1 1 Introduction 1.1 The plant cell wall One major difference between organisms assigned to the plant and the animal kingdom is that the plant cell is surrounded by a network of polymers which is called the plant cell wall. Robert Hooke, one of the early microscopists, identified in the 1660s cell wall structures in samples of cork and coined later the term “cell” for the enclosed compartment. Three hundred and fifty years later our comprehensive understanding of structure, function and metabolism of cell walls is far from being complete. The cell wall in plants represents the outer border to the environment or the interface between adjacent cells and thereby controls cell adhesion (Cosgrove, 1997) and cell to cell communication. Approximately 40 distinct cell types can be found on average in a plant and in turn their walls require different structures to accommodate their various functions. The cell wall resists internal turgor pressure (Bacic et al., 1998) and determines cell shape. Plant cell walls are a physical barrier protecting the protoplast against pathogen infection (Darvill et al., 1980). Furthermore, the cell wall can harbour physiologically active signaling compounds (Ridley et al., 2001) or defense molecules such as inhibitor proteins which prevent carbohydrate degradation by microbial glycoside hydrolases (Lionetti et al., 2007). Additionally it can be a source of storage carbohydrates which are mobilised and further metabolised upon germination (Buckeridge et al., 1992; Franco et al., 1996; Santos et al., 2004). Porosity, pH, charge and salt concentration of cell walls are determined by their compositions which in turn influence apoplastic water transport as well as solute and nutrient storage and uptake. Three substructures can be distinguished in terms of their developmental emergence. The middle lamella is synthesised by two dividing cells during cell plate formation (Matar and Catesson, 1988) whereas primary and secondary cell walls are products of individual cells. The middle lamella consists mainly of pectins. The primary cell wall in dicotyledonous plants is composed of cellulose, hemicelluloses and pectins approximately accounting for a third each whereas 1- 5% is made of proteins (Cosgrove, 1997). It is a highly hydrated (75-80% water; Cosgrove, 1997) composite material in which the loadbearing cellulose- hemicellulose network is embedded in gel-like matrix consisting of pectins. Due to its composite character the primary cell wall can maintain cell shape while it is flexible enough to allow tropic movements and cell expansion during growth. After elongation and growth have ceased secondary wall formation occurs. The secondary cell wall consists mainly of crystalline cellulose and due to impregnation with phenolic lignins becomes water-repellent and stiff allowing water transport in heights to over hundred meters and thereby to resist pressures of >100 MPa. In all development stages proteins, either structural or enzymatic, are associated with the wall. Structural hydroxyproline rich proteins (HRGPs),
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1 Introduction
1.1 The plant cell wall
One major difference between organisms assigned to the plant and the animal kingdom is that the plant cell is surrounded by a network of polymers which is called the plant cell wall. Robert Hooke, one of the early microscopists, identified in the 1660s cell wall structures in samples of cork and coined later the term “cell” for the enclosed compartment. Three hundred and fifty years later our comprehensive understanding of structure, function and metabolism of cell walls is far from being complete. The cell wall in plants represents the outer border to the environment or the interface between adjacent cells and thereby controls cell adhesion (Cosgrove, 1997) and cell to cell communication. Approximately 40 distinct cell types can be found on average in a plant and in turn their walls require different structures to accommodate their various functions. The cell wall resists internal turgor pressure (Bacic et al., 1998) and determines cell shape. Plant cell walls are a physical barrier protecting the protoplast against pathogen infection (Darvill et al., 1980). Furthermore, the cell wall can harbour physiologically active signaling compounds (Ridley et al., 2001) or defense molecules such as inhibitor proteins which prevent carbohydrate degradation by microbial glycoside hydrolases (Lionetti et al., 2007). Additionally it can be a source of storage carbohydrates which are mobilised and further metabolised upon germination (Buckeridge et al., 1992; Franco et al., 1996; Santos et al., 2004). Porosity, pH, charge and salt concentration of cell walls are determined by their compositions which in turn influence apoplastic water transport as well as solute and nutrient storage and uptake. Three substructures can be distinguished in terms of their developmental emergence. The middle lamella is synthesised by two dividing cells during cell plate formation (Matar and Catesson, 1988) whereas primary and secondary cell walls are products of individual cells. The middle lamella consists mainly of pectins. The primary cell wall in dicotyledonous plants is composed of cellulose, hemicelluloses and pectins approximately accounting for a third each whereas 1-5% is made of proteins (Cosgrove, 1997). It is a highly hydrated (75-80% water; Cosgrove, 1997) composite material in which the loadbearing cellulose-hemicellulose network is embedded in gel-like matrix consisting of pectins. Due to its composite character the primary cell wall can maintain cell shape while it is flexible enough to allow tropic movements and cell expansion during growth. After elongation and growth have ceased secondary wall formation occurs. The secondary cell wall consists mainly of crystalline cellulose and due to impregnation with phenolic lignins becomes water-repellent and stiff allowing water transport in heights to over hundred meters and thereby to resist pressures of >100 MPa. In all development stages proteins, either structural or enzymatic, are associated with the wall. Structural hydroxyproline rich proteins (HRGPs),
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which are highly glycosylated, can be found in the wall. HRGPs (reviewed by Farrokhi et al., 2006) can be split into Arabinogalactan proteins (AGPs), extensins and proline rich proteins (PRPs) which are thought to be involved in a variety of developmental processes such as xylem formation (Dolan et al., 1995; Motose et al., 2004), secondary cell wall thickening (Schindeler et al., 1995) and programmed cell death (Gao and Showalter, 2000). The enzymatic wall proteins contain expansins, glycosyl hydrolases and transgylcosylases which catalytically modify cell wall carbohydrates. It is thought that wall polysaccharide metabolism plays an important role in cell elongation and response to pathogens. The primary component of all cell walls is cellulose, the most abundant polymer on earth. Cellulose is a homopolymer consisting of β-1,4-linked glucose units, forming a crystalline cellulose microfibrile (Somerville, 2006). Cellulose is synthesized at the plasma membrane by rosette like structures (Delmer, 1999; Brown and Saxena, 2000). During the self-assembly process of the microfibrile the crystallinity can circumstantially be broken. Perfect alignment might be prevented by e.g. trapping other surrounding cell wall polymers between the different chains of the nascent microfibril. These amorphous parts of cellulose are more susceptible to cell wall degrading enzymes and chemicals but may also function as interface between cellulose and matrixpolysaccharides, such as pectins and hemicelluloses. Primary cell walls are also rich in another major group of components, the pectins. All pectic polysaccharides contain D-galacturonic acid (GalA) and thus present polyanions. Pectins modulate wall porosity (Baron-Epel et al., 1998), pH, degree of hydration and ion strength since they have the ability to bind metal ions. Furthermore, they are involveld in cell adhesion (Iwai et al., 2000). They are the most soluble part of the wall and can be easily extracted by hot water, chelators or mild alkaline solutions e.g. Na2CO3 (Cosgrove, 1997). Structurally the pectic network is devided into homogalacturonan (HG), rhamnogalacturonan I (RGI) and rhamnogalacturonan II (RG II) (O`Neill et al., 1980). HG consist of an unbranched chain of α-1,4-linked galacturonic acid residues. The carboxyl moiety at C6 is thought to be highly methylesterified (70-80%) during secretion (Willats et al., 2001). The degree of esterification and the presence of cations such as Ca2+ influence the gelling properties. Cell wall located pectin methylesterases (PME) cleave some of the methyl groups to initiate binding of the carboxylate ions to Ca2+ (Carpita and McCann, 2000). RGI is composed of a repeating disaccharide α -1, 2-D-Rha-α-1, 4-D-GalA. About one third of the carboxyl groups of the GalA units are acetylated and about half of the C4 hydroxyl groups of Rha units are linked to α-1, 5-L-arabinans or β-1, 4-D-galactans but the degree of substitution depends heavily on species and tissue. RGII consists of HG backbone which is highly decorated with complex structures consisting of unusual sugars.
