Introduction

Plant roots are dynamic organs that interface with the soil. They provide plants with water and nutrients from the soil. This root-soil interface also involves roots secreting a varity of high and low molecular weight compounds, known as root exudate. This exudate includes organic acids, phytohoromones, sterols, proteins, and polysaccharides and has been well documented (Oades 1978; McNear 2013). Secreting this exudate uses a substantial amount of energy for example, Zea mays uses between 20% and 25% of its photosynthetic product (McNear 2013). The polysaccharides of this exudate, known as root mucilage, are a neglected area of research. Mucilage is the gelatinous substance which surrounds the root caps and epidermal layers behind the caps, and consists of cell wall-related polysaccharides and glycoproteins (Bacic et al. 1986; Moody et al. 1988; Read and Gregory 1997; Figure 1). Cell wall-related polysaccharides such as pectin and arabinogalactan proteins (AGPs) present in mucilage may be responsible for the gelatinous property of mucilage (McCulley 1999). The mechanisms underlining how these polysaccharides are secreted remains unclear; however, two possible avenues of research are under investigation. Root caps are subject to mechanical forces as they penetrate through the soil, and thereby, cells are regularly lysed, releasing their components (Figure 1; Read and Gregory 1997). However, other research indicates that mucilage is actively secreted through specialised root cap cell walls (Guinel and McCully 1986). Regardless of the method of secretion, some root cap cells or border cells are programmed to detach and continue to proliferate in the region surrounding the cap. These border cells rapidly increase mucilage production and form a defence network trapping and deterring infectious bacteria (Driouich et al. 2013; Iijima et al. 2004).

Figure 1. The root-soil interface is a highly dynamic environment with numerous fungi, bacteria, and fauna interactions (Adapted from: Dennis et al. 2010). Mucilage is secreted from the root caps and epidermal layers behind the caps. Where mucilage is located, mycorrhizal fungi are located, and uptake the secreted carbon from plants. Sloughed off root cap cells or border cells are released into the soil, secreting additional mucilage. Mucilage is also released from lysed cells. Lysed cells also attract soil dwelling insects, which feed on the released cellular components.

Monosaccharide linkage analysis of secreted root polysaccharides

To determine the composition of secreted polysaccharides prior to this investigation, monosaccharide linkage analysis was undertaken. For monosaccharide linkage analysis to occur, the free hydroxyl groups on the polysaccharides are methylated. After methylation, the glyosidic linkages holding the constituent monosaccharides together within the polysaccharides are hydrolysed using acid, producing individual monosaccharides (Lindberg 1972). After hydrolysis, the hydroxyls groups that are at the positon involved in the linkage are not methylated, thus the position of the linkage can be identified. The monosaccharides must then be reduced to open their rings. These open rings are then acetylated with the unmethylated hydroxyl groups forming a stable acetyl group. The partially methylated hydroxyls that were originally free, and the partially acetylated hydroxyls that were originally involved in the linkage are identified by gas chromatography (GC) combined with mass spectrometry (MS) (Pettolino et al. 2012). The GC retention times and the MS spectra are used to identify how the monosaccharides are linked to form polysaccharides. One of the major limitations of this technique is that experimenters must infer the presence of polysaccharides (Table 2; Pettolino et al. 2012). Many investigations have used Zea mays as standard to study mucilage, as Z. mays forms large droplets of mucilage form its root caps. Early investigations which analysed the monosaccharides secreted by using Z. mays roots revealed that galactose, xylose, arabinose, rhamnose and glucose were present (Table 1; Chabound 1983; Moody et al. 1988). Other species have been investigated which have shown that mucilage composition differs; forming unique monosaccharide profiles (Table 1).

Abbreviations for Table 1 (below) are as follows: arabinose – Ara, fucose – Fuc, galactose – Gal, galacturonic acid – GalA, glucose – Glu, glucuronic acid – GlcA, mannose – Man, rhamnose – Rha, uronic acid – UroA, and xylose – Xyl.

Monosaccharide analyses have provided a detailed understanding of the monosaccharides that form the secreted polysaccharides of studied plants. These analyses demonstrate that plants have a unique combination of these monosaccharides, which form profiles (Table 1). From these monosaccharide profiles, galactose, arabinose, and uronic acid rank high in dicotyledons, whereas in Poaceae glucose, galactose, and xylose rank high.

Cell wall-related polysaccharides in mucilage

Pectin purified from fruit can create gels with the addition of calcium; this also occurs within cell walls and the middle lamella of plant cells, which adheres plant cells together (Jarvis 1982). The gelatinous nature of mucilage also serves to act as an anti-desiccant, which protects Melastoma malabathricum root caps from drying out (Watanabe et al. 2008). AGPs are glycoproteins which have a high water binding capacity, forming highly viscous gels, which some research indicates causes the gelling of mucilage; AGPs have been detected in several species, including Z. mays (Table 2; Knee et al. 2001). Other studies indicate that pectin is responsible for the gelatinous properties of mucilage due to the role of pectin within the cell wall matrix (Read and Gregory 1997). It is conceivable that both AGPs and pectin form the gel within mucilage and are present in differing amounts, forming distinct gelatinous configurations in different species.

Comparison of secreted root polysaccharides from dicotyledons and Poaceae

The primary cell walls of flowering plants can be placed into two basic categories in regards to their biochemistry, Type I and Type II. Type I cell walls, found in dicotyledons and non-commelinoid monocotyledons contain equal levels of cellulose microfibrils and xyloglucan (Albersheim et al. 2010). This cellulose-xyloglucan network is embedded within a pectin matrix, which is rich in homogalacturonans, rhamnogalacturonans I and II. Heteromannan is also present within the matrix of Type I cell walls (Ishii 1997; Fry 2004). Type II cell walls found in commelinoid monocotyledons, which includes the order of Poales and within that order the family of Poaceae or true grasses, contain less xyloglucan than cellulose. Glucuronoarabinoxylan and mixed-linkage glucans are the major glycans that cross-link the cellulose microfibrils instead of xyloglucan (Nishitani and Nevins 1991). Type II cell walls also contain less pectin and heteromannan, and higher levels of phenylopropanoids, and arabinoxylan networks (Kato et al. 1982).

