By continuing to browse this site you agree to us using cookies as described in About Cookies
Notice: Wiley Online Library will be unavailable on Saturday 7th Oct from 03.00 EDT / 08:00 BST / 12:30 IST / 15.00 SGT to 08.00 EDT / 13.00 BST / 17:30 IST / 20.00 SGT and Sunday 8th Oct from 03.00 EDT / 08:00 BST / 12:30 IST / 15.00 SGT to 06.00 EDT / 11.00 BST / 15:30 IST / 18.00 SGT for essential maintenance. Apologies for the inconvenience.
Gas films on hydrophobic surfaces of leaves of some wetland plants can improve O2 and CO2 exchange when completely submerged during floods. Here we investigated the in situ aeration of rhizomes of cordgrass (Spartina anglica) during natural tidal submergence, with focus on the role of leaf gas films on underwater gas exchange. Underwater net photosynthesis was also studied in controlled laboratory experiments. In field experiments, O2 microelectrodes were inserted into rhizomes and pO2 measured throughout two tidal submergence events; one during daylight and one during night-time. Plants had leaf gas films intact or removed. Rhizome pO2 dropped significantly during complete submergence and most severely during night. Leaf gas films: (1) enhanced underwater photosynthesis and pO2 in rhizomes remained above 10 kPa during submergence in light; and (2) facilitated O2 entry from the water into leaves so that rhizome pO2 was about 5 kPa during darkness. This study is the first in situ demonstration of the beneficial effects of leaf gas films on internal aeration in a submerged wetland plant. Leaf gas films likely contribute to submergence tolerance of S. anglica and this feature is expected to also benefit other wetland plant species when submerged.
Submergence tolerance in plants has been classified into two main response types – an escape response involving shoot elongation or a quiescence response to conserve energy until water recedes (Bailey-Serres & Voesenek 2008). Both response types also involve many other traits, such as aerenchyma formation, tolerance of tissue hypoxia and anoxia, protection against free radicals, formation of adventitious roots, and traits enabling at least some photosynthesis when under water (Laan & Blom 1990; Bailey-Serres & Voesenek 2008; Colmer & Voesenek 2009). Well-developed aerenchyma is crucial for efficient aeration of submerged organs and tissues (Armstrong 1979; Jackson & Armstrong 1999; Colmer 2003). Here we focus on underwater photosynthesis and internal aeration via aerenchyma, and in particular the role in these two processes of gas films retained on submerged leaves.
Leaf gas films are a micro-layer, initially of trapped air, on the hydrophobic surface(s) of leaves when submerged. A gas film enlarges the water-gas interface and may also allow stomata to remain open when under water (Pedersen, Rich & Colmer 2009), and so is a feature that facilitates O2 and CO2 exchange between leaves and surrounding waters. Leaf gas films contribute to submergence tolerance of rice (Raskin & Kende 1983; Beckett et al. 1988; Pedersen et al. 2009). Pedersen et al. (2009) showed in laboratory experiments that the internal partial pressure of O2 (pO2) of submerged rice is dependent on the presence of leaf gas films, both during light and dark periods. Removal of leaf gas films during light periods caused root pO2 to drop as a consequence of reduced underwater photosynthesis as CO2 uptake from the water would have been reduced. During darkness, the removal of the gas films also resulted in a significant drop in root pO2 because of decreased O2 entry via the leaves. Interestingly, gas films also occur on leaves of other wetland species (Colmer & Pedersen 2008b), including S. anglica (observed during a study of foliar N uptake; Bouma et al. 2002). Thus, leaf gas films, in addition to aerenchyma and other traits, may contribute to the ability of S. anglica to grow in frequently inundated lower salt marsh areas. S. anglica provides an interesting case study to further elucidate the role of leaf gas films in submergence tolerance, as this species experiences frequent tidal submergences, in contrast with longer-lasting floods that have been the focus of most work on plant submergence tolerance (Bailey-Serres & Voesenek 2008).
