understanding and manipulating primary cell walls in plant...
TRANSCRIPT
Understanding and manipulating primary cell
walls in plant cell suspension cultures
Felicia Leijon
Doctoral Thesis in Biotechnology
School of Engineering Sciences in Chemistry, Biotechnology and Health
Royal Institute of Technology
Stockholm, Sweden
2019
© Felicia Leijon, Stockholm, 2019
TRITA-CBH-FOU-2019:2
ISBN 978-91-7873-074-2
KTH School of Engineering Sciences in Chemistry, Biotechnology and Health
Royal Institute of Technology
AlbaNova University Center
106 91 Stockholm
Sweden
Printed by Universitetsservice US-AB, Stockholm 2019
i
Abstract The cell wall is required for many aspects of plant function and
development. It is also an accessible and renewable resource utilized both
in unrefined forms and as raw material for further development.
Increased knowledge regarding cell wall structure and components will
contribute to better utilization of plants and the resources they provide.
In this thesis aspects of the primary cell wall of Populus trichocarpa and
Nicotiana tabacum are explored.
In Publication I a method for isolation and biochemical
characterization of plant glycosyltransferases using a spectrophotometric
or a radiometric assay was optimized. The radiometric assay was applied
in Publication II where the proteome of the plasmodesmata isolated
from P. trichocarpa was analyzed. Proteins identified belonged to
functional classes such as “transport”, “signalling” and “stress responses”.
Plasmodesmata-enriched fractions had high levels of callose synthase
activity under ion depleted conditions as well as with calcium present.
The second part of the thesis comprises the alteration of the cell wall of N.
tabacum cells and A. thaliana plants through in vivo expression of a
carbohydrate binding module (CBM) (Publication III). In tobacco this
resulted in cell walls with loose ultrastructure containing an increased
proportion of 1,4-β-glucans. The cell walls were more susceptible to
saccharification, possibly due to changes in the structure of cellulose or
xyloglucan. Arabidopsis plants showed increased saccharification after
mild pretreatment, suggesting that heterologous expression of CBMs is a
promising method for cell wall engineering. In Publication IV cellulose
microfibrils (CMFs) and nanocrystals (CNCs) were extracted from the
transgenic cells. CNC preparation resulted in higher yields and longer
CNCs. Nanopapers prepared from the CMFs of the CBM line
demonstrated enhanced strength and toughness. Thus, changes to the
ordered regions of cellulose were suggested to take place due to CBM
expression.
Keywords: Callose synthase, carbohydrate-binding module, cell wall
engineering, cellulose microfibril, cellulose nanocrystal,
glycosyltransferase, mass spectrometry, plasmodesmata, Populus,
primary cell wall, radiometric assay, spectrophotometric assay.
ii
Sammanfattning
Cellväggen har en avgörande roll för många aspekter av funktion och
utveckling hos växtcellen. Den utgör en tillgänglig och förnybar resurs
både i sin obearbetade form och som råmaterial för vidare förädling.
Ökad kunskap rörande cellväggens struktur och sammansättning skulle
därför bidra till bättre nyttjande av växter och de råvaror de
tillhandahåller. I den här avhandlingen undersöks olika aspekter av den
primära cellväggen hos Populus trichocarpa och Nicotiana tabacum.
I Publikation I optimeras en spektrofotometrisk och en radiometrisk
metod för biokemisk karakterisering av gykosyltransferaser från växter.
Den radiometriska metoden tillämpas sedan i Publikation II där
proteomet hos plasmodesmata från P. trichocarpa celler kartlades.
Identifierade proteiner tillhörde funktionella klasser så som ”transport”,
”signalering” och ”stress”. Plasmodesmata anrikade membranfraktioner
hade förhöjda nivåer av kallos-syntas aktivitet både vid frånvaro av joner
och med kalcium.
Den andra delen av avhandlingen består av in vivo uttryck av en
kolhydratbindande modul (CBM) i tobak och Arabidopsis plantor för att
introducera förändringar i cellväggen (Publikation III). Detta
resulterade i cellväggar med porös ultrastruktur innehållande mer 1,4-β-
glukaner. Cellväggarna var lättare att bryta ner till kolhydratmonomerer
vilket kan bero på att strukturen hos cellulosa eller xyloglukan förändrats.
Arabidopsis plantor var också lättare att bryta ner efter en mild
förbehandling vilket tyder på att heterologt uttryck av CBM är en lovande
metod för att manipulera cellväggen. I Publikation IV extraherades
mikrofibriller av cellulosa och nanokristallin cellulosa från de transgena
tobakscellerna. Framställningen av nanokristallin cellulosa från celler
som uttryckte CBM gav högre utbyte och kristallerna var längre.
Nanopapper tillverkat från mikrofibrillerna visade på ökad styrka och
seghet. Detta tolkades som att uttrycket av CBM hade lett till
förändringar i de organiserade regionerna av cellulosa.
Nyckelord: Glykosyltransferas, kallos-syntas, kolhydratbindande modul,
masspektrometri, cellulosa mikrofibriller, nanokristallin cellulosa,
plasmodesmata, Populus, primär cellvägg, radiometrisk analys,
spektrofotometrisk analys.
iii
List of Publications
Publication I:
Brown, C., Leijon, F., and Bulone, V. (2012). Radiometric and
spectrophotometric in vitro assays of glycosyltransferases involved in
plant cell wall carbohydrate biosynthesis. Nat. Protoc. 7, 1634-1650.
Publication II:
Leijon, F., Melzer, M., Zhou, Q., Srivastava, V., and Bulone, V. (2018).
Proteomic analysis of plasmodesmata from Populus cell suspension
cultures in relation with callose biosynthesis. Front. Plant Sci. 9, 1681.
Publication III:
Leijon, F.*, Melida, H.*, Melzer, M., Larsson, T., Srivastava, V., Gomez,
L., Guerriero, G., McQeen-Mason, S., and Bulone, V. The effect of
carbohydrate-binding modules (CBMs) on plant cell wall properties: an in
vivo approach. Manuscript.
Publication IV:
Butchosa, N., Leijon, F., Bulone, V., and Zhou, Q. (2018). Stronger
cellulose microfibrils network structure through the expression of
cellulose-binding modules in plant primary cell walls. Submitted to
Cellulose.
*Both authors contributed equally to the work.
iv
The Author´s Contribution
Publication I:
Felicia Leijon performed optimization experiments.
Publication II:
Felicia Leijon enriched membrane fractions, performed activity assays,
product characterization and prepared samples for mass-spectrometric
analysis. She also performed data analysis and contributed to the writing
of the manuscript.
Publication III:
Felicia Leijon performed plant transformation, extraction of cell wall and
cell wall carbohydrates, growth studies, xyloglucan assay, localisation
studies, PCR, qPCR and contributed to the writing of the manuscript.
Publication IV:
Felicia Leijon performed plant transformation and preparation of plant
cell cultures. She also contributed to the extraction of cellulose and
preparation and testing of the nanopapers.
v
List of Abbreviations
BcsA Bacterial cellulose synthase
CAZy Carbohydrate active enzymes
CalS Callose Synthase
CBM Carbohydrate-binding module
CesA Cellulose synthase
CipA Cellulosome-integrating protein A
CMF Cellulose microfibril
CNC Cellulose nanocrystal
CNF Cellulose nanofibril
DP Degree of polymerization
DRM Detergent-resistant microdomain
ER Endoplasmic reticulum
GC-MS Gas chromatography–mass spectrometry
GH Glycoside hydrolase
GRP Glycine-rich protein
Gsl Glucan synthase like
GT Glycosyltransferase
HPAEC High-performance anion-exchange chromatography
HRGP Hydroxyproline-rich glycoprotein
LPMO lytic polysaccharide monooxygenase
MS Mass spectrometry
NAD Nicotinamide adenine dinucleotide
NDP Nucleoside diphosphate
PRP Proline-rich protein
PM Plasma membrane
PD Plasmodesmata
ROP Regulation of cell polarity protein
SEL Size exclusion limit
SuSy Sucrose synthase
TC Terminal complex
TMD Transmembrane domain
UGT UDP-glucose transferring protein
WT Wild type
vi
Contents 1. Introduction ...................................................................................... 1
1.1. The plant cell wall ............................................................................ 2
1.1.1. Cell wall structure................................................................... 3
Middle lamella and pectins ................................................................ 3
Primary cell wall ................................................................................. 3
Secondary cell wall ............................................................................ 5
1.1.2. Cell wall growth ...................................................................... 6
1.1.3. The plasma membrane and the plasmodesmata and their
relationship with the cell wall .................................................. 7
Plasma membrane ............................................................................. 7
Plasmodesmata ................................................................................. 7
1.1.4. Model organisms used for studying the plant cell wall .......... 9
Plant cell suspension cultures ......................................................... 11
1.2. Cell wall polysaccharides - Structure and function ................... 12
1.2.1. Cellulose .............................................................................. 13
Nanocellulose .................................................................................. 15
1.2.2. Callose ................................................................................. 17
1.2.3. Xyloglucan ........................................................................... 18
1.3. Biosynthesis of cell wall polysaccharides .................................. 21
1.3.1. Classification and properties of glycosyltransferases .......... 21
1.3.2. Biochemical studies of GT activities in vitro ........................ 23
1.3.3. Cellulose synthase complex ................................................ 24
1.3.4. Callose synthase complex ................................................... 28
1.4. Carbohydrate-binding modules ................................................... 31
1.4.1. CBM/ligand interaction ......................................................... 31
1.4.2. CBM applications ................................................................. 33
1.4.3 CBM3 from CipA of Clostridium thermocellum .................... 34
vii
2. Present investigation .................................................................... 37
2.1. Aim of the present investigation ................................................. 37
2.2. Rationale, Results and Conclusion .............................................. 38
2.2.1. Publication I: In vitro glycosyltransferase assays ................ 38
Rationale ......................................................................................... 38
Results............................................................................................. 39
Conclusion ....................................................................................... 41
2.2.2. Publication II: The Populus plasmodesmata proteome ....... 41
Rationale ......................................................................................... 41
Results............................................................................................. 42
Conclusion ....................................................................................... 43
2.2.3. Publication III: The effect of carbohydrate-binding modules
(CBMs) on plant cell wall properties: an in vivo approach .. 44
Rationale ......................................................................................... 44
Results............................................................................................. 45
Conclusions ..................................................................................... 47
2.2.4. Publication IV: Stronger cellulose microfibrils network
structure through in vivo expression of CBMs ..................... 47
Rationale ......................................................................................... 47
Results............................................................................................. 48
Conclusions ..................................................................................... 50
2.3 Concluding remarks and future perspectives .............................. 51
3. Acknowledgments ......................................................................... 55
4. References ..................................................................................... 57
1
1. Introduction
This section aims to give a general background to the field of plant cell
walls, their carbohydrate components and the enzymes responsible for
their synthesis. To contextualize the individual publications further, short
reviews are also given on the structure and function of plasmodesmata,
cellulose nanofibrils and carbohydrate binding modules.
This thesis is comprised of four publications and is divided into two
distinct parts; the first part is more fundamental as it includes the
optimization of two methods for assaying glycosyltransferases and the
application of one of these assays in research regarding the cell wall
spanning structure plasmodesmata. The second is on the engineering of
cell wall properties by transgene expression and the subsequent
characterisation of the cell wall and cellulose properties thereof. The two
parts presented share a common thread in the investigation of the
properties of the primary cell wall of suspension cultured plant cells.
2
1.1. The plant cell wall
From an environmental point of view cell walls are an attractive raw
material as they are renewable, recyclable and their cultivation in the
form of forest represents a large carbon sink. Both the agricultural and
forest industries rely on high-throughput production to make relatively
low-cost bulk products. They also generate a great deal of waste, largely
cell wall based, which could be redirected into production of high-value
applications. Development of such new cell wall based materials and
products have been a major focus of the scientific community and
research and development sectors for the last decade. This has brought a
growing need for deeper knowledge of all aspects of cell walls to be able to
use the raw material they provide in the most efficient way. Basic
research covering cell wall structure, function and regulation is important
in order to provide the strongest foundations for future applications.
As plants are stationary beings, they have a need to withstand and adapt
to changing and adverse conditions. The plant cell wall helps them do this
through providing a structure that is recalcitrant yet still has a highly
adaptable organisation. The cell wall surrounds the plant cell; it is a
physical first line of defence, structural load bearer, mediator of cell-cell
adhesion and definer of shape. As it has many roles to fill it is not
surprising that there is great variation in the composition and shape of
the cell wall. Depending on the cell type, developmental stage and
species, the cell wall is adapted to meet the plants requirements.
Furthermore, walls within one cell can vary locally in structure,
showcasing how precisely regulated the architecture of the cell wall is.
The plant cell wall is mainly made up of polysaccharides (and lignin in the
case of secondary cell walls), unlike bacterial cell walls where
peptidoglycans are the main constituents, fungi that have a large amount
of glucans in addition to some chitin and animal cells that mostly lack cell
walls. The current plant cell wall model consists of a cellulose network
crosslinked by hemicelluloses within a pectic matrix (Figure 1A) (Carpita
and Gibeaut, 1993; Cosgrove, 2005). In certain specialized cells there is
also a secondary cell wall present. While descriptions of plant cell primary
and secondary walls often focus on structural and compositional
differences one should keep in mind that a distinction can also be made
on the developmental state of the cell, where the primary cell wall
3
surrounds the protoplasts of growing cells and therefore needs to be
dynamic and adaptable, while the secondary cell wall is more static as it is
found in cells where cell enlargement has ceased.
However, when discussing the finer details of composition and structure
it is important to keep in mind that the model is assembled from results
from many fields of research and that it is not necessarily an absolute
faithful representation of the in vivo situation. In view of the variation
found in cell wall structure it is also important to remember that the
model varies a great deal depending on what species, tissue and cell type
it represents.
1.1.1. Cell wall structure
Middle lamella and pectins
The middle lamella is a pectin-rich region that sticks adjoining cells
together (Figure 1A and 1B). The pectic polysaccharides
(homogalacturonan, rhamnogalacturonan I and rhamnogalacturonan II)
are hypothesised to form domains through covalent and non-covalent
linkages, leading to a highly hydrated matrix. Depending on the tissue
and its developmental stage, the type of polysaccharides and level of
crosslinking in the pectic matrix varies (Vincken et al., 2003). Ca2+
bridges are responsible for the establishment of a 3D network of
homogalacturonan, and boron bridges are formed between
rhamnogalacturonan II molecules (Morris et al., 1982; Ishii et al., 1999;
Vincken et al., 2003). This matrix is also present in the primary cell wall
where it encases the other cell wall polysaccharides. The network is
important both for cell wall mechanical properties and for the porosity of
the matrix.
Primary cell wall
The primary cell wall is formed by a load-bearing cellulose network cross
linked and coated with hemicelluloses (Figure 1A). The matrix is further
intermeshed with pectic polysaccharides that are hydrated, allowing for
the cell wall to comprise a large aqueous gel fraction; about 75% of the
matrix is water (Cosgrove, 1997). The aqueous phase allows for transport
of solutes within the wall and plays an important role in enabling
extensibility of the wall, allowing the cell to expand. The primary cell wall
4
is therefore flexible, but at the same time needs to be strong enough to
control the size and shape of the cell and withstand the intracellular
turgor pressure. It is generally thin (50-200 nm) (Albersheim et al.,
2011), in comparison to the secondary cell wall that can become several
micrometers thick (Doblin et al., 2010).
Figure 1. Model of the plant primary cell wall (A) (adapted with permission from Sticklen,
2008).Transverse section of cells forming secondary cell walls showing cell wall hierarchy
(B). Cellulose microfibril orientation in primary cell wall (P) is random while secondary cell
walls contain layers (S1, S2, S3) with differently oriented cellulose microfibrils (C).
5
A typical primary cell wall contains, in dry weight, about 40% pectins,
30% hemicelluloses, 25% cellulose and the remainder is protein (Taiz and
Zeiger, 2010). These numbers are however variable depending on species
and cell type. Typically, the primary cell walls of higher plants have been
divided into two different structure types: Type 1 is rich in hemicellulose
(essentially xyloglucans) and pectic polysaccharides and is present in
most angiosperms, while Type 2 is found in monocots and contains a
reduced amount of xyloglucan and pectins in favour of mixed-linkage
glucans and glucuronoarabinoxylans (Carpita and Gibeaut, 1993). The
cellulose content of primary cell walls is lower compared to secondary cell
walls and consists of both crystalline forms and substantial amounts of
less ordered cellulose (Sturcová et al., 2004; Thomas et al., 2013).
The primary cell wall further consists of a fraction of proteins. These can
be divided into proteins, glycoproteins and proteoglycans. The insoluble
protein group includes structural proteins such as hydroxyproline-rich
glycoproteins (HRGPs), glycine-rich proteins (GRPs) and proline-rich
proteins (PRPs) and the soluble proteins include enzymes, lectins and
transport proteins. As the cell wall forms part of the cell´s protection
from pathogens it also contains defence proteins and is an important
source of signalling molecules (reviewed in John et al., 1997; Lagaert et
al., 2009).
Secondary cell wall
In connection with the end of cell growth and differentiation, a second
cell wall can be deposited between the primary cell wall and the plasma
membrane (Figure 1B). This secondary cell wall is structurally and
functionally specialized, conveying the rigidity and strength to the cell
wall that is needed for upright growth and transportation of water within
the plant.
The secondary cell wall is divided into three layers (S1-3) defined by the
microfibril angle of the cellulose microfibrils within each layer (Figure 1B
and 1C), leading to a resilient structure. The secondary cell wall is
composed of approximately 40-80% cellulose, 10-40% hemicelluloses,
the remaining 5-25% being lignin, with a small amount of proteins
(Kumar et al., 2016). In angiosperms the hemicelluloses are for the most
part glucuronoxylans and glucomannans (except in monocots where
glucuronoarabinoxylans are dominant), and in gymnosperms they are
6
galactoglucomannans and glucuronoarabinoxylans (Scheller and Ulvsko,
2010). A substantial constituent of the secondary cell wall is the phenolic
polymer lignin. It has high molecular weight and a complex and
heterogeneous structure formed mainly of monomers of p-
hydroxycinnamyl alcohols (monolignols) (reviewed in Vanholme et al.,
2010).
1.1.2. Cell wall growth
In order for the cell to grow the primary cell wall must expand. The great
morphological variation seen in plant cells is achieved by utilizing
variations in structure and stress distribution within the cell wall and
localized enzymatic activity. The orientation of the microfibrils
determines to a certain degree in what direction cell expansion takes
place. Cells are inclined to grow perpendicular to the aligned fibrils,
leading to anisotropic growth (Probine and Preston, 1962; Kerstens et al.,
2001; Suslov and Verbelen, 2006). The ways in which a cell grows is
called either diffuse growth (reviewed in Braidwood et al., 2014) where
the cells expand equally in all directions, or tip growth (reviewed in
Rounds and Bezanilla, 2013), which is an expansion of the cell in one
direction only. Tip growth relies on weakening of the cell wall by
acidification and expansion through the formation of a Ca2+ gradient
together with the action of cytoskeletal components and vesicular
trafficking. Pollen tubes and root hairs are cell types representative of tip
growth. Diffuse growth or an intermediate form of the two growth types is
found in most other cell types.
Due to its high water content and viscoelastic properties, the primary cell
wall allows for expansion while being able to counterbalance the turgor
pressure from the cell interior. Plant cells expand by loosening the cell
wall structure, a process called stress relaxation, while simultaneously
allowing the turgor pressure in the cell to press the protoplast outwards
against the cell wall (Cosgrove, 2005). The cell wall matrix is then forced
to give way and expand outwards. As this is taking place, new cell wall
material is being synthesized, reinforcing the new wall structure and
preventing the wall from thinning through stretching. This expanding
movement of the cell wall components is believed to be due to cellulose
microfibrils and their associated polymers separating or sliding from each
7
other (Marga et al., 2005). The phenomenon is termed polymer creep.
Cell wall loosening is hypothesised to be achieved by modification of
polymer cross-linking. Extensibility of the cell wall may be altered by both
cell wall modifying enzymes, such as endo-transglycosylase/hydrolase
enzymes (Takeda et al., 2002; Park et al., 2003; Miedes et al., 2011; Park
and Cosgrove, 2012b) and by other cell wall-loosening agents such as
expansins. These are a group of small proteins that bind carbohydrates
(similar to carbohydrate binding modules, see section 1.4.) and loosen
noncovalent interactions between polysaccharides under low pH
conditions, leading to so-called acid growth (reviewed by Cosgrove, 2015).
1.1.3. The plasma membrane and the plasmodesmata and their
relationship with the cell wall
Plasma membrane
The plasma membrane (PM) consists of a phospholipid bilayer, which
also comprises glycolipids, sterols and proteins. It is a diffusional barrier
separating the outside of the cell from the cytosolic inner content. The
membrane upholds gradients important for many cell functions and
contributes vesicle transportation to and from the cell´s interior.
The protein portion of the membrane is found either embedded in the
bilayer or attached to the surface through post-translational
modifications. These proteins are involved in many important cellular
functions, for instance signal transduction, transport and stress response.
Enzymes involved in the synthesis of cell wall polysaccharides such as
cellulose and callose are also embedded in the PM (see section 1.3.3. and
1.3.4.).
Plasmodesmata
The plasmodesmata (PD) are membrane lined pores stretching across the
cell walls of adjacent cells, enabling exchanges between cells through the
cytoplasmic continuum (Figure 2) (Roberts and Oparka, 2003). The PMs
of the PD-connected cells are interlinked through the channels and coat
the entirety of the structure. A membranous central rod, the
desmotubule, crosses the PD and connects endoplasmic reticulum (ER) in
the adjacent cells (Overall et al., 1982). Between the PM and the
desmotubule spoke-like and globular structures, possible cytoskeletal
8
Figure 2. Overview of plasmodesma localization in the cell. The insert illustrates the
plasmodesmata structure and callose turnover at plasmodesmata through the synergistic
action of enzymes belonging to the callose synthases and β-1,3-glucanases (after Maule et
al., 2012).
proteins or cross-linking proteins have been observed (Robards, 1968;
Ding et al., 1992; Overall and Blackman, 1996; Nicolas et al., 2017). These
are hypothesised to be part of the mechanism that regulates the size
exclusion limit (SEL) of the structure and may aid in the opening and
closing of the pore or control separation of the membrane components
(Roberts and Oparka, 2003; Nicolas et al., 2017). Observations by
electron microscopy estimate the PD diameter to be of 20-50 nm (Ehlers
and Kollman, 2001) and in most growing tissues the SEL of the structure
has been found to be between 30-50 kDa (Crawford et al., 2000; Kim et
al., 2005). The PD structure, simple or branched, varies between cell
types and differentiation stages (reviewed in Ehlers and Kollman, 2001).
Recent results by Nicolas et al. (2017) further indicate that the
ultrastructure of the PD, for example pore diameter and protein
structures in the pore, are also dependent on the differentiation stage of
the cell and its age.
