Ethene (ethylene) production in the marine macroalga Ulva (Enteromorpha) intestinalis L. (Chlorophyta, Ulvophyceae): effect of light-stress and co-production with dimethyl sulphide

Authors

  • INA PLETTNER,

    Corresponding author
    1. School of Environmental Sciences, University of East Anglia, Norwich NR4 7TJ, United Kingdom
      Ina Plettner, (current address) University of Bremen, Marine Botany – FB 2, Postfach 330440, 28334 Bremen, Germany. E-mail: plettner@uni-bremen.de
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  • MICHAEL STEINKE,

    1. School of Environmental Sciences, University of East Anglia, Norwich NR4 7TJ, United Kingdom
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  • GILL MALIN

    1. School of Environmental Sciences, University of East Anglia, Norwich NR4 7TJ, United Kingdom
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Ina Plettner, (current address) University of Bremen, Marine Botany – FB 2, Postfach 330440, 28334 Bremen, Germany. E-mail: plettner@uni-bremen.de

ABSTRACT

Ethene (ethylene; H2C = CH2) is one of a range of non-methane hydrocarbons (NMHC) that affect atmospheric chemistry and global climate. Ethene acts as a hormone in higher plants and its role in plant biochