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First published online July 27, 2007; 10.1104/pp.107.102624 Plant Physiology 145:266-276 (2007) © 2007 American Society of Plant Biologists Effect of Secondary Metabolites Associated with Anaerobic Soil Conditions on Ion Fluxes and Electrophysiology in Barley Roots1,[C]School of Agricultural Science and Tasmanian Institute of Agricultural Research, University of Tasmania, Hobart, Tasmania 7001, Australia
The effects of secondary metabolites produced by waterlogged soils on net K+, H+, and Ca2+ fluxes were studied in the mature zone of roots of two barley (Hordeum vulgare) cultivars contrasting in their waterlogging (WL) tolerance using the noninvasive microelectrode ion flux measuring technique. In WL-sensitive variety ‘Naso Nijo’, all three lower monocarboxylic acids (formic, acetic, and propionic acids) and three phenolic acids (benzoic, 2-hydroxybenzoic, 4-hydroxybenzoic acids) caused a substantial shift toward steady K+ efflux, accompanied by an immediate net influx of H+. Detrimental effects of secondary metabolites on K+ homeostasis in root cells were absent in WL-tolerant ‘TX’ variety. Root treatment with Mn2+ caused only a temporary K+ loss that returned to the initial level 10 min after treatment. Phenolic acids slightly increased Ca2+ influx immediately after treatment, while other metabolites tested resulted in transient Ca2+ efflux from the root. In the long-term (24 h) treatment, all metabolites tested significantly reduced K+ uptake and the adverse effects of phenolic acids were smaller than for monocarboxylic acids and Mn2+. Treatment with monocarboxylic acids for 24 h shifted H+ from net efflux to net influx, while all three phenolic acids did not cause significant effects compared with the control. Based on results of pharmacological experiments and membrane potential measurements, a model explaining the effects of secondary metabolites on membrane transport activity is proposed. We also suggest that plant tolerance to these secondary metabolites could be considered a useful trait in breeding programs.
Owing to the anaerobic metabolism of plants or microbes, significant accumulation of toxic substances occurs in waterlogged soil (Lynch, 1977  ; Tanaka et al., 1990; Armstrong and Armstrong, 2001). Materials potentially toxic to plants that are accumulated in flooded soils include reduced manganese, iron, hydrogen sulfide, various organic acids, and ethylene (Armstrong and Gaynard, 1976). Surprisingly, to date, all breeding programs aimed at improving waterlogging (WL) tolerance in plants have focused almost exclusively on preventing oxygen loss or improving its transport to, or storage in the root (Jackson and Armstrong, 1999). Plant tolerance to these secondary metabolites has never been considered a useful trait in breeding programs.
The type and amount of organic acids produced depends upon the fermentive character of the microflora, the type and amount of organic materials added, and on the prevailing soil conditions (Rao and Mikkelsen, 1977
The extent to which the accumulation of toxic metabolites is causally linked to observed deficiencies of macronutrients in waterlogged soils is not clear. Energy deficiency, caused by lack of O2, reduces the availability of many essential nutrients including nitrogen, phosphorus, sulfur, and also most of the trace elements (Drew, 1988
Thus far, most reports have dealt with the analysis of the overall changes in ion content in plant tissues or with monitoring the kinetics of nutrient depletion in a growth solution (Glass, 1973
The major bulk of experiments were performed on the WL-sensitive variety ‘Naso Nijo’, where much stronger responses were expected. Accordingly, most of results below refer to ‘Naso Nijo’ roots unless specified otherwise.