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Figure 1.1: Cell wall model modified from Pauly. Proteins are left out for reasons of simplification. The cell wall can be divided into three structurally and compositionally different entities (cellulose, hemicelluloses and pectins). Both hemicelluloses and pectins can be further subdivided. Xyloglucan, galactomannan and arabinoxylan represent hemicelluloses whereas pectins can be divided into homogalacturonan (HG), rhamnogalacturonan I (RGI) and rhamnogalacturonan II (RGII).
In the cell wall 14 different sugars can be found most of which are present in the side chains of RGII. Borate has the ability to connect apiose residues in RGII side chains and thus allow RGII side chains to form a structure which is discussed to have besides the cellulose-hemicellulose network also loadbearing abilities (O´Neill et al., 2001).
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1.2 Xyloglucan structure and properties
1.2.1 Fine structure of xyloglucan
Another major group of wall polysaccharides, the hemicelluloses, are due to their structural similarity non-covalently connected to cellulose microfibrils via hydrogen bonds (McQueen and Cosgrove, 1994) and form the loadbearing cellulose-hemicellulose network. By definition hemicelluloses are polymers which can be extracted from cell walls by chaotropic agents like alkali but not by mild buffers, hot water or chelators (Hayashi and McLachlan, 1984; O´Neill and York, 2003) and thereby comprises xyloglucans, xylans, mannans and derivatives of them. Hemicelluloses share the presence of β-D-pyranosyl backbone as a common feature. However, unlike cellulose they do have side chains which prevent crystallisation. Xyloglucan (XyG) interlocks cellulose microfibrils by spanning the distance between adjacent fibres and places them in proper spatial arrangement. Xyloglucan has been found in the primary cell walls of all higher plants that have been investigated to date (Popper and Fry, 2004). However, the percentage and the fine structures vary among plant species (McNeill et al., 1984) and even among cell wall domains within a single cell. XyG comprises 10–25% of the cell walls of dicots and non-graminaceous monocots and is the most prominent hemicellulose in these walls (McNeill, 1984). Grasses (Poaceae), in contrast contain only 6% XyG (Carpita, 1996; Gibeaut et al., 2005). XyG is built of β-1,4-linked D-gluco-pyranosyl residues creating a backbone to which several α-D-Xylp substituents are attached at O6. These xylosyl residues can be further substituted with e.g. β-1,2-linked D-Galp before additional α-1,2-L-Fucp residues are attached. To unambiguously describe XyG structures a concise nomenclature was developed (Tab. 1.1; Fry et al., 1993; Hantus et al., 1997; Ray et al., 2004). According to this nomenclature a XyG molecule is named after partitioning the backbone into segments of single glucosyl residues and their attached side chain. For instance an unsubstituted bare backbone glucosyl residue is abbreviated with “G” while a glucosyl backbone unit which is further substituted with an α-1,2-linked D-Xylp residue is encoded by “X”. In addition, XyG can be O-acetylated at various positions (Kiefer et al., 1989) which is indicated by underscored letters (Jia et al., 2005). Digestion of xyloglucan with β-1,4-endoglucanase (EG) revealed a repetitive block structure (York et al., 1990) of XyGOs mostly consisting of oligosaccharides with a cellotetraose backbone structure. Two types of XyG have been classified according to the number of xylosyl substitutions. The “XXXG”-type in which approximately 75% of the backbone residues are highly branched (Vincken et al., 1997) comprises 10–25% of the primary cell walls of the gymnosperms, dicotyledonous plants exceptional Laminales (e.g. peppermint, basil) and Solanales (potato, tomato, tobacco), and the non-graminaceous
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monocotyledonous plants (Hoffman et al., 2005). In the “XXGG”-type, which is typical for solanaceous plants, only about half the backbone is substituted. The main repetitive motifs in the dicotyledonous “XXXG”-type are XXXG, XXFG, and XLFG but their percentage may vary depending on the species. Examples of these structures are presented in figure 1.2. Interestingly, the xylose on the non-reducing end of this type of oligosaccharides has so far never been found to be further substituted. In many of those species the β-D-Galp residues often contain O-acetyl substituents (L, F) (Kiefer et al., 1989; Lerouxel et al., 2002; Maruyama et al., 1996; York et al., 1988). The O-acetyl substituent is mainly found on the O-6 position, but has been shown to migrate in aqueous solutions to the O-3 or O-4 position (Pauly et al., 1999a; Kiefer et al., 1990). Table 1.1: One letter code represents unambiguously xyloglucan structures according to Fry et al. (1993) and extended by Hantus et al. (1997) and Ray et al. (2004). Underlined letters indicate that O-acetyl moieties are attached. -[1-β-D-Glcp-4]n- represents polymeric backbone residue. Code contains side chain and backbone block with n=1.
Code Represented structure G -[1-β-D-Glcp-4]n- G Ac(1 6) -[1-β-D-Glcp-4]n- X α-D-Xylp(1 6)-[1-β-D-Glcp-4]n- L β-D-Galp(1 2)-α-D-Xylp(1 6)-[1-β-D-Glcp-4]n- L Ac(1 6)-β-D-Galp(1 2)-α-D-Xylp(1 6)-[1-β-D-Glcp-4]n- F α-L-Fucp(1 2)-β-D-Galp(1 2)-α-D-Xylp(1 6)-[1-β-D-Glcp-4]n- F α-L-Fucp(1 2)-β-D-Galp-(1 2)Ac(1 6)-α-D-Xylp(1 6)-
[1-β-D-Glcp-4]n- J α-L-Galp(1 2)-β-D-Galp(1 2)-α-D-Xylp(1 6)-[1-β-D-Glcp-4]n- A α-D-Xylp(1 6)-[1-β-D-Glcp-4]n-(2 1)-α-L-Araf B α-D-Xylp(1 6)-[1-β-D-Glcp-4]n-(2 1)- β-D-Xylp C α-D-Xylp(1 6)-[1-β-D-Glcp-4]n-(2 1)-β-D-Xylp-(3 1)-α-L-Araf S α-L-Araf(1 2)- α-D-Xylp(1 6)-[1-β-D-Glcp-4]n- S Ac(1 5)-α-L-Araf(1 2)- α-D-Xylp(1 6)-[1-β-D-Glcp-4]n- T β -L-Araf(1 3)- α-L-Araf(1 2)- α-D-Xylp(1 6)-[1-β-D-Glcp-4]n- U β-D-Xylp(1 2)-α-D-Xylp(1 6)-[1-β-D-Glcp-4]n-
The general use of alkaline solutions to extract XyG from cell walls (Joseleau et al., 1992; O’Neill and Selvendran, 1983) removes O-acetates, obscuring their
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presence in most species. Despite the preponderance of XyGOs with cellotetraose backbones, minor amounts of XyGOs with a cellotriose (XXG, Pauly et al., 2001a) or with a cellopentaose backbones has been found in the seeds of the tropical plant Hymenaea coubaril L. (XXXXG; Buckeridge et al., 1997) or in morning glory (Ipomea purpurea, XXGGG) (Hoffman et al., 2005). Such XyG might originate from metabolism in the wall or an altered biosynthetic mechanism and might have different conformational properties (O’Neill and York, 2003). The “XXGG”-type XyG comprise 10–15% of the walls of the dicots Laminales and Solanales (Hoffman et al., 2005; Vincken et al., 1996, 1997; York et al., 1996). It is not only distinct from its block structure but it can also be considered as arabino-XyG (S, T; Tab. 1.1) since no fucosyl residues are attached. In addition to the O-acetyl groups on the β-Galp residue (L), the α-Araf -residue at the O-5 position (S) and even some of the backbone β-Glcp can be O-acetylated (G) (Jia et al., 2005). The O-acetyl groups seem to occur most often in the third position from the non-reducing end, as in XXGG, suggesting that the O-acetyl group might substitute the Xyl residue in the “XXXG”-type xyloglucan. XyGOs such as XXGGG have also been found in some of those species (Hoffman et al., 2005; Jia et al., 2005). “XXGG”-type XyG are also present in graminaceous monocots such as cereals or grasses (Sims et al., 2000) but α-D-Xylp (X) or O-acetyl substituents (G) are the only substituents. Other predominant XyGOs are cellapentaose or cellohexaose units (XXGGG, XXGGGG; Jia et al., 2005). Because of their low degree of substitution, these XyGs are insoluble in water and neutral buffers (Akiyama and Kato, 1982a,b; Kato et al., 1982a,b). Barley (Hordeum vulgare L.) meristematic cells seem to be enriched in XyG before the onset of enhanced β-D-glucan and glucuronoarabinoxylan synthesis during elongation (Carpita, 1996; Carpita et al., 2001; Fincher et al., 2005) and low amounts (10%) of β-D-Galp-containing side chains (L) have been found in endosperm tissue (Shibuya and Misaki 1978). Although cell cultures of Festuca and Zea (Smith and Fry, 1991) were able to integrate α-L-Fucp containing exogenously fed oligosaccharides by transglycosylases into their polymers fucose has not been detected in any naturally occurring XyG prepared from the cell walls of graminaceous plants so far.
