Organic anions in the rhizosphere of Al-tolerant and Al-sensitive wheat lines grown in an acid soil in controlled and field environments.
Aluminium (Al) toxicity at low soil pH is a major problem in much of the arable land around the world (von Uexkull and Mutert 1995). Plants have a suite of mechanisms to deal with this hostile soil condition, including exudation of organic anions from roots (Ryan et al. 2001). An example of this mechanism has been intensively studied in the near-isogenic lines of wheat ET8 and ES8 (>95% genetic similarity, differing in the Altl locus) (Fisher and Scott 1983; Delhaize et al. 1993, 2004). ET8, the tolerant line, releases high concentrations of malate from the root tips in the presence of Al, which is thought to complex with monomeric [Al.sup.3+] within the rhizosphere of the tip to prevent toxicity (Ryan et al. 1995).
To date, organic anion exudation, particularly malate, has been quantified for ET8 and ES8 lines in solution culture using excised tips of wheat seedlings (Ryan et al. 1995; Kataoka et al. 2002). To our knowledge, malate efflux has not been quantified in the field for these lines, although organic anions have been detected in soil for other wheat lines (Pearse et al. 2006), and for other species including Lupinus albus (Dinkelaker et al. 1989) and Banksia integrifolia (Grierson 1992). Solution culture experiments of exudation processes avoid the chemical and biological complexity in soil. This complexity presents challenges for sampling organic anions in situ at the rhizosphere scale, due to the potential for microbial (biodegradation) and chemical (sorption) removal of these compounds from the soil solution (Jones 1998; Jones et al. 2003).
As a significant constraint to in situ organic anion sampling from soil-grown plants has always been access to the rhizosphere, and the availability of methods for localised collection, this study investigated different rhizosphere sampling methods for quantification of organic anions around the roots of ET8 and ES8 wheat plants in soil systems. A controlled environment experiment evaluated the use of anion exchange membranes for organic anion sampling at discrete locations along plant roots, and a field experiment measured rhizosphere organic anions at various developmental stages over a growing season, when grown in a highly acidic soil. A malate sorption experiment was also conducted to determine how much malate may be bound to the soil by biological or chemical processes, and how much can be desorbed, to provide context to the organic anion concentrations in the root-adhered soil and bulk soil samplings from the field experiment.
Materials and methods
Controlled environment experiment
Topsoil from a yellow Kurosol (Isbell 1996) was collected from near Chiltern, in Victoria, Australia (36[degrees]S, 146[degrees]E). The soil was air-dried, sieved, and packed into PVC pots (150 mm diameter, 300 mm depth) and wet up to 75% of field capacity. The soil pH was 5.33, as measured in a 1 : 5 soil: 0.01M Ca[Cl.sub.2] suspension. Two wheat plants (ET8 or ES8) were planted into each pot, with each treatment replicated 3 times in a randomised block design. Plants were watered with a 1% nutrient solution (Hoagland and Arnon 1938) to the same water content by weighing the pots and determining the amount of water lost.
Plants were sampled at the 6-leaf stage by laying the pot on its side and cutting away a section of the PVC pot without disturbance to the plant or soil. Three nodal roots were gently exposed and sampled non-destructively with anion exchange membranes placed on tip and mature root regions. While previous studies have focussed on sampling of the seminal roots, the selection of nodal roots arose from the observation that these roots exhibited greater growth and development than the seminal roots at the time of sampling (6-leaf stage).