There have been many investigations which have analysed the monosaccharide profiles of several dicotyledon and Poaceae species. These investigations have suggested that the cell wall polysaccharides of dicotyledons and Poaceae reflect their secreted polysaccharides (Bacic et al. 1986; Moody et al. 1988; Table 2). This seems obvious, yet no direct evidence is available to suggest that mucilage originates from cell walls. Some research provides differing views of the monosaccharide composition of the secreted polysaccharides. After hydrolysing the secreted polysaccharides of Z. mays and L. angustifolius, one study determined that there were high levels of glucose within both plants (Read an Greggory 1997), although this contrasted other research (Chabound 1983; Bacic et al. 1986; Table 1), where low quantities of glucose were identified.

Investigations have yielded differing views on the polysaccharides secreted by roots, due to the limitations of monosaccharide linkage analysis. Research using P. sativum root mucilage suggests that xyloglucan was secreted in low amounts (Knee et al 2001). Whereas, Poaceae mucilage contain higher levels of AGPs compared to dicotyledons (Read and Gregory 1997). Other research suggests that AGPs are high within both dicotyledon and Poaceae mucilages (Rasse et al. 2005). Furthermore, when seed coats are exposed to water they can secrete large quantities of mucilage (Arsouski et al. 2010), which may interfere with the monosaccharide linkage analysis. This seed coat mucilage is rich in rhamnogalacturonan, which has not been detected in some research where direct pipetting of mucilage droplets has occurred, or the use of mature specimens (Read and Gregory 1997; Chabound 1983; Bacic et al. 1986).

Mechanisms and origin of the secretion of root mucilage

It remains to be determined whether mucilage polysaccharides are actually derived from the cell walls or are derived directly from the Golgi apparatus. The Golgi apparatus produces the hemicelluloses, pectin and glycoproteins that form cell walls (Cosgrove 2005). The origin of these secreted polysaccharides remains a mystery, with two theories proposed. The first theory suggests that mucilage polysaccharides are secreted through lysed cell wall components (Bacic et al. 1986). As roots penetrate the soil, cells are continually replaced as continual friction causes root cap cells to frequently lyse (McNear 2013). The second theory suggests that cells apoplastically or passively secretes mucilage into the soil through their cell walls (Foster 1982). If mucilage is apoplastically secreted, the mechanisms which help to guide the polysaccharides through the complex cell wall structure are unknown. Fluorescence microscopy investigations have attempted to provide an answer to the secretion of mucilage, and indicate an active means of secretion through cell walls (Figure 2; McCully and Sealy 1996). This is regardless of the possibility of sloughed off root cap cells releasing their cell wall polysaccharides as they lyse. One investigation suspended cut Z. mays roots for two hours and observed droplets of mucilage forming at the root caps (Read and Gregory 1997). Interestingly, Z. mays mucilage was treated with α-amylase, in a monosaccharide linkage analysis, which hydrolysed the bonds holding long-chains of glucose together. This revealed the presence of starch in Z. mays (Bacic et al. 1986). Starch is only located within the plastids of cells, and particularly in the amyloplasts (Bechtel and Wilson 2003), indicating that mucilage was derived from lysed cell walls. One study detected small amounts of lignin in Z. mays mucilage (Mary et al. 1992), further supporting that mucilage originated from lysed cells.

Research suggests that mucilage is solely secreted from the root caps and epidermal cells behind the caps (Bacic et al. 1986; Read and Gregory 1997). Other research indicates that mucilage secretion also occurs at the root hairs (Rasse et al. 2005). The use of the AGP specific binding reagent, β-glycosyl Yariv reagent which binds to the links between the D-galactose backbone of AGP, was bound to the root caps, root hairs and epidermal layers along the root caps of cryostat secretions of Z. mays (Bacic et al. 1986). Under fluorescence microscopy, α-L-arabinofuranosyl monoclonal antibody (MAb) bound to the same location as the reagent (Bacic et al. 1986), thus confirming that AGPs were actively secreted not just at the root caps but at the root hairs and epidermal layers behind the caps. Conversely, other studies suggest mucilage secretions only occur at the root caps (McNear 2013; Walker et al. 2003). Interestingly, one investigation suggested that L. sativum and P. sativum do not secrete gelatinous droplets on their root caps unlike their Poaceae counterparts, Z. mays and T. aestivum, which secrete high levels of AGPs, and form large gelatinous droplets on their root caps (Narasimhan et al. 2003).

Monoclonal antibodies are highly specific molecular probes that can identify and quantify polysaccharides from plant cell walls. Secreted polysaccharides are cell wall-related, and since MAbs are already available to probe a diversity of cell wall polysaccharides, MAbs would be able to directly quantify these secreted polysaccharides. These MAbs would also remove the need to infer the presence of polysaccharides, which was previously undertaken by hydrolysing the monosaccharides necessary for linkage analysis.

Objectives

The aim of this investigation is to study the polysaccharides secreted by plant roots using MAbs, and to determine their function.

• Determine the polysaccharides secreted by Arabidopsis thaliana, Lupinus angustifolius, and Pisum sativum to represent dicotyledons and Triticum aestivum, Zea mays, and Oryza sativa to represent Poaceae. Three different sampling systems will be used: root washing, liquid media, and agar to determine their major polysaccharides secreted.

• Determine if the Secreted Polysaccharide Profiles (SPP) of Arabidopsis thaliana and Triticum aestivum roots are altered when plants are grown in different light, dark, and nutrient regimes, and in the presence of abiotic factors including salt, aluminium, and cadmium ions.

• Determine the locations of polysaccharide secretion along the roots of A. thaliana.

Materials and Methods

Plant preparation and media

Arabidopsis thaliana (L.) Columbia wild type, Lupinus angustifolius (L.), Pisum sativum (L.), Triticum aestivum Var. ‘Cadenza’ (L.), Zea mays (L.) ‘Earlibird’ F1 hybrid and Oryza sativa (L.) seeds (University of Leeds, UK) were sterilised using 10% household bleach (Unilever Domestos Original Bleach, Dublin, Ireland) mixed with distilled water in Falcon tubes. During this process, seeds were left on a Platform 8 rocker (Stuart Scientific, London, UK) at 30 Rev/min for 10 mins. Subsequently, seeds were rinsed with sterilised distilled water 6 times within an Airstream laminar air flow cabinet.