Underwater photosynthesis can supply submerged terrestrial plants with O2 produced endogenously, but with large diurnal variations, and also provide carbohydrates for plant metabolism (Laan & Blom 1990). When completely submerged during darkness, O2 available to plants is limited to that which enters via diffusion from the surrounding water. Consequently, internal pO2 declines because of respiration and anoxia can develop in the roots, until pO2 rises again during light periods, reaching a new quasi-steady state between O2 production and respiration/loss to the environment (Waters et al. 1989; Colmer & Pedersen 2008a; Pedersen, Malik & Colmer 2010). Leaf gas films enhance O2 uptake by submerged leaves when in darkness and also increase underwater net photosynthesis when light is available (Colmer & Pedersen 2008b; Pedersen et al. 2009), and some of this O2 moves via aerenchyma to the roots (Pedersen et al. 2009). In a study of leaf gas films on partially submerged rice, Raskin & Kende (1983) suggested that gas films also function as a snorkel promoting a non-throughflow convection of air to the submerged parts of the plant; however, this was later shown to be untenable (Beckett et al. 1988).
We tested three main hypotheses, using S. anglica and short tidal submergence:
1root and rhizome pO2 decline upon submergence because of constraints in O2 exchange with the surrounding water;
2root and rhizome pO2 during submergence increase when light is available to shoots, because of transport of O2 produced by underwater photosynthesis to belowground organs; and
3leaf gas films enhance root and rhizome pO2 during submergence, both during dark and light periods, owing to improved exchange of O2 and CO2 between leaves and the water (O2 entry during darkness; CO2 entry for photosynthesis during light periods).
This study constitutes the first in situ (i.e. field) measurements of the role of leaf gas films on internal plant aeration during complete submergence.
MATERIALS AND METHODS
Study site and plant materials
Skallingen in Ho Bay, Western Jutland (N 55.536943°, E 8.256730°), Denmark, was chosen as the site for in situ studies and for collection of plants used in laboratory experiments. The bay has a large population of S. anglica, which is often inundated at each high tide, twice each 24 h. In this population, the largest plants were approximately 40 cm tall and the longest leaf laminas were approximately 20 cm. Plants for laboratory experiments were collected as turfs (approximately 20 cm by 20 cm wide and 25 cm deep, sediment blocks containing plants) and transported in plastic bags the same day to the laboratory in Hillerød, Denmark. Plants for biomass measurements were collected whole by digging up complete turfs and then separating them into components before oven drying for 48 h at 60 °C.
In situ pO2 dynamics in rhizomes
Intra-plant pO2 dynamics in rhizomes of S. anglica were followed in situ at Ho Bay. An 8-channel picoamperemeter (PA8000, Unisense, Aarhus, Denmark) connected to a laptop running Picolog (Pico Technology Ltd, Cambridgeshire, UK) for data logging, and one three-channel underwater picoamperemeter with built-in data logger (PA3000UP-OP, Unisense) were used. 50 µm tip diameter O2 microelectrodes (OX-50UW, Unisense) were used to measure inside the rhizomes and 500 µm tip diameter electrodes were used to measure O2 in the water and sediment. The electrodes were calibrated immediately before use in water at air equilibrium (20.6 kPa O2) and in anoxic water with dithionite (0 kPa O2).
Small areas of sediment within selected patches of S. anglica were gently excavated until a rhizome was found. The rhizome was further exposed using water to wash away the sediment. Micromanipulators with microelectrodes were mounted on aluminium stands fixed in the sediment, and the microelectrodes were positioned into rhizomes using changes in signal to detect the surface of the rhizomes following the procedure of Borum et al. (2006) and Pedersen, Vos & Colmer (2006). Oxygen microelectrodes were inserted approximately 1000 µm into the rhizomes and sediment was then gently added to again cover the rhizome and electrode until both were under at least 4 cm of sediment. When using this procedure, the bio-geochemical profiles in marine sediments are re-established within 1 h (Pedersen, Binzer & Borum 2004). Other electrodes were positioned 4 cm into the sediment, at the sediment surface and at or just above the canopy top. Six electrodes were placed in six different plants, three controls and three treatments. Patches of 8–12 shoots were isolated from nearby patches of shoots by cutting to a depth of 25 cm with a large knife in a circle around each patch – this was done as otherwise extensive numbers of neighbouring shoots would likely have needed dilute Triton X brushing to remove all leaf gas films that could facilitate O2 entry along the measured rhizome – and this was not feasible with the time constraint imposed by a rising tide. The treated plants were, just prior to tidal inundation, brushed with Triton X (0.1% v/v dilution in bay water) to reduce surface hydrophobicity so that gas films did not form upon submergence. Two tide cycles were monitored, one during day light and one during night-time. Control plants remained the same but the treatment plants were changed after the first tide cycle to avoid any complications from potential lingering effects of brushing leaves with dilute Triton X on the treated plants.
