Rhizome phyllosphere oxygenation in Phragmites and other species in relation to redox potential, convective gas flow, submergence and aeration pathways

Authors


Author for correspondence: J. Armstrong Tel: +44 1482 465527 Fax: +44 1482 465458 Email: j.armstrong@hull.ac.uk

Summary

  • • Underground rhizomes of emergent aquatic macrophytes are important for perennation, vegetative spread, competition and anchorage. In four species we examined the potential for the development of oxidized phyllospheres around rhizome apical buds, similar to the protective oxygenated rhizospheres around roots.
  • • Redox potentials and polarographic measurements of radial oxygen loss were recorded using platinum cathodes around the apical buds. The aeration pathway from atmosphere to phyllosphere was investigated anatomically and by applied pressurized gas flow.
  • • Redox potentials increased by +400, +45, +200 and +340 mV around rhizome apices of Phragmites australis, Oryza rhizomatis, Carex rostrata and Glyceria maxima, respectively. Radial oxygen loss from rhizome apices of Phragmites was increased by convective gas flow through the rhizome and by shoot de-submergence, and decreased by resistances applied within the aeration pathway and by shoot submergence.
  • • We conclude that oxygen passes via internal gas-space connections between aerial shoot, rhizome and underground buds and into the phyllosphere regions via scale-leaf stomata and surfaces on the buds. We suggest that oxidized phyllospheres may protect rhizome apices against phytotoxins in waterlogged soils, just as oxidized rhizospheres protect roots.

Introduction

The release of oxygen to the rhizosphere from laterals and the subapical regions of adventitious roots of wetland plants has been the subject of many studies. Much is known of the distribution and quantification of oxygen flux along root systems (Armstrong, 1979; Sorrell et al., 2000; Colmer, 2003) and the factors promoting them, for example, gas space formation (Jackson & Armstrong, 1999; Evans, 2004), root permeability (Armstrong et al., 2000; De Simone et al., 2003) and convective throughflows (Armstrong & Armstrong, 1990; Armstrong et al., 1992). Others have described the promotion of oxidations within the rhizosphere (Begg et al., 1994; Kirk & Bajita, 1995) and the protection against phytotoxins that such oxidations confer on the vulnerable permeable regions of the root system (Mendelssohn & Postek, 1982; Lee et al., 1999; Pedersen et al., 2004). Recent papers have also shown that factors such as anoxia (Colmer et al., 1998; Colmer, 2002) and phytotoxins (Armstrong & Armstrong, 2001a, 2005) can induce impermeability to oxygen in normally permeable regions, while Soukup et al. (2002), De Simone et al. (2003) and Enstone & Peterson (2005) have reported on the physical and chemical nature of such impermeability. Armstrong et al. (1999) showed decreased radial oxygen loss (ROL) to the rhizosphere in Phragmites by experimentally submerging the shoot system. Also, ROL can fluctuate diurnally in submerged plants because of daytime photosynthesis (Waters et al., 1989; Pedersen et al., 2006).

In many wetland species, for example Phragmites australis, Typha angustifolia, Carex rostrata, Glyceria maxima, Eriophorum angustifolium, water lilies (e.g. Nuphar and Nelumbo) and perennial, rhizomatous rice Oryza rhizomatis, the underground system is composed of not only extensive roots but also rhizomes buried in permanently flooded mud. Some rhizomes, as in Phragmites, will grow to depths of > 1 m. While roots are essential for anchorage and absorption of water and mineral nutrients, rhizomes are also important for anchorage, act as perennating organs (Dykjová & Hradecka, 1976), and enable the plant to spread vegetatively (Haslam, 1969; Granéli et al., 1992) and compete with other species (Hellings & Gallagher, 1992). Rhizomes bear plentiful buds and root primordia, are organs of food storage, and can survive when roots and aerial shoots have died. The rhizomes of many species, such as Schoenoplectus lacustris, Typha latifolia and P. australis have been shown to withstand considerable periods of anoxia (Crawford & Brandle, 1987) and some, such as those of the former two species, will even exhibit bud extension under anoxia. The rhizomes of these and other species, particularly Acorus calamus, also have metabolic pathways for internally detoxifying phytotoxins such as sulphide and excess ammonia absorbed from the waterlogged soils (Brändle, 1991; Fürtig et al., 1996; Weber & Brändle, 1996).

Although it is well known that the rhizomes of wetland plants are highly porous, to date, comparatively little has been documented on the surface permeability of rhizomes to oxygen. Armstrong & Boatman (1967) noted zones of iron oxidation around rhizome apices of E. angustifolium in anoxic soils. Also, more recently, it was shown for Phragmites that reduced methylene blue dye becomes oxidized around the rhizome apices, but not around the mature subapical parts, and that gas will bubble from the apex when an excised rhizome end is exposed to a pressurized gas stream (Armstrong & Armstrong, 1988). The mature parts of the Phragmites rhizome appear to be impermeable to oxygen. There is no oxidation of methylene blue dye around these regions, which have epidermal and subepidermal cell layers with lignified and suberized cell walls, while the epidermis itself is covered by a waxy cuticle (Armstrong & Armstrong, 1988).