Three sampling systems were used to screen the secreted polysaccharides of plants: root washing, agar, and liquid culture. For root washing, an array of 26 MAbs which screened the liquid after washing were used (Table 3); all seeds were grown on BG11 (Blue Green; Rippka et al. 1979) liquid medium with no sucrose, on soaked filter paper, for 168 h. Moist filter papers were placed into 10 cm x 10 cm square Petri dishes with microporous tape around the lid. All plants were grown in a growth room with a photoperiod of 16 h and a constant temperature of 20ᵒC. All plants were 8 cm from the light source with a light intensity of 110 µmol m^-2 s^-1.

For agar grown plants, 40 A. thaliana seeds were grown on BG11 without sucrose and 0.9% plant agar, for 96 h. Autoclaved BG11 medium (50 mL) was poured into square Petri dishes and left for 20 mins prior to seed sowing. A. thaliana seeds were placed in two lines near the top and middle of the medium. All plants were subjected to 12 h photoperiod, a constant temperature of 20◦C, with a light intensity of 110 µmol m-2 s-1. After growth, seedlings underwent root washing, where their roots were dipped into 1.8 mL of distilled water for 1 hr. Roots were then removed, and 200 µL of x10 Phosphate Buffered Saline (PBS) was added to the solution in preparation for Enzyme-linked Immunosorbent Assay (ELISA).

For liquid culture, A. thaliana and T. aestivum were grown in liquid autoclaved BG11, with no sucrose. Forty T. aestivum seeds were placed into each 250 mL conical flasks with of 50 mL of BG11. For A. thaliana, 50 mg seeds were added to each flask. For plants grown without nutrient, 50 mL of autoclaved distilled water was added to each flask. A. thaliana was incubated for 96 h and T. aestivum was incubated for 120 h in an Orbi-Safe environmental shaker (Thermo Scientific, London, UK). Twelve hour photoperiod with a constant temperature of 22ᵒC, and a rotation of 135 g was used as standard, with 40 µmol m^-2 s^-1 recorded. For plants grown in the dark, flasks were incubated within another orbital shaker without light, but retaining the same temperature and rotation. For the time series growth analysis, daily sampling (1.8 mL) of A. thaliana and T. aestivum media were undertaken within a laminar air cabinet using sterilised tips. After samples were taken, 1.8 mL of sterilised distilled water or BG11 was added to flasks to prevent loss of media. Both A. thaliana and T. aestivum samples were centrifuged at 3,893 g for 10 mins prior to adding 200 µL of x10 PBS, for the ELISA. For abiotic factors analysis, 1 M solutions of aluminium chloride (AlCI₃), cadmium chloride (CdCI₂), and sodium chloride (NaCI) were made, and aliquots were added to liquid BG11 making 50 µM to 800 µM of AlCI₃ and CdCI₂, and between 50 mM to 800 mM of NaCI concentrations prior to adding seeds. The abiotic factors analysis used the same environmental parameters as outlined for liquid culture. Controls were also used which were absent of salt concentrations. ELISAs were used to detect the amounts of xyloglucan secreted.

Analysis of growth

A. thaliana and T. aestivum root lengths were determined along with A. thaliana plant fresh weight, and T. aestivum rot fresh weights. Root lengths were measured using a digital calliper (Tresena, Essen, Germany); by each end of the calliper at the top of the root near the seed coat and at the bottom near the root cap. An Explorer Pro balance (Ohaus, Nanikon, Switzerland) was utilised to weigh individual T. aestivum roots, and A. thaliana total plant fresh weights. Prior to weighing, seedlings were patted dried with paper towels. The balance was reset to zero, for every measurement.

ELISA of secreted root polysaccharides

To determine the identity of the secreted polysaccharides, and their quantities MAbs were used within a series of ELISAs. Aliquots (100 µL per well) of PBS were added to 96 well microtitre plates (Thermo Scientific, Roskilde, Denmark), excluding the top row. Into the top row, a 1.8 mL of sample with 200 µL of x10 PBS was added at 125 µL per well. From the top row, 25 µL was taken up and titrated (1 in 5 dilution) down the rows, excluding the last row. The plates were left over-night at 4ᵒC to incubate. Plates were washed in tap water and shaken dry. The plates were blocked using 5% Marvel Original milk powder (w/v; Premier Foods, London, UK) in x1 PBS, 300 µL per well. The plates were then wrapped in tin foil and incubated at room temperature (RT) for 2 h. Washing was repeated after incubation and a 1:10 dilution of MAb (University of Leeds, UK) was used at 100 µL per well (Table 3). MAbs were added to block; plates were covered and then incubated at RT for 1 h. Plates washed and shaken dry. A 1:1,000 dilution of anti-rat horseradish peroxidase (HRP) secondary antibody (Sigma-Aldrich, Dorset, UK), was mixed in block; 100 µL per well was added and incubated for 1 h at RT. Plates washed and shaken dry. The following substrate was followed: 18 mL distilled water, 2 mL of 1M sodium acetate pH 6.0, 200 µL of tetramethyl benzidine (10 mg/mL in Dimethyl sulfoxide) and 20 µL of 6% (v/v) hydrogen peroxide which was added just before adding substrate to wells. Substrate (100 µL per well) was added and left for 8 mins before adding 50 µL of 2.5 M sulphuric acid per well. Absorbances were read at 450 nm on a MultiSkan FC spectrometer (Thermo Scientific, London, UK).