Temperature and light were measured using two pendant loggers (HOBO Pendant Temp/Light Data Logger UA-001–08 Onset Computer Corporation, Pocasset, MA, USA), one buried 4 cm in the sediment and the other placed on the sediment surface. A water level data logger (HOBO U20, Bourne, MA, USA) was used to measure the water depth. Water pH was measured using a pH electrode (Mettler Toledo LO T403-M8, Greiffen see, Switzerland) connected to a pH meter (Radiometer pHM, Willich, Germany). Data were recorded from low tide to low tide. Water samples were brought back to the laboratory to measure salinity (YSI Salinity 30 M/10FT, Yellow Springs, OH, USA) and alkalinity by Gran titration (Stumm & Morgan 1996).
Root pO2 dynamics – laboratory experiments
S. anglica collected from the field were transferred into hydroponic culture medium. Cuttings were washed from the sediments and mounted in holes in bucket lids with shoot bases held using foam so that roots and rhizomes were in culture medium inside the buckets. The composition of the nutrient solution was (in mm): Na+, 275; Cl-, 275; K+, 3.5; Ca2+, 2.0; Mg2+, 1.0; NO3-, 6.6; SO42−, 1.0; H2PO4-, 0.5; Mn2+, 0.0045; Zn2+, 0.004; Cu2+, 0.0015; BO33−, 0.023; MoO42−, 0.00005; FeEDTA, 0.0375. There was no forced aeration (i.e. no bubbling) of the medium. The buckets were covered with aluminium foil to prevent light entry and algae growth. Cuttings were grown for at least 4 weeks before being used in experiments.
For use in laboratory experiments, cuttings were transferred from the hydroponics into a horizontal chamber (length 26 cm, width 7 cm) made from Perspex (Colmer & Pedersen 2008a). The plant was positioned so that roots, rhizome and the shoot base were inside the chamber in previously deoxygenated half-strength artificial seawater [Smart & Barko (1985) solution with added NaCl (275 mm), KHCO3 (2.2 mm), pH was 9.1] and the medium also contained 0.1% (w/v) dissolved agar, to prevent convective movement. The shoot was outside the root-rhizome chamber as it emerged through a hole in the side sealed with blu-tac putty (Bostik, Riverside, England) to make the seal water tight. The root-rhizome chamber was placed in a larger transparent container, thus allowing artificial seawater to be added to submerge the horizontally placed shoot protruding from the root-rhizome chamber but without disturbing the roots and rhizome. Initially the shoots were in air. The shoots were then submerged using artificial seawater at 30 ppt NaCl (513 mm; see underwater photosynthesis measurements for composition) bubbled with air. After near steady state, water was temporarily lowered, the adaxial leaf side brushed with Triton X (0.05%) and then submerged again. Temperatures were 17 to 19 °C. During light periods, photosynthetically active radiation (PAR) (400–500 µmol m−2 s−1) was provided to the shoots by halogen lights and a fluorescent work lamp.
Clark-type O2 microelectrodes with a guard cathode and tip diameter of 25 µm (OX-25, Unisense A/S, Aarhus, Denmark) were used. Electrodes were calibrated immediately before use in water at air equilibrium (20.6 kPa O2) and in anoxic water with dithionite (0 kPa O2). A micromanipulator (MM5, Märzhäuser, Wetzlar, Germany) was used to insert the tip of the microelectrode, 200–300 µm into a root 2–4 cm from the root-rhizome junction. The microelectrode was connected to a multimeter (MicroSensor Multimeter, Unisense A/S) and the output was logged every 60 s on a computer using SensorTrace basic (Unisense A/S).
Underwater net photosynthesis
Two leaf lamina segments (∼2.5 cm in length) from the middle of each leaf were excised using a razor blade. One was used as the control (with gas film) and the other was used as the treatment in which gas film formation was prevented, by light brushing five times, on the adaxial side (the hydrophobic side), with a fine paintbrush soaked in 0.05% Triton X in incubation medium (composition given next), then washed for 5 s, three times, in medium without Triton X. This treatment prevented formation of a gas film on the adaxial lamina surface when submerged (cf. Colmer & Pedersen 2008b).