Death of underground and emergent vertical rhizomes can be a common feature of dieback of Phragmites and associated with high levels of phytotoxins such as sulphide and the lower organic acids in the sediments (Kovacs et al., 1989; Armstrong et al., 1996d; Cíkováet al., 1999; Armstrong & Armstrong, 2001b). The rhizome apices of the reed appear to be quite vulnerable to attack from such phytotoxins (Armstrong et al., 1996a; Armstrong & Armstrong, 1999) possibly because of their permeable nature. Fürtig et al. (1996) also showed that Phragmites rhizomes can be damaged by sulphide. Therefore it seems reasonable to suggest that perhaps, like roots, rhizome apices may normally be protected from low levels of such phytotoxins by the presence of oxidized phyllospheres.

The aim of this paper is to elucidate the permeable nature of rhizome apices by testing the following hypotheses on a range of morphologically contrasting emergent macrophytes: P. australis, the perennial rice O. rhizomatis, C. rostrata and G. maxima.

  • 1Rhizome apices are permeable to oxygen and induce oxidized phyllosphere regions in a waterlogged soil (similarly to oxidized rhizospheres around fine laterals and the apices of adventitious roots).
  • 2Atmospheric oxygen passes via the plant's interconnecting gas-space system from shoot to rhizome and diffuses from rhizome apices via stomata on the scale leaves covering the meristem.
  • 3Oxygen diffusion from mature parts of the rhizome is prevented or curtailed because of hypodermal suberization, epidermal cuticularization, an absence of stomata, and aerenchyma channels of withered scale leaves sealed by callus.
  • 4Radial oxygen loss from rhizome apex to phyllosphere can be (i) reduced by physical resistance to gas flow or callus in the aeration system, (ii) increased by convective air flow within the rhizome, (iii) reduced by shoot submergence, and (iv) affected by the gas regime around the aerial shoot.

Materials and Methods

Plant material and soil/nutrient media

Phragmites australis  Horizontal rhizomes of Phragmites australis Cav. ex Steud. (approx. 200–300 mm length, 10–15 mm outer diameter) collected from the banks of the Humber estuary, UK in October 1997, were either used immediately or stored for a few days in polythene bags in a cold room at 5°C. A soil slurry, rich in organic matter, was collected from the floor of a shallow freshwater lake and kept in 12-l buckets with a surface layer of water 50–100 mm deep for 3 d in the laboratory at 18–23°C before being used to cover the rhizomes; the buckets were covered in black polythene to exclude light and prevent algal growth.

Additionally, 4–6-month-old Phragmites plants were used, previously raised from seed and grown in a waterlogged loam in 0.25-l pots. These plants each had four to five shoots (approx. 200–400 mm shoot height; 100 mm total rhizome length) and were used intact for measurements on both vertical and horizontal rhizomes. The plantlets were secured with culms emergent and the rhizomes submerged in quarter-strength stagnant Hoagland's solution in glass tubes (400 mm height, 50 mm diameter). The tubes were sheathed in black polythene, and black polystyrene beads were poured on to the liquid surface to a depth of approx. 20 mm to exclude the light. The plants were kept for up to 1 wk in a growth room at 18°C and 35–55% RH, with constant cool fluorescent lighting from the side to give an even distribution over the shoot system of 80–100 µmol m−2 s−1 PAR. At the beginning of the experiments, the lengths of horizontal and vertical rhizomes were approx. 20–60 mm and 25–45 mm, respectively.

Perennial rice, Carex and Glyceria  Plants used were perennial rice [Oryza rhizomatis (URU WEE) Vaughan's], Carex rostrata Stokes and Glyceria maxima Holmberg. The rice was raised from seed and grown in 12-l buckets during the summer in a glasshouse (16–29°C); cuttings of Carex and Glyceria were taken from plants in tanks of waterlogged loam. These cuttings and individual rice tillers each had a shoot, rhizome, underground buds and roots. Original shoot heights were approx. 200–350 mm. After washing the underground parts, the dead roots were trimmed away. Plants were transferred to glass tubes as for Phragmites, then grown on in stagnant Hoagland's solution (quarter-strength for rice; full-strength for Carex and Glyceria) for 2–3 wk to promote rhizome production from the growth of buds. The lengths of horizontal rhizomes for rice, Carex and Glyceria were then approx. 10–15, 40–50 and 25 mm, and of the vertical rhizomes approx. 20, 30–60 and 25–80 mm, respectively. (In rice the buds grew horizontally for only a few millimetres before turning upwards to form vertical rhizomes.)

Redox potential in phyllospheres (around rhizome apices) in waterlogged soil

Phragmites excised rhizomes  Each horizontal rhizome (120–200 mm length) was freshly cut, and platinum wire electrodes (50 mm wire length, 0.37 mm diameter) were coiled spirally around the rhizome apex (see Fig. 7a). In some cases, additionally, the first coil of a subapical electrode was positioned at approx. 15–20 mm behind the rhizome apex. Care was taken to ensure the electrodes did not damage the rhizome tips, and that the connecting wires to the electrometer were slack so that the electrodes could move as the rhizomes grew in length. The rhizomes were placed inside a glass tank (600 mm length, 120 mm width, 200 mm height) each with its cut end connected by rubber tubing to a plastic tube that passed through the end wall of the tank. Outside the tank the plastic tubes were each attached to a T-piece so that conditions could be varied at the cut end of the rhizome. Each rhizome was tied with string to a glass support to anchor it on the floor of the tank, which contained two rhizomes. The rhizomes were then covered in soil slurry to a depth of approx. 80 mm; the depth of water above the sediment was approx. 50 mm. Similar platinum-wire electrodes as controls were inserted at a depth of 70–80 mm in the sediment, in positions remote from the rhizomes. Each platinum electrode was used in conjunction with a silver–silver chloride reference electrode (Eh + 199 mV) the tip of which was positioned beneath the soil surface. Redox potentials in the phyllospheres around rhizome tips and in the bulk soil were measured using a high-impedance electrometer. Once the rhizomes and electrodes had been positioned, the top and walls of the tank were covered in black polythene to exclude light, preventing algal growth and possible photosynthesis of the rhizomes.

