Chapter 1: Introduction

1.1 The root and soil environment

1.1.1 Role of root exudates

Plant roots are highly dynamic organs, which serve as the sole interface between the plant and the soil. The root-soil interface not only serves to gather water and nutrients for the plant but involves a complex series of interactions with soil-dwelling microbiota, and the soil itself. These below ground interactions are fundamental for maintaining soil structure, soil ecology and plant growth (Kibblewhite et al. 2008; Lehmann and Kleber 2015), which are vital to the terrestrial biosphere. As a part of these complex interactions with the soil, plant roots release a range of low and high molecular weight compounds into the soil, which include: organic acids, amino acids, phenolics, enzymes, deoxyribonucleic acid (DNA), proteins, sugars, and polysaccharides (Neumann and Romheld 2001; Bais et al. 2006; Dennis et al. 2010). The exuded organic compounds have been demonstrated to be widespread amongst plants. However, it is unknown what proportion each molecule makes of the total root exudate. This array of released molecules is known as root exudate (Oades 1978; Foster 1982; McCully 1999).

Root exudates form the basis of a multifaceted response that plants may use to bioengineer the surrounding soil known as the rhizosphere. Root exudates can alter rhizosphere chemistry to gain more nutrients for the plant by acidifying or changing the redox conditions through chelating agents such as phytosiderophores, which can liberate nutrients, particularly iron that are bound to soil particles. Root exudates can also aid in mycorrhizal fungi associations, increasing the uptake of phosphorous and nitrogen for plants (Haichar et al. 2014), sequester toxic metals (particularly AI3+), and form the initial point of contact for invading pathogens before they enter the root body through degrading the cell walls (Badri and Vivanco 2009; McNear 2013; Baetz and Martinoia 2014). Furthermore, exudates released by a competitor plant have been shown to induce defence mechanisms in other species through genetic cues (Bais et al. 2006; Semchenko et al. 2014). However, the identity of these genetic cues and how they are recognised by the plant remain undetermined. Through scattered evidence, the organic substances and their proportions within root exudate are thought to be unique to a particular plant species, reflecting the biochemistry of their cells (Bacic et al. 1986; Moody et al. 1988; McCully and Sealey 1996; Read and Gregory 1997; Narasimhan et al. 2003; Biedrzycki et al. 2010).

Root exudates are believed to be released by two mechanisms, the first mechanism, is the passive transportation of exudates across the cellular membranes of the root cells into the surrounding soil due to osmotic differences between the soil and the cell (Bais et al. 2006). The second mechanism is through pre-programmed detached root cells, border and border-like cells (Driouich et al. 2013; Mravex et al. 2017). These border cells detach as the root caps penetrate through the soil and remain alive for up to 90 days, secreting root exudates independently of the root body (Miyasaka and Hawes 2001; Driouich et al. 2013). The exudates from these cells can blanket the root caps and protect them from friction as they burrow deeper into the soil (Figure 1.1; Ray et al. 1988; Bengough and McKenzie 1997; McCully 1999). Border cells are also at the forefront of the defence network of the plant. These cells can form cellular traps with actively released DNA that attract pathogens, engulfing them before invading the root body (Hawes et al. 1998; Gunawardena and Hawes 2002).

1.1.2 Role of root mucilage

Polysaccharides, long repeating subunits of sugars, and glycoproteins, proteins encapsulated in polysaccharide, are released by roots and form the high molecular weight components of root exudate. These components relate to the cell wall components of the root body. Released polysaccharides and glycoproteins form a thick mucilaginous layer surrounding the root caps and tips of plants (Schwartz 1883; Chaboud 1983; Bacic et al. 1986; Moody et al. 1988). These released polysaccharides and glycoproteins are collectively referred to as root mucilage. This root mucilage can be visualised as it forms a viscous droplet on exposed root caps and tips (Guinel and McCully 1986; Bacic et al. 1986). No other root structures have been reported to be involved in the release of these components. The mechanisms involved in the release of root mucilage from the root caps and tips remain unclear. The leading hypothesis is that they derive from sloughed off root cap and tip cells that lyse, releasing their cell wall components into the rhizosphere, due to continual friction whilst penetrating through soil (Read and Georgy 1997). As roots penetrate further into the soil, root caps are faced with an extraordinary amount of pressure (>7 kg/cm2; McNear 2013). Mucilage serves to lubricate the root caps to reduce the pressure faced by the caps as they penetrate through deeper layers of soil (Guinel and McCully 1986; Read and Gregory 1997; Figure 1.1).

Similar to that of the low molecule weight compounds of root exudate, root mucilage has also been implicated as a part of the defence network of plants, serving as a bait molecule, attracting beneficial microorganisms and sequestering heavy metals including aluminium, cadmium and lead (Morel et al. 1986; Ray et al. 1988; Dennis et al. 2010). Root mucilage that envelops root caps and tips also provides protection from desiccation, ensuring that they remain hydrated (Greenland 1979; McCully and Sealey 1996). It has even been proposed that root mucilage could bind soil particles together to form aggregates (Ray et al. 1988; Morel et al. 1991; Traore et al. 2000). This aggregation effect strengthens and maintains the root-soil interface otherwise known as the rhizosheath, which in turn increases water and nutrient acquisition for the plant, and enhances soil quality by increasing water infiltration and soil pore size, and thus aeration (Oades 1984; Hartnett et al. 2012; Sun et al. 2015). Some species of grass have been demonstrated to have some regulation over the thickness of their rhizosheath. During periods of drought some grasses can increase the thickness of their rhizosheath in an attempt to secure the uptake of water from the soil (Hartnett et al. 2012).

Research has alluded to the identity of the polysaccharides present within the root mucilages of several species including maize (Zea mays), wheat (Triticum aestivum), cowpea (Vigna unguiculata) and lupin (Lupinus angustifolius). These studies indicate the presence of arabinogalactan-protein (AGP), pectin, heteroxylan and xyloglucan within the root mucilages of the plants studied (Bacic et al. 1986; Moody et al. 1988; Guinel and Sealey 1996; Read and Gregory 1997). The release of these polysaccharides and glycoproteins consumes a large proportion of the photosynthetically fixed-carbon by plants. Research suggests that these macromolecules can consume up to 40% of the photosynthetic production of a plant (Newman 1985; McNear 2013). Although, the proportion of photosynthetically fixed-carbon released as mucilage remains controversial with numbers ranging between 10% and 40% (Walker et al. 2003; Jones et al. 2007). The release and amount of mucilage is thought to be dependent on a plethora of factors including; species, climate, nutrient availability, and the biological, chemical and physical properties of the local soil (Miki et al. 1980; Jones et al. 2009). There are many unanswered questions about the released polysaccharides and glycoproteins, and it is an area of research which is neglected as it is a hidden process of a plant, despite the high energy demand of releasing these molecules.

Diagram of root-soil interaction

Figure 1.1 I Overview of the complex interactions of the rhizosphere, taken from Dennis et al. 2010

Roots release an array of low and high molecular weight substances, known as root exudate, which may be involved in nutrient acquisition, plant defence, attraction of beneficial microbiota, and the modification of the structure of the rhizosphere. Some of the released polysaccharides and glycoproteins form a thick gel surrounding root caps and tips, known as root mucilage.