Net photosynthesis under water, by lamina segments (with or without gas films), was measured using the method described by Sand-Jensen, Pedersen & Nielsen (1992), with some modifications. The glass bottles used were 25 mL, and two glass beads were added to ensure mixing as the bottles rotated inside the illuminated water bath at 20 °C (cf. Colmer & Pedersen 2008b); one lamina segment was placed in each bottle. PAR inside the glass bottles was 550 µmol m−2 s−1 (measured using a 4π US-SQS/L Wals, Effeltrich, Germany).
The incubation medium was based on the general purpose culture medium described by Smart & Barko (1985) and contained (in mm): Ca2+, 0.62; Mg2+, 0.28; Cl-, 1.24; SO42−, 0.28; but also KHCO3 and NaCl were added so that alkalinity was 2.2 mm and salinity 30 ppt (513 mm) NaCl. The dissolved O2 concentration in the incubation was set at 50% of air equilibrium, by bubbling 1:1 volumes of N2 and air; this procedure was applied to prevent increase in O2 above air equilibrium levels during measurements that might have led to photorespiration and thus decreased net photosynthesis (Colmer & Pedersen 2008b). Because the glass bottles were incubated in light immediately after adding the lamina segments there was no risk of tissue hypoxia as O2 would have been produced. The pH was 8.04, which resulted in approximately 15 µM CO2, and leaves were unable to use HCO3- as a carbon source (own unpublished data).
Following incubations of known duration, dissolved O2 concentration in each bottle was measured using an O2 microelectrode (OX-500, Unisense A/S) connected to a picoamperemeter (PA2000, Unisense). The electrode was calibrated as described previously. Dissolved O2 concentrations in bottles prepared and incubated in the same way as described previously, but without lamina segments, served as blanks. The projected area of each lamina segment was measured using a leaf-area meter (LI-3000). Five replicates for controls and treatments were measured.
Underwater dark respiration
Underwater dark respiration was measured using the same method as underwater net photosynthesis, but the incubations were in darkness. O2 concentration in the medium commenced at air equilibrium (20.6 kPa, 237 µM O2 at 20 °C, 30 ppt NaCl) and declined during incubation to an average of 204 µM O2.
Tissue porosity and leaf gas film thickness
Porosity (% gas spaces per unit tissue volume) was measured for leaves, rhizomes and roots, by determining plant tissue buoyancy before and after vacuum infiltration of the gas spaces with water (Raskin 1983), using the equations as modified by Thomson et al. (1990). Triton X at 0.05% was used to remove surface gas films on leaf segments, and care was taken to ensure no external gas was trapped between tissue segments. Leaves, rhizomes and roots were cut into 50 mm segments for the measurements; only mid-leaf segments were used for leaf porosity measurements. To estimate the leaf gas film thickness, gas film volume was measured and related to leaf area. Buoyancy of leaf segments under water with gas films intact was determined, and then gas films were removed by brushing with 0.05% Triton X and the measurements were repeated. Leaf segment projected areas were measured using a leaf-area meter (LI-3000, Li-Cor, Lincoln, NE, USA).
Leaf surface hydrophobicity (or non-wettability), specific leaf area and belowground-to-aboveground dry mass ratio
Surface hydrophobicity was assessed by measuring the contact angle of a 5 mm3 droplet of water on the leaf surfaces (Adam 1963; Brewer & Smith 1997). Lamina segments were held flat using double-sided tape. Droplets were applied to the lamina of 10 replicate leaves, five on the adaxial side and five on the abaxial side, and photographed at ×16 magnification using a horizontally positioned dissecting microscope (Leica MS5, Solms, Germany) and digital camera. The droplet contact angles were measured on a computer running ImageJ (ImageJ v.1.43U, National Institutes of Health, Bethesda, MD, USA).
Specific leaf area (SLA) of lamina was measured by determining the area (LI-3000, Li-Cor) and dry mass of samples. Belowground-to-aboveground dry mass ratio was determined by separating roots and rhizomes from the shoots and drying each fraction for 48 h at 60 °C, recording dry mass, and calculating the ratio.
Scanning electron micrograph pictures of leaf laminas were taken using a field emission scanning electron microscope (JEOL JSM-6335F, Peabody, MA, USA) using the approach of Madsen (2009).