Figure 1.2: Example of a typical “XXXG”-type xyloglucan structure which can be typically found in Arabidopsis is depicted (-XLFGXXLG-). The unsubstituted glucosyl residue marked with arrows can be attacked and hydrolysed by various enzymes such as endoglucanases or xyloglucan endotransglycosylase/ hydroylases (XTHs) (see ´Xyloglucan metabolism´).
1.2.2 Properties of xyloglucan
Xyloglucan properties are highly dependent on length, molecular weight and ultrastructure but information of polymersequence is rarely available hindering the analysis of potential microdomains. Microscopic observations revealed that cellulose microfibrils have a spatial distance of about 16-40 nm and that they are connected by tethers (McCann et al., 1990; Satiat-Jeunemaitre et al., 1992). Electron micrographs derived from deep etched cell wall specimen (Itoh and Ogawa, 1997; McCann et al., 1990, 1992) and from artificially assembled composites of bacterial cellulose and xyloglucan showed similar organization of microfibrils and cross linking xyloglucan fibers (Whitney et al., 1995) which in turn suggested that firstly network adhesion might be driven by abiotic processes and secondly that xyloglucan is one cross linking polymer. Xyloglucans are able to coat cellulose microfibrils in muro as it was shown by antibody staining of xyloglucans (Vian et al., 1992). An even stronger labeling could be observed in the cross linking domains (Baba et al., 1994). XyG binds to cellulose in vitro in a pH-dependent manner (Hayashi et al., 1987) suggesting that the polymers are associated by hydrogen bonds. Alkaline solutions are capable of loosening these hydrogen bonds and release xyloglucans from cell wall material (Itoh and Ogawa, 1997; McCann et al., 1990). Alkaline extracted xyloglucans can have a length of about 200-500 nm and reach molecular weights of >300 kDa (McCann et al., 1990, Carpita and McCann, 2000).
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Therefore they have conceivable sizes to span the distance between microfibrils and to adhere in longer, lateral stretches to their surface and form the load-bearing network. Further evidence for a strong association between cellulose and XyG is derived from suspension-cultured tomato cells which were grown in the presence of 2,6-dichlorobenzonitrile (DCB), a herbicide inhibiting cellulose biosynthesis (Shedletzky et al., 1990). In these walls cellulose was virtually absent and thus due to a lack of a scaffold almost all XyG was secreted into the medium (Shedletzky et al., 1992). Xyloglucan can in vitro efficiently bind to cellulose in the presence of arabinogalactans, pectins or even other β-glucans providing another line of evidence that XyG is the major cross linking polymer (Hayashi et al., 1987). The side chain substitution seems to play an important role for solubility and for the binding capacity in vitro since tamarind xyloglucan binds less efficiently to cellulose than unsubstituted cellodextrin (Hayashi et al., 1994a, b). The binding efficiency positively correlates with the degree of backbone polymerisation (Hayashi et al., 1994c). While exclusively xylosylated (GXXXG) xyloglucan has a critical length of at least five backbone residues for aggregation already twelve glucosyl residues are necessary to attach galactosylated XyGOs to cellulose (Vincken et al., 1995). In addition, removal of terminal galactosyl residues strengthened gel formation and allowed self-association of xyloglucan (Reid et al., 1988). As indicated by antibody and extraction studies earlier on (Carpita and Gibeaut, 1993; Hayashi et al., 1994) various xyloglucan domains can be distinguished. Pauly et al. (1999a) analysed depectinated cell wall material of pea to show that approximately one third of XyG can be released by utilizing a xyloglucan-specific endoglucanase (XEG, Pauly et al., 1999b) while additionally one third was subsequent extracted by alkaline solutions. Treatment of remaining material with an endo-glucanase (cellulase) which digests preferentially the amorphous regions of cellulose set the remnant of xyloglucan free. Based on this data, it was postulated that XyG can be divided in an enzyme-accessible (XEG-soluble), cellulose coating (alkali-soluble) and a cellulose encased domain (cellulase-soluble). The enzyme accessible domain is also most likely the part which can be modified by endogenous plant enzymes during cell wall remodeling whereas the interface between cellulose microfibrils and XyG tethers is loosened by expansins.
1.3 Biosynthesis and Metabolism
1.3.1 Biosynthesis
Matrix polysaccharides such as xyloglucan are thought to be synthesized in the Golgi apparatus. For instance, immunolabeling analysis utilizing two antibodies against exclusively xylosylated or fucosylated xyloglucan respectively revealed that the xylosylated core structure (XXXG) is predominantly synthesized in the
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medial and trans Golgi membranes while the later attachment of fucosyl residues occurs in the trans Golgi network (reviewed by Carpita and McCann, 2000). Biochemically, the Golgi localized glycosyltransferases and synthases (Scheible and Pauly, 2004) use activated nucleotide sugars that are present in the cytosol or the Golgi (Zhang and Staehelin, 1992) to form a nascent xyloglucan before the polysaccharide is exported to the extracellular matrix by secretory vesicle shuttling (Staehelin and Moore, 1995). Activated nucleotide sugars are ultimately provided by reactions from products of the Calvin cycle, their activation by the attachment of nucleoside diphosphates (NDPs) and the conversion of NDP-sugars into each other by enzymes of the sugar interconversion pathway. All these steps occur either in the cytoplasm (Amor et al., 1995; Usadel et al., 2004; Seifert et al., 2002) or partially in the Golgi apparatus (Darvill et al., 1980; Haper et al., 2002; Burget et al., 2003; Molhoj et al., 2004). Xyloglucan synthesis requires UDP-D-glucose, UDP-D-galactose, UDP-D-xylose and GDP-L-fucose in most dicots as activated precursor molecules and their synthesis and interconversions are biochemically well established (Dörmann and Benning, 1998; Reiter and Vanzin, 2001; Harper and Bar-Peled, 2002; Kobayashi et al., 2002; Pattathil et al., 2005; Seifert et al., 2004; Seifert, 2004; Barber et al., 2006). Initially, our knowledge of these pathways was obtained from isolated Arabidopsis mutants, whose xyloglucan structure was altered. Reiter et al. (1997) identified murus1 (mur1) plants which exhibited a complete absence of cell wall associated fucose including xyloglucan. Bonin et al. (1997) cloned the MUR1 gene and identified it as a GDP-D-mannose-4,6-dehydratase (GMD2) catalysing the first step in the de novo synthesis of GDP-L-fucose in vitro. Eight allelic mur1 mutants were identified. Plants which lacked or showed reduced MUR1 activity were dwarfed and had a 50 to 200 fold reduction in the content of cell wall associated fucose in shoots while they only displayed 40% reduction in roots (Reiter et al., 1993). However, terminal fucosyl residues located in xyloglucan side chains (XXFG) were replaced by unusual α-L-galactosyl moieties attached to the regular β-D-galactosyl residues (XXJG) (Pauly et al., 2001). Apart from the appropriate nucleotide sugars the biosynthesis of xyloglucan requires at least five enzymatic reactions and thus glycosyltransferases (GTs) for the XXXG-type xyloglucan of Arabidopsis, namely a backbone building beta-D-glucan synthase, an alpha-D-xylosyltransferase, a beta-D-galactosyltansferase, and an alpha-L-fucosyltransferase as well as an acetyl-transferase to modify the galactosyl moiety. Based on structural similarities of the of cellulose and hemicellulose glucan chains it was assumed that glucan synthases synthesizing XyG backbone could be found in one of the eight Cellulose Synthase Like (CSLA to CSLH) gene families (Richmond and Somerville, 2000; Hazen et al., 2002). Indeed, a recent report by Cocuron et al., (2007) showed that a member of the CSLC family,
