A multi-polysaccharide complex is released by the roots of cereal

4.1 Introduction

Our understanding of the biochemical properties of polysaccharides released by plant roots remains largely incomplete. Difficulty in collecting sufficient material from roots has remained a limiting factor in deciphering the biochemistry and structure of these polysaccharides. In order to explore micro-structures, polysaccharide complexes, within the cell wall matrix highly sensitive techniques have been developed, namely, Epitope Detection Chromatography (EDC) amongst other immunochemical assays. EDC utilises chromatography, including anion-exchange chromatography and size-exclusion chromatography, to elute fractions of polysaccharides, which are then probed with a range of monoclonal antibodies through ELISA (Cornuault et al. 2014). A gradient of sodium chloride is used during anion-exchange EDC to separate polysaccharides by charge. For instance polysaccharides that elute prior to sodium chloride have no charge as no salt was required to elute them. Highly charged polysaccharides, including pectin require high amounts of salt to be eluted (Figure 4.1). During size-exclusion EDC polysaccharides are eluted by size. Large polysaccharides have less interaction with the column matrix as they cannot pass through the small pores, thus eluting early. Small polysaccharides and oligosaccharides are able to pass through the pore spaces and thus elute later.

EDC can also be used to explore intra-molecular interactions, sub-families of polysaccharides, namely pectin, and multi-polysaccharide complexes. This technique can also be used on the nanogram and microgram scales, which is useful for deciphering polysaccharide-polysaccharide and polysaccharide-protein links as well as multi-polysaccharide complexes that may be present as minor components of the cell wall architecture (Cornuault et al. 2015). In addition to EDC, sandwich-ELISA can be used to explore co-linked polysaccharides. Within this technique, a CBM is used to coat a microtitre plate, which binds to one co-linked partner. Samples containing this hypothesized intra-molecular interaction are then added to the plate. MAbs are then used to probe the sample (Cornuault and Knox 2014). If there is a link between the partners then there will be a signal. These tools when combined form a powerful approach to determine the presence of these nanogram to microgram scale multi-polysaccharides complexes.

This chapter explores the biochemical properties of polysaccharides released into the concentrated hydroponate of wheat cv. Cadenza (hereon referred to as Cadenza) using these highly sensitive techniques, with a focus on anion-exchange EDC. A survey on the root cell walls as well as the polysaccharides released by wheat roots was conducted. To confirm the observations made by the antibody-based approach a glycan sample was provided to the Complex Carbohydrate Research Centre (CCRC) to undertake monosaccharide composition and monosaccharide linkage analyses. The biochemical properties of other cultivars of wheat, Avalon and Skyfall were compared to Cadenza along with other species of cereal, maize and barley, and eudicotyledons, pea, rapeseed and tomato, and a basal land plant, liverwort.

Figure 4.1 I Schematic diagram that shows anion-exchange EDC

A salt gradient (0.6 M) is run after a sample containing polysaccharides is injected into an anion-exchange column. Within the column, the column matrix is positively charged, which attracts the polysaccharides that have a negative charge due to the presence of carboxyls. As the salt gradient is increased, the chloride ions, which are negatively charged, elute the polysaccharides that are retained within the column. This elution is caused by the chloride ions outcompeting with the polysaccharides bound to the matrix. Polysaccharides that are slightly negative are eluted with little salt, and polysaccharides that are highly negative are eluted with high amounts of salt. The resulting fractions are collected and then transferred into 96 microtitre plates. The microtitre plates are subsequently developed using the ELISA protocol.

4.2 Results

4.2.1 Four released epitopes co-eluted during anion-exchange EDC

To explore the biochemistry of the released polysaccharides, Cadenza concentrated hydroponate (50 µg) was suspended in 20 mM sodium acetate buffer (Figure 4.2). The buffer with the hydroponate was injected into a weak 1 mL anion-exchange chromatography column. A step gradient of sodium chloride (0.6 M) was used to elute the polysaccharides within the concentrated hydroponate, into 1 mL fractions. Aliquots (40 µL) of 1 M sodium carbonate were then placed into each fraction to increase the pH. This facilitated the binding of the molecules within the sample to the wells of a microtitre plate for ELISA. MAbs were used to screen the wells of the ELISA plate. There were two peaks of LM25 (xyloglucan) neutral (arrow 1) which eluted before the salt gradient, and acidic (arrow 2) which eluted as the salt gradient was increased. Within the second acidic peak of xyloglucan epitopes, there was a clear co-elution of the AGP, extensin and xylan epitopes (Figure 4.2, A). When comparing the commercial standard of xyloglucan, tamarind seed xyloglucan, with the xyloglucan in the concentrated hydroponate, the xyloglucan within the hydroponate appeared to be acidic (Figure 4.2, B). As tamarind seed xyloglucan eluted prior to the salt gradient it indicated that the commercial standard was neutral unlike the xyloglucan from the hydroponate which required ~300 mM NaCI to elute from the column.

Figure 4.2 I Anion-exchange EDC analysis of polysaccharides released from the roots of Cadenza

Fifty micrograms of concentrated Cadenza hydroponate was injected into a 1 mL weak anion-exchange chromatography column. Aliquots of 100 µL (containing ~5 µg) of each collected fraction were assayed for each MAb. A step gradient of 0.6 M of NaCI was used (A). Two forms of xyloglucan, neutral (1) and acidic (2), were detected from the released polysaccharides. A co-elution of AGP, extensin and xylan was present within the second peak of xyloglucan (2) which was acidic. Data shown are a mean of three biological replicates. (A). Tamarind seed xyloglucan (100 ng) was injected into the same 1 mL anion-exchange column (B). The tamarind seed xyloglucan eluted before the salt gradient, indicating that the polysaccharide was neutral (B). Data shown are a mean of three biological replicates. ELISA absorbance values were determined by measuring the absorbance at 450 nm.

To examine the co-elution in more detail, samples (50 µg) of Cadenza concentrated hydroponate were passed through an anion-exchange chromatography column (1 mL). Samples were eluted using a delayed gradient of sodium chloride (0.6 M). This gradient had a steeper increase in salt compared to the gradient previously utilised (Figure 4.2) in an attempt to tease a part the neutral and acidic fractions. The neutral (arrow 1) and acidic (arrow 2) forms of xyloglucan remained for longer within the column when using the delayed gradient of salt. The acidic form of xyloglucan was retained for longer within the column compared to using the step gradient. Additionally, the amount of salt needed to elute the co-elution remained the same, ~300 mM. The co-elution of AGP, extensin and xylan epitopes also remained within the acidic form of xyloglucan (Figure 4.3). The signal of LM11, which binds to the epitopes of xylan, had slightly increased compared to the other gradient used (Figure 4.2, A and Figure 4.3). This increase was observed throughout the three biological replicates. Delaying the gradient of salt divided the neutral and acidic peaks, perhaps reducing molecular crowding on the ELISA plates, which probably account for this increase in signals.

Figure 4.3 I The co-elution of AGP, extensin, xylan and xyloglucan was retained for longer within the column when the gradient of salt was delayed

Epitope detection chromatogram of 50 µg concentrated Cadenza hydroponate. The chromatogram reveals that the two forms of xyloglucan; neutral xyloglucan (fractions 0-16; 1) and an acidic xyloglucan (fractions 40-60; 2) that was detected using LM25 was retained in the column for longer before the salt gradient. The co-elution of extensin (LM1), AGP (LM2) and xylan (LM11) also remained stable when delaying the salt gradient. The acidic peak of xyloglucan was retained for longer within the anion-exchange column under the longer neutral gradient. Data shown are a mean of three biological replicates. ELISA absorbance values were determined by measuring the absorbance at 450 nm.

4.2.3 Acidic and neutral forms of xyloglucan were isolated

Using the delayed gradient of salt, the neutral and acidic forms of xyloglucan could be isolated (Figure 4.4). Samples of concentrated hydroponate (3 mg) were suspended in 20 mM sodium acetate buffer, and injected into a larger, 15 mL weak anion-exchange chromatography column. Fractions of 15 mL were collected, and assayed using LM1 (extensin), LM2 (AGP), LM11 (xylan) and LM25 (xyloglucan). Increasing the throughput by 15x enabled the isolation of these two forms of xyloglucan. The co-elution remained with what appeared to be the acidic xyloglucan (Figure 4.4, B). Furthermore, similar patterns of the epitopes within the co-elution remained stable after isolation. The neutral form of xyloglucan also remained unchanged when compared to previous assays (Figure 4.2, A and Figure 4.4, A). Further examination of the anion-exchange chromatograms (Figure 4.2, Figure 4.3 and Figure 4.4), including shallower salt gradients (not included), revealed that the co-elution of REC1 was eluted with a salt concentration of ~300 mM. This co-elution of four epitopes, AGP, extensin, xylan and xyloglucan from the roots of Cadenza is referred to as Root Exudate Complex 1 (REC1).

Figure 4.4 I Isolation of the neutral and acidic forms of xyloglucan

Anion-exchange EDC of 5 µg isolated neutral fractions (A) and 5 µg acidic fractions (B). Three milligrams of concentrated wheat hydroponate was injected into a 15 mL column. After collecting, samples were freeze-dried, dialysed and re-run through anion-exchange EDC. Fifty micrograms of isolated fractions were injected into a 1 mL column for this analysis. After the re-run both the neutral fractions (A) and acidic fractions (B) were detected independently. This co-elution of AGP (LM2), extensin (LM1), xylan (LM11) and xyloglucan (LM25) within the second acidic peak is referred to Root Exudate Complex 1 (REC1). Each data point is a mean of three biological replicates. ELISA absorbance values were determined by measuring the absorbance at 450 nm.