1.2 Related-cell wall components of released polysaccharides

1.2.1 The plant cell wall

Cell walls maintain the structural integrity of plant cells (Albersheim et al. 2010), and act as a decentralised skeleton which maintains the integrity of plant’s shape, and prevents it from collapsing under its own weight. Cell walls that are formed of a highly heterogeneous combination of polysaccharides (Figure 1.2; Bacic et al. 1988; Keegstra 2010; Fry 2011). There are two types of cell walls present in plants, primary and secondary. The primary cell walls are a flexible barrier that encapsulates each cell within plants. Primary cell walls provide mechanical support to each cell, maintain cell shape, regulate internal turgor pressure, pressure caused by water influx within the cell, and protect against pathogens, dehydration and other environmental factors (Somerville et al. 2004). The secondary cell wall, forms between the plasma membrane and primary cell wall of supporting tissues including sclerenchyma (fibrous cells), which develops after a cell has constructed its primary cell wall and has stopped expanding (Carpita and McCann 2000). Secondary cell walls provide mechanical support to many land plants. The secondary cell wall consists of cellulose microfibrils that form in parallel with non-pectic polysaccharides for instance, xylan and xyloglucan which crosslink with them (Carpita and McCann 2002). One of the key differences between primary and secondary cell walls is the inclusion of lignin, which is a phenolic that crosslinks with polysaccharides, making secondary cell walls less permeable to water and more rigid. Primary and secondary cell walls contain cellulose, non-pectic polysaccharides and pectin, although in different proportions (Carpita and McCann 2002; Albersheim et al. 2010).

In primary cell walls, cellulose microfibrils are extremely tough supporting structures which are crosslinked with non-pectic polysaccharides; examples include xylan, xyloglucan, heteromannan, and mixed-linkage glucans which are only found in Poaceae cell walls. These cellulose-non-pectic polysaccharides crosslinks form a strong interconnected framework (Cosgrove 2005; Knox 2008; Burton et al. 2010). This framework is embedded in a pectin matrix, which adds to the plasticity of cell walls (Figure 1.2; Cosgrove 2005). Pectin is a heterogeneous polysaccharide that is formed of four domains; homogalacturonan (HG), rhamnogalacturonan II (RG-II), xylogalacturonan, and rhamnogalacturonan I (RG-I). Pectin is rich with galacturonic acid making this complex macromolecule highly acidic. This acidity attracts calcium ions that bind the HG together, forming a gelatinous pectin matrix (Somerville et al. 2004). The outer layer of cell walls in plants are layered with pectin, forming the middle lamella, which acts like flexible gum, adhering neighbouring cells together and allowing them to stretch (Somerville et al. 2004; Albersheim et al. 2010). Cell walls are found across the entire plant kingdom with degrees of variation on the type and proportion of each polysaccharide.

Diagram of the plant cell wall

Figure 1.2 I Current model of primary cell wall matrix, based on known Arabidopsis configuration, taken from Somerville et al. 2004

Three major polysaccharides are present in primary cell walls, cellulose, non-pectic polysaccharides and pectin. Cellulose microfibrils are woven together to form microfibrils, which provide the tensile strength of the cell wall. The non-pectic polysaccharides, including xyloglucan (XG), form interwoven tethers linking the cellulose microfibrils together. This cellulose-non-pectic polysaccharide framework is embedded within a pectin matrix.

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1.2.2 Major polysaccharides within plant cell walls

Cellulose is the most abundant polysaccharide on Earth. It is a ubiquitous polysaccharide, accounting for a third of the total mass of a typical plant (Somerville 2006). Cellulose is a homogenous polysaccharide formed of linear chains of many thousands of 1,4-β-linked glucose residues (Figure 1.3). Cellulose microfibrils are formed of cellulose chains that are hydrogen bonded in parallel to each other (Somerville et al. 2004; Zhang et al. 2017). Cellulose microfibrils are the key structural support of plant cell walls, and are found in numerous everyday products such as paper, textiles and insulation.

Non-pectic polysaccharides are a broad description of any non-cellulosic cell wall polysaccharides; for instance, heteroxylan, xyloglucan, heteromannan and mixed-linkage glucan. Heteroxylan is a heterogeneous polysaccharide, which has a backbone of 1,4-β-linked xylose residues (Figure 1.3). Heteroxylan can be in three main forms depending on the inclusion of arabinosyl or glucuronyl side chains, glucuronoxylan, arabinoxylan and glucuronoarabinoxylan (GAX). When heteroxylan has a single 1,2-α-linked glucuronic acid residue linked to the backbone, for about every 10 xylosyl residues, it is glucuronoxylan (Albersheim et al. 2010). Glucuronoxylan is a charged form of heteroxylan that is mainly found in the secondary walls of eudicotyledons such as in birchwood (Albersheim et al. 2010; Hao and Mohnen 2014). When heteroxylan has been substituted with α-L-arabinofuranose residues, attached at positions 2, 3 or 2,3 on the backbone, it is arabinoxylan. Arabinoxylan is particular abundant in the secondary cell walls of plants, and within the endosperm of cereal grains (Albersheim et al. 2010). GAX is similar to its counterpart arabinoxylan but includes a single glucuronic acid residue for every 5 to 6 xylosyl residues on its backbone (Albersheim et al. 2010; Hao and Mohnen 2014). GAX is only present within the cell walls of commelinid monocotyledons such as Bromeliads and Poaceae. The GAX in the secondary cell wall of Poaceae contain fewer side chains than the GAX found in primary cell walls, which is believed to result in a more robust GAX-cellulose interaction (Vogel 2008; Burton et al. 2010). Heteroxylan is a structural supporting polysaccharide within the cell wall, forming hydrogen bonds on cellulose microfibrils (Simmons et al. 2016).

Xyloglucan comprises of a backbone of 1,4-β-linked glucose residues carrying various side chains, such as 1-α-linked xylose, 1,2-β-linked galactose, 1,2-α-L-arabinofuranose. Within cereals xyloglucan can be fucosylated through a 1,2-α-linked L-fucose linked to a 1,2-β-linked galactose residue. The backbone, which is the same as cellulose, is responsible for the strong crosslinking between xyloglucan and cellulose microfibrils within cell walls (Knox 2008; Scheller and Ulskov 2010; Burton et al. 2010; Figure 1.3). One feature of xyloglucan is that there is a unique regularity in the distribution of the side chains, which occurs in different species. This results in repetitive subunits on the structure of xyloglucan. This variation in branching patterns is denoted by a one letter code, for instance, XXXG represents three 1-α-linked xylose residues followed by one 1,4-β-linked glucose residue which are all linked to the 1,4-β-linked glucose backbone (Hayashi 1989; York et al. 1996; Figure 1.3). The primary cell walls of a range of eudicotyledons, non-cereal monocotyledons and gymnosperms contain fucosylated xyloglucans with a XXFGF structure. Whereas in Solananceae, which includes potato, fucose residues are rarely present but have short arabinofuranosyl side chains (Hayashi 1989; York et al. 1996; Albersheim et al. 2010).