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AtCSLC4, which was heterologously expressed in Pichia pastoris encodes a protein synthesising β-1,4-glucans. Two members of a family of xylosyltransferases (AtXTs, now renamed to AtXXT, Cavalier et al., 2008) has been shown to attach xylosyl residues to β-1,4-glucose oligomers and thereby producing xyloglucan oligosaccharides (XyGOs) (Faik et al., 2002; Cavalier and Keegstra, 2006). Initially Faik and co-workers (2002) identified seven Arabidopsis genes sharing sequence similarities to an α-xylosyltransferase derived from microsomal pea preparations. These genes encoding closely related glycosyltransferases were annotated as galactosyltransferases and grouped into the CAZy glycosyl transferase (GT) family 34 (www.cazy.org/fam/GT34.html). AtXXT1 and AtGT2 (now AtXXT2, Cavalier and Keegstra, 2006; Cavalier et al., 2008) were grouped into one class since they shared 83% identical amino acids and were capable of transferring xylosyl moieties to glucose oligomers. While AtGT3 to AtGT5 had 75-80% identical amino acids in common and clustered in a second group AtGT6 and AtGT7 built a third group with 75% identical amino acids. AtXXT1 and AtXXT2 were proven to use cello-pentaose and cello-hexaose as substrates for xyloglucan synthesis with a more efficient usage of cello-hexaose. Cavalier and Keegstra (2006) could show that both enzymes used preferentially the fourth glucosyl residue from the reducing end of the substrate molecule as an attachment site for xylosylation in vitro and thereby producing GXGGG and GGXGGG respectively. In addition a second xylosyl residue was added when the amount of bare glucan chains was significantly reduced during the enzyme reactions resulting in GXXGG and GGXXGG. A minor proportion of tri-xylosylated GXXXGG could also be detected and led to the assumption that this molecule was rather synthesized by the subsequent addition of xylosyl residues using GXGGGG as primer molecule than an alternating enzyme mechanism (GGXGGG GGXXGG GXXXGG). Both enzymes showed thereby redundancy in the synthesis of xyloglucan oligosaccharides in vitro. Thus, if additional GTs are necessary to add the third xylosyl residue next to the non-reducing end in vivo is questionable. Until now none of the other enzyme members of family GT34 (AtGT3 to AtGT7) have been biochemically proven to transfer xylosyl residues to XyG but a recently published report about knockout (K.O.) plants of AtGT5 (now XXT5, Zabotina et al., 2008) demonstrated XXT5 to be a putative xylosyltransferase located in the Golgi apparatus and showed Atxxt5 plants displaying a root hair phenotype while being xylose deficient. The generation of double K.O. plants revealed that AtXXT1 and AtXXT2 indeed act on xyloglucan in planta since these plants contain hardly any xyloglucan (Cavalier et al., 2008), Interestingly, co-expression of Arabidopsis xyloglucan specific α-xylosyltransferase 1 (AtXXT1) and AtCSLC4 in Pichia resulted in production of larger β-1,4-linked glucan chains than synthesised by clones exclusively expressing CSLC4 (Cocuron et al., 2007). This suggested either a direct or an indirect interaction of the AtCSLC4 glucan synthase. Previously
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Hayashi et al. (1984) observed an interdependent effect of the UDP-glucose/UDP-xylose ratio on the generation of different building blocks in membrane preparations of soybeans. They could show that a dominant concentration of UDP-glucose in vitro promoted the formation of the pentasaccharide XXG whereas when the ratio was reversed and UDP-xylose was more abundant predominately the heptasaccharide (XXXG) were produced. Already in 1981 Hayashi and Matsuda (Hayashi and Matsuda, 1981a) observed that xyloglucan chain elongation required a concurrent addition of xylose and glucose units in vitro. Those findings have made them formulate the hypothesis that the XyG glucansynthase forms a complex or is at least closely associated with the xylosyltransferase initiating side chain substitution. β-D-galactose is the second moiety on a XyG side chain. Xyloglucan contains two galactose residues on position two and three counted from the reducing end. Therefore it seems likely that at least two different galactosyltransferases with varying substrate specificities are necessary. Earlier identified murus3 (mur3) mutants which showed a significant reduction in cell wall associated fucose (Reiter et al., 1997) were cloned and further characterized by Madson et al. (2003). MUR3 encoded by the locus At2g20370 is a residue-specific GT which adds galactose to the first xylosyl residue next to the reducing end (XXLG) and provides the basis for the attachment of fucose which explains its reduction in cell walls fucose (50%, Reiter et al., 1997) and galactose (50%, Madson et al., 2003) content in the mur3 mutant. Biochemical and phenotypical observations of mur3 shoots revealed that xyloglucan structure was strongly impaired and virtually no fucosylated or double galactosylated XyGOs were present (Madson et al., 2003). All XXLG-type structures were instead replaced by XLXG which accounts for 45% of the XyGOs in mur3. In contrast, these XyGOs occurring only in minor proportions in wild type (3%). GUS expression and RT-PCR analysis showed that MUR3 is expressed in all major organs and the consistent biochemical phenotype indicated that no genetic redundancy could be observed. mur3 plants showed besides collapsed trichome papillae no obvious phenotype and the tensile strength of their inflorescence stems was wild type-like when grown in continuous light. Peña et al. (2004) observed swelling of epidermal cells close to hypocotyls hook and suggested that impaired xyloglucan structure reduced the tensile strength of etiolated seedlings to 40% of wild type level. Based on sequence homologies MUR3 resided in CAZy family GT47 (Henrisatt et al., 2001). Family GT47 comprises 39 genes in Arabidopsis which cluster in four clades (A to D, Zhong and Ye, 2003). MUR3 together with ten other GTs (GT11 to GT20) were grouped into clade A and promoter:GUS fusions of them revealed that AtGT18 (At5g62220) was expressed throughout whole seedlings (root, hypocotyl, cotyledons, first true leaves) and in inflorescences (stem, flowers) resembling MUR3 expression pattern. Transgenic plants containing promoter-Gus Atgt13 (At2g32740) fusion constructs displayed a similar pattern but showed a gradient of staining in roots including highest expression in older
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parts and no staining at all in the fast growing root tip (Li et al., 2004). T-DNA mutants of Atgt13 and Atgt18 displayed a significant reduction of about 10-14% in cell walls galactose content respectively, and thus made them considerable candidates for glycosyltransferases introducing the second galactosyl residue (XLXG) into xyloglucan. In Arabidopsis exclusively the first side chain next to the reducing end can terminate with a fucosyl residue (XXFG) whereas to date only the undecasaccharide (XFFG) isolated from sycamore cells showed fucosylation on the second side chain next to the reducing end (Hisamatsu et al., 1991). Pulse-chase experiments revealed that side chain extension with galactose and fucose appears on a preformed xylosylated XyG backbone and that it takes place in the Golgi stacks since no further xyloglucan biosynthesis could be detected in Golgi vesicles (Camirand and MacLachlan, 1986; Camirand et al., 1987). Therefore it is conceivable that the fucose moiety is added by a unique α-1,2-fucosyl transferase in Arabidopsis. Indeed, murus2 (mur2) mutants showing a decrease of fucosylation of approximately 50% in above ground cell wall polymers. Similar to mur3 no obvious phenotypes in the growth habit and no change in tensile strength of inflorescence stems could be observed in mur2 plants (Vanzin et al., 2002). 44% of the XyGOs appeared to be double galactosylated (XLLG) in xyloglucan derived from mur2 shoots (Vanzin et al., 2002) or derived from leaves of etiolated seedlings (Peña et al., 2004) whereas XLLG accounted for only 4% of total XyGOs extracted from wild type. Cloning of the gene revealed that MUR2 encodes an α-L-fucosyltransferase acting on xyloglucan and a more detailed cell wall analysis showed that less than 2% of xyloglucan remained fucosylated (Vanzin et al., 2002). Earlier on Perrin et al. (1999) cloned the same gene (AtFUT1) following a different approach. Perrin and co-workers isolated alpha-fucosyltransferase activity from microsomal preparation of pea (PsFuT1, Faik et al., 2000) and identified based on deduced amino acid sequences an ortholog in Arabidopsis which were subsequently expressed in mammalian COS cells and its function was biochemically proven. AtFUT1/MUR2 and PsFUT1 belong to family GT37 and share 62% sequence homology (Henrisatt and Davies, 2000). Additional nine members (AtFUT2 to AtFUT19) of fucosyltransferase family GT37 have been identified in Arabidopsis but all of them share less sequence homology to AtFUT1 than AtFUT1 to PsFUT1. Furthermore, none of them was active against xyloglucan (Sarria et al., 2001). In Arabidopsis O-acetylation of XyG occurs at the galactosyl moiety of the first side chain close to the reducing end (Pauly, 1999). While mainly the hydroxyl group at C6 is converted into an ester bond the acetate might migrate under certain conditions in vitro forming linkages with the hydroxyl group of C3 and C4 (Pauly, 1999). Microsomal preparations of suspension cultured potato stem cells were capable of transferring acetates to cell wall polymers by feeding 14C-radiolabeled acetyl-CoA (Pauly and Scheller, 2000) suggesting that O-