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4.2.4 Size-exclusion EDC demonstrated high heterogeneity within the sizes of the polysaccharides in REC1

To examine the sizes of the polysaccharides released by Cadenza roots (REC1), and within its root body, size-exclusion EDC was undertaken that used a different column matrix. Isolated REC1 (100 µg; Figure 4.4) as well as extracted polysaccharides from the root body (50 µg) were injected into a 120 mL size exclusion column, and eluted using 1 M NaCI. The resulting elutions were collected into 96 1 mL fractions. Aliquots (40 µL) of 1 M sodium carbonate were then placed into each fraction to increase the pH, which helped to facilitate the binding of the molecules within the sample to microtitre plate wells for ELISA. The larger the molecule, the lower amount of interaction that molecule will have with the column as it percolates, thus large molecules will elute and appear early on the chromatogram. From the analysis, there was a large range of sizes within the polysaccharides of REC1 (Figure 4.5, A) and the root body (Figure 4.5, B). In particular, there was a wide size range of xyloglucan molecules that were in REC1 when compared to the commercial standard, tamarind seed xyloglucan (Figure 4.5). This was also reflected by the range of xylan molecules detected released by wheat roots. There is a small signal of LM2 (AGP), which eluted just before fraction 40. There is also a large signal of LM1 (extensin), between fractions 40 and 65 (Figure 4.5). When examining the root body, there were two clear peaks of xylan, which was reflected by a lower signal trace of xyloglucan (Figure 4.5, B). There were no signals from LM1 and from LM2 in the root body. The xyloglucan that was contained within the root body and the hydroponate considerably varied to tamarind seed xyloglucan (Figure 4.5). Unexpectedly, the chromatographic trace of tamarind seed xyloglucan (Figure 4.5, B) appeared similar to that of the anion-exchange EDC. This shows that the xyloglucan within tamarind seed tested was very high in size (Figure 4.2).

Figure 4.5 I Complex heterogeneity in polysaccharides size is present within REC1 and the root body

Size-exclusion EDC revealed a large range of differentially sized AGP (LM2), extensin (LM1), xylan (LM11) and xyloglucan (LM25) molecules in REC1 (A) and root body (B). This range of differentially sized polysaccharides also differs between what is present within the root cell walls (B) compared to what is released (A). Tamarind seed xyloglucan (100 ng) screened using LM25 was also injected into the size exclusion column with the same data inserted onto the above. Tamarind seed xyloglucan (grey) is a large molecule that contains fewer smaller domains compared to the hydroponate (A) and root body (B). Samples of the acidic co-elution (100 µg) and root body cell wall extract (50 µg) were each dissolved into 1 mL of high salt buffer, and injected into a 120 mL size exclusion column. The resulting fractions were directly probed with the MAbs. Each data point is a mean of two biological replicates. A gel filtration calibration kit containing five protein standards was used to calculate KDa values displayed. ELISA absorbance values were determined by measuring the absorbance at 450 nm, V0 = void volume.

4.2.5 Total carbohydrate analysis revealed the presence of undetected glycan within the anion-exchange EDC

To determine the total glycan content of the concentrated hydroponate of Cadenza, a Phenol-Sulphuric Acid assay was carried out on fractions of the anion-exchange EDC analysis (Figure 4.3). For this assay, 50 µL from each of the 96 fractions collected during anion-exchanged EDC were removed. Aliquots (50 µL) of concentrated sulphuric acid and 5% (w/w) phenol in dH₂O (30 µL) were sequentially added. Glucose was used to convert the absorbance units into total µg per fraction. When all fractions of the anion-exchange EDC were totalled, there was 300 µg of glycan detected. As there was 300 µg used for the anion-exchange EDC analysis, this indicates that the total concentrated hydroponate of Cadenza is formed of glycan (Figure 4.6). Within the neutral region of the chromatogram there was a high detection of total carbohydrate. This total carbohydrate was present within the neutral xyloglucan fractions (1-15), and between the neutral xyloglucan fractions and REC1 fractions, marked X (Figure 4.6). This second region of total neutral glycan (16-35) did not follow the MAb peaks on the chromatogram trace. There was also a third peak of total carbohydrate (40-60), which was within the acidic region of the chromatogram (Figure 4.6).

Figure 4.6 I Total carbohydrate analysis of the concentrated hydroponate of Cadenza using the anion-exchange EDC system

Aliquots (50 µL) of concentrated hydroponate were collect from each 1 mL fraction from the same anion-exchange EDC assay within Figure 4.3. An overlay of the anion-exchange EDC results from Figure 4.3 has been added. The Phenol-Sulphuric Acid fractions were read using the absorbance at 450 nm. From this analysis, there were three forms of total carbohydrate. Fractions 1 to 15, and fractions 16-35 show high concentrations of carbohydrate within the neutral region. The second neutral form of total carbohydrate does not match the two peaks detected within the EDC (X). The third form of total carbohydrate, between fractions 40 and 60, reveals the presence of acidic carbohydrates. Data are a mean of three biological replicates.

4.2.6 Sandwich-ELISA confirms potential linkages of REC1

To further explore if the co-elution of the major polysaccharides released by the roots of Cadenza, as seen in anion-exchange EDC, were linked a sandwich-ELISA was undertaken. Microtitre plates were coated with a xylan-specific CBM (CBM2b1-2; McCartney et al. 2006), which also bound to the xylan released by Cadenza (Figure 4.7, A). Other polysaccharides that were linked with the xylan released by Cadenza would bind to the CBM-coated wells, generating MAb signals. The MAbs, LM1 (extensin), LM2 (AGP), LM11 (xylan) and LM25 (xyloglucan) along with the secondary anti-rat HRP were used to reveal the presence of any linked polysaccharides. Negative controls, wells with no CBM2b1-2, were directly compared to wells with CBM2b1-2 to verify the signal for a possible link between polysaccharides. The sandwich-ELISA used 10 µg/mL of the isolated acidic fractions (Figure 4.7, B), and determined that the major epitopes released by Cadenza were linked as a part of a putative multi-polysaccharide complex (Figure 4.7, B).

The signals from the top MAbs in the wells containing CBM2b1-2 were all significantly higher compared to the negative controls (Two-Sample T-Test, T= 31.43, P= <0.05). Therefore, supporting the presence of REC1 (Figure 4.7, B). The signal from LM11 confirmed that the CBM2b1-2 was binding to xylan.

Figure 4.7 I Sandwich-ELISA confirms the presence of REC1

Schematic diagram representing a sandwich-ELISA analysis (A). The xylan-specific carbohydrate binding module, CBM2b1-2, is coated onto a 96 well microtitre plate. After coating, a sample of hydroponate is incubated on the CBM2b1-2 coated plate. Any xylan present within a multi-polysaccharide complex binds to the CBM. The primary anti-rat MAb along with the secondary anti-rat HRP antibody are incubated onto the CBM coated wells revealing the presence of other epitopes that are bound to the xylan epitopes (A). The sandwich-ELISA demonstrates that AGP, extensin, xylan and xyloglucan epitopes were a part of a complex (B). CBM2b1-2, which binds to xylan, was used to coat the microtitre plates, negative = no CBM control. Concentrated hydroponate (10 µg/mL) was used for the analysis (B). Each data point is a mean of three biological replicates. ELISA absorbance values were determined by measuring the absorbance at 450 nm. Standard deviation bars are shown; asterisks indicates significant difference (**P= 0.001 and ***P= 0.0001).

4.2.7 The epitope profile on anion-exchange EDC of the root body differs to that of the hydroponate

In order to see if REC1 was present within the root body of Cadenza, the biochemical properties of the cell walls were explored. To determine the profile of the root body, root cell walls were extracted. Roots were frozen by liquid nitrogen and freeze-dried. Fifteen milligrams of freeze-dried root material was dehydrated by adding increasing amounts of EtOH (v/v; 70%, 80%, 90% and 100%), and then acetone (100%) and methanol and chloroform (2:3), which removed the non-polysaccharide components of the root body. After removing these soluble materials, 1 mg of root cell walls was extracted using 4 M KOH and 1% sodium borohydride. This final material was assayed using LM1 (extensin), LM2 (AGP), LM11 (xylan) and LM25 (xyloglucan). For the EDC analysis 50 µg was injected into an anion-exchange column, and for the sandwich-ELISA analysis 10 µg/mL was used. The EDC chromatogram revealed that there was a range of xylan forms within the cell walls of Cadenza roots (Figure 4.8). Within the first peak of xylan, there is a co-elution of neutral xyloglucan epitopes. Within the second peak, there is a co-elution of AGP, extensin, xylan and xyloglucan epitopes, which is similar to that of REC1 that was detected within the concentrated hydroponate (Figure 4.2). There are no co-elutions of the major epitopes within the other peaks of xylan. The sandwich-ELISA analysis demonstrates that the putative multi-polysaccharide complex may also present within the root cell walls (Figure 4.8, 2). Signals from the MAbs in the wells containing CBM2b1-2 were all significantly higher to that of the negative controls (Two-Sample T-Test, T= 27.33, P= <0.05), supporting the presence of REC1 within the root body of Cadenza (Figure 4.4). The signal from LM11 confirmed that the CBM2b1-2 was binding to xylan.

Figure 4.8 I EDC and sandwich-ELISA analyses of Cadenza root cell walls reveal a range of acidic forms of xylan

Fifty micrograms of root cell wall material was injected into a 1 mL anion-exchange chromatography column for EDC. Aliquots of 100 µL (containing ~5 µg) of each collected fraction were assayed for each MAb (A). The chromatogram revealed a peak of LM25 that was co-eluting with AGP (LM2), extensin (LM1) and xylan (LM11; 2). Five clear peaks of LM11 dominate the chromatogram, which range from neutral to highly acidic (A). Data are a mean of three biological replicates. ELISA absorbance values were determined by measuring the absorbance at 450 nm. The sandwich-ELISA reveals that REC1 released by wheat roots is also present within the root body but at potentially lower levels compared to the concentrated hydroponate (B). CBM2b1-2, which binds to xylan, was used to coat the microtitre plates, negative = no CBM control. Root cell wall material (10 µg/mL) was used for the analysis (B). The antibody LM11 which binds to xylan was included as a control for CBM2b1-2. There was strong binding of xyloglucan (LM25) to xylan (CBM2b1-2) along with weaker binding of AGP (LM2) and extensin (LM1) to xylan (CBM2b1-2; B). Each data point is a mean of three biological replicates. ELISA absorbance values were determined by measuring the absorbance at 450 nm. Standard deviation bars are shown; asterisks indicates significant difference (*P= <0.05, **P= 0.001 and ***P= 0.0001).