Generally, xyloglucan does not carry a charge, however, there have been reports that demonstrate the presence of a charged form of xyloglucan within the cell walls of non-vascular land plants such as liverworts (Peña et al. 2008), and in the root hairs of Arabidopsis (Peña et al. 2012a). This rare xyloglucan has xylose side chains that contain a 1,2-β-linked galactosyluronic residue, which gives this xyloglucan a charge (Peña et al. 2012a). This galacturonic acid residue typically forms a xyloglucan structure where a 1,2-β-linked galacturonic acid residue is linked to a 1-α-linked xylose, which is in turn linked to the 1,4-β-linked glucose backbone. Within the root hairs of Arabidopsis a 1,2-α-linked L-fucose residue is able to link with the galactosyluronic residue (Peña et al. 2012a). Despite the variability within the structure of xyloglucan, xyloglucan has been demonstrated to be widespread throughout plants and is considered unique to plants (Fry 1989; Peña et al. 2008).

Mannans are a highly heterogeneous polysaccharide, which have a backbone of 1,4-β-linked mannose residues (Figure 1.3). This heteromannan can be in four forms, mannan only formed on 1,4-β-linked mannose residues, glucomannan, galactomannan and galactoglucomannan (Albersheim et al. 2010). Glucomannan consist of an 1,4-β-linked mannose and 1,4-β-linked glucose backbone. Glucomannan is a minor polysaccharide within the primary cell walls of eudicotyledons and cereals (Scheller and Ulskov 2010). In galactomannan there is a 1,4-β-linked mannose backbone which has side chains of 1,6-α-linked galactose. In galactoglucomannan the backbone consist of mannose and glucose residues, which has side chains of 1,6-α-linked galactose. These side chains of galactose residues are mostly linked to the mannose on the backbone but some galactose can linked to the glucose on the backbone (Benova-Kaosova et al. 2006; Goubet et al. 2009). Within the secondary cell walls of gymnosperm there is a galactose: glucose: mannose ratio of 1:1:3 for the composition of heteromannan. Whereas, this ratio is 1:1 (glucose: mannose) within the primary cell wall of plants. Furthermore, the proportion of galactose is much lower in primary cell walls (Benova-Kaosova et al. 2006; Scheller and Ulskov 2010). Heteromannans are widely spread throughout the plant kingdom, functioning as a structural support as well as a means of storage (Goubet et al. 2009).

Mixed-linkage glucans are only found in cereals and ferns. Mixed-linkage glucans are formed of a mixture of 1,4-β-linked and β-1,3 linkage glucose residues (Woodward et al. 1983). Each segment of mixed-linkage glucan contains between two and three continuous 1,4-β-glucan residues attached by a single 1,3-β-glucan residue (Woodward et al. 1983; Buckerigde et al. 1999; Figure 1.3). These 1,3-β-glucan linkages twist the structure of the polysaccharide, which gives the mixed-linkage glucan more flexibility as it prevents the aggregation of cellulose hydrogen bonds. These linkages also increase the solubility of mixed-linkage glucan compared to a continuous chain of 1,4-β-glucan residues, which forms cellulose (Kiemle et al. 2014; Figure 1.3). Mixed-linkage glucans are most abundant in the cell walls of the endosperm, and young tissues (Kiemle et al. 2014).

Pectin is a complex heterogeneous macromolecule formed of four domains: HG, xylogalacturonan, RG-I and RG-II. Pectin is a highly acidic polysaccharide due to the backbone containing an abundance of 1,4-α-linked galacturonic acid residues (Caffall and Mohnen 2009; Figure 1.3). The ratios of the pectin domains is highly variable, with HG being the most abundant, constituting typically 65% of the total polysaccharide, RG-I constitutes between 20% and 35%, RG-II and xylogalacturonan each constituting less than 10% (Zandleven et al. 2007; Mohnen 2008). HG consists of a backbone of 1,4-α-linked galacturonic acid residues. A number of galacturonic acid residues can be methyl esterified on the carboxyls or acetylated on the hydroxyls (Carpita and McCann 2000; Figure 1.3). The backbone of galacturonic acids enables HG to form gels by crosslinking divalent ions such as calcium. Modifications, methyl and acetyl esterification, influence the gelling properties of pectin by lowering the density of the gels (Cosgrove 2005). Similar to HG, xylogalacturonan is formed of a backbone of 1,4-α-linked galacturonic acid with side chains of 1,3-β-linked xylose attached to the hydroxyl of the galacturonic acid residues (Zandleven et al. 2007; Mohnen 2008; Figure 1.3).

RG-I consists of a backbone of alternating 1,2-α-linked rhamnose and 1,4-α-linked galacturonic acid residues. RG-I is a highly branched polysaccharide with many terminal residues of galactosyl and arabinosyl residues on the hydroxyls of the terminal rhamnosyl residue (Figure 1.3). The 1,4-α-linked galacturonic acid backbone is frequently acetylated on the hydroxyls (Moller et al. 2008; Verhertbruggen et al. 2009). The backbone of RG-I can also be substituted on the C-4 position with 1,5-α-linked arabinans, 1,5-α-linked galactans or arabinogalactans (Carpita and Gibeaut 1993).

RG-II is considered to be the most complex component of the cell wall that consists of 12 monosaccharides linked by 20 different types of glycosidic linkages as determined by enzyme treatment (Ralet et al. 2008; Lee et al. 2013). RG-II typically consists from eight 1,4-α-linked galacturonic acid residues within its backbone. Some of the galacturonic acid on the backbone can be methylated (Figure 1.3). The number of side chains of RG-II is highly dependent on the species (Ishii et al. 1999; Harholt et al. 2010). Rare sugars including D-apiose are also present on RG-II, which makes RG-II hard to degrade by cell wall enzymes. RG-II is also a highly conserved structure of plant cell walls, which uniquely contains a borate (BO3) that gives RG-II the ability to link with other RG-II (Kobayashi et al. 1996; Roach et al. 2012). Pectin is a gelling agent that is primarily involved in forming the matrix of the cell wall. Commercial uses of pectin vary but take advantage of its gelling properties, and are used as binding agents in food including jams (Crandall and Wicker 1986).

Cell walls also contain various proteins, which have enzymatic or structural roles. Cell wall proteins have many functions including signalling, remodelling or constructing the architecture of the cell wall, and defence (Albersheim et al. 2010). Structural cell wall proteins can be classified into three groups based on their amino acid content, glycine-rich proteins, proline-rich proteins and hydroxyproline-rich proteins. One group of proteins, AGPs are rich in both proline and hydroxyproline (Clarke et al. 1979; Kieliczewski 2001). AGPs are typically formed of 90% glycan and 10% protein (Bacic et al. 1997). The protein domain is typically heavily glycosylated and varies in length between 5 KDa and 30 KDa, and rich in alanine and hydroxyproline amino acids (Bacic et al. 1997; Kieliczewski 2001). The glycan domain of AGP is formed of arabinogalactan that is O-linked to the hydroxyproline residues in the protein core. The glycan backbone of AGP is typically formed of 1,3 and 1,6-β-linked galactose residues with side chains of a single 1,6-β-linked galactose that holds 1,3-α-linked L-arabinose residues, 1,6-β-glucuronic acid residues and a single 1,4-α-linked L-arabinose residues (Ellis et al. 2010; Figure 1.3). The presence of glucuronic acids can vary between species making the glycoprotein more charged, for instance, the AGP in gum Arabic is rich in glucuronic acid making the polysaccharide highly acidic (Gane et al. 1995). AGPs have a variety of functions from cell differentiation, reproduction, cell signalling and programmed cell death (Bacic et al. 1997; Kieliczewski 2001; Cosgrove 2005; Pereira et al. 2013).