Figure 7.

Spiral oxygen electrode sensor and anatomical features. (a) Phragmites australis: excised rhizome with platinum wire cathode around apex. Bar, 15 mm. (b) Phragmites australis: transverse section of apical bud of vertical rhizome, stained with phloroglucinol and concentrated hydrochloric acid to indicate lignification (red). Note concentric scale leaves with aerenchyma channels (A); the abaxial epidermal wall thickening is far greater than on the adaxial side. Bar, 600 µm. (c) Phragmites australis: longitudinal section of apical bud of horizontal rhizome. Black indicates gas-transport connections; D, developing nodal diaphragm; P, developing pith cavity; S, scale leaf. Bar, 600 µm. (d) Oryza rhizomatis: transverse section of apical bud of vertical rhizome. Note concentric scale leaves with aerenchyma channels, A; the abaxial epidermal wall thickening is far greater than on the adaxial side. Bar, 600 µm. (e) Oryza rhizomatis: longitudinal section of apical bud of horizontal rhizome. Black indicates gas-transport connections; D, developing nodal diaphragm; P, developing pith cavity; arrows, trabeculae (diaphragms) within scale leaf. Bar, 600 µm. (f) Phragmites australis: sliver from abaxial surface of scale leaf epidermis of vertical rhizome apex. Note stomata (arrows). Bar, 50 µm. (g) Phragmites australis: transverse section of scale leaf to show porous trabecula (diaphragm) T, composed of stellate parenchyma, across aerenchyma channel; cf. (e). Bar, 130 µm.

Perennial rice, Carex, Glyceria and Phragmites plants  Each plant was removed from the Hoagland's solution, the rhizomes and roots were washed with tap water, and coiled platinum wire electrodes fitted over rhizome apices, as described previously. The lead from each electrode was secured to the plant, and the rhizomes and roots were submerged in soil slurry together with fibre tips of the Ag–AgCl electrodes; here the depth of water above the slurry was approx. 30 mm. The shoots were emergent and the rhizome apices were submerged to a depth of at least 40 mm within the slurry.

Radial oxygen loss (ROL) to the phyllospheres (from rhizome apices)

Coiled Pt wire cathodes used in conjunction with Ag–AgCl anodes (as above) were attached to polarographs and polarized at an appropriate voltage to ensure all O2 molecules reaching the cathode were electrolytically reduced to water. At equilibrium, the electrolysis current was used to calculate the O2-diffusion rate at the cathode surface. Unlike the cylindrical cathodes used previously by Armstrong (1979), these coiled cathodes, because of their open nature, could not be used to assess the rate of O2 per unit area of rhizome tip. Here ROL was expressed in terms of O2 diffusion rate (ng min−1).

Excised rhizomes (Phragmites)  Electrodes and rhizomes were submerged in slurry as for redox measurement.

Intact plants (all species)  Electrodes were applied to rhizome tips as described previously for whole plants. The plants were replaced in the glass tubes with roots and rhizomes submerged in stagnant agar (0.05% w/v containing 7 mm KCl), previously degassed by autoclaving and bubbling with nitrogen. The tubes had special rubber bungs to anchor the cathode connections and Ag–AgCl anodes and the base of the emergent shoot, and to minimize leakage of oxygen from the atmosphere into the agar. The rhizome apices were at least 40 mm below the surface of the agar. Control cathodes were positioned as far from the rhizomes as possible. For ROL measurements in relation to shoot submergence, special acrylic tubes were used (see below).

Altering resistance to gaseous diffusion

Excised Phragmites rhizomes were attached to either (a) a callused node (from a damaged rhizome) using rubber tubing or (b) a 2-µl glass microcap capillary (33 mm length; resistance to gaseous diffusion = 0.272 × 105 s cm−3) using Terostat. In each case the free end of the system connected to the atmosphere.

Humidity-induced convective gas flow

Excised rhizomes  Humidity-induced convective flow (HIC) in an intact plant was simulated. The cut end of the rhizome (submerged under at least 60 mm depth of soil slurry) was attached via a T-piece to the cut end of the rhizome of a leafy Phragmites plant, which induced the flow (approx. 1 m shoot height). The other end of the T-piece was connected via rubber tubing to a senesced efflux culm (1.5 m height) with the apex snapped off. To permit only diffusive transport to the rhizome via the senesced culm, the flow from the leafy culm was blanked off.

Rates of convective flow from the living shoot before and after the experiment were measured by attaching a soap-film flow meter between the cut end of the excised rhizome and the efflux culm.

Whole plants: varying the rate of HIC  Plants were transferred to troughs of soil slurry; shoots were either in air (56–57% RH) to promote HIC, or surrounded by a polythene bag so that convection was slowed down as RH increased to 94–95%. Venting of the convective flow was through the cut end of a rhizome, as the plants were < 1 yr old and had no dead efflux culms. The RH was measured with a humidity meter (HMP.32UT; Vaisala, Helsinki, Finland); convective flow rates were measured by connecting the flow meter to the cut end of the rhizome. The roots and rhizomes of these plants were submerged under at least 40 mm depth of soil slurry.