4.2.8 REC1 was not disrupted after xylanase and xyloglucanase digests

As REC1 putatively contains xylan and xyloglucan it was decided to degrade them in order to explore their potential linkages, and how removing them would affect the behaviour of REC1 on the chromatographic system. Xylan and xyloglucan were selected as there are a range of enzymes, xylanase and xyloglucanase, which are available to degrade them unlike the other domains of REC1, AGP and extensin. Aliquots (50 µL) containing 50 µg of concentrated hydroponate was dissolved in 100 mM sodium acetate buffer. Hydroponates were incubated at 40°C and 60°C for 2 h with β-xylanase (480 U) and xyloglucanase (40 U). After digesting, samples of hydroponate were assayed with the MAbs. When the enzymes were added to the concentrated hydroponate their respective targets were successfully digested. This was shown by the absence of the signals of LM11 (xylan), and LM25 (xyloglucan; Figure 4.9). Surprisingly the co-elution of the AGP, extensin, xylan and xyloglucan epitopes remained unaffected after the enzyme treatments, which digested the xylan and xyloglucan domains of REC1 (Figure 4.9). This demonstrates that degrading the xylan and xyloglucan did not affect the co-elution that forms REC1, suggesting that they were not acidic domains.

Figure 4.9 I Xylanase and xyloglucanase digests of Cadenza concentrated hydroponate

Concentrated hydroponate (50 µg) was dissolved in sodium acetate (100 mM) buffer along with BSA (1 mg/mL). Enzymes, β-Xylanase M1 (60 µg/mL), which degrades (1,4)-β-D-xylosidic linkages in xylan and xyloglucanase (120 µg/mL), which degrades 1,4-β-D-glucosidic linkages in xyloglucan were added to 1 mL of sample for 10 mins. Signals of LM11 (xylan) and LM25 (xyloglucan) were not detected when their respective enzymes were added showing that the treatments were effective. Each treatment had a maximum absorbance of 1.0 au. A 1:10 dilution of antigen was used to get signals of 1.0 au for LM25. Each mark on the y axis represents 0.5 au relative to each group. Each data point is a mean of three biological replicates. Relative ELISA absorbance values were determined by measuring the absorbance at 450 nm.

4.2.9 Co-elutions of the major epitopes were detected within the hydroponates of other wheat cultivars

After investigating some aspects of the biochemical properties of REC1 released by Cadenza, two other cultivars of wheat were analysed. Wheat cultivars, Avalon and Skyfall, were grown using the same parameters and hydroponic system as used for Cadenza. These cultivars were chosen as they are widely grown winter varieties whereas, Cadenza is an older and less popular spring variety (Kumar et al. 2011). After two weeks of hydroponic culture, the hydroponates were concentrated, dialysed and freeze-dried. Fifty micrograms of Avalon and Skyfall concentrated hydroponates were then assayed using EDC. The chromatograms reveal that were are two forms of xyloglucan, neutral and acidic released into the hydroponates of Avalon and Skyfall. Within the neutral form of xyloglucan no other epitopes were detected from the initial survey (Figure 4.10). The fractions containing the second, acidic form of xyloglucan contained a co-elution of AGP, extensin and xylan (Figure 4.10), as previously observed in the hydroponate of Cadenza (Figure 4.3).

When the salt gradients are delayed the acidic peaks of LM25 (xyloglucan), along with the co-elution were retained for longer within the anion-exchange column (Figure 4.10). This demonstrates that the co-elution was unaffected when the salt gradient was delayed (Figure 4.10), as previously observed in Cadenza (Figure 4.3). The highest signal detected from the MAbs was from LM25 followed by LM1, LM11 and LM2 (Figure 4.10). There were increases in the signals of the MAbs within the co-elution of AGP, extensin, xylan and xyloglucan when the gradient of salt was delayed for both Avalon and Skyfall (Figure 4.10). This was also observed from the hydroponate of Cadenza (Figure 4.3), perhaps due to reducing molecular crowding. Additionally, there were slight differences in the hydroponic profiles between the cultivars, for example, the signals of LM1 were higher, ranking the second highest, within the concentrated hydroponates of Avalon and Skyfall (Figure 4.10) compared to Cadenza where LM11 was the second highest signal (Figure 4.3). These slight differences suggest that Avalon and Skyfall released a similar molecule to REC1, which contained different components.

Figure 4.10 I Two forms of xyloglucan are released by the wheat cultivars Avalon and Skyfall as determined using Anion-Exchange EDC

Concentrated hydroponates of Avalon and Skyfall (50 µg) were injected into a 1 mL anion-exchange chromatography column. Five micrograms of this was assayed for each MAb. A step gradient of 0.6 M of Na⁺CI^- was used (figures on the left) followed by a long neutral gradient of 0.6 M of Na⁺CI^- (figures on the right). Two forms of xyloglucan (LM25), neutral and acidic were detected in both cultivars along with a co-elution of AGP (LM2), extensin (LM1) and xylan (LM11) within the acidic form of xyloglucan (figures on the left). When the gradient was altered to have a longer neutral step the co-elution of LM1, LM2, LM11 and LM25 were retained in the column for longer (figures on the right). Each data point is a mean of three biological replicates. Relative ELISA absorbance values were determined by measuring the absorbance at 450 nm.

4.2.10 REC1-like molecule is released by the other cultivars of wheat

As a co-elution of AGP, extensin, xylan and xyloglucan was determined within the hydroponates of Avalon and Skyfall, which was similar to Cadenza, a sandwich-ELISA was undertaken to determine if they were linked. This was to determine if the co-elution of epitopes, previously determined were linked (Figure 4.10). Microtitre plates were coated with the xylan-specific carbohydrate-binding module, CBM2b1-2 (McCartney et al. 2006). LM1 (extensin), LM2 (AGP), LM11 (xylan) and LM25 (xyloglucan) were used to screen the microtitre wells with and without the CBM coating. Isolated acidic fractions (from Figure 4.10; 10 µg/mL) were added to each well. The signals with the CBM coating were all significantly higher to that of the wells that lacked CBM (Figure 4.11); for Avalon (Two-Sample T-Test, T= 36.47, P= <0.05), and for Skyfall (Two-Sample T-Test, T= 24.49, P= <0.05). These significant differences demonstrate that these cultivars of wheat release molecules similar to REC1. The signal from LM11 confirmed that the CBM2b1-2 was binding to xylan (Figure 4.11).

Figure 4.11 I Sandwich-ELISA reveals REC1-like macromolecules are released by Avalon and Skyfall

CBM2b1-2, which binds to xylan, was used to coat the microtitre plates, negative = no CBM control. Ten micrograms a millilitre of concentrated hydroponate was used. The sandwich-ELISAs reveal that xyloglucan (LM25), AGP (LM2) and extensin (LM1) epitopes were strongly bound to xylan (CBM2b1-2). LM11 that binds to the epitopes of xylan had the highest signal, confirming that the CBM was binding to xylan. Each data point is a mean of three biological replicates. ELISA absorbance values were determined by measuring the absorbance at 450 nm. Standard deviation bars are shown; asterisks indicates significant difference (**P= 0.001 and ***P= 0.0001).

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4.2.11 Similar hydroponic profiles were observed between wheat and other cereals

After uncovering a putative multi-polysaccharide complex released by wheat roots, other species of cereal were grown hydroponically to investigate the taxonomic occurrence of REC1 and REC1-related molecules. Barley and maize hydroponates were initially screened with 29 MAbs and found to contain the four epitopes, similar to that of wheat (Table 3.1 and Table 3.2). After concentrating, 50 µg of each species hydroponate was suspended in 20 mM sodium acetate buffer, and injected into a 1 mL weak anion-exchange chromatography column. Two gradients were used to explore alterations in the elution of the epitopes. The concentrated hydroponates of both barley and maize contained two forms of xyloglucan; neutral and acidic (Figure 4.12). Furthermore, both concentrated hydroponates contained a co-elution of AGP, extensin and xylan with the second acidic form of xyloglucan (Figure 4.12). There were slight differences in the signal strengths of each MAb within the co-elution. For barley, extensin epitopes ranked the highest followed by xyloglucan, xylan and AGP epitopes. Whereas, for maize, xyloglucan epitopes ranked the highest followed by xylan, AGP and extensin. These differences suggest that barley and maize release a macromolecule similar to REC1. When the salt gradient was extended the co-elution was retained for longer within the column (Figure 4.12). The rankings of the MAb signals remained the same during the delayed gradient of salt.

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Figure 4.12 I Two forms REC1-like macromolecules were detected within the hydroponates of barley and maize

Fifty micrograms of concentrated hydroponate of barley and maize were injected into a 1 mL anion-exchange chromatography column. Five micrograms of this was assayed for each MAb. A step gradient of 0.6 M of Na⁺CI^- was used (figures on the left) followed by a long neutral gradient of 0.6 M of Na⁺CI^- (figures on the right). Two forms of xyloglucan (LM25), neutral and acidic were detected in both species along with a co-elution of AGP (LM2), extensin (LM1) and xylan (LM11) within the acidic form of xyloglucan. Relative ELISA absorbance values were determined by measuring the absorbance at 450 nm.

4.2.12 Sandwich-ELISA evidence of a REC1-like macromolecule released by other cereals

As there was a co-elution of the four major epitopes detected within the hydroponates of barley and maize, a sandwich-ELISA was undertaken to determine if these co-eluting polysaccharides were linked. Microtitre plates were coated with CBM2b1-2 (McCartney et al. 2006), a xylan-specific binding agent. Samples of both barley and maize isolated acidic fractions (from Figure 4.12; 10 µg/mL) were incubated on the plates. Wells with no CBM coating were used as a negative control. MAbs were used to reveal the presence of any linked polysaccharides. The sandwich-ELISAs reveal that the four polysaccharide epitopes were linked as a part of a REC1-like complex, released by the roots of both barley and maize (Figure 4.13). The MAb signals were all significantly higher than their respective controls; for barley (Two-Sample T-Test, T= 27.43, P= <0.05), and for maize (Two-Sample T-Test, T= 20.66, P= <0.05). LM11 signals, in both species hydroponate, were significantly higher to their respective controls, indicating that the CBM was indeed binding to xylan (Figure 4.13). REC1 (Figure 4.3) and similar macromolecules to REC1 (Figure 4.13) have been detected within the concentrated hydroponates of three cultivars of wheat and three species of cereal (Figure 4.7 and Figure 4.11), which not only supports the release of REC1 macromolecules but highlights its potential biological importance as these macromolecules are being released by a range of grasses.

Figure 4.13 I Sandwich-ELISA analyses reveal the presence of REC1-like component released by barley and maize roots

CBM2b1-2, which binds to xylan, was used to coat the microtitre plates, negative = no CBM control. Ten micrograms a millilitre of concentrated hydroponate was incubated on the microtitre plates. The sandwich-ELISAs show for both barley and maize, xyloglucan (LM25), extensin (LM1) and AGP (LM2) were bound to the xylan-specific CBM, CBM2b1-2. For both species hydroponate, LM11 strongly bound to CBM2b1-2, confirming that the CBM was binding the xylan. Each data point is a mean of three biological replicates. Relative ELISA absorbance values were determined by measuring the absorbance at 450 nm. Standard deviation bars are shown; asterisks indicates significant difference P= <0.05.