Another example of a structural cell wall protein are extensins, which are a type of hydroxyproline-rich proteins, which contain low amounts of repeating serine amino acids (Borner et al. 2002). These chains of repeating one serine and four hydroxyprolines form a rod-like shape of extensin. Extensin are glycosylated proteins with short (3-4 residues) repeating chains of 1,4, 1,2 1,2 and 1-3-α-linked L-arabinose residues that are attached to the hydroxyprolines. Attached to the serine is a single 1,3-β-linked galactose residue (Borner et al. 2002; Figure 1.3). Extensins are widespread throughout the plant kingdom, and are a key component that is responsible for cell wall rigidity. Production of extensins can be induced as a wounding response (Hirsinger et al. 1997).

Diagram of major plant polysaccharides

Figure 1.3 I Schematic of the major cell wall polysaccharides, adapted from Burton et al. 2010 and Liang et al. 2010

The major macromolecules of the cell wall can be placed into four categories: cellulose, non-pectic polysaccharides, pectin and glycoprotein. Cellulose is formed of a 1-4-β-D-glucan backbone. When the backbone of cellulose has been decorated with xylose, galactose and fucose residues it forms the non-pectic polysaccharide, xyloglucan. When the backbone of cellulose is twisted by having a 1-3-β-D-glucan substitute, forming mixed-linkage glucan. Xylan is a heterogeneous non-pectic polysaccharide, which generally has a backbone formed of 1-4-β-D-xylose with residues of arabinose and glucuronic acid residues. Hetromannan is another heterogeneous non-pectic polysaccharide that generally has a backbone of mannose and glucuronic acid residues that is decorated with galactose residues. Pectin is a complex macromolecule formed of four domains, RG-I, HG, xylogalacturonan (XGA) and RG-II. The backbone of pectin is rich in galacturonic acid residues. AGP are formed of a protein core rich in hydroxyproline that has been enclosed in a glycan rich in galactose, arabinose, glucuronic acid and rhamnose residues. Extensin is a rod shaped hydroxyproline-rich glycoprotein, which contains high amounts of arabinoses with one galactose residue.

Based on cell wall biochemistry and architecture, angiosperm or flowering plants can be placed into two basic categories, Type I and Type II. Type I primary cell walls are found in eudicotyledons and non-commelinid monocotyledons, which include the families of Solananceae and Brassicaceae. Type I cell walls contain cellulose microfibrils and high amounts of xyloglucan crosslinks (Figure 1.2; Carpita and Gibeaut 1993), which are embedded within a pectic matrix (Ishii 1997). Type II cell walls, found in grasses such as Poaceae, which include wheat, oat (Avena sativa) and barley (Hordeum vulgare), contain cellulose microfibrils that are encased within a matrix of GAX and mixed-linkage glucans, with trace levels of xyloglucan crosslinks (Nishitani and Nevins 1989; Carpita 1996; Vogel 2008). In addition, Type II cell walls contain less pectin and glycoproteins such as AGP, and contain higher crosslinks between lignin and other polysaccharides, which form rigid interconnections when the cell has stopped expanding (Iijima et al. 1990; Burton et al. 2010). An example of a phenolic crosslink in Type II cell walls is feruloylated arabinoxylan. Feruloylated arabinoxylan is formed of arabinoxylan that has been decorated by ferulic acid, which mediates linkages between arabinoxylan chains and between arabinoxylan and lignin in grasses (Chateigner-Boutin et al. 2016).

1.2.3 Cell wall multi-polysaccharide complexes

The current view of the cell wall matrix is that the polysaccharides are separate structures that are tethered together non-covalently, forming a complex framework (Gibeaut and Carpita 1994; Keegstra 2010; Cosgrove and Park 2012). Recently, this view has begun to change, as some protein, non-pectic and pectic polysaccharides have been shown to form covalent complexes. This finding adds an additional layer of complexity to the cell wall architecture of plants. This type of bond can also form linkages with other polysaccharides, therefore, forming multi-polysaccharide complexes. Covalent attachments between polysaccharide-protein were initially hypothesized in the 1990s (Ryden and Selvendran 1990), but very little evidence had been uncovered until covalent bonds between xyloglucan-pectin (RG-I) were reported in Arabidopsis callus cells (Renard et al. 1997; Duan et al. 2004; Popper and Fry 2008). It was proposed that the negatively charged pectin domain of this xyloglucan-pectin complex promoted the integration of xyloglucan into the cell wall matrix (Popper and Fry 2008). After the synthesis of xyloglucan-pectin, within the Golgi apparatus, the complex remained highly stable, being transported to the cell wall through vesicles until its integration into the cell wall (Popper and Fry 2008). It was also observed that approximately 30% of this complex was released into the medium of the cells in the form of sloughed cell wall components (Popper and Fry 2008). However, the function of this complex remains unclear. It is assumed that the xyloglucan-pectin complex aided in cell wall assembly and acted as structural support within the cell wall. Potential linkages of xylan-pectin and xylan-AGP have also been reported when using immunochemical techniques (Cornuault et al. 2015). Another form of polysaccharide complex has also been described, where xylan could fold onto the surface of cellulose microfibrils through hydrogen bonding in a two-fold helical screw formation (Simmons et al. 2016). This folding, similar to the xyloglucan-pectin, is also assumed to contribute to the structural integrity of the cell wall.

Evidence for a covalently linked multi-polysaccharide complex had not been detected until four years ago. Research using Arabidopsis (Arabidopsis thaliana) callus cells has uncovered a multi-polysaccharide complex, Arabinoxylan Pectin Arabinogalactan-Protein (APAP1; Tan et al. 2013), which was released into the liquid medium. This complex contains an AGP core with branching subdomains of covalently attached arabinoxylan and RG-I (Tan et al. 2013). The AGP core is acting as a cross-linker holding the pectic and arabinoxylan subdomains together (Tan et al. 2013). Within the cell wall, this crosslinking could provide a continuous network of cell wall polysaccharides and structural proteins, thereby increasing the strength of the cell wall. However, this multi-polysaccharide complex was only found to be a minor component of the cell wall when compared to the other polysaccharide components of the callus cells. On further analysis, the pectic and arabinoxylan subdomains were shorter within the complex that was exuded compared to the complex that was detected within the cell wall (Tan et al. 2013). This suggests that this multi-polysaccharide complex was tightly woven through hydrogen bonding within the fabric of the cell wall matrix. Although, the mechanisms underpinning how the subdomains are assembled into a complex, and the location of its assembly both remain undetermined. One hypothesis suggests that the subdomains are formed in the Golgi apparatus and translocated to the plasma membrane where it is assembled before being integrated into the cell wall matrix (Tan et al. 2013).

