic during the day.
Involvement of diurnal aspect in the control of lignin biosynthesis is plausible
due to the fact that optimal biomass accumulation in the woody tissues should be
coordinated with resource availability and starch turnover (Rogers et al., 2005).
Additional levels of regulation
The enzyme catalysing the first step of lignin biosynthesis, PAL, controls the flux
into the pathway. PAL is known to be under post-translational control. Kelch-
repeat F-box (KFB) proteins physically interact with PAL isozymes and mediate
their proteolysis via the ubiquitin-26S proteasome pathway (Zhang et al., 2013).
Furthermore, the Populus trichocarpa COMT is regulated by reversible
phosphorylation, enabling the cell to adjust the metabolic flux to developmental
and environmental needs (Wang et al., 2015).
Other factors affecting the flux through the pathway are the compartmentalization
of pathway intermediates as well as their inhibitory activity towards specific
biosynthetic enzymes, and the access to cofactors and co-substrates like
shikimate, ATP and S-adenosylmethionine (reviewed by Vanholme et al., 2019).
3.4.2. Transport of the lignin monomers
The detection of only scarce pools of free monolignols by targeted metabolomics
(Jaini et al., 2017) suggests that, upon synthesis, the lignin monomers are rapidly
transported across the plasma membrane for polymerization in the secondary cell
wall. Three different transport mechanisms have been proposed: exocytosis via
ER-Golgi derived vesicles, passive diffusion, and export via membrane-bound
transporters and proton-coupled antiporters (Meents et al., 2018). Observations
of early autoradiographic studies using radiolabelled phenylalanine suggested
that monolignols are present in the rough ER, Golgi and plasma membrane
associated vesicles (Pickett-Heaps, 1968; Fujita and Harada, 1979; Takabe et al.,
1985). The more recent findings of Kaneda et al. (2008) showed that the
phenylalanine incorporation was primarily into proteins rather than monolignols.
Hence the contribution of the ER-Golgi-vesicle mediated exocytosis as a lignin
monomer transport mechanism remains unclear. The small size of monolignols
may facilitate their passive diffusion across membranes. This theory was
24
supported by in vitro observation of the diffusion of lignin precursor analogs into
liposomes and lipid bilayer discs (Boija et al., 2007; 2008), but there is no in vivo
evidence for passive diffusion yet. According to this theory, the concentration
gradient of monolignols between the cytosol and the cell wall would determine
the diffusion rate. The secondary cell wall localized lignin-polymerizing laccases
and peroxidases may create a monolignol sink and an energetically favourable
gradient by the quick polymerization of free monolignols (Perkins et al., 2019).
In addition, Vermaas et al. (2019) demonstrated that most lignin-related
compounds can passively cross membranes at adequate translocation rate to meet
the need of the polymerization machinery.
The third mechanism for lignin monomer transport across the plasma membrane
to the cell wall is mediated by ATP-binding cassette (ABC) transporters and
proton-coupled antiporters (PCA) (Miao and Liu, 2010; Kaneda et al., 2011;
Alejandro et al., 2012; Tsuyama et al., 2013). Based on co-expression with
phenylpropanoid biosynthetic genes, several ABC transporters were identified as
candidates in monolignol export (Ehlting et al., 2005). To date only one,
ABCG29, has been demonstrated as a monolignol transporter (Alejandro et al.,
2012). ABCG29 is a candidate exporter for p-coumaryl alcohol units which are
however only found in small amounts in the angiosperm secondary cell wall.
Yeast cells expressing the Arabidopsis ABCG29 were capable of p-coumaryl
alcohol transport, while the lack of ABCG29 in Arabidopsis increased the
sensitivity to this alcohol (Alejandro et al., 2012). Furthermore, ABCG11,
ABCG22, ABCG29 and ABCG36 are co-expressed with MYB58 in Arabidopsis
tracheary element cell cultures, indirectly suggesting their involvement in
lignification (Takeuchi et al., 2018). In addition, proton-coupled antiporters were
proposed as a transport mechanism of lignin precursors in both angiosperms and
gymnosperms. Membrane vesicles prepared from the differentiating xylem
displayed coniferin (the glucoside of coniferyl alcohol) transport activity in an
ATP-dependent fashion (Tsuyama et al., 2013). Such conserved, proton-gradient
dependent transport mechanism was demonstrated for p-glucocoumaryl alcohol
as well (the glucoside of p-coumaryl alcohol) (Tsuyama et al., 2019). Currently,
the passive diffusion of monolignols without a need for transporters seems to be
the strongest theory for monolignol movement across the plasma membrane
(Vermaas et al., 2019), however this needs to be confirmed with more in vivo
studies.
25
3.4.3. Lignin polymerization
Laccases and peroxidases
Polymerization of lignin typically follows the deposition of polysaccharides
during secondary cell wall formation. It has been postulated that the
polysaccharide matrix provides a scaffold for lignin polymerization (Tobimatsu
and Schuetz, 2019). In the cell wall, each monolignol produces the monomer
radical form by losing one H atom. The free radicals then assemble by oxidative
coupling, forming the lignin polymer. Laccases and peroxidases are the
phenoloxidases implicated in the activation of monolignols to radicals (Fig. 4).
Upon polymerization, monolignols become syringyl (S), guaiacyl (G) and p-
hydroxyphenyl (H) lignin units (Boerjan et al., 2003).
Laccases are multi-copper containing extracellular glycoproteins that oxidize
phenolic, inorganic and aromatic amine substrates, using molecular oxygen (O2)
as a secondary substrate. A large number of functionally redundant laccase
isoforms are present in angiosperms. Out of the 17 laccases found in Arabidopsis,
reverse genetic studies showed that LACCASE4 (LAC4), LAC11, LAC15 and
LAC17 have roles in lignin polymerization. While LAC15 is required for seed
coat lignification (Liang et al., 2006), LAC4, LAC11 and LAC17 are involved in
the lignification of tracheary elements and xylem fibers (Berthet et al., 2011;
Zhao et al., 2013). Several members of the LAC gene family are under the post-
transcriptional control of miRNAs. miRNA397 targets members of the LAC gene
family in Populus trichocarpa, Chinese pear, flax (Linum usitatissimum) and
hemp (Behr et al., 2019). Moreover, the Arabidopsis miRNA857 and
miRNA397b regulate LAC7 and LAC4 transcripts, respectively (Zhao et al.,
2015; Wang et al., 2014).