4.2.13 Some evidence of co-eluting polysaccharide epitopes from eudicotyledons

Exploring the biochemical properties of cereal hydroponates revealed that AGP, extensin, xylan and xyloglucan were linked to form REC1. The major epitopes in the hydroponates of eudicotyledons (Table 3.2 and Figure 3.5) were examined to determine if these polysaccharides were linked to form a similar complex. Commonly cultivated eudicotyledon crops, pea, rapeseed and tomato, were grown in a 50:50 mixture of vermiculite and perlite. Seedlings were transferred into hydroponics for two weeks. After two weeks, their hydroponates were concentrated. Quantities of concentrated hydroponate, 100 µg of tomato, 200 µg of pea and rapeseed, were suspended in sodium acetate buffer, and injected into an anion-exchange column. Subsequent sandwich-ELISAs followed using 10 µg/mL of each concentrated hydroponate. The chromatograms of each eudicotyledon revealed highly diverse relative levels of the four major polysaccharide epitopes, AGP (LM2), extensin (LM1), xylan (LM11) and xyloglucan (LM25; Figure 4.14). From anion-exchange EDC, no such strong co-elutions were evident within the hydroponates of the eudicotyledons compared to the cereals (Figure 4.14). For all sandwich-ELISAs, LM11 was significantly higher compared to wells that contained no CBM2b1-2 (McCartney et al. 2006), demonstrating that CBM was binding to xylan.

The chromatogram of pea hydroponate reveals the presence of three forms of xyloglucan epitopes, neutral, acidic and highly acidic. There appears to be slight indications of three co-elutions of AGP, extensin and xylan with the three forms of xyloglucan (Figure 4.14). When the sandwich-ELISA was undertaken, the wells coated with CBM2b1-2 were all significantly higher compared to their respective controls (Two-Sample T-Test, T= 28.27, P= <0.05). However, when combined with the low signals on the chromatogram, there is weak evidence of a REC-like complex within the hydroponate of pea (Figure 4.14).

The concentrated hydroponate of rapeseed contained two distinct peaks of xyloglucan, neutral and acidic. The acidic form of xyloglucan was lower compared to the neutral form. This acidic peak of xyloglucan maybe present due to the root hairs of Arabidopsis, which is within the same family as rapeseed (Brassicaceae), containing an acidic form of xyloglucan, which contains a galacturonic acid residue (Peña et al. 2012a). There were also two peaks of xylan one of which slightly co-eluted with the acidic xyloglucan (Figure 4.14). AGPs were also present but did not co-elute with the other polysaccharide (Figure 4.14). A very low signal from LM1 (extensin) was present across the chromatogram. The sandwich-ELISA indicated that AGP, extensin and xylan could be linked as a part of a complex (Two-Sample T-Test, T= 45.53, P= 0.05). However, the signals from the sandwich-ELISA are only just significant, and since there are little evidence of co-eluting polysaccharides it cannot be confidently established that these polysaccharides were linked.

The chromatogram of tomato hydroponate reveals the presence of two forms of xyloglucan epitopes, neutral and acidic. There was a lower signal of LM25 (xyloglucan) detected within the acidic region compared to the neutral region of the chromatogram (Figure 4.14). There was a large signal of LM2 (AGP), which dominates the chromatogram of the concentrated hydroponate of tomato. Xylan was also present in a highly acidic form, eluting just before the AGP. There was a small indication that the four major polysaccharides may be linked by the weak signals that appear just before fraction 60. However, no clear co-elutions were evident across the chromatogram. This result was reflected by the lack of a strong signal of LM1, LM2 and LM25 within the sandwich-ELISA (Figure 4.14). The only signal that was significantly higher to that of the respective control is LM11, which demonstrates that CBM2b1-2 is binding to xylan.

Figure 4.14 I Anion-exchange EDC analysis of released polysaccharides from pea, rapeseed and tomato differs to that of grasses

Two hundred micrograms concentrated hydroponate of pea and rapeseed, and 100 µg of tomato were injected into a 1 mL anion-exchange chromatography column (figures on the left). Twenty micrograms of pea and rapeseed, and 10 µg of tomato hydroponates were assayed for each MAb; a long neutral gradient of 0.6 M of Na⁺CI^- was used. There are three weak co-elutions between AGP (LM2), extensin (LM1), xylan (LM11) and xyloglucan (LM25) within the pea hydroponate. Within the hydroponate of rapeseed, AGP (LM2) and xylan (LMM11) weakly to co-elute within the acidic region. AGP (LM2) dominates the hydroponate of tomato followed by two forms of xyloglucan and xylan (LM11). CBM2b1-2, which binds to xylan, was used to coat the microtitre plates of pea and tomato concentrated hydroponate (figures on the right). Ten micrograms a millilitre of concentrated hydroponate was incubated on the microtitre plates. Negative = no CBM control; standard deviation bars are shown. The sandwich-ELISAs show that xyloglucan (LM25), AGP (LM2) and extensin (LM1) binds to xylan (CBM2b1-2) within the pea hydroponate. The sandwich-ELISA demonstrates that the released polysaccharides from rapeseed, AGP (LM2) and extensin (LM1) bind to xylan (LM11). For tomato, no co-elution was demonstrated. Each data point for both the EDC and sandwich-ELISA results were a mean of three biological replicates. Relative ELISA absorbance values were determined by measuring the absorbance at 450 nm. Asterisks indicates significant difference (*P= <0.05 and **P= 0.001).

4.2.14 Rhizoids of liverwort release an putative AGP-xyloglucan complex designated REC2

The opportunity arose to examine the liquid medium of liverwort, which is an early diverging form of land plant. The ancestors of these liverworts were the first forms of plants to colonise the land over 470 millions of years ago (Kenrick and Crane 1997; Bateman et al. 1998). Comparing the biochemical properties of the liquid medium of liverworts to the concentrated hydroponates of more recent plants has provided a greater insight into the polysaccharides released by plants. Liverworts (Marchantia polymorpha) were grown and the biochemical proprieties of their exudates examined. Ten liverwort gemmae, cup-like appendage that form clones, were grown on agar with full BG11 medium for 30 days. After 30 days, gemmae were removed and the surrounding agar finely cut, and placed into an Eppendorf tube. Deionised water was placed into the tube, and the mixture agitated overnight to remove the exudates released into the agar. This water gel extract was then screened with 29 MAbs only revealing the presence of LM2, which bind to the epitopes of AGP and LM25 that binds to the epitopes of xyloglucan. There were no signals detected from LM11 (xylan), JIM7 and LM19 (HG), and LM6 (RG-I; work was undertook by Ms Bev Merry). After initially screening the agar gel extracts of liverworts, a tissue print was carried out using a nitrocellulose sheet, which was undertaken by Ms Sue Marcus. The nitrocellulose was probed with LM25, and revealed that xyloglucan was released along the rhizoids of liverworts as well as the rhizoid tips. Higher signals of LM25 (xyloglucan) were detected at the tips of rhizoids (Figure 4.15, D). Signals of LM2 (AGP) were also found to be higher at the tips of the rhizoids.

To further explore the biochemical properties of the AGP and xyloglucan released by liverwort into liquid medium, anion-exchange EDC and sandwich-ELISA were undertaken. After the liverworts have been grown in agar for a month they were removed and the agar extracted with dH2O. One millilitre of this wash was directly injected into a weak anion-exchange column (1 mL). An additional 1 mL of this wash was also incubated in microtitre wells coated with a xyloglucan- and cellulose-binding CBM, CBM3a (Blake et al. 2006). The anion-exchange EDC clearly shows two distinct forms of xyloglucan, neutral and acidic, released by the rhizoids of liverwort (Figure 4.15, B). The chromatogram also reveals a clear co-elution of AGP with the second acidic form of xyloglucan. This suggests that there is a possible complex formed of AGP and xyloglucan that was being released by the liverworts (Figure 4.15, B). This possible link between AGP and xyloglucan was confirmed by the strong MAb signals with the CBM3a coated wells, within the sandwich-ELISA (Two-Sample T-Test, T= 19.37, P= <0.05; Figure 4.15, C). Once determining the presence of a REC2, a basic phylogenetic analysis was carried out based on taxonomy, which determined that the last common ancestor between liverworts and cereals, which release these multi-polysaccharide complexes, lived over 400 million years ago (Figure 4.15, A). This indicates that releasing these putative complexes may be wide spread across the plant kingdom. Further examination of the anion-exchange chromatograms, revealed that the co-elution of AGP-xyloglucan was eluted with a salt concentration of ~500 mM. It was observed that the amount of salt required to elute REC2 (Figure 4.15, C) was more than what it was required to elute wheat REC1 (Figure 4.7, B) and REC1-like complexes released from the roots of barley and maize (Figure 4.12). This was most likely due to the presence of an acidic xyloglucan, which contains galacturonic acid residues found within liverworts (Peña et al. 2008).

Figure 4.15 I Analysis of liverwort (Marchantia polymorpha) agar gel extracts reveals a REC1-like complex, REC2

Phylogenetic tree analysis of the species used within this study including liverwort (A). The last common ancestor of the species used was the clade Embryophyta, which were the first land plants that evolved from green algae, specifically the Charophyte, which occurred 470 million years ago, indicated by red circle. This phylogenetic tree was created using the Phylot phylogenetic tree taxonomy generator (A). One millilitre of agar gel extract was injected into a 1 mL anion-exchange chromatography column. One hundred microliters of liverwort gel extract were assayed for each MAb. A long neutral gradient of 0.6 M of Na+CI- was used (B). A clear co-elution of AGP (LM2) and xyloglucan (LM25) can be seen between fractions 58 and 80. Two peaks of xyloglucan (LM25) are also visible (B). CBM3a, which binds to xyloglucan-cellulose, was used to coat the microtitre plates of liverwort gel extracts (C). One millilitre of liverwort gel extract was incubated on the microtitre plates. Negative = no CBM control; standard deviation bars are shown (C). The sandwich-ELISA clearly demonstrates that AGP (LM2) and xyloglucan (LM25) are bound to each other in a REC1-like complex (C). Each data point for both the EDC and sandwich-ELISA results were a mean of three biological replicates. ELISA absorbance values were determined by measuring the absorbance at 450 nm. Asterisks indicates significant difference P= <0.001.