Epitope detection chromatography

Prior to chromatography, A. thaliana and T. aestivum liquid media samples were dialysed for 72 h and freeze dried. A. thaliana (250 µg) and T. aestivum (45 µg, and 12.5 µg for xyloglucan analysis) mucilage extracts were diluted in 2.5 mL of dH₂O (distilled water). After diluting, 2.5 mL of the samples were injected into a weak anion-exchange chromatography column (1 mL Hi-trap ANX FF; GE Healthcare, London, UK) using a BioLogic LP system (Biorad, Hempstead, UK). Samples were eluted at 1 mL per minute with sodium acetate buffer (20 mM) at pH 4.5 from 0 to 17 mins. An increase in salt concentration to 50 mM of sodium acetate (pH 4.5) for a further 17 mins with a linear gradient from 0% to 100% with NaCI (0.6 M) to 48 mins was used. This was followed by 8 mins of 50 mM sodium acetate with 600 mM NaCI (elution sample 1) (Cornuault et al. 2014). Forty eight 1 mL fractions were retained. Injection loop (5 mL) and columns were washed with 10 mL of NaOH (0.1 M) between sample injections. Columns were then equilibrated and injection loops washed with 10 mL of acetate buffer (20 mM) before the next injection. The fractions were adjusted to pH 7.0 by supplementing 40 µL of Na₂CO₃ (1 M), and 100 µL aliquots were incubated in microtitre plate wells overnight at 4ºC for subsequent ELISA protocol (Cornuault et al. 2014).

Analysis of secreted polysaccharides using nitrocellulose sheets

To determine the location of secreted polysaccharides along the roots, A. thaliana was grown on BG11 with 0.9% agar for 2 weeks. Nitrocellulose membrane (8 cm x 4 cm; Whatmann GMbH, Dassel, Germany) was laid on top agar, with the plants removed, and left overnight at 4ºC. The nitrocellulose was blocked with 5% milk powder, x1 PBS, and sodium azide (0.0025% w/v) and incubated while shaking (30 Rev/min) at RT for 2 h. Nitrocellulose was then washed with tap water 3 times and left in x1 PBS for 5 mins for 3 times. The following MAbs were used: LM2, LM8, LM19, LM21 and LM25, and were added to the nitrocellulose at 1:5 dilution and incubated for 1 h on the shaker. A control without MAbs was also used. Washing the nitrocellulose was repeated. Anti-rat HRP was added to nitrocellulose at 1 in 1,000 dilution to 5% of milk power and x1 PBS for 1 h at RT while shaking. Washing was then repeated. The colour reagent was followed: 25 mL of dH₂O, 5 mL of chloronaphthol in 100% ethanol (5 mg/mL) and 30 µL of 6% (v/v) hydrogen peroxide. The nitrocellulose was then placed in-between blotting paper for 1 hr, and visualised using an Epson SX400 flatbed scanner (Epson, Hampstead, UK) with a contrast ratio of 168:71.

Statistical analysis

Root length, fresh weight, and dark, light, nutrient regimes, and abiotic factor data were analysed to determine any significant differences. Data were analysed using Minitab 17 statistical software (Minitab, Coventry. UK). Descriptive statistics were analysed, followed by the normal distribution using the Anderson-Darling test and equal variance using the Levene’s test. This led to the One-way Independent ANOVA or its non-parametric equivalent, the Krusal-wallis test. In order to determine where significant differences lay post-hoc, Tukey or Mann-Whitney U tests were used. GraphPad Prism 6 software (GraphPad, California, US) was used to generate all figures.

Results

MAbs can be used to detect secreted polysaccharides

Cell wall specific MAbs were used to screen the secreted polysaccharides from plant roots, from three different sampling systems root washing, liquid media, and agar media. These MAbs detected AGP, pectin, heteromannan, and xyloglucan as the major polysaccharides secreted by roots (Table 4).

An abundance of xyloglucan was secreted by T. aestivum and Z. mays roots

Plants were grown on moist filter paper for 168 h, and had their roots dipped into distilled water prior to ELISA screening. A. thaliana was grown on agar for root washing. The levels of pectin were high within the dicotyledons screened (Table 4). LM25 that binds to xyloglucan was also detected in high amounts, ranking first for A. thaliana, second for P. sativum, and seventh for L. angustifolius. Xyloglucan was ranked between first and second within the Poaceae (Table 4). LM8 that binds to xylogalacturonan was detected within L. angustifolius and P. sativum, ranking fourth and fifth (Table 4). LM5 and LM6 that both bind to rhamnogalacturonan I were ranked higher in dicotyledons; however they were harder to detect within the Poaceae. LM16 epitope of arabinan, a component of rhamnogalacturonan I was higher in dicotyledons. Within Poaceae, epitopes of pectin were generally harder to detect. LM11 and LM12 epitopes of xylan were ranked high within the secreted polysaccharides of Poaceae. The Dicotyledons contain high amounts of pectin, heteromannan and xyloglucan within their cell walls, which was reflected by the secreted polysaccharides detected. Furthermore, Poaceae cell walls contain high levels of xylan, which also reflects their secreted polysaccharides detected. However, there was an abundance of xyloglucan within T. aestivum and Z. mays, which does not reflect the low levels of xyloglucan in their cell walls.

Detection of epitopes from root washing and liquid culture were similar

The SPP of A. thaliana and T. aestivum were similar when probing the liquid media, and liquid from root washing. This verifies that the sampling methods used can detect similar polysaccharides secreted. Liquid cultured A. thaliana contained more MAb signals compared to root washing, with LM25 that binds to xyloglucan doubling in absorbance. High levels of LM1 that bind to extensin were detected at 1.05 au. More MAbs that bind to the epitopes of homogalacturonan were present within the liquid cultured A. thaliana. MAbs from liquid cultured T. aestivum had similar absorbance values to root washing (Table 5).

Growing T. aestivum in dark without nutrients, increased root lengths and root fresh weights while the dark decreased A. thaliana root lengths

A. thaliana was grown in liquid growth medium for 96 h and T. aestivum for 120 h. After these times, the root lengths of both plants were measured as well as the fresh weights and root fresh weights for T. aestivum. The root lengths of A. thaliana significantly decreased when grown in the dark, and in distilled water compared to 12 h of light (Mann-Whitney U, W=1365, P=<0.05). Within the dark treatments there were no significant difference in root growth (Mann-Whitney U, W=918.5, P=0.946; Figure 2, A). The fresh weights of A. thaliana were unaffected when grown under the different treatments (Kruskal-Wallis, H=5.5, P=0.139; Figure 2, B).