Another group of phenoloxidases expressed in lignifying tissues are the heme
containing class III peroxidases. These enzymes polymerize lignin monomers in
vitro and their secondary substrate is hydrogen peroxide (H2O2). Similar to
laccases, peroxidases are encoded by a multigene family. Arabidopsis has 73
class III peroxidases. Only a subset of these enzymes has been implicated in
lignification of different cell types: PEROXIDASE2 (PRX2), PRX4, PRX25,
PRX47, PRX52, PRX64, PRX66, PRX71 and PRX72 (Tokunaga et al., 2009;
Herrero et al., 2013; Lee et al., 2013; Fernández-Pérez et al., 2015; Shigeto et al.,
2013; 2015). Peroxidases display different affinity towards the lignin precursors.
For example, some only use sinapyl alcohol as a substrate (Østergaard et al.,
2000), while others can catalyze the polymerization of sinapyl and coniferyl
alcohol as well (Demont-Caulet et al., 2010).
26
Several studies suggest that laccases and peroxidases do not act redundantly, but
rather in a complementary manner (Lee et al., 2013; Zhao et al., 2013). Laccases
are expressed in the Casparian strip forming endodermis, but they do not
contribute to the lignification of the Casparian strip (Rojas-Murcia et al., 2020).
Lignin polymerization in the Casparian strip is dependent on PRX64 (Lee et al.,
2013). The establishment of functional TEs involves both phenoloxidase
families, and it has been proposed that laccases and peroxidases may act
sequentially in TE lignification (Sterjiades et al., 1993). Barros et al. (2015)
discussed three hypothetical models how the phenoloxidase activities could be
determined by different substrate specificities, distinct spatio-temporal
expression or differential complex formation. Complementary activity of
laccases and peroxidases is supported by the subcellular localization study which
found that fluorescently-tagged LAC4 is localized in all secondary cell wall
layers of fibers and vessels, while PRX64 is found only in the middle lamella and
cell corners of fibers (Chou et al., 2018). Recently, Hoffmann et al. (2020)
demonstrated that, depending on the cell type and the developmental stage,
specific LACs and PRXs colocalized within specific cell wall domains. High
resolution localization analyses in Arabidopsis revealed the presence of LAC4,
LAC17, and PRX72 in the thick secondary cell wall of xylem vessel elements
and fibers, whereas LAC4, PRX64, and PRX71 were localized to fiber cell
corners. Early in fiber development LAC4 was briefly expressed in the fiber cell
corners, possibly having a role in the initiation of lignification.
The establishment of the characteristic patterned secondary cell walls requires the
spatial restriction of lignification. Once in the apoplast, monolignols are highly
mobile (Lion et al., 2017; reviewed by Tobimatsu and Schuetz, 2019), while
fluorescence recovery after photobleaching (FRAP) analysis revealed that LAC4
is immobile, likely being anchored to the secondary cell wall (Schuetz et al.,
2014; Chou et al., 2018). PRXs are localized throughout the whole cell wall, but
their oxidative activity is specific to the lignifying regions (Hoffmann et al.,
2020). Presumably, the controlled production of reactive oxygen species and their
relative amount in the apoplastic region restrict PRX activity (Hoffmann et al.,
2020).
Altogether, it seems clear that the patterning of lignin deposition is determined
by the localization of the phenoloxidases and the availability of the monolignols
and the reactive oxygen species (Tobimatsu and Schuetz, 2019; Hoffmann et al.,
2020).
27
Figure 4. Monolignol transport and the formation of the lignin polymer. The
monolignols are moved outside the cell through passive diffusion across the plasma
membrane, by vesicle-mediated transport or by ABC transporters and proton-coupled
antiporters (PCA). O2 and H2O2 are the secondary substrates of phenoloxidases. O2
can freely diffuse from the air, while H2O2 is generated by the sequential activity of
NADPH oxidase and superoxide dismutase (SOD). Laccases and peroxidases
activate the monolignols to radicals. The free monolignol radicals react with a radical
in a growing lignin polymer via oxidative coupling.
Secondary substrates
The secondary substrates of the phenoloxidases, O2 and H2O2, are necessary for
the production of the lignin monomer radicals. O2 availability is not limited, since
oxygen can diffuse from the air and it is therefore present in the xylem sap
(Gansert, 2003). H2O2 has a very short half-life (10-9 – 10-3 s) (D’Autréaux and
Toledano, 2007), implying its continuous production in lignifying tissues. H2O2
is produced by the sequential activity of two enzymes: nicotineamide adenine
dinucleotide phosphate hydrogen oxidase (NADPH oxidase) and superoxide
dismutase (SOD) (Fig. 4) (Ogawa et al., 1997). NADPH oxidase catalyzes the
production of apoplastic superoxide from cytoplasmic NADPH (Sagi and Fluhr,
2006), and SOD derives O2 and H2O2 from the toxic superoxide radicals (Liochev
and Fridovich, 1994). In the xylem, H2O2 production was localized to tracheary
elements and parenchyma cells (Ros Barceló et al., 2005; Gómez Ros et al.,
2006). Furthermore, SOD was shown to be present in xylem vessels, fibers and
28
parenchyma cells (Karpinska et al., 2001), indicating that all xylem cell types are
a source of phenoloxidase secondary substrates.
Assembly of the polymer
The dehydrogenation of the monolignols by phenoloxidases is followed by cross
coupling to the growing lignin polymer. Presumably, the monomer-polymer
coupling reactions are the result of a redox shuttle system. Önnerud et al. (2002)
identified manganese as a radical carrier. Upon oxidation by phenoloxidases, the
manganese ions diffuse to the lignification site in the cell wall, where the
monolignols and the growing end of the lignin polymer are oxidized. Manganese
and calcium are both homogenously distributed in wood; the latter forms calcium
pectate at the lignification initiation sites, in the middle lamella and the cell
corners. Calcium was proposed to form calcium superoxide complexes that
oxidize the growing lignin polymer (Boerjan et al., 2003).