Liverworts were grown for 7 days on agar with no nutrients by Ms Sue Marcus. Liverwort was imaged using a compound microscope (D, left). After growth, nitrocellulose sheets were placed onto the agar without the liverwort. The nitrocellulose sheet was then probed with LM25 (xyloglucan). The nitrocellulose print revealed the location of xyloglucan released along the rhizoids and tips of rhizoids (D, right). Scale = 1 mm; T = thallus.

4.2.15 REC1 contains monosaccharides that are abundant within the four major polysaccharides determined by MAbs

In order to verify the observations made within the immunochemical analyses, a preparation of REC1 was provide to the Complex Carbohydrate Research Centre (CCRC) for monosaccharide composition and monosaccharide linkage analyses. These biochemical analyses determined the major monosaccharides within the preparation. The CCRC hydrolysed the sample using 2 M trifluoroacetic acid, then reduced the sample using sodium borohydride (10 mg/mL) prepared in 1 M ammonium hydroxide, and finally, acetylated the sample using 9:1 (v/v) methanol and acetic acid. The resulting alditol acetates were analysed using gas chromatography combined with mass spectrometry. From the analysis, the sample was found to contain, in descending order of abundance, mannose, glucose, xylose, arabinose, rhamnose, galactose and fucose (Figure 4.16). The most abundant monosaccharide was mannose, forming 30.6% of the sample analysed (Figure 4.16), which was not previously determined by mannan-specific MAbs (Table 3.1). No acidic monosaccharide residues were detected within the analysis, which is also an unexpected result due to previous immunochemical analyses that suggest the presence of glucuronic acid present in AGP.

Figure 4.16 I Total ion chromatogram and summary of monosaccharide composition analysis of REC1

The preparation of REC1 (300 µg) was hydrolysed with 2 M trifluoroacetic acid for 4 h at 100°C. REC1 was then reduced with sodium borohydride (10 mg/mL) prepared in 1 M ammonium hydroxide for 1 h at RT. The preparation was then acetylated using 9:1 (v/v) methanol in acetic acid for 1 h at RT. The resulting alditol acetates were analysed using total ion chromatography combined with mass spectrometry. Inositol (20 µg) was used as an internal standard. The analysis reveals that mannose had the largest proportion (%) compared to the other monosaccharides within REC1. Glucose had the second highest proportion of REC1 followed by xylose, arabinose, rhamnose, galactose and fucose. This work was undertaken at the CCRC.

4.2.16 Monosaccharide linkage analysis reveals a high diversity of glycosidic linkages within REC1

To further explore the biochemistry of the REC1 preparation, the CCRC undertook a monosaccharide linkage analysis to determine which glycosidic linkages were present. From these linkages, signatures of polysaccharides were determined, which could verify the presence of the polysaccharides detected by the MAbs. The sample was suspended in DMSO, and thoroughly agitated for a week. Aliquots (200 µL) of 1 M sodium hydroxide and methyl iodide (100 µL) were used to methylate the free hydroxyls on the polysaccharides within the preparation. The preparation was then hydrolysed using 2 M trifluoroacetic acid, and reduced using sodium borohydride (10 mg/mL) in 1 M ammonium hydroxide. The resulting partially methylated alditol acetates were analysed using gas chromatography combined with mass spectrometry, which was undertaken at the CCRC. The chromatogram revealed a high diversity of glycosidic linkages within the preparation (Figure 4.17). The major linkages were formed of mannose (35.6%), glucose (34.7%) and xylose (9.5%), which verified with the monosaccharide composition analysis (Figure 4.16). There was a particular abundance of 1,4-linked mannose, which forms the backbone of mannan. There was also an abundance of 1,4-linked glucose residues, which forms the backbone of many plant cell wall polysaccharides including cellulose, xyloglucan and mixed linkage glucan, amongst others. The third most abundant linkage was 1,3-linked glucose, which is present within mixed linage glucan and callose (Figure 4.17).

On further examination, many linkages were uncovered that were present within the four major polysaccharides previously determined by the MAbs. Strong signatures of AGP and extensin, terminal, 3-, 4-linked Arap, terminal, 5-linked Araf, terminal Galp, terminal Rhap, were detected. Strong signatures of xylan, 4-linked Xylp, and xyloglucan, terminal Fucp, terminal Galp, 4- and 4,6-linked Glcp, terminal and 2-linked Xylp were detected. Other polysaccharide signatures were also detected including arabinan, cellulose, galactomannan, glucomannan, mannan, mixed linkage glucan, RG-I and RG-II (Table 4.1). Possible signatures of fungal cell wall polysaccharides may have also been detected within the sample of REC1. Many other linkages were present within the preparation of REC1, which could not be assigned to a clear polysaccharide signature (Table 4.1).

Figure 4.17 I Total ion chromatogram of monosaccharide linkage analysis of REC1

The sample (1 mg) was suspended in DMSO and agitated for 1 week. Methylation was undertaken by using 200 µL of 1 M sodium hydroxide, and then 100 µL methyl iodide. The sample was then hydrolysed using 400 µL trifluoroacetic acid at 121ºC. This preparation was then reduced using 1.2 mL sodium borodeuteride (10 mg/mL) prepared in 1 M ammonium hydroxide. The resulting partially methylated alditol acetates were analysed using gas chromatography combined with mass spectrometry. There was a high amount and diversity of linkages within the preparation. A large amount of 1,4-linked mannose, which forms the backbone of mannan was determined. The next most abundant linkage was 1,4-linked glucose that forms the backbone of many plant cell wall polysaccharides including, cellulose, xyloglucan, mixed linkage glucan and others. The third most abundant linkage was 1,3-linked glucose, which is present in mixed linkage glucan. Many other linkages involving arabinose, fucose, galactose, rhamnose and xylose that were detected are found in the four polysaccharides, AGP, extensin, xylan and xyloglucan, determined by the MAbs. This work was undertaken at the CCRC.

Table 4.1 I Summary of monosaccharide linkage composition of REC1

The sample (1 mg) was suspended in DMSO and agitated for 1 week. Methylation was undertaken by using 200 µL of 1 M sodium hydroxide, and then 100 µL methyl iodide. The sample was then hydrolysed using 400 µL 2 M trifluoroacetic acid at 121ºC. The sample was then reduced using 1.2 mL sodium borodeuteride (10 mg/mL) prepared in 1 M ammonium hydroxide. The resulting partially methylated alditol acetates were analysed using gas chromatography combined with mass spectrometry. This work was undertaken at the CCRC. The major linkages uncovered in the preparation were mannose (pyranose) based including 1,4-linked mannose, which forms the backbone of mannan. The other mannose linkages were not determined within plants. Glucose-based linkages were the second most abundant linkages found in REC1. In particular, 1,4-linked glucose was uncovered in the sample that forms the backbone of many polysaccharides including xyloglucan, cellulose, glucomannan and mixed linkage glucan. Another glucose linkage was 1,4,6-glucose which is found in xyloglucan. Other linkages uncovered within the preparation of REC1 were rhamnose-based, which can be found in AGP and rhamnogalacturonan, arabinose-based linkages found in AGP and extensin, and xylose-based linkages which form the backbone of xylan, and the decorations on xyloglucan. Abbreviations stand for: Ara, arabinose; Fuc, fucose, Gal, galactose; Glc, glucose; Man, mannose; Rha, rhamnose; Xyl, xylose; P, pyranose, F, furanose. All values (Mol %) ≥3 have been underlined. Any deduced linkage that cannot be assigned a clear polysaccharide signature has been marked (-). Signatures of fungal cell wall polysaccharides (*) may have been detected within the sample (Pettolino et al. 2009).

4.2.17 There was no mannan detected when screening REC1 with a wide range of mannan-specific probes

To explore further the presence of mannan within REC1, as deduced by the monosaccharide composition and monosaccharide linkage analyses, a range of mannan-specific probes (LM21, CCRCM-170; Pattathil et al. 2010 and CBM27; Boraston et al. 2003) were used to screen a sample of REC1. Other MAbs specific to AGP (LM2), extensin (LM1), xylan (CCRCM-108; Pattathil et al. 2010, LM11, LM27 and LM28) and xyloglucan (LM25) were included as these polysaccharides have been detected before. Additionally, callose (1,3-β-D-glucan; Meikle et al. 1991) and mixed linkage glucan (1,3:1,4-β-D-glucan; Meikle et al. 1994) specific MAbs were included. The screen revealed the presence of AGP, extensin, xylan and xyloglucan (Table 4.2). There was also a slight signal from CCRCM-108 which recognises the epitopes of methylated xylan. Furthermore, there was no detection of callose, mixed linkage glucan or mannan (Table 4.2). The lack of signals from the mannan-specific MAbs appears to contrast the monosaccharide composition and monosaccharide linkage analysis (Table 4.1 and Table 4.2).

Table 4.2 I No mannan was detected when screening REC1 with a range of mannan-specific probes through ELISA

After screening a sample of REC1 (10 µg/mL) through ELISA with a range of molecule probes specific to mannan (CCRCM-170, LM21 and CBM27), there was no mannan detected. This contrasts with the high levels of mannan detected within the monosaccharide composition and monosaccharide linkage analyses. The mannan detected may be novel to the roots of Cadenza or fungi cell walls. Callose (1,3-β-glucan) and mixed linkage glucan (1,3:1,4-β-glucan) were also not detected within REC1. AGP (LM2), extensin (LM1), xylan (CCRCM-108, LM11 and LM27) and xyloglucan (LM25) where detected within REC1. All LM and CCRCM antibodies were used at a 1:10 dilution, CBM was used at a concentration of 10 µg/mL, and callose and mixed linkage glucan antibodies were used at a 1:1,000 dilution. ELISA absorbance values were determined by measuring the absorbance at 450 nm. Data are a mean of two biological replicates. A cut off point of 0.1 au was removed, any values below this level were regarded as having no signal (-).