The root growth of T. aestivum significantly increased when grown in the dark with distilled water compared to the other treatments (Mann-Whitney U, W=133, P=<0.05). When T. aestivum was grown within the other treatments there were no significant differences (Kruskal-Wallis, H=15.93, =P<0.05). Growing T. aestivum in the dark with distilled water significantly increased the root fresh weights (Mann-Whitney U, W=771.5, P=<0.05). However, the other treatments did not significantly affect the root fresh weights of T. aestivum (Kruskal-Wallis, H=74.01, P<0.05). Prior to the growth analysis, 24 h sampling of the liquid media was undertaken to determine the rate of secretion of the major polysaccharides AGPs, pectin, heteromannan and xyloglucan.

Xyloglucan secretion in A. thaliana was light-dependent

To determine the rate of polysaccharide secretion from A. thaliana and T. aestivum roots, both plants were grown for 192 h in liquid growth medium within an orbital shaker. Every 24 h, 1.8 mL of medium was removed and analysed using an ELISA.

When A. thaliana was grown in 12 h light with BG11, LM2 that binds to AGPs gradually increased until it peaked at 96 h with an absorbance of 1.3 au. The detection of AGPs then gradually declined until 192 h. When grown in the dark with distilled water, the detection of AGP significantly decreased between 24 h and 168 h (Figure 5, A; Kruskal-Wallis H=18.44, P=<0.05). At 192 h, there was a significant reduction between 12 h light with BG11, and 12 h light with distilled water (Figure 3, A-C; Man-Whitney U, W=103, P=0.133). LM19 that binds to homogalacturonan gradually increased until 96 h, when it plateaued until 192 h in 12 h light with BG11. In 12 light with distilled water homogalacturonan formed a bell-shaped curve. In the dark with distilled water, this bell shape became left-skewed (Figure 5, B). From 24 h to 48 h all the treatments remained unchanged. At 72 h, homogalacturonan was significantly reduced in the dark, and 12 h light in distilled water (Mann-Whitney U, W=99, P=0.002). Homogalacturonan was significantly higher in the 12 h light and BG11 compared with the other treatments, between 96 h and 192 h (Mann-Whitney U, W=117, P=0.006). In the dark and distilled water, homogalacturonan was significantly lower compared with BG11 treatments, this continued until 192 h. At 192 h, BG11 treatments, and 12 h light with distilled water were not significant from each other (Figure 3, D-F; Mann-Whitney U, W=54, P=0.006). LM21 that binds to heteromannan was at its highest within 12 h light and BG11 compared with the other treatments. There were no significant differences between all the treatments, until 72 h (Mann-Whitney U, W=103, P=0.133). At 72 h, heteromannan in 12 h light with BG11 was significantly higher compared with the other treatments, continuing until 192 h (Mann-Whitney U, W=45, P=<0.05). After 192 h, dark with BG11, and 12 h light with distilled water were not significantly different. At 96 h, dark in distilled water became significantly lower compared with light with BG11 (Figure 5, C; Mann-Whitney U, W=45, P=<0.05).

LM25 that binds to xyloglucan was reduced within the dark treatments, yet was unaffected by the lack of nutrients. Twenty-four hours after planting, the detection of xyloglucan was not significant. After 48 h, xyloglucan in the 12 h light treatments were significantly higher compared with the dark treatments (Mann-Whitney U, W=45, P=<0.05). Seventy-two hours after planting, xyloglucan was not significantly different within the light treatments. The dark treatments remained significantly lower until 192 h (Mann-Whitney U, W=45, P=<0.05). After 192 h, 12 h light with BG11 significantly increased compared with 12 h light with distilled water, continuing until 168 h were 12 h light with and without BG11 was not significant (Figure 3, J-L).

To summarise, the exclusion of light significantly lowered the detection of LM2 that binds to AGPs (Figure 3, A and B), and LM25 that binds to xyloglucan (Figure 3, J and K) within the liquid growth media of A. thaliana. The detection of LM19 that binds to homogalacturonan, and LM21 that binds to heteromannan were also reduced when A. thaliana was grown in the dark. This was not as significant as the other polysaccharides. The exclusion of nutrients had less impact on the secretion of AGPs, pectin, heteromannan and xyloglucan.

Xyloglucan secretion remained high when T. aestivum was grown without light and nutrient

Twenty-four hours after planting, LM2 that binds to AGPs remained unchanged in all the treatments. After 48 h, 12 h light with distilled water had the highest level of AGPs. There was a significant decrease in AGPs in the dark and distilled water (Figure 5, D; Mann-Whitney U, W=126, P=<0.05). At 72 h, 12 h light with BG11 was significantly higher in LM2 compared with the other treatments (Mann-Whitney U, W=45, P=<0.05). Twelve hours light with distilled water was higher than the dark treatments. At 96 h, the light treatments had the highest level of LM2 that binds to AGPs, which continued until 192 h (One-way ANOVA, F=40.3, P=<0.05). Additionally, 12 h light with distilled water decreased emerging with the dark treatments. At 24 h, LM19 that binds to homogalacturonan was significantly higher in the dark treatments, relative to the other treatments, continuing until 96 h (Figure 5, E; Mann-Whitney U, W=126, P=<0.05). Twelve hours light with BG11 had the highest levels of homogalacturonan compared with 12 h light and distilled water, and dark with BG11, continuing until 96 h. At 96 h, dark with BG11 had the lowest level of homogalacturonan (Figure 4, D-F; One-way ANOVA, F=82.13, P=<0.05). At 120 h, both 12 h light with BG11 and dark with distilled were significantly higher compared with the other treatments, continuing until 192 h (Mann-Whitney U, W=116, P=<0.05). At 192 h, dark with BG11 was not significant compared with 12 h light with BG11, and dark with distilled water.

LM21 that binds to heteromannan was significantly higher in the 12 h light with BG11 (One-way ANOVA, F=131, P=<0.05). At 48 h, 12 h light with distilled water had the highest level of heteromannan, with dark and 12 h light with distilled water having the lowest level (Figure 5, G; Mann-Whitney U, W=126, P=<0.05). At 72 h, 12 h light with BG11 had the highest amount of heteromannan compared with other treatments. At 96 h, dark with distilled water had the highest amount. After 120 h, 12 h light with BG11 had the highest amount of heteromannan alongside light with BG11 and dark with 12 h light with distilled water, continuing until 192 h (Figure 4, G-E; Mann-Whitney U, W=92, P=<0.05).

