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Physiology and Development |
Department of Biological Sciences, University of Hull, Hull, UK, HU6 7RX
Received for publication August 22, 2000. Accepted for publication January 18, 2000.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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Key Words: lignification • organic acids • Phragmites • phytotoxins • radial oxygen loss • rhizosphere • rice • root permeability
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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; Armstrong, Armstrong, and Van der Putten, 1996
; Armstrong et al., 1996b
; Armstrong and Armstrong, 1999
). These include stunting and death of roots and shoots, failure of laterals to emerge, bud death, premature senescence of shoots, blockages within the gas space and vascular systems, lower levels of starch in rhizomes and cell wall lignification and suberization of surface cell layers of laterals and apical regions of adventitious roots. The lower the pH of the rooting medium, the greater is the concentration of the lipid-soluble, toxic, undissociated organic acid molecules (Hollis, 1967
; Jackson and Taylor, 1970
; Rao and Mikkelsen, 1977
; Lynch, 1978, 1982
) and the more severe are the symptoms (Armstrong and Armstrong, 1999
) Commonly, mixtures of many organic acids can occur in the field, including die-back sites. They are produced from the decaying underground parts of the plant (Kovacs et al., 1989
; Armstrong and Armstrong, 1999
) and may be generated from other sources of decaying organic matter. For example, at a eutrophic site in the Czech Republic, bud death and reed decline have been associated with a strongly reducing organic layer in the surface of the lake sediment (Cízková, Strand, and Lukavská, 1996
) and with mixtures of organic acids in interstitial waters (Cízková et al., 1999
). So far, however, there has been no direct evidence that cocktails of dilute organic acids, at a comparatively high pH of 6 (as can occur in the field) and such that the concentration of each acid is innocuous, can be harmful to the reed. In this study, the concentration of each acid in a cocktail was 1 mmol/L at pH 6, with the undissociated concentration being 
0.06 mmol/L. It has previously been shown that monocarboxylic acids individually at this concentration are not toxic to Phragmites (Armstrong and Armstrong, 1999
).
Various diseases of rice have also been associated with toxicity due to sulphide and organic acids. For example, in Akiochi, or "autumn decline," symptoms include early flowering, discoloration of roots, and progressive decline of the plant. The lower organic acids, including acetic, propionic, and butryic have been shown variously to reduce the uptake of P, K, Si, Mn, Mg, Ca, and NH4-N (Mitsui et al., 1954
; Takijima, Shiojima, and Arita, 1960
; Tanaka and Navasero, 1967
; Rao and Mikkelson, 1977
) and to reduce root respiration (Tanaka and Navasero, 1967
). Rao and Mikkelson found that individually these acids at 10 mmol/L and pH4 caused the death of seedlings within 1–2 d; at 1 mmol/L (pH 4.6) the seedlings showed signs of desiccation. Increased organic acid toxicity at lower pH has also been shown to apply to rice seedlings (Tanaka and Navasero, 1967
; Rao and Mikkelsen, 1977
). So far, however, there appears to have been little documentation of any anatomical effects of organic acids on rice.
In wetland plants, the passage of atmospheric oxygen via the internal gas space system of the plant to the underground organs is vital to maintain aerobic metabolism, while the radial diffusion of oxygen out of the permeable parts of the root system into the rhizosphere protects those vulnerable parts by promoting oxidations of potential phytotoxins, e.g., FeII, MnII, sulphide, and organic acids commonly existing in flooded soils (Armstrong, 1970
; Trolldenier, 1988
; Conlin and Crowder, 1989
; Armstrong, Armstrong, and Beckett, 1992
; Begg et al., 1994
; Saleque and Kirk, 1995
; Wang and Peverley, 1999
). Currently, there is still much interest in the efflux of oxygen from the roots of Phragmites, Typha, and other aquatic macrophytes, which aids water purification in both natural and artificial reed beds by inducing nitrification in the rhizosphere (Reddy, Patrick, and Lindau, 1989
). Furthermore, radial oxygen loss (ROL) from root to rhizosphere inhibits methanogenesis (Oremland, 1988
; Conrad, 1989
), promotes CH4 oxidations within the rhizospheres (Epp and Chanton, 1993
; Gilbert and Frenzel, 1995
), and can thereby reduce potential efflux of CH4 from the plants by 34% for Phragmites (Grunfeld and Brix, 1999
) and by >80–90% for rice (Holzapfel-Pschorn, Conrad, and Seiler, 1986
; Frenzel, Rothfuss, and Conrad, 1992
). Natural and cultivated wetlands contribute 40–50% of total emissions of CH4 to the atmosphere, accounting for 7–9% of global warming. In vegetated wetlands >90% of CH4 emitted from rice paddies (Banker et al., 1995
) and 62% from Phragmites-dominated habitats (Grunfeld and Brix, 1999
) can pass into the atmosphere via the root 
internal gas space shoot pathway; in certain species this is augmented during the growing season by convection (Chanton et al., 1993
; Sorrell and Boon, 1994
; Whiting and Chanton, 1996
; Grunfeld and Brix, 1999
). Thus, factors affecting root permeability, ROL, and the porosity of the gas-space system of the plant are clearly important. So far, there appears to have been no published work investigating the possible effects of phytotoxins on root permeability to oxygen and its radial loss to the rhizosphere.