Anion exchange membranes (BDH Laboratory Supplies, No. 55164 2S) were cut into pieces 1 by 2mm, soaked in deionised water (d[H.sub.2]O) for 24 h, loaded with 4 exchanges of 0.5 M NaHC[O.sub.3], and stored in 0.1 M NaCl until use. Membrane pieces were gently placed on exposed root tips (the apical 2-5 mm of the root which was free of adhered soil) and mature root regions (>50mm behind the tip enclosed in a rhizosheath) of nodal roots. After 10min the membranes were removed (optimum exposure time determined by prior validation experiments), immediately placed in 0.5 M HCl, and stored at 4[degrees]C for several hours with gentle shaking to ensure elution of organic anions from the membrane surface. An aliquot of the HCl solution was centrifuge-filtered (0.22 [micro]m filter) at 2800g for 5 min and frozen until analysis by high pressure liquid chromatography (HPLC). A small bulk soil sample was taken from approximately the same depth as the sampled nodal root tips. The soil samples were moistened to approximately 1 g soil to 0.7 mL d[H.sub.2]O (with duplicate samples dried and weighed to account for actual moisture contents), shaken for 10min, and centrifuged at 2800g for 10min, and then the supernatant was centrifuge-filtered (0.22/[micro]m filter) at 2800g for 10 min and frozen until HPLC analysis (see below for HPLC analysis details).
To determine elution efficiency and detection limit for malate of the membrane method, 2 membrane pieces (2 by 2 mm) were added to 1.5-mL centrifuge tubes with 10-300 [micro]M malate for 10 min. Pieces were then removed with forceps, shaken to remove excess solution, eluted with 0.5 M HCl for 10 min, filter-centrifuged (0.22 [micro]m) at 2800g, and immediately analysed by HPLC.
There was near complete recovery of malate added at 10 [micro]M (98%). However, as the concentration of added malate increased, the recovery level decreased, stabilising at 42% recovery at 50-300 [micro]M malate addition (P < 0.05).
A field site was established near Chiltem, in Victoria, Australia (36[degrees]S, 146[degrees]E), on an acidic, yellow Kurosol (Isbell 1996) in May 2002. Soil at the field experiment site was more acidic than that collected for the controlled environment experiment previously discussed. The wheat was sown using regional district practices. Each treatment was replicated 5 times in a randomised block design, with a plot size of 4 by 1 m. Most roots were restricted to the top 100mm, as at this depth an abrupt texture change occurred (clay increased from 9.0 to 34%), causing a hard layer that severely impeded root growth.
Plants were sampled at the 4- and 6-leaf stages, and at flowering. The plants were dug up (8 plants per plot), and returned to the laboratory. The roots were excised from the shoots, bulked together from each plot, and shaken to remove adhered soil ('root-adhered soil'), from which organic anions were extracted (described below). Bulk soil was also collected from each plot, and treated in the same way as the root-adhered soil. The intact roots were washed in a volume of 0.2 mM Ca[Cl.sub.2] which was twice the root fresh mass. This 'root washings' solution was also analysed for organic anions (described below). After washing, the roots were stored in 100% ethanol at 4[degrees]C until stained with a 2% methyl violet dye solution, and scanned for total root length, using a root scanning system (Delta-T Devices Ltd, UK). Shoots were dried at 60[degrees]C for 24h and weighed. At the end of the growing season all plots were harvested and grain yield determined.
For the root-adhered and bulk soil, 0.5 g was weighed in duplicate into sterile 1.5-mL centrifuge tubes, wet with 0.35 mL HPLC-grade water (BDH Laboratory Supplies), and shaken for 10min. The solution pH was measured with a micro-probe (Radiometer, XC161) before centrifuging at 2800g for 10 min. The supernatant was removed and centrifuge-filtered (0.22-[micro]m filter) at 2800g for 10min. For the 'root washings', duplicate 0.5-mL aliquots were centrifuged and filtered in the same way as the soil solutions. The root-adhered soil solution, bulk soil solution and root washings were frozen immediately after centrifugation at -20[degrees]C until HPLC analysis for organic anions (see below).