Changing the gas regime

Excised rhizomes (Phragmites)  Slow flows (100 cm2 min−1) of N2, air or O2 from gas cylinders were circulated through the T-piece attached to the cut end of the rhizome.

Whole plants  For rice, Carex and Glyceria, shoots were each enclosed in a polythene bag through which slow flows (100 cm2 min−1) of N2 or air were circulated from gas cylinders.

Shoot-system submergence

Whole Phragmites plants were transferred initially to acrylic tubes (60 mm diameter, 250 mm length). These were then fitted with a succession of similar, but open-ended tubes (200 mm length) to increase the head space and facilitate submergence of the shoot system in deoxygenated agar. The same rubber bungs as described for the glass tubes were used to anchor the plant and electrodes in the bottom tube. To prevent shoot and possibly rhizome photosynthesis, the growth room was kept in darkness except for a dim light used when modifying the apparatus or to take ROL readings. As an added precaution, tubes were covered in black polythene.

Experiments

The following experiments were each performed two to four times using different rhizomes or plants.

(1) Phyllosphere redox potentials in soil slurry  Effects of duration of rhizome submergence, diffusive resistance, internal gas regime and convective flow.

  • (a) Submergence of excised Phragmites rhizomes: with the cut ends of the rhizomes connected to the outside air, redox potentials were monitored for 114 h after submergence.
  • (b) Diffusive resistance: a resistance to gas diffusion (a 2 µl microcap 2.72 × 104 s cm−3) was connected at the cut end of a Phragmites rhizome, the free end being open to the air. The resistance was inserted for 40 h, removed for 5 h, inserted again for 37 h, and then removed.
  • (c) Changes of gas regime at the cut ends of Phragmites rhizomes: air was applied for approx. 330 min, followed by N2 (24 min), then air (860 min), back to N2 (66 min) and finally O2.
  • (d) Changes of gas regime around the shoots of intact rice, Carex and Glyceria plants: here shoots were in air for at least 18 h, followed by 2–2.25 h in N2, and then returned to the air for 18 h.
  • (e) Humidity-induced convective flow rates in intact Phragmites plants: redox values were measured when convective flow rates were 0.4, 0.08 and again 0.4 cm3 min−1.

(2) Radial oxygen loss to Phragmites phyllospheres  Effects of gas-flow resistance, convective flow and shoot submergence. In experiments 2a and 2b the excised rhizomes were submerged in soil slurry; in experiment 2c the roots and rhizomes of intact plants were submerged in deoxygenated agar.

  • (a) A callused rhizome node forming a resistance to gas flow was attached to the cut end of a Phragmites rhizome for 55 h and subsequently removed.
  • (b) A convective flow of air was induced across the cut ends of excised Phragmites rhizomes. The first period of convection was approx. 5.25 h, followed by 14 h without convection, then 21 h with convection, after which convection was stopped.
  • (c) Shoot submergence. Initially all culms were emergent for approx. 7.75 h. The shoot system was then submerged for approx. 6 h, with only the tips of the leaves emergent. The water level was then dropped for 14 h to expose three-quarters of the height of the shoot system, and finally the water level was dropped to its original value to expose all the shoots fully again. The ROL was measured from the tips of both vertical and horizontal rhizomes.

(3) Test for radial oxygen loss from mature parts of rhizomes  On intact plants of Glyceria, Carex and Phragmites, with shoots emergent and rhizomes and roots submerged in deoxygenated agar, ROL to electrodes coiled around mature parts of rhizomes was investigated as described previously. The plants were kept in the dark to prevent photosynthesis of the rhizomes. Unfortunately, it was difficult to investigate rice because the subapical region of the rhizome was too short to take a coiled electrode and it was covered in roots, from which ROL seemed likely.

Anatomy

For each species, hand-cut transverse and longitudinal sections of the apical 5–10 mm of vertical and horizontal rhizomes, and epidermal slivers from the adaxial and abaxial surfaces of scale leaves surrounding rhizome apices, were examined using an Olympus BX40 photomicroscope. Transverse and longitudinal sections of leaf sheath, stem and rhizome, and of the shoot–rhizome junctions, were examined for signs of gas connections. Sections were stained with phloroglucinol and concentrated hydrochloric acid (confirmed with aniline hydrochloride) to indicate lignification, or with Sudan III to detect suberization/cuticularization (Gurr, 1965). The latter was also investigated in transverse sections of rhizomes by looking for yellow autofluorescence in blue light.

For each species, attempts were made to syringe air under pressure (> 1 kPa) through the cut end of an emergent shoot of an intact plant, via the rhizome and out of submerged buds. The rhizome buds and roots were under water. Using similar methods, gas-space connections were investigated between leaf sheath and stem, and between stem and rhizome.

Results

Experiments

Apart from experiments 1d and 1e, it was not possible to combine the results for replicates because of the slightly different natures of the plants, soil/agar samples and durations of experiments. The results presented are typical examples; the replicates all produced similar patterns.

(1) Phyllosphere redox potentials

(a) Phragmites: effects of duration of rhizome submergence. Initially, all electrodes read approx. 300 mV because of oxygenation of the slurry while pouring it over the rhizomes, but after 114 h redox values around apical and subapical parts of the rhizome were 425 and 78 mV, respectively, whereas the bulk soil was at −70 mV (Fig. 1).

Figure 1.