1.2 Plant cell wall polysaccharide analysis

1.3.1 Monosaccharide linkage analysis

For monosaccharide linkage analysis to be undertaken, polysaccharides within a sample are methylated, typically using methyl iodide, uncovering all open hydroxyls on the polysaccharide (Lisec et al. 2006). The sample is then hydrolysed using a strong acid, trifluoroacetic acid, which breaks down the glycosidic linkages holding the polysaccharide together (Björndal et al. 1970; Lindberg 1972), thereby, generating monosaccharides. The hydroxyls that were involved in the glycosidic linkages remain unmethylated. The free monosaccharides are then reduced, typically with sodium hydroxide, to open their rings. Once reduced, these open rings are then acetylated with the unmethylated hydroxyl groups, which were involved in linkage, forming stable acetyl groups (Hakan et al. 1970; York et al. 1985). The samples are then analysed by gas chromatography, liquid chromatography or gel chromatography that is combined with mass spectrometry (Hakan et al. 1970; Pettolino et al. 2012). NMR can be used instead of mass spectrometry to explore in more detail the conformational structure within polysaccharides (Pettolino et al. 2012), rather than molecular weights. To identify monosaccharides linkages, the retention times from the chromatography separation are combined with the molecular weights determined from mass spectrometry (Bauer 2012; Pettolino et al. 2012). Altogether, the hydroxyls linking the polysaccharide and types of glycosidic links can be determined, and therefore, the identity of the polysaccharide can be inferred (Björndal et al. 1970; Lisec et al. 2006; Pettolino et al. 2012). Using this established technique, many studies have determined the identity of the polysaccharides released by plant roots and within the cell walls of many species. However, fully understanding the biochemistry of plant cell walls using monosaccharide linkage analysis is challenging and time consuming. Moreover, these physio-chemical methods require large amounts of material for analysis, typically a minimum of 1 mg is needed (Moller et al. 2008; Pattathil et al. 2012). This can be a limiting factor of deciphering smaller components of the cell wall.

1.3.2 Monoclonal antibodies and Carbohydrate Binding Modules

A complementary approach to detect a range of cell wall-related polysaccharides and glycoproteins are monoclonal antibodies (MAb) and carbohydrate-binding modules (CBMs), which are highly specific and sensitive molecular probes. When a foreign substance or antigen (for example bacteria) enters an animal’s body, the white blood cells or lymphocytes within the lymphatic system, which acts like a blood filter, detects the antigen and in response increases the production of antibodies (Medzhitov and Janeway 1997). These antibodies recognise and bind to the foreign substance for other lymphocytes to target them (Willats et al. 2002).

MAbs used to detect cell wall polysaccharides are cloned from a single lymphocyte (removed from the spleen which stores them) that generates antibodies in response to a polysaccharide (antigen) that has been injected into an animal such as mice or rat (Willats et al. 2000; Willats et al. 2002). The antibodies generated from these lymphocytes solely bind to one epitope of that antigen (Moller et al. 2008; Lee et al. 2011). An epitope is defined as a specific oligosaccharide motif on a polysaccharide, usually between 3 and 6 monosaccharides in length (Lee et al. 2011). For instance, the MAb LM2 binds to the epitopes of β-glucuronic acid which is present on AGP (Smallwood et al. 1996; Yates et al. 1996). These MAbs are highly versatile, and are used within many in situ and in vitro immuno-assays as diagnostic tools including: Enzyme Linked Immuno-sorbent Assay (ELISA), immunoblotting and chromatography. There is also a large library of MAbs specific to many different types of glycan and glycoprotein available from a variety of sources including: Leeds Monoclonal (LM), John Innes Monoclonal (JIM), Complex Carbohydrate Research Centre Monoclonal (CCRCM) and Monoclonal Antibody Centre, Cambridge (MAC).

As an alternative to MAbs, CBMs are also highly specific molecular probes, which derive from microbial cell wall polysaccharide hydrolases. These molecular probes are non-catalytic protein domains from bacterial carbohydrate-active enzymes such as glycoside hydrolases, which specifically bind to their target polysaccharide (Boraston et al. 2004; Knox 2008), comparable to that of MAbs. There is a diverse range of CBMs available on the Carbohydrate-Active enZYmes database that bind to many polysaccharides such as CBM2b1-2, specific to xylan (McCartney et al. 2006), CBM3a specific to xyloglucan and cellulose (Blake et al. 2006), which are used within many immuno-assays including ELISA, as with MAbs.

Most techniques that do not use MAbs or CBMs, such as gas chromatography and monosaccharide linkage analysis, require large amounts of material, between 1 mg and 10 mg for one run (Pettolino et al. 2012). This need for large amounts is problematic to dissecting small tissues or samples containing low concentrations of polysaccharides. These MAbs and CBMs are highly advantageous at exploring polysaccharides contained in low amounts within cell walls, and can be used on mass to generate cell wall profiles. Molecular probes are also widely used to dissect the architecture of cell walls. The cell walls of tissues can be sequentially solubilised using alkali treatments for example, sodium carbonate (Na2CO3) and potassium hydroxide (KOH) to release the components of the cell wall. Cation collators are also used to release polysaccharides within the cell walls that are bound by cations for instance, Cyclohexane Diamine Tetraacetic Acid (CDTA) binds to Ca2+ that holds pectic polysaccharides (specifically HG) together (Cornuault et al. 2014; Pose et al. 2015). Upon treatment, either MAbs or CBMs can efficiently identify which polysaccharides have been released through immuno-based techniques including ELISA. Using the outlined technique, research that solubilised cell walls with CDTA and KOH uncovered potential linkages between pectin and xylan in potato tubers (Solanum tuberosum), and xylan and AGP in commercial oat grain cell walls (Cornuault et al. 2015).

MAbs and CBMs can also be used for in situ fluorescence labelling to reveal locations of specific polysaccharides within the cell wall (Knox 2008). This fluorescence labelling has enabled a plethora of studies exploring the architecture, composition, plasticity and re-modelling of the cell walls in many tissues including: leaves and stems (Hanford et al. 2003; Blake et al. 2006; Verhertbruggen et al. 2009; McCartney et al. 2009), pollen (van Aelst and van Went 1992), fruits (Ordaz-Ortiz et al. 2009) and roots (Willats et al. 2004) within a wide range of species. These MAbs have enabled researchers to reveal where each major polysaccharide is within the cell wall matrix with precision, which had previously been unavailable. Prior to the development of MAbs and CBMs, a limited number of dyes were used to localise cell wall components. However, these dyes were not as specific or quantitative as MAbs. For instance, Calcofluor white binds to 1-4-β-D-glucan which forms cellulose but this also forms the backbone of xyloglucan; as a result the dye could bind to both polysaccharides (Herth and Schnepf 1980). Consequently this led to difficulties interpreting their signals. As a direct result of the development of MAbs and CBMs, many new techniques have been established such as, high-throughput glycan microarrays. These microarrays use small dots of cell wall extract that are placed onto nitrocellulose sheets, and are screened using a large library of MAbs or CBMs (Moller et al. 2008). This allowed the rapid identification of polysaccharides within an extract, which was previously unavailable. Additionally as these microarrays use MAbs or CBMs, small components of the cell wall could be assayed.

1.4 Profiles of polysaccharides released by roots

A limited number of studies examining polysaccharides released by plant roots suggest a close relationship between the polysaccharides present within the root body and polysaccharides released by roots (Bacic et al. 1986; Guinel and McCully 1986; Moody et al. 1988; Read and Gregory 1997). Investigations using monosaccharide linkage analysis have determined that wheat and maize released high amounts heteroxylan, and low amounts of xyloglucan and pectic polysaccharides (Chaboud, 1983; Chaboud and Mireille 1984; Bacic et al. 1986; Table 1.1). Whereas, cowpea cress (Lepidium sativum), and Indian rhododendron (Melastoma malabathricum) released trace amounts of xyloglucan, and high proportions of pectic polysaccharides including, RG-I (Moody et al. 1988; Sims et al. 2000; Watanabe et al. 2008). The released polysaccharides are similar to their respective eudicotyledons and grasses cell wall biochemistries (Vogel 2008). However, one investigation using callus cells of Timothy-grass (Phleum pratense) found that there was more arabinogalactan-protein (AGP) and xyloglucan exuded by cells compared to that of their cell wall (Sims et al. 2000). This differential in released and cell wall polysaccharides may be due to the undifferentiated state of the cells. It is probable that these polysaccharides would be released in similar proportions to that of the cell wall. However, there is no evidence that polysaccharides are released in equal proportions. Many studies have not directly detected these polysaccharides, and as a result they cannot calculate their proportions in and outside of the cell wall. For a summary of work to date see Table 1.1.