At the end of the growing season, 4 bulk soil samples were taken from each experimental plot and combined for each soil depth (0-25, 25-50, 50-100mm). These were dried at 40[degrees]C for 24h and ground (<2 mm diameter). Soil pH was measured in a 1:5 soil : 0.01 M Ca[Cl.sub.2] suspension. To measure [Al.sup.3+], soil samples were moistened to field capacity for 24 h. Soil solutions were obtained by centrifugal drainage of the moist soils and filtered through a 0.025-[micro]m membrane filter (Millipore, VSWP) to avoid contamination from Al-containing microparticulates in the measurement of monomeric [Al.sup.3+] (Menzies et al. 1991), which was determined by a short-term colourimetric assay using pyrocatechol violet (Kerven et al. 1989). The soil pH and [Al.sup.3+] values are presented in Table 1.
Malate sorption/desorption experiment
Topsoil (0-100 mm) from the field site was dried at 40[degrees]C and sieved to < 2 mm diameter. Duplicate 2.5-g samples of soil were weighed into 70-mL acid-washed screw-top polyethylene containers, with 25 mL of L(-) malic acid (Sigma-Aldrich) added at 4 concentrations (0, 2.5, 5, 10 [micro]mol/g). These were shaken for 9 time periods (0, 5, 15, 30, 45min, 1, 2, 4, 8h) by end-over-end shaking at 29 r.p.m. After shaking, a 1-mL aliquot of solution was removed, filtered through a 0.22-[micro]m filter, and frozen at -20[degrees]C. This sample was analysed for residual malate by HPLC to determine the loss of malate to soil chemical and biological processes. The remaining suspended soil was washed onto a Whatman No. 1 filter and air-dried overnight. Duplicate 0.5-g subsamples of air-dried soil were weighed into sterile 1.5-mL micro-spin centrifuge tubes (Eppendorf), wet up with 0.35mL d[H.sub.2]O, shaken on an oscillating shaker for 10min at 20[degrees]C, and centrifuged for 10mix at 1400g. The supernatant was removed, centrifuge-filtered (0.22 [micro]m) for 10 min at 2800g, and frozen at -20[degrees]C until analysed by HPLC to determine the potential for malate desorption.
Organic anion analysis by HPLC
All frozen samples were removed from the -20[degrees]C freezer 20 min before injection (samples were not concentrated) into a Waters 501 HPLC system (Waters Pty Ltd, USA), equipped with a HPX-87H Aminex column (Bio-Rad Laboratories, USA) to detect organic anions according to their retention times and absorbance peaks at 212nm. The mobile phase was degassed 0.008 N [H.sub.2]S[O.sub.4] (filtered under vacuum to < 0.22 [micro]m), at a flow rate of 0.6 mL/min and run time of 20 min. Standards (Bio-Rad Laboratories, USA) of oxalic acid, citric acid, malic acid, aconitic acid, succinic acid, formic acid, and acetic acid were used to calibrate the system and generate standard curves to calculate the concentrations of the samples. Oxalate results were not reported due to lack of detection. HPLC grade water filtered to 0.22 [micro]m was used in all HPLC applications.
GENSTAT[TM] version 8 (GENSTAT 8 Committee 2005) was used to determine treatment effects by 1- and 2-way analysis of variance (ANOVA), with linear regression used to determine relationships between variables.
Controlled environment experiment
Organic anions at root tip and mature root surfaces were detected with the anion exchange membrane technique (Fig. 1). The root tips of ES8 and ET8 had similar organic anion profiles, with no significant differences in exudation of citrate, malate, aconitate, succinate, formate, or acetate between the ES8 and ET8line. There were also no statistically significant differences in the concentrations of citrate, malate, aconitate, succinate, or formate detected at the 2 different sampling regions. However, acetate concentrations were significantly higher when measured on the mature regions than on the tips (P<0.05; Fig. 1). Of the organic anions measured, succinate was detected at the greatest concentrations; malate, aconitate, formate, and acetate levels were similar; while citrate concentrations were very low.
The concentrations of exudates measured from root tips and mature regions indicate that poor absorption of organic anions onto anion exchange membranes was not a constraint to collection, as the elution efficiency of these membranes (as measured by malate) is greatest at low anion concentrations.