GraphPad Prism 5 (GraphPad Software Inc., http://www.graphpad.com) was used for data analysis and statistics including two-way anova with a Bonferroni post hoc and Student t-test to compare means. Mean variations are given as Standard Error Measurements (±SEM); probability level 0.05. Data for the laboratory experiments in Table 2 were squareroot transformed before testing for significant differences between mean values.
Table 2. Influence of leaf gas films on rhizome (field) or root (laboratory) pO2 (kPa) in Spartina anglica with shoots in air or shoots completely submerged
Controls are plants with leaf gas films intact (abbreviated as +GF in lower part of the Table); treated plants had leaf gas films removed (abbreviated as –GF in lower part of the Table) with 0.1% Triton X just prior to inundation in the in situ experiments. In the laboratory experiments, the leaf gas films were removed (0.05% Triton X) after quasi-steady state had been achieved. Significant differences between the two treatments at each condition were tested by Student's t-test on in situ data. Two-way anova was used on square root transformed data for the laboratory study. Data are mean ± SEM; significance levels are P ≤ 0.05 (n = 3).
In situ experiments
Rhizome pO2 (kPa) shoots in air
18.3 ± 1.0a
18.2 ± 1.1a
12.4 ± 0.8a
12.3 ± 0.8a
Rhizome pO2 (kPa) shoots submerged
10.6 ± 0.8a
6.0 ± 0.4b
5.0 ± 0.6a
1.4 ± 0.0b
Shoots in air
Root pO2 (kPa)
13.0 ± 1.4a
10.2 ± 0.7a
4.8 ± 1.0b
9.9 ± 1.4a
3.5 ± 1.5b
0.3 ± 0.1c
Surface hydrophobicity, tissue porosity and leaf gas film volume of S. anglica
Plant characteristics are summarized in Table 1. Water droplet contact angle was measured on the adaxial and abaxial sides of S. anglica leaves to determine the surface hydrophobicity. A droplet contact angle of ≥110° is considered non-wettable and therefore hydrophobic (Brewer & Smith 1997). The adaxial leaf side had an average droplet contact angle of 148° and the abaxial leaf side had an average contact angle of 53°. We presume that a non-wettable leaf surface is a prerequisite for the formation of leaf gas films. This supports our observations that gas films only form on the adaxial side of S. anglica leaves. Interestingly, all stomata are located on the adaxial side of S. anglica leaves (Maricle et al. 2009). Average gas film thickness was 50 mm, which is consistent with data for this feature also on lamina of rice (Pedersen et al. 2009).
Table 1. Leaf characteristics and tissue porosity of Spartina anglica
Hydrophobicity was measured using the water droplet contact angle (Brewer & Smith 1997). Tissue porosity and gas film volume were both determined as change in buoyancy after vacuum infiltration of water or removal of gas films, respectively (Pedersen et al. 2009). Gas film thickness is for the adaxial side, as the abaxial side does not have a gas film and was determined using gas film volume and leaf area. Belowground-to-aboveground dry mass ratio is from field-collected S. anglica. Hydroponically grown cuttings of S. anglica had 0.92 ± 0.16 (n = 6) belowground-to-aboveground dry mass ratio (cuttings had been trimmed of most rhizomes).
Porosity is the % of gas volume per unit tissue volume and was measured to evaluate the potential for gas diffusion in leaves, rhizomes and roots. The lowest porosity was measured in the lamina at 7.4%. The leaf sheath, rhizome and roots had porosities (%) of 32, 54 and 37, respectively.
S. anglica in situ O2 dynamics in rhizomes
The sediment at Ho Bay consisted of fine sand and clay and it stayed waterlogged throughout the study period with scattered puddles that remained until the next high tide. Sediments at 4 cm depth stayed anoxic during the 24 h it was monitored (i.e. both at high and low tide; data not shown). During high tide, water column O2 concentrations were higher during the daylight tide at 256 to 371 µM O2 (air equilibrium = 249.3 µm at 20 °C and 22 ppt NaCl) than during the night tide with 212 to 228 µm O2. Alkalinity was 2.2 mequiv. L−1, salinity 22 ppt, and with pH varying between 7.80 in the morning and 8.45 during the afternoon.