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4.3 Discussion

4.3.1 Released polysaccharides have a complex biochemistry and potential linkages

There has been a renewed effort to explore the cell wall architecture with a particular focus on how polysaccharides form continues interconnected networks. The current view of the cell wall matrix is that cell wall constituents have a singular function (Keegstra 2010; Cosgrove and Park 2012). However, this concept is increasingly becoming more nuanced as more polysaccharides have been shown to be linked (Renard et al. 1997; Duan et al. 2004; Popper and Fry 2008) some of which have been shown to form a multi-polysaccharide complex (Tan et al. 2013). For the first time, this investigation has used EDC and sandwich-ELISA to explore the biochemical properties of polysaccharides released by roots. Using these tools, AGP, extensin, xylan and xyloglucan, were demonstrated to be linked as a part of a multi-polysaccharide complex released by cereal roots. Uncovering this released complex has expanded our understanding of polysaccharides released by roots. Prior to this investigation released polysaccharides were thought to be released as individual polysaccharides rather than the more interconnected structures extrapolated within this research. As it is believed that polysaccharides released from roots come from the cell walls of the root body (Bacic et al. 1986; Moody et al. 1988), they reflect the widening understanding of the cell wall architecture.

The initial EDC screens revealed the presence of two forms of xyloglucan, neutral and acidic, within the concentrated hydroponate of wheat (Figure 4.2 and Figure 4.3). The majority of literature suggests that xyloglucan is neutral (Somerville et al. 2004; Cosgrove 2005; Knox 2008; Albersheim et al. 2010). Acidic forms of xyloglucan have been determined, which include galacturonic acid residues, within basal land plants including liverworts (Peña et al. 2008), and within the root hair of Arabidopsis (Peña et al. 2012a). Acidic xyloglucan has not been previously determined in or being released by other plants particularly, cereals. The xyloglucan released by wheat as determined by this investigation may be novel or may not be acidic. The hydroponates of cereal indicate that there is a link between AGP, extensin, xylan and xyloglucan (Figure 4.3 and Figure 4.12). A comparison between the standard tamarind seed xyloglucan, which is neutral, was compared to the xyloglucan released by wheat roots (Figure 4.2). This comparison confirmed that the released xyloglucan was acidic. When taking all of this into account, the acidic xyloglucan uncovered in the concentrated hydroponate may be being retained within the anion-exchange column due to linkages with AGP. An AGP macromolecule was detected co-eluting with the acidic xyloglucan. This AGP contained a glucuronic acid residue, inferred by the binding of LM2, which specifically recognises glucuronic acid within AGP. The presence of glucuronic acid makes AGP released by plants highly acidic (Smallwood et al. 1996; Yates et al. 1996). AGP characteristically consists of a small protein core (typically between 5-30 KDa) that has a series of highly heterogeneous arabinogalactans sidechains, which represents 90-95% of the total glycoprotein (Stacey et al. 1990; Showalter 2001; Geshi et al. 2013). This makes AGP a complex molecule, which may have attached the xyloglucan to the matrix within the anion-exchange column until enough salt had eluted the AGP, thus releasing the xyloglucan (Figure 4.2 and Figure 4.4). This indicates that the xyloglucan would remain neutral but is held in the column by the AGP, thus giving the appearance of an acidic xyloglucan. This AGP may act as a crosslinker within REC1, comparable to APAP1 (Tan et al. 2013), which holds the subdomains of extensin, xylan and xyloglucan together. To confirm this hypothesis degrading the AGP with galactanases that hydrolysed the β-1,3- and β-1,6-Galp links that formed the backbone (Ellis et al. 2010), or using β-glucuronidase to target the β-glucuronic acid residue, as detected by LM2 (Smallwood et al. 1996; Yates et al. 1996) would be a key experiment. If degrading the AGP disrupts the co-elution of REC1 during anion-exchange EDC, then this would confirm the hypothesis. As a consequence, AGP could be the key polysaccharide within REC1.

Size-exclusion EDC (Figure 4.5) reveals a highly complex heterogeneous matrix of polysaccharides released from wheat roots. REC1 may not be comparable to APAP1 that has a defined AGP core that linked to the arabinoxylan and pectin subdomains (Tan et al. 2013). REC1 could be formed of different subpopulations of xyloglucan and xylan, which are linked to an AGP protein core with an extensin subdomain. Instead of neatly defined subdomains; REC1 may be formed of an aggregate of differentially sized polysaccharides. The chromatograms of the root body and hydroponate differed (Figure 4.5, A and B), suggesting the polysaccharides may aggregate after they are released by wheat roots forming these elongated traces. The xyloglucan released by wheat roots differed from the commercial standard, tamarind seed xyloglucan. The tamarind seed xyloglucan eluted from the column at the same time as the xyloglucan within REC1 (Figure 4.5, A). However, the tamarind seed xyloglucan was fully eluted at fraction 60, whereas, the xyloglucan from REC1 continued until it eluted at fraction 80. This suggests that the xyloglucan released by wheat roots differs in size compared to the commercial sample.

When comparing polysaccharides that were contained within the root body to the hydroponate, xylan had the strongest signal, revealing two populations (Figure 4.5, B). This has been determined in previous work, which shows that xylan is highly abundant and heterogeneous in both the primary and secondary cell walls of grass species (Ebringerová and Heinze 2000; Vogel 2008; Simmons et al. 2016). The trace of xyloglucan on the anion-exchange chromatogram of the root body had a similar pattern to xylan, which suggests that there could be two populations of xyloglucan within the roots of wheat. Perhaps, containing higher amounts of a particular polysaccharide increases the chances of that polysaccharide becoming more diverse in size and structure. The factor that determines the size of each polysaccharide produced in a plant cell is unknown. There could be a great variation in the size of each polysaccharide, therefore, the differently sized polysaccharides could play a specific function or have a specific binding affinity to other polysaccharides.

The binding of LM11 within the concentrated hydroponate of wheat suggests the presence of heteroxylan (Figure 4.2). Furthermore, there was no signal of LM10 within the hydroponates of the cereal crops (Table 3.1 and Figure 3.3). LM10 is known to bind to the non-reducing terminal of xylan (Ruprecht et al. 2017). There was no binding of LM12, which indicates that the released xylan lacked ferulic acid, therefore, indicating that released xylan was neutral (McCartney et al. 2005). This is further supported by the lack of a signal from LM28 (Table 3.1), which binds to the epitopes of glucuronoxylan, which is an acidic form of xylan (Cornuault et al. 2015). If the released xylan was neutral then the signal of LM11 should have eluted earlier within the anion-exchange chromatogram, similar to that of the acidic xyloglucan. This further supports the hypothesis that xylan is linked to an acidic molecule (AGP). When samples of concentrated hydroponate were treated with xylanase and xyloglucanase there were no alterations within the co-elution (Figure 4.9). This verifies that xylan and xyloglucan do not affect the acidity of REC1.

When the onset of the salt gradient was delayed during anion-exchange EDC, the co-elution of the polysaccharides was retained for longer within the column (Figure 4.3). Additionally, there was no separation of the co-elution or any new peaks described. This demonstrates that the polysaccharides within this co-elution were strongly enough linked to be teased apart from the neutral peak of xyloglucan by delaying the gradient of salt. If there was a weaker link between the polysaccharides the co-elution would have been disrupted as the salt gradient was altered. More detailed research using a shallow gradient, which do not have a sudden increase in salt (similar appearance to an exponential growth curve) is required to determine if the co-elution remains stable.

The total carbohydrate assay uncovered that there was a complex glycan content in the concentrated hydroponate of Cadenza. The neutral (fractions 0-25) and acidic (fractions 39-45) clusters of glycan generally confirm the observations made in anion-exchange EDC (Figure 4.3). However, there was a second mass of neutral glycan, which was unexpected. This second mass of neutral glycan had not been detected by the MAbs. This suggests that there could be another body of glycans being released by the wheat roots that are not detected by MAbs (Figure 4.6). This factor may play an important role in the polysaccharide matrix released by roots. There was a slight increase in MAb signals that were within the co-elution as the gradient of salt was delayed. This increase in signal suggests that this factor could potentially be masking the binding of the polysaccharides within this co-elution. More research exploring the total glycan and protein content is required to identify and characterise this mass of neutral glycan.

4.3.2 Complex heterogeneity occurred within the linkages of REC1

The monosaccharide composition and monosaccharide linkage analyses detected residues and linkages that are found in AGP, extensin, xylan and xyloglucan (Table 4.1) detected by the immunochemical assays. The detection of similar polysaccharide signatures verifies the observations made from the MAbs, which determined the presence of AGP, extensin, xylan and xyloglucan within the concentrated hydroponate of Cadenza. One surprise from these structural tests was the absence of acidic residues, particularly glucuronic acid found in AGP and galacturonic acid found in acidic xyloglucan (Figure 4.16). Previous work within this study determined that REC1 was acidic through the binding of LM2, which specifically recognises the epitopes of β-linked glucuronic acid present in AGP (Table 3.1 and Figure 3.3). Furthermore, the results from the anion-exchange chromatography highlighted the acidic property of REC1 (Figure 4.2). Past investigations have also determined the presence of glucuronic acid within the liquid medium of wheat (Moody et al. 1988). The investigation uncovered that glucuronic acid formed 24% (total Mol) of the wheat exudate (Moody et al. 1986). The medium of maize has also been demonstrated to contain high levels of glucuronic acid (Bacic et al. 1986). Furthermore, galacturonic acid has also been detected within the root mucilage of wheat, representing a minor component (3% of the total monosaccharides detected; Moody et al. 1988). There are only two sources of galacturonic acid within the concentrated hydroponate of Cadenza as determined by MAbs, xyloglucan and xylan. Galacturonic acid found in xyloglucan has only been detected within the root hairs of Arabidopsis (Peña et al. 2012a) and basal plants such as liverworts (Peña et al. 2008). Additionally, galacturonic acid is also present within heteroxylans, glucuronoxylan and GAX, which are only present within cereals. This absence of galacturonic acid within the concentrated hydroponate of Cadenza indicates that the xylan and xyloglucan released by Cadenza are neutral. The discrepancy between this research and past investigations with the monosaccharide composition analysis (Figure 4.16) may be due to issues related to the methods employed by the CCRC or may suggest that REC1 is not acidic but could be highly sticky within the anion-exchange column.