The structure of the highly branched lignin polymer forms by the cross-coupling
of monomer and oligomer radicals through condensed (C-C) linkages (5-5
biphenyl, β-β resinol, β-5 phenylcoumaran, β-1 diarylpropane) and non-
condensed (C-O-C) ether linkages (β-O-4 β-aryl-ether, 5-O-4 biphenyl ether)
(Terashima and Fukushima, 1988). The coupling of two monolignols,
dimerization, preferentially occurs at the β positions, resulting in β-O-4, β-β and
β-5 bonds. The β-O-4 ether bond is generated most frequently (Vanholme et al.,
2010). The formed dimer is then dehydrogenated in order to couple with the next
monomer radical. The gradual extension of the polymer with one radical is
through endwise polymerization. Most of the 5-5 and 5-O-4 bonds are formed by
coupling of preformed oligomers (Boerjan et al., 2003). 5-5 linkages can be
observed in the coupling of G-lignin oligomers (Vanholme et al., 2010). Lignin
with high G unit content is more highly crosslinked, more rigid and more
hydrophobic than lignin rich in S units. Interestingly, if only β-O-4, β-β and β-5
bonds were present in the polymer, it would result in a completely linear structure
(Bonawitz and Chapple, 2010). The abundance of the various linkage types is
highly dependent on species, on the availability of each lignin subunit and on the
cell type. Also, the polymer length is determined by the availability and reactivity
of different lignin monomer radicals (Ralph et al., 2008). In Populus tremula x
alba the average length of a linear lignin chain may be between 13 and 20 units
(reviewed by Vanholme et al., 2010).
The initiation site for lignin polymerization is generally at the cell corner of the
middle lamella and in the S1 region of the secondary cell wall. Several hypotheses
29
have been put forward about what defines the exact location where
polymerization begins. Dirigent proteins were proposed to be the landmarks for
lignin initiation sites (Davin and Lewis, 2000). Currently ENHANCED
SUBERIN1/DIRIGENT10 (ESB1/DIR10) is the only dirigent protein which has
been shown to be required for the formation and lignification of the Casparian
strip (Hosmani et al., 2013). In grasses ferulates, wall-bound phenolics, can
dimerize or polymerize by forming ester-to-ether linkages to cross-link lignins
and polysaccharides (Ralph et al., 1995). These phenolics can provide initiation
sites for lignin polymerization (reviewed by Ralph et al., 2004). Furthermore
glycine-rich proteins and pectin-bound peroxidases were also postulated to form
nucleation sites (Boerjan et al., 2003).
3.4.4. Cell-autonomous vs. non-cell-autonomous lignification
Lignin biosynthesis and deposition are tightly regulated but at the same time
flexible and dynamic processes, occurring in response to specific developmental,
metabolic or environmental changes. A better understanding of the mechanisms
regulating lignification is necessary to enable lignin engineering (Ragauskas et
al., 2014). Lignification of certain cell types can occur in cell-autonomous as well
as non-cell-autonomous fashion (Barros et al., 2015). It was suggested early on
that lignification of the vessel walls continues after their cell death since they
differentiate and die rapidly to become functional in water transport (Stewart,
1966; Donaldson, 2001). The post-mortem, non-cell-autonomous lignification
would require the cooperative action of vessel-neighbouring, living cells. In line
with this theory, several genes encoding for enzymes from the lignin biosynthetic
pathway are not exclusively expressed in TEs, but also in the cambium and the
xylem parenchyma cells (Hu et al., 1998; Chen et al., 2000; Lacombe et al., 2000;
Lauvergeat et al., 2002; Baghdady et al., 2006; Blokhina et al., 2019).
Additionally, metabolomics analysis of Norway spruce tracheids and ray
parenchyma cells revealed the presence of monolignols and glycoconjugates in
both cell types (Blokhina et al., 2019). The xylogenic Zinnia elegans cell culture
system, containing a mixture of TEs and non-lignifying parenchymatic cells, has
been practical to dissect the cooperative aspect of lignification. The dead TEs in
this in vitro system continued to lignify thanks to the action of the living
parenchymatic cells (Hosokawa et al., 2001; Pesquet et al., 2013). Manipulation
and analyses of the cell culture media revealed that monolignols are supplied to
the TEs from the parenchymatic cells through the medium (Hosokawa et al.,
2001). Lignin-biosynthetic genes are expressed in the parenchymatic cells which
therefore can both synthesize and transport monolignols and reactive oxygen
30
species to the TEs (Pesquet et al., 2013). The limitation of the cell culture system
is the artificial setup and the lack of developmental and tissue context.
Smith et al. (2013) demonstrated in planta that vessel lignification is a
cooperative process where the lignin monomers are supplied to the lignifying
cells by the neighbouring cells. Lignin biosynthesis was suppressed in lignifying
xylem cells by an artificial microRNA targeting CCR1 driven by the CESA7
promoter. Still, the vessel elements lignified normally, thanks to the contribution
of the neighbouring xylem parenchyma cells. However, when lignin biosynthesis
was specifically blocked in the parenchyma cells the vessels exhibited collapsed,
irregular xylem phenotype (Smith et al., 2017). The cell wall of these partially
collapsed vessels was not completely devoid of lignin, suggesting that the vessels
initiate cell-autonomous lignification while they are still alive. Moreover, non-
native monolignol conjugates were expressed in the parenchyma cells and these
compounds were incorporated into the cell wall (Smith et al., 2017). Later in stem
development xylary fibers also contribute lignin precursors to vessel lignification
but less than the parenchyma cells, and this lignin pool is not essential for proper
vessel formation (Smith et al., 2017). Xylary fibers also have the ability to
undergo non-cell-autonomous lignification, while interfascicular fibers fully
lignify in a cell-autonomous manner (Smith et al., 2013). The vessel-
neighbouring fibers of Populus tremula exhibited lignin composition
intermediate between that of vessels and fibers, having relatively high levels of
G lignin units (Gorzsás et al., 2011). It is conceivable that the function of the
xylem cell and its tissue context determines the type of lignification the cell
undergoes.