Prior to analysis, polysaccharides released from roots had to be isolated. There were three widely employed methods. One method involved teasing roots from the soil to collect root mucilage in situ over a number of days (Morel et al. 1986; Mounier et al. 2004). Another method involved growing seedlings (defined as ≤7 days old) on moist filter paper in the dark (McCully and Sealey 1996; Read and Gregory 1997; Narasimhan et al. 2003) or in the light (Ray et al. 1988) and directly pipetting the mucilage that was secreted from the root caps of seedlings (Read and Gregory 1997). Alternatively, seedlings could be submerged in sterile liquid media and grown on (Bacic et al. 1986; Moody et al. 1988; Osborn et al. 1999). However, submerging seedlings in liquid medium may result in difficulties interpreting data as the whole plant has been submerged, not just the roots. Furthermore, plants grown using liquid culture must be agitated to oxygenate the medium. It has been documented that this agitation can damage plants particularly the leaves, which may release particular cell wall components such as extensin as a stress response (Smallwood et al. 1995). As yet, no research has used hydroponics to collect the polysaccharides released from the roots of plants.

Table of plant mucilage compositions

1.5 Mechanisms involved in the release of polysaccharides and from roots

The mechanisms underpinning the release of polysaccharides and glycoproteins by roots have yet to be determined. However, there are two hypotheses that have been proposed. The first hypothesis proposes that released polysaccharides originate from lysed root cap cells (Read and Gregory 1997). As roots penetrate through the soil, root cap cells and tips are regularly lysed due to the continual friction that the soil exerts on the root caps. When the cells undergo lysis they release the polysaccharides that were contained within the cell wall matrix (Read and Gregory 1997; Iijima et al. 2002; Iijima et al. 2004). The second hypothesis proposes that the released polysaccharides are actively secreted through root cell walls (Guinel and McCully 1986). Plant polysaccharides are synthesised in the Golgi apparatus, and then are packaged into vesicles. The vesicles fuse to the plasma membrane and are deposited to the cell wall (Northcote and Pickett-Heaps 1966). The hypothesis suggests that the polysaccharides to be released travel through the cell wall matrix until they are secreted through specialised pores within the cell wall (Guinel and McCully 1986). However, no direct evidence for these specialised pores or the active secretion of polysaccharides has been observed. The active transportation of polysaccharides through this complex matrix would demand high amounts of energy. It has been shown that as roots penetrate through the soil; root cap cells are regularly lysed, thereby releasing their components which are made of polysaccharides that are sticky (Millar et al. 1989; Iijima et al. 2004). These gummy polysaccharides would then form the mucilage that surrounds the caps where they had originated (Bacic et al. 1986; Morel et al. 1987).

Root border cells are cells that are programmed to detach themselves from the root cap, and divert all their energy reserves into the production of released polysaccharides (Hawes et al. 1998; Cannsean et al. 2012; Mravec et al. 2017). Prior to their separation, the cement holding cell walls together, pectin, is degraded through the enzyme polygalacturonase (Driouich et al. 2007; Durand et al. 2009). The process of separation also loosens the cell wall matrix, resulting in the solubilisation of cell walls (Hadfield and Bennet 1998). It was proposed that the degradation of pectin (HG, XGA and RG-I) and arabinan within the middle lamella were involved in the detachment of cells on the root caps, which form root border cells (Willits et al. 2004; Mravec et al. 2017). It was also suggested that pectic polysaccharides could significantly contribute to root mucilage. Using a MAb that specifically binds to XGA (LM8; Willats et al. 2004), it was uncovered that XGA was secreted into the root mucilage of pea through either secretory vesicles or by aggregates (Mravec et al. 2017). After detachment, root border cells continually form new polysaccharides to replace their cell wall matrix, which is continually being lost to the rhizosphere (Figure 1.1; Stephenson and Hawes 1994; Driouich et al. 2007; Driouich et al. 2013). Once root border cells become free, the enzymatic activity that resulted in their detachment continues to degrade attached cells. However, these activities are not as effective, which results in semi-detached cells with a loosened cell wall matrix (Hawes et al. 2002; Driouich et al. 2013). Once loosened, the weakened cells could release their components into the rhizosphere. The semi-detached root cap cells would then have to produce more polysaccharides, to ensure their structural integrity, to replace their cell wall matrix, which was being lost to the rhizosphere.

Some research indicates that polysaccharides are not solely released from the root caps and tips of plants. It was hypothesized that these polysaccharides are also released from the epidermal layers behind the root caps and tips (Guinel and McCully 1986). An AGP-specific binding reagent, β-glycosyl Yariv reagent, and an AGP-specific antibody, α-L-arabinofuranosyl were used to stain the roots of maize, and could bind to the root caps, tips and epidermal layers behind the caps and tips (Bacic et al. 1986). Perhaps, as the root caps penetrate through the soil, epidermal cells are also subjected to mechanical pressures but to a lesser amount. This idea was supported by the lower levels of AGP that were detected being exuded from the epidermal layers compared to the root caps and tips (McCully and Sealey 1996; Read and Gregory 1997). The areas (how far along the root epidermis) that are involved in the release of polysaccharide behind the caps and tips remain unknown. Abundant on the epidermal layer of roots are root hairs, which are single celled elongated outgrowths. The primary role for root hairs is to increase the surface area of roots to secure the rhizosheath, and to extract water and nutrients from the soil (Gilroy and Jones 2000; Hartnett et al. 2012; Huang et al. 2017). Root hairs have not been observed to release molecules such as cell wall-related polysaccharides; with no research being undertaken on root hairs with regards to exudation.

Despite the mechanisms underpinning secretion, pectic polysaccharides, specifically homogalacturonan, and AGP have been detected from the root caps of several crop species such as maize and wheat (Table 1.1; Bacic et al. 1986; Guinel and McCully 1986; Morel et al. 1987; Moody et al. 1988; Read and Gregory 1997). Within plants, pectic polysaccharides are involved with the adhesion of plant cells (Willats et al. 2004; Driouich et al. 2007); AGPs are also present on the surface of cells (Albersheim et al. 2010). Since both these polysaccharides are present on the cellular surface, it is probable that they would be present in root mucilage. Pectic polysaccharides and AGP have a high water binding capacity that can form highly viscous gels within the cell wall matrix (Albersheim et al. 2010). Due to their gelling properties these polysaccharides are used as food additives to produce jams and gums. It is reasonable to suggest that these gelling agents result in the gelatinous properties of root mucilage.