Rhizome pO2 was measured in natural stands of S. anglica during two consecutive tidal submergence events; one in daylight and one during night-time. To evaluate the importance of leaf gas films for rhizome pO2 during inundation causing complete submergence, the formation of leaf gas films was prevented by brushing the leaves in three patches with 0.1% v/v Triton X in bay water, while three other patches served as controls. During low tide, rhizome pO2 of plants with shoots in air varied from 18.4 to 21.8 kPa and there was no significant difference in rhizome pO2 between the treatment and control plants. Inundation resulted in a dramatic drop of the rhizome pO2 in both control and treatment plants, but rhizome pO2 remained significantly higher in plants with leaf gas films intact, both when in light (Fig. 1) as well as in darkness (Fig. 2). When inundated in daylight (Fig. 1), rhizome pO2 of S. anglica decreased from ∼18 kPa prior to submergence, to ∼10 kPa (lowest mean pO2 value before pO2 starts to rise after re-exposure to air when tide receded) with leaf gas films intact, but declined further to 6.0 kPa with leaf gas films removed (i.e. pO2 with leaf gas films intact was 1.8-fold higher than when gas films were removed; Table 2). During night (Fig. 2), inundation caused pO2 in the rhizomes to decrease from 12.4 to 5.0 kPa in plants with intact gas films, but dropped much lower for plants without gas films to 1.4 kPa (i.e. rhizome pO2 was 3.7-fold higher in plants with leaf gas films; Table 2). Interestingly, when shoots were in air, rhizome pO2 was significantly higher during daylight hours than during the night (Figs 1 & 2, Table 2).
S. anglica O2 dynamics in roots – laboratory experiments
Root pO2 was measured in six hydroponically grown S. anglica plants under controlled laboratory conditions to verify the in situ data and to evaluate the influence of leaf gas films on root pO2. We measured root pO2 during submergence of plants with or without leaf gas films, both in light and darkness (Fig. 3). Root pO2 values when the shoots were in air and in light ranged from 9.5 to 16.4 kPa, and in darkness from 8.4 to 12.4 kPa (Table 2), with mean values not significantly different between light and dark regimes. Submergence during light with leaf gas films intact decreased root pO2 to 78% of initial values with an average of 10.2 kPa, whereas removal of the leaf gas films further reduced root pO2 to 37% of initial values with shoots in air, resulting in an average root pO2 of 4.8 kPa in submerged plants without leaf gas films. Submergence during dark resulted in a decrease to 35% of initial values and removal of leaf gas films further reduced it to 2.5% of initial values, resulting in an average root pO2 of 0.3 kPa for plants submerged without leaf gas films.
Underwater net photosynthesis
Underwater net photosynthesis of leaves with or without gas films was measured at light saturation (550 µmol photons m−2 s−1) with 15 µm free CO2 and 2.2 mm KHCO3 (Table 3). Leaves with gas films had a 2.7-fold higher underwater net photosynthesis (0.54 µmol O2 m−2 s−1) compared with leaves without gas films (0.20 µmol O2 m−2 s−1). The rates of underwater net photosynthesis compare with photosynthesis in air by leaves of S. anglica of 12.5 µmol m−2 s−1 (Mallott et al. 1975) so that underwater net photosynthesis with leaf gas films was 4% and without leaf gas films 2% of rates in air.
Table 3. Underwater net photosynthesis and dark respiration of Spartina anglica leaf segments with or without gas films
With gas film
Without gas film
Leaves were excised just prior to use from plants collected from the Ho Bay field site. Experiments were conducted at 20 °C and 550 µmol photons m−2 s−1 of PAR. The medium used was artificial sea water based on a culture medium from Smart & Barko (1985) with 513 mm NaCl added (30 ppt). Alkalinity was 2.2 mequiv. L−1 and contained 15 µm CO2. Underwater respiration was measured in water at 84–100% air equilibrium. SLA is given in Table 1, so as to enable conversions to a dry mass basis if needed. Significant differences were tested by Student's t-test. Data are mean ± SEM; significant levels are P ≤ 0.05 (n = 5).
PAR, photosynthetically active radiation; SLA, specific leaf area.