Early in the evolution many plant taxa did not contain parenchyma cells within
the xylem, possibly relying exclusively on cell-autonomous lignification. The
ability of the parenchymatic cells to cooperatively supply lignin precursors to the
water-transporting vessels may be a more recently evolved trait (Smith et al.,
2017). However, the evolution of the rays might not be the only determinant of
non-cell-autonomous lignification, since some of the fibers also have the ability
to take part in the process. Altogether, the reason for the appearance of non-cell-
autonomous lignification is not clear. This developmental strategy of the land
plants could have evolved to cope with the increasing tension pressure created by
water transport and with the increase in the height and weight of the growing
plant body. Continuous fortification of the cell walls throughout the plant’s
lifespan certainly allows more efficient and undisturbed flow of the xylem sap
through the vascular system. Furthermore, non-cell-autonomous lignification
31
could enrich the ability of xylem elements to adapt to changing environment and
pathogens.
3.4.5. Lignin as a resource for biotechnological applications
The secondary cell walls are a rich source of lignocellulosic plant biomass, with
the potential to partially replace fossil fuel usage. The effectiveness at which such
biomass pool can be utilized depends greatly on the chemical properties and
ultrastructure of the cell wall (Meents et al., 2018). Currently, the large-scale
industrial use of plant biomass is limited due to high processing costs. Lignin is
the major barrier to efficient extraction of cellulose fibers and to saccharification
for production of pulp and liquid biofuel, respectively (Baucher et al., 2003; Zeng
et al., 2014), and its removal is pricey, laborious and environmentally hazardous.
Lignin also curtails the nutritional value of crops meant for human consumption
or livestock feed (Wang and Frei, 2011). Unfortunately, extreme environmental
conditions, which will become more common due to climate change, enhance
lignin biosynthesis (Moura et al., 2010; Ployet et al., 2018; Behr et al., 2019).
Biotechnological modifications aim to improve lignocellulose utilization by
increasing the calorific values, the extractability of cell wall polymers, as well as
saccharification and fermentation yields (Boerjan et al, 2003; Weng et al., 2008;
Mansfield, 2009; Novaes et al., 2010; Vanholme et al., 2010). In field crops,
lignin content could be decreased by traditional breeding methods, natural and
induced mutations, and transgenesis. Ideally, one would like to reduce the content
of the lignin polymer to a level where efficient saccharification could be achieved
without pretreatment. Notably, the pretreatment of the lignocellulosic biomass
imposes major environmental hazards. However, the straightforward reduction of
the total lignin content by genetic engineering has frequently led to reduced
overall biomass production even under controlled growth conditions (Piquemal
et al., 1998; Franke et al., 2002; Leplé et al., 2007; Voelker et al., 2010).
Typically, reduction of lignin affects negatively plant fitness and productivity
(Wagner et al., 2009). Nonetheless, it was demonstrated in Arabidopsis, tobacco
(Nicotiana tabacum) and Populus tremula x alba that changes in the ratio of
certain lignin units do not compromise plant performance and the water-
conducting capacity of the vessel elements. Specifically, increased S/G ratio
could improve the bioprocessing properties of wood without having a negative
influence on plant growth. (Franke et al., 2000; Huntley et al., 2003; Sibout et
al., 2002; Stewart et al., 2009; Studer et al., 2011). The methoxy groups of S-
lignin present at the 3 and 5 positions of the aromatic ring enhance reactivity to
32
thermal and catalytic fragmentation. In the native lignin polymer, the β-ether
bonds are the most labile, and their frequency is higher in the lignin polymer
when there is increased S unit content. Consequently, the S unit rich lignin has
less 5-5 and β-5 bonds, which makes it more easily extractable (Mahon and
Mansfield, 2019). Increasing the S/G ratio of the lignin polymer is however not
feasible in softwood species which do not have S-type lignin. Nevertheless,
Wagner et al. (2015) demonstrated that S-type monolignol units could be
generated in Pinus radiata by metabolic engineering of the phenylpropanoid
pathway. Furthermore, esther linkages were introduced in Populus alba ×
grandidentata by expressing a non-native monolignol transferase; this
modification improved the digestibility of the lignin polymer (Wilkerson et al.,
2014). Altogether, the plasticity of the lignin biosynthetic pathway is a promising
asset to generate easily exploitable biomass.
In the xylem of Arabidopsis, along with most angiosperms, fibers contain the
bulk of the secondary cell walls. In Arabidopsis the lignin content of the
interfascicular fiber cell walls can be significantly reduced without affecting plant
fitness. Growth is not at all or only hardly compromised as long as vessels lignify
normally (Smith et al., 2017). Using synthetic biology tools Yang et al. (2013)
managed to direct lignin biosynthesis to vessels and increase the polysaccharide
deposition in the fiber cell walls. This combined strategy resulted in higher sugar
yields. Tissue-specific alteration of the lignin profile, with the aim to maintain
vessel integrity, is a feasible biotechnological approach for biomass amelioration.
The biosynthesis of an alternative lignin monomer, curcumin, was successfully
introduced into Arabidopsis, resulting in a polymer more prone for chemical
changes. Plants containing curcumin and phenylpentanoids in their cell walls
exhibited normal growth and greatly increased saccharification efficiency,
indicating the potential of non-native monomers as a way to improve biomass
properties (Oyarce et al., 2019).
Likely, crosslinking and interaction between lignin and the cell wall carbohydrate
matrix limits the sugar release efficiency after enzymatic saccharification (Mahon
and Mansfield, 2019). For instance, xylan might form linkages with lignin via the
methyl-glucuronic acid ([Me]GlcA) branching groups, since the genetic removal
of [Me]GlcA lead to increased xylose and glucose release upon enzymatic
hydrolysis (Lyczakowski et al., 2017). Such findings demonstrate that
downstream applications may be eased by genetically distancing the cell wall
polymers from each other.