After 24 h of planting, LM25 that binds to xyloglucan remained unchanged until the signal from LM25 significantly increased in the 12 h light and BG11 (Mann-Whitney U, W=45, P=<0.05). After 48 h, light treatments had the highest levels of xyloglucan, continuing to 96 h (Figure 4, J-L). By 96 h, dark with BG11 had the lowest amount of xyloglucan; continuing until 192 h (Mann-Whitney U, W=126, P=<0.05). At 120 h, dark with distilled water had overtaken 12 h light with BG11 to have the highest amount of xyloglucan, this continued until 168 h (Figure 5, H). At 192 h, light with distilled water was significantly higher than other treatments (Mann-Whitney U, W=45, P=<0.05).

To summarise, the exclusion of light reduced the levels of LM2 that binds to AGPs (Figure 4, A and B), and LM25 that binds to xyloglucan (Figure 4, J and K) within the liquid growth media of T. aestivum. The detection of LM19 that binds to homogalacturonan, and LM21 that binds to heteromannan slightly decreased when T. aestivum was grown in the dark. When T. aestivum was grow without nutrients there were little impacts on the detection of the secreted polysaccharides.

Abiotic factors decreased the detection of xyloglucan secreted from T. aestivum roots

To explore abiotic factors affecting xyloglucan secretion, T. aestivum was grown in sodium chloride, aluminium chloride and cadmium chloride. As the concentration of the salts was increased, the amounts of xyloglucan detected declined (Figure 6, top row). There were significant decreases in the detection of xyloglucan when T. aestivum was grown between 100 µM and 150 µM of AlCI₃. This decline plateaued until 400 µM where there was another significant decrease at 450 µM (Mann-Whitney U, W=126, P=<0.05). When T. aestivum was grown in CdCI₂, there was a significant decrease in the detection of xyloglucan at 25 µM. After this decrease, xyloglucan did not significantly decline until 150 µM, 175 µM and 200 µM. The detection of xyloglucan plateaued until 400 µM where there was another significant decrease at 800 µM (Mann-Whitney U, W=126, P=<0.05). When T. aestivum was grown in NaCI, there were significant declines in the detection of xyloglucan between 0 mM and 50 mM, 75 mM and 200 mM, 300 mM and 400 mM (Mann-Whitney U, W=124, P=<0.05). The root growth of T. aestivum significantly decreased as the concentrations of aluminium, cadmium and sodium chloride increased; this was particularly noticeable when T. aestivum was grown in sodium chloride (Mann-Whitney U, W=165, P=<0.05; Figure 6, A).

Cell wall-related polysaccharides were deferentially secreted along the roots of A. thaliana using a novel technique

A. thaliana was grown on agar for two weeks, nitrocellulose sheets were then placed on top of the agar with the plants removed. These nitrocellulose sheets were probed using MAbs to reveal specific cellular locations of polysaccharide secreted along the roots of A. thaliana. The no MAb negative control, where primary antibodies were excluded from the printing protocol, lacked a signal. LM2 epitope of AGPs was clearly seen along the main root, root cap and along the lateral roots (Figure 6). The most concentrated regions of LM2 were at the lower half of the main root. LM8 that binds to xylogalacturonan was detected along the main root. No LM8 was detected at the lateral roots or caps (Figure 6). LM19 that binds to homogalacturonan was clearly visible along the main root and root cap. LM19 was also visible at the tips of the lateral roots in the upper part of the root structure (Figure 6). The MAb LM21 that binds to heteromannan was visible at the hypocotyl, and the location of the leaves and lateral roots. LM21 was also faintly visible along the main root (Figure 6). The MAb LM25 that binds to xyloglucan was highly visible across the entire root structure. Concentrated regions of LM25 were shown primarily at the tips of the lateral roots and root cap (Figure 6).

There was a co-elution of AGPs, extensin and xyloglucan within A. thaliana and T. aestivum along with three peaks of xyloglucan

To explore the structure of the polysaccharides secreted by plant roots, A. thaliana and T. aestivum exudate samples were injected into a weak anion-exchange column. A salt gradient was used to elute the polysaccharides, separating them by charge. An ELISA identified the polysaccharides contained within each fraction. There were three peaks of LM25 that binds to xyloglucan in both A. thaliana and T. aestivum (Figure 8). In A. thaliana, there was a co-elution between fractions 20 and 30 with LM2 that binds to AGPs, LM1 that binds to extensin, and the second peak of LM25 that binds to xyloglucan (Figure 8, A). There was a further elution of LM25, and LM19 that binds to homogalacturonan between fractions 30 and 40 (Figure 8, A). These co-elutions and peaks of MAb were reduced, and eluted at similar fractions when A. thaliana was grown in the dark (Figure 8, B). There was one neutral peak of LM21 that binds to heteromannan in A. thaliana grown in light (Figure 8, A). LM8 that binds to xylogalacturonan co-eluted with LM19 that binds to homogalacturonan in A. thaliana (Figure 8, A-B). Within T. aestivum, LM1 that binds to extensin, LM2 that binds to AGPs, and the second peak of LM25 that binds to xyloglucan, co-eluted between fractions 20 and 30 (Figure 8, C). When T. aestivum was grown in the dark this co-elution and the MAb peaks were reduced, and eluted at similar fractions (Figure 8, D). The co-elutions that were detected with AGPs, extensin and xyloglucan from A. thaliana and T. aestivum show that these polysaccharides were interacting within a possible polysaccharide complex. The three peaks of LM25 demonstrate that xyloglucan secreted by these plants was in three different forms: neutral, acidic and pectic.

Discussion

MAbs are useful tools to study secreted polysaccharides

This investigation has demonstrated that cell wall specific MAbs can be used to directly detect the secreted polysaccharides of plant roots. Using different sampling systems, MAbs have detected similar polysaccharides and amounts that were secreted from the plants. The major polysaccharide secreted by A. thaliana and T. aestivum were AGPs, pectin, heteromannan and xyloglucan.