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acetylation takes place in the Golgi and that acetyl-CoA is the donor. There are also two lines of evidence which inferred that fucosylation is correlated with O-acetylation. First, although overexpression of AtFUT1 did not affect XyG fucosylation it did enhance O-acetylation of XyG (Perrin et al., 2003). In addition, seed storage XyG e.g. from tamarind is neither fucosylated nor O-acetylated also indicating that one or both side chain substitutions might be a steric hindrance for tight sphere packing of storage carbohydrates. Interestingly when fucosylation was impaired, such as in light grown mur1 suspension cultured cells, O-acetylation of XXLG was enhanced in comparison to WT and even di-acetylated oligosaccharides (XXLG + 2Ac, XXJG + 2Ac) could be observed (Pauly et al., 2001a). This suggested that over-fucosylation may promote the attachment of O-acetyl moieties but the lack of it does not prevent their generation especially since in mur1 cells the unusual α-L-Galp residue can occupy the α-L-Fucp position (XXJG) in the XyG side chain. After XyG synthesis has ceased in the Golgi apparatus (Hayashi and Matsuda, 1981a; Hayashi et al., 1984; Camirand and MacLachlan, 1986; Camirand et al., 1987; Tamura et al., 2005) the cell wall building blocks are secreted (Robinson et al., 1976; Taiz et al., 1983; Kawasaki, 1981) probably by actin guided (Boevink et al., 1998; Nebenfuhr et al., 1999; Nebenfuhr and Staehelin, 2001; Tamura et al., 2005) vesicle trafficking to the apoplast. Analysis of katamari mutants (kam, Tamura et al., 2005) impaired in an N-terminal cytosolic domain of the xyloglucan galactosyltransferase MUR3/KAM indicated an interaction of that particular Golgi located enzyme and actin filaments. In the extracellular matrix xyloglucans adsorb to the surface of cellulose microfibrils in a self assembly process and interact due to a complex contribution of van der Waals, dispersion and electrostatic forces as well as hydrogen bonds. These interactions induce loosening of cellulose crystallinity at the microfibril´s surface (Hanus and Mazeau, 2006). The free not linked parts can be accessed by XyG modifying enzymes such as XTHs and other glycosidases which may alter binding properties. The alkaline sensitive interface between cellulose and XyG is considered to be the point of expansin action. Other domains may even have been entrapped during cellulose microfibril formation and probably do probably not resemble the physiologically active part of the network (Pauly et al., 1999b). After deposition the cell wall is not just an inert network after polysaccharide deposition; instead it is a physiologically active compartment whose polymer structures such as xyloglucans undergo dynamic, controlled processes of remodelling and modifications in a way that the wall can resist the high turgor pressure and yet allows flexibility during cell elongation. Each individual step from synthesising a fully functional cell wall metabolising enzyme, transporting it to the right location at the right time and making it to proceed its reaction in a synergistic manner requires a highly complex network of interactions but also
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provides the cell with regulatory elements which probably in most of the following cases are far more complex as described here.
Xyloglucan endo-transgylcosylase/hydrolases are capable of hydrolysing a xyloglucan chain and re-joining it to (another) xyloglucan acceptor molecule. This reaction can be imagined as a micro-grafting process (Fry et al., 1992) of polymers which allows extension of xyloglucan chains tethering cellulose microfibrils without destabilisation of the whole network. In Arabidopsis 33 XTH genes have been identified yet and their respective proteins were classified in CAZy family GH16 (Henrisatt and Davies, 1997). XTHs can hydrolyse hemicellulosic backbones if an un-substituted glucose residue in the glucan chain is exposed (Fig. 1.2). This reaction seems to happen regardless of side chain substitution patterns and is the so called xyloglucan endohydrolase function (XEH) of XTHs. It is noteworthy that endo-glucanases (family GH9, www.CAZy.org/fam/GH9.html) produce similar reaction products but that mechanism and enzyme topology is clearly distinct from those of XTHs and that XET activity of XTHs probably evolved from an ancestral XTH to accomplish specialized functions during cell wall remodelling events such as fruit ripening, seed germination or tissue elongation (Baumann et al., 2007). The newly formed reducing terminus of the hydrolysed xyloglucan chain is then transferred to the non-reducing end of an acceptor molecule (Fry et al., 1992), referred to as endo-transglycosylation reaction (XET, Rose et al., 2002). In theory, three reactions depending on the molecular size of donor and acceptor molecule can occur. First, donor and acceptor are both polymers (Smith and Fry, 1991), secondly the donor is a polymer and the acceptor is an oligomer (Nishitani and Masuda, 1982; Talbott and Ray, 1992) or at last both acceptor and donor molecules are XyGOs (Fanutti et al., 1993, Fanutti et al., 1996). The XEH type of reaction can also be envisioned as the polymer-oligomer variant in which water simply serves as acceptor molecule (Rose et al., 2002). Within the family diverse substrate specificities, physiological conditions and transcript profiles have been observed coordinating XTH activity. Activity studies of pea stems confirmed various XyGOs as suitable acceptor molecules but with descending affinity (XLLG>XXXG>XXFG>XXG) suggesting that xyloglucan side chain motifs have an influence on substrate binding (Fry et al., 1992a, Lorence and Fry, 1993). As apoplastic proteins most of the XTHs have a narrow pH optimum typically between 5 and 6 (Campell and Braam, 1999a). The pH of apoplastic solutions derived from azuki bean epicotyls was measured in a range of pH 6.2 to 6.6 (Nishitani and Tominaga, 1991). Since auxin mediated cell wall acidification is able to quickly reduce the pH of about one unit (Jacobs and Ray, 1976) XET activity might be enhanced and elongation promoted. While AtXTH2 to AtXHT7 were expressed in various organs most of the other XTHs showed an
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organ specific pattern such as AtXTH17 to AtXTH20 were exclusively expressed in roots (Yokohama and Nishitani, 2001b). Even though the latter were specific transcribed in roots, tissues of expression were substantially different, such as e.g. AtXTH19 was preferentially expressed in the root apex and the elongating zone while AtXTH20 was mainly expressed in the basal part and in the vascular tissues of the roots (Vissenberg et al., 2005). The variance of expression profiles can also be exemplified by comparison of AtXTH22 and AtXTH23 whose transcript levels are influenced by a variety of stimuli. AtXTH22 was excited by mechanical stimuli (Braam et al., 1992), brassinosteroids (Xu et al., 1995 and 1996) and auxin (Goda et al., 2004) whereas expression of AtXTH23 was induced by auxin (Goda et al., 2004), brassinolide and gibberellic acid (Yokohama and Nishitani, 2001). Thus, variations in XTH enzyme complements with their specific functions may modulate plant growth and morphogenesis during developmental or due to external biotic and abiotic factors. Already early on Nishitani (1997) speculated that some XTHs are involved in auxin dependent elongation processes while other members mediate molecular grafting of cell walls which are no longer expanding (Nishitani and Vissenberg, 2006). In rice 29 open reading frames of XTHs with differential expression patterns have been identified (Yokoyama et al., 2004) and some of them exclusively showed expression in elongating tissues indicating a strong correlation between XET activity and expansion. However, xyloglucan is less abundant in rice than in dicots and is believed to play a minor role as cross linking polymer (Yokoyama et al., 2004; Yokoyama and Nishitani, 2004). Thus, in Poales at least some XTHs may act on other