33
Lignin is often referred to as the recalcitrant in the cell wall we all want to get rid
of, but this rudimentary view should be revised. Lignin is also a highly attractive
polymer due to its high energy content and structure that could be utilized. A
sustainable and profitable biorefinery concept requires maximal and diversified
use of the feedstock (Liao et al., 2020). The approaches to turn lignin into a
resource include the generation of homogenous and linear polymers consisting
exclusively of caffeyl alcohol (C-lignin) for carbon-fiber production, introduction
of valuable novel monomers like tricin or increasing the aldehyde group content
(Ralph et al., 2019; Mahon and Mansfield, 2019). Tricin is a flavonoid which can
only function as polymerization initiator, and it can be used to generate shorter
polymers (Lan et al., 2015). Aldehyde-rich lignin has less non-covalent
interactions with hemicelluloses and in turn it is more suitable for downstream
applications, such as saccharification (Zhao et al., 2013; Carmona et al., 2015;
Van Acker et al., 2017).
Recent advances in lignin valorization include its use in thermoplastics,
blends/composites, as carbon fiber precursors, heavy metal absorbents and
nanoparticles, as wells as the incorporation of lignin-based copolymers into
biomedical materials (reviewed by Wang et al., 2020). These emerging fields
provide diverse novel opportunities for the utilization of the lignocellulosic
biomass.
Plant biomass represents a valuable and multiuse resource. Enhancing the value
and usability of lignin by exploiting natural variations and the potentials of
genetic engineering may enable the establishment of sustainable biorefineries.
34
Research aims and objectives
The aim of my thesis is to identify and characterize novel regulators of
lignification, using Arabidopsis thaliana as a model system. Lignification is a
multistep process, and the focus of these investigations is on the biosynthesis of
lignin monomers. The overall goal was to address the regulation of lignin
biosynthesis from previously less-explored angles, such as non-cell-autonomous
lignification and the timing of lignin biosynthesis, and provide new molecular
tools for the field.
To address the research goal, a variety of methods was employed, such as cell
wall chemistry analyses, genetic, transcriptomic, microscopic and
microspectroscopic tools. Consequently, three potential regulators of
lignification were identified. In Paper I the first known molecular regulator in
non-cell-autonomous lignification is revealed. To investigate the molecular
network of this novel regulator a large-scale protein-interactor screen was
performed (Paper II). The screen yielded several candidate genes, and based on
cell wall chemistry analysis and literature studies two of them were selected for
further investigation. The candidates were characterized in detail in connection
to lignin biosynthesis (Paper II, Paper III). These findings will help to better
understand secondary cell wall development, provide knowledge pool which can
be transferred into economically important species and potentially be used for
biotechnological purposes in the future.
35
Results and discussion
PIRIN2 is a non-cell-autonomous regulator of S-type lignin
accumulation
A PIRIN (PRN) protein was earlier suggested to play a role in non-cell-
autonomous lignification in Zinnia elegans xylogenic cell cultures (Pesquet et al.,
2013). The concept of non-cell-autonomous lignification implies that xylem
elements are assisted by their neighbouring cells to get their cell walls fully
lignified (Hosokawa et al., 2001; Pesquet et al., 2013; Smith et al., 2013; De
Meester et al., 2018). While this theory has been around for many years, the
molecular players have remained unknown. The initial findings by Pesquet et al.
(2013) prompted us to investigate the PRN gene family and their involvement in
lignification in Arabidopsis.
Promoter activity analyses revealed that out of the four Arabidopsis PRN genes,
PRN2, PRN3 and PRN4 are active in the xylem of seedlings and mature plants
(Paper I, Fig. 1). The PRN2 promoter exhibited a very specific pattern restricted
to the xylem cells neighbouring the vessel elements. Similarly, the promoter of
the PRN2 homologue in Populus was active in the ray cells next to the vessel
elements (Paper I, Fig. S2). The localization of PRN2 next to the vessel elements
was confirmed by confocal laser scanning microscopy imaging of the PRN2:GFP
fusion protein (Paper I, Fig. 2). The promoter activity and protein localization
results revealed that PRN2 is present in the cells surrounding the vessel elements,
but not in the vessel elements themselves. These provided the first indication that
PRN2 is likely to be involved in vessel differentiation or function. Since vessel
elements undergo programmed cell death early on, they might need the support
of neighbouring cells for complete differentiation or to function properly in
different processes.
Next, we performed pyrolysis-gas chromatography/mass spectrometry (py-
GC/MS) on a set of prn single and double mutants to assess their possible role in
lignification (Paper I, Fig. 3). Besides determining total lignin content, py-
GC/MS is also able to distinguish between G-, S- and H-type lignin, which are
important indicators of the processability of the cell wall material. All prn2
mutants exhibited increased S-type lignin content and consequently significantly
higher S/G-type lignin ratio (Paper I, Fig. 3, 4). The effect of PRN2 on the S-
type lignin content of the secondary cell walls was confirmed by two-dimensional
nuclear magnetic resonance spectroscopy (Paper I, Fig. 4).
36
Upon observing the vessel-neighbouring expression pattern of PRN2 and
chemical changes at the whole tissue level, we set on to investigate the lignin
profile on a cellular level. Raman and FT-IR microspectroscopy allowed in situ,
high-resolution chemotyping of individual cell walls (Gorzsás et al., 2011; 2017)
of vessels and fibers next to the predicted PRN2-expressing cells in prn2. The
microspectroscopic analyses revealed chemical changes that correlated with S-
type lignin, G-type lignin, lignin content and with structural properties of the
lignin polymer in the hypocotyl and in the vascular bundle of the inflorescence
stem of the prn2 mutants (Paper I, Fig. 5). Smith et al. (2017) demonstrated that
interfascicular fibers lignify predominantly in a cell-autonomous manner.
Interestingly, the cell wall chemistry of the interfascicular fibers was not altered
in prn2, suggesting the absence of non-cell-autonomous lignification in this
region, or that PRN2 does not play a role there. Altogether, these findings confirm
that PRN2 controls the non-cell-autonomous lignification of xylem elements.