Net K+ uptake of about 60 nmol m–2 s–1 was measured from mature epidermal root cells of 3-d-old seedlings in control (steady-state) conditions. Addition of phenolic compounds (benzoic acid, 2-hydroxybenzoic acid, 4-hydroxybenzoic acid; 200 µM working concentration) and volatile monocarboxylic organic compounds (formic acid, acetic acid, propionic acid; 10 mM working concentration) rapidly decreased net K+ influx (Fig. 1 ). Among the three different phenolic acids, 2-hydroxybenzoic acid and 4-hydroxybenzoic acid caused much more adverse effects on K+ uptake compared with benzoic acid (Fig. 1A), completely arresting net K+ uptake within 10 min after treatment. All three volatile monocarboxylic organic acids not only arrested K+ influx but also caused significant (P < 0.001) K+ efflux from barley (Hordeum vulgare) roots, with an effect increasing in the following sequence: propionic acid >> acetic acid > formic acid (Fig. 1B). This effect was specific and not related to changes in osmolality of the experimental solution, as isotonic treatment with KCl, NaCl, or Na gluconate caused no K+ efflux from barley roots (data not shown). Adding 300 mg L–1 Mn2+ caused an almost instantaneous reduction of K+ uptake, which quickly (within 5 min) returned to its initial value (Fig. 1C). In general, the effect of monocarboxylic organic acids on root K+ fluxes was significantly stronger than one caused by phenolic acids.
Transient H+ Fluxes
Net H+ efflux of 10 to 15 nmol m–2 s–1 was measured in control (steady-state) conditions from barley roots. Application of all secondary metabolites significantly (P < 0.01) affected H+ fluxes (Fig. 2
). An immediate and significant shift toward net H+ uptake was observed in response to all phenolic and monocarboxylic organic acids tested (Fig. 2, A and B), while in the case of Mn2+ treatment, net H+ efflux was significantly (P < 0.01) reduced (Fig. 2C). Among phenolics, the effect followed the ranking: 4-hydroxybenzoic acid
Transient Ca2+ Fluxes Net zero Ca2+ flux was measured in control (steady-state) conditions. Root treatment with phenolic acids led to a gradual and prolonged increase in net Ca2+ uptake (Fig. 3A ). Such a slowness of response may be indicative of a cytosolic mode of action. No significant (P < 0.05) difference between the effects of various phenolic acids was found. Adding 10 mM monocarboxylic organic acid to the bath, however, caused an immediate and substantial Ca2+ efflux from barley roots (Fig. 3B). A similar trend was observed for Mn2+ treatment (Fig. 3C). In both cases, net Ca2+ flux recovered to its original (zero) value within 10 to 15 min (Fig. 3, B and C).
Genotypical Difference No significant (P < 0.05) difference in initial (steady-state) flux levels was found for any ions measured between two contrasting genotypes (WL-sensitive ‘Naso Nijo’ and WL-tolerant ‘TX’; Fig. 4 ). However, transient flux kinetics differed substantially between genotypes. The most striking difference was observed for K+ flux (Fig. 4A). Contrary to WL-sensitive ‘Naso Nijo’ genotype, lower monocarboxylic (acetic) acid treatment did not cause any net K+ loss in the WL-tolerant ‘TX’ variety. Moreover, ‘TX’ roots even increased net K+ uptake 20 min after treatment with 10 mM acetic acid (Fig. 4A), while similar treatment caused a very substantial K+ loss from the roots of WL-sensitive ‘Naso Nijo’ variety. Also significantly reduced was ‘TX’ net Ca2+ uptake in response to 2-hydroxybenzoic treatment compared with ‘Naso Nijo’ (Fig. 4C). No clear difference was evident between genotypes in terms of H+ flux responses for either acetic or 2-hydroxybenzoic treatment (Fig. 4B). Treatment with Mn2+, however, has significantly reduced H+ efflux in WL-sensitive ‘Naso Nijo’ variety (compared with control) but was not significant in WL-tolerant ‘TX’ variety (Fig. 4B).
Long-Term Ion Flux Responses K+ uptake was significantly reduced in ‘Naso Nijo’ roots after 24 h treatment with all secondary metabolites tested (Fig. 5A ). Root treatment with phenolic compounds (benzoic acid, 2-hydroxybenzoic acid, 4-hydroxybenzoic acid) caused a significant (P < 0.01) decrease in net K+ uptake. No significant (P < 0.05) difference between the effects of various phenolic compounds was found. In monocarboxylic acid treated roots, K+ fluxes were shifted to substantial (–40 to –100 nmol m–2 s–1) net efflux. Among them, acetic acid and propionic acid caused more severe effects than formic acid. Mn2+ treatment also caused net K+ efflux. In general, the adverse effects of phenolic acids were smaller than the other four treatments.