During linkage analysis, the CCRC reported that there were issues with hydrolysing the sample. This may have been due to freeze-drying the sample, which has been previously reported to cause difficulties in rewetting (Pettolino et al. 2012). Perhaps, the glucuronic acid within the sample was insoluble or was somehow degraded during the analysis. Several gradients of salt were used through anion-exchange chromatography to analyse REC1 which all demonstrated that REC1 was eluting within the acidic region (Figure 4.2 and Figure 4.3). REC1 could be sticking to the anion-exchange columns until sufficient salt has dislodged the complex and eluted it. More work using different column matrixes is required to verify if REC1 is being retained in the column due to the complex being acidic or highly sticky. Using a cation-exchange column or different anion-exchange matrixes may help to expose if REC1 is acidic or highly sticky. Moreover, isolating more REC1 is required to undertake more structural tests that may further elucidate the acidity of the complex.

Previous examinations of root mucilage have detected similar monosaccharide residues and linkages, as well as their amounts to that of this investigation (Table 4.1; Bacic et al. 1986; Moody et al. 1988). Signatures of AGP (terminal Araf, 6-linked Galp, 3,6-linked Galp and terminal Rhap), xylan (4-linked Xylp) and xyloglucan (terminal Galp, terminal Glcp, 4-linked Glcp, terminal and 2-linked Xylp) have also been detected within the liquid medium of wheat seedlings (Moody et al. 1988). As well as these signatures, previous research has determined the presence of linkages that cannot be assigned a clear polysaccharide signature, similar to this investigation. Past research examining the sterile liquid medium of wheat determined the presence of 6-linked Glcp and 2,4-linked Glcp (Moody et al. 1988). These unassigned linkages confirm the observations made in this investigation. Research examining the linkages within the media of other cereal crops such as maize and rice have also determine similar signatures of AGP, xylan and xyloglucan (Table 1.1; Bacic et al. 1986; Chaboud and Rougier 1984; Moody et al. 1988; Read and Gregory 1997). Other linkages uncovered within the liquid medium of maize seedlings grown in the dark, have also been detected within this investigation, namely 3-linked Fucp, 2,3-linked Galp, 3,6-linked Galp and 2,3-Manp (Table 1.1; Bacic et al. 1986). Furthermore, 3,4-linked Galp was detected within the liquid medium of cowpea seedlings (Table 1.1; Moody et al. 1988; Read and Gregory 1997). Although these linkages have been uncovered within several studies, how they fit together remains unknown.

This investigation has detected novel signatures of arabinan, cellulose (could also be xyloglucan; Figure 1.3), callose, extensin, galactomannan, mannan, mixed linkage glucan, RG-I and RG-II (Table 4.1) within the hydroponate of wheat, which have not previously been determined (Table 1.1). Much more research is needed to fully understand how these linkages are connected to determine which structures (oligo- or polysaccharide) they form. From the monosaccharide linkage analysis (Table 4.1), the preparation of REC1 is more heterogeneous and complex than initially thought. Considerably more research using NMR-based detection techniques are required to figure out how these linkages fit together. Is REC1 a complex or a highly heterogeneous aggregate or matrix of cell wall polysaccharides released by roots? Why is this complex so heterogeneous? These are amongst the many questions that remain unanswered about REC1.

Another unexpected result of the monosaccharide composition and linkage analyses was the high amount of mannose and 1,4-linked Manp linkages, which form the backbone of mannan (Figure 4.16; Figure 4.17; Table 4.1). From antibody-based approaches (Table 3.1 and Figure 3.3) there was no detection of mannan. Furthermore, there was an additional screen of a sample of REC1 using several mannan-specific molecular probes, which also did not detected any mannan (Table 4.2). This may suggest that there were issues with the methods employed by the CCRC or that the mannose residues and linkages are heavily masked by other molecules within the preparation. Screening a sample of REC1 with a range of mannan-specific MAbs and CBM revealed there was no signal of mannan (Table 4.2). This suggests that the mannan detected within the monosaccharide composition and monosaccharide linkage analyses may be novel to the exudate of Cadenza or fungi cell walls (Pettolino et al. 2009). Another explanation is that there was an issue with the structural analysis. Furthermore, the mannose residues and linkages may be modified in such a way that the mannan-specific MAbs could not recognise or that the mannose in REC1 is novel. There were also no signals from the callose and mixed linkage glucan MAbs when screening REC1 (Table 4.2). This may suggest that the deuced 1,3-linked Glcp (Table 4.1) may not belong to either callose or mixed linkage glucan but may be a linked novel in REC1.

Several mainly glucose and mannose linkages, for instance 3,4-linked Glcp, 2,4,6-linked Glcp, 2-linked Manp, 4-linked Manp, 2,3-linked and Manp, were detected within the preparation of REC1. These signatures may be from fungal cell wall polysaccharides (Pettolino et al. 2009) or from other possible sources of contamination, including from starch or paper (Pettolino et al. 2012). It may also be conceivable that these linkages (except for 4-linked Manp) are novel in plants. Wheat were grown in non-sterile conditions, which could have been contaminated with fungi, bacteria or a combination. Research exploring the major linkages within the cell walls of Rhynchosporium secalis, which causes barley leaf scald, determined that there were high levels of mannose and glucose linkages (Pettolino et al. 2009), which were also uncovered within this investigation (Table 4.1). It is known that barley leaf scald can infect wheat by entering the roots and causing symptoms such as greyish-green spots, dry lesions and chlorosis, primarily on the leaves (Pettolino et al. 2009). However, such symptoms were not apparent on the leaves of the wheat grown for this study. It is conceivable that other fungi or bacteria could have infected the hydroponics system without presenting any obvious indications of infection. This may account for the high amounts of mannose and mannose-based linkages detected. The MAbs may not bind to the epitopes of a mannan-like molecule released by fungi as they have been raised against plant cell wall polysaccharides. Within other research that grew wheat seedlings in sterile conditions, these linkages were not detected (Moody et al. 1988). One exception to the possible fungi cell wall linkages was 2,3-linked Manp, which had been detected within the sterilised liquid medium of maize (Bacic et al. 1986). How these possible fungal cell wall polysaccharides interact with the polysaccharides released from roots remains unclear. Future hydroponic systems would need to be grown in sterile conditions in order to prevent possible contamination issues.

When combining the signatures of polysaccharides detected within this investigation (Table 4.1), linkages described from past research (Bacic et al. 1986; Moody et al. 1988), and linkages present in the cell walls of barley leaf scald fungi (Pettolino et al. 2009) there are three linkages that have yet to be explained, 2,4-Glcp (1.9% of total linkages in REC1), 2,4,6-linked Manp (0.3%) and 2,3-linked Rhap (0.3%). These linkages appear to be specific to the REC1 preparation, and novel to wheat. Perhaps, these linkages play an important role within this putative complex.

4.3.3 Multi-polysaccharide complexes may share similar characteristics

Our understanding of the architecture of cell walls is becoming more complex with more emerging technologies used to explore it. The current paradigm of classifying cell wall polysaccharides into structurally defined groups, each with a single function (Keegstra 2010; Cosgrove and Park 2012) is increasingly becoming redundant. Recently, some cell wall polysaccharides have been found to be linked as a part of a multi-polysaccharide complex, APAP1 (Tan et al. 2013). Covalent linkages of xyloglucan and pectin have also been shown to occur (Duan et al. 2004; Popper and Fry 2008; Cornuault et al. 2015). These investigations have established a new understanding of cell wall architecture. Evidence thus far suggests that these multi-polysaccharide complexes may constitute a minor proportion of the cell wall (Tan et al. 2013). Furthermore, these complexes may be tissue-specific or specific to developmental stages of a plant. These complexes may also have crucial functions within the mechanical properties and dynamics of the cell wall matrix. The current understanding of these multi-polysaccharide complexes remains limited, with little evidence of their role, location and biosynthesis.

REC1 is the first multi-polysaccharide complex released by plant roots to be identified. The first complex uncovered in Arabidopsis callus, APAP1, is formed of an AGP core with covalently attached subdomains of RG-I and arabinoxylan (Tan et al. 2013). Prior to the discovery of these complexes, inter-glycan links between pectic polysaccharides and structural proteins were theoretically possible (Ryden and Selvendran 1990). Only until recently, covalent links between pectic polysaccharides and xyloglucan (Duan et al. 2004; Popper and Fry 2008), and hydrogen links between xylan and cellulose (Simmons et al. 2016) had been observed. Potential linkages of xylan-pectin and xylan-AGP have also been reported using immunochemical techniques (Cornuault et al. 2015). One major similarity between REC1 and APAP1 is the presence of AGP and xylan subdomains. Within APAP1, the AGP acts as a crosslinker, which forms the core of the complex, holding the other polysaccharide subdomains together (Tan et al. 2013). This was confirmed by undertaking a knockdown of the gene that formed the AGP core. Once the knockdown had occurred, APAP1 was not detected (Tan et al. 2013). It is conceivable that the AGP within REC1 is also acting as a crosslinker molecule, holding the other subdomains together. Further evidence is shown by the delay in the elution of xyloglucan within the anion-exchange column when the extended salt gradient used. It was postulated that the AGP was retaining the other polysaccharides within the column (Figure 4.2 and Figure 4.3).

A limitation of APAP1 is that it is lacking in significance as it was only detected within the callus cells of Arabidopsis rather than the whole plant. Exploring the biochemistry of callus cells, which do not solely occur in the wild, may not be representative of whole plants. Whereas, REC1 appears to be a major factor released by roots. Growing cells of these plants may have affected the biosynthesis, mechanisms of release, and location of secretion. It would be interesting to see if the presence of APAP1 is altered within and being released from Arabidopsis plants rather than callus cells. This study detected REC1 by growing crop species in hydroponics, which may have also affected the release of these polysaccharides. Hydroponics remains an unnatural place for these crops to be grown in. It would be interesting to grow wheat in soil, or extract polysaccharides from soils that have had wheat growing in, with strong alkalis to see if REC1 can be detected.

4.3.4 REC1 may be constructed in the cell wall matrix

The biosynthesis of these multi-polysaccharide complexes remains unclear, however, it was speculated that the subdomains of APAP1 were produced in the Golgi apparatus (Tan et al. 2013). The biosynthesis of xylan and pectin occurs within the Golgi apparatus (Gibeaut and Carpita 1994; Reiter 2002). The synthesis of APAP1 would have to occur spatially and temporally in sync, as AGP is synthesised in both the endoplasmic reticulum (protein domain) and Golgi apparatus (glycan domain; Fincher et al. 1983; Gladys 1998). The biosynthesis of APAP1 probably occurs in situ within the cell wall matrix. This would require less energy as the whole structure would not need to be passed through the plasma membrane to get into the cell wall. The pectic and arabinoxylan subdomains may become attached to the AGP core within the cell wall matrix through enzymatic (transglycanase) activities (A. Galloway, 2017. Email to L. Tan, 9 May). A comparable process could also occur for the biosynthesis of REC1 (Figure 4.3 and Figure 4.7) and REC2 (Figure 4.15).