cross linking matrix polysaccharides such as arabinoxylan. This idea might be emphasised by the finding that the genome of poplar, as a representative of wooden species, encompasses 41 XTHs and 11 of those grouped into class III showing both XET and XEH activity while the rest clustered in class I and II and exclusively exhibited XET activity (Geisler-Lee et al., 2006). In poplar 4-O-methylglucuronoxylan is a major hemicellulose which is deposited in primary as well in secondary cell walls during wood formation (Mellerowicz et al., 2001) and can be considered as one reasonable substrate for XTH activity. But it has to be noted that Geisler-Lee et al. (2006) found expression of AtXXT1 orthologs during wood formation suggesting that xyloglucan biosynthesis may have occurred during secondary wall deposition. In addition, to date all analysed XTHs which exhibiting XET function were shown to use XyGOs with a backbone of ≥Glc4 as acceptor molecules (Rose et al., 2004). Other endo-β-1,4-glucanases belong to CAZy family GH5, GH9 and GH10 (www.CAZy.org) and so far 13, 25 and 12 genes have been identified in Arabidopsis in these three families respectively. While family GH5 encodes mainly endo-β-1,4-mannanses and GH10 preferentially endo-β-1,4-xylanases (for review Minic, 2008) family GH9 represents endo-β-1,4-glucanases/cellulases and 22 of them are predicted to be located at the plasma
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membrane or to be secreted to the apoplast (Minic and Jouanin, 2006). Functions could be assigned to the KORRIGAN (kor, Nicol et al., 1998; Robert et al., 2005) and CELLULASE (CEL, Yung et al., 1999; Shani et al., 2006) sub-groups which rather suggested them to be involved in cellulose synthesis and xylem development respectively than in xyloglucan metabolism. None of the genes clustering in these families has been proven to act on xyloglucan yet.
1.3.3 Xyloglucan mobilisation/degradation
Besides maintaining structural functions xyloglucan can also serve as storage (amyloid) carbohydrate such as in the cotelydons of the tropical plants nasturtium (Tropaeolum majus L.; Edwards et al., 1985), tamarind (Tamarindus indica; Kooiman, 1957) or Copaifera langsdorfii (Buckeridge et al., 1992). The mobilisation of the sugars requires degradation of the polymer and all aforementioned plants contain only galacto-xyloglucan lacking terminal fucosyl residues and therefore a complete breakdown of the XyG can be achieved by subsequent action of β-D-galactosidase, α-D-xylosidase, β-D-glucosidase and endo-β-1,4-D-glucanase. The ratio of XXXG, XLXG, XXLG and XLLG building blocks varies in terms of species, developmental stage and even origin of collected plant material (Buckeridge et al., 1992) but in all of them XLLG is the most abundant XyGO inferring a central role for terminal galactose in compact packing of amyloid xyloglucans. Furthermore, β-galactosidase activity isolated form nasturtium preferred intact polymeric XyG as substrate (Edwards et al., 1988) to cleave the galactosyl residue near to the non-reducing end [(XLLG)n XXLG(XLLG)n-1]. Fanutti et al. (1991) could separate α-xylosidase activity from germinating nasturtium seeds and reported a binding preference for XXLG (Km 0.32 mM) over XXXG (Km 0.62 mM) whereas the capacity of turn-over was half as big for XXLG as for XXXG. They could also demonstrate that only oligomers and not polymers were hydrolysed suggesting that additional backbone degrading endo-β-1,4-glucanase activity was necessary prior to xylose mobilisation. Edwards and colleagues (Edwards et al., 1986) separated endo-β-1,4-glucanase activity from nasturtium cotyledons to homogeneity and found it to be preferentially active against amyloid galacto-xyloglucan and in-turn suggested its participation in sugar mobilisation. Later on Fanutti et al. (1996) demonstrated that this enzyme contained both XET and XEH activity depending on the availability of substrates and their corresponding fine structures which infers a potential role not only in carbohydrate mobilisation but also in the biosynthesis of tight packed amyloid xyloglucan. β-D-glucosidase activity is required as last step for complete breakdown of xyloglucan and it was isolated from nasturtium in 1998 by Crombie and co-workers (Crombie et al., 1998). According to its protein sequence the enzyme was grouped into glycoside hydrolase family 3 (Henrisatt, 1991, www.CAZy.org/fam/GH3). In their study they showed that this enzyme had exo-activity against xyloglucan
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oligosaccharides removing the glucose moiety from the non reducing end of the backbone. Galactosyl residues next to the cleavage site (e.g. GLXG) prevented hydrolysis while xylosylation (GXLG) had no effect suggesting tightly coordinated actions of endo-β-1,4-glucanase, β-D-galactosidase, α-D-xylosidase and β-D-glucosidase during carbohydrate mobilisation. A synergistic and controlled action of a set of xyloglucan degrading enzymes is also required during growth processes such expansion in which the cell wall and especially the tethering xyloglucan polymers needed to be remodelled. Already in 1989 O´Neill et al. purified a protein from pea seedlings exhibiting α-D-xylosidase activity (O´Neill et al., 1989). The α-D-xylosyl residue was cleaved from the non-reducing terminus under acidic conditions (optimum pH 4.9-5.1). They could show that the substrate specificity of the enzyme was more pronounced for XXFG than for XXG or XXXG but the release of the xylosyl residue was catalysed with nearly the same velocity for all tested XyGOs. In Arabidopsis three genes have been identified sharing sequence homologies representative for family GH31 (reviewed in Minic and Jouanin, 2006) which encloses several α-xylosidases. Sampedro et al. (2001) cloned one of the putative α-D-xylosidase (AtXYL1) and could correlate an increased level of expression and higher enzyme activity to younger leaves indicating a developmentally regulated level of transcription and an involvement in expansion. Lorence and Fry (1993) reported that xylosylation at the non-reducing terminus of xyloglucan is essential for transglycosylation (XET) activity extracted from pea and bean leaves. Therefore, it was inferred that α-xylosidase action may deprive suitable XTH substrates and thus regulate elongation processes in concert with XTHs (Fry, 1995). In contrast to the amyloid system, galactosidase activity isolated from apoplastic fluids of Arabidopsis seedlings preferred to digest the non-reducing galactose moiety from fucogalacto-xyloglucan oligosaccharides (XXLG XXXG, XLFG XXFG) (Iglesias et al., 2006). Taken together, these data resemble natural substrate specificities of plant β-galactosidases involved in different physiological processes such mobilisation of amyloid xyloglucan and the derivatisation of structural cell wall xyloglucan during elongation. To date, only few reports have dealt with the identification of an α-1,2-L-fucosidase activity and their corresponding genes in plants. Augur et al. (1993) isolated fucosidase activity from pea seedlings and claimed in 1995 (Augur et al., 1995) that it was related to a 20 kDa protein encoded by the PsFUC1 gene but Tarragó and co-workers could not gain any activity from heterologously expressed PsFUC1 which was expressed in the different systems E.coli, insect cells and Arabidopsis (Tarragó et al., 2003). Additionally overexpression in Arabidopsis did not lead to any enhanced fucosidase activity. They separated a 55 kDa protein from pea seedlings co-eluting with FUC1 which showed fucosidase activity and also substantial sequence homology to FUC1. However, a similar pair of proteins has been identified and cloned in Arabidopsis (de la Torre et al., 2002). While
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heterologous AtFUC1 was only active against 2´-fucosyl-lactinol, the identified AtFXG1 cleaved specifically terminal fucose from XXFG. All activities necessary to completely catabolise the side chains of structural XyGOs (α-1,2-fucosidase, β-1,2-galactosidase, α-1,6-xylosidase and β-1,4-glucosidase) have been detected in the apoplastic fluids of Arabidopsis seedlings (Iglesias et al., 2006) and some of the underlying genes have been identified (XYL1, At1g68569, CAZy familiy GH31; FXG1, At1g67830, CAZy family GH0). In addition, proteomic and molecular approaches supported the idea that the cell wall proteome undergoes various post-translational modifications such as phosphorylation and glycosylation and that it may be regulated by the processing of proteases and GHs (Minic, 2008).