The monocarboxylic acid treatments shifted H+ from net efflux to net influx (Fig. 5B). Mn2+ treatment reduced H+ efflux to around zero. Among the phenolic acids, 2-hydroxybenzoic acid and 4-hydroxybenzoic acid did not cause significant (P < 0.05) changes to H+ fluxes, while benzoic acid slightly reduced H+ efflux (Fig. 5B). Phenolic acids caused significant (P < 0.05) net Ca2+ efflux from roots pretreated for 24 h (Fig. 5C). Formic and acetic acids also slightly reduced net Ca2+ uptake, while propionic acid and Mn2+ did not significantly (P < 0.05) affect Ca2+ fluxes (Fig. 5C).
Effects of various channel blockers and metabolic inhibitors on ion fluxes kinetics were studied using one chemical from each group (specifically, acetic acid, 2-hydroxybenzoic acids, and Mn2+).
None of the inhibitors used significantly affected the initial Ca2+ flux after 1 h of incubation (data not shown). However, La3+ and Gd3+ (two known nonselective cation channel [NSCC] blockers; Demidchik et al., 2002
Initial K+ uptake in barley roots was strongly suppressed by TEA+ (Fig. 7A ). All these inhibitors were also efficient in reducing the magnitude of K+ flux response to 2-hydroxybenoic (Fig. 7B) and acetic (Fig. 7C) acids and Mn2+ (Fig. 7D; significant at P < 0.05). Both Gd3+ and La3+ (two known NSCC channel blockers; Demidchik et al., 2002 ) also significantly reduced initial K+ uptake and the magnitude of K+ flux response to all chemicals tested (Fig. 7; significant at P < 0.05).
Root pretreatment in 1 mM vanadate (a known inhibitor of the plasma membrane [PM] H+-ATPase) shifted the initial H+ flux from net efflux to net influx (Fig. 8 ), while no significant effect of TEA+, La3+, or Gd3+ on initial H+ flux was observed. Vanadate treatment also significantly (P < 0.01) reduced the magnitude of H+ flux changes in response to all treatments tested (data not shown).
Membrane Potential Responses and H+-ATPase Activity The average membrane potential in the mature zone of barley roots was –133.9 ± 2.0 mV in control. Phenolic compounds caused substantial membrane depolarization (as illustrated in Fig. 9A for the treatment with 200 µM 2-hydroxybenzoic acid), stabilizing at –90 mV level approximately 10 min after the treatment was applied. Membrane potential kinetics in response to other compounds was not measured.
Contrary to the short-term effects of 2-hydroxybenzoic acid, 24 h treatment with 200 µM phenolics caused significant (P < 0.01) hyperpolarization of the membrane potential (Fig. 6B). The largest hyperpolarization effect was found in roots treated with 4-hydroxybenzoic acid. All three monocarboxylic acids and Mn2+ treatments induced significant (P < 0.001) long-term depolarization of membrane potential (Fig. 6B). Results of membrane potential measurements were consistent with direct estimation of ATP hydrolytic activity from PM vesicles isolated from the microsomal fraction of barley roots (Fig. 9C). No significant (P < 0.05) difference was found in ATP hydrolytic activity between control samples and samples treated with either acetic acid or Mn2+. At the same time, the PM vesicles from roots treated with 2-hydroxybenzoic acid had about 40% higher ATP hydrolytic activity compared with control roots (significant at P < 0.05; Fig. 9C).