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1.6 Soil organic matter and aggregate status

1.6.1 Soil organic matter

Root mucilage is continually released into the rhizosphere. It is hypothesized that root mucilage can influence the abundance of soil aggregates (Metha et al. 1960; Tisdall and Oades 1982; May et al. 1993). This form of bioengineering ensures that the roots have a strong interface with the soil so that they can extract the necessary resources required for plant growth. Soil is a vital component of the terrestrial biosphere, which supports ecosystems that are crucial to all land-based life. Soil is a highly complex and dynamic substrate, which is estimated to contain between 1010 and 1026 microorganisms per gram (Prosser 2015). As well as holding a multitude of life, soil also supports all crop production that the world population depends on (Prosser 2015). Prior to land-dwelling life, the land would have been formed of fragmented rock caused by atmospheric chemical weathering, and mechanical processes (Huggett 1998; Bateman et al. 1998).

The structure and composition of soil is complex, and highly heterogeneous. Soil particles can be placed into three basic categories, clay particles (≤2 µm), silt particles (between 2 µm and 60 µm) and sand particles (between 60 µm and 2,000 µm; Day 1965). Various concentrations of these particles exist in nature, which generate different soil types that have distinctive characteristics (Brown 2008). For instance, if a soil contains a high proportion of sand particles, the soil becomes more efficient at draining excess water. However, this results in the soil being poor at retaining water (Day 1965). Plants grown in this soil type will require frequent irrigation compared to soils with high proportions of clay and silt particles, which retain more water but are less effective at draining excess water (Lal 1997). The smaller particles in soil including silt and clay, collectively have a greater surface area compared to larger particles such as sand. The larger surface area allows the smaller particles to hold onto higher amounts of water (Cheshire 1979; De Booth et al. 1984; Brown 2008). Additionally, soils that contain higher amounts of aggregates have greater porosity. Soils with larger and more frequent pores have a greater capacity to bind to water due to the larger surface area, which would otherwise drain away (Lal 1997). This increase in water holding capacity is also influenced by the levels of organic matter within the soil. A larger amount of organic matter in soils also increases the retention of water (Kibblewhite et al. 2008). Having a high water holding capacity results in more water being available to plants.

One of the most complex facets of soil is the organic matter. One of the largest contributors of soil organic matter is from decaying plant matter such as leaves and roots. Other sources include excretions and decaying matter from animals and microbes (Kibblewhite et al. 2008; McNear 2013). These sources of organic matter significantly contribute to the stability of soils by gluing soil particles together forming aggregates (Cheshire 1979; De Booth et al. 1984; Ebringerová and Heinze 2000; Carrizo et al. 2015). Without this glue, the structure of the soil would breakdown and become easily eroded. The gradual decay of organic matter in soils is mostly caused by microbes, turning freshly fallen leaves into humus. Humus is the dark brown or black substance found in the upper layer of soil (0-30 cm depth), where the soil is very crumbly and biologically active (Brown 2008). Soil-dwelling invertebrates, including earthworms regularly churn up this humus with the mineral fragments of the soil. Humus has many beneficial aspects to soil, from maintaining the structure of soil through particle aggregation, thereby increasing porosity, water infiltration and nutrient cycling (Haynes and Francis 1993; Lal 1997; Kibblewhite et al. 2008).

The decomposition of biomass in soil by microorganisms results in the release of carbon dioxide due to microbial respiration. Only a small proportion of carbon is retained within soil through the formation of humus. Humus is highly resistant to decomposition, whereas freshly laid plant debris is more prone to decomposition (Cheshire 1979; De Booth et al. 1984; McNear 2013). As a result of this deposition and release of carbon, soils potentially hold a vast amount of carbon. Some estimates suggest there is up to 2,500 billion tonnes of carbon within soils, up to 80% of global organic carbon (Houghton 2007; Kibblewhite et al. 2008). This sink is increasingly under scrutiny to develop more accurate climate models and as a tool in climatic imitation strategies.

Humin is said to be the principal component of humus, which is a collective term used to describe a heterogeneous mixture of macromolecules, derived from decaying organic matter in soils (Cheshire 1979; Cheshire 1990). Humin is said to be the most resistant organic substance within humus to breakdown. It is assumed that this humin derives from plant cell wall components of decaying matter (Haynes and Francis 1993; Lal 1997). However, this has not yet been demonstrated. Isolated humin, derived from alkali treatments (using KOH) of humus, could form aggregates between the mineral fragments of soil with organic matter, which were water stable (Metha et al. 1960; Chenu and Guerif 1989; Cheshire 1990). However, the term humin remains vague with no characterisation of the macromolecules within humin, or a standard method of isolation. Furthermore, the effects of humin in aggregating soil remains controversial with issues of repeatability in published work due to the heterogeneity of the humus sampled, and the resulting humin extracted (Lehmann and Kleber 2015).

1.6.2 Polysaccharide-derived soil aggregates

It has been proposed that decomposing organic matter from plant debris, bacterial, algal and fungal debris and secretions can form water stable aggregates in soil (Oades, 1984; Six et al. 1999; Carrizo et al. 2015). Polysaccharides are thought to be the adhesive binding soil particles together through weak hydrogen bonds between the carboxyls (negatively charged), and to a lesser degree the hydroxyls on the released molecules and the cations (typically Ca2+, K+ and Mg2+) that adsorb to negatively charged surface of clay minerals (Figure 1.4; Foster 1981; Traore et al. 2000; Olsson et al. 2011). Clay particles (defined as: ≤2 µm) are formed from amorphous Fe and AI oxides and hydroxides, or from naturally occurring sheet-like structures formed from repeating silicate or aluminium oxide units (Greenland 1965; Fitz Patrick 1993). Without the presence of organic deposits, the binding of clay particles is reduced, highlighting the importance of organic residues in the formation of aggregates (Oades 1993; Rasse et al. 2005). In addition to hydrogen bonds, polysaccharides may even interact with clay particles through electrostatic bonds or Van der Waal interactions (Cheshire et al. 1985; Cheshire 1990). Once more polysaccharides become available, more clay particles can bind together forming microaggregates (defined as: ≤250 µm), which then adhere to other microaggregates and larger particles, forming larger more stable macroaggregates (defined as: ≥250 µm; Tisdall and Oades 1982; Vaughan and Malcolm 1985; Foster 1981; Tisdall 1994).

Diagram of how soil sticks to together

Figure 1.4 I Schematic of polysaccharide-clay binding leading to the formation of microaggregates

Clay particles have a mostly negative charge which attracts cations (typically Ca2+, K+ and Mg2+) in soil through electrostatic interactions. These cations then form a bridge with the negatively charged carboxyl and hydroxyl groups present on polysaccharides released by plant roots. After sticking, polysaccharides bond other clay particles together, bridging the gaps between each particle; thus increasing soil particle aggregation.

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Within most fertile soils, the presence of acidic polysaccharides is well documented (Edwards and Bremner 1967; Clapp and Davis 1970; Cheshire 1990). Acidic polysaccharides are more charged, containing higher levels of carboxyls, compared to their neutral counterparts, and therefore, possess a stronger capacity to bind to clay particles (Figure 1.4; Edwards and Bremner 1967; Cheshire 1990). This increases the capacity for polysaccharide-clay particle binding. Neutral polysaccharides, derived from bacterial exopolysaccharides have been shown to have weak capacity for polysaccharide-clay particle binding (Clapp and Davis 1970; Cheshire 1990). Using transmission electron microscopy, these neutral polysaccharides, could readily bind to the flat surface of clay particles but did not contain any carboxyls for clay binding (Olness and Clapp 1973; Chenu and Guerif 1989). The stability of aggregates is thought to be proportional to the molecular weight of the polysaccharides (Cheshire 1990). The larger the molecule the more carboxyls there are to form more polysaccharide bridges that link clay particles, thus stabilising their attachment (Figure 1.4; Chenu and Guerif 1989).