Observations between the composition of Poaceae cell walls and secreted polysaccharides

Dicotyledons cell walls contain high amounts of pectin, xyloglucan and heteromannan within their cell walls (Vogel 2008). This was reflected by the SPP detected (Table 4). After screening A. thaliana, JIM7 that binds to homogalacturonan, and LM25 that binds to xyloglucan were only detected from root washing. One study screened L. angustifolius and P. sativum cell walls using JIM7, LM8, and LM25 and found that homogalacturonan, xylogalacturonan, and xyloglucan were heavily featured (Willats et al. 2004). In this study, JIM7, LM8, and LM25 were also featured within their secreted polysaccharides. This confirms that the cell wall polysaccharides in dicotyledons match what would be expected if their secreted polysaccharides derived from their cell walls (Vogel 2008; O’Neill and York 2003; Ishii 1997).

Within Poaceae, MAbs that bind to pectin were low, matching their cell walls, which only contain trace amounts. LM11 and LM12, which bind to xylan, were high in the SPP of T. aestivum and Z. mays (Table 4). This matched the profile of Poaceae cell walls (O’Neill and York 2003). Furthermore, low levels of heteromannan were detected, again matching the profile of Poaceae cell walls. However, LM25 that binds to xyloglucan was highly detected within Poaceae secreted polysaccharides. This was surprising as xyloglucan only represents up to 5% of Poaceae cell wall dry weight (Ishii 1997; O’Neill and York 2003). This suggests that xyloglucan may be secreted directly from the Golgi apparatus, independent of the cell walls; supporting previous research (McCully and Sealy 1996; Foster 1982). Mucilage may even be formed from subsets of secreted polysaccharides with xyloglucan having a higher diffusion rate into the soil, leaving pectin and AGPs to form the gel, which surrounds the root caps. The reason behind this segregated secretion of xyloglucan from other mucilage polysaccharides remains unclear.

When T. aestivum was grown in liquid culture, the first six MAbs detected matched the MAbs screened from root washing (Table 5). However, there were elevated levels of LM1 that binds to extensin in the media. The MAb LM1 that binds to extensin is known to be secreted from the leaves as a wound response (Smallwood et al. 1995). This suggests that the leaves were subjected to damage whilst the plants were agitated. After screening T. aestivum root washed liquid, extensin was still detected. High levels of LM19 instead of JIM7 were evident in A. thaliana, which both bind to homogalacturonan. LM25 that binds to xyloglucan was also detected, matching the profile of a dicotyledon cell wall (Vogel 2008). The MAbs signals in both liquid media and root washed liquid were comparable, validating both techniques. Since more A. thaliana seeds were utilised in liquid culture, it was expected that more MAb signals would be detected.

Root growth depended on the size of the seed endosperm

When in the dark, A. thaliana root growth significantly decreased, whereas T. aestivum root length increased. It has been observed that the root length of L. angustifolius and Z. mays grown in the dark extend, and it is for this reason why other research grew their plants in the dark in order to directly collect mucilage (Read and Gregory 1997; Read and Gregory 1999). T. aestivum responded as expected by increasing root lengths. However, A. thaliana root length decreased when grown without light. This may be due to the size of A. thaliana endosperm which holds less carbohydrates compared to T. aestivum, thus roots cannot grow as long without light.

Daily monitoring of the major secreted polysaccharides from A. thaliana and T. aestivum roots

For the first time, the rate of polysaccharides secreted from A. thaliana and T. aestivum roots have been determined over an eight day period. The polysaccharides detected from A. thaliana grown in light with BG11 peaked at 96 h: LM25 that binds to xyloglucan peaked at 168 h. After plateauing, xyloglucan detection decreased, perhaps by enzyme activity. This activity is contained within the root exudate (Dennis et al 2010; Moody et al. 1988; Ray et al. 1988). When A. thaliana was grown in the dark, polysaccharides secreted decreased. This decrease was less noticeable in light with distilled water. When grown in the dark, the detection of secreted polysaccharides was less despite the presence of nutrients. Deprived of light, plants cannot produce carbohydrates through photosynthesis (Albersheim et al. 2010), thus their polysaccharide secretion would naturally decrease. Nutrient deficiency did not greatly impact upon the detection of A. thaliana secreted polysaccharides. These time series have demonstrated that LM2 that binds to AGPs and LM19 that binds to homogalacturonan peaked at 96 h, whereas in the dark these peaks are unclear. LM21 that binds to heteromannan also peaked at 96 h. This had shifted later to 144 h and 192 h in the dark, suggesting a slower rate of secretion. LM25 that binds to xyloglucan rapidly declined in the dark; peaking earlier which indicates a faster rate of secretion. When A. thaliana was grown in the dark, the root lengths declined. The declines in secreted polysaccharides; together with deceases in root length suggests a link between root growth and the rate of polysaccharide secretion.

When deprived of light, AGPs, pectin and heteromannan secreted had decreased within T. aestivum. However, xyloglucan increased when T. aestivum was subjected to dark with distilled water. This observation was further puzzling when T. aestivum was grown in dark with BG11; there was a reduction in xyloglucan. There was a clear peak of LM25 that binds to xyloglucan at 96 h in light with BG11, which shifted to 120 h in the dark with distilled water, again shifting to 144 h in the dark with BG11. These peak delays indicate a slower rate of xyloglucan secretion. However, AGPs and heteromannan shifted forward; this indicated an earlier rate of secretion. When T. aestivum was grown in the dark with BG11 and distilled water, root lengths increased. Since there was an increase of xyloglucan in the dark, together with an increase in root length, it is probable that there is a link. When T. aestivum was grown in varying concentrations of NaCI, AlCI₃, and CdCI₂ the levels of xyloglucan detected significantly decreased. This was unsurprising, as the root lengths declined as well as the germination rates, leaving fewer plants and surface area for polysaccharides to be secreted.

