Understanding and Manipulating Primary Cell Walls in Plant Cell Suspension Cultures

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

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

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.

i 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.

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.

iii 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.

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

v 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

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

vii

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.

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

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

3 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).

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

5 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

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

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

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.

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).

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.

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).

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- 4 linked polymer made up by glucopyranosyl residues in the C1 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

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.

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

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

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

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

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).

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).

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).

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).

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).

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

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

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.

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

27 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

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

29 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.

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

31 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).

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.

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).

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).

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).

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.

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.

Figure 12. Overview of radiometric and spectrophotometric assays for glycosyltransferases (Reprinted with permission from Brown et al., 2012).

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.

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

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).

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.

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.

30 WT

20 CBM3 material.hour

10 nmolof sugars released/g

0 0.5M NaOH 0.5M NaOH No NaOH Tobacco Arabidopsis Arabidopsis

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,

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.

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

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).

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 Modulus Tensile strength Strain-to-failure Work-of-fracture (g cm-3) (GPa) (MPa) (%) (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.

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.

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

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.

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.

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.

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.

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.

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.

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.

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.

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.

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,

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.

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.,

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.

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

81 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.