This study seeks to test three hypotheses: (1) that cocktails of the lower monocarboxylic organic acids, where the concentration of each acid is innocuous, may nevertheless be harmful to Phragmites, (2) that rice responds to organic acid toxicity in ways similar to Phragmites by inducing cell wall thickening in the hypodermal layers of adventitious root apices and in the epidermis of laterals, and blockages in the internal gas space and vascular systems, and (3) that in rice and Phragmites, organic acids, by inducing cell wall thickening in normally permeable regions of the root system, thereby also induce impermeability to oxygen and reduce ROL to the rhizospheres.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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100 mm), which had been raised from seed collected from plants along the Humber estuary, were used as the source of plant material.
Rice
Seeds of cv. Norin 36 were germinated in the summer on moist tissue in shallow trays covered in polythene in a propagating frame with natural light; T = 20°C. Germination occurred in 1 wk and the trays were then transferred to the open bench in the glass house for 3–4 wk until the shoots were 6–8 cm high. The seedlings were then transplanted into buckets (volume = 15 L) containing moist John Innes compost No. 2 (John Innes Institute, Norwich, UK); the water table was gradually raised over 2 wk, until the soil was flooded to a depth of 2 cm. Conditions were: T = 16°C (minimum) and 29°C (maximum), with natural light, and an 18-h day length.
Treatments
Phragmites
The old roots were trimmed back to 1–2 cm and old, dead stems shortened to 100–130 mm and the plants were transferred to black polythene-covered glass tubes (height = 400 mm; diameter = 50 mm) containing 25% Hoagland's solution to resume root and shoot growth under growth room conditions; the shoots received continuous lighting from the side; T = 18°C; photosynthetically active radiation (PAR) = 80–100 µmol·m–2·s–1. The plants were secured in the tubes with the old culms emergent and the rhizomes submerged under 40 mm of culture solution; they were arranged randomly in line, parallel to a bank of lights, and their positions changed every alternate day to ensure that, as far as possible, they experienced identical external conditions.
After 12–14 d, when the roots were 1–30 mm and shoots 100–150 mm long, plants of similar size were selected and divided into three groups of eight plants each; their rhizomes and roots were immersed, as described above, in the following culture solutions, each having as the base 25% Hoagland's solution in stagnant agar (0.05% w/v) degassed by autoclaving: (1) control, containing no acids; (2) cocktail 1, containing a mixture of five lower volatile monocarboxylic organic acids (acetic, propionic, n-butyric, iso-butyric, and caproic) each at 1 mmol/L concentration (total undissociated concentration = 0.35 mmol/L); (3) cocktail 2, containing seven acids (formic, acetic, propionic, n-butyric, iso-butyric, caproic, and valeric) each at 1 mmol/L concentration (total undissociated concentration = 0.42 mmol/L). The pH of each culture medium was 6, adjusted by the addition of NaOH solution and using a portable pH meter (Camlab Ltd., Cambridge, UK). Half of the plants from each treatment received only one dose of the acid medium, which was left unchanged between days 1 and 5 for adventitious root ROL and anatomical assays. For the remaining plants, the media were renewed every alternate day; these plants were used for monitoring growth and senescence.