Polysaccharide adherence to clay particles is believed to prevent enzymatic degradation by soil microorganisms (De Booth et al. 1984; Cheshire 1990). However, the carboxyls and hydroxyls that are not bound to the cations on clay particles may be vulnerable to enzyme degradation (Cheshire et al. 1983). This would reduce aggregate stability. The gradual degradation of the glycosidic bonds within the polysaccharides would correspond to a gradual loss of aggregate stability (De Booth et al. 1984). One of the key factors for polysaccharide-derived aggregates is the viscosity and the solubility of the polysaccharide. For aggregate formation to occur, polysaccharides within soils must be water-soluble so that they can diffuse to the clay particles.

Limited evidence has shown that root mucilage sourced directly from the root caps of maize in situ was able to form stable soil aggregates when added to fresh soil (Tisdall and Oades 1982; Morel at al. 1991; Watt et al. 1993). Once formed, these aggregates caused by maize root mucilage were found to be water stable, as determined by wet sieving. Wet sieving fractionates soil into defined sizes through sieves that are shaken and flooded with tap water (Brown 2008). When sodium periodate (NaIO4), which oxidises the glycosidic bonds within polysaccharides, was introduced to the soil the isolated mucilage was rapidly degraded. There was also a significant reduction in the abundance of soil aggregates (Tisdall and Oades 1982). Other studies using a modelled form of root mucilage, polygalacturonic acid (commercial standard of pectin), uronic acid and glucose, have also demonstrated an increase in the abundance of soil aggregates (Metha et al. 1960; Reid and Goss 1981; May et al. 1993). However, this modelled root mucilage was less effective compared with the isolated maize root mucilage (Tisdall and Oades 1982; Morel at al. 1991; Watt et al. 1993). This suggests that as well as decaying matter, plants could be continually adding carbon into soils without the need for decay. The identities of the polysaccharides present within the isolated root mucilage were not determined nor the total glycan concentrations of each root mucilage collected. These initial investigations show a potential new source of carbon that could be being continually released into soils. Although the proportion of a plant’s carbon which is generated and then released into the soil remains hotly debated with some figures suggesting between 10% and 40% (Walker et al. 2003; Jones et al. 2007).

An investigation determined that cellulose, from decaying matter, was a strong source of soil macroaggregates (Mizuta et al. 2015). As cellulose is not charged (containing no carboxyls) contact between the hydroxyls, which have a weaker bind affinity to the cations on the silica sheets of clay, makes cellulose a poor microaggregator (Cheshire et al. 1979; Cheshire 1990). This suggests that not all polysaccharides released by decaying plant matter may be responsible for aggregate formation. The identity of polysaccharides released from decaying matter, which can increase micro- and macroaggregates remains undetermined. Furthermore, there has been no study to determine the binding affinities of the polysaccharides known to be released from plants, in the form of root mucilage, to clay particles. The only major known polysaccharides released by roots, in the form of root mucilage, are AGP and pectin (Bacic et al. 1986; Ray et al. 1998, Osborn et al. 1999). It may be possible that these polysaccharides are involved soil aggregation.

Soil microbiota such as bacteria, fungi and algae exude exopolysaccharides into the bulk soil, which is outside the influence of roots (Oades 1978; Tisdall and Oades 1982; Oades 1993). These microorganisms release a matrix of extracellular polysaccharide that they can also potentially aggregate soil mineral particles (Griffiths and Burns 1972; Tisdall 1994; Flemming and Wingender 2010). It is believed that the porosity caused by microaggregates in soil forms a refuge for microorganisms, which are vulnerable to predation (Rillig et al. 2015; Rillig et al. 2017). Additionally the interior of these microaggregates have been show to differ to that of the bulk soil, forming microenvironments favourable for microbe growth (Rillig et al. 2017. Therefore, it is in the interests of the microorganisms to maintain microaggregates.

It is hypothesized that microorganisms release exopolysaccharides so that they can adhere to microaggregates, and even help to form them as a secondary effect by binding to clay particles (Griffiths and Jones 1965; Griffiths and Burns 1972; Rillig 2004). Interestingly, microorganisms including rhziobacteria (Rhizobium leguminosarum and R. fluorescens) and arbuscular mycorrhizal fungi (Glomus perpusillum) have been demonstrated to use root mucilage isolated from maize and pea as a source of energy (May et al. 1993; Knee et al. 2001; Jones et al. 2000; Gunina and Kuzyakov 2015; Sun et al. 2015). In effect these microorganisms were found to increase their production of exopolysaccharides when their medium supplemented with maize root mucilage. This increase in secretion could enhance the abundance of soil microaggregates.

There is only one clear example of an exopolysaccharides released from a microorganism that forms soil aggregates. The bacteria Xanthomonas campestri has been reported to release xanthan to form an extracellular matrix, adhering cells together in order to shuttle resources required growth (Czarnes et al. 2000; Chang et al. 2015). When xanthan, which is formed of long repeating subunits of glucose, mannose and glucuronic acid was collected from cultures of Xanthomonas and added to soil, it was demonstrated to increase the abundance of microaggregates (Chang et al. 2015). The increase in microaggregates was shown to be comparable to that of isolated root mucilage, modelled root mucilage, uronic acid and glucose which were also added to soil (Metha et al. 1960; Reid and Goss 1981; Tisdall and Oades 1982; Morel at al. 1991; May et al. 1993). Another similarity to root-derived aggregates is that bacterial-derived aggregates remain stable for weeks after the bacteria have undergone senescence (Griffiths and Jones 1965; Chang et al. 2015). This aggregation effect increases the uptake of water and nutrients from the soil to bacteria, as in the case of roots, and stabilises the microaggregates that bacteria inhabit (Cheshire 1979). Another example of an exopolysaccharide from a microorganism that has been shown to cause soil aggregate is the disputed glycoprotein glomalin (Wright and Upadhayya 1997). Some believe glomalin is secreted by the hyphae and spores of arbuscular mycorrhizal fungi (Glomus spp; Rillig 2004; Rillig et al. 2015). It is suggested that glomalin adheres the hyphae network to the surface of roots as well as the surrounding soil resulting in microaggregates (Rillig 2004; Spohn and Giani 2010; Rillig et al. 2015). However, glomalin has not been defined and nothing as yet is known about its component parts or structure.

1.5 Aims and objectives

To date, the hydroponic medium of plants has never been probed using MAbs. This investigation aims to develop an understanding of the biochemistry and function of polysaccharides released from the roots of crop species, with a focus on wheat (Triticum aestivum). A hydroponic system was developed in order to isolate the polysaccharides in preparation for immunochemical analysis. There are three key objectives that were followed throughout this investigation:

1. Identify the polysaccharides released from the roots of a variety of crops using a hydroponic system and monoclonal antibodies.

2. Study the biochemistry of the major polysaccharides released by crop species.

3. Determine possible functions of polysaccharides released by roots in relation to soil aggregate status.

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