The bulk soil of the controlled environment experiment contained detectable concentrations of succinate, formate, and acetate (Fig. 1). Succinate was the dominant organic anion measured, at concentrations at least 10-fold greater than formate and acetate (P<0.001).
Shoot biomass did not differ between ET8 and ES8 at the 4- and 6-leaf stages, but was greater for ET8 at flowering (P<0.05; Fig. 2). There were no significant differences in root length or grain yield between ET8 and ES8 (Fig. 2).
Succinate was the only organic anion detected in the bulk soil, with concentrations increasing from 0.033 [micro]mol/g at 4-leaf, 0.174 [micro]mol/g at 6-leaf, and 0.426 [micro]mol/g at flowering stages (P < 0.001).
A range of organic anions was detected in the root-adhered soil and root washings (Fig. 3), with detection and concentrations dependent on the collection method (root-adhered soil and root washings) and sampling time. In the root-adhered soil we found no evidence of differences in organic anion concentrations between ES8 and ET8, including malate.
In the root washing samples, aconitate and succinate concentrations were significantly greater in ES8 at the 4-leaf stage than in ET8 (P<0.05). Acetate concentrations in root washings were also significantly greater in ES8 than ET8 at both the 4- and 6-leaf stages (P < 0.05). However, there were no differences in malate concentrations between ES8 and ET8. There was a significant effect of sampling time/plant age on organic anions measured in the root washings (Fig. 3), with concentrations of citrate, malate, aconitate, and succinate all decreasing at flowering (P < 0.05). Acetate concentrations in ES8 also significantly decreased with increased plant age (P < 0.005).
Significant relationships were measured between malate concentrations and concentrations of the other organic anions except formate (P < 0.001; Fig. 4). As malate concentrations increased, so too did concentrations of citrate, aconitate, succinate, and acetate (Fig. 4). Although evident in both root-adhered soil and root washings, this trend was most significant in the root washing samples.
[FIGURE 1 OMITTED]
Organic anion concentrations measured in the root-adhered soil and root washings were also significantly related for malate ([R.sup.2] = 0.55, P < 0.001) and all other organic anions except formate (P < 0.001). For example, if malate concentrations in rhizosphere soil increased, malate concentrations in root washings also increased. This indicates that although different units of measure were used for each sampling method (expressed per root length or adhered soil), each organic anion reported a similar process/chemistry that was due to the roots and/or root-associated organisms.
Malate sorption/desorption experiment
Malate sorption increased with time, with greatest sorption after shaking for 8 h (Fig. 5). The malate sorbed after 8 h decreased with increasing initial malate concentration. At 2.5 [micro]mol/g malate addition, 85% malate was adsorbed, while at 10 [micro]mol/g addition, only 42% was adsorbed. None of the malate added could be re-extracted using the method outlined.
[FIGURE 2 OMITTED]
[FIGURE 3 OMITTED]
This study has shown that several methods can be used for the collection of organic anions from the rhizosphere of wheat plants. The data presented from the controlled environment experiment demonstrate the application of anion exchange membranes for non-destructive collection of organic anions from discrete regions of soil-grown plants (Fig. 1). A similar approach to localised collection has been used previously, whereby moist filter paper was used as the absorbent, applied to different sections of the root (Zhang et al. 2002). However, comparative experiments showed filter papers to have a poorer elution efficiency than anion exchange membranes (data not shown). The benefits of using anion exchange membranes include ease of application, control over the sampling region, the ability to sample growing plants in situ, and the range of organic anions which can be collected., There are, however, 2 considerable shortcomings inherent in the use of anion exchange membranes or similar materials for localised exudate sampling in soil-grown plants. These are (i) the high degree of variability in the concentrations of anions collected across a root system (as evident in Fig. 2), and (ii) the lack of simultaneous exposure of the root tip to both the membrane and the associated stimulant to exudation (e.g. [Al.sup.3+]).