Secondary Metabolites Toxicity and WL Tolerance in Barley
All seven secondary metabolites associated with anaerobic soil conditions inhibited root elongation in 24 h treatments (data not shown), highlighting their detrimental effects on root metabolism. They also caused significant alterations in root membrane-transport activity even in the presence of oxygen. The most significant was a pronounced shift toward K+ efflux, caused by both phenolics and monocarboxylic acids (Fig. 1). In the case of monocarboxylic acids, the result was a very substantial K+ loss measured from the roots of WL-sensitive ‘Naso Nijo’ (Fig. 1B). Such K+ loss has been previously reported from barley roots in response to salinity (Chen et al., 2005
In this study, we extend these findings to plant WL tolerance. The WL-tolerant ‘TX’ was capable not only of completely preventing net K+ loss after acetic acid treatment (Fig. 4A) but even slightly enhancing net K+ uptake by roots under stress conditions. Also less affected (compared with WL-sensitive ‘Naso Nijo’) was K+ uptake in response to phenolics (Fig. 4A). These data support the idea that WL tolerance in barley is conferred not only by differences in root anatomy (high percentage of aerenchyma in ‘TX’ genotype; Pang et al., 2004
Early reports of Glass (1974)
Among the three phenolics, the effects of 2-hydroxybenzoic and 4-hydroxybenzoic acids on K+ flux were larger than effects caused by benzoic acid (Fig. 1). Earlier, Glass (1973)
Under the conditions of this experiment (pH 5.5), most phenolic acids in solution will be in the dissociated form. This undissociated acid concentration can be calculated according to the Henderson-Hasselbalch equation:
As shown in Table I , the amount of undissociatd acids was relatively low and comprised 4.7%, 0.3%, and 8.7% for benzoic, 2-hydroxybenzoic, and 4-hydroxybenzoic acids, respectively. Therefore, no obvious correlation between the magnitude of effect and the amount of dissociated compound was found.
The mechanisms by which phenolic compounds control K+ transport across the PM remain elusive. Based on the fact that removal of phenolics caused a rapid recovery of K+ reabsorption, Glass (1974) suggested a direct effect on cell membranes. No specific details were offered though. Our data reported in this study suggests that major voltage-dependent K+-transporting systems may be key players. This is supported by the observed immediate membrane depolarization (Fig. 9A). Moreover, our data suggest that both increased H+ (Fig. 2A) and Ca2+ (Fig. 3A) uptake could contribute to this depolarization as depicted in Figure 10
.
It is traditionally believed that most phenolic acids cross the cell membrane in an undissociated form by passive diffusion (Jackson and St. John, 1980 ). However, recent cloning and functional characterization of the MCT1 family of transporters suggest that uptake of both monocarboxylic acids and benzoic acid occur via the H+-coupled cotransport mechanism, at least in animal systems (Kido et al., 2000). We suggest that a similar scenario is also applicable to plant tissues. As undissociated phenolic acid is electrically neutral, not only will increased net H+ flux be generated as a result of such activity (consistent with our H+ flux data; Fig. 2A), but also a substantial membrane depolarization is expected (Fig. 9). Such a depolarization will affect intracellular K+ homeostasis by reducing K+ uptake via inward-rectifying K+ channels and enhancing K+ efflux via depolarization-activated outward-rectifying K+ channels (Maathuis and Sanders, 1996; Fig. 10), explaining the rapid shift toward net K+ efflux after phenolics application (Fig. 1A). As both of these channels are TEA+ sensitive, inhibition of K+ efflux by TEA+ in our experiments (Fig. 7) is consistent with this model.
Several Ca2+-permeable channels may mediate Ca2+ uptake into root cells; each of these may contribute to the observed Ca2+ influx after phenolics application. Of special interest may be depolarization-activated Ca2+ channels (Thion et al., 1998
Once inside the cell, permeated phenolic acids dissociate and acidify the cytosol (Guern et al., 1986
Theoretically, membrane hyperpolarization observed after long-term phenolic treatment (Fig. 9B) was expected to reverse the detrimental effects of metabolites on K+ transport. However, this was not the case, and K+ uptake after 24 h of treatment with phenolic acids was significantly (P < 0.05) lower than in the control (Fig. 5A). The answer may lie in the fact that net Ca2+ uptake measured soon after treatment (Fig. 3A) may result in a substantial elevation in cytosolic free Ca2+. Patch-clamp experiments on guard cells suggest that the inward K+ current is greatly reduced by elevating [Ca]cyt to micromolar concentrations (Schroeder and Hagiwara, 1989
It should be also mentioned that some benzoic acid derivatives [e.g. 5-nitro-2-(3-phenylpropylamino) benzoic acid] were found to be potent inhibitors of anion channels (Roberts, 2006
Similar to phenolics, lipid-soluble undissociated forms of the volatile monocarboxylic acids are often regarded as the most toxic (Jackson and Taylor, 1970
These authors also suggested that changes in membrane lipid composition might be responsible for the observed leak of K+ and Ca2+ from roots treated with monocarboxylic acids. However, it is highly unlikely that such a non-ion-specific change in general membrane permeability may occur almost immediately (within 1 min) after the treatment, as resolved by the MIFE system for K+ efflux in our experiments (Fig. 1B). Such changes in permeability are usually associated with a change in membrane lipid components (Jackson and Taylor, 1970 ; Glass, 1974; Jackson and St. John, 1980), the latter process most likely operating on a slower time scale. Importantly, the above changes in membrane permeability are believed to be nonspecific (Glass and Dunlop, 1974), thus, a mirror image kinetics for K+ efflux and Ca2+ uptake (according to electrochemical potential for each ion) should be observed. This was obviously not the case in our study. While the K+ leak gradually increased with time (Fig. 1B), Ca2+ efflux was short lived and returned back to control values within 10 to 15 min after treatment. This suggests that fluxes of these two ions are mediated by different transport systems, and thus cannot be attributed to a general change in membrane permeability.
A plausible alternative explanation may be offered. Similar to our model, phenolics, monocarboxylic acids are transported into the cytosol most likely in an undissociated form (Kido et al., 2000
Contrary to the effect of phenolics, monocarboxylic acids did not cause any substantial increase in Ca2+ uptake (Fig. 3, A and B, respectively), indicating a specificity of regulation of Ca2+ signaling by these secondary metabolites. The effect of specific blockers of the Ca2+-ATPase (CPA and thapsigargin) on acetic-induced transient Ca2+ efflux was rather small (Fig. 6B), suggesting a relatively minor role for the PM Ca2+ pump in this process and pointing toward a possible mediation of Ca2+ efflux by the PM Ca2+/H+ exchanger. At the same time, Ca2+ flux responses to monocarboxylic acids were completely blocked by either Gd3+ and La3+ (Fig. 6, B and C), as well as strongly inhibited by TEA+. The latter results may suggest that a substantial component of Ca2+ efflux may originate from the K+/Ca2+ Donnan exchange in the cell wall (Shabala and Newman, 2000
The above scenario is further supported by the results of long-term experiments (Fig. 5). While a substantial K+ leak was measured 24 h after treatment with monocarboxylic acids (Fig. 5A), no significant (P < 0.05) Ca2+ leak was found (Fig. 5C). Thus, it is highly unlikely that general changes in membrane permeability were involved as was suggested by Jackson and coauthors (Jackson and Taylor, 1970 In summary, this study shows that secondary metabolites associated with waterlogged soil conditions adversely affect root nutrient uptake and that the perturbation to root ionic homeostasis is much stronger in WL-sensitive genotypes. Accordingly, we suggest that tolerance to these stresses should be targeted in any program to breed crops for WL tolerance.
Plant Material and Growth Conditions
Two barley (Hordeum vulgare) varieties, WL-sensitive ‘Naso Nijo’ and WL-tolerant ‘TX9425’ (Pang et al., 2004
Net fluxes of H+, Ca2+, and K+ were measured using the noninvasive MIFE technique (University of Tasmania, Hobart, Australia). Details on fabrication and calibration of H+, Ca2+, and K+ ion selective microelectrodes have been described previously (Shabala et al., 1997
Two major groups of organic acids, namely monocarboxylic acids and phenolic acids, were chosen for experiments (Table I and II). These are the most widely reported compounds associated with anaerobic soil conditions (Lynch, 1977
One hour before measurement, 5 mL basic salt medium (BSM) solution (0.1 mM CaCl2, 0.2 mM KCl, pH 5.5 unbuffered) was added to a plexiglass measuring chamber (100 mm long, 30 mm deep, and 4 mm wide). A seedling was taken from the growth container and placed immediately into the chamber. The root was immobilized in the horizontal position by fine Teflon partitions 5 mm above the floor of the chamber as described in Pang et al. (2006) In transient experiments, steady-state fluxes were measured for 5 min, then 5 mL of BSM solution containing a double concentration of an appropriate chemical was added into the chamber, and the measurement continued for a further 30 min. Solution pH was adjusted to 5.5 in advance using NaOH/HCl, and no substantial changes in Ca2+ or Mn2+ activity was caused by addition of any of organic acids. About 2 min is required for unstirred layer conditions to be reached. This period of time was discarded from the analysis and appears as a gap in the figures. For measurement of the long-term effects of secondary metabolites on root ion fluxes and membrane potential the components studied were added to the growth plastic container (basic solution) 24 h before measurement. The final concentrations of phenolic acids (benzoic acid, 2-hydroxybenzoic acid, and 4-hydroxybenzoic acid) were 200 µM, volatile monocarboxylic organic acids (formic acid, acetic acid and propionic acid) were 10 mM, Mn2+ (added as MnSO4 salt) was 300 mg L–1; all these concentrations were selected based on previous literature reports showing they are physiologically relevant. Solution pH was adjusted to 5.5 (using HCl/NaOH) in all treatments and monitored continuously by the pH microelectrode. Solutions were aerated continuously during the 24-h treatment period.