The biosynthesis of plant polysaccharides occurs within the Golgi apparatus, with protein synthesis occurring within the rough endoplasmic reticulum (Albersheim et al. 2010). After production, these glycans and glycoproteins are packaged into vesicles and transported to the plasma membrane mediated through the protein complex exocyst for cell wall integration (Guinel and McCully 1986; Somerville et al. 2004; Synek et al. 2014). It is probable that the subdomains of REC1 are constructed on the plasma membrane, either as a continual conveyer or en bloc, after vesicles have delivered sufficient supplies of glycans. It would be less energetically favourable for these complexes to be passed through the plasma membrane, and then transported and integrated into the cell wall architecture. This would be like transporting a skyscraper from a construction site to a large city, and integrating it. It would be less resource intensive to transport smaller modules of a skyscraper to a large city, and fixing the modules together in situ. It would be interesting to explore the energy demands of REC1 exudation on plants.

As roots penetrate through deeper layers of soil they are subjected to ever increasing friction. This friction causes root cap cells to undergo lysis (Read and Gregory 1997; Iijima et al. 2002). As well as lysed cells, some cells are pre-programmed to detach themselves and actively release polysaccharides (Hawes et al. 1998; Driouich et al. 2013). Detaching border cells results in the loosening of cell walls of neighbouring cells. Cells with loosened cell walls actively produce a continual amount of polysaccharides, to ensure their structural integrity (Hawes et al. 2002; Driouich et al. 2013). As a consequence, it is reasonable to suggest that these means are the ways in which REC1 is released. Although the mechanisms underpinning the release of these multi-polysaccharide complexes remains uncertain. As cell detachment as not been observed from liverworts, it may be that lysed rhizoid cells result in the release of REC2. Nonetheless, rhizoids have not previously been determined to release molecules, and more research is required to further develop our understanding of liverwort exudates.

4.3.5 Pea and rapeseed may release a multi-polysaccharide complex that could be related to REC1

There were striking differences between the chromatograms of the concentrated hydroponates of eudicotyledons and cereals (Figure 4.3, Figure 4.10 and Figure 4.14). These differences suggest that plant roots release a unique mixture of polysaccharides. There was a range of pectic polysaccharides detected within the hydroponates of pea, rapeseed and tomato. This may reflect the high levels of pectin within the cell walls of eudicotyledons compared to cereals of which pectin is a minor component (Table 3.2). Pectin may have contributed to the biochemical diversity of the released polysaccharides from eudicotyledons. Pectic polysaccharides may be influencing the released polysaccharides assayed. Additional anion-exchange EDCs are required to explore the biochemistry of the released pectin, and if they are linked with the other released polysaccharides. Despite the lack of strong evidence of a multi-polysaccharide complex released by eudicotyledons, there were small hints that these polysaccharides could be linked (Figure 4.14). Additionally, there were some similarities between eudicotyledon and cereal released polysaccharides. There were generally two forms of xyloglucan released; neutral and acidic (Figure 4.14). This was particularly noticeable in the concentrated hydroponate of rapeseed, which is related to Arabidopsis that contains an acidic xyloglucan within its root hairs (Peña et al. 2012). Further biochemical exploration will clarify if these polysaccharides are linked as a complex or not.

4.3.6 Releasing multi-polysaccharide complexes may occur across land plants

Early land plants would have been confronted with high water stress, low nutrient ability and serve weather conditions compared to that of living in the primordial oceans (Field et al. 2015; Mitchell et al. 2016). Despite the long evolutionary distance between modern-day plants and the relatives of early land plants (~470 million years; liverworts), much of their cell wall matrix composition remains similar to that of Type I primary cell walls (Popper and Fry 2003; Del Bem and Vincentz 2010; Popper et al. 2011). Despite the similarities in cell wall composition, the function of rhizoids and root hairs greatly differs. Rhizoids were thought to serve only to anchor the plants into the primordial soils, whereas, root hairs are primarily involved in water and nutrient uptake, as well as anchorage (Jones and Dolan 2012; Arteaga-Vazquez 2015). Furthermore, it has been uncovered that the root hairs of Arabidopsis also released an acidic xyloglucan (Peña et al. 2012), comparable to that of xyloglucan in liverwort cell walls (Peña et al. 2008). As of yet it has not been demonstrated that the rhizoids of liverworts are involved in releasing cell wall components. This investigation uncovered that liverwort rhizoids release cell wall-relate polysaccharides, which form an AGP-xyloglucan complex, REC2. This finding may lead to a change in our understanding of the mechanics of rhizoids, as initial work present here suggests that they can release polysaccharide. By screening agar gel extracts of liverworts with 29 MAbs, AGP and xyloglucan, as detected by LM2 and LM25 respectively, ranked the highest (work undertaken by Ms Bev Merry). Furthermore, using nitrocellulose tissue prints the areas releasing these molecules were confirmed to be released by the rhizoids. Higher levels of xyloglucan were observed being released at the tips of the rhizoids (Figure 4.15, D). This observation is comparable to the polysaccharides released from the root caps and tips of higher plants (Bacic et al. 1986; Morel et al. 1988; Read and Gregory 1997; Watanabe et al. 2000). Perhaps, mechanisms which underpin the release of these polysaccharides in liverworts are comparable to higher plants (Figure 4.18). Interestingly, within the initial screen of the liverwort agar extract there were no xylan and pectin detected, which are components of the cell walls in liverworts (Konno et al. 1987; Popper and Fry 2003). The reason for the lack of pectin and xylan within the agar extract remains undetermined.

Liverwort xyloglucan is acidic due to the presence of galacturonic acid (Peña et al. 2008), which accounts for the second acidic peak of xyloglucan within anion-exchange EDC (Figure 4.15, B). However, there was a neutral form of xyloglucan detected. This neutral form of xyloglucan is also a new finding as the xyloglucan within liverwort cell walls was thought to be only acidic (Peña et al. 2008). It is interesting to observe that in cereal and liverwort exudates that there are consistently two forms of xyloglucan, neutral and acidic (Figure 4.3 and Figure 4.15, B). There could be a larger polysaccharide undetectable by MAbs screened, which is highly neutral, eluting a part of the xyloglucan. More work is required to understand liverwort exuded polysaccharides, and to characterise the structure of REC2 through monosaccharide composition and monosaccharide linkage analyses. This may be problematic as liverworts are slow growing, and it was found that they do not grow well within a hydroponic system. As AGP and xyloglucan may be linked within the hydroponates of cereal and the exudates of liverwort, it may be that releasing these polysaccharide complexes is a trait of land plants. The last common ancestor of both cereal and liverworts were the first land plants to have emerged from Charophyte (green algae), over 470 million years ago (Kenrick and Crane 1997; Sorensen et al. 2010; Fangel et al. 2012; Figure 4.15, A). Perhaps, the first land plants that released their cell wall components were more able to colonise the land by developing a primitive root-soil interface. This interface would make it more effective to extract water and nutrients from the primordial soil. It would be interesting to trace the evolutionary development of REC1 by exploring the exudates of other basal plants such as hornworts, and other types of plants such as horsetails, ferns, cycads and gymnosperm such as conifers (Figure 4.18). Furthermore, cross-referencing the evolutionary development of AGP, extensin, xylan and xyloglucan with their release would also be of interest.

Figure 4.18 I Schematic of REC1 detected across land plants

REC1 was uncovered within the concentrated hydroponate of Cadenza. Further screening revealed the presence of a REC1-like molecule within the concentrated hydroponate of other wheat cultivars, Avalon and Skyfall. Expansion of the screening also revealed the presence of a REC1-like molecule released by the roots of barley and maize. Some evidence exists on a REC1-like molecule which may be released by the roots of pea, tomato and rapeseed. Liverworts were determined to release an AGP-xyloglucan putative complex, REC2, which may relate to REC1. Could other species across the plant kingdom from mosses to cycads release a complex similar to that of REC1?

The last common ancestor of liverworts and Arabidopsis was the first land plants, which emerged from fresh water green algae (Charophyte) over 470 million years ago (Kenrick and Crane 1997; Sorensen et al. 2010). Prior to the colonisation of land, green algae were the dominant photosynthesising lifeforms. These algae had similar cell wall polysaccharides to that of early land plants, and modern-day plants. However, one major difference is that these green algae lacked xyloglucan (Fry 1989; Popper and Fry 2003). Taxonomic studies have determined that xyloglucan first appeared within the earliest land plants, particularly, liverworts (Popper and Fry 2003). This suggests that xyloglucan may have given early land plants an advantage that permitted the colonalisation of the land (Fry 1989; Popper and Fry 2008; Del Bem and Vincentz 2010). The acidic form of xyloglucan may be a relic of this pre-adaptive advantage or it may still play an important role for the establishment of roots in soils.

4.4 Conclusion

The current understanding of cell wall architecture is changing with the discovery of polysaccharide linkages, and multi-polysaccharide complexes. These changes reflect the growing complexity of released polysaccharides. Root Exudate Complex 1 was detected within the hydroponate of cereal. This complex is possibly formed of four subdomains, an AGP core with extensin, xylan and xyloglucan attachments. REC1 was initially uncovered when two forms of xyloglucan, neutral and acidic, were detected within the hydroponate of wheat. The charge of the AGP macromolecule may have been retaining the other subdomains within the anion-exchange column until enough salt eluted the AGP. This indicates that the xyloglucan was in fact neutral, and not acidic. Digesting the xylan and xyloglucan subdomains had no effects on the REC1 co-elution. This further supports the hypothesis that AGP is the major component of REC1. By extending the salt gradient, the REC1 fractions could be isolated for structural characterisation. The monosaccharide composition and linkage analyses verified the presence of the four major polysaccharides determined by the MAb. Furthermore, the linkage analysis demonstrated that REC1 contained a highly diverse mixture of linkages, some of which were novel to wheat. REC1 may be constructed within the cell wall matrix, and released through lysed root cap cells, actively released by border cells or lost from loosened cell walls of weakened root cells. The rhizoids of liverworts released an AGP-xyloglucan complex, REC2. By uncovering two multi-polysaccharide complexes in plants placed at opposite ends of the plant kingdom, these complexes may be wide spread in plants playing a vital function for plants.

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