Phragmites: effects of soil flooding on redox potentials (mV) in phyllospheres around the apex (•) and subapical region (○) of excised horizontal rhizome and in bulk soil (▴). The cut end of the rhizome was connected to the atmosphere.

(b) Phragmites: effect of 2-µl microcap gas diffusion resistance. Insertion of the microcap resulted in a fall in redox potential around the rhizome apex from approx. +420 to +75 mV over 40 h (Fig. 2); removal of the resistance resulted in an almost immediate response and a rise in redox to the original value during the subsequent 5 h; repetition of the procedures produced similar effects. The redox potentials remote from the rhizomes declined from 0 to −90 mV.

Figure 2.

Phragmites: effects on redox potential (mV) in phyllosphere around the apex of excised horizontal rhizome (•) and in the bulk soil (▴) of insertion and removal of microcap resistances in the rhizome aeration path. The cut end of the rhizome was connected to the atmosphere.

(c) Phragmites: effects of changes in gas regime on phyllosphere redox potentials. With the cut end of the rhizome connected to still air, apical and subapical electrodes were reading approx. +90 to +95 mV, respectively (Fig. 3). When the end of the rhizome was exposed to nitrogen, readings for apical and subapical electrodes fell to +60 and −72 mV, respectively, within 24 min. The more rapid decrease for the subapical electrode was possibly caused by an initially narrower oxygenated rhizosphere in this region compared with the apex. On exposing the rhizome end to air, the readings rose to +177 and +115 mV, respectively, within 90 min. Similar effects were repeated on exposing the rhizome end to nitrogen and then to oxygen rather than air, but here, as expected, readings rose to values higher than with air. The readings for the control electrodes remained virtually constant at approx. −80 to −90 mV throughout the experiment.

Figure 3.

Phragmites: effects on redox potential (mV) in phyllospheres around the apex (•) and subapical region (○) of excised horizontal rhizome and in the bulk soil (▴) of applying N2, air or O2 at the cut end of the rhizome.

(d) Oryza, Glyceria and Carex: effects of changes in gas regime around shoots on phyllosphere redox potentials. For all three species, after the emergent shoots had been in air for 18 h, the phyllosphere redox potentials were more positive by 45, 234 and 338 mV, respectively, for perennial rice, Carex and Glyceria than the sediment remote from the submerged buds (Table 1). When the shoots were in nitrogen, phyllosphere redox potential started to become more reducing within 5–10 min, and after 2–2.25 h values had fallen by 36, 179 and 56 mV for rice, Carex and Glyceria, respectively. After re-exposing the shoots to air, redox potentials became more positive again, and after 18 h values resembled the original values in air. Redox measurements for control electrodes remained comparatively constant throughout the experiment, between −207 and −227 mV. The experiment indicated that these species also induce positive redox values in the phyllosphere regions, probably resulting from ROL from submerged buds.

Table 1.  Effects of air and nitrogen around emergent shoots on redox potentials developed in the phyllospheres around rhizome apices submerged in waterlogged soil
Redox potentials (mV)Oryza rhizomatisCarex rostrataGlyceria maxima
ControlsPhyllosphereControlsPhyllosphereControlsPhyllosphere
  1. Means ± SE, n = 5. Within columns, letters denote significant differences: Mann–Whitney rank sum test, P ≤ 0.029.

  2. For Carex and Glyceria data were for both vertical and horizontal rhizomes; for Oryza only vertical rhizomes were used.

Shoots in air after 18 h−223 ± 5.5a−178 ± 12b−221 ± 4.8a  13 ± 55d−217 ± 3a121 ± 14f
After shoots 2–2.25 h in N2−227 ± 4.5a−214 ± 8c−220 ± 4.5a−166 ± 17e−207 ± 12a 65 ± 18g
After shoots returned to air for 18 h−226 ± 3.9a−144 ± 14b−221 ± 4.7a  12 ± 67d−211 ± 9a137 ± 16f

(e) Phragmites (intact plants): effects of convective flow on phyllosphere redox potentials. Increased relative humidity around Phragmites shoots decreased the rate of humidity-induced convection (Table 2). After the shoots had been in comparatively dry air (56–57% RH) for 3 h, convective flow rates were 0.4 cm3 min−1, whereas rates decreased to 0.084 cm3 min−1 after 2.5 h exposure to humid air (94–95% RH). The higher convection rate was resumed by putting shoots in drier air again. With the higher convection rate, phyllosphere redox values were +67 to +28 mV compared with −55 mV at the lower convection rate. The plants used were young (300–500-cm shoot height) and produced relatively slow convective flow rates; even so, the effect of increased flows over short time periods on phyllosphere redox potentials was obvious. Redox values for the bulk soil were −212 to −216 mV.

Table 2. Phragmites: effects on phyllosphere redox potentials (around rhizome apices submerged in waterlogged soil) of changing convective flow rates by varying relative humidity around aerial shoots
RH (%)Convective flow rate through plants (cm3 min−1)Phyllosphere redox potentials (mV)Control electrodes bulk soil (mV)
  1. Redox measurements were taken after shoots had initially been in dry air (56% RH) for 3 h, then placed in humid air (94% RH) to reduce the convection rate for 2.5 h, and finally returned to drier air (57% RH) for 2.5 h.

  2. Means ± SE; n = 5–6. Within columns, letters denote significant differences according to t-test (P = 0.014) or Mann–Whitney rank sum test (P = 0.008).