In the plant, PD play an important role for cell-to-cell trafficking of
solutes through the cytoplasmic sleeve that spans the pore. Proteins,
small RNAs, hormones and metabolites can all pass through the
cytoplasmic sleeve (De Storme and Geelen, 2014). Transport possibly also
takes place through lateral movement in the desmotubule lumen and
membrane (Grabski et al., 1993; Cantrill et al., 1999; Guenoune-Gelbart
et al., 2008; Barton et al., 2011). Since PD are the only cell-cell gateways
in the otherwise recalcitrant cell wall, they are prime points for pathogen
movement within the host plant. Viruses use proteins involved in PD
9
regulation to move between cells (reviewed by Benitez-Alfonso et al.,
2010).
The polysaccharide callose (β-1,3-glucan, see section 1.2.2.) has been
shown to be involved in the regulation of non-selective flow through the
PD. It is deposited and hydrolyzed from the neck region of the structure
in a dynamic manner, leading to the physical opening and closing of PD
and a change in conductivity (Figure 2) (reviewed in Zavaliev et al., 2011;
de Storme and Geelen, 2014). Members of the callose synthase enzyme
family (also called GSLs, see section 1.3.4.), have been found to be
involved in callose deposition at the PD (Guseman et al., 2010; Vaten et
al., 2011; Xie et al., 2011; Cui et al., 2016). Callose removal by hydrolysis
and subsequent opening of the pore is accomplished by β-1,3-glucanases
(Levy et al., 2007; Benitez-Alfonso et al., 2013; Zavaliev et al., 2013).
Callose deposition at plasmodesmata is predominantly linked to response
to hormonal signalling or biotic and abiotic stress (Schultz et al., 1995;
Radford et al., 1998; Citovsky et al., 1998; Sivaguru et al., 2000; Bilska
and Sowinski, 2010; Cui et al., 2016). Changes to PD conductivity through
callose deposition have been shown to be affected by auxin levels (Han et
al., 2014), regulated by salicylic acid accumulation (Lee et al., 2011; Wang
et al., 2013; Cui et al., 2016) and by accumulation of reactive oxygen
species (Benite-Alfonso et al., 2009 a, b; Oide et al., 2013). Regulation of
PD closure may also in part be Ca2+ dependent as PD closure is induced
by micromolar changes in intercellular calcium concentration (Tucker
and Boss, 1996; Holdaway-Clarke et al., 2000; Sager and Lee, 2014).
Therefore it is likely that the control of PD aperture is modulated by a
complex regulatory system.
1.1.4. Model organisms used for studying the plant cell wall
The complexity of the cell wall means that studying it is difficult and that
it is essential to have general theoretical models of the system in order to
simplify discussion, but also that there is a need to use model organisms
with representative cell walls. A range of factors have determined which
organisms have been selected depending on what exactly is of interest to
study.
10
In the following section a short description of each of the three model
organisms used in this thesis is given.
Populus trichocarpa (black cottonwood) and Populus tremula x
tremuloides (hybrid aspen) have emerged as model organisms for
hardwood trees. P. trichocarpa was the first tree with a sequenced
genome, due to the fact that its genome is relatively small (~500 Mbp,
Tuskan et al., 2006) in comparison to the softwood model organism pine
whose genome is 50 times larger (Wullschleger et al., 2002). Laboratory
work using Populus is facilitated by its fast growth and the availability of
a range of tools for its manipulation and genetic transformation (Meilan
and Ma, 2006). To some extent, Populus is related to Arabidopsis
(Jansson and Douglas, 2007) and their cell walls share many
characteristics (Chaffey et al., 2002; Bhalerao et al., 2003), making much
of the knowledge gained from the study of Arabidopsis valid for Populus
as well. However, to study the processes of wood development (xylem
formation, maturation, tension and compression wood) and responses to
seasonal variation such as cambial dormancy/activity and early- and
latewood formation a tree model, such as Populus, is needed.
Arabidopsis thaliana has been the most widely used plant model
organism and has therefore come to be used for studies of cell wall
biosynthesis, even though it largely lacks the woody tissue normally
associated with secondary cell wall formation. The reason for its extensive
use is a combination of traits that make Arabidopsis well suited as a
model organism: it can easily be cultivated under laboratory conditions,
has a short generation time, a relatively small genome and was the first
plant with a sequenced genome (The Arabidopsis Genome Initiative,
2000; Koornneef and Meinke, 2010). This has facilitated the
development of many molecular biology tools used for its study.
Nicotiana tabacum, and its close relative Nicotiana benthamiana are two
varieties of the tobacco family commonly used in studies of plant
physiology, fundamental biological processes and molecular biology
applications (Nagata et al., 1992). Aside from being an important crop,
supporting a profitable industry, tobacco is interesting as it provides a
good model for studies of plant-pathogen interactions (Goodin et al.,
2008) and is transformable (Mayo et al., 2006; Sparkes et al., 2006).
11
Plant cell suspension cultures
The method of cultivating plant cells suspended in defined liquid growth
media allows for the study of a system where the cell population is less
complex than in a whole plant where cell differentiation, tissue
differences and recalcitrant tissues complicate certain studies (Figure 3).
Suspension cultured cells also have the advantage of growing fast
compared to whole plants, and therefore they can readily and regularly
provide relatively large amounts of starting material for further studies.
Growth conditions can be tightly controlled and nutrients and hormones
are rapidly and evenly distributed to all cells in the culture (Mustafa et al.,
2011). A wide variety of plant tissues can be propagated as cell
suspensions. For many types of suspension cultures, there are established
protocols for transformation allowing genetic manipulation of the cells.
Furthermore, because most plant cells are totipotent it is often possible to
regenerate plants from cultured cells.
Figure 3. Populus trichocarpa cell suspension cultures.
12
1.2. Cell wall polysaccharides - Structure and function
The plant cell builds all the structurally and functionally diverse
polysaccharides it needs from a rather limited number of
monosaccharides. In aqueous solutions, the hydroxyl group of the C-4 or
C-5 of the sugar monomers reacts with the aldehyde or ketone group
leading to ring formation. Further, the configurations of the
monosaccharides are described as L or D according to the Fischer
nomenclature.
In oligo- and polysaccharides, the monosaccharides are further linked to
one another through glycosidic linkages. These are named by the carbons
involved in the linkage (e.g, 1→3, 1→4, etc) and divided into α- or β-
configuration depending on the stereochemistry of the anomeric carbon.
An oligosaccharide is a shorter chain of sugars (typically <10 residues)
while a polysaccharide consists of longer chains. Most sugar chains have a
reducing and non-reducing end giving the chain a polarity. The non-
reducing end has its terminal sugar fixed in a ring form through the
formation of the glycosidic bond to the subsequent sugar in the chain
(Figure 4A). At the reducing end the sugar can occur in the ring form or
liner form as the aldehyde or ketone group of the terminal sugar is not
permanently fixed into a ring. This is important in relation to
polysaccharide synthesis and degradation, and also analysis, as the
different ends of the chains have different chemical reactivity.
The type and amount of polysaccharides present in the different parts of
the cell wall vary between species and also between cell types. The
possibility to vary polysaccharide length, degree of polymerization (DP),
and position and configuration of bonds formed allows the cell to adapt
the carbohydrate portion of the cell wall. In addition to the
polysaccharides discussed in section 1.1, there are a number of less
common saccharides which have specialized functions within the cell.
Polysaccharides are hypothesized to form cell walls by self-assembly,
whereby organized structures are spontaneously formed by self-
aggregation, and by enzyme-mediated assembly, where catalytic proteins
assist in linking the cell wall components together (Cosgrove, 1997).
In the following sections three polysaccharides of importance for this
thesis are presented in more detail (cellulose, callose and xyloglucan).
13
1.2.1. Cellulose
Cellulose is the most abundant natural polymer found on earth and is
synthesised by plants, algae, bacteria, oomycetes and some animals. It
has a long history of human use and remains an attractive resource as it is
accessible and renewable. The major source of cellulose used in industry
is plants, foremost trees, where it is present in the primary and secondary
cell walls as microfibrils (nm/µm scale). In the cell it interconnects with
other components of the wall, adding strength and structural rigidity to
the matrix. The excellent load-bearing ability of cellulose is one of the
reasons trees can grow so tall. Cellulose is relatively resistant to both
chemical extraction and processing, and to enzymatic degradation as it is
chemically stable and insoluble.
The chemical composition and structure of cellulose is simple in
comparison to many other polysaccharides. It is an unbranched β-1,4-
linked polymer made up by glucopyranosyl residues in the 4C1 chair
formation (Figure 4A). The disaccharide cellobiose is often viewed as the
repeating unit of cellulose since each glucose residue in the chain is
oriented 180° relative to the previous residue.
The linear structure of cellulose and its many hydroxyl groups allow for
packing of the individual chains into sheets that are then stacked to form
crystals and microfibrils. Sheets are held together through inter and intra
hydrogen bonding between the free hydroxyl groups on the cellulose
molecules (Nishiyama et al., 2002, 2003). Hydrogen bonding and van der
Waals interactions between the individual sheets then lead to a stacking
of sheets that finally form microfibrils (Jarvis, 2003; Wada et al., 2004).
Cellulose can occur in both recalcitrant crystal forms and as more
accessible non-crystalline cellulose. Models of microfibril organisation
suggest that cellulose is for the most part found as a mixture of the two
forms, with stretches of crystalline cellulose interrupted by less organised
regions (Figure 4B) and that the cross section of the microfibril may also
have a varying degree of crystallinity with a central crystal core (reviewed
in Nishiyama, 2009).
Depending on the unit cell parameter of the crystals, cellulose can be
divided into four different forms called allomorphs (I-IV). Cellulose I
(native cellulose) is the form commonly found in nature. It has its
cellulose chains oriented parallel to each other (Hieta et al., 1984; Chanzy
14
Figure 4. Cellulose hierarchical structure. The structure of cellulose with the repeating unit
cellobiose in brackets (A). Ultrastructure of cellulose microfibrils with alternating ordered
and disordered regions (B) (after Moon et al., 2011).
and Henrissat, 1985; Koyama et al., 1997). It is further subdivided into
allomorphs Iα and Iβ depending on how the glucan chains are organized
within the crystals (Atalla and Vanderhart, 1984). Bacterial and algal
cellulose contain a higher amount of allomorph Iα (Sugiyama et al., 1991),
while cellulose Iβ is more abundant in higher plants (Atalla and
Vanderhart, 1984). The reason for this is unknown. Cellulose II has its
chains oriented in an anti-parallel manner, giving a more energetically
favourable structure (Kolpak and Blackwell, 1976; Stipanovic and Sarko,
1976; Langan et al., 2001). With a few exceptions (Sisson, 1938; Kuga et
al., 1993; Saxena et al., 1994; Shibazaki et al., 1998) this form of cellulose
is not found in living organisms but typically produced from chemical
treatments of cellulose I. Cellulose IV is a disorganized form of cellulose I
(Wada et al., 2004; Newman, 2008), also naturally occurring in some
plant primary cell walls and micro-organisms (Chanzy et al., 1979; Bulone
et al., 1992; Helbert et al., 1997), but to a much lesser extent than
cellulose I. Cellulose IV may also be formed by converting cellulose I or II
into cellulose III by chemical treatment and then further into cellulose IV
by thermal treatment.
15
The DP is the number of glucose monomers in a single cellulose chain. It
is an important factor as the properties of cellulose partly change with
length and length distribution within a sample. The DP varies a great deal
depending on the source of the material and what extraction method has
been used to recover the fibres, as different extraction procedures affect
depolymerisation differently. In the primary cell wall of growing cells
cellulose is found mainly in two fractions ranging in DP from 250-500 to
up to 2500-4000 (Blaschek et al., 1982; Franz and Blaschek, 1990). In the
secondary cell wall, the DP is considered to be between 10000-15000
(Albersheim et al., 2011). In wood pulp, a common industrial raw
material, the DP is in the range 300-1700 (Klemm et al., 2005).
Cellulose is synthesized by enzymatic complexes embedded in the PM.
Newly synthesized microfibrils are deposited in the cell wall as they are
formed and intermesh with other cell wall constituents. Cellulose
biosynthesis is further detailed in section 1.3.3.
Nanocellulose
Cellulose fibres can be disintegrated into nanoscale fragments, which are
grouped into cellulose nanofibrils (CNFs) and cellulose nanocrystals
(CNCs) (Figure 5). When extracted from wood CNFs are approximately
0.5-10 µm long and 4-100 nm wide and CNC are approximately 50-500
nm long and 3-5 nm wide (Moon et al., 2011). CNFs are most commonly
produced by defibrillation, the mechanical or chemomechanical
treatment of cellulose fibres or aggregates of nanofibers, resulting in
cellulose nanofibrils (Turbak et al., 1983; Paakko et al., 2007; Saito et al.,
2006). These elements contain both crystalline and non-organized
regions of cellulose.
Mechanical isolation of CNFs results in the presence of hydroxyl groups
on the nanofibril surface. Through chemical pretreatments, carboxyl or
carboxymethyl groups can also be introduced on the surface of CNFs
before their isolation (Saito et al., 2006; Wågberg et al., 2008). CNCs are
prepared through acid hydrolysis of fibres, resulting in the removal of the
non-crystalline sections, while leaving the crystalline cellulose portions
intact (Figure 5) (Ranby, 1949, 1951; Marchessault et al., 1961). The acid
hydrolysis used to produce CNCs may, in some conditions, functionalise
their surfaces, most commonly with sulphate esters (Marchessault et al.,
1961). The surface functionalities on both CNFs and CNCs can then be
16
Figure 5. Cellulose nanoparticle isolation. Cellulose microfibril orientation in primary cell
walls (P) is random while secondary cell walls contain layers (S1, S2, S3) with differently
oriented microfibrils. Cellulose microfibrils can be isolated from the cell wall and further
processed into CNFs and CNCs (after Postek et al., 2011).
further exploited to covalently or non-covalently graft molecules or
functional groups to the surface in order to alter fibril characteristics and
tailor properties (Araki et al., 2001; Ljungberg et al., 2005; Zhou et al.,
2005, 2009; Ifuku et al., 2007; Littunen et al., 2011; Wang et al., 2011;
Fujisawa et al., 2013; Zhang et al., 2014).
If cellulose microfibrils (CMFs) (with one dimension in the nanoscale) are
instead produced by industrial biosynthesis, modifications can take place
as the microfibril is formed or shortly after. Modifications are made by
introducing chemicals or proteins to the growth medium of the cellulose
producing organism (Brown et al., 1982; Haigler et al., 1982; Shpigel et al.
1998a; Yamanaka et al., 2000). An alternative method is through the
addition or alteration of genes which can lead to the production of novel
proteins that further interact with or affect the structure of the CMF
(Kawano et al, 2002; Shoseyov et al., 2003; publication IV of this thesis).
Alterations to CMF structure could possibly also be imposed by altering
the biosynthetic machinery responsible for the production of the cellulose
fibrils.
Due to the differences in cellulose biosynthesis and cellulose synthase
organisation in different organisms, the characteristics and properties of
the final cellulose structure vary (Brown et al., 1996). Therefore
17
extraction methods and final applications of cellulose nanoparticles to a
certain degree differ depending on the source of material (Moon et al.,
2011). Applications for cellulose nanomaterials can currently be divided
into two main types: materials made from cellulose CNFs and/or CNCs,
such as nanopapers, hydrogels, aerogels and foams, or composite
materials where cellulose nanoparticles are used as reinforcement fillers
(reviewed in Moon et al, 2011). Cellulose from plant secondary cell walls
is most commonly used for applications, while primary cell wall cellulose
is so far essentially of academic interest.
1.2.2. Callose
Callose is a linear form of β-1,3-glucans derived from plants (Figure 6). In
fully mature cell walls callose is a minor contributor to the overall
carbohydrate content. Pollen tube walls are the only identified higher
plant tissues where callose is the main structural polysaccharide (Stone
and Clarke 1992; Li et al., 1999) and may have a load-bearing role (Parre
and Geitman, 2005). However, during the life cycle of a plant, callose is
an important component which is deposited under specific natural events
and subsequently degraded by β-1,3-glucanases, making its presence
mostly transient. For example, it is found in the cell plate during
cytokinesis (Hong et al., 2001b; Thiele et al., 2009), in guard cells
(Apostolakos et al., 2009) and in reproductive tissues such as the cell wall
during pollen tube growth and in the pollen mother cell (Rae et al., 1985;
Schlupmann et al., 1994; Owen and Marakoff, 1995; Ferguson et al.,
1998). It is also deposited in the cell wall as a response to stress related
scenarios, such as wounding and pathogen attack (Kauss, 1985; Jacobs et
al., 2003; Nishimura et al., 2003), and is important in the PD (see section
1.1.3.). The precise function of callose is, in many of these cases, not well
understood.
β-1,3-Glucans in plants are predominantly linear, although a small
amount of β-1,6-branches has been reported in some instances (Huwyler
et al., 1978; Maltby et al., 1979; Hayashi et al., 1987; Hoffman and Timell
1970, 1972; Rae et al, 1985). There are numerous other β-1,3-glucans
from natural sources, all displaying varying branching and length.
Curdlan, a high molecular weight bacterial equivalent to callose is
commonly used for the study of linear β-1,3-glucans. Like cellulose
18
Figure 6. Structure of callose, a linear β-1,3-glucan derived from embryophytes.
β-1,3-glucans are homopolymers of glucose units (Figure 6). However, the
monomers are β-1,3-linked which leads to a twisted chain and thereby the
formation of triple helices when the polymer aggregates (Pelosi et al.,
2003; Stone and Clark, 1992). Hydrogen bonding takes place both within
helices in the individual callose chains and in helix-helix interactions
(Bluhm and Sarko, 1977; Marchessault et al., 1977; Deslandes et al., 1980;
Chuah et al., 1983; Young et al., 2000 Pelosi et al., 2006). β-1,3-Glucans
are found in nature as soluble, gel forming or insoluble crystalline
structures. The properties of the β-1,3-glucans in solution and gel form
depend on the size and conformation of the polymer, which in turn
depend on the biological origin and extraction method used.
There is currently no detailed information on the in vivo molecular
structure of callose from plants or the nature of its interactions with other
cell wall carbohydrates due to the effects that extraction methods can
have on the structure of the polymer. Callose is synthesised by PM-
located enzymes termed callose synthases. Further details on callose
biosynthesis are presented in section 1.3.4.
1.2.3. Xyloglucan
Xyloglucan has a β-1,4-linked glucan backbone, which is decorated to
varying degrees with short oligosaccharide branches, composed of xylose,
further substituted by galactose, fucose and/or arabinofuranose (Figure
7). The interval between substitutions on the backbone and the type of
branching is often repetitive, leading to a backbone motif of four glucose
residues with substitutions on the first two or three of these (Vincken et
al., 1997). The composition of the side chains and complexity of the
19
decoration patterns varies depending on species and tissue (Hoffman et
al., 2005; Peña et al., 2008; Hsieh and Harris, 2009). The branched
nature of the polysaccharide leads to a less ordered structure compared to
cellulose. Xyloglucan therefore does not form crystalline microfibrils.
Xyloglucan binds to the surface of cellulose microfibrils and has dual
opposing roles, either coating the fibres thereby preventing interactions
between them or, to a lesser estemate, binding and crosslinking the
cellulose (Hayashi, 1989; Nishitani, 1998, Bootten et al., 2004; reviewed
in Park and Cosgrove, 2015; Zheng et al., 2018). This contributes to the
distribution of the microfibrils in the cell wall matrix and can lessen fibril
aggregation (Anderson 2010). Mutational studies in Arabidopsis that
generate plants with no xyloglucan, lead to remarkably minor growth
defects, but altered mechanical properties and changes to cellulose
biosynthesis (Cavalier et al., 2008; Park and Cosgrove, 2012a; Xiao et al.,
2016). Xyloglucan can also form covalent links with other polysaccharides
such as pectic polysaccharides (Popper and Fry, 2008).
Figure 7. Xylogluco-oligosaccharide with common side chains. Side chains are, from left to
right: 1. an α‐1,6 xylosyl substitution; 2. the xylosyl can be further substituted by a β‐1,2
linked galactosyl residue, which in turn can be substituted by an α‐1,2‐L‐fucosyl residue (3).
20
Biosynthesis of the xyloglucan backbone has been shown to be catalyzed
by a membrane-bound enzyme that is a member of the cellulose synthase
like gene family C (Cocuron et al., 2007). The glucan chain is then
decorated by glycosyltransferases (see section 1.3.) that synthesise the
side chains (Perrin et al., 1999; Faik et al., 2002; Vanzin et al., 2002;
Madson et al., 2003; Li et al., 2004; Peña et al., 2004; Cavalier et al.,
2008; Zabotina et al., 2008, 2012). Synthesis takes place in the Golgi
apparatus, as is believed to be the case for all hemicelluloses and pectins
(Scheller and Ulvskov, 2010). The polysaccharides are then transported
in vesicles to the apoplast and incorporated in the cell wall.
Xyloglucan has been found in all land plants studied (Popper and Fry,
2003, 2004; Peña et al., 2008). It is the principal hemicellulose found in
the primary cell wall of dicots, constituting approximately 20-25% (w/w)
of the cell wall polysaccharides. The primary cell wall in conifers contains
10% xyloglucan whereas Poaceae (grasses) primary walls comprise 2-5%
xyloglucan (Scheller and Ulvskov, 2010). In some plants, xyloglucan is
also a storage polysaccharide in seeds (Gidley et al., 1991; Buckeridge et
al, 2000; Buckeridge, 2010).
21
1.3. Biosynthesis of cell wall polysaccharides
The enzymes that contribute to the formation and regulation of the
dynamic cell wall structure are classified as glycosyltransferases (GTs),
glycoside hydrolases (GHs), transglycosylases, glycan phosphorylases and
polysaccharide lyases. Together these enzymes interplay to allow for the
formation of the carbohydrate portion of the plant cell wall and its
remodelling to meet the specific requirements of plant developmental
needs and growth stages. GHs are involved in carbohydrate turnover,
both in the cell wall and in processing of storage carbohydrates. This is
achieved either by the complete degradation of carbohydrates or through
the modification of length and decoration patterns. As their name
indicates, GHs catalyse the breakage of glycosidic bonds in carbohydrates
through hydrolysis. This class of enzymes is generally better characterized
than GTs.
Plant cell wall polysaccharides are synthesized by GTs. Cellulose and
callose are synthesized in the PM by large complexes and extruded into
the cell wall while hemicelluloses and pectins are synthesized in the Golgi
apparatus, either as fully formed cell wall carbohydrates or as precursors
that are then exported to the apoplast by exocytosis. Both cellulose and
callose synthase complexes are discussed in detail later in this chapter.
1.3.1. Classification and properties of glycosyltransferases
GTs either have an inverting or retaining mode of action, meaning that
the configuration of the bond formed is either inverted or retained
relative to the bond of the leaving group in the sugar donor (Figure 8)
(Sinnott et al., 1990). Furthermore, the mode of catalysis is either
processive or non processive depending on whether several sugars are
added sequentially or the product is released immediately after the
monosaccharide transfer reaction has taken place (Saxena et al., 1995).
Glycosidic bond formation is catalysed using activated sugar donors, such
as nucleoside diphosphate sugars (Lairson et al., 2008). Bond formation
between sugars is most common, leading to polysaccharide products, but
acceptor substrates can also be proteins, lipids, nucleic acids or small
molecules and therefore GTs can also synthesize the sugar components
of, e.g, glycoproteins and glycolipids (Wagner and Pesnot, 2010).
22
While GHs present diverse types of structures, the GTs have a more
reduced repertoire of folds, possibly due to common ancestry (Lairson et
al., 2008). GTs that use nucleotide sugars as donors have either a GT-A
fold, with two tightly associated domains, one nucleotide binding and one
acceptor binding forming a twisted β-sheet bordered by α-helices (Bourne
and Henrissat, 2001; Charnock and Davies, 1999; Unligil and Rini,
2000), or a GT-B fold, with two similar but more flexibly linked
Rossmann-like domains with the active site in the cleft between them (Hu
and Walker, 2002; Vrielink et al., 1994). The GT-A fold was recently
confirmed for the GT-domain of a bacterial cellulose synthase (BcsA)
(Morgan et al., 2013) and modelling suggests that plant cellulose
synthases have a similar fold (Sethaphong et al., 2013).
GTs are listed and classified in the CAZy (Carbohydrate Active enZymes)
database together with other carbohydrate active enzymes and domains
(Lombard et al., 2014). Currently there are over 477000 GTs sorted into
106 families based on their amino acid sequence similarity. Both catalytic
machinery and mode of action tend to be conserved within the families.
Figure 8. The inverting and retaining mechanisms of glycosyltransferases give rise to two
stereochemically different products. (Reprinted with permission from Tvaroška, 2015).
23
1.3.2. Biochemical studies of GT activities in vitro
The degree to which the GTs have been biochemically characterized
varies greatly between different families. Biochemical characterization of
GTs has been hampered by the fact that many are large membrane-bound
enzymes, often with a high number of transmembrane domains and
typically part of unstable multiprotein complexes. Other possible
complications include the need for post-translational modifications,
correct folding, and risk of denaturation. Biochemical characterization of
heterologously expressed GTs is further complicated by the fact that the
many common expression systems utilize hosts that have their own GTs,
thus creating background activity.
Biochemical assays of plant glucan synthases have mainly been done
either by spectrophotometric assays or by quantifying the incorporation
of radioactively labelled substrates as, for example, reported in
publication I of this thesis. The product can then be characterized by
specific enzymatic degradation, chemical linkage analysis and/or sugar
analysis using gas chromatography–mass spectrometry (GC-MS). Assays
must take into account that the product formed may be water soluble or
insoluble, depending on the length and branching of the carbohydrate
formed. Finding the right conditions for assaying enzyme activity is also
crucial as multiple factors such as co-factors, ions, acceptors and
activation by proteolytic cleavage can affect activity. Members of the GT-
A type enzymes are particularly dependent on the coordinated binding of
a divalent metal ion such as Mg2+, Mn2+ or Ca2+ and in many the
coordination of the ion is established by the aspartates of the concerved
DxD-motif (Wiggins and Munro, 1998; Breton et al., 1998). The choice of
detergent for solubilization of membrane-bound enzymes is also a crucial
step as GTs typically are unstable upon extraction from the membrane.
Some detergents also act as activator and enhance enzyme activity (Li et
al., 1997; Lai Kee Him et al., 2001).
The recently characterized recombinant cellulose synthase from Populus
has a requirement for reconstitution into a lipid bilayer in order to retain
activity (Purushotham et al., 2016). For the cellulose and callose
synthases, specialized lipid environments in the membrane may be of
importance for activity as Detergent-Resistant Microdomains (DRMs)
fractions have been found to be enriched in glycan synthase activities
(Bessueille et al., 2009; Briolay et al., 2009; Srivastava et al., 2013).
24
DRMs can be prepared from the PM through the isolation of the lipid
fraction (and their associated proteins) resistant to solubilisation by non-
ionic detergents (Rietveld and Simons 1998; London and Brown, 2000).
Due to the enrichment of sphingolipids, sterols and phospholipids with
saturated fatty acids in the preparation from plants a distinct membrane
environment is created (Mongrand et al., 2004; Borner et al., 2005; Laloi
et al., 2007; Bessueille et al., 2009), which is believed to harbour
specialized protein populations linked to cellular processes such as
carbohydrate synthesis, signal transduction, apoptosis and endocytosis
(reviewed in Mongrand et al., 2004). It is debatable if DRMs correspond
to an actual transient in vivo membrane microdomain structure within
the PM (Lichtenberg et al., 2005; Tanner et al., 2011), but they
nevertheless offer the opportunity to study proteins that segregate within
a specific lipid environment.
1.3.3. Cellulose synthase complex
The enzymes that catalyse the formation of cellulose from UDP-glucose
are named cellulose synthases (CesAs). Higher plants usually have large
CesA gene families, with Arabidopsis having 10 CesAs (Richmond and
Somerville, 2000) and Populus trichocarpa 18 (Djerbi et al., 2005). The
CesAs belong to GT family 2 (GT2) according to the CAZy classification.
They are processive enzymes using an inverting mechanism to catalyze
the formation of a cellulose chain. The product is polymerized from the
non-reducing end (Koyama et al., 1997; Lai Kee Him et al., 2002). CesAs
are located in the PM with multiple transmembrane domains that span
the membrane, allowing for the final product to be extruded into the
extracellular space as it forms (Figure 9A). The newly formed cellulose
chains probably spontaneously aggregate into microfibrils (Delmer, 1999;
Guerriero et al., 2010). Modelling of a cotton CesA on the previously
determined structure of the bacterial cellulose synthase shows that
elongation of the glucan chain is likely initiated on the cytoplasmic side of
the PM and that the chain is then translocated through a channel formed
by transmembrane domains (Morgan et al., 2013; Sethaphong et al.,
2013; Slabaugh et al., 2014).
The cellulose synthases form complexes located in the PM. These were
identified by freeze-fracture experiments in algae and named terminal
25
complexes (TCs) as they were observed at the tip of growing microfibrils
(Brown and Montezinos, 1976). TCs exhibit a variation of spatial
arrangements in the PM and give rise to fibrils with different dimensions
depending on the species they are found in (Okuda et al., 1994; Brown et
al., 1996).
In plants, TCs are organised in hexagonal supramolecular structures,
called “rosettes”, with six subunits each that, up until recently, were
believed to contain six putative CesAs each (Figure 9A) (Mueller and
Brown, 1980; Brown, 1996; Kimura et al., 1999). Assuming each CesA
produces one cellulose chain, the number of cellulose chains in a
microfibril can never be more than the number of CesAs in the TC
synthesizing it. It is therefore tempting to speculate that each plant TC
would make a microfibril of 36 glucan chains. Primary cell wall
microfibrils have however been found to consist of 18-24 chains (Bootten
et al., 2004; Newman et al., 2013; Thomas et al., 2013; Wang and Hong,
2016), raising the question of whether some CesA proteins in the complex
are inactive or that part of the protein complex is composed of accessory
proteins (reviewed Guerriero et al., 2010). In both the primary and
secondary cellulose synthase complexes, the ratio between the different
CesA isoforms has been shown to be 1:1:1, allowing for either three or up
to six CesA per subunit (Gonneau et al., 2014; Hill et al., 2014).
Confirming these findings, recent transmission electron microscopy
results combined with computational studies point toward 18 cellulose
synthases contributing to the rosette structure in plants (Figure 9A)
(Nixon et al., 2016). These are presumably arranged in a trimeric
organization, giving rise to a microfibril composed of 18 cellulose chains
per rosette (Nixon et al., 2016).
In Arabidopsis, which is the organism that has been most commonly used
for studying the cellulose synthase complexes of plants, cellulose in the
primary and secondary cell walls is likely made by different sets of CesAs.
Mutational studies (reviewed by Endler and Persson, 2011) have shown
that AtCesA1, AtCesA3 and AtCesA6 all contribute to the formation of
cellulose in the primary cell wall matrix (Caño Delgado and Penfield,
2003; Ellis and Turner, 2001; Desprez et al., 2007,) and most likely form
homo- and heterodimers with each other (Desprez et al., 2007). While
AtCesA1 is necessary for cellulose synthesis (Beeckman and Przemeck,
2002), AtCesA3 and AtCesA6 knockout mutants are still able to produce
26
Figure 9. Hypothetical models for the cellulose synthase complex (A) (after Guerriero et al.,
2010; Sethaphong et al., 2013; Nixon et al., 2016) and the callose synthase complex (B)
(after Verma and Hong, 2001). Many aspects of how the complexes work and what
accessory proteins they interact with are unknown. Proteins believed to be associated with
the complexes are represented as ovals. KOR, KORRIGAN endoglucanase; CSI1, Cellulose
Synthase-Interactive protein 1; COB, COBRA; KOB, KOBITO; TED6, Tracheary Element
Differentiation-related genes 6; SuSy, Sucrose Synthase; ROP, Regulation Of cell Polarity
protein; ANN, Annexin; UGT, UDP-Glucose Transferase.
27
cellulose to some degree (Fagard et al., 2000). Some redundancy has
been shown to exist between AtCesA5, AtCesA2 and AtCesA6, allowing
the former two to replace the latter in forming functional complexes with
AtCesA1 and AtCesA3 (Persson et al., 2007).
In the secondary cell wall AtCesA4, AtCesA7 and AtCesA8 form a complex
where AtCesA4 is crucial for formation of the complex and mutation of
any individual gene leads to defective secondary cell walls (Taylor et al.,
2000, 2003). These three CesAs have also been found to make up an
intact cellulose synthase complex isolated through dual epitope tagging
(Atanassov et al., 2009).
It is tempting to speculate that the reason for the occurrence of multigene
families is that CesAs are active during different developmental stages or
in specific tissues. However, a certain amount of redundancy has been
found among several of the different isoforms of CesA (Persson et al.,
2007; Endler and Person 2011). Carroll et al. (2012) showed that all
primary and secondary cell wall CesAs are able to interact with each other
and that a primary CesA could partially compensate for the loss of a
secondary CesA and vice versa, proving that some mixed complexes are
active. The recently biochemically characterized recombinant CesA8 from
hybrid aspen shows that a single subunit is also able to form microfibrils
in vitro (Purushotham et al., 2016). Interestingly this CesA is Mn2+
dependent and has no or very little activity in the presence of Ca2+ or
Mg2+ (Purushotham et al., 2016), which is contradictory to CesA activity
from solubilized enzyme preparations from other sources that require
Ca2+ and Mg2+ (Colombani et al., 2004; Lai Kee Him et al., 2002;
Cifuentes et al., 2010; Okuda et al., 1993; Kudlick et al., 1996, 1997).
CesAs contain the conserved motif D,D,D,QxxRW that is common to
many processive GTs (Saxena et al., 1995). For example, both the closely
related cellulose-synthase-like gene products and the curdlan synthase of
Agrobacterium tumefaciens, also belonging to the GT2 family, share this
motif (Stasinopoulos et al., 1999; Richmond and Somerville, 2000). The
D,DxD motif is found in most processive and non-processive GTs, leading
to the hypothesis that they are of importance for the catalysis while the
D,QxxRW motif found towards the C-terminal part of the protein is
present predominantly in processive GTs and may therefore be involved
in processivity (Saxena et al., 1995). Mutations of the D,Q,R and W
28
residues of this motif, however, show that these amino acids are also
essential for enzyme activity in Gluconacetobacter xylinus (now known as
Komagataeibacter xylinus), as are the aspartates of the D,DxD motif
(Saxena and Brown, 1997; Saxena et al., 2001). The crystal structure of
BcsA of Rhodobacter sphaeroides revealed that the binding site for the
acceptor sugar is in part formed by the QxxRW motif (Morgan et al.,
2013). The same structure also showed that two of the D,DxD aspartic
acids were found to be involved in coordinating the UDP of the substrate,
while the third may correspond to the catalytic base (Morgan et al., 2013).
The N-terminal end of plant CesAs are involved in the formation of
homo- and heterodimers in vitro through a zinc finger domain with four
conserved CxxC motifs (Kurek et al., 2002).
The substrate of cellulose synthases, UDP-glucose, is synthesized in the
cytoplasm by sucrose synthase (SuSy) and UDP-glucose
pyrophosphorylase. Both these proteins have been suggested to be
accessory proteins to the cellulose synthase complex, supplying it with its
substrate (Figure 9A). Heterologous overexpression of a membrane-
bound SuSy from cotton led to an increased amount of cellulose
(Coleman et al., 2009). In addition to SuSy, several other plant proteins
have, through mutant analysis, been found to have an effect on cellulose
production. Among these are the β-1,4-glucanase Korrigan, KOBITO,
COBRA, TED6 and CSI1, among others (Nicol et al., 1998; Schindelman
et al., 2001; Sato et al., 2001; Pagant et al., 2002; Roudier et al., 2005;
Endo et al., 2009; Takahashi et al., 2009; Gu et al., 2010; Song et al.,
2010). However, precise functions of these proteins remain unclear.
1.3.4. Callose synthase complex
Callose synthases (CalS) are GTs that use an inverting mechanism to
catalyze the formation of linear β-1,3-glucan chains from UDP-glucose.
The gene family consists of 12 genes in Arabidopsis and in Populus
trichocarpa there are at least 13 genes currently annotated as callose
synthases (Hong et al., 2001b; publication II of this thesis). All gene
products belong to the GT48 family in the CAZy classification together
with β-1,3-glucan synthases from other organism (Coutinho et al., 2003;
Lombard et al., 2014). The annotations CalS or Glucan Synthase Like
(GSL) are both found in the literature and it is worth noting that the
29
numbering of the genes is not consistent between the two annotation
systems (see section 1.1.3.).
Callose and cellulose synthases share features such as being PM located,
having large cytosolic domains and using the same substrate to catalyze
the formation of linear polysaccharides (Figure 9B). However callose
synthases are almost twice as long as cellulose synthases, approximately
2000 amino acids versus 1000 amino acids, and have up to twice the
number of predicted transmembrane domains (TMDs) (10-16 vs 8)
(Doblin et al., 2001; Hong et al., 2001b; Ostergaard et al., 2002;
Somerville, 2006; Ellinger and Voigt, 2014; McNamara et al., 2015).
While studies of gene knock outs and down regulation of the CalS genes
show that they are required for callose deposition at various stages of cell
development, these experiments do not demonstrate that they are the
catalytic subunits (Enns et al., 2005; Dong et al., 2005; Nishikawa et al.,
2005). Callose synthases lack the D,DxD and D,QxxRW motifs, involved
in catalysis and processivity, which are characteristic of the members of
the cellulose synthase super family (GT2) of plants and the curdlan
synthase of Agrobacterium (Saxena et al., 1995; Stasinopoulos et al.,
1999; Verma and Hong, 2001; Karnezis et al., 2003). Although the
corresponding motifs have not yet been identified for callose synthase,
the protein may still be the catalytic subunit as the large cytosolic domain
of the CalS protein has a high similarity to FKS, the β-1,3-glucan synthase
of yeast (Cui et al., 2001; Doblin et al., 2001; Hong et al., 2001b;
Ostergaard et al., 2002). The lack of the motifs typically found to be
involved in the binding of the substrate suggests that UDP-glucose might
not be bound directly by the callose synthase (Verma and Hong, 2001).
Instead, substrate binding could possibly be carried out by a protein
associated to the callose synthase, such as the UDP-glucose transferase
(UGT) (Hong et al., 2001a; Verma and Hong, 2001).
As callose synthases are large integral PM proteins with a high number of
TMDs, it has instead been proposed that they may be a pore forming
structure used for the translocation of the carbohydrate chain across the
membrane, the associated catalytically active protein remaining
unidentified (Brownfield et al., 2009). Possibly, several callose synthases
would be involved in one complex in analogy with the cellulose synthase
rosette structures (Brownfield et al., 2009). Large complexes have been
30
found associated to newly synthesized β-1,3-glucan polysaccharide chains
(Bulone et al., 1995; Lai Kee Him et al., 2002) and both barley and
spinach GSLs have been identified as components of substantial
complexes (Li et al., 2003; Kjell et al., 2004). These complexes are
hypothesized to be formed by callose synthases possibly in association
with other proteins such as UGT, Regulation of cell polarity protein
(ROP), and SuSy (Figure 9B) (Amor et al., 1995; Hong et al., 2001a+b).
Annexin has also been found in callose synthase enriched preparations
and was proposed to be involved in linking the callose synthase to the
cytoskeleton (Andrawis et al., 1993).
Callose synthase activity has been confirmed in a number of cases from
detergent extracts from different plant tissues and cell suspension
cultures (Wu et al., 1991; Okuda et al., 1993; Bulone et al., 1995; Turner et
al., 1998; Lai Kee Him et al., 2001; Li et al., 2003; Colombani et al., 2004;
Bessueille et al., 2009; Cifuentes et al., 2010; Vaten et al., 2011). The type
of detergents that lead to the highest activity differs between callose
synthases and may reflect a requirement for specific membrane
compositions for the different forms of the enzyme. Callose synthase
activity has been shown to be either Ca2+ dependent or independent. The
nature of the interaction between Ca2+ and the enzyme complex is
unknown. The dependent form, which is far more studied than the
independent form, has been linked to the depolarization of the membrane
in connection to wounding (Kauss, 1985; Kohle et al., 1985; Fredrikson
and Larsson, 1992; Schlupmann et al., 1993). The Ca2+ independent
activity has so far only been found in pollen tube cells (Schlupmann et al.,
1993) and in plasmodesmata enriched extracts, as reported in publication
II of this thesis.
31
1.4. Carbohydrate-binding modules
Carbohydrate active enzymes such as glycoside hydrolases often have
carbohydrate binding non-catalytic protein domains as part of their
multimodular structure. These domains typically promote the interaction
between enzyme and substrate, thereby enhancing enzymatic
degradation, and are termed carbohydrate-binding modules (CBMs).
Information regarding identified and characterized CBMs is found in the
CAZy database (Lombard et al., 2014) where CBMs are grouped in 83
sequence based families often showing conserved ligand specificity and
protein fold within the families. The structures of the CBMs have also
given rise to a classification in fold families depending on their protein
fold (Boraston et al., 2004). The most common fold is that of the β-
sandwich made up by two β-sheets with a coordinated metal ion
(Boraston et al., 2004; Oliveira et al., 2015).
CBMs are believed to enhance the efficiency of their associated enzyme by
targeting the enzyme to a defined region within a complex carbohydrate
substrate, holding the enzyme in close proximity to the substrate for a
prolonged time and thereby increasing the local concentration of the
enzyme (Bolam et al., 1998; Hervé et al., 2010). Controversially, some
CBMs were also suggested to disrupt the packed surfaces of otherwise
inaccessible carbohydrates, allowing for enzymatic degradation (Din et
al., 1991; Southall et al., 1999; Gourlay et al., 2012). Members of the
CBM33 family with ligand disruptive abilities have been shown to be
redox-active enzymes and have since been reclassified as lytic
polysaccharide monooxygenases (LPMOs) (Vaaje-Kolstad et al., 2010;
Horn et al., 2012).
1.4.1. CBM/ligand interaction
Together CBMs cover a huge range of binding specificities, discriminating
not just the type of carbohydrates but also their length and organization.
CBMs have been grouped into three types (A, B and C) based on the
topology of their ligand interacting site and hence the ligands
characteristics and the function of the CBM (Figure 10) (Boraston et al.,
2004; Gilbert et al., 2013). The interaction between the CBM and their
ligands takes place through non-covalent bonds such as hydrogen bonds
32
and aromatic stacking interactions between the residues present in the
two surfaces (reviewed in Gilbert et al., 2013).
Type A CBMs have a flat hydrophobic binding site characterized by the
presence of several aromatic amino acids that interact with the crystalline
surfaces of cellulose or chitin (Linder et al., 1995; Xu et al., 1995; Tormo
et al., 1996). They have very poor, if any, affinity to soluble ligands
(Bolam et al., 1998).
Type B CBMs bind individual polysaccharides internally, as endo-type
carbohydrate active enzymes, and generally have a preference for chains
of four residues or longer (Boraston et al., 2004). This type of CBMs is so
far the most common. The interaction site is a grove or cleft, open in both
ends to allow the accommodation of the ligand. Multiple subsites along
the structure hold the sugar chain and specificity is often dependent on
hydrogen bonds and the correct positioning of aromatic residues
(Johnson et al., 1996; Simpson et al., 2000).
Oligosaccharides too short to be bound by type B CBMs interact with type
C CBMs. These have a pocket shaped interaction site and are therefore
limited to binding short oligosaccharides or the ends of glycans (as exo-
acting enzymes) (Boraston et al., 2001; Nootenboom et al., 2002; Abbott
et al., 2008).
Figure 10. CBMs are grouped into type A, B and C based on how they interact with their
ligands (after www.CAZypedia.org).
33
CBMs have been discovered that are promiscuous in their binding,
allowing for interaction with multiple ligands (Charnock et al., 2002; van
Bueren et al., 2005; van Bueren and Boraston, 2007; Boraston et al.,
2003) or that have multiple binding sites (Boraston et al., 2001; Henshaw
et al., 2004). Carbohydrate active enzymes can also contain more than
one CBM allowing for either stronger interaction with one type of ligand
or for interaction with several different ligand types, depending on the
specificity of the CBMs (Bolam et al., 2001; Boraston et al., 2003).
1.4.2. CBM applications
The highly specific interaction between CBMs and their ligands has been
exploited in a number of applications (reviewed in Oliviera et al., 2015).
The use of recombinant CBMs is common for applications as is some level
of protein engineering on the CBMs, such as enhancement of their CBMs
binding abilities or fusion to other proteins.
Fusion proteins are used as probes in microscopy for visualization of the
carbohydrate components of the cell wall (McCartney et al., 2004; Ding et
al., 2006) and as affinity tags for purification of recombinant proteins
(Shpigel el al., 1998b; Kavoosi et al., 2004) or for immobilization of
proteins or whole cells on carbohydrate surfaces (Francisco et al., 1993;
Lehtio et al., 2001). The efficiency of enzymatic reactions can also be
enhanced by engineering CBMs, altering or enhancing substrate
recognition by the enzyme (Limon et al., 2001; Ravalason et al., 2009).
CBMs can be utilized in the material field for the modification of fibres or
for crosslinking carbohydrate materials (Cavaco–Paulo et al., 1999;
Kitaoka et al., 2001; Levy et al., 2002; Gustavsson et al., 2004). Most
commonly this has been done by addition of recombinant CBMs to a
process, but attempts to modify the fibre or plant cell wall in planta as it
is biosynthesized have also been made (Safra-Dassa et al., 2006; Obembe
et al., 2007; Nardi et al., 2015; Perini et al., 2017). Growth effects, cell
wall impact and fibre properties through in vivo expression of CBMs are
explored in publications III and IV of this thesis.
34
1.4.3 CBM3 from CipA of Clostridium thermocellum
The carbohydrate-binding module 3 (CBM3) used in publications III and
IV of this thesis originates from the non-catalytic scaffolding protein
cellulosome-integrating protein A (CipA) found in the cellulosome of the
bacterium Clostridium thermocellum (Gerngross et al., 1993). The
cellulosome with its associated hydrolases is tethered to the cellulose
substrate through the CipA protein and its CBM (Kruus et al., 1995;
Demain et al., 2005).