For measuring ROL from fine laterals, an additional group of nine plants was grown in 25% Hoagland's solution until adventitious roots were 180–290 mm long and the longest laterals 15–18 mm. (It was not possible to use plants from the main treatments as, here, the laterals from the cocktails were much shorter than those of the controls.) The cocktails were added during the period of ROL measurement (see relevant section).
Rice
When the shoots were 30 cm high, individual tillers were removed, the roots cut back to 1–2 cm, washed, and each was transferred into a glass tube with the shoot base and root system submerged, as described for Phragmites, and kept in the growth room (T = 22°C; PAR = 80–100 µmol·m–2·s–1); here the nutrient medium was 25% Yoshida solution (Yoshida et al., 1976
), renewed on alternate days.
When the longest new roots were 100 mm long, the plants were divided into two groups of six, for separate treatments, the base of each rooting medium being 25% Yoshida solution (Yoshida et al., 1976
) in 0.05% w/v agar, degassed by autoclaving: (1) controls, at pH 4.5, without the addition of acetic acid; (2) 1.5 mmol/L acetic acid at pH 4.5 undissociated concentration = 1.05 mmol/L. For half the plants from each treatment the media were left unchanged between days 1 and 5 for root ROL and initial anatomical assays. The media of the remaining plants were changed every alternate day until day 14 for final anatomical examinations.
Growth
For Phragmites, shoot growth was recorded on at least nine shoots per treatment on each of the 4 d preceding treatments and during the first 4 d of the treatment period; adventitious root growth increments were measured until day 37 by marking the positions of the tips of selected roots (at least 17 per treatment) on the glass container and measuring the distances between marks using digital callipers (RS Components Ltd., Corby, Northamptonshire, UK). The growth of new adventitious roots and of the thick laterals from the cocktail 1 treatment was also monitored as were the onset of shoot senescence and the percentage of shoot senescence, viz. (number of senescing leaves/total number of leaves) x 100, the latter being measured on day 22.
Anatomy
On alternate treatment days 1–6, for both species, transverse sections were made of selected adventitious roots and laterals from plants used in the ROL measurements. Further studies were made on Phragmites after 10 d, and on rice plants after 14 d of continuous exposure to acetic acid and on the controls, including transverse sections of rhizome, root-rhizome junctions, and adventitious roots. Phragmites sections and entire laterals from both species were stained with phloroglucinol and concentrated hydrochloric acid to detect lignification (confirmed with aniline hydrochloride) and were occasionally stained with Sudan 111 to detect suberization. Sections of rice adventitious roots and rhizomes were left unstained as there was comparatively little lignification. Specimens were photographed using an Olympus BX40 photomicroscope (Olympus Optical Co. Ltd., Tokyo, Japan).
Radial oxygen loss from roots
Phragmites
Radial oxygen loss along adventitious roots was measured using sleeving cylindrical Pt cathodes in conjunction with Ag-AgCl anodes, after the method of Armstrong and Wright (1975)
. Radial oxygen loss profiles were taken from 3 mm subapically to the points at which laterals emerged and were measured on at least two roots from each plant per treatment, between days 3 and 5 after the start of the treatment period. This time was chosen because apical root wall lignification and suberization in the cocktail treatments were detectable at this stage (see RESULTS: ANATOMY). For ROL measurements, the control or treatment media were replaced with freshly deoxygenated 0.05% w/v aqueous agar containing 1/4-strength Hoagland's solution.
The ROL from lateral roots was detected by means of bare Pt wire cathodes (length = 50 mm; diameter = 0.37 mm), used in conjunction with Ag-AgCl anodes. The cathode was loosely coiled around the adventitious root in the region where the laterals were 10–15 mm long. The ROL was measured with the roots in freshly deoxygenated control-type medium for at least 8 h; this medium was then drained from the base of the tube so as not to disturb the electrode and then either cocktail 1 or 2 was added and ROL measurements were resumed. Electrodes were also inserted in the media in positions remote from the roots to measure background O2 diffusion rates.
It was necessary to confirm, if possible, that reduced ROL from adventitious roots and laterals was a function of root wall impermeability and not related to an absence of oxygen within the adventitious roots. After ROL measurements were completed on Phragmites from the toxin treatments, apical 30-mm regions of adventitious roots were removed, with the plants still intact, and the sleeving Pt electrode moved around the cut end, to detect O2 diffusing from the cut end. Also, in both species, attempts were made to blow air, via a stem base, through the rhizome and out of the cut ends of roots submerged under water.