Next, we were curious to address the mode of function of PRN2. The human PRN
protein interacts with several different transcription factors and acts as a
transcriptional co-regulator (Wendler et al., 1997; Dechend et al., 1999; Liu et
al., 2013B). The subcellular localization of PRN2 to the nucleus and the
cytoplasm was demonstrated earlier (Zhang et al., 2014). Thus, we hypothesized
that PRN2 might regulate S-type lignin biosynthesis by the transcriptional control
of lignin-biosynthetic genes. Notably, several lignin-biosynthetic genes and
secondary cell wall-related transcription factors had increased expression in prn2
mutants, and the PRN2-overexpressing lines showed opposite tendencies for
many genes (Paper I, Fig. 6). F5H1 encodes the enzyme required for S-type
lignin biosynthesis. Interestingly, out of all genes analyzed F5H1 displayed the
most consistent changes in expression, with significantly higher expression level
in the prn2 mutants and lower in the PRN2-overexpressing lines. In conclusion,
the gene expression analyses uncovered the function of PRN2 as a transcriptional
suppressor of lignin-biosynthetic genes and secondary cell wall-related
transcription factors.
PRN2 had the most consistent and pronounced effect on S-type lignin
accumulation. Presumably, PRN2 suppresses S-type lignin biosynthesis by
regulating the expression of F5H1, a key S-type lignin-specific biosynthetic gene.
This, together with the vessel-neighbouring expression of PRN2, suggest that
PRN2 is a molecular factor ensuring that vessel elements acquire their typical, G-
type lignin enriched cell walls which are more resistant to the mechanical forces
created by water transport.
37
HISTONE MONOUBIQUITINATION2 (HUB2) affects the
lignin composition of xylem vessels
PRN2 that was described as a suppressor of S-type lignin accumulation in
is not a transcription factor, and has to interact with other proteins in the
regulation of lignin-biosynthetic genes. To identify the molecular partners of
PRN2, we performed a large-scale yeast two-hybrid screen. We retrieved 18
protein interactor candidates (Paper II, Dataset 1 available online at Dryad).
To identify interactors which affect lignin content or composition, py-GC/MS
analysis was performed on T-DNA insertion mutants of the PRN2 interactor
genes. From all genotypes analyzed, the two knockout lines of HUB2 exhibited
significantly lower S-type lignin level and increased G-type lignin content (Paper
II, Fig. 1). These findings implied a previously unreported, lignin-related
function for HUB2.
HUB2 is an E3 ubiquitin ligase catalyzing the histone H2B monoubiquitination
of the chromatin (H2Bub1). HUB2 is involved in various plant developmental
processes, e.g. flowering time (Cao et al., 2008; Xu et al., 2009), circadian clock
(Himanen et al., 2012) and pathogen resistance (Zou et al., 2014; Zhang et al.,
2015; Zhao et al., 2020). Cotton GhHUB2, the functional homolog of AtHUB2,
was implicated in secondary cell wall formation and lignification (Feng et al.,
2018). Based on the PRN2-HUB2 protein interaction finding, we addressed the
possible role of HUB2 in lignification in Arabidopsis.
The protein interaction between PRN2 and HUB2 was confirmed by both in vitro
and in vivo approaches (Paper II, Fig. 2). Next, we analyzed the promoter
activity of HUB2 in the xylem of mature stems and hypocotyls. The HUB2
promoter was active in different xylem cell types, including the PRN2-expressing
cells (Paper I, Fig. 1J-L), but was absent from the vessel elements (Paper II,
Fig. 3C-E).
The initial mutant screen was followed up by genetic interaction study through
py-GC/MS analysis of hypocotyl samples of hub2 and prn2 single, and hub2 prn2
double mutants (Paper II, Fig. 3). Unlike the hub2 single mutants, the hub2 prn2
double mutants exhibited higher S-type lignin content compared to WT,
suggesting that prn2 is epistatic to hub2. The G-type lignin content of hub2 and
the hub2 prn2 double mutants was significantly higher compared to WT, while
the effect of prn2 on G-type lignin content was varying. The opposite effect of
HUB2 and PRN2 on S-type lignin accumulation was confirmed in mature parts
of the inflorescence stem (Paper II, Fig. 4).
38
To elucidate the effect of HUB2 on individual xylem cell walls, Raman and FT-
IR microspectroscopic chemotyping of hub2 mutant hypocotyls was performed
(Paper II, Fig. 5, Fig. S2). The in situ chemical profiling corroborated the py-
GC/MS results obtained at the whole tissue level, revealing elevated levels of G-
type lignin, and consequently lower S/G ratios. Altogether, these findings
indicate that HUB2 promotes S-type lignin accumulation at the expense of G-
type lignin.
HUB2 is part of the enzyme complex mediating H2Bub1 chromatin
modifications in Arabidopsis. Hence, we hypothesized that HUB2, together with
PRN2, affect lignification by mediating the H2Bub1 levels of lignin-biosynthetic
genes. Strikingly, the ChIP-qPCR profiling of prn2 did not reveal differences in
H2Bub1 levels of selected lignin-biosynthetic genes (Paper II, Fig. S3). These
results implied that PRN2 and HUB2 act through an H2Bub1-independent
mechanism (Paper I). To address the effect of HUB2 and PRN2 on gene
expression, transcriptomic analysis was performed on mature hypocotyls of hub2
and prn2 single, and hub2 prn2 double mutants (Paper II, Fig. 6). Since PRN2
and HUB2 had distinctly opposite effect on S-type lignin content, we focused on
the transcriptional changes which could underlie such cell wall chemistry
changes. Strikingly, F5H1 which encodes the enzyme specific for S-type lignin
biosynthesis was up-regulated in prn2 and hub2 prn2, but down-regulated in
hub2. These changes in F5H1 expression correlated with the S-type lignin content
of the genotypes. Taken together, our results suggest that PRN2 and HUB2
modulate S-type lignin levels through the transcriptional regulation of F5H1. The
protein interaction and genetic analyses indicate that PRN2 and HUB2 function
together, whereby PRN2 acts downstream of HUB2 in controlling S-type lignin
accumulation.
HUB2 is likely to stimulate S-type lignin biosynthesis in most xylem cells, except
vessel elements, where it is not expressed (Brady et al., 2007). In contrast, the S-
type lignin-suppressor role of PRN2 is concentrated around the vessel elements.