CBM3 preferably binds to crystalline cellulose (Morag et al., 1995;
Georgelis et al., 2012; Ruel et al., 2012), specifically to the hydrophobic
face of the crystal (Lehtio et al., 2003; Ding et al., 2006; Dagel et al.,
2011). Hernandez-Gomez (2015) also reported a week interaction with
tamarind xyloglucan, however with a 50-100 times lower affinity than for
cellulose. CBM3 does not bind cello- or xyloglucan oligosaccharides
(Hernandez-Gomez et al., 2015).
The CBM3 is a small protein of ~150 amino acids. The structure has been
determined to consist of a jelly-roll topology, made up of a nine-stranded
β sandwich (Figure 11) (Tormo et al., 1996). Ligand interaction is
proposed to take place by hydrogen bonding and aromatic stacking
interactions through amino acids found in a planar array on the surface of
the protein (Tormo et al., 1996). Mutation of the five linearly arranged
amino acids leads to loss of cellulose binding, while single mutations do
not abolish binding (Figure 11, right) (Hernandez-Gomez et al., 2015). A
groove located on the opposite side of the protein containing several
conserved residues forms a potential second ligand interaction site
(Tormo et al., 1996). The precise function of this site has however not
been determined.
The Clostridium thermocellum CBM3 has been used extensively in
research for the labelling of crystalline cellulose (Blake et al., 2006; Kljun
et al., 2011; Ruel et al., 2012; Zheng et al., 2018). It has a ~50% sequence
identity with CBM3 from Clostridium cellulovorans, which is also a well
characterized CBM (Goldstein et al., 1993).
35
Figure 11. The structure of CBM3 from C. thermocellum viewed looking down three (1,2,3)
cellulose chains (upper left) and along a single cellulose chain (lower left). Proposed
residues (yellow) involved in interaction with the cellulose chains in the cellulose surface
(white) viewed from above (right) (Adapted with permission from Tormo et al., 1996).
36
37
2. Present investigation
2.1. Aim of the present investigation
The overall aim of this thesis is to contribute to a better understanding of
the plant primary cell wall structure and its components. As stated in the
introduction, this thesis has two distinct parts. The first part covers the
optimization of two methods for determining the enzymatic activity of the
glycosyltransferases important for cell wall biosynthesis (Publication
I). The radiometric method was then applied in the study of the
proteome and the associated callose synthase activity of the
plasmodesmata, a vital cell wall structure (Publication II).
In the second part, an attempt to introduce structural changes to the cell
wall aiming at increasing the amount of sugars that can be extracted for
bioethanol production is presented. The in vivo effect of a carbohydrate
binding module on the structure of the primary cell wall was investigated
(Publication III) and the properties of nanocellulose particles derived
from the engineered cell walls were then characterized (Publication
IV).
38
2.2. Rationale, Results and Conclusion
2.2.1. Publication I: In vitro glycosyltransferase assays
Rationale
GTs play an important role in the formation of cell wall carbohydrates
and so far the functionally characterized representatives from this group
are few (Dhugga et al., 2004; Burton et al., 2006; Cocuron et al., 2007;
Liepman et al., 2005; Doblin et al., 2009; Purushotham et al., 2016).
Biochemical characterization of the GTs involved in cell wall biogenesis
would not only contribute to a better understanding of the overall process
of the formation of this important structure, but information gained could
also be exploited in the protection of crops, both by reinforcing the cell
wall of crops and by targeting GTs in pathogenic organisms such as fungi,
bacteria and oomycetes.
Possible ways of functionally characterizing GTs in plants has been
indirect through gain-of-function approaches (Burton et al., 2006; Doblin
et al., 2009) or through the study of mutants (reviewed in Doblin et al.,
2010). The use of oligosaccharide acceptors labelled with fluorophores
has also been used for the characterization of several GTs (Ishii et al.,
2002, 2004; Konishi et al., 2006; Lee et al., 2012). This method is
however limited by the need to separate the products by high-
performance anion-exchange chromatography (HPAEC) which means
that water-insoluble products cannot be monitored and that the
throughput is low. The large fluorescent group may also obstruct the
interaction of the enzyme with its acceptor. Additional methods applied
to the characteriszation of GTs from other sources include monitoring the
pH change that takes place as a result of the GT reaction (Deng and
Cheng, 2004), chromatographic methods following the change in amount
of donor nucleotidesugar or product nucleoside diphosphate using UV
detection (Kopp et al., 2007) or mass spectrometry (Yang et al., 2005).
A generally applicable method for assaying and biochemically
characterizing GTs is therefore desirable. Furthermore, a standardised
method for assaying GTs facilitates correct interpretation and
comparisons of results within the scientific community.
39
Results
Two methods for assaying GTs were optimized and are presented in detail
in this publication with a focus on the assay of plant GTs (Figure 12). The
spectrophotometric method uses a coupled enzymatic reaction where the
nucleoside diphosphate (NDP) produced by the GT reaction is
phosphorylated to a nucleoside triphosphate (NTP) by pyruvate kinase
(Gosselin et al., 1994). The pyruvate released from this reaction is then
used as a substrate by lactate dehydrogenase and converted to lactate. In
this process the reduced form of nicotinamide adenine dinucleotide
(NADH) is oxidised to NAD+, a conversion that can be monitored through
measurement of the absorbance at 340 nm. The GT reaction is thus
coupled to a change in NAD+ levels and indirectly monitored.
In the radiometric assay, the amount of incorporated radioactive
monosaccharide in the produced polysaccharide is measured after
removal of the excess of radiolabelled NDP-sugar. Depending on whether
the product is ethanol-insoluble or soluble the means of removal are
different. Ethanol-insoluble polysaccharides are retained on a glass-fibre
filter and any remaining substrate is washed away. If the product is
ethanol-soluble or displays a broad size distribution, the excess of NDP-
sugar is removed using an anion-exchange chromatography column
(Egelund et al., 2006).
Both methods can be used for assaying processive and non-processive
GTs and for enzymes producing soluble and insoluble products. The
radiochemical method has the further advantage that it can distinguish
between soluble and insoluble products. It can also be used for relatively
impure enzyme sources such as membrane fractions or detergent
extracts, while the enzymes used in the spectrophotometric method are
sensitive to impurities that may be present in such crude extracts. For
spectrophotometric assays of GTs, the procurement of a highly enriched
and pure enzyme source is consequently crucial. The radiometric assay is
therefore more applicable to initial screening for activities or
investigations of uncharacterized enzymes. A drawback with the
radiometric assay is that with the use of radiolabelled substrates comes
the need for careful attention to lab security protocols and requirements
for dedicated lab spaces and special waste management.
40
Figure 12. Overview of radiometric and spectrophotometric assays for glycosyltransferases
(Reprinted with permission from Brown et al., 2012).
41
Conclusion
This study gives a comprehensive yet easy-to-follow protocol for
determining the enzymatic activity of GTs using two alternative methods.
The choice of the method for biochemical characterization of an enzyme
must take into account the source and purity of the enzyme and the
expected level of activity.
2.2.2. Publication II: The Populus plasmodesmata proteome
Rationale
The plasmodesmata is a membrane lined pore-like structure that has an
important role in regulating the flow of material between plant cells, and
it acts as a signalling hub and point of control of pathogen spread
(reviewed in Roberts and Oparka, 2003). While the qualitative proteome
of Arabidopsis PD has been previously published (Fernandez-Calvino et
al., 2011) a second, semi-quantitative, proteome would further contribute
to identifying proteins highly enriched in PD and to expanding our
understanding of the different functional roles that the structure plays in
plants.
The β-1,3-glucan callose is involved in both the transient and more
permanent closure of the PD (reviewed in De Storme and Geelen, 2014).
While the negative effect of callose deposition on conductivity of the PD
has been demonstrated (Radford et al., 1998; Sivaguru et al., 2000; Rinne
et al., 2005; Levy et al., 2007; Guseman et al., 2010; Vaten et al., 2011),
callose synthase activity in PD enriched samples has to our knowledge
never been assayed. Further we found it interesting to investigate if the
callose synthase activity found in PD enriched samples was calcium
dependent. The reasoning behind this is that fluctuations in cytoplasmic
Ca2+ levels have been connected to the regulation of PD closure (Tucker
and Boss, 1996; Holdaway-Clark et al., 2000). Since callose synthase
activity occurs in two forms, both Ca2+ dependent (more commonly,
Kauss, 1985; Köhle et al., 1985; Schlupmann et al., 1993; Colombani et al.,
2004) and independent (Schlupmann et al., 1993), calcium levels may be
one possible regulatory way to control callose deposition and cell-to-cell
movement through PD.
42
Results
Two membrane fractions, microsomal and plasmodesmata enriched,
were isolated from cell suspension cultured P. trichocarpa cells through
enzymatic degradation of the cell wall followed by differential
centrifugation to enrich the membrane fraction of the PD. In total 1113
unique proteins were identified from the PD sample by mass
spectrometry (MS) analysis. From these, 201 were further shown to be
highly enriched in the PD fraction using spectral counting and strict
filtering criteria.
Proteins identified by MS were cross-referenced with databases and
processed using online available software to determine predicted
properties such as protein topology, post-translational modifications,
cellular localization and involvement in biological processes. Results
showed that the proteins enriched in the PD fraction mainly represent the
following functional classes: signal transduction, transporters,
intracellular trafficking, stress related, metabolism and unknown
function. Many of the proteins, such as remorin, callose synthases,
plasmodesmata located proteins, tetraspanins and reversibly glycosylated
polypeptides were previously reported to be PD located in other
organisms. This highlights the similarities in PD protein composition
between plants and also contributes to validating the isolation of PD. A
large proportion of the PD proteins were intrinsic membrane proteins.
This fraction further had a subpopulation with distinctly longer TMDs
compared to proteins from the microsomal fraction.
MS data showed three callose synthases to be among the highly enriched
PD proteins, GSL5, 10 and 12. Other highly enriched proteins also related
to callose metabolism were four predicted β-1,3-glucanases, possibly
involved in callose hydrolysis at the PD, therby restoring PD conductivity
(Levy et al., 2007; Zavaliev et al., 2013), and four “purple” acid
phosphatases suggested to be involved in
phosphorylation/dephosphorylation of the callose synthases (Kaida et al.,
2009). In vitro assays showed callose synthase activity found in PD
fractions was highly enriched compared to the microsomal fraction, thus
indicating the successful enrichment of PD (Figure 13). Callose synthase
activity in the PD fractions included the production of both insoluble and
soluble products, but the insoluble product was proportionally more
abundant (Figure 13). Furthermore, the activity was found to be both
43
cation dependent and independent, in contrary to most forms of studied
callose synthase activities that are dependent on Ca2+ (Figure 14).
Conclusion
This study identifies the first semiquantitative proteome of PD from a
tree species, the model organism P. trichocarpa. Out of 1113 identified
proteins in the PD fraction, 201 proteins where enriched. Forty-three
percent were previously identified as PD located according to the TAIR
database, leaving over half of the proteins as potentially new tree PD
specific proteins. The PD enriched proteins represent several functional
protein groups which highlights the importance of these structures in
different cell functions, i.e., signalling, stress responses and trafficking
and transport. Callose synthase activity in PD enriched samples was
found in both a cation dependent and independent form. The detection of
this novel form of cation independent activity at the PD will hopefully
contribute to the ongoing exploration of PD regulation.
Figure 13. Callose synthase activity measured as nmol of glucose incorporated into insoluble
and soluble products. Significant differences between MF and PEF samples were found
(columns marked by asterisks) (p<0,001 for both insoluble and soluble samples, Students t-
test). Bars indicate standard deviation. Microsomal fraction (MF), cell wall (CW),
plasmodesmata-enriched fraction (PEF). (Reprinted with permission from Leijon et al.,
2018).
44
Figure 14. Remaining callose synthase activity measured under ion depleted conditions
(p<0,001 for both insoluble and soluble samples, Students t-test). Bars indicate standard
deviation. Microsomal fraction (MF), plasmodesmata-enriched fraction (PEF). (Reprinted
with permission from Leijon et al., 2018)
2.2.3. Publication III: The effect of carbohydrate-binding modules
(CBMs) on plant cell wall properties: an in vivo approach
Rationale
Making use of waste material from forest and agricultural industries for
production of bioethanol in order to meet the world increasing need for
alternatives to fossil fuels has in part been limited by the high energy
consumption of this process. Increasing the yield in bio-refineries and
minimizing the need for chemical and heat pretreatment of raw materials
in order to make lignocellulose more accessible would lead to a more
efficient bioethanol production.
Through in planta expression of a carbohydrate-binding module, a
heterologous non-catalytic cellulose binding protein, we hypothesized
that cellulose crystallinity could be affected, leading to a cell wall more
efficiently degraded to its sugar monomers. Cell suspension cultures of
Nicotiana tabacum and Arabidopsis thaliana plants expressing CBM3
were generated using Agrobacterium-mediated transformation. Cellulose
and hemicellulose fractions were extracted from these cell walls for
further analysis.
45
Results
The transformed tobacco cells expressing CBM3 displayed an elongated
morphology compared to the wild type (WT) cells. Further investigation
of the cross section of the cell wall using TEM revealed a disaggregated
outer cell wall layer (Figure 15).
Saccharification efficiency of the CBM3 line showed a 36% increase using
a NaOH pretreatment (Figure 16). As saccharification efficiency has been
coupled to the degree of cellulose crystallinity cellulose characterization
was performed. The analysis indicated no changes in the amount of
crystalline cellulose (Updegraff), nor to the cellulose crystallinity (solid-
state NMR). As the anticipated alterations to cellulose were notably
absent, a more thorough cell wall analysis was undertaken.
Monosaccharide composition showed increased amounts of uronic acids
and a moderate decrease in galactose and arabinose content in the CBM3
expressing cells, pointing toward changes in the pectic acide composition.
Using mild hydrolytic conditions (TFA release) an 80% increase in the
amount of non-cellulosic glucose was detected. An increase was also
Figure 15. Cell wall ultrastructure of wild-type (WT) and cells expressing CBM3 (CBM3)
visualized by transmission electron microscopy. Type A cell walls are without adjacent cell
wall and type B cell walls have an adjoining cell.
46
Figure 16. Enzymatic saccharification of the BY-2-transformed cell line and Arabidopsis
plants. Pretreatment of cell walls was performed with or without 0.5 M NaOH for 30 min at
90ºC and the remaining material hydrolyzed using an enzyme mixture with a 4:1 ratio of
Celluclast and Novozyme 188. The released reducing sugars were quantified using the 3-
methyl-2-benzothiazolinonehydrazone method (Gomez et al., 2010).
observed for fully hydrolysed cell wall samples, but to a much lesser
extent. Further analysis of the hemicellulose fraction (H2SO4 treatment)
was undertaken to determine if the increase in glucose amount was due to
changes to the hemicellulose or non-crystalline form of cellulose. Linkage
analysis of the hemicellulose fraction showed a 30% increase in 1,4-linked
glucose for the transformed line. Quantification of xyloglucan using the
Kooiman assay showed a 13% increase for the CBM3 line. Linkage
analysis also revealed a reduction in 1,2-linked xylose, terminal xylose
and terminal arabinofuranose units indicating that a change to
xyloglucan substitutions may have resulted from the transgene
expression.
Arabidopsis thaliana plants expressing CBM3 showed no macroscopic or
microscopic phenotypical changes. Cell wall analysis revealed a 16 %
decrease in crystalline cellulose (Updegraff), but only minor changes to
the overall monosaccharide composition. Cell walls from the CBM3 line
pretreated with NaOH had a lower saccharification than the WT line,
0
10
20
30
40
50
60
0.5M NaOHTobacco
0.5M NaOHArabidopsis
No NaOHArabidopsis
nm
ol o
f su
gars
re
leas
ed
/g
mat
eri
al. h
ou
r
WT
CBM3
47
while a second saccharification test omitting the NaOH from the
pretreatment resulted in overall lower sugar yields for both samples but
an increase of released sugars by 79% for the CBM3 line (Figure 16). We
speculate that this is the result of the harsher pretreatment removing the
more accessible cell wall components.
Conclusions
In vivo expression of CtCBM3 caused morphological changes to the
tobacco cells and had a positive effect on the saccharification of the
primary cell walls of suspension cultured cells. Although the cell wall was
thoroughly characterized, it could not be determined exactly what
structural changes caused the increase in degradability. We see two
possible explanations to the increase in 1,4-β-glucans found in both the
cell wall and hemicellulose fractions: either the amount of non-crystalline
cellulose is affected or modifications to the amount and structure of
xyloglucan is taking place. Overall our results indicate that the increased
saccharification is due to other factors than changes in cellulose
crystallinity. The increased saccharification without chemical
pretreatment together with the absence of detrimental phenotypes in
Arabidopsis plants expressing the CBM3, suggests that this approach to
plant cell wall engineering for increased saccharification is a promising
option.
2.2.4. Publication IV: Stronger cellulose microfibrils network
structure through in vivo expression of CBMs
Rationale
CBM3 from Clostridium thermocellum has previously been shown to
bind to crystalline cellulose (Lehtio et al., 2003; Hervé et al., 2010; Dagel
et al., 2011; Ruel et al., 2012). While publication III presents a
characterization of the changes to the primary cell wall that resulted from
the overexpression of the CBM3 it was not clear what impact it had on the
properties of the cellulose. Therefore the effect of the in vivo presence of
CBM3 on cellulose was investigated through the further characterization
of CMFs and CNCs. Nanopaper was also produced in order to better
understand the mechanical properties of the resulting material.
48
Results
FE-SEM showed that cellulose was extracted in the form of “cell ghosts”,
i.e. essentially flattened cells, and that an increased aspect ratio was
visible in the CBM3 expressing cells when compared to WT.
Light transmittance measurements indicated that 3 minutes of
ultrasonication was enough for the complete defibrillation of cellulose
from both the WT and CBM3 lines. Compared to the treatment used for
CMF preparation from primary cell walls from other sources this method
was much milder (Dinand et al., 1996; Dufresne and Vignon, 1998;
Niimura et al., 2010). As the samples resulting from the CBM3 line
demonstrated higher light transmittance than the WT, this would indicate
that the CMFs in this sample were less aggregated. Comparison of the WT
and CBM3 CMFs structure performed by TEM and AFM showed that the
fibres were completely individualized, with a width of 2-3 nm, and had a
probable length of several micrometres. No morphological differences
between the samples could be found.
After acid hydrolysis of the extracted cellulose the average length of the
CBM3-CNCs (201 nm) was found to be significantly higher than that of
WT-CNCs (122 nm) (Figure 17a and b). The yield after hydrolysis was also
higher, 24% and 11% for CBM3 and WT respectively, indicating that the
cellulose extracted from the CBM3 expressing cells resists acid hydrolysis
better and therefore may contain more ordered cellulose chains.
Stress-strain curves (Figure 18) resulting from the mechanical testing of
papers prepared from the different CMFs showed that the CBM3 line
paper had higher mechanical properties, both regarding tensile strength
and ductility (Table 1). The CBM3 line paper presents a twice as high
work-to-fracture value compared to its WT counterpart (Table 1). These
enhanced properties were attributed to the higher ordered domains. FE-
SEM imaging of the tensile-fractured surface of the nanopaper derived
from the CBM3 line showed a more porous structure with multiple CMFs
pulled out of the surface (Figure 19b). This suggests that the stronger
nanofibrillar network formed by the CMFs from the CBM expressing cells
had altered deformation behaviour. The higher strain-to-failure and more
apparent porosity observed was likely due to more sliding taking place
between the laminated sheets of the paper. When extrapolated to the
primary cell wall, the properties of the microfibril network may give rise
49
to a cell wall with enhanced toughness and which is more inclined to
stretch, explaining the elongated morphology of the transformed cells.
Figure 17. Images of CNCs and histograms with their corresponding length distributions.
WT line (a), CBM3 line (b). (Adapted with permission from Butchosa Robles, 2014).
Figure 18. Stress-strain curves of nanopapers made from CNFs isolated from wild type (WT)
and CBM3 (CBM) lines (Reprinted with permission from Butchosa Robles, 2014).
50
Table 1. Mechanical properties of nanopapers. Papers were prepared with CNFs extracted
from WT or CBM3 expression lines (Adapted with permission from Butchosa Robles,
2014).
Sample Density
(g cm-3)
Modulus
(GPa)
Tensile strength
(MPa)
Strain-to-failure
(%)
Work-of-fracture
(MJ m-3)
WT 1.42 ± 0.01 8.3 ± 0.5 143 ± 8 2.3 ± 0.4 192 ± 46
CBM3 1.41 ± 0.03 9.3 ± 0.2 198 ± 9 3.6 ± 0.4 438 ± 72
Figure 19. Cross-sections of the tensile-fractured surface of nanopapers cast from CMFs
from the WT line (a) and CBM3 line (b) (Adapted with permission from Butchosa Robles,
2014).
Conclusions
In conclusion, this work showed that overexpressing the carbohydrate
binding module 3 from Clostridium thermocellum in Nicotiana tabacum
suspension cultured cells resulted in a higher CNC yield and CNCs with
an increased length. The nanopaper prepared from CMFs extracted from
the same cells displayed enhanced toughness, demonstrated through
higher tensile strength and strain-to-failure values. We propose that this
is because cellulose from the CBM3 expressing line has CMFs with
ordered regions containing longer stretches of cellulose of a higher order
compared to the WT line, and that this result in a stronger cellulose
microfibril network.
51
2.3 Concluding remarks and future perspectives
The overall aim of this thesis was to study aspects of primary cell wall in
plant cell suspension cultures, a model system used in our group for the
study of enzymes involved in cell wall biosynthesis. Specifically, we were
interested in the protein composition of membrane fractions derived
from plasmodesmata and the GTs localized to this structure. We also
wanted to produce cell walls with enhanced susceptibility to enzymatic
degradation through introducing changes to cellulose structure at the
level of fibril formation.
The detailed methods description of how to assay GTs, found in
Publication I, facilitates biochemical characterization of enzymes
putatively responsible for catalysing some of the major cell wall
polysaccharides in primary cell walls. It is our hope that the methods
paper through outlining the protocol for two alternative GT activity
assays in a structured and easy to follow manner will contribute to the
successful biochemical characterization of additional GTs.
Further studies of the proteins related to the cell wall were performed in
Publication II were MS was used to determine the proteins enriched in
membrane fractions of the cell wall spanning structure plasmodesmata.
Comparison of the previously known proteome of the PD of suspension
cultured Arabidopsis cells to the PD proteome from Populus trichocarpa
suggests that certain protein families and functional classes are recurring
in these structures. The availability of a second PD proteome will
hopefully facilitate selection of interesting PD-specific proteins for further
studies, through conditional mutants and localization experiments from
the complex and most likely shifting protein population of the PD. The
characterization of a monocot PD proteome (tentatively maize) may also
reveal differences in PD composition among higher plants.
The work-flow for determining the PD proteome could be expanded to
identify PD proteins associated to mechanical, chemical or pathogen
related stress as the growth conditions of the cell suspension cultures are
easy to manipulate.
52
The finding that callose synthase activity in PD-enriched membranes is
not dependent on Ca2+ for activity is interesting both in the context of the
PD and more generally with respect to callose synthases. Further
investigations into the optimal conditions for PD-derived callose synthase
activity may contribute to shed light on the role of Ca2+ in regulation of
callose deposition at the PD. To which degree the different callose
synthases found to be enriched in PD fractions contribute to the
measured activity and if their cation dependency varies is an additional
aspect worth exploring.