Rice
The ROL from the apical and subapical regions of adventitious roots was measured using the same method as that described above for Phragmites. Measurements were taken 3–4 d after the roots had received a single dose of 1.5 mmol/L acetic acid at pH 4.5; for the control roots the medium was also at pH 4.5.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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5 mm long. However, cocktail 1 was not sufficiently toxic to prevent the growth of new adventitious roots, which appeared in two flushes; also thick laterals of indefinite length and resembling adventitious roots were produced on adventitious roots that had stopped growing. We have previously noticed this effect in response to butyric acid (Armstrong and Armstrong, 1999
); here this may well have been in response to n- and iso butyric acids in cocktail 1.
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Rice
Growth of roots and shoots was not measured in this study; as mentioned in the introduction, it has been documented that organic acids, including acetic, can inhibit the growth of rice. In this study, also, it was noted that in plants exposed to acetic acid for 2 wk, adventitious and lateral root growth was much inhibited; there was also death of adventitious root apices and death of laterals that had emerged prior to treatment and premature shoot senescence.
Anatomy
Cell wall lignification, as detected using phloroglucinol and hydrochloric acid was invariably confirmed with aniline hydrochloride and, in all the cases tested, was associated with some degree of suberization.
Phragmites
In the controls no lignification or suberization was detected throughout the experimental period in the apical 20 mm of adventitious roots or in the epidermis of laterals. However, after only 24–30 h in cocktail 2 and 2–3 d in cocktail 1, there were some signs of lignification and suberization of the walls of the surface cell layers of the apical 10 mm of adventitious roots, including those of the root cap and of the epidermis of fine laterals. The effects became more pronounced as the experiment proceeded, especially with the cocktail 2 treatment (Fig. 2A–D, J–L). Although in the first 3–4 d the cortex and stele were apparently relatively unaffected, thereafter, particularly with cocktail 2, some of the cortical intercellular gas spaces and protoxylem and phloem became occluded, cortical cells became lignified, and there was premature lignification of the hypodermal layers and stele (Fig. 2E–G).
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; Votrubova and Pechácková, 1997). These same effects have been described for Phragmites in relation to responses to phytotoxins (Armstrong, Armstrong, and Van der Putten, 1996
; Armstrong, Afreen-Zobayed, and Armstrong, 1996
; Armstrong and Armstrong, 1999
). Neither the adventitious nor lateral roots of the controls showed any of these effects, even after 4 wk.
Radial oxygen loss from roots
Adventitious root apices
The profiles of ROL from the apical regions varied for roots of differing lengths and according to their positions on the rhizome. Therefore, although at least six roots per treatment were tested, it was only possible to group together roots from the various treatments where these parameters were similar. However, within a treatment, the ROL profile patterns were the same. Measurements further up the roots were prevented by the emergence of laterals. For both Phragmites and rice, when a pressurized air flow was applied to shoot bases after ROL measurements had been completed, air freely bubbled from submerged roots whose apices had been excised, thus indicating a free diffusive pathway from shoot to root at this stage.
Phragmites
After 3–5 d of treatment, ROL values for control adventitious roots for the apical 2–30 mm were high, 63–84 ng O2·cm–2·min–1; but declined to zero at 60 mm behind the apex (Fig. 4, Table 1). For the cocktail 1 treatment the apical ROL values were variable, but the highest were 30–78 ng O2·cm–2·min–1 for the apical 4 mm; thereafter, they fell sharply to zero at 30 mm behind the apex. For some roots from this treatment, apical values did not exceed 6 ng O2·cm–2·min–1. However, roots from the cocktail 2 treatment were even more affected; apical ROL values were consistently very low: 0–12 ng O2·cm–2·min–1, and reached zero, at 6 mm from the apex. In all treatments oxygen freely diffused from cut apical regions, indicating that there was adequate internal oxygen transport down the roots.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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). The stunting of adventitious and lateral roots, slowing down of shoot growth and premature cell wall thickening in surface layers of roots were apparent within the first 4 d of treatment with both cocktails, but more obvious with the more toxic cocktail 2, which contained two more acids than cocktail 1. Also, the onset of premature shoot senescence occurred 2 wk earlier with cocktail 2.