56–57  0.4 ± 0.06d   67 ± 29b−211.8 ± 4.4a
94–950.084 ± 0.03e−54.8 ± 25c−212.2 ± 6.2a
56–58  0.4 ± 0.06d 28.2 ± 8.5b  −216 ± 7.5a

(2) Radial oxygen loss to phyllosphere

(a) Phragmites: effect of callus on ROL from horizontal rhizome apices. A single callused rhizome node in the aeration pathway resulted in an immediate response and a decrease in ROL from the rhizome apex from approx. 22 to 0.1 ng O2 min−1 over 44 h (Fig. 4); removal of the callused node after 64 h resulted in an immediate rise in ROL. The control electrode remote from the rhizome registered zero ROL throughout the experiment.

Figure 4.

Phragmites: effects on oxygen diffusion rate from horizontal rhizome apex to phyllosphere (•) of insertion and removal of callused rhizome node in the rhizome aeration path. The cut end of the rhizome was connected to the atmosphere. ▴, O2 values for bulk soil.

(b) Phragmites: effect of convective flow of air (pressurized ventilation) on ROL from horizontal rhizome apices. The application of a ‘reed-plant-induced’ convective flow of air (4 cm3 min−1) across the cut end of an excised Phragmites rhizome resulted a rise in ROL from the rhizome apex from approx. 2 to 2.6 ng O2 min−1 over 5.3 h (Fig. 5). After stopping convection for approx. 14 h (during which time ROL remained fairly constant), resumption of convection resulted in a further increase in ROL from 2.6 to a maximum of 3.2 ng O2 min−1 over 10 h. Stopping convection then caused an immediate response and fall in ROL. The diffusion current for the control electrode was consistently < 0.05 ng O2 min−1, indicating that there was virtually no oxygen in the soil remote from the rhizome.

Figure 5.

Phragmites: effects on oxygen diffusion rate to phyllosphere of ± convection across the end of an excised horizontal rhizome (•). Natural humidity-induced convection was supplied by a leafy Phragmites plant. O2 diffusion rate for bulk soil < 0.05 ng min−1.

(c) Phragmites: effects of shoot submergence on ROL from apices of horizontal and vertical rhizomes. The ROL from the vertical rhizome apex was consistently higher than from the horizontal rhizome apex (e.g. initially approx. 17 and 6 ng O2 min−1, respectively; Fig. 6), the latter being further from the source of oxygen in the emergent shoot system. After almost complete submergence, ROL for both rhizome apices fell immediately, reaching 0 ng O2 min−1 over 6.2 h. A three-quarter de-submergence of the shoot system induced a rapid rise in ROL; after 14.6 h values were 12.2 and 8 ng O2 min−1, respectively, for vertical and horizontal rhizomes. Subsequent complete de-submergence of the shoot system produced further increases in ROL to 17.2 and 11 ng O2 min−1, respectively. The final value for the horizontal rhizome apex (11 ng O2 min−1) was higher than at the beginning of the experiment; this may have been caused by a change in the position of the spiral electrode. When the shoot system was completely exposed to the air, there may have been some convective flow taking place. The readings of the control electrode remote from rhizomes and roots were close to zero throughout the experiment.

Figure 6.

Phragmites: effects of shoot submergence and de-submergence on oxygen diffusion rate from submerged rhizome apices to phyllospheres. •, Apex of horizontal rhizome; ○, apex of vertical rhizome. O2-diffusion rate in agar remote from rhizome apices and roots was close to zero throughout.

The results show that when gas exchange with the atmosphere is prevented during shoot submergence in darkness, ROL from rhizome apices falls to virtually zero.

(3) Radial oxygen loss from mature rhizomes

Phragmites, Carex and Glyceria: virtually no ROL was detected from mature parts of rhizomes of Phragmites and from Carex rhizomes in general. However, in most Glyceria rhizomes investigated there was some ROL from the rhizome surface, even 20 cm behind the apex, but the flux was far less than from the apex itself. It may have been that these rhizomes were not yet fully mature; it was difficult to find lengths of living, overwintered Glyceria rhizomes as they appeared to rot easily and to be attacked by microorganisms. There was evidence of iron oxidation on the surfaces.

Anatomy

In all species except Carex, with the excised rhizome apex under water, air could be syringed under pressure (approx. 1–2 kPa) in through the cut end of a mature part of the rhizome, to bubble out of the apex and sometimes from between the overlapping scale leaves themselves. By similar methods, interconnecting internal gas-space connections were found between leaf sheath, stem and rhizome. These connections, linking the stomata on aerial shoots with the scale-leaf aerenchyma and stomata on rhizome apices, were also visible microscopically in longitudinal and transverse sections.

Transverse sections of the apical buds of vertical and horizontal rhizomes of all species showed the typical concentric arrangement of the scale leaves around the apical meristem (Fig. 7b,d). Between overlapping scale leaves there are small spaces, commonly leading to a small ‘pore’ at the bud apex. Stomata could be found chiefly on abaxial but rarely on adaxial surfaces of the scale leaves (Fig. 7f), but in Carex and Glyceria the stomata were far more sparse than in the other species. In all species there were quite large stomata on the abaxial surfaces of the tips of the scales. The stomata are subtended by intercellular spaces that connect to the scale leaf aerenchyma channels; the latter, like those of the aerial leaf sheaths, are interrupted by porous, transverse diaphragms or trabeculae (Fig. 7e,g). The scale-leaf aerenchyma leads to the aerenchyma channels (or intercellular spaces in the case of rice) of the rhizome apex; in rice, Phragmites and Glyceria there are nodal connections, via radial channels of high porosity, linking these with the developing pith cavity. These are similar to connections between leaf sheath aerenchyma and pith cavity of the Phragmites culm (Armstrong et al., 1996c). In Carex the rhizome has no pith cavity, but radially arranged cortical lysigenous aerenchyma channels are present. The gas-space system of the apical bud is continuous with that of the mature part of the rhizome behind the bud. Examples of these gas-space connections are visible in longitudinal sections of rhizome apices (Fig. 7c,e), where gas spaces within scale leaves connect with those of the developing pith cavity.