Pretreatment with inhibitors was carried out when the root was transferred to the measuring chamber. Orthovanadate (an inhibitor of P-type ATPase), TEACl (a putative K+ channel blocker), GdCl3 and LaCl3 (NSCCs blockers), and CPA and thapsigargin (specific Ca2+-ATPase inhibitors) were used to modify the activity of selected PM transporters. These inhibitors were mixed with the basic solution (0.2 mM KCl, 0.1 mM CaCl2) to achieve their final concentrations that were as follows: vanadate, 1 mM; TEA+, 10 mM; Gd3+, 50 µM; La3+, 200 µM; CPA, 50 µM; thapsigargin, 5 µM. After 1 h pretreatment in the appropriate inhibitor, transient ion flux responses to one of the secondary metabolites were measured, as described above (still in the presence of inhibitor in the bath solution).
The roots of intact barley plants were mounted in a measuring chamber and the roots were gently secured in a horizontal position with small plastic blocks. Experimental conditions were the same as those for the ion flux measurement. The plant was allowed to stabilize for 60 min. Measurements of the electrical potential difference (Vm) across the root-cell membranes were made in the root mature zone, 1 to 2 cm from the root tip essentially as described by Cuin and Shabala (2005)
Barley was grown in vermiculite for 7 d in the dark. The vermiculite was watered with basic nutrient solution containing 0.2 mM KCl and 0.1 mM CaCl2 (BS). The roots were washed carefully and rinsed with BSM. Around 10 g roots (fresh weight) were taken into BS, with one of 20 mM acetic acid in BSM (pH 5.5), 200 µM 2-hydroxybenzoic acid in BSM (pH 5.50), or 300 mg/L MnSO4 in BSM (pH 5.5) added for 30 min. Roots were then homogenized in 200 mL ice-cold homogenization buffer (50 mM MOPS, 5 mM EDTA, 330 mM Suc, 0.6% polyvinylpyrrolidone, 5 mM ascorbate, 5 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride). PM was isolated from the mirosomal fraction (30,000 g) by partitioning at 4°C at an aqueous polymer two-phase system (9 g + 3 g) composed of 6.2% Dextran D1037 (Sigma), 6.2% PEG3350 (Sigma), 330 mM Suc, 5 mM potassium phosphate pH 7.8, 3 mM KCl, 0.1 mM EDTA, and 1 mM dithiothreitol (Larsson et al., 1994
We are grateful to Dr. A. Fuglsang (University of Copenhagen) for her valuable advice on H+-ATPase hydrolytic assay experiments and Mrs. Julie Harris (University of Tasmania) for her kind assistance with using ultracentrifuge. Received May 21, 2007; accepted July 23, 2007; published July 27, 2007.
1 This work was supported by Grain Research and Development Corporation (M.Z. and N.M.) and Australian Research Council (S.S.) grants. 
2 Present address: School of Plant Biology (MO84), University of Western Australia, 35 Stirling Highway, Crawley 6009, Western Australia, Australia. The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Sergey Shabala (sergey.shabala{at}utas.edu.au).
[C] Some figures in this article are displayed in color online but in black and white in the print edition. www.plantphysiol.org/cgi/doi/10.1104/pp.107.102624 * Corresponding author; e-mail sergey.shabala{at}utas.edu.au.
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2-hydroxybenzoic acid > benzoic acid. For monocarboxylic organic acids, responsiveness of H+ flux followed the ranking: formic acid > acetic acid > propionic acid. 