The toxicity of an organic acid cocktail will be directly related to the number and concentrations of acids present in the rooting medium and inversely related to pH. In the present study, it can be predicted that at a slightly lower pH of 5.9, rather than 6, cocktail 1 would have been as toxic as cocktail 2. The pH of the rhizosphere must also influence the toxicity of organic acids. It has already been demonstrated that the pH of the rhizospheres of adventitious roots and laterals can be lower than that of the bulk soil, sometimes by more than two pH units, e.g., in rice (Begg et al., 1994
; Saleque and Kirk, 1995
), in Phragmites (Conlin and Crowder, 1989
), and in Cyperus involucratus, Eleocharis sphacelata, and Juncus ingens (Sorrell and Orr, 1993
). Similarly we have found (Armstrong and Armstrong, 1999
) that the rhizospheres around the apices of young adventitious roots of Phragmites can be at pH 6.6–6.7, while the rooting medium is at pH 7.7.
It is interesting that rice responded anatomically to acetic acid as does Phragmites to the lower organic acids in general, and to sulphide (Armstrong, Afreen-Zobayed, and Armstrong, 1996
; Armstrong, Armstrong, and Van der Putten, 1996
; Armstrong and Armstrong, 1999
), and in the present study to cocktails of dilute organic acids. These responses included premature cell wall thickening and some lignification of the normally permeable regions of the root system, occlusion within the vascular systems and the intercellular gas spaces of adventitious roots, and proliferations of callus that partly blocked the gas spaces within root–rhizome junctions and rhizomes. In rice, however, unlike Phragmites, there was comparatively little premature lignification, except in the epidermis of the laterals. The intercellular cortical spaces and vascular elements became occluded with yellowish-brown substances that darkened with time. Soukop et al. (Charles University, Prague, unpublished data) have found that this type of occlusion in Phragmites contains polysaccharide gums. Acetic acid has been shown to cause leakiness of cell membranes (Van Overbeek and Blondeau, 1954
; Jackson and Taylor, 1970
) and loss of ions from roots (Lee, 1976
); we suggest, therefore, that such occlusions in both rice and Phragmites are due in part to phytotoxin damage to cell membranes. We interpret all of these anatomical symptoms as defense responses (Friend, 1981
; Asada and Matsumoto, 1987
) to prevent further ingress and spread of the toxins within the plants and possibly fungal infection. In Phragmites a predisposition to fungal infection has been associated with phytotoxin damage (Armstrong, Afreen-Zobayed, and Armstrong, 1996
; Armstrong and Armstrong, 1999
). Insofar as we are aware, this is the first documentation of anatomical effects of acetic acid in rice. Since Akiochi disease of rice has also been linked to sulphide toxicity (e.g., Park and Tanaka, 1968
), it would be interesting to know whether these same anatomical symptoms can be induced in rice by sulphide as they are in Phragmites (Armstrong, Afreen-Zobayed, and Armstrong, 1996
).
For both Phragmites and rice, reductions of ROL from adventitious root apices and fine laterals were apparent at early stages of treatment. Since the internal aeration pathways allowed pressurized gas flow from the shoots to the apices of adventitious roots, we associate the reduced ROL with lowered permeabilities to oxygen induced by atypical cell wall lignification and suberization in the epidermis of the laterals and of the hypodermal layers of adventitious root apices; in some cases these were sufficient to reduce ROL to zero. At later stages, blockages that develop extensively within the gas spaces of adventitious roots, root–rhizome junctions, and rhizomes will impede longitudinal and lateral transport of oxygen to the laterals and to and within the adventitious root apex and contribute to reduced ROL to the rhizospheres and reduced respiratory oxygen supply. Reduced permeability and ROL from lateral root zones are likely to be particularly serious, since it is here that the oxidized rhizospheres are most extensive (Armstrong and Armstrong, 1988
) and the surface areas for absorption of water and nutrients are very large. One could therefore predict that reduced permeability to oxygen will be accompanied by reduced uptake of water and nutrients. This is supported by A. Soukopp (Charles University, Prague, unpublished data), who found that root wall lignification reduced permeability to water and iodate ions in Phragmites.