[FIGURE 4 OMITTED]
[FIGURE 5 OMITTED]
Using anion exchange membranes, a selection of organic anions was detected on the root tips and mature regions of nodal roots from plants grown in the controlled environment. However, there were no differences in organic anion profiles between the ES8 and ET8 wheat lines. Although exudation of malate is thought to be the mechanism conferring AI tolerance in the field for ET8, leading to better root extension and increased yields, this response has only been measured when the ET8 is challenged with high Al in solution culture experiments (Delhaize et al. 1993; Basu et al. 1994; Ryan et al. 1995).
This experiment validated the use of anion exchange membranes for localised collection of organic anions from roots, as demonstrated by the suite of anions measured. However, the relatively high pH of this soil (pH 5.33) and the separation of root from Al in the soil at the time of sampling suggest that the environment did not challenge the wheat plants sufficiently to evaluate differences in rhizosphere malate concentrations between the ES8 and ET8 lines.
The field experiment provided an opportunity to investigate the performance of ET8 and ES8 in a highly acidic soil over a complete growing season. Although the soil was highly acidic, with high levels of [Al.sup.3+] in solution, the only physiological difference observed between ES8 and ET8 was increased shoot biomass in ET8 at flowering. However, there was a trend of increased root length and greater grain yield in the ET8 wheat line. Previous field studies have also measured greater ET8 shoot growth after the 6-leaf stage and greater grain yield in ET8 than ES8 (e.g. Tang et al. 2002).
The field experiment utilised 2 established methods of organic anion collection: rhizosphere soil and root washings. Although a range of organic anions was detected with both methods, neither of these methods detected differences in organic anion profiles, including malate, between ES8 and ET8.
Malate may have been released from the roots of ET8 in response to [Al.sup.3+], but may become immediately sorbed to soil surfaces. In the field, the highest level of malate measured in root-adhered soil was 2.58 [micro]mol malate/g soil. In the malate sorption experiment, when 5.0 [micro]mol/g of malate was added, only 2.32 [micro]mol/g of malate was retained in solution after 8 h (Fig. 5). Therefore, the malate in the root-adhered soil may be <50% of the malate initially exuded. This is supported by malate sorption isotherms by Jones et al. (1996b), who demonstrated that malate preferentially binds to solid phase sorption sites rather than metals in solution. This also suggests that the efficiency of malate complexation as a mechanism for Al detoxification in a soil environment may be quite low. Rapid consumption of malate by soil organisms may also reduce detectable malate concentrations, as previously shown by Jones et al. (1996b).
Furthermore, the soil pH may not have been low enough to promote sufficiently high A1 concentrations and malate efflux. The soil in the field was pH 3.9, similar to the pH of solution experiments (pH 4.2) which stimulated efflux of malate from the roots of ET8 (Ryan et al. 1995). However, the soil solution [Al.sup.3+] measured in the field experiment (31 [micro]M) was in the low range of concentrations used in solution culture (200 [micro]M [Al.sup.3+]); malate exudation at 31 [micro]M [Al.sup.3+] was only half of that measured at 200 [micro]M [Al.sup.3+] (Ryan et al. 1995). Although the concentrations of [Al.sup.3+] in the field may be phytotoxic to wheat (Slattery et al. 1995), the [Al.sup.3+] measured in soil solution may not have been high enough to trigger a detectable malate exudation response in the ET8 line in soil, as measured by the methods used here.
A proportion of the released malate may also have chelated Al within the apoplast of the root, a possibility put forward by Kinraide et al. (2005).
The range of organic anions detected in the rhizosphere of ES8 and ET8 was different from that of the bulk soil. The age of the wheat plant also had a significant effect on rhizosphere organic anions, likely due to changes in plant physiology associated with phenology, and with root development and aging. Various organic anions were detected around the root in synchrony (Fig. 4), possibly because they are all related to activities of the TCA cycle and other metabolic processes (Basu et al. 1994) and diffuse away from the root along a concentration gradient. The concentration and range of organic anions in the rhizosphere decreased by the time of flowering, possibly due to reduced root growth during the reproductive stage, reduced plant-available soil moisture, or a reallocation of metabolites for grain-filling (Yang et al. 2001).