Underwater net photosynthesis (µmol O2 m−2 s−1)
0.54 ± 0.04a
0.20 ± 0.03b
Underwater dark respiration (µmol O2 m−2 s−1)
0.53 ± 0.01a
0.50 ± 0.01a
Underwater O2 uptake in darkness
Underwater respiration by lamina segments with or without gas films was measured when in water near air equilibrium O2. There was no significant difference in respiration of leaf segments with (0.53 µmol O2 m−2 s−1) and without (0.50 µmol O2 m−2 s−1) gas films when in water with O2 at near air equilibrium (Table 3).
The present study evaluated the influence of leaf gas films on internal aeration of rhizomes of a salt marsh plant (S. anglica) in the field during tidal submergence, both in daylight and during the night. We also evaluated, in laboratory experiments, underwater net photosynthesis, tissue porosity, root O2 status and other features of S. anglica, as related to submergence tolerance. We found that, when challenged by complete submergence lasting for several hours: (1) pO2 in rhizomes and roots showed substantial declines both in light and dark submergence events; (2) underwater net photosynthesis was stimulated by the presence of leaf gas films and that higher underwater photosynthesis translated into higher pO2 in belowground tissues; and (3) night-time O2 uptake from the surrounding water, and thus internal pO2, was significantly increased by the presence of leaf gas films. The eco-physiological implications of these key findings are discussed next.
Root and rhizome aeration of S. anglica is negatively influenced by tidal inundation
During submergence, internal aeration of tissues can be restricted because of constraints on O2 uptake, primarily because of the slow diffusion of gases in water compared with in air (Armstrong 1979). The constraints on O2 uptake caused by tidal inundation resulted in declines in rhizome pO2 of S. anglica (Figs 1 & 2) because the O2 supply from shoots could not keep up with the demand in the roots and rhizomes that rely on O2 supplied by the shoot when in anoxic sediment (Revsbech et al. 1980). Declines in O2 were also found in the shoot base of Spartina alterniflora during tidal submergence (Gleason & Zieman 1981). The rhizome of S. anglica has well-developed lacunae and a large central hollow pith resulting in high tissue porosity (Table 1) facilitating gas phase diffusion of O2 from the shoot to the rhizome and further into the roots that also contain aerenchyma. We found that during natural tidal submergence, S. anglica was able to maintain oxic rhizomes both during day and night, albeit at a pO2 level below atmospheric. Following submergence, quasi-steady state levels were reached within a few hours, indicating that even if floods lasted longer, rhizomes would likely have continued to receive O2. Our laboratory experiments showed that during complete submergence, S. anglica was also able to keep the upper part of the roots oxic over a prolonged period, as long as leaf gas films were present.
In the field situation, pO2 in belowground tissues of S. anglica while the shoot was in air was higher during daytime than at night-time. A likely explanation could be that the increased pO2 in the rhizomes was derived from photosynthesis in the leaf sheaths, which could be using CO2 from the sediment (Hwang & Morris 1992; Winkel & Borum 2009; Pedersen et al. 2011). Thus, any O2 produced would elevate pO2 in the rhizome particularly if stomata are few on the abaxial side of the sheath facing outside so that gas exchange with the surrounding air would be restricted and so downwards movement of O2 is promoted. Alternatively, there could be a pressure-driven mass flow of air occurring from the shoots and along the rhizome during the day when shoots are in air, although it is uncertain if S. anglica has such flows. Pressure-driven flows have been shown to occur in several wetland plants (Brix, Sorrell & Schierup 1996; Grosse, Armstrong & Armstrong 1996; Sorrell, Brix & Orr 1997) and Hwang & Morris (1991) showed that shoots of S. alterniflora are capable of hygrometric pressurization, although orders of magnitudes lower than pressures reported in Phragmites australis (Brix et al. 1996). If S. anglica is also capable of hygrometric pressurization, and if a through-flow pathway is present, rhizome pO2 would then be higher during the daytime when the shoots were in air. Any pressure-driven mass flow, if present, would cease upon complete submergence of the shoots.
Leaf gas films enhance in situ pO2 in rhizomes during tidal inundation
By possessing leaf gas films, root and rhizome pO2 of S. anglica is enhanced during tidal inundation. During the day, leaf gas films enable pO2 in the rhizome to remain relatively high when submerged (10 kPa). Removal of gas films resulted in declines in rhizome pO2, but these were still oxic (6 kPa). During night, rhizome pO2 levels for plants with leaf gas films declined to 5 kPa, but without leaf gas films pO2 in the rhizomes dropped to 1.4 kPa. Whether this drop in pO2 has adverse effects on growth and cell functioning is uncertain but the lower pO2 in rhizomes would have resulted in even lower pO2 in the roots (further from O2 source), as demonstrated in the laboratory experiments during dark (Fig. 3b). Our data showed that leaf gas films were critical for the supply of O2 to rhizomes and roots during complete submergence, especially at night when no light is available for photosynthesis.