In all species, on scale leaves around rhizome apices the abaxial epidermal walls become increasingly thickened and cuticularized with maturity, but this is far less pronounced in Glyceria. On the adaxial side, however, this cuticularization and thickening develop later (Fig. 7b,d), and in Carex and Glyceria buds they are patchy or almost absent except in the outermost scale. As the bud grows and the scale leaves mature, the epidermal walls become lignified and suberized, their aerenchyma channels become sealed with callus and the scales eventually wither and die, existing as remnants at the rhizome nodes. The development of callus is similar to the sealing of the aerial leaf sheath aerenchyma found during shoot senescence in Phragmites (Armstrong et al., 1996b). Anatomically, the possible pathways for oxygen to pass from rhizome apices to the phyllosphere regions appear to be via scale-leaf stomata and possibly via noncuticularized abaxial and adaxial surfaces of the scale leaves.

In mature rhizomes there are lignified subepidermal fibres; the thickened cuticularized epidermal walls are also lignified but, compared with Phragmites, these cell-wall deposits are not so pronounced in Carex and rice, and are patchy in Glyceria. No stomata were observed on rhizome internodes. In rice, Phragmites and Glyceria the rhizome pith cavity (and in Carex the cortical lysigenous aerenchyma) connects with that of the aerial stem, with the leaf sheath aerenchyma channels and, in turn, with the leaf sheath stomata. Thus in all the species studied there is an internal gas-space connection between the atmosphere and the rhizome scale-leaf aerenchyma and stomata.

Discussion

Phyllosphere redox potentials around rhizome apices were consistently more positive than those of the rooting medium by 170–500 (Phragmites), 45–80 (rhizomatous rice), 230 (Carex), and 340 mV (Glyceria) (Table 2; Figs 1–3). Similarly, for Phragmites ROL to the phyllospheres was higher than O2-diffusion rates in the bulk soil (Figs 4–6). This indicated that oxygen diffuses from rhizome apices, inducing oxidized phyllospheres similarly to the production of oxidized rhizospheres around roots, and confirms the first hypothesis.

For all four species, pressurized air-flow and anatomical studies revealed interconnections between leaf stomata, gas spaces within leaves, stems and rhizomes, and stomata on the scale leaves of rhizome apices. This, together with the detection of ROL to the phyllosphere and the manipulations of oxygen concentrations at the rhizome base or around shoot systems, confirms the second hypothesis, that atmospheric O2 can diffuse along this pathway within the plant and out into the phyllosphere. The patchy cuticularization and relative lack of cell-wall thickening of the adaxial epidermis of young, inner scale leaves could indicate that oxygen may diffuse indirectly through these layers into the phyllosphere. In Carex and Glyceria there was also patchy cuticularization of the abaxial scale-leaf epidermis. These observations could account for the substantial phyllosphere oxidation observed in Carex and Glyceria (Table 1), while scale leaf stomata were comparatively scarce. Our conclusions concerning the pathway by which oxygen diffuses from the rhizome pith cavity via gas spaces of the submerged apex into the phyllosphere are summarized in Fig. 8.

Figure 8.

Phragmites australis, Oryza rhizomatis, Carex rostrata and Glyceria maxima: suggested pathways of oxygen diffusion from vertical or horizontal rhizome into underground bud and the phyllosphere.

Around mature/maturing parts of rhizomes, the absence of ROL and oxidizing power (Phragmites) or reduction in ROL relative to the apex (Carex and Glyceria) correlates with cuticularization, epidermal and hypodermal thickenings, and the absence of stomata on the rhizome wall. In Phragmites, callus in the bases of withered scale leaves apparently prevented oxygen from diffusing from the scales. In view of the strong epidermal and hypodermal thickening and thick cortex of the mature rice rhizome, it seems very unlikely that there would be ROL from here. These findings partly lend support to the third hypothesis, but it seems that some types of mature rhizomes, as in Glyceria, may permit some degree of ROL.

In Phragmites, ROL from the rhizome apex to the phyllosphere was reduced by applied resistances to gas flow, by N2 around the aerial parts, and by shoot submergence; conversely, ROL to the phyllosphere was increased by convective air flow through the rhizome, and by O2 around the aerial shoot. These findings confirmed the fourth hypothesis. When gas exchange with the atmosphere was prevented during shoot submergence, ROL from rhizome apices fell to virtually zero. Similar effects with shoot submergence were found for ROL from root apices in Phragmites (Armstrong et al., 1999), E. angustifolium (Gaynard & Armstrong, 1982) and Halosarcia (Pedersen et al., 2006).

In Phragmites, rhizome nodes with callused diaphragms proved to be very effective resistances to diffusive aeration and virtually impermeable to O2 (experiment 2a); they no doubt prevent diffusive transport to O2 to both roots and underground buds, which form blind-ending parts of the underground system. Callused nodes also prevent convective flow (Armstrong et al., 1996b) and, although useful in terms of sealing wounds, preventing flooding of damaged underground parts and almost certainly in preventing the spread of phytotoxins, they must sometimes be responsible for the death of roots and buds by blocking gas transport from the aerial parts.