It seems therefore that PRN2 interacts with HUB2 to prevent its S-type lignin
stimulatory function in the neighbourhood of the vessels to ensure their correct
lignin composition.
39
PIB is a potential modulator of the diurnal timing of lignin
biosynthesis
In Arabidopsis xylem, PRN2 suppresses S-type lignin accumulation in a non-cell-
autonomous manner (Paper I) through a mechanism that is at least in part
mediated by suppression of HUB2 function (Paper II). PRN2, however, interacts
with several other proteins (Paper II) which might also participate in the
regulation of vessel lignification. One of these PRN2 interacting proteins is the
CRYPTOCHROME2-interacting basic helix-loop-helix3 (CIB3)/PRN2-
interacting basic helix-loop-helix (PIB) transcription factor (Paper II). The
human PRN protein interacts with transcription factors and functions as a
transcriptional co-regulator, suggesting that PRN2 might interact with PIB in the
regulation of lignin-biosynthetic genes. In addition, the paralog of CIB3,
CIB2/PIB-LIKE, was previously implicated in secondary cell wall formation
(Oikawa et al., 2010). Thus, we addressed the role of PIB in the regulation of
lignification in Arabidopsis.
First, we confirmed the protein interaction between PRN2 and HUB2 in vitro by
yeast growth and enzymatic assays, and in vivo by co-immunoprecipitation assay
(Paper III, Fig. 1). Next, the promoter activity of PIB was analyzed in 5-day-old
seedlings and in 6-week-old plants (Paper III, Fig. 2). The PIB promoter was
active in different tissues throughout the plant, for example in the parenchymatic
cells around the vessel elements in the phase I secondary xylem and in the
vascular tissues of the leaves. Protein localization analysis by confocal laser
scanning microscopy revealed the presence of PIB in the nucleus (Paper III, Fig.
2F). The results of the promoter activity and protein localization analyses imply
the involvement of PIB in a variety of plant developmental processes, including
vascular development.
PRN2 suppresses S-type lignin accumulation (Paper I). To assess the effect of
PIB on lignification, lignin content and composition were determined. Neither
py-GC/MS analysis at the whole tissue level (Paper III, Fig. 3), nor FT-IR
microspectroscopy at the cellular level revealed any changes in the lignin profile
of pib cell walls (Paper III, Fig. S3). Since PIB-LIKE is co-regulated with xylan
biosynthetic genes (Oikawa et al., 2010), the monosaccharide composition of the
pib secondary cell walls was also determined (Paper III, Fig. S4), but no
significant changes could be detected.
Next, gene expression was analyzed by RNA-sequencing of pib-1, pib-like-1,
prn2-2, pib-4 pib-like-2 and pib-1 prn2-2 mutant lines at two timepoints, night
40
(darkness) and day (light) (Paper III, Fig. 4). Two timepoints were selected to
capture any light-dependent changes in gene expression since PIB and PIB-LIKE
interact with the CRY2 photoreceptor (Liu et al., 2013C). The expression of
lignin-biosynthetic genes was significantly higher in pib during the night.
Furthermore, differentially expressed genes (DEGs) belonging to the circadian
rhythm, plant-type cell wall organization and response to blue light gene ontology
(GO) terms were overrepresented in pib during the day. Taken together, it is
plausible that PIB modulates lignin biosynthesis in a diurnal fashion, potentially
to synchronize lignin biosynthesis with resource availability.
Next, we followed the diurnal expression of lignin-biosynthetic genes over a 24-
hr cycle in the roots of the pib mutant. The diurnal expression of C3H1 was
slightly delayed in pib, but the difference was not significant compared to the
Col-0 WT. The expression level of PAL1 and 4CL1 was significantly lower in
pib during the night, but the other lignin-biosynthetic genes did not exhibit
significant changes in their expression (Paper III, Fig. 5). These results suggest
that PIB has only minor effects on the diurnal expression profile of lignin-
biosynthetic genes. However, it is plausible that alterations on a few genes, such
as C3H1, influence the overall timing of lignin biosynthesis. In addition,
functional redundancy between PIB and PIB-LIKE as well as tissue-specific
function complicate the monitoring of any transcriptomic changes at the whole
organ level. Nevertheless, to test the ability of PIB to regulate lignin-biosynthetic
genes, we performed transient expression analysis in tobacco protoplasts. The
transactivation assay demonstrated that PIB can suppress the promoter activity of
most lignin-biosynthetic genes to varying degrees (Paper III, Fig. 6).
In conclusion, PIB, the protein interactor of PRN2, modulates the diurnal
expression of lignin-biosynthetic genes via transcriptional repression. The exact
mechanism and the molecular network of PIB as a diurnal transcriptional
modulator remain to be elucidated. Nonetheless, we propose that PIB affects the
diurnal timing of lignin biosynthesis via transcriptional repression.
41
Conclusions and perspectives
Even though lignification of the plant secondary cell walls has been extensively
studied for decades novel molecular players involved in the process are
continuously identified. This work uncovered three proteins which influence
lignin biosynthesis at the transcriptional level and constitute novel aspects in the
regulation of lignification.
First, PRN2 was identified and characterized as the first known molecular
regulator of non-cell-autonomous lignification in Arabidopsis. PRN2 is
expressed in the cells around the vessel elements and affects the cell wall
composition of the neighbouring vessels and fibers. The promoter activity of the
ortholog in hybrid aspen, PttPRN2, is also next to the vessel elements, suggesting
a similar role for PRN2 in trees. By modulating the expression of PRN2 lignin
composition can be altered without compromising plant health and fitness. This
makes PRN2 and non-cell-autonomous lignification promising tools for
biotechnology. For example, the restriction of lignin biosynthesis to the PRN2-
expressing cells might result in low-lignin lignocellulosic biomass without a yield
penalty.
Furthermore, the molecular network of PRN2 was broadened by the
characterization of two protein interactors. The chromatin modifying enzyme
HUB2 promotes S-type lignin biosynthesis. The exact mechanism of HUB2
function in lignin biosynthesis remained unclear but it did not seem to include
chromatin modification. Single-cell and cell-type-specific techniques will enable
further investigation of the mechanism of PRN2 and HUB2 in lignification.