1.4 Roles of xyloglucan in plant growth and development
XyG is an important structural element of the wall and it is conceivable that its biosynthesis and metabolism is controlled during cells life to maintain its functional role. To ensure correct cell wall remodelling and to avoid failure sensing of cell wall derived oligosaccharides allow the plant cell to monitor and to react on certain biotic or abiotic circumstances such as pathogen attack. XXFG was shown to inhibit endogenous (Warneck and Seitz, 1993) and auxin promoted (York et al., 1984) growth of pea stems at nanomolar concentrations. Feeding pea stem sections with polymeric xyloglucan did suppress growth while feeding the oligosaccharide XXFG at millimolar concentrations promoted elongation (Takeda et al., 2002). Furthermore, attachment of a neighbouring galactosyl residue (XLFG) abolished the growth promoting effect (McDougall and Fry, 1989a). Thus, it might be speculated that α-fucosidase and/or subsequent degradation of XyGOs produced biologically active carbohydrate signaling molecules (oligosccharins), such as XLLG and XXXG which also displayed positive growth effects in millimolar concentrations. One possibility is that simply more effective XET acceptor molecules were present (XLLG>XXXG>XXFG, as mentioned in the XTH paragraph, Fry et al., 1992, Lorence and Fry, 1993) which promoted growth. Additional evidence to favour a “XET-” over an “oligosaccharin- theory” in terms of expansion is given by the fact that mur2 and mur3 mutants have altered XyGO structures especially lacking terminal fucosyl residues without showing an effect in growth performance under laboratory conditions (Reiter et al., 1997). O-acetylation might influence hydrolytic degradation properties of xyloglucan side chains or might play a role in the fine tuning of acceptor molecules involved in transglycosylation events. But no experimental data has been obtained in vivo or in vitro. Additionally, XyGOs have shown oligosaccharin related effects which are not consistent with the “XET”-theory. For instance xyloglucan derived
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oligosaccharides can alter plant morphogenesis. Small fragments like the pentasaccharide FG are able to induce adventitious roots in the absence of auxin (2,4-dichlorophenoxyacetic acid, 2,4-D) using cultured wheat embryos as model system. In contrast, the co-presence of auxin and FG enhanced callus formation suggesting a hormone-like action (Pavlova et al., 1992). Cell wall deconstruction by pathogenic enzymes releases XyGOs which in turn can act as elicitors and induce the accumulation of antimicrobial phytoalexins (Pavlova et al., 1996). Expansins are a group of cell wall proteins which are able to induce flexibility without destroying cell wall integrity. For instance heat inactivated cell walls when incubated with expansins showed enhanced relaxation after applying a constant load which completely restored acid-induced wall extension (McQueen-Mason et al., 1992). However, expansins do not hydrolytically cleave any wall polymer (Yuan et al., 2001). Interestingly, Yuan et al. (2001) identified a fungal β-1,4-endoglucanase from Trichoderma reesi (Cel12A, GH12, Henrissat et al., 1998, www.cazy.org/index.html) which were able to increase cell wall plasticity like expansins but in a mechanistically different manner. Cel12A also mimicked features of auxin-induced wall relaxation and putatively hydrolysed β-1,4-glucan linkages in the xyloglucan backbone after a lag phase. In contrast, treatments with other endoglucanases led to breakage of tissues over time whereas relaxation in the presence of expansin were apparent in less than a minute and maintained wall integrity even after prolonged incubation times. Pectins and other hemicelluloses, like glucomannans, were tested to be cooperation partners in vitro but only artificial composites of bacterial cellulose and xyloglucans were found to have increased extension properties after treatment with expansins (Whitney et al., 2000) suggesting them to be the most likely natural substrate. Three lines of evidence supported the H-bond character of the tight XyG-cellulose attachment (reviewed by Cosgrove, 2005). First, lower pH promoted expansin induced expansion in inactivated cell walls while neutral or mild alkaline conditions did not show any effect which makes them suitable mediators for the acid growth. Second, urea enhanced progressively expansin catalysed H-bond disruption whereas thirdly extension was slowed down in the presence of D2O in comparison to H2O (McQueen-Mason and Cosgrove, 1994). Thus, it is assumed that expansins work at the alkaline extractable domain of xyloglucan by loosening progressively the H-bonds at the xyloglucan-cellulose interface and thereby allow a process during expansion which can be envisioned as ´polymer creeping´ (Yuan et al., 2001; Marga et al., 2005). Interestingly, overexpression studies revealed controversial data. For instance, overexpression of an endogenous expansin in Arabidopsis led to increased growth of petioles and leaves (Cho and Cosgrove, 2000) while enhanced expression of a tomato expansin resulted in decreased fruit size (Brummell et al., 1999). In Arabidopsis 38 open reading frames have been detected which encode expansin-like proteins (Li et al., 2003) and the spatial
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and temporal appearance of members of that gene family seem to be correlated to meristematic tissues (Reinhardt et al., 1998; Cho and Cosgrove, 2002; Sampedro and Cosgrove, 2005; http://www.bio.psu.edu/expansins).