The specific composition of the PD membrane with an enrichment in
sterols and sphingolipids with long chain saturated fatty acids (Grison et
al., 2015) suggests that the PD membranes may contain domains
resembling DRMs. Furthermore, DRMs extracted from the PM of P.
trichocarpa cells were shown to contain a sub-population of proteins
likely originating from the PD (Srivastava et al., 2013). Studies of the PD-
derived DRM fraction using the traditional DRM isolation protocol
(Triton X-100 solubilization and centrifugation on a sucrose gradient)
was hindered by the small volumes of PD-enriched membranes that are
typically obtained with the current isolation protocol. If this obstacle
would be overcome, investigations of the proteome of PD-derived DRM
fractions would be an interesting contribution to the field.
In Publications III and IV, focus was given to the polysaccharide
components of the cell wall. In order to further understand what features
of the cell wall are of importance for digestibility, cell wall engineering
aimed to introduce changes to cellulose structure was undertaken and a
thorough cell wall characterization was performed.
Engineering of cell walls for more efficient degradation to sugar
monomers allowed the production of plants with increased
saccharification, without the requirement for chemical pre-treatment of
the cell wall. While this is a promising result, the approach would have to
be tested further in a plant more adapted to large scale biomass
production than Arabidopsis. Optimization of the pretreatment
conditions would also be needed to increase the overall sugar yield.
As no conclusive answer could be given to why CBM expression in
tobacco results in higher saccharification, a more detailed
53
characterization of the cellulose fraction would be of interest. Answers
may also be found through further investigations of the pectin
components of the cell wall or by characterizing the xyloglucan structure.
While we succeeded in the engineering of plant cell walls for more
efficient release of sugar monomers (Publication III), further
investigations in Publication IV on the properties of the resulting
nanoparticles and materials were needed to form a hypothesis regarding
the effect of heterologous CBM3 expression on cellulose structure. The
paper produced from CMFs derived from the engineered cells exhibited
enhanced mechanical properties. CNC preparation from transgenic cell
walls resulted in higher yields and longer CNCs, suggesting that the
plants expressing CBM3 may contain a different arrangement of the
ordered regions in cellulose.
The possibility to change the properties of cellulose nanofibers through
interfering with cellulose biosynthesis is an interesting concept. This will
be further explored through different approaches as the process of
biosynthesis and the enzymes involved in synthesising cellulose become
better characterized.
As primary cell walls are not a practical source of cellulose for scale up
and applications, it would be interesting from a material science
perspective to determine whether the effects observed in this study
persists when applied to secondary cell wall cellulose biosynthesis, for
example in a tree species.
54
55
3. Acknowledgments
This thesis has been a long time in the making, which means that I have
had the privilege to meet many fine people during my years at the
division of Glycoscience. Thank you all!
I started of doing my master thesis at the department and then got a PhD
position. Thank you to my main supervisor Professor Vincent Bulone for
giving me the opportunity to pursue a PhD, for always taking time to
answer questions and discuss results, and for you enthusiasm and
positive view on science.
Thank you to my co-supervisor Dr Vaibhav Srivastava who has been a
great supervisor and discussion partner during the final years of my
studies. I would also like to thank my other colleagues who have
supervised me in the lab; Erik, Gea, Johanna and Sara. I am very thankful
that you took the time to teach me the hands-on aspects of science. A
special thank you to Erik and Johanna for taking so good care of me when
I was new in the group.
The publications that comprise this thesis would have never been if it was
not for the expertise and hard work of my fantastic collaborators. I would
especially like to thank Hugo, Núria and Christian.
Thank you to Lauren for giving many valuable suggestions regarding the
plasmodesmata project and for proofreading this thesis.
I am also grateful to Anna for sharing her great knowledge of plant cell
cultures. All the little tips and tricks have been much appreciated through
the years.
Thank you to Ela and Annie for help in the lab and to Nina for
administration.
Finally, I am grateful to my grandmother Lillian and my mother Julia for
introducing me to the wonders of plants as a child. I am sure that my
green fingers and love of plants is inherited and that I would not have
chosen this field of research if it was not for you.
56
57
4. References
Abbott, D.W., Eirin-Lopez, J.M., and Boraston, A.B. (2008) Insight into ligand
diversity and novel biological roles for family 32 carbohydrate-binding modules. Mol. Biol.
Evol., 25, 155–167.
Albersheim, P., Darvill, A., Roberts, K., Sederoff, R., and Staehelin, A. (2011)
Plant cell walls: From chemistry to biology. Garland Science, New York.
Amor, Y., Haiglerm, C.H., Johnson, S., Wainscott, M., and Delmer, D.P. (1995) A
membrane-associated form of sucrose synthase and its potential role in synthesis of
cellulose and callose in plants. Proc. Natl. Acad. Sci. USA, 92, 9353–9357.
Anderson, C.T., Carroll, A., Akhmetova, L., and Somerville, C. (2010) Real‐time
imaging of cellulose reorientation during cell wall expansion in Arabidopsis roots. Plant
Physiol., 152, 787–796.
Andrawis, A., Solomon, M., and Delmer, D.P. (1993) Cotton fiber annexins: a
potential role in the regulation of callose synthase. Plant J. 3, 763–772.
Apostolakos, P., Livanos, P., Nikolakopoulou, T.L., and Galatis, B. (2009) The
role of callose in guard cell wall differentiation and stomatal pore formation in the fern
Asplenium nidus. Ann. Bot. 104, 1373–1387.
Araki, J., Wada, M., and Kuga, S. (2001) Steric stabilization of a cellulose microcrystal
suspension by poly(ethylene glycol) grafting. Langmuir, 17, 21–27.
Atalla, R.H., and Vanderhart, D.L. (1984) Native cellulose: a composite of two distinct
crystalline forms. Science, 223, 283–285.
Atanassov, I.I., Pittman, J.K., and Turner, S.R. (2009) Elucidating the mechanisms
of assembly and subunit interaction of the cellulose synthase complex of Arabidopsis
secondary cell walls. J. Biol. Chem., 284, 3833–3841.
Barton, D.A., Cole, L., Collings, D.A., Liu, D.Y., Smith, P.M., Day, D.A., and
Overall, R.L. (2011) Cell-to-cell transport via the lumen of the endoplasmic reticulum.
Plant J. 66, 806–817.
Beeckman, T., and Przemeck, G. (2002) Genetic complexity of cellulose synthase A
gene function in Arabidopsis embryogenesis. Plant Physiol., 130, 1883–1893.
Benitez–Alfonso, Y., Cilia, M., San Roman, A., Thomas, C., Maule, A., Hearn, S.,
and Jackson, D. (2009) Control of Arabidopsis meristem development by thioredoxin-
dependent regulation of intercellular transport. Proc. Natl. Acad. Sci. USA, 106, 3615–3620.
Benitez–Alfonso, Y., and Jackson, D. (2009) Redox homeostasis regulates
plasmodesmal communication in Arabidopsis meristems. Plant Signal. Behav., 4, 655–659.
Benitez–Alfonso, Y., Faulkner, C., Ritzenthaler, C., and Maule, A.J. (2010)
Plasmodesmata: gateways to local and systemic virus infection. Mol. Plant–Microbe
Interact., 23, 1403–1412.
58
Benitez–Alfonso, Y., Faulkner, C., Pendle, A., Miyashima, S., Helariutta, Y.,
and Maule, A. (2013) Symplastic intercellular connectivity regulates lateral root
patterning. Dev. Cell, 26, 136–147.
Bessueille, L., Sindt, N., Guichardant, M., Djerbi, S., Teeri, T.T., and Bulone, V.
(2009) Plasma membrane microdomains from hybrid aspen cells are involved in cell wall
polysaccharide biosynthesis. Biochem. J., 420, 93–103.
Bhalerao, R., Nilsson, O., and Sandberg, G. (2003) Out of the woods: forest
biotechnology enters the genomic era. Curr. Opin. in Biotechnol., 14, 206–213.
Bilska, A., and Sowinski, P. (2010) Closure of plasmodesmata in maize (Zea mays) at
low temperature: a new mechanism for inhibition of photosynthesis. Annals of Botany, 106,
675–686.
Blake, A.W., McCartney, L., Flint, J.E., Bolam, D.N., Boraston, A.B., Gilbert,
H.J., and Knox, J.P. (2006) Understanding the biological rationale for the diversity of
cellulose–directed carbohydrate–binding modules in prokaryotic enzymes. J. Biol. Chem.,
281, 29321–29329.
Blaschek, W., Koehler, H., Semler, U., and Franz, G. (1982) Molecular weight
distribution of cellulose in primary cell walls. Planta, 154, 500–555.
Bluhm, T.L., and Sarko, A. (1977) The tripel helical structure of lentinan, a linear ß (1–
3)–glucan. Can. J. Chem., 55, 293–299.
Bolam, D.N., Ciruela, A., McQueen–Mason, S., Simpson, P., Williamson, M.P.,
Rixon, J.E., Boraston, A., Hazlewood, G.P., and Gilbert, H.J. (1998) Pseudomonas
cellulose–binding domains mediate their effects by increasing enzyme substrate proximity.
Biochem. J., 331, 775–781.
Bolam, D.N., Xie, H., White, P., Simpson, P.J., Hancock, S.M., Williamson,
M.P., and Gilbert, H.J. (2001) Evidence for synergy between family 2b carbohydrate
binding modules in Cellulomonas fimi xylanase 11A. Biochemistry, 40, 2468–2477.
Bootten, T.J., Harris, P.J., Melton, L.D., and Newman, R.H. (2004) Solid–state
13C–NMR spectroscopy shows that the xyloglucans in the primary cell walls of mung bean
(Vigna radiata L.) occur in different domains: a new model for xyloglucan–cellulose
interactions in the cell wall. J. Exp. Bot., 55, 571–583.
Boraston, A.B., McLean, B.W., Guarna, M.M., Amandaron–Akow, E., and
Kilburn, D.G. (2001) A family 2a carbohydrate–binding module suitable as an affinity tag
for proteins produced in Pichia pastoris. Protein Expr. Purif., 21, 417–423.
Boraston, A.B., Kwan, E., Chiu, P., Warren, R.A., and Kilburn D.G. (2003)
Recognition and hydrolysis of noncrystalline cellulose. J. Biol. Chem., 278, 6120–6127.
Boraston, A.B., Bolam, D.N., Gilbert, H.J., and Davies, G.J. (2004) Carbohydrate–
binding modules: fine–tuning polysaccharide recognition. Biochem. J., 382, 769–781.
Borner, G.H.H., Sherrier, D.J., Weimar, T., Michaelson, L.V., Hawkins, N.D.,
MacAskill, A., Napier, J.A., Beale, M.H., Lilley, K.S., and Dupree, P. (2005)
Analysis of detergent–resistant membranes in Arabidopsis. Evidence for plasma membrane
lipid rafts. Plant Physiol., 137, 104–116.
59
Bourne, Y., and Henrissat, B. (2001) Glycoside hydrolases and glycosyltransferases:
families and functional modules Curr. Opin. Struct. Biol., 11, 593–600.
Braidwood, L., Breuer, C., and Sugimoto, K. (2014) Mybody is a cage: mechanisms
and modulation of plant cell growth. New Phytol., 201, 388–402.
Breton, C., Bettler, E., Joziasse, D.H., Geremia, R.A. and Imberty, A. (1998)
Sequence-function relationships of prokaryotic and eukaryotic galactosyltransferases. J.
Biochem., 123, 1000-1009.
Briolay, A., Bouzenzana, J., Guichardant, M., Deshayes, C., Sindt, N.,
Bessueille, L., and Bulone, V. (2009) Cell wall polysaccharide synthases are located in
detergent–resistant membrane microdomains in oomycetes. Appl. Environ. Microbiol., 75,
1938–1949.
Brown, R.M., and Montezinos, D. (1976) Cellulose microfibrils: visualization of
biosynthetic and orienting complexes in association with the plasma membrane. Proc. Natl.
Acad. Sci. USA, 73, 143–147.
Brown, R.M., Haigler, C., and Cooper, K. (1982) Experimental induction of altered
non–microfibrillar cellulose. Science, 218, 1141–1142.
Brown, R.M. (1996) The biosynthesis of cellulose. Part A. J. Macromol. Sci., 33, 1345–
1373.
Brownfield, L., Doblin, M., Fincher, G.B., and Bacic, A. (2009) Biochemical and
molecular properties of biosynthetic enzymes for (1,3)-β-glucans in embryophytes,
chlorophytes and rhodophytes. In chemistry, biochemistry, and biology of 1-3 β-glucans and
related polysaccharides. Academic Press, Amsterdam. pp. 283–326.
Buckeridge, M.S., dos Santos, H.P., and Tine, M.A.S. (2000) Mobilisation of storage
cell wall polysaccharides in seeds. Plant Physiol. Biochem, 38, 141–156.
Buckeridge, M.S. (2010) Seed cell wall storage polysaccharides: Models to understand
cell wall biosynthesis and degradation. Plant Physiol., 154, 1017–1023.
van Bueren, A.L., Morland, C., Gilbert, H.J., and Boraston, A.B. (2005) Family 6
carbohydrate binding modules recognize the non–reducing end of beta–1,3–linked glucans
by presenting a unique ligand binding surface. J. Biol. Chem., 280, 530–537.
van Bueren, A.L., and Boraston, A.B. (2007) The structural basis of alpha–glucan
recognition by a family 41 carbohydrate–binding module from Thermotoga maritima. J.
Mol. Biol., 365, 555–560.
Bulone, V., Chanzy, H., Gay, L., Girard, V., and Fevre, M. (1992) Characterisation
of chitin and chitin synthase from the cellulosic cell wall fungus Saprolegnia monoica. Exp.
Mycol., 16, 8–21.
Bulone, V., Fincher, G., and Stone, B.A. (1995) In vitro synthesis of a microfibrillar
(1–3)–beta–glucan by a ryegrass (Lolium multiflorum) endosperm (1–3)–beta–glucan
synthase enriched by product entrapment. Plant J., 8, 213–225.
60
Burton, R.A., Wilson, S.M., Hrmová, M., Harvey, A.J., Shirley, N.J., Stone,
B.A., Newbigin, E.J., Bacic, A., and Fincher, G.B. (2006) Cellulose synthase‐like CslF
genes mediate the synthesis of cell wall (1,3;1,4)‐β‐d‐glucans. Science, 311, 1940–1942.
Butchosa Robles, N. (2014) Tailoring cellulose nanofibrils for advanced materials.
Doctoral Thesis. KTH Royal Institute of Technology, ISBN 978–91–7595–329–8.
Caño Delgado, A., and Penfield, S. (2003) Reduced cellulose synthesis ‐ invokes
lignification and defense responses in Arabidopsis thaliana. Plant Cell, 34, 351–362.
Cantrill, L.C., Overall, R.L., and Goodwin, P.B. (1999) Cell–to–cell communication
via plant endomembranes. Cell. Biol. Int., 23, 653–661.
Carpita, N.C., and Gibeaut, D.M. (1993) Structural models of primary‐cell walls in
flowering plants ‐ consistency of molecular‐structure with the physical‐properties of the cell
wall during growth. Plant J., 3, 1–30.
Carroll, A., Mansoori, N., Li, S., Lei, L., Vernhettes, S., Visser, R.G.F.,
Somerville, C., Gu, Y., and Trindade, L.M. (2012). Complexes with mixed primary
and secondary cellulose synthases are functional in Arabidopsis plants. Plant Physiol., 160,
726–737.
Cavaco–Paulo, A., Morgado, J., Andreaus, J., and Kilburn, D. (1999) Interactions
of cotton with CBD peptides. Enzyme Microb. Technol., 25, 639–643.
Cavalier, D.M., Lerouxel, O., Neumetzler, L., Yamauchi, K., Reinecke, A.,
Freshour, G., Zabotina, O.A., Hahn, M.G., Burgert, I., Pauly, M., Raikhel, N.V.,
and Keegstra, K. (2008) Disrupting two Arabidopsis thaliana xylosyltransferase genes
results in plants deficient in xyloglucan, a major primary cell wall component. Plant Cell,
20, 1519–1537.
Chaffey, N., Cholewa, E., Regan, S., and Sundberg, B. (2002) Secondary xylem
development in Arabidopsis: a model for wood formation. Physiol. Plant., 114, 594–600.
Chanzy, H., Imada, K., Mollard, A., Vuong, R., and Barnoud, F. (1979)
Crystallographic aspects of sub–elementary cellulose fibrils occurring in the wall of rose
cells cultured in vitro. Protoplasma, 100, 303–316.
Chanzy, H., and Henrissat, B. (1985) Undirectional degradation of valonia cellulose
microcrystals subjected to cellulase action. FEBS Lett., 184, 285–288.
Charnock, S.J., and Davies, G J. (1999) Structure of the nucleotide-diphospho-sugar
transferase, SpsA from Bacillus subtilis, in native and nucleotide-complexed forms.
Biochemistry, 38, 6380-6385.
Charnock, S.J., Bolam, D N., Nurizzo, D., Szabó, L., McKie, V.A., Gilbert, H.J.,
and Davies, G J. (2002) Promiscuity in ligand–binding: The three–dimensional structure
of a Piromyces carbohydrate–binding module, CBM29–2, in complex with cello– and
mannohexaose. Proc. Natl. Acad. Sci. USA, 99, 14077–14082.
Chuah, C.T., Sarko, A., Deslandes, Y., and Marchessault, R.H. (1983) Triple–
helical crystalline structure of curdlan and paramylon hydrates. Macromolecules, 16, 1375–
1382.
61
Cifuentes, C., Bulone, V., and Emons, A.M.C. (2010) Biosynthesis of callose and
cellulose by detergent extracts of tobacco cell membranes and quantification of the polymers
synthesized in vitro. J. Integr. Plant Biol., 52, 221–233.
Citovsky, V., Ghoshroy, S., Tsui, F., and Klessig, D. (1998) Non–toxic concentrations
of cadmium inhibit systemic movement of turnip vein clearing virus by a salicylic acid–
independent mechanism. Plant J., 16, 13–20.
Cocuron, J.C., Lerouxel, O., Drakakaki, G., Alonso, A.P., Liepman, A.H.,
Keegstra, K., Raikhel, N., and Wilkerson, C.G. (2007) A gene from the cellulose
synthase–like C family encodes a β–1,4 glucan synthase. Proc. Natl. Acad. Sci. USA, 104,
8550–8555.
Coleman, H.D., Yan, J., and Mansfield, S.D. (2009) Sucrose synthase affects carbon
partitioning to increase cellulose production and altered cell wall ultrastructure. Proc. Natl.
Acad. Sci. USA, 106, 13118–13123.
Colombani, A., Djerbi, S., Bessueille, L., Blomqvist, K., Ohlsson, A., Berglund,
T., Teeri, T.T., and Bulone, V. (2004) In vitro synthesis of (1→3)–β–D–glucan (callose)
and cellulose by detergent extracts of membranes from cell suspension cultures of hybrid
aspen. Cellulose, 11, 313–327.
Cosgrove, D.J. (1997) Assembly and enlargement of the primary cell wall in plants. Annu.
Rev. Cell Dev. Biol., 13, 171–201.
Cosgrove, D.J. (2005) Growth of the plant cell wall. Nat. Rev. Mol. Cell Biol, 6, 850–861.
Cosgrove, D.J. (2015) Plant expansins: diversity and interactions with plant cell walls.
Curr. Opin. Plant Biol., 25, 162–172.
Coutinho, P.M., Deleury, E., Davies, G.J., and Henrissat, B. (2003) An evolving
hierarchical family classification for glycosyltransferases. J. Mol. Biol., 328, 307–317.
Crawford, K.M., and Zambryski, P.C. (2000) Subcellular localization determines the
availability of non–targeted proteins to plasmodesmatal transport. Curr. Biol., 10, 1032–
1040.
Cui, X.J., Shin, H.S., Song, C., Laosinchai, W., Amano, Y., and Brown, R.M.
(2001) A putative plant homolog of the yeast β–1,3–glucan synthase subunit FKS1 from
cotton (Gossypium hirsutum L.) fibers. Planta, 213, 223–230.
Cui, W., and Lee, J.Y. (2016) Arabidopsis callose synthases CalS1/8 regulate
plasmodesmal permeability during stress. Nat. Plants, 2, 16034.
Dagel, D.J., Liu, Y–S., Zhong, L., Luo, Y., Himmel, M.E., Xu, Q., Zeng,Y., Ding,
S–Y., and Smith,S. (2011) In Situ imaging of single carbohydrate–binding modules on
cellulose microfibrils. J. Phys. Chem., 115, 635–641.
Delmer, D.P. (1999) Cellulose biosynthesis: Exciting times for a difficult field of study.
Annu. Rev. Plant Physiol. Plant Mol. Biol., 50, 245–276.
Demain, A.L., Newcomb, M., and Wu, J.H.D. (2005) Cellulase, Clostridia, and
Ethanol. Microbiol. Mol. Biol. Rev., 69, 124–154.
62
Deng, C., and Chen, R.R. (2004) A pH-sensitive assay for galactosyltransferase. Anal.
Biochem., 330, 219–226.
Deslandes, Y., Marchessault, R.H., and Sarko, A. (1980) Triple–helical structure of
(1→3)–d–glucan. Macromolecules, 13, 1466–1471.
Desprez, T., Juraniec, M., Crowell, E.F., Jouy, H., Pochylova, Z., Parcy, F.,
Höfte, H., Gonneau, M., and Vernhettes, S. (2007) Organization of cellulose synthase
complexes involved in primary cell wall synthesis in Arabidopsis thaliana. Proc. Natl. Acad.
Sci. USA, 104, 15572–15577.
Dhugga, K.S., Barreiro, R., Whitten, B., Stecca, K., Hazebroek, J., Randhawa,
G.S., Dolan, M., Kinney, A.J., Tomes, D., Nichols, S., and Anderson, P. (2004)
Guar seed β–mannan synthase is a member of the cellulose synthase super gene family.
Science, 303, 363–366.
Din, N., Gilkes, N.R., Tekant, B., Miller, R.C., Warren, A.J., and Kilburn, D.G.
(1991) Non–hydrolytic disruption of cellulose fibers by the binding domain of a bacterial
cellulase. Bio–Technol., 9, 1096–1099.
Dinand, E., Chanzy, H., and Vignon, M.R. (1996) Parenchymal cell cellulose from
sugar beet pulp: preparation and properties. Cellulose, 3, 183–188.
Ding, B., Turgeon, R., and Parthasarathy, M.V. (1992) Substructure of freeze–
substituted plasmodesmata. Protoplasma, 169, 28–41.
Ding, S.Y., Xu, Q., Ali, M.K., Baker, J.O., Bayer, E.A., Barak, Y., Lamed, R.,
Sugiyama, J., Rumbles, G., and Himmel, M.E. (2006) Versatile derivatives of
carbohydrate–binding modules for imaging of complex carbohydrates approaching the
molecular level of resolution. Biotechniques, 41, 435–443.
Djerbi, S., Lindskog, M., Arvestad, L., Sterky, F., and Teeri, T.T. (2005) The
genome sequence of black cottonwood (Populus trichocarpa) reveals 18 conserved cellulose
synthase (CesA) genes. Planta, 221, 739–746.