Cellular location of cell wall-related secreted polysaccharides

There have been many reports about the location of mucilage secretion. The majority of research suggests that mucilage is solely secreted at the root caps. Other research suggests that mucilage secretion is more widespread, including, the epidermal layers behind the caps and lateral roots. However, this research has precisely detected mucilage polysaccharides to determine the location of secretion. It is apparent that different mucilage polysaccharides are secreted in different locations, thus mucilage is differentially secreted along the entire root system. LM2 that binds to AGPS was viable along the main root and at the lateral roots. LM8 that binds to xylogalacturonan was visibly punctate along the root, whereas in other research LM8 was located to P. sativum root caps (Figure 7; Willats et al. 2004). LM25 that binds to xyloglucan was secreted along the entire root structure particularly at the root cap, and lateral root tips. Since xyloglucan was detected in high levels along the roots it is reasonable to suggest that xyloglucan is secreted from root hairs, as well as root caps and lateral tips. One study uncovered that the cell walls of A. thaliana root hairs contain an acidic form of xyloglucan (Pena et al. 2012). This acid xyloglucan may be secreted along the roots but not at the root caps. The reasons why polysaccharides are secreted differentially remains undetermined.

AGPs, extensin, and xyloglucan interacted in A. thaliana and T. aestivum

When A. thaliana was grown in light with liquid BG11, extensin and xyloglucan co-eluted. This co-elution indicates that these polysaccharides interacted or are a part of a polysaccharide complex. Within T. aestivum, AGPs, extensin and xyloglucan also co-eluted, suggesting that they also interact. LM8 that binds to xylogalacturonan and LM19 that binds to homogalacturonan co-eluted within A. thaliana. This indicates that the pectin secreted by A. thaliana roots contained less detectable xylogalacturonan than homogalacturonan. Since these polysaccharides are secreted, less adhesion properties to the cells may be required for their secretion. Previous studies suggest that xylogalacturonan is involved in cell adhesion of suspended cultures of Daucus carota (Willats et al. 2004; Kikuchi et al. 1996).

Interestingly, there were three forms of xyloglucan in both A. thaliana and T. aestivum. These three peaks suggest that there were neutral xyloglucan, acidic xyloglucan, and pectic-xyloglucan. The peaks of xyloglucan were also descending in size, indicating a larger amount of neutral residues, fewer acidic and fewer pectic residues. One study identified an acidic xyloglucan was specific to root hair cell walls of A. thaliana (Pena et al. 2012); this possible interaction has not been demonstrated within T. aestivum. It is probable that the second peak of xyloglucan was this acidic xyloglucan. Probing the agar of A. thaliana with nitrocellulose, revealed the presence of xyloglucan across the entire root system; this suggested the involvement of root hairs in the secretion of xyloglucan. Perhaps this observation was the secretion of this acidic xyloglucan. This pectic-xyloglucan interaction was demonstrated in Rosa sp. cells (Thompson and Fry 2000). Prior to this study, xyloglucan had not previously been shown to be in three different forms, occurring together or in the secreted polysaccharides of A. thaliana and T. aestivum.

Future work

There still remain many unanswered questions about secreted polysaccharides, including the origins and mechanisms of secretion, legacy, function, and their interaction with the soil, soil-dwelling insects, fungi, and bacteria. Research using polygalacturonic acid (PGA) as a model for mucilage and Z. may mucilage increased the stabilisation of aggregate size within soils (Traore et al. 2000; Morel et al. 1991). To examine soil aggregation stability, standards representing the major polysaccharides secreted including gum arabic, PGA, and xyloglucan from tamarind seed, would be added to sandy, loamy, and slit soils in varying concentrations. This analysis would determine if these polysaccharides have an effect on soil aggregate sizes. Furthermore, using these polysaccharide standards to grow various symbiotic and parasitic soil-dwelling bacteria, fungi, and insects would determine if these micro-organisms could metabolise them. Growing A. thaliana in sterilised soil and extracting the secreted polysaccharides may uncover the level of solubility of these polysaccharides, using an ELISA. Furthermore, analysing the polysaccharides secreted by A. thaliana mutant XXT1/2, which lacks xyloglucan in cell walls, may determine a role of xyloglucan for micro-organism growth. If micro-organism growth is minimal or not evident, secreted polysaccharides from the wild type could be introduced to see if these micro-organisms recover their growth.

The Epitope Detection Chromatography analysis revealed several co-eluting polysaccharides within A. thaliana and T. aestivum. Characterising these co-elutions using a sandwich ELISA would determine if these polysaccharides are interacting. Further experiments could occur to determine how these polysaccharides may be linked. Nitrocellulose sheets could be used to probe the roots of T. aestivum, to determine if the secreted polysaccharides were differentially secreted. Growing T. aestivum and A. thaliana in different light, dark, and nutrient regimes would determine any alterations in the cellular location of secreted polysaccharides. Expanding the root washing screen on other plants would develop a SPP database. This database would enable a better understanding of how SPP composition differs between genera and perhaps species.

Conclusion

The secreted polysaccharides from A. thaliana, L. angustifolius, P. sativum, Z. mays and T. aestivum roots were screened, which uncovered distinct differences in their secreted polysaccharides. The Poaceae species secreted xylans and less pectin compared to their dicotyledon counterparts. Surprisingly, there was an abundance of xyloglucan secreted by T. aestivum and Z. mays, as well as the dicotyledons. Xyloglucan may in actual fact be a subset of mucilage, which is highly diffuse while pectin and AGPs remain at the root caps forming a viscous gel. When A. thaliana and T. aestivum were grown in the dark and with nutrient, secreted polysaccharides signals were reduced along with root lengths. In particular, the detection of xyloglucan in the liquid media was light-dependent. Secreted polysaccharides were also reduced along with root growth, when T. aestivum was grown in varying salt, aluminium and cadmium concentrations. This suggests a link between root length and the amounts of secreted polysaccharides. A. thaliana secreted polysaccharides were differentially secreted across the root system. AGPs were visible along the main root and ends of the lateral roots. Pectin was visible along the main root and root cap. Heteromannan was detected on the hypocotyl and the leaves. Xyloglucan was secreted across the entire root system, particularly at the root cap and lateral tips. The high levels of xyloglucan along the roots of A. thaliana may suggest that the root hairs secreted xyloglucan.

There were three different forms of xyloglucan detected, which were secreted from the roots of A. thaliana and T. aestivum. These three forms included a neutral xyloglucan, an acidic xyloglucan, and a pectic-xyloglucan. These three forms of xyloglucan have been shown to occur within the cell walls but have not been shown to occur together or within secreted polysaccharides. Finally, AGPs, extensin and xyloglucan co-eluted in A. thaliana and T. aestivum, suggesting that they may interact as a part of a polysaccharide complex.

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