The measured rhizosphere organic anions may have been derived from the plant roots or from soil organisms. Several of the organic anions, such as acetate, formate, and succinate, are metabolised by rhizosphere microorganisms (Stevenson 1967; Vance et al. 1996). The synthesis of organic anions by microorganisms is supported by the presence of succinate in non-rhizosphere 'bulk' soil samples. The dramatic increase in succinate in bulk soil at the flowering stage of sampling is of particular interest, due to the dry soil conditions at this time (soil moisture data not shown). The succinate measured at flowering may have been released upon microbial cell lysis induced by the onset of dry conditions.
In conclusion, rhizosphere organic anions have been linked with tolerance to Al toxicity (Delhaize et al. 1993; Basu et al. 1994; Ma et al. 2001; Li et al. 2002) and deficiencies of phosphorus (Strom et al. 2002; Veneklaas et al. 2003; Dong et al. 2004), iron (Jones et al. 1996a), and zinc (Rengel 2002). The exudation of organic anions from wheat has also been associated with cadmium uptake (Cieslinski et al. 1998) and copper toxicity (Nian et al. 2002). These studies highlight the inherent difficulty in assigning causal factors responsible for stimulating organic anion exudation in complex soil systems, whereby more than one stimulant may be active at any time. Therefore, the challenge for further soil-based research into organic anion exudation is not only to identify the dominant stress that has triggered the response (i.e. Al toxicity, P deficiency), but also to clearly differentiate between plant-and microbially derived contributions. This study highlights the inherent difficulty in detecting organic anion exudation in complex soil systems, even with 3 methods.
Spatial detection and quantification of organic anions in rhizosphere and bulk soil systems is an issue which is yet to be resolved with current techniques. Synchrotron-based X-ray technology is one approach which may be able to determine the organic anions present in these systems and the complexes formed in situ, at a resolution not previously possible. Such applications may provide major advances in our knowledge of the specific roles of organic anions in soil systems.
This work was funded through the Science, Technology and Innovation (STI) program of the Department of Innovation, Infrastructure and Regional Development (DIIRD), Victoria, and from the GRDC for MW. We are grateful to Dr Caixan Tang of La Trobe University for providing the seed, Greg Codes for the access to the field site, and Dr Maartin Hens, CSIRO Plant Industry, for assistance with the anion-exchange membrane method. Thanks also to Dr Erik Veneklaas, University of Western Australia, for advice on collecting root leachates and reviewing the manuscript, and Drs Matt Denton and Kirsten Barlow, Department of Primary Industries, Victoria, and Dr Alan Richardson, CSIRO Plant Industry for reviewing the manuscript.
Manuscript received 19 September 2007, accepted 13 February 2008
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C. R. Schefe (A,D), M. Watt (B), W. J. Slattery (C), and P. M. Mele (A)
(A) Primary Industries Research Victoria (PIRVic), Department of Primary Industries, Rutherglen Centre, RMB 1145, Rutherglen, Vic. 3685, Australia.
(B) CSIRO Plant Industry, GPO Box 1600, Canberra, ACT 2601, Australia.
(C) Australian Greenhouse Office, GPO Box 621, Canberra, ACT 2601, Australia.
(D) Corresponding author. Email: firstname.lastname@example.org
Table 1. The pH and [Al.sup.3+] values for the field experiment Samples were collected in each plot, with results averaged across each depth Soil solution Depth (mm) Soil pH (Ca[Cl.sub.2]) [Al.sup.3+], ([micro]M) 0-25 4.14 22.6 25-50 3.83 33.9 50-100 3.74 36.1
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|Author:||Schefe, C.R.; Watt, M.; Slattery, W.J.; Mele, P.M.|
|Publication:||Australian Journal of Soil Research|
|Date:||May 1, 2008|
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