Underwater photosynthesis allows S. anglica to maintain roots and rhizomes with a supply of O2 when the shoots are completely submerged. While underwater photosynthesis was only 4% of photosynthesis in air (Table 3; Mallott et al. 1975), the O2 produced elevates pO2 of roots and rhizomes above levels when submerged in the dark. In situ, we found that even if the water was muddy and light attenuation was high (Fig. 1), underwater photosynthesis was still sufficient to raise rhizome pO2 above levels when in darkness (Fig. 2). The decline in pO2 after the initial rise (Fig. 1) could be explained by a depletion of internal CO2 until a new steady state is achieved between CO2 supply and consumption in photosynthesis. The production of O2 was insufficient to maintain rhizome pO2 at a level similar to those prior to submergence, presumably because of constraints on CO2 uptake from the water that had between 5 and 26 µm CO2. Light attenuation was likely of less importance than restricted CO2 supply as light availability was above 1500 µmol photons m−2 s−1 at the sediment surface prior to inundation, after which it fell to around 300 µmol photons m−2 s−1 (Fig. 1).
Leaf gas films enhance underwater O2 uptake during dark
Leaf gas films can enhance dark uptake of O2, and particularly at low O2 availability (Colmer & Pedersen 2008b). In the present study at air equilibrium O2, leaf gas films did not benefit underwater dark respiration by leaf segments of S. anglica (Table 3). During the in situ measurements, water O2 concentrations varied at night between 213 and 228 µm O2 (15.6–16.6 kPa O2), which is 72–77% of air equilibrium. Colmer & Pedersen (2008b) have shown that leaf segments of P. australis are capable of sustaining underwater O2 uptake down to external O2 concentrations of 60 µm if the leaves have intact gas films; below that concentration, diffusion limitations restrict O2 uptake. When the gas films were removed from P. australis, underwater O2 uptake was diffusion limited even at air saturation (20 °C = 284 µM O2). This was not the case for S. anglica; O2 uptake by leaf segments with or without gas films was not O2 limited when in water near air equilibrium. Gas films on leaves of S. anglica were, however, important for root and rhizome aeration during submergence. Whole plant dark O2 uptake was not measured. However, as leaf gas films enhanced root and rhizome pO2 during dark, this demonstrates that leaf gas films alleviate a bottleneck that exists for O2 entry via leaves.
The mechanism by which the leaf gas films function has been described as an enlargement of the water-gas interface (cf. plastrons of insects, Hebets & Chapman 2000), but the possibility of stomata to remain open under water and thereby to continue to function as a conduit for gas distribution from the gas films to the inside of the plant has also been put forward (Colmer & Pedersen 2008b). Scanning electron micrograph images show that the abaxial side of S. anglica leaves is much smoother and has very few stomata compared with the heavily ridged adaxial side with more than 99% of the stomata (Fig. 4, Table 1, Maricle et al. 2009). This coincides with leaf gas films only being present on the adaxial side of the leaves. While this is speculative, for S. anglica to have almost all stomata located on the side that has a gas film when under water, could be of adaptative value to submergence tolerance.
Leaf gas films enhance rhizome pO2 during tidal inundation in natural stands of S. anglica. Enhanced underwater photosynthesis because of leaf gas films not only elevates internal pO2, but the gas film also facilitates O2 entry during submergence in darkness. In addition to the leaf gas films, large volumes of aerenchyma in sheaths, rhizomes and roots are adaptive traits possessed by S. anglica to enhance internal aeration and thus growth despite frequent inundation in lower salt marshes.
This work was supported by The Danish Council for Independent Research grant no. 09–072482 and by a University of Western Australia International Postgraduate Research Scholarship to Anders Winkel. We thank Mette Cristine Schou Frandsen for the excellent scanning electron micrograph pictures and Jens Borum for useful advice and constructive comments on the manuscript. We also thank the referees for useful criticisms and especially one of the referees for very valuable contributions to improve the final version of the manuscript.