Doblin, M.S., Melis, L.D., Newbigin, E., Bacic, A., and Read, S.M. (2001) Pollen
tubes of Nicotiana alata express two genes from different beta–glucan synthase families,
Plant Physiol., 125, 2040–2052.
Doblin, M.S., Pettolino, F.A., Wilson, S.M., Campbell, R., Burton, R.A., Fincher,
G.B., Newbigin, E., and Bacic, A. (2009) A barley cellulose synthase–like CSLH gene
mediates (1,3;1,4)–b–D–glucan synthesis in transgenic Arabidopsis. Proc. Natl. Acad. Sci.
USA, 106, 5996–6001.
Doblin, M., Pettolino, F., and Bacic, A. (2010) Plant cell walls: the skeleton of the
plant world. Funct. Plant Biol., 37, 357–381.
Dong, X., Hong, Z., Sivaramakrishnan, M., Mahfouz, M., and Verma, D.P.S.
(2005) Callose synthase (CalS5) is required for exine formation during microgametogenesis
and for pollen viability in Arabidopsis. Plant J., 42, 315–328.
Dufresne, A., and Vignon, M.R. (1998) Improvement of starch film performances using
cellulose microfibrils. Macromolecules, 31, 2693–2696.
63
Egelund, J., Petersen, B.L., Motawia, M.S., Damager, I., Faik, A., Olsen, C.E.,
Ishii, T., Clausen, H., Ulvskov, P., and Geshi, N. (2006) Arabidopsis thaliana RGXT1
and RGXT2 encode Golgi–localized (1,3)–α–d–xylosyltransferases involved in the synthesis
of pectic rhamnogalacturonan–II. Plant Cell, 18, 2593–2607.
Ehlers, K., and Kollmann, R. (2001) Primary and secondary plasmodesmata: structure,
origin, and functioning. Protoplasma, 216, 1–30.
Ellinger, D., and Voigt, C.A. (2014) Callose biosynthesis in Arabidopsis with a focus on
pathogen response: what we have learned within the last decade. Ann. Bot., 114, 1349–1358.
Ellis, C., and Turner, J.G. (2001) The Arabidopsis mutant cev1 has constitutively active
jasmonate and ethylene signal pathways and enhanced resistance to pathogens. Plant Cell,
13, 1025–1033.
Endler, A., and Persson, S. (2011) Cellulose synthases and synthesis in Arabidopsis.
Mol. Plant, 4, 199–211.
Endo, S., Pesquet, E., Yamaguchi, M., Tashiro, G., Sato, M., Toyooka, K.,
Nishikubo, N., Udagawa–Motose, M., Kubo, M., Fukuda, H., and Demura, T.
(2009) Identifying new components participating in the secondary cell wall formation of
vessel elements in Zinnia and Arabidopsis. Plant Cell, 21, 1155–1165.
Enns, L.C., Kanaoka, M.M., Torii, K.U., Comai, L., Okada, K., and Cleland, R.E.
(2005) Two callose synthases, GSL1 and GSL5, play an essential and redundant role in plant
and pollen development and in fertility. Plant Mol. Biol., 58, 333–349.
Fagard, M., Desnosm, T., Desprez, T., Goubet, F., Refregier, G., Mouille, G.,
McCann, M., Rayon, C., Vernhettes, S., and Höfte, H. (2000) PROCUSTE1 encodes
a cellulose synthase required for normal cell elongation specifically in roots and dark–grown
hypocotyls of Arabidopsis. Plant Cell, 12, 2409–2423.
Faik, A., Price, N.J., Raikhel, N.V., and Keegstra, K. (2002) An Arabidopsis gene
encoding an α–xylosyltransferase involved in xyloglucan biosynthesis. Proc. Natl. Acad. Sci.
USA, 99, 7797–7802.
Ferguson, C., Teeri, T.T., Siika–aho, M., Read, S.M., and Bacic, A. (1998) Location
of cellulose and callose in pollen tubes and grains of Nicotiana tabacum. Planta, 206, 452–
460.
Fernandez-Calvino, L., Faulkner, C., Walshaw, J., Saalbach, G., Bayer, E.,
Benitez-Alfonso, Y., and Maule, A. (2011) Arabidopsis plasmodesmal proteome. PLoS
ONE 6:e18880
Francisco, J.A., Stathopoulos, C., Warren, R. A., Kilburn, D.G., and Georgiou,
G. (1993) Specific adhesion and hydrolysis of cellulose by intact Escherichia coli expressing
surface anchored cellulase or cellulose binding domains. Bio/Technology, 11, 491–495.
Franz, G., and Blaschek, W. (1990) Cellulose. In: Methods in plant biochemistry.
Academic Press, London, vol 2, 291–322.
Fredrikson, K., and Larsson, C. (1992) Activators and inhibitors of the plant plasma
membrane 1,3–β–glucan synthase. Biochem. Soc. Trans, 20, 710–713.
64
Fujisawa, F., Saito, T., Kimura, S., Iwata, T., and Isogai, A. (2013) Surface
engineering of ultrafine cellulose nanofibrils toward polymer nanocomposite materials.
Biomacromolecules, 14, 1541–1546.
Georgelis, N., Yennawar, N.H., and Cosgrove, D.J. (2012) Structural basis for
entropy–driven cellulose binding by a type–A cellulose–binding module (CBM) and
bacterial expansin. Proc. Natl. Acad. Sci. USA, 109, 14830–14835.
Gerngross, U.T., Romaniec, M.P., Kobayashi, T., Huskisson, N.S., and Demain,
A.L. (1993) Sequencing of a Clostridium thermocellum gene (cipA) encoding the
cellulosomal SL–protein reveals an unusual degree of internal homology. Mol. Microbiol., 8,
325–334.
Gidley, M.J., Lillford, P.J.D., Rowlands, W., Lang, P., Dentini, M., Crescenzi,
V., Edwards, M., Fanutti, C., and Reid, J.S.G. (1991) Structure and solution
properties of tamarind–seed polysaccharide. Carbohydr. Res., 214, 299–314.
Gilbert, H.J., Knox, J.P., and Boraston, A.B. (2013) Advances in understanding the
molecular basis of plant cell wall polysaccharide recognition by carbohydrate–binding
modules. Curr. Opin. Struct. Biol., 23, 669–677.
Goldstein, M.A., Takagi, M., Hashida, S., Shoseyov, O., Doi, R.H., and Segel,
I.H. (1993) Characterization of the cellulose–binding domain of the Clostridium
cellulovorans cellulose–binding protein A. J. Bacteriol., 175, 5762–5768.
Gomez, L.D., Whitehead, C., Barakate, A., Halpin, C., and McQueen-Mason,
S.J. (2010) Automated saccharification assay for determination of digestibility in plant
materials. Biotechnol. Biofuels, 3, 23.
Gonneau, M., Desprez, T., Guillot, A., Vernhettes, S., and Höfte, H. (2014)
Catalytic subunit stoichiometry within the cellulose synthase complex. Plant Physiol., 166,
1709–1712.
Goodin, M.M., Zaitlin, D, Naidu, R.A., and Lommel, S.A (2008) Nicotiana
benthamiana: Its history and future as a model for plant–pathogen interactions. Mol. Plant
Microbe Interact., 21, 1015–1026.
Gosselin, S., Alhussaini, M., Streiff, M.B., Takabayashi, K., and Palcic, M.M.
(1994) A continuous spectrophotometric assay for glycosyltransferases. Anal. Biochem.,
220, 92–97.
Gourlay, K., Arantes, V., and Saddler, J.N. (2012) Use of substructure–specific
carbohydrate binding modulesto track changes in cellulose accessibility and surface
morphology during the amorphogenesis step of enzymatic hydrolysis. Biotechnol. Biofuels,
5, 51.
Grabski, S., Defeijter, A.W., and Schindler, M. (1993) Endoplasmic–reticulum
Forms a dynamic continuum for lipid diffusion between contiguous soybean root–cells.
Plant Cell, 5, 25–38.
Grison, M.S., Brocard, L., Fouillen, L., Nicolas, W., Wewer, V., Dörmann, P.,
Nacir, H., Benitez–Alfonso, Y., Claverol, S., Germain, V., Boutté, Y., Mongrand,
65
S., and Bayer, E.M. (2015) Specific membrane lipid composition is important for
plasmodesmata function in Arabidopsis. Plant Cell, 27, 1228–1250.
Gu, Y., Kaplinsky, N., Bringmann, M., Cobb, A., Carroll, A., Sampathkumar, A.,
Baskin, T.I., Persson, S., and Somerville, C.R. (2010) Identification of a cellulose
synthase–associated protein required for cellulose biosynthesis. Proc. Natl. Acad. Sci. USA,
107, 12866–12871.
Guenoune–Gelbart, D., Elbaum, M., Sagi, G., Levy, A., and Epel, BL. (2008)
Tobacco mosaic virus (TMV) replicase and movement protein function synergistically in
facilitating TMV spread by lateral diffusion in the plasmodesmal desmotubule of Nicotiana
benthamiana. Mol. Plant Microbe. Interact., 21, 335–345.
Guerriero, G., Fugelstad, J., and Bulone, V. (2010) What do we really know about
cellulose biosynthesis in higher plants? J. Integr. Plant Biol., 52, 161–175.
Guseman, J.M., Lee, J.S., Bogenschutz, N.L., Peterson, K.M., Virata, R.E., Xie,
B., Kanaoka, M.M., Hong, Z., and Torii, K.U. (2010) Dysregulation of cell–to
cellconnectivity and stomatal patterning by loss–of–function mutation in Arabidopsis
CHORUS (GLUCAN SYNTHASE–LIKE 8). Development, 137, 1731–1741.
Gustavsson, M.T., Persson, P.V., Iversen, T., Hult, K., and Martinelle, M. (2004)
Polyester coating of cellulose fiber surfaces catalyzed by a cellulose–binding module–
Candida antarctica lipase B fusion protein. Biomacromolecules, 5, 106–112.
Haigler, C.H., White, A.R., Brown, R.M., and Cooper, K.M. (1982) Alteration of in
vivo cellulose ribbon assembly by carboxymethylcellulose and other cellulose derivatives. J.
Cell Biol., 94, 64–69.
Han, X., Hyun, T., Zhang, M., Kumar, R., Koh, E.J., Kang, B.H., Lucas, W.J.,
and Kim, J.Y. (2014) Auxin–callose–mediated plasmodesmal gating is essential for tropic
auxin gradient formtion and signaling. Dev. Cell, 28, 132–146.
Hayashi, T., Read, S., Bussell, J., Thelen, M.P., Lin, F.–C., Brown, R.M.J., and
Delmer, D. (1987) UDP–glucose: (1,3)–beta–glucan synthases from mung bean and
cotton. Plant Physiol., 83, 1054–1062.
Hayashi, T. (1989) Xyloglucans in the primary cell wall. Annu. Rev. Plant Physiol. Plant
Mol. Biol., 40, 139–168.
Helbert, W., Sugiyama, J., Ishihara, M., and Yamanaka, S. (1997) Characterization
of native crystalline cellulose in the cell walls of Oomycota. J. Biotechnol, 57, 29-37.
Henshaw, J.L., Bolam, D.N., Pires, V.M., Czjzek, M., Henrissat, B., Ferreira,
L.M., Fontes, C.M., and Gilbert, H.J. (2004) The family 6 carbohydrate binding
module CmCBM6–2 contains two ligand–binding sites with distinct specificities. J. Biol.
Chem., 279, 21552–21559.
Hernandez–Gomez, M.C., Rydahl, M.G., Rogowski, A., Morland, C., Cartmell,
A., Crouch, L., Labourel, A., Fontes, C.M., Willats, W.G., Gilbert, H.J., and
Knox, J.P. (2015) Recognition of xyloglucan by the crystalline cellulose–binding site of a
family 3a carbohydrate–binding module. FEBS Lett., 589, 2297–2303.
66
Hervé, C., Rogowski, A., Blake, A.W., Marcus, S.E., Gilbert, H.J., and Knox, J.P.
(2010) Carbohydrate–binding modules promote the enzymatic deconstruction of intact
plant cell walls by targeting and proximity effects. Proc. Natl. Acad. Sci. USA, 4, 15293–
15298.
Hieta, K., Kuga, S., and Usuda, M. (1984) Electron staining of reducing ends evidences
a parallel‐chain structure in Valonia cellulose. Biopolymers, 23, 1807–1810.
Hill, J.L., Hammudi, M.B., and Tien, M. (2014) The Arabidopsis cellulose synthase
complex: a proposed hexamer of CESA trimers in an equimolar stoichiometry. Plant Cell,
26, 4834–4842.
Hoffmann, G.C., and Timell, T.E. (1970) Isolation of a β 1–3 glucan (laticinan) from
compression wood of Larix laricinia. Wood Sci. Technol., 4, 159–162.
Hoffmann, G.C., and Timell, T.E. (1972) Polysaccharides in compression wood of
tamarack (Larix laricina). 1. Isolation and characterization of laricinan, an acidic glucan.
Svensk Papperstidning, 75, 135–141.
Hoffman, M., Jia, Z.H., Peña, M.J., Cash, M., Harper, A., Blackburn, A.R.,
Darvill, A., and York, W.S. (2005) Structural analysis of xyloglucans in the primary cell
walls of plants in the subclass Asteridae. Carb. Res., 340, 1826–1840.
Holdaway–Clarke, T.L., Walker, N.A., Hepler, P.K., and Overall, R.L. (2000)
Physiological elevations in cytoplasmic free calcium by cold or ion injection result in
transient closure of higher plant plasmodesmata. Planta, 210, 329–335.
Hong, Z., Zhang, Z., Olson, J.M., and Verma, D.P. (2001a) A novel UDP–glucose
transferase is part of the callose synthase complex and interacts with phragmoplastin at the
forming cell plate. Plant Cell, 13, 769–779.
Hong, Z., Delauney, A.J., Verma, D.P., Park, S., Baker, J.O., Himmel, M.E.,
Parilla, P., and Johnson, D.K. (2001b) A cell plate–specific callose synthase and its
interaction with phragmoplastin. Plant Cell, 13, 755–768.
Horn. S.J., Vaaje–Kolstad, G., Westereng, B., and Eijsink, V.G. (2012) Novel
enzymes for the degradation of cellulose. Biotechnol. Biofuels, 5, 45.
Hsieh, Y.S.Y., and Harris, P.J. (2009) Xyloglucans of monocotyledons have diverse
structures. Mol. Plant, 2, 943‐965.
Hu, Y., and Walker, S. (2002) Remarkable structural similarities between diverse
glycosyltransferases. Chem. Biol., 9, 1287–1296.
Huwyler, H.R., Franz, G., and Meier, H. (1978) β–1,3–Glucans in the cell walls of
cotton fibres (Gossypium arboreum L.). Plant Sci. Lett., 12, 55–62.
Ifuku, S., Nogi, M., Abe, K., Handa, K., Nakatsubo, F., and Yano, H. (2007)
Surface modification of bacterial cellulose nanofibers for property enhancement of optically
transparent composites: Dependence on acetyl–group DS. Biomacromolecules, 8, 1973–
1978.
67
Ishii, T., Matsunaga, T., Pellerin, P., O'Neill, M.A., Darvill, A., and Albersheim,
P. (1999) The plant cell wall polysaccharide rhamnogalacturonan II self–assembles into a
covalently cross–linked dimmer. J. Biol. Chem., 274, 13098–13104.
Ishii, T., Ichita, J., Matsue, H., Ono, H. and Maeda, I. (2002) Fluorescent labeling of
pectic oligosaccharides with 2–aminobenzamide and enzyme assay for pectin. Carbohydr.
Res., 337, 1023–1032
Ishii, T., Ohnishi–Kameyama, M. and Ono, H. (2004) Identification of elongating β–
1,4–galactosyltransferase activity in mung bean (Vigna radiata) hypocotyls using 2–
aminobenzaminated 1,4–linked β–d–galactooligosaccharides as acceptor substrates. Planta,
219, 310–318.
Jacobs, A.K., Lipka, V., Burton, R.A., Panstruga, R., Strizhov, N., Schulze–
Lefert, P., and Fincher, G.B. (2003) An Arabidopsis callose synthase, GSL5, is required
for wound and papillary callose formation. Plant Cell, 15, 2503–2513.
Jansson, S., and Douglas, C.J. (2007) Populus: a model system for plant biology. Annu.
Rev. Plant Biol., 58, 435–458.
Jarvis, M. (2003) Chemistry: cellulose stacks up. Nature, 426, 611–612.
John, M., Röhrig, H., Schmidt, J., Walden, R., and Schell, J. (1997) Cell signalling
by oligosaccharides. Trends Plant Sci., 2, 111–115.
Johnson, P.E., Tomme, P., Joshi, M.D., and McIntosh, L.P. (1996) Interaction of
soluble cellooligosaccharides with the N–terminal cellulose–binding domain of
Cellulomonas fimi CenC. 2. NMR and ultraviolet absorption spectroscopy. Biochemistry,
35, 13895–13906.
Kaida, R., Satoh, Y., Bulone, V., Yamada, Y., Kaku, T., Hayashi, T., and Kaneko,
T.S. (2009) Activation of β-Glucan synthases by wall-bound purple acid phosphatase in
tobacco cells. Plant Physiol. 150, 1822–1830.
Karnezis, T., Epa, V.C., Stone, B.A., and Stanisich, V.A. (2003) Topological
characterization of an inner membrane (1→3)–beta–D–glucan (curdlan) synthase from
Agrobacterium sp. strain ATCC31749. Glycobiology, 13, 693–706.
Kauss, H. (1985) Callose biosynthesis as a Ca2+–regulated process and possible relations to
the induction of other metabolic changes. J. Cell Sci., 2, 89–103.
Kavoosi, M., Meijer, J., Kwan, E., Creagh, A. L., Kilburn ,D. G., and Haynes, C.
A. (2004) Inexpensive one–step purification of polypeptides expressed in Escherichia coli
as fusions with the family 9 carbohydrate–binding module of xylanase 10A from T.
maritima. J. Chromatogr. B Analyt. Technol. Biomed. Life Sci., 807, 87–94.
Kawano, S., Tajima, K., Kono, H., Erata, T., Munekata, M., and Taka, M. (2002)
Effects of endogenous endo–beta–1,4–glucanase on cellulose biosynthesis in Acetobacter
xylinum ATCC23769. J. Biosci. Bioeng., 94, 275–281.
Kerstens, S., Decraemer, W.F., and Verbelen, J-P. (2001) Cell walls at the plant surface behave mechanically like fiber-reinforced composite materials. Plant Physiol., 127, 381–385.
68
Kim, I., Cho, E., Crawford, K., Hempel, F.D., and Zambryski, P.C. (2005) Cell–
to–cell movement of GFP during embryogenesis and early seedling development in
Arabidopsis. Proc. Natl. Acad. Sci. USA, 102, 2227–2231.
Kimura, S., Laosinchai, W., Itoh, T., Cui, X., Linder, C., and Brown, R. (1999)
Immunogold labeling of rosette terminal cellulose–synthesizing complexes in the vascular
plant Vigna angularis. Plant Cell, 11, 2075–2086.
Kitaoka, T., and Tanaka, H. (2001) Novel paper strength additive containing cellulose–
binding domain of cellulase. J. Wood Sci., 47, 322–324.
Kjell, J., Rasmusson, A.G., Larsson, H., and Widell, S. (2004) Protein complexes of
the plant plasma membrane resolved by Blue Native PAGE. Physiol. Plant, 121, 546–555.
Klemm, D., Heublein, B., Fink, H.P., and Bohn, A. (2005) Cellulose: fascinating
biopolymer and sustainable raw material. Angew. Chem. Int. Ed. Engl., 44, 3358–3393.
Kljun, A., Benians, T.A.S., Goubet, F., Meulewaeter, F., Knox, J.P., and
Blackburn, R.S. (2011) Comparative analysis of crystallinity changes in cellulose I
polymers using ATR–FTIR, X–ray diffraction, and carbohydrate–binding module probes.
Biomacromolecules, 12, 4121–4126.
Kohle, H., Jeblick, W., Poten, F., Blaschek, W., and Kauss, H. (1985) Chitosan–
elicited callose synthesis in soybean cells as a Ca2+–dependent process. Plant Physiol., 77,
544–551.
Kolpak, K.J., and Blackwell, J. (1976) Determination of the structure of cellulose II.
Macromolecules, 9, 273–278.
Konishi, T., Ono, H., Ohnishi–Kameyama, M., Kaneko, S. and Ishii, T. (2006)
Identification of a mung bean arabinofuranosyltransferase that transfers arabinofuranosyl
residues onto (1,5)–linked α–l–arabino–oligosaccharides. Plant Physiol., 141, 1098–1105.
Koornneef, M., and Meinke, D. (2010) The development of Arabidopsis as a model
plant. Plant J., 61, 909-921
Kopp, M., Rupprath, C., Irschik, H., Bechthold, A., Elling, L., and Müller, R.
(2007) SorF: A glycosyltransferase with promiscuous donor substrate specificity in vitro.
ChemBioChem, 8, 813-819.
Koyama, M., Helbert, W., Imai, T., Sugiyama, J., and Henrissat, B. (1997)
Parallel–up structure evidences the molecular directionality during biosynthesis of bacterial
cellulose. Proc. Natl. Acad. Sci. USA, 94, 9091–9095.
Kruus, K., Lua, A.C., Demain, A.L., and Wu, J.H. (1995) The anchorage function of
CipA (CelL), a scaffolding protein of the Clostridium thermocellum cellulosome. Proc. Natl.
Acad. Sci. USA, 92, 9254–9258.
Kudlicka, K., Lee, J.H., and Brown, R.M. (1996) A comparative analysis of in vitro
cellulose synthesis from cell–free extracts of mung bean (Vigna radiata, Fabaceae) and
cotton (Gossypium hirsutum, Malvaceae). Am. J. Bot., 83, 274–284.
69
Kudlicka, K., and Brown, R.B. (1997) Cellulose and callose biosynthesis in higher plants
(I. Solubilization and separation of (1–>3)–and (1–>4)–β–glucan synthase activities from
mung bean). Plant Physiol., 115, 643–656.
Kuga, S., Takagi, S., and Brown, R.M. (1993) Native folded–chain cellulose II.
Polymer, 34, 3293–3297.
Kumar, M., Campbell, L. and Turner, S. (2016) Secondary cell walls: biosynthesis and
manipulation. J. Exp. Bot., 67, 515–531.
Kurek, I., Kawagoe, Y., Jacob–Wilk, D., Doblin, M., and Delmer, D. (2002)
Dimerization of cotton fiber cellulose synthase catalytic subunits occurs via oxidation of the
zinc–binding domains. Proc. Natl. Acad. Sci. USA, 99, 11109–11114.
Lagaert, S., Belien, T., and Volckaert, G. (2009) Plant cell walls: Protecting the barrier
from degradation by microbial enzymes. Semin. Cell Dev. Biol., 20, 1064–1073.
Lai Kee Him, J., Pelosi, L., Chanzy, H., Putaux, J.L., and Bulone, V. (2001)
Biosynthesis of (1–>3)–β–D-glucan (callose) by detergent extracts of a microsomal fraction
from Arabidopsis thaliana. Eur. J. Biochem, 268, 4628–4638.