In rice, reductions in root permeability and ROL and impedances in the gas transport pathways produced by the acetic acid treatment could help to explain some of the symptoms of organic-acid-induced Akiochi, including reduced root respiration and nutrient uptake, and the decreased ability of the roots to oxidize iron, reported by Takijima (1965)
. The vascular blockages reported in the present study could also help to explain the reduced uptake of nutrients in Akiochi mentioned earlier.
It should be noted, however, that in wetland plants, including Phragmites and rice, the lignification of hypodermal layers in maturing subapical parts of adventitious roots, accompanied by reduced ROL to the rhizosphere, is a well-known, natural phenomenon, coinciding with the development of aerenchyma and laterals (Luxmoore, Stolzy, and Letey, 1970
; W. Armstrong, 1971; J. Armstrong, 1992; Colmer et al., 1998
; Armstrong et al., 2000). The latter are freely permeable to ROL and divert oxygen from the gas spaces within the main root. Thus, the normal development of impermeable layers in the basal and subapical regions of the root wall must be useful and help to conserve oxygen in the main root for supplying the laterals and the apical parts of the adventitious root. However, epidermal lignification of laterals and of the apical hypodermal layers of adventitious roots are atypical, since these are normally absorptive regions for salt and water uptake. Also, being permeable, they are vulnerable to attack from soil-borne phytotoxins, and so it is important that they are protected by oxidizing rhizospheres, as mentioned in the introduction.
Critically high accumulations of phytotoxins, arising from perhaps excessive fertilization or other means of eutrophication, can probably set in motion a complex series of reactions, and we present a scheme (Fig. 7) showing some of the possible interactions. We suggest that (a) the toxins will penetrate the rhizospheres, where the O2 will be rapidly consumed during biological and chemical oxidation of reduced toxins such as Fe2+, Mn2+, and S2– and in the microbial oxidation of organic acids; (b) if the metabolic breakdown of the phytotoxins in the rhizosphere is incomplete, the toxins will penetrate the vulnerable laterals and adventitious root apices, which will react by becoming less permeable, thus resulting in a diminution of ROL; (c) additionally, there may be a temporary proliferation of aerobic organisms in the rhizosphere stimulated by the presence of the phytotoxin substrates; (d) effects (a)–(c) will cause the shrinkage of the rhizospheres, thus bringing the source concentrations of phytotoxins closer to the roots, which may become overwhelmed. Reduced ROL from the roots will also result from blockages in the gas spaces of roots and rhizomes for diffusive transport, and in some species, e.g., Phragmites, of rhizomes for convective transport of oxygen. The shrunken rhizospheres will be ineffective in protecting the roots, resulting in an overall stunting of the root systems, a factor that will also contribute to a general reduction in rhizosphere oxidation. In extreme cases of dieback, root, rhizome, and bud death occur and finally localized death and decay of the plants, leading to further release of phytotoxins.
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) and methanotrophs (Gilbert and Frenzel, 1995
), and increases in anaerobic microbes, including methanogenic bacteria. Rates of nitrification will decrease, whereas methanogenesis will increase, especially if acetate is among the phytotoxins present (Rothfuss and Conrad, 1993
). Additionally reduced root permeability and biomass, shoot senescence, and blockages in the xylem will tend to decrease water uptake by the plants, and this will promote higher water tables and thereby increase rates of denitrification and methanogenesis. In relation to the latter, Sorrell et al. (1997)
found that rates of methanogenesis were far higher at dieback sites than at healthy reed sites. Moreover, dieback sites tended to remain waterlogged and methanogenic during dry periods, whereas healthy sites became dry and inactive. They conclude that "reed wetlands may not be great generators of CH4 while the plants are healthy and actively growing, but become more intense CH4 producers when the plants senesce and die" (p. 1984). If decreased root permeability to oxygen is accompanied by diminished permeability to CH4, as seems likely, then, despite increased methanogenesis, less CH4 may pass along the root rhizome shoot pathway into the atmosphere, due to decreases in root permeability and biomass and reduced diffusive gas transport and, in some cases, e.g., Phragmites, convective flow through the plant. Here, phytotoxin damage has been correlated with reduced convective flow at reed dieback sites, due to blockages in the gas space system and premature shoot senescence (Armstrong et al., 1996a
). The possible effects of phytotoxins on CO2 production and emission from wetlands have not been discussed here, but they may be similar to those effects on CH4.