1.5 Isolation and structural analysis of xyloglucan
The cell wall is a highly complex and dynamic network consisting of 14 different monosaccharides, three classes of carbohydrates (cellulose, hemicelluloses, pectins), hosting probably more than 20-30 different types of linkages and various structural and enzymatic (glyco-)proteins. Microscopy is a powerful tool to understand cytological scenarios but its help is limited by resolution and in revealing molecular interactions. Even though electron microscopic observations of in vitro cellulose-hemicellulose interactions indicated that xyloglucan can tether cellulose microfibrils by forming networks similarly looking like those observed in muro (Carpita and McCann, 2000) a direct evidence for xyloglucan being the cross linking polymer is still missing. A detailed analysis of cell wall polymers often requires various extraction, purification and enrichment steps which in turn may alter the fine structure of a class of polymer but surely disorders the macromolecular organisation which was present in situ. Laser-microdissection allows sampling of single cells or even cell wall domains (e.g. outer wall of epiderimal cells) but is very labour intensive. Therefore the collected material used is often a mixture of specialised cell walls derived from a number of different cell types. Mostly alcohol insoluble residue (AIR) is a starting material to extract a certain polymer out of the cell wall. Chelators such as CDTA can be used to break Ca2+ bridges existing between homogalacturonan (HG) strands and solubilise a part of the pectic network. The release of hemicelluloses which adhere to cellulose microfibrils requires harsher alkali (1M, 4M KOH or NaOH) conditions to loosen H-bonds. A drawback of using alkali is the removal of functional groups from hemicelluloses, such as acetyl-esters, and thereby preventing the analysis of the ´naturally´ occurring structure of these polymers. Strong acids can be utilised to break crystalline parts of cellulose fibrils by hydrolysing glycosidic linkages. Purified and/or heterologously expressed hydrolytic enzymes derived from a number of variable sources (e.g. microorganism or plants, www.CAZy.org) are a powerful tool to decompose polymers in building blocks as they are present in the wall without perturbing functional groups. A vast number of enzymes specific for certain domains of polymers are commercially available [e.g. endo-polygalacturonases, cellulases or xyloglucan specific endo-glucanase (XEG; EC 3.2.1.151)] and used to digest the cell walls prior to purification. Once polymers or oligomers are extracted they can be separated and analysed by chromatographic means. Size exclusion chromatography (SEC) for example allows the separation of polymers
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according to their molecular mass (Bauer et al., 1973; Hayashi et al., 1984) and detection can be achieved by refractive index. Furthermore, High Performance Anion Exchange Chromatography (HPAEC) coupled with pulsed amperometric detection (PAD) can be utilised for separation and analysis of xyloglucan oligomers. HPAEC combines the power of Affinity and the speed of High Performance Liquid Chromatography (HPLC) but for PA-detection carbohydrates need to be eluted under alkali conditions which again impair the analysis of functional O-acetyl-groups (Hilz et al., 2006). An advantage of HPAEC-PAD is its high reproducibility in quantification of XyGOs and the potential to separate structural isomers such as XXLG and XLXG (Lerouxel et al., 2002). Polysaccharide analysis using carbohydrate gel electrophoresis (PACE) can be used to analyse sugar mono- and oligomers. Another method, the capillary electrophoresis (CE), is an advanced and miniaturised technique which can be facilitated in an automated and high through put fashion (Immerzeel and Pauly, 2006). In both PACE and CE detection is achieved by labeling the sample with fluorophores prior to analysis additionally allowing an absolute quantification. Although some of these chromatographic methods are very useful purification techniques in terms of carbohydrate analytics all of them share the disadvantage that they require suitable and pure standards. Analysis facilitating traditional chromatographic methods are sometimes time consuming since a HPLC separation can take up to 2 hours. Enzymatic hydrolysis of polymers coupled with MALDI-TOF mass spectrometry made it possible to acquire oligosaccharide mass profiles within minutes and thereby can be applied in a high throughput fashion. It takes advantage of the regularity with which plants synthesise their cell wall carbohydrates and the specificity of available enzymes. It is suitable to access semiquantitative differences in a sample versus a control data set (Lerouxel et al., 2002) and only low amounts of cell wall material are necessary to determine structural differences. Due to its mild extraction and ionisation techniques functional groups such as O-acetyl-esters can be analysed. A drawback is that structural isomers can not be distinguished (XXLG/XLXG) and oligosaccharides with a degree of polymerisation <3-5 can not always be properly assigned due to an increased noise ratio in lower mass ranges. Additionally statistical means are always required to cement fingerprints derived from spectroscopic techniques (Möller et al., 2006). Fragmentation of analytes using tandem mass spectrometry can be used to determine the remaining mass and gives insights about the substituents that was fragmented off. Nuclear magnetic resonance (NMR) spectroscopy can not only be applied to fingerprint but also to quantify certain compounds. Furthermore it is a powerful tool to analyse unknown components since it provides detailed structural information (Ralph et al., 2001; Bootten et al., 2003; Perrin et al., 2003) and is a non-destructive technique. Arising problems using NMR can be its low throughput and its insensitivity since substantial amounts of up to 100 mg are necessary to analyse complex samples. Since for the usage of NMR no further
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sample preparation is required the intact material can be applied to subsequent traditional sugar analytics. Qualitative and quantitative monosaccharide composition can be achieved facilitating gas chromatography and mass detection (GC-MS) analysis. In order to make inert sugar molecules volatile derivatisation steps are required. Therefore carbohydrate polymers are hydrolysed into monosaccharides which are subsequently reduced and converted into their corresponding volatile alditol acetates (AA) (Albersheim et al. 1967, Mankarios et al. 1979). Information about linkages within a polymer can be gained if the sample has been partially methylated prior to acid hydrolysis. Subsequent conversion to partially methylated AA (PMAA) similar to AA derivatisation allows GC-MS separation and analysis. In the recent years the production of a variety of antibodies (AB; Plant Probes Leeds, UK; CarboSource, Athens, USA) and carbohydrate binding modules (CBMs; McCartney et al., 2006) which recognize specific sugar epitopes made it feasible to develop carbohydrate arrays (Willats et al., 2002) suitable to screen cell wall alterations in a high throughput fashion (Møller et al., 2007). Utilization of ABs and CBMs further allows the study of carbohydrate distribution in muro and to gain insights into the cell wall itself. But the availability of pure standards and the lack of conformational information of the natural occurring epitopes are often the limitations for a detailed characterization of the antibodies which has to be solved in the future.
1.6 Biotechnological applications of cell walls
Plant cell walls fulfill various biological functions during plant live but besides that they are used by mankind for a variety of different applications. Cell walls mainly account for dietary fibers in human nutrition (McNeil et al., 1984) and some of the wall integrated compounds are believed to protect against colon cancer or diabetes (Carpita and McCann, 2000). Substances within oat and barley brans might lower serum cholesterol (Terpstra et al., 2002) and prevent consumers from coronary heart disease. XyG walls are remodeled during fruit ripening (Brummell et al., 1999, for review see Brummell, 2006) and thus their composition has an impact on durability of fruits and vegetables as well as on shelf life and processing ability of plant based products (Bruening and Lyons, 2000). Furthermore, packaging material of sustainable “plastics” is a field of future oriented products. Therefore cell wall derived sugars can be converted into robust but biodegradable polymers or ingredients for glues (Lapasin and Pricl, 1995; Nawrath et al., 1994). Plant cell walls are good filling, packaging and insulating material which protects e.g. housing against heat and cold (Lapasin and Pricl 1995). Plant fibres, essentially consisting of cellulose microfibrils coated with hemicelluloses, and can be manufactured to ropes or textiles and in turn they are used to tighten and wrap bodies as they were
1 Introduction
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designed for by nature. World wide cotton production is estimated of about 20 Mt per year (http://www.plantcultures.org) with China being the largest producer (~6.3 Mt in 2005) but probably also one of the biggest consumers of cotton fibres (www.wikipedia.org). While the cotton industry is led mostly by China, central Asian republics (India, Pakistan) and the USA the pulp and paper business is dominated by North America, Northern Europe and Japan. Pulp and paper production is accompanied with high energy consumption and greatly relies on the extractability of cell wall polymers. To reduce increasing energy costs these heavy industries generate substantial amounts of their demand by burning un-favored byproducts such as liquor waste solids (www.epa.gov). Breeding and genetic approaches aiming lignin reduction in woods would thereby lead on the one hand to an eased extraction but on the other hand to a loss of energy. Industries energy demand is coupled to growing markets and an increasing world population which can be especially exemplified by developing countries. Optimistic predictions estimate that fossil energy resources will last for the next ~100 years without considering increase in demand and in costs. The biggest proportion of biomass produced on earth is made of plant cell wall polymers (Prade et al., 1999) and thus it displays the greatest source of renewable energy worldwide (Reiter, 2002). Approximately 1011 tons cellulose per year is naturally synthesized which can at least be partially used in an energy mixture. In the recent years attempts to replace petroleum based consumables has been discussed and initiatives to investigate cell wall derived products has been started (www.epobio.net). One of the foci is the exploration of the feasibility of replacing fossil fuels by biogas or bioethanol derived from saccharified material (Ragauskas et al., 2006; Gray et al., 2006). For instance, the USA agreed on that ~28 billion liters gasoline have to be supplied from renewable resources by 2012. Furthermore, it was estimated that the USA have the ability to produce around 50 billion liters of bioethanol per year using cornstarch but using nutritional carbohydrates from food plants will cause conflicts. Therefore, alternative feed stocks, such as cell walls from energy plants, have to be investigated which can assist feeding worlds energy demand (Möller et al., 2006). In conclusion, studying cell wall metabolism is not only of growing importance from the scientific point of view but also because many natural products are already made of plant cell walls or contain compounds extracted from them. Finally, the demand of plant cell walls as a sustainable resource will increase in the next decades and the intelligent usage will require sophisticated knowledge about polymer biochemistry and about the genetic and physiological control of their metabolism.