Lai Kee Him, J., Chanzy, H., Müller, M., Putaux, J.L., Imai, T., and Bulone, V.
(2002) In vitro versus in vivo cellulose microfibrils from plant primary wall synthases:
structural differences. J. Biol. Chem., 277, 36931–36939.
Lairson, L.L., Henrissat, B., Davies, G.J., and Withers, S.G. (2008)
Glycosyltransferases: structures, functions, and mechanisms. Annu. Rev. Biochem., 77,
521–555.
Laloi, M., Perret, A–M., Chatre, L., Melser, S., Cantrel, C., Vaultier, M–N.,
Zachowski, A., Bathany, K., Schmitter, J–M., Vallet, M., Lessire, R., Hartmann,
M–A., and Moreau, P. (2007) Insights into the role of specific lipids in the formation and
delivery of lipid microdomains to the plasma membrane of plant cells. Plant Physiol., 143,
461–472.
Langan, P., Nishiyama, Y., and, Chanzy, H. (2001) X–ray structure of mercerized
cellulose II at 1 Å resolution. Biomacromolecules, 2, 410–416.
Lee, J–Y., Wang, X., Cui, W., Sager, R., Modla, S. Czymmek, K., Zybaliov, B.,
van Wijk, K., Zhang, C., Lu, H., and Lakshmanan, V. (2011) A plasmodesmata–
localized protein mediates crosstalk between cell–to–cell communication and innate
immunity in Arabidopsis. Plant Cell, 23, 3353–3373.
Lee, C., Zhong, R., and Ye, Z.–H. (2012) Arabidopsis family GT43 members are xylan
xylosyltransferases required for the elongation of the xylan backbone. Plant Cell Physiol.,
53, 135–143.
Lehtio, J., Wernerus, H., Samuelson, P., Teeri, T. T., and Stahl, S. (2001) Directed
immobilization of recombinant staphylococci on cotton fibers by functional display of a
fungal cellulose–binding domain. FEMS Microbiol. Lett., 195, 197–204.
Leijon, F., Melzer, M., Zhou, Q., Srivastava, V., and Bulone, V. (2018). Proteomic
analysis of plasmodesmata from Populus cell suspension cultures in relation with callose
biosynthesis. Front. Plant Sci., 9, 1681.
70
Levy, I., Nussinovitch, A., Shpigel, E., and Shoseyov, O. (2002) Recombinant
cellulose crosslinking protein: a novel paper–modification biomaterial. Cellulose, 9, 91–98.
Levy, A., Erlanger, M., Rosenthal, M., and Epel, B.L. (2007) A plasmodesmata–
associated β–1,3–glucanase in Arabidopsis. Plant J., 49, 669–682.
Li, H., Bacic, A., and Read, S.M. (1997) Activation of pollen tube callose synthase by
detergents. Evidence for different mechanisms of action. Plant Physiol., 114, 1255–1265.
Li, H., Lin, Y.K., Heath, R.M., Zhu, M.X., and Yang, Z.B. (1999) Control of pollen
tube tip growth by a Rop GTPase–dependent pathway that leads to tip–localized calcium in
flux. Plant Cell, 11, 1731–1742.
Li, J., Burton, R.A., Harvey, A.J., Hrmova, M., Wardak, A.Z., Stone, B.A., and
Fincher, G.B. (2003) Biochemical evidence linking a putative callose synthase gene with (1
–>3)–β–D–glucan biosynthesis in barley. Plant Mol. Biol., 53, 213–225.
Li, X., Cordero, I., Caplan, J., Molhoj, M., and Reiter, W.D. (2004) Molecular
analysis of 10 coding regions from Arabidopsis that are homologous to the MUR3
xyloglucan galactosyltransferase. Plant Physiol., 134, 940–950.
Lichtenberg, D., Goñi, F.M., and Heerklotz, H. (2005) Detergent–resistant
membranes should not be identified with membrane rafts. Trends Biochem. Sci., 30, 430–
436.
Liepman, A.H., Wilkerson, C.G., and Keegstra, K. (2005) Expression of cellulose
synthase–like (Csl) genes in insect cells reveals that CslA family members encode mannan
synthases. Proc. Natl. Acad. Sci. USA, 102, 2221–2226.
Limon, M.C., Margolles–Clark, E., Benitez, T., and Penttila, M. (2001) Addition of
substrate–binding domains increases substrate–binding capacity and specific activity of a
chitinase from Trichoderma harzianum. FEMS Microbiol. Lett., 198, 57–63.
Linder, M., Mattinen, M., Kontteli, M., Lindeberg, G., Ståhlberg, J.,
Drakenberg, T., Reinikainen, T., Pettersson, G., and Annila, A. (1995)
Identification of functionally important amino acids in the cellulose‐binding domain of
Trichoderma reesei cellobiohydrolase I. Protein Sci., 4, 1056–1064.
Littunen, K., Hippi, U., Johansson, L. S., Österberg, M., Tammelin, T., Laine,
J., and Seppala, J. (2011) Free radical graft copolymerization of nanofibrillated cellulose
with acrylic monomers. Carbohyd. Polym., 84, 1039–1047.
Ljungberg, N., Bonini, C., Bortolussi, F., Boisson, C., Heux, L., and Cavaillé,
J.Y. (2005) New nanocomposite materials reinforced with cellulose whiskers in atactic
polypropylene: effect of surface and dispersion characteristics. Biomacromolecules, 6,
2732–2739.
Lombard, V., Ramulu, H.G., Drula, E., Coutinho, P.M., and Henrissat, B. (2014)
The carbohydrate–active enzymesdatabase (CAZy) in 2013. Nucleic Acids Res., 42, 490–
495.
London, E., and Brown, D.A. (2000) Insolubility of lipids in triton X–100: physical
origin and relationship to sphingolipid/cholesterol membrane domains (rafts). Biochim.
Biophys. Acta., 1508, 182–195.
71
Madson, M., Dunand, C., Li, X.M., Verma, R., Vanzin, G.F., Calplan, J., Shoue,
D.A., Carpita, N.C., and Reiter, W.D. (2003) The MUR3 gene of Arabidopsis encodes a
xyloglucan galactosyltransferase that is evolutionarily related to animal exostosins. Plant
Cell, 15, 1662–1670.
Maltby, D., Carpita, N.C., Montezinos, D., Kulow, C., and Delmer, D.P. (1979) β–
1,3–Glucan in developing cotton fibers. Structure, localization and relationship of synthesis
to that of secondary wall cellulose. Plant Physiol., 63, 1158–1164.
Marchessault, R.H., Morehead, F.F., and Joan Koch, M. (1961) Some
hydrodynamic properties of neutral suspensions of cellulose crystallites as related to size
and shape. Journal of colloid science, 16, 327–344.
Marchessault, D.H., Deslandes, H., Ogawa, K., and Sundarajan, P.R. (1977) X–
ray diffraction data for β–(1 - 3)–D–glucan. Can. J. Chem., 55, 300–303.
Marga, F., Grandbois, M., Cosgrove, D.J., and Baskin, T.I. (2005) Cell wall
extension results in the coordinate separation of parallel microfibrils: evidence from
scanning electron microscopy and atomic force microscopy. Plant J., 43, 181–190.
Maule, A., Faulkner, C., and Benitez–Alfonso, Y. (2012) Plasmodesmata “in
Communicado”. Front. Plant Sci., 3, 30.
Mayo, K.J, Gonzales B.J., and Mason, H.S. (2006) Genetic transformation of tobacco
NT1 cells with Agrobacterium tumefaciens. Nat. Protoc., 3, 1105–1111.
McCartney, L., Gilbert, H.J., Bolam, D.N., Boraston, A.B., and Knox, J.P. (2004)
Glycoside hydrolase carbohydrate–binding modules as molecular probes for the analysis of
plant cell wall polymers. Anal. Biochem., 326, 49–54.
McNamara, J.T., Morgan, J.L., and Zimmer, J. (2015) A molecular description of
cellulose biosynthesis. Annu. Rev. Biochem., 84, 895–921.
Meilan, R., and Ma, C. (2006) Poplar (Populus spp.). In: Wang K. (eds) Agrobacterium
Protocols Volume 2. Methods in Molecular Biology, vol 344. Humana Press.
Miedes, E., Zarra, I., Hoson, T., Herbers, K., Sonnewald, U., and Lorences, E.P.
(2011) Xyloglucan endotransglucosylase and cell wall extensibility. J. Plant Physiol., 168,
196–203.
Mongrand, S., Morel, J., and Laroche, J. (2004) Lipid rafts in higher plant cells:
purification and characterization of triton X–100–insoluble microdomains from tobacco
plasma membrane. J. Biol. Chem., 279, 36277–36286.
Moon, R. J., Martini, A., Nairn, J., Simonsen, J., and Youngblood, J. (2011)
Cellulose nanomaterials review: structure, properties and nanocomposites. Chem. Soc. Rev.,
40, 3941–3994.
Morag, E., Lapidot, A., Govorko, D., Lamed, R., Wilchek, M., Bayer, E. A., and
Shoham, Y. (1995) Expression, purification, and characterization of the cellulose–binding
domain of the scaffoldin subunit from the cellulosome of Clostridium thermocellum. Appl.
Environ. Microbiol., 61, 1980–1986.
72
Morgan, J.L.W., Strumillo, J., and Zimmer, J. (2013) Crystallographic snapshot of
cellulose synthesis and membrane translocation. Nature, 493, 181–186.
Morris, E.R., Powell, D.A., Gidley, M.J. and Rees, D.A. (1982) Conformations and
interactions of pectins: I. Polymorphism between gel and solid states of calcium
polygalacturonate. J. Mol. Biol., 155, 507-516.
Mueller, S.C., and Brown, R.M., (1980) Evidence for an intramembranous component
associated with a cellulose microfibril synthesizing complex in higher plants. J. Cell Biol.,
84, 315–326.
Mustafa, N.R., Ward de Winter, W., van Iren, F., and Verpoorte, R. (2011)
Initiation, growth and cryopreservation of plant cell suspension cultures. Nat. Protoc., 6,
715–742.
Nagata, T., Nemoto, Y., and Hasezawa, S. (1992) Tobacco BY–2 cell line as the “HeLa”
cell in the cell biology of higher plants. Int. Rev. Cytol., 132, 1–30.
Nardi, C.F., Villarreal, N.M., Rossi, F.R., Martínez, S., Martínez, G.A., and
Civello, P.M. (2015) Overexpression of the carbohydrate binding module of strawberry
expansin2 in Arabidopsis thaliana modifies plant growth and cell wall metabolism. Plant
Mol. Biol., 88, 101–117.
Newman, R.H. (2008) Simulation of X-ray diffractograms relevant to the purported
polymorphs cellulose IVI and IVII. Cellulose, 15, 769–778.
Newman, R.H., Hill, S.J., and Harris, P.J. (2013) Wide–angle X–ray scattering and
solid–state nuclear magnetic resonance data combined to test models for cellulose
microfibrils in mung bean cell walls. Plant Physiol., 163, 1558–1567.
Nicol, F., His, I., Jauneau, A., Vernhettes, S., Canut, H., and Hofte, H. (1998) A
plasma membrane–bound putative endo–1,4–β–D–glucanase is required for normal wall
assembly and cell elongation in Arabidopsis. EMBO J., 17, 5563–5576.
Nicolas, W.J., Grison, M.S., Trepout, S., Gaston, A., Fouche, M., Cordelieres,
F.P., Oparka, K., Tilsner, J., Brocard, L., and Bayer, E.M. (2017) Architecture and
permeability of post–cytokinesis plasmodesmata lacking cytoplasmic sleeves. Nat. Plants, 3,
17082.
Niimura, H., Yokoyama, T., Kimura, S., Matsumoto, Y., and Kuga, S. (2010) AFM
observation of ultrathin microfibrils in fruit tissues. Cellulose, 17, 13–18.
Nishikawa, S.I., Zinkl, G.M., Swanson, R.J., Maruyama, D., and Preuss, D.J.
(2005) Callose (β–1,3 glucan) is essential for Arabidopsis pollen wall patterning, but not
tube growth. BMC Plant Biol., 5, 22.
Nishimura, M.T., Stein, M., Hou, B.H., Vogel, J.P., Edwards, H., and
Somerville, S. C. (2003) Loss of a callose synthase results in salicylic acid–dependent
disease resistance. Science, 301, 969–972.
Nishitani, K. (1998) Construction and restructuring of the cellulose‐xyloglucan framework
in the apoplast as mediated by the xyloglucan related protein family ‐ A hypothetical
scheme. J. Plant Res., 111, 159–166.
73
Nishiyama, Y., Langan, P., and Chanzy, H. (2002) Crystal structure and hydrogen
bonding system in cellulose Iβ from synchrotron X-ray and neutron fiber diffraction. J. Am.
Chem. Soc., 124, 9074–9082.
Nishiyama, Y., Sugiyama, J., Chanzy, H., and Langan, P. (2003) Crystal structure and hydrogen bonding system in cellulose Iα from synchrotron X-ray and neutron fiber diffraction. J. Am. Chem. Soc., 125, 14300–14306.
Nishiyama, Y. (2009) Structure and properties of the cellulose microfibril. J. Wood Sci.,
55, 241–249.
Nixon, B.T., Mansouri, K., Singh, A., Du, J., Davis, J.K., Lee, J.G., Slabaugh, E.,
Vandavasi, V.G., O’Neill, H., Roberts, E.M., Roberts, A.W., Yingling, Y.G., and
Haigler, C.H. (2016) Comparative structural and computational analysis supports
eighteen cellulose synthases in the plant cellulose synthesis complex. Sci. Rep., 6, 28696.
Notenboom, V., Boraston, A.B., Williams, S.J., Kilburn, D.G., and Rose, D.R.
(2002) High–resolution crystal structures of the lectin–like xylan binding domain from
Streptomyces lividans xylanase 10A with bound substrates reveal a novel mode of xylan
binding. Biochemistry, 41, 4246–4254.
Obembe, O.O., Jacobsen, E., Timmers, J., Gilbert, H., Blake, A.W., Knox, J.P.,
Visser, R.G.F., and Vincken, J.P. (2007) Promiscuous, non–catalytic, tandem
carbohydrate–binding modules modulate the cell–wall structure and development of
transgenic tobacco (Nicotiana tabacum) plants. J. Plant Res., 120, 605–617.
Oide, S., Bejai, S., Staal, J., Guan, N., Kaliff, M., and Dixelius, C. (2013) A novel
role of PR2 in abscisic acid (ABA) mediated, pathogen–induced callose deposition in
Arabidopsis thaliana. New Phytol., 200, 1187–1199.
Okuda, K., Li, L., Kudlicka, K., Kuga, S., and Brown, R.M. Jr. (1993) β–Glucan
synthesis in the cotton fiber. I. Identification of β–1,4– and β–1,3–glucans synthesized in
vitro. Plant Physiol., 101, 1131–1142.
Okuda, K., Tsekos, L., and Brown, R.M. (1994) Cellulose microfibril assembly in
Erythrocladia subintegra Rosenv.: An ideal system for understanding the relationship
between synthesizing complexes (TCs) and microfibril crystallization. Protoplasma, 180,
49–58.
Oliveira, C., Carvalho, V., Domingues, L., and Gama, F.M. (2015) Recombinant
CBM–Fusion Technology — Applications Overview. Biotechnol. Adv., 33, 358–369.
Ostergaard, L., Petersen, M., Mattsson, O., and Mundy, J. (2002) An Arabidopsis
callose synthase. Plant Mol. Biol., 49, 559–566.
Overall, R.L., Wolfe, J. and Gunning, B.E.S. (1982) Intercellular communication in
Azolla roots. I. Ultrastructure of plasmodesmata. Protoplasma, 111, 134–150.
Overall, R.L., and Blackman, L.M. (1996) A model of the macromolecular structure of
plasmodesmata. Trends Plant Sci., 1, 307–311.
Owen, H.A., and Makaroff, C.A. (1995) Ultrastructure of microsporogenesis and
microgametogenesis in Arabidopsis thaliana (L.) Heynh. ecotype Wassilewskija
(Brassicaceae) Protoplasma, 185, 7–21.
74
Paakko, M., Ankerfors, M., Kosonen, H., Nykanen, A., Ahola, S., Österberg, M.,
Ruokolainen, J., Laine, J., Larsson, P.T., Ikkala, O., and Lindström, T. (2007)
Enzymatic hydrolysis combined with mechanical shearing and high–pressure
homogenization for nanoscale cellulose fibrils and strong gels. Biomacromolecules, 8,
1934–1941.
Pagant, S., Bichet, A., Sugimoto, K., Lerouxel, O., Desprez, T., McCann, M.,
Lerouge, P., Vernhettes, S., and Höfte, H. (2002) KOBITO1 encodes a novel plasma
membrane protein necessary for normal synthesis of cellulose during cell expansion in
Arabidopsis. Plant Cell, 14, 2001–2013.
Park, Y.W., Tominaga, R., Sugiyama, J., Furuta, Y., Tanimoto, E., Samejima,
M., Sakai, F., and Hayashi, T. (2003) Enhancement of growth by expression of poplar
cellulase in Arabidopsis thaliana. Plant J., 33, 1099–1106.
Park, Y.B., and Cosgrove, D.J. (2012a) Changes in cell wall biomechanical properties in
the xyloglucan‐ deficient xxt1/xxt2 mutant of Arabidopsis. Plant Physiol., 158, 465–475.
Park, Y.B., and Cosgrove, D.J. (2012b) A revised architecture of primary cell walls
based on biomechanical changes induced by substrate-specific endoglucanases. Plant
Physiol., 158, 1933–1943.
Park, Y.B., and Cosgrove, D.J. (2015) Xyloglucan and its interactions with other
components of the growing cell wall. Plant Cell Physiol., 56, 180–194.
Parre, E., and Geitmann, A. (2005) More than a leak sealant: the mechanical properties
of callose in growing plant cells. Plant Physiol., 137, 274–286.
Pelosi, L., Imai, T., Chanzy, H., Heux, L., Buhler, E., and Bulone, V. (2003)
Structural and morphological diversity of (1→3)–β– D–glucans synthesized in vitro by
enzymes from Saprolegnia monoica. Comparison with a corresponding in vitro product
from blackberry (Rubus fruticosus). Biochemistry, 42, 6264–6274.
Pelosi, L., Bulone, V., and Heux, L. (2006) Polymorphism of curdlan and (1→3)–β–d–
glucans synthesized in vitro. A 13C CP–MAS and x–ray diffraction analysis. Carbohydr.
Polym., 66, 199–207.
Peña, M.J., Ryden, P., Madson, M., Smith, A.C., and Carpita, N.C. (2004) The
galactose residues of xyloglucan are essential to maintain mechanical strength of the
primary cell walls in Arabidopsis during growth. Plant Physiol., 134, 443‐451.
Peña, M.J., Darvill, A.G., Eberhard, S., York, W.S., and O'Neill, M.A. (2008)
Moss and liverwort xyloglucans contain galacturonic acid and are structurally distinct from
the xyloglucans synthesized by hornworts and vascular plants. Glycobiology, 18, 891‐904.
Perini, M.A., Sin, I.N., Villarreal, N.M., Marina, M., Powell, A.L., Martínez,
G.A., and Civello, P.M. (2017) Overexpression of the carbohydrate binding module from
Solanum lycopersicum expansin 1 (Sl–EXP1) modifies tomato fruit firmness and Botrytis
cinerea susceptibility. Plant Physiol. Biochem., 113, 122–132.
Perrin, R.M., DeRocher, A.E., Bar–Peled, M., Zeng, W., Norambuena, L.,
Orellana, A., Raikhel, N.V., and Keegstra, K. (1999) Xyloglucan fucosyltransferase,
an enzyme involved in plant cell wall biosynthesis. Science, 284, 1976–1979.
75
Persson, S., Paredez, A., Carroll, A., Palsdottir, H., Doblin, M., Poindexter, P.,
Khitrov, N., Auer, M., and Somerville, C.R. (2007) Genetic evidence for three unique
components in primary cell–wall cellulose synthase complexes in Arabidopsis. Proc. Natl.
Acad. Sci. USA, 104, 15566–15571.
Popper, Z.A., and Fry, S.C. (2003) Primary cell wall composition of bryophytes and
charophytes. Ann. Bot., 91, 1‐12.
Popper, Z.A., and Fry, S.C. (2004) Primary cell wall composition of pteridophytes and
spermatophytes. New Phytol., 164, 165–174.
Popper, Z.A., and Fry, S.C. (2008) Xyloglucan‐pectin linkages are formed intra‐
protoplasmically, contribute to wall‐assembly, and remain stable in the cell wall. Planta,
227, 781–794.
Postek, M.T., Vladar, A., Dagata, J., Farkas, N., Ming, B., Wagner, R., Raman,
A., Moon, R. J., Sabo, R., Wegner, T. H., and Beecher, J. (2011) Development of the
metrology and imaging of cellulose nanocrystals. Meas. Sci. Technol., 22, 024005.
Probine, M.C., and Preston, R.D. (1962) Cell growth and the structure and mechanical
properties of the wall in internodal cells of Nitella opaca. II. Mechanical properties of the
walls. J. Exp. Bot., 13, 111–127.
Purushotham, P., Cho, S.H., Díaz–Moreno, S.M., Kumar, M., Nixon, B.T.,
Bulone, V., and Zimmer, J. (2016) A single heterologously expressed plant cellulose
synthase isoform is sufficient for cellulose microfibril formation in vitro. Proc. Natl. Acad.
Sci. USA, 113, 11360–11365.
Radford, J.E., and White, R.G. (1998) Localization of a myosin–like protein to
plasmodesmata. Plant J., 14, 743–750.
Rae, A.L., Harris, P.J., Bacic, A., and Clarke, A.E. (1985) Composition of the cell
walls of Nicotiana alata Link et Otto pollen tubes. Planta, 166, 128–133.
Ranby, B.G. (1949) Aqueous Colloidal Solutions of Cellulose Micelles. Acta. Chem. Scand.,
3, 649–650.
Ranby, B.G. (1951) Fibrous macromolecular systems. Cellulose and muscle. The colloidal
properties of cellulose micelles. Discuss. Faraday Soc., 11, 158–164.
Rietveld, A., and Simons, K. (1998) The differential miscibility of lipids as the basis for
the formation of functional membrane rafts. Biochim. Biophys. Acta, 1376, 467–479.
Richmond, T.A., and Somerville, C.R. (2000) The cellulose synthase superfamily.
Plant Physiol., 124, 495–498.
Rinne, P.L.H., van den Boogaard, R., Mensink, M.G.J., Kopperud, C.,
Kormelink, R., Goldbach, R., and van der Schoot, C . (2005) Tobacco plants
respond to the constitutive expression of the tospovirus movement protein NSM with a
heat–reversible sealing of plasmodesmata that impairs development. Plant J., 43, 688–707.
Robards, A.W. (1968) Desmotubule—a Plasmodesmatal Substructure. Nature, 218, 784.
76
Roberts, A.G., and Oparka, K.J. (2003) Plasmodesmata and the control of symplastic
transport. Plant Cell Environ., 26, 103–124.