Although high phytotoxin levels will probably tend to reduce potential methanogenesis because of diminished root exudation (Whiting and Chanton, 1993
; Minoda and Kimura, 1994
) correlated with blockages in the phloem and decreased root permeability and biomass, it seems likely that the overall effect of phytotoxins will be to increase rates of methanogenesis. However, since aquatic macrophytes are important emitters of CH4, the question of the effects of phytotoxins on the overall long-term emissions of CH4 from wetlands would appear to merit further investigation.
It would obviously be of great interest to know, from further practical investigations and modelling, the extent to which the reactions indicated in Fig. 7 and other related effects might take place. They are clearly relevant to the health and survival of reed and of other emergent aquatic macrophytes such as rice in localities that are prone to accumulations of phytotoxins, to the role of reed in the phytopurification of waste waters by artificial wetlands, and to the emissions of CH4 and other greenhouse gases via such plants from wetlands.
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FOOTNOTES |
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2 Author for correspondence (w.armstrong{at}biosci.hull.ac.uk
; FAX:
01482 465458).
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LITERATURE CITED |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONLITERATURE CITED |
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———, F. Afreen-Zobayed W. Armstrong 1996 Phragmites die-back: sulphide- and acetic acid-induced bud and root death, lignifications, and blockages within the aeration and vascular systems. New Phytologist 134: 601-614[CrossRef][ISI]
———, andW. Armstrong 1988 Phragmites australis—a preliminary study of soil-oxidising sites and gas transport pathways. New Phytologist 108: 373-382[CrossRef][ISI]
———, and ———. 1999 Phragmites die-back: toxic effects of propionic, butyric and caproic acids in relation to pH. New Phytologist 142: 201-218[CrossRef][ISI]
———, ———, I. B. Armstrong G. R. Pittaway 1996a Senescence, and phytotoxin, insect, fungal and mechanical damage: factors reducing convective gas-flows in Phragmites australis. Aquatic Botany 54: 211-216[CrossRef]
———, ———, andP. M. Beckett 1992 Phragmites australis: Venturi- and humidity-induced convections enhance rhizome aeration and rhizosphere oxidation. New Phytologist 120: 197-207[CrossRef][ISI]
———, ———, andW. H. Van der Putten 1996 Phragmites die-back: bud and root death, blockages within the aeration and vascular systems and the possible role of phytotoxins. New Phytologist 133: 399-414[CrossRef][ISI]
———, ———, Z. Wu F. Afreen-Zobayed 1996b A role for phytotoxins in the Phragmites die-back syndrome?. Folia Geobotanica et Phytotaxonomica 31: 127-142
Armstrong W. 1970 Rhizosphere oxidation in rice and other species: a mathematical model based on the oxygen flux component. Physiologia Plantarum 23: 623-630[CrossRef]
———. 1971 Radial oxygen losses from intact rice roots as affected by distance from the apex, respiration and waterlogging. Physiologia Plantarum 25: 192-197[CrossRef]
———, D. Cousins J. Armstrong D. W. Turner P. M. Beckett 2000 Oxygen distribution in wetland plant roots and permeability barriers to gas-exchange with the rhizosphere: a microelectrode and modelling study with Phragmites australis. Annals of Botany 86: 687-703
———, andE. Wright 1975 The theoretical basis for the manipulation of flux data obtained by the cylindrical platinum electrode technique. Physiologia Plantarum 35: 21-26[CrossRef]
Asada Y. J. Matsumoto 1987 Induction of disease resistance in plants by a lignification-inducing factor. In S. Nishimura, C. P. Vance, and N. Doke [eds.], Molecular determinations of plant diseases, 223–231. Science Society of Japan Press, Tokyo, Japan
Banker B. C. H. K. Kludze D. P. Alford R. D. Delaune C. W. Lindau 1995 Methane sources and sinks in paddy rice soils: relationship to emissions. Agriculture, Ecosystems and Environment 53: 243-251[CrossRef]
Begg C. B. M. G. J. D. Kirk A. F. Mackenzie H.-U. Neue 1994 Root-induced iron oxidation and pH changes in the lowland rice rhizosphere. New Phytologist 128: 469-477[CrossRef][ISI]
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) received no organic acids, N = 7, bars = ±1 SE. Cocktail 1 treatment (
, 
, and 
) and cocktail 2 treatment (
= acetic acid treatment (N = 3); bars = ±1 SE