Finally, we report that the diurnal timing of lignin biosynthesis is modulated by
the PRN2-interacting PIB protein. The diurnal regulation of lignin biosynthesis
was previously unexplored, here we present the first potential modulator of this
process. The effect of PIB on the gene expression changes of lignin-biosynthetic
genes could be light-dependent. The exact molecular mechanism needs to be
elucidated in the future, addressing the possible functional redundancy between
PIB and its paralog PIB-LIKE.
HUB2 and PIB are likely to be part of two independent, PRN2-containing protein
complexes. It is plausible that the activity of the complexes is spatially or
temporally separated. However, the possibility that all three proteins belong to a
single complex cannot be excluded. The finding of additional molecular partners
would help uncover the exact mechanism and makeup of the PRN2-dependent
42
protein complex(es). In Arabidopsis gene duplication events within the PRN gene
family and the sub-family containing PIB and PIB-LIKE, may have contributed
to functional redundancy, thus hampering molecular genetics studies.
Interestingly, in hybrid aspen there are only two PRN genes and one PIB/PIB-
LIKE ortholog. Using hybrid aspen to further study these genes could help to
counteract redundancy, while also gaining understanding on the regulation of
lignification in economically more relevant species.
Figure 5. PRN2, HUB2 and PIB regulate lignin biosynthesis.
In the PRN2-expressing cells next to the vessel elements, PRN2 suppresses the
expression of lignin-biosynthetic genes, and thus monolignol biosynthesis. In
particular, PRN2 negatively regulates S-type lignin biosynthesis by blocking the
promoting effect of HUB2 on F5H1 expression. In the fiber cells where only HUB2
is expressed (PRN2 is not), HUB2 stimulates S-type lignin biosynthesis through the
transcriptional regulation of F5H1. PIB modulates the diurnal expression of the
lignin-biosynthetic genes. Whether PRN2, HUB2 and PIB are part of a single protein
complex or two separate complexes is open for future research.
43
Acknowledgements
My dear PhD supervisor, Hannele Tuominen, I am very grateful to you for these
past five years. It has been a bumpy road, as it is always the case in scientific
research, but the finish line is here. I want to thank you the freedom you gave me
to follow my curiosity, but also taking on the guidance when there was no space
to wonder. I very much appreciate your patience, support, sense of humor and
caring.
I am thankful for the two supervisors I had during my master’s projects: Delphine
Gendre and Kristoffer Jonsson. I learnt a lot from both of you, the time spent
under your guidance prepared me for my PhD research.
I want to thank my colleagues who worked with me in shared projects, or who
helped me in any other way. Thanks to Bo Zhang, Pál Miskolczi, Sacha
Escamez, Carolin Seyfferth, Yin Wang and Maxime Chantreau for all your
expertise, support, and patience.
Furthermore, I want to acknowledge all members of the HT group, Angela,
Anna, Bernard, Caro, Hardy, Isura, Marta, Max, Mikko, Sacha, Shruti and
Yin. I am really grateful for the helpful, supportive, friendly and fun atmosphere
of our group.
I want to thank the three project students I got to work with over the years, Inkeri
Soppa, Hannah Ohm and Sara Ölmelid. I really appreciate all your hard work.
I am grateful to the members of my reference group, László Bakó, Markus
Schmid, Maria Eriksson, Rishi Bhalearo and Urs Fischer for their advises and
that they followed my PhD journey.
A significant amount of time and energy was spent on teaching at bachelor’s and
master’s level courses. I really enjoyed these challenges, but it would not have
been the same without my amazing teaching partners. Thanks to Bernard
Wessels, Daria Chrobok, Jean Claude Nzayisenga, Kerstin Richau, Mikko
Luomaranta and Yan Ji.
Thanks to Junko Takahashi-Schmidt for all the help with the various cell wall
chemistry analyses, and very importantly for her endless patience, hard work and
for the fun moments we shared.
44
I want to express my gratitude to Jan Karlsson, Janne, for all his technical
support and patience. I apologize if my questions were sometimes close to drive
you crazy.
I am very grateful for the work of the personnel in the greenhouse and in the
transgenic facility. Without them our research would not progress, their work is
essential and much appreciated.
I am really glad that we kicked off the Greener UPSC group and I hope that it
will be a long-lasting movement. Thanks to Anne B, Sonja, Nico, Sara R,
Domenique, Alex, Noemi, Lill and Loïc for joining in and I wish you all the
best.
I want to wish everyone at UPSC happy and successful years to come. In my
opinion UPSC is special, since everybody contributes in some way for the
betterment of the work environment, be it greenhouse issues or afterwork events.
Thanks for everything!
To all my friends, in Umeå, in Sweden, in my home and in other corners of the
world: perhaps sometimes unknowingly, but you guys helped me so much along
the way. I am very grateful that I have you in my life.
My special thanks to Karolin, Julia, Hanna néni, Eszter and Szabi for the times
we shared in Umeå and everything else.
I cannot thank enough to my extended family for all their love, support and so
much more.
Az én nagy családom, Annamária, Nénje, Robi, Ancsi és Dóri, tudom azt, hogy
nélkületek ez nem sikerült volna. Nagyon hálás vagyok mindenért.
I want to thank from the bottom of my heart to my parents, who always let me
follow my own path. You are my role models. You have made a difference in so
many people’s life, I am very proud of you.
Drága szüleim! Szívből köszönöm, hogy mindig hagyjátok, hogy a saját utam
járjam. Ti vagytok a példaképeim. Olyan sok emberen segítetek, nagyon büszke
vagyok rátok. Köszönöm, hogy ti mindig bíztok bennem, támogattok és velem
vagytok. Remélem, hogy ez még nagyon sokáig így lesz.
My husband, Anirban, you shared every day of this journey with me, literally
from day one. You made me laugh when things were not great, or when they
were. I learnt so much from your wisdom, professionalism, and straight talk. You
made these years really special, thank you so much for everything!
45
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