Roudier, F., Fernandez, A.G., Fujita, M., Himmelspach, R., Borner, G.H.H.,
Schindelman, G., Song, S., Baskin, T.I., Dupree, P., Wasteneys, G.O., and
Benfey, P.N. (2005) COBRA, an Arabidopsis extracellular glycosyl–phosphatidyl inositol–
anchored protein, specifically controls highly anisotropic expansion through its involvement
in cellulose microfibril orientation. Plant Cell, 17, 1749–1763.
Rounds, C.M., and Bezanilla, M. (2013) Growth mechanisms in tip–growing plant cells.
Annu. Rev. Plant Biol., 64, 243–265.
Ruel, K., Nishiyama, Y., and Joseleau, J.P. (2012) Crystalline and amorphous
cellulose in the secondary walls of Arabidopsis. Plant Sci., 193–194, 48–61.
Safra–Dassa, L., Shani, Z., Danin, A., Roiz, L., Oded Shoseyov, O., and Wolf, S.
(2006) Growth modulation of transgenic potato plants by heterologous expression of
bacterial carbohydrate–binding module. Mol. Breed., 17, 355–364.
Sager, R., and Lee, J–Y. (2014) Plasmodesmata in integrated cell signalling: insights
from development and environmental signals and stresses. J. Exp. Bot., 65, 6337–6358.
Saito, T., Nishiyama, Y., Putaux, J.L., Vignon, M., and Isogai, A.. (2006)
Homogeneous suspentions of individualized microfibrils from TEMPO–catalysed oxidation
of native cellulose. Biomacromolecules, 7, 1687–1691.
Sato, S., Kato, T., Kakegawa, K., Ishii, T., Liu, Y.G., Awano, T., Takabe, K.,
Nishiyama, Y., Kuga, S., Nakamura, Y., Tabata, S., and Shibata, D. (2001) Role of
the putative membrane–bound endo–1,4–betaglucanase KORRIGAN in cell elongation and
cellulose synthesis in Arabidopsis thaliana. Plant Cell Physiol., 42, 251–263.
Saxena, I.M., Kudlicka, K., Okuda, K., and Brown, R.M. (1994). Characterization of
genes in the cellulose–synthesizing operon (acs operon) of Acetobacter xylinum:
implications for cellulose crystallization. J. Bacteriol., 176, 5735–5752.
Saxena, I.M., Brown, R.M., Fevre, M., Geremia, R., and Henrissat, B. (1995)
Multidomain architecture of β–glycosyl transferases: implications for mechanism of action.
J. Bacteriol., 177, 1419–1424.
Saxena, I.M., and Brown, R.M. (1997) Identification of cellulose synthase(s) in higher
plants: Sequence analysis of processive β–glycosyltransferases with the common motif 'D,
D, D35Q(R,Q)XRW'. Cellulose, 4, 33–49.
Saxena, I.M., Brown, R.M., and Dandekar, T. (2001) Structure–function
characterization of cellulose synthase: relationship to other glycosyltransferases.
Phytochemistry, 57, 1135–1148.
Scheller, H.V., and Ulvskov, P. (2010) Hemicelluloses. Annu. Rev. Plant Biol., 61, 263–
289.
Schindelman, G., Morikami, A., Jung, J., Baskin, T.I., Carpita, N.C.,
Derbyshire, P., McCann, M.C., and Benfey, P.N. (2001) COBRA encodes a putative
GPI–anchored protein, which is polarly localized and necessary for oriented cell expansion
in Arabidopsis. Gene Dev., 15, 1115–1127.
77
Schlupmann, H., Bacic, A., and Read, S. (1993) A novel callose synthase from pollen
tubes of Nicotiana. Planta, 191, 470–481.
Schlupmann, H., Bacic, A., and Read, S.M. (1994) Uridine diphosphate glucose
metabolism and callose synthesis in cultured pollen tubes of Nicotiana alata Link et Otto.
Plant Physiol., 105, 659–670.
Schulz, A. (1995). Plasmodesmal widening accompanies the short-term increase in
symplasmic phloem unloading in pea root tips under osmotic stress. Protoplasma, 188, 22–
37.
Sethaphong, L., Haigler, C.H., Kubicki, J.D., Zimmer, J., Bonetta, D., DeBolt,
S., and Yingling, Y.G. (2013) Tertiary model of a plant cellulose synthase. Proc. Natl.
Acad. Sci. USA, 110, 7512–7517.
Shibazaki, H., Saito, M., Kuga, S., and Okano, T. (1998). Native Cellulose II
production by Acetobacter Xylinum under physical constraints. Cellulose, 5, 165–173.
Shoseyov, O., Levy, I., Shani, Z., and Mansfield, S.D. (2003) Modulation of wood
fibers and paper by cellulose–binding domains. Applications of Enzymes to Lignocellulosics,
855, 116–131.
Shpigel, E., Roiz, L., Goren, R., and Shoseyov, O. (1998a) Bacterial cellulose–binding
domain modulates in vitro elongation of different plant cells. Plant Physiol., 117, 1185–1194.
Shpigel, E., Elias, D., Cohen, I.R., and Shoseyov, O. (1998b) Production and
purification of a recombinant human hsp60 epitope using the cellulosebinding domain in
Escherichia coli. Protein Expr. Purif., 14, 185–191.
Simpson, P.J., Xie, H., Bolam, D.N., Gilbert, H.J., and Williamson, M.P. (2000)
The structural basis for the ligand specificity of family 2 carbohydrate–binding modules. J.
Biol. Chem., 275, 41137–4142.
Sinnott, M.L. (1990) Catalytic mechanisms of enzymatic glycosyl transfer. Chem. Rev.
Washington, DC, U.S, 90, 1171–1202.
Sisson, W.A. (1938) The existanve of mercierized cellulose and its orientation in Halicystis
as indicated by x–ray diffraction analysis. Science, 87, 350.
Sivaguru, M., Fujiwara, T., Šamaj, J., Baluška, F., Yang, Z., Osawa, H., Maeda,
T., Mori, T., Volkmann, D., and Matsumotoet, H. (2000) Aluminum–induced1→3–
β–d–glucan inhibits cell–to–cell trafficking of molecules through plasmodesmata. A new
mechanism of aluminium toxicity in plants. Plant Physiol., 124, 991–1006.
Slabaugh, E., Davis, J.K., Haigler, C.H., Yingling, Y.G., and Zimmer, J. (2014)
Cellulose synthases: new insights from crystallography and modeling. Trends Plant Sci., 13,
1360–1385.
Somerville, C. (2006) Cellulose synthesis in higher plants. Annu. Rev. Cell Dev. Biol., 22,
53–78.
Song, D., Shen, J., and Li, L. (2010) Characterization of cellulose synthase complexes in
Populus xylem differentiation. New Phytol, 187, 777–790.
78
Southall, S.M., Simpson, P.J., Gilbert, H.J., Williamson, G., and Williamson,
M.P. (1999) The starch–binding domain from glucoamylase disrupts the structure of
starch. FEBS Lett., 447, 58–60.
Sparkes, I.A., Runions, J., Kearns, A., and Hawes, C. (2006) Rapid, transient
expression of fluorescent fusion proteins in tobacco plants and generation of stably
transformed plants. Nat. Protoc., 1, 2019–2025.
Srivastava, V., Malm, E., Sundqvist, G., and Bulone, V. (2013) Quantitative
proteomics reveals that plasma membrane microdomains from poplar cell suspension
cultures are enriched in markers of signal transduction, molecular transport, and callose
biosynthesis. Mol. Cell. Proteomics, 12, 3874–3885.
Stasinopoulos, S.J., Fisher, P.R., Stone, B.A., and Stanisich, V.A. (1999) Detection
of two loci involved in (1–>3)–β–glucan (curdlan) biosynthesis by Agrobacterium sp.
ATCC31749, and comparative sequence analysis of the putative curdlan synthase gene.
Glycobiology, 9, 31–41.
Sticklen, M.B (2008) Plant genetic engineering for biofuel production: towards affordable
cellulosic ethanol. Nat. Rev. Genet., 9, 433–443.
Stipanovic, A., and Sarko, A. (1976) Packing analysis of carbohydrates and
polysaccharides. 6. Molecular and crystal structure of regenerated cellulose II.
Macromolecules, 9, 851–857.
Stone, B.A., and Clarke, A.E. (1992) Chemistry and biology of (1−>3)–β–glucans. La
Trobe University Press, Victoria, Australia, 431–491.
De Storme, N., and Geelen, D. (2014) Callose homeostasis at plasmodesmata:
molecular regulators and developmental relevance. Front. Plant Sci., 5, 138.
Sturcová, A., His, I., Apperley, D.C., Sugiyama, J., and Jarvis, M.C. (2004)
Structural details of crystalline cellulose from higher plants. Biomacromolecules, 5, 1333–
1339.
Sugiyama, J., Vuong, R., and Chanzy. H. (1991) Electron diffraction studyon the two
crystalline phases occurring in native cellulose from an algalcell wall. Macromolecules, 24,
4168–4175.
Suslov,D. and Verbelen, J-P. (2006). Cellulose orientation determines mechanical
anisotropy in onion epidermis cell walls. J. Exp. Bot., 57, 2183–2192.
Taiz, L., and Zeiger, E. (eds) (2010). Plant Physiology, 5th ed, Sinauer Associates.
Takahashi, J., Rudsander, U.J., Hedenstrom, M., Banasiak, A., Harholt, J.,
Amelot, N., Immerzeel, P., Ryden, P., Endo, S., Ibatullin, F.M., Brumer, H., del
Campillo, E., Master, E.R., Scheller, H.V., Sundberg, B., Teeri, T.T., and
Mellerowicz, E.J. (2009) KORRIGAN1 and its aspen homolog PttCel9A1 decrease
cellulose crystallinity in Arabidopsis stems. Plant Cell Physiol., 50, 1099–1115.
Takeda, T., Furuta, Y., Awano, T., Mizuno, K., Mitsuishi, Y., and Hayashi, T.
(2002). Suppression and acceleration of cell elongation by integration of xyloglucans in pea
stem segments. Proc. Natl. Acad. Sci. USA, 99, 9055–9060.
79
Tanner, W., Malinsky, J., and Opekarová, M. (2011) In plant and animal cells,
detergent–resistant membranes do not define functional membrane rafts. Plant Cell, 23,
1191–1193.
Taylor, N.G., Laurie, S., and Turner, S.R. (2000) Multiple cellulose synthase catalytic
subunits are required for cellulose synthesis in Arabidopsis. Plant Cell, 12, 2529–2540.
Taylor, N.G., Howells, R.M., Huttly, A.K., Vickers, K., and Turner, S.R. (2003)
Interactions among three distinct CesA proteins essential for cellulose synthesis. Proc. Natl.
Acad. Sci. USA, 100, 1450–1455.
The Arabidopsis Genome Initiative (2000) Analysis of the genome sequence of the
flowering plant Arabidopsis thaliana. Nature, 408, 796–815.
Thiele, K., Wanner, G., Kindzierski, V., Jurgens, G., Mayer, U., Pachl, F., and
Assaad, F.F. (2009) The timely deposition of callose is essential for cytokinesis in
Arabidopsis. Plant J., 58, 13–26.
Thomas, L.H., Forsyth, V.T., Sturcova, A., Kennedy, C.J., May, R.P., Altaner, C.
M., Apperley, D.C., Wess, T.J., and Jarvis, M.C. (2013) Structure of cellulose
microfibrils in primary cell walls from collenchyma. Plant Physiol., 161, 465–476.
Tormo, J., Lamed, R., Chirino, A.J., Morag, E., Bayer, E.A., Shoham, Y., and
Steitz, T.A. (1996) Crystal structure of a bacterial family–III cellulose–binding domain: a
general mechanism for attachment to cellulose. EMBO J., 15, 5739–5751.
Tucker, E.B., and Boss, W.F. (1996) Mastoparan–induced intracellular Ca2+ fluxes may
regulate cell–to–cell communication in plants. Plant Physiol., 111, 459–467.
Turbak, A.F., Snyder, F.W., and Sandberg, K.R. (1983) Microfibrillated cellulose. In
U.S. Patent 4374702 A: 1983.
Turner, A., Bacic, A., Harris, P.J., and Read, S.M. (1998) Membrane fractionation
and enrichment of callose synthase from pollen tubes of Nicotiana alata Link et Otto.
Planta, 205, 380–388.
Tuskan, G., Difazio, S., Jansson, S., Bohlmann, J., Grigoriev, I., Hellsten, U.,
Putnam, N., Ralph, S., Rombauts, S., Salamov, A., Schein, J., Sterck, L., Aerts,
A., Bhalerao, R.R., Bhalerao, R.P., Blaudez, D., Boerjan, W., Brun, A., Brunner,
A., Busov, V., Campbell, M., Carlson, J., Chalot, M., Chapman, J., Chen, G.L.,
Cooper, D., Coutinho, P.M., Couturier, J., Covert, S., Cronk, Q., Cunningham,
R., Davis, J., Degroeve, S., Déjardin, A., Depamphilis, C., Detter, J., Dirks, B.,
Dubchak, I., Duplessis, S., Ehlting, J., Ellis, B., Gendler, K., Goodstein, D.,
Gribskov, M., Grimwood, J., Groover, A., Gunter, L., Hamberger, B., Heinze,
B., Helariutta, Y., Henrissat, B., Holligan, D., Holt, R., Huang, W., Islam–
Faridi, N., Jones, S., Jones–Rhoades, M., Jorgensen, R., Joshi, C., Kangasjärvi,
J., Karlsson, J., Kelleher, C., Kirkpatrick, R., Kirst, M., Kohler, A., Kalluri, U.,
Larimer, F., Leebens–Mack, J., Leplé, J.C., Locascio, P., Lou, Y., Lucas, S.,
Martin, F., Montanini, B., Napoli, C., Nelson, D.R., Nelson, C., Nieminen, K.,
Nilsson, O., Pereda, V., Peter, G., Philippe, R., Pilate, G., Poliakov, A.,
Razumovskaya, J., Richardson, P., Rinaldi, C., Ritland, K., Rouzé, P., Ryaboy,
D., Schmutz, J., Schrader, J., Segerman, B., Shin, H., Siddiqui, A., Sterky, F.,
80
Terry, A., Tsai, C.J., Uberbacher, E., Unneberg, P., Vahala, J., Wall, K.,
Wessler, S., Yang, G., Yin, T., Douglas, C., Marra, M., Sandberg, G., Van de
Peer, Y., and Rokhsar, D. (2006) The genome of black cottonwood, Populus trichocarpa
(Torr. & Gray). Science, 313, 1596–1604.
Tvaroška, I. (2015) Atomistic insight into the catalytic mechanism of glycosyltransferases
by combined quantum mechanics/molecular mechanics (QM/MM) methods. Carbohydr.
Res., 403, 38–47
Unligil, U.M., and Rini, J.M. (2000) Glycosyltransferase structure and mechanism.
Curr. Opin. Struct. Biol., 10, 510–517.
Vaaje–Kolstad, G., Westereng, B., Horn, S.J., Liu, Z., Zhai, H., Sorlie, M., and
Eijsink, V.G. (2010) An oxidative enzyme boosting the enzymatic conversion of
recalcitrant polysaccharides. Science, 330, 219–222.
Vanholme, R., Demedts, B., Morreel, K., Ralph, J., and Boerjan, W. (2010) Lignin
biosynthesis and structure. Plant Physiol., 153, 895‐905.
Vanzin, G.F., Madson, M., Carpita, N.C., Raikhel, N.V., Keegstra, K., and
Reiter, W.D. (2002) The mur2 mutant of Arabidopsis thaliana lacks fucosylated
xyloglucan because of a lesion in fucosyltransferase AtFUT1. Proc. Natl. Acad. Sci. USA, 99,
3340–3345.
Vaten, A., Dettmer, J., Wu, S., Stierhof, Y–D., Miyashima, S., Yadav, S.R.,
Roberts, C.J., Campilho, A., Bulone, V., Lichtenberger, R., Lehesranta, S.,
Mähönen, A.P., Kim, J.Y., Jokitalo, E., Sauer, N., Scheres, B., Nakajima, K.,
Carlsbecker, A., Gallagher, K.L., and Helariutta, Y. (2011) Callose biosynthesis
regulates symplastic trafficking during root development. Dev. Cell., 21, 1144–1155.
Verma, D.P., and Hong, Z. (2001) Plant callose synthase complexes. Plant Mol. Biol, 47,
693–701.
Vincken, J.P., York, W.S., Beldman, G., and Voragen, A. (1997) Two general
branching patterns of xyloglucan, XXXG and XXGG. Plant Physiol., 114, 9–13.
Vincken, J.P., Schols, H.A., Oomen, R.J.F.J., McCann, M.C., Ulvskov, P.,
Voragen, A.G.J., and Visser, R.G.F. (2003) If homogalacturonan were a side chain of
rhamnogalacturonan I. Implications for cell wall architecture. Plant Physiol., 132, 1781–
1789.
Vrielink, A., Ruger, W., Driessen, H.P., and Freemont, P.S. (1994). Crystal
structure of the DNA modifying enzyme β-glucosyltransferase in the presence and absence
of the substrate uridine diphosphoglucose. EMBO J., 13, 3413–3422.
Wada, M., Chanzy, H., Nishiyama, Y., and Langan, P. (2004). Cellulose IIII crystal structure and hydrogen bonding by synchrotron X-ray and neutron fiber diffraction. Macromolecules, 37, 8548–8555.
Wada, M., Heux, L., and Sugiyama, J. (2004) Polymorphism of cellulose I family:
reinvestigation of cellulose IV. Biomacromolecules, 5, 1385–1391.
81
Wagberg, L., Decher, G., Norgren, M., Lindström, T., Ankerfors, M., and Axnas,
K. (2008) The build–up of polyelectrolyte multilayers of microfibrillated cellulose and
cationic polyelectrolytes. Langmuir, 24, 748–795.
Wagner, G.K. and Pesnot, T. (2010) Glycosyltransferases and their assays.
ChemBioChem, 11, 1939–1949.
Wang, M., Olszewska, A., Walther, A., Malho, J. M., Schacher, F.H.,
Ruokolainen, J., Ankerfors, M., Laine, J., Berglund, L. A., Österberg, M., and
Ikkala, O. (2011) Colloidal ionic assembly between anionic native cellulose nanofibrils and
cationic block copolymer micelles into biomimetic nanocomposites. Biomacromolecules, 12,
2074–2081.
Wang, X., Sager, R., Cui, W., Zhang, C., Lu, H., and Leea, J–Y. (2013) Salicylic acid
regulates plasmodesmata closure during innate immune responses in Arabidopsis. Plant
Cell, 25, 2315–2329.
Wang, T., and Hong, M. (2016) Solid–state NMR investigations of cellulose structure
and interactions with matrix polysaccharides in plant primary cell walls. J. Exp. Bot., 67,
503–514.
Wu, A., Harriman, R.W., Frost, D.J., Read, S.M., and Wasserman, B.P. (1991)
Rapid enrichment of CHAPS–solubilized UDPglucose: (1,3)–b–glucan (callose) synthase
from Beta vulgaris L. by product entrapment. Entrapment mechanisms and polypeptide
characterization. Plant Physiol., 97, 684–692.
Wullschleger, S.D., Jansson, S., and Taylor, G. (2002) Genomics and forest biology:
Populus emerges as the perennial favorite. Plant Cell, 14, 2651–2655.
Xiao, C., Zhang, T., Zheng, Y., Cosgrove, D.J. and Anderson, C.T. (2016)
Xyloglucan deficiency disrupts microtubule stability and cellulose biosynthesis in
Arabidopsis, altering cell growth and morphogenesis. Plant Physiol., 170, 234–249.
Xie, B., Wang, X.M., Zhu, M.S., Zhang, Z.M., and Hong, Z.L. (2011) CalS7 encodes
a callose synthase responsible for callose deposition in the phloem. Plant J., 65, 1–14.
Xu, G–Y., Ong, E., Gilkes, N.R., Kilburn, D.G., Muhandiram, D.R., Harris–
Brandts, M., Carver, J–P., Kay, L.E., and Harvey, T.S. (1995) Solution structure of a
cellulose–binding domain from cellulomonas fimi by nuclear magnetic resonance
spectroscopy. Biochemistry, 34, 6993–7009.
Yamanaka, S., Ishihara, M., and Sugiyama, J. (2000) Structural modification of
bacterial cellulose. Cellulose, 7, 213–225.
Yang, M., Brazier, M., Edwards, R. and Davis, B .G. (2005). High‐throughput mass‐
spectrometry monitoring for multisubstrate enzymes: determining the kinetic parameters
and catalytic activities of glycosyltransferases. ChemBioChem, 6, 346-357.
Young, S.H., Dong, W.J., and Jacobs, R.R. (2000) Observation of a partially opened
triple–helix conformation in 1→3–β–glucan by fluorescence resonance energy transfer
spectroscopy. J. Biol. Chem., 275, 11874–11879.
Zabotina, O.A., van de Ven, W.T.G., Freshour, G., Drakakaki, G., Cavalier, D.,
Mouille, G., Hahn, M.G., Keegstra, K., and Raikhel, N.V. (2008) Arabidopsis XXT5
82
gene encodes a putative α–1,6–xylosyltransferase that is involved in xyloglucan
biosynthesis. Plant J., 56, 101–115.
Zabotina, O.A., Avci, U., Cavalier, D., Pattathil, S., Chou, Y.H., Eberhard, S.,
Danhof, L., Keegstra, K., and Hahn, M. G. (2012) Mutations in multiple XXT genes of
Arabidopsis reveal the complexity of xyloglucan biosynthesis. Plant Physiol., 159, 1367–
1384.
Zavaliev, R., Ueki, S., Epel, B.L., and Citovsky, V. (2011) Biology of callose (β–1,3–
glucan) turnover at plasmodesmata. Protoplasma, 248, 117–130.
Zavaliev, R., Levy, A., Gera, A., and Epel, B.L. (2013) Subcellular dynamics and role
of Arabidopsis β–1,3–glucanases in cell–to–cell movement of tobamoviruses. Mol. Plant–
Microbe Interact., 26, 1016–1030.
Zhang, Z., Sebe, G., Rentsch, D., Zimmermann, T., and Tingaut, P. (2014)
Ultralightweight and flexible silylated nanocellulose sponges for the selective removal of oil
from water. Chem. Mater, 26, 2659–2668.
Zheng, Y., Wang, X., Chen, Y., Wagner, E. and Cosgrove, D.J. (2018). Xyloglucan
in the primary cell wall: assessment by FESEM, selective enzyme digestions and nanogold
affinity tags. Plant J., 93: 211-226.
Zhou, Q., Greffe, L., Baumann, M.J., Malmström, E., Teeri, T.T., and Brumer
H. (2005) Use of xyloglucan as a molecular anchor for the elaboration of polymers from
cellulose surfaces: A general route for the design of biocomposites. Macromolecules, 38,
3547–3549.
Zhou, Q., Brumer, H., and Teeri, T.T. (2009) Self–organization of cellulose
nanocrystals adsorbed with xyloglucan oligosaccharide−poly(ethylene glycol)−polystyrene
triblock copolymer. Macromolecules, 42, 5430–5432.