In Phragmites, humidity- and Venturi-induced convective flows have been shown to aerate the rhizome system more effectively than diffusion alone (Armstrong et al., 1992). By conveying fresh air to the root–rhizome junctions, they also facilitate diffusive aeration of roots and greater oxygen release to the rhizosphere regions around fine laterals and the apices of adventitious roots than is possible if aeration of the rhizomes is by diffusion alone. As both roots and rhizome apices are blind ends of the underground system, it was not surprising that humidity-induced convective flow in Phragmites also promoted diffusive aeration of rhizome apical buds, as evidenced by increased phyllosphere ROL and redox potentials. Convective flows have not been found so far in O. rhizomatis, Carex and Glyceria (unpublished data), but in species that do exhibit significant convections, such as Nuphar (Dacey, 1981), Nelumbo (Mevi-Schutz & Grosse, 1988), Phragmites (Armstrong & Armstrong, 1990), Typha (Bendix et al., 1994), Eleocharis (Sorrell & Boon, 1994) and others (Brix et al., 1992), and especially in those with deeply growing rhizomes, phyllosphere oxidation enhanced by convective flows could be beneficial. Increased ROL to the phyllospheres induced by convections should enable these species to colonize more reducing sediments than would otherwise be possible and, through enhancement of apical bud aeration, enable horizontal rhizomes to grow longer and deeper, and vertical rhizomes to emerge from greater depths of water. All these effects may well confer competitive advantages and enable more widespread colonization. This is indicated by the work of Vretare & Weisner (2000), who found that in Phragmites, when convective flows are permitted, rhizome growth in terms of length, numbers and biomass is increased. They also showed (Vretare Strand & Weisner, 2002) that for plants growing in organic sediment, convective flows promoted greater O2 concentrations in stem bases and greater uptake of certain minerals.

Epidermal and hypodermal cell-wall thickening and the waxy cuticle of the scale leaves no doubt protect rhizome apices from abrasion, pathogens, phytotoxins and sometimes desiccation. However, the presence of stomata could make them vulnerable. In roots, there is much evidence that the release of oxygen to the rhizosphere regions offers protection to permeable and vulnerable parts of the root system against phytotoxins commonly found in reducing waterlogged soils, such as sulphides, FeII, MnII and the lower volatile organic acids (Mendelssohn & Postek, 1982; Lee et al., 1999; Pedersen et al., 2004). Oxidation of these compounds to less-toxic forms can occur either through the activities of aerobic microorganisms that colonize the oxygenated rhizospheres, or by direct chemical oxidation (Begg et al., 1994; Kirk & Bajita, 1995). The release of oxygen to the phyllospheres could fulfil a similarly protective function for rhizome apices to that of ROL from roots.

Root and rhizome apices of Phragmites can die when exposed experimentally to sulphide (Armstrong et al., 1996a) and the lower volatile organic acids (Armstrong & Armstrong, 1999). Sometimes in waterlogged soils these toxins are at comparatively high concentrations and can induce a dieback syndrome (Armstrong et al., 1996d; Armstrong & Armstrong, 2001a, 2001b). It is likely that here there is insufficient detoxification directly and by the phyllosphere microorganisms, and phytotoxins may enter the rhizome apices via the scale-leaf stomata and/or via permeable, noncuticularized developing scale leaves, resulting in death of the buds. Sulphide may enter in the gas or liquid phase; organic acids, being oily in nature and reducing the surface tension of the water, may enter the scale-leaf stomata in the liquid phase. Toxins may also pass through noncuticularized surfaces of scale leaves; they may also enter the roots and be transported to the buds via the vascular system, and in the case of sulphide and organic acids, also via the gas-space system (Pedersen et al., 2004). Although rhizomes have a greater internal capacity than roots to detoxify sulphide (Furtig et al., 1996), it may be that rhizome apices do not have as great an ability as the mature parts.

It would be interesting to see if metabolites diffuse from rhizome apices, encouraging the growth of aerobes within the phyllosphere regions, as occurs within the rhizospheres around roots. It has been noted that in roots, oxidations within rhizospheres occur quickly, but that re-reduction of the soil can be comparatively slow (personal observation; Jensen et al., 2005). We suggest that the same will apply to phyllosphere oxidations and could be beneficial to the tips during dark periods if shoots are partially submerged (Pedersen et al., 2006); this could also benefit roots emerging from near the tip, as they will tend to grow into a less-toxic medium.

In this study, the possibility that some phyllosphere oxidation could take place around dormant lateral or apical buds on the rhizome has not been examined. However, it is likely that factors that enhance or decrease growing apical bud aeration will similarly affect dormant bud aeration.

In conclusion, we suggest that in rhizomatous emergent aquatic macrophytes colonizing reducing sediments, effective aeration of rhizome apices is necessary to promote aerobic respiration, perennation and competitive vegetative growth, and that ROL from apices to the phyllospheres is important to protect the apices from phytotoxins. In those species capable of increasing the extent and degree of phyllosphere oxygenation through convective flows, these advantages will be enhanced.

Acknowledgements

We thank Mr Vic Swetez of the University of Hull's Botanic Gardens for growing the plants and the International Rice Research Institute, Los Baños, the Philippines for supplying the rhizomatous rice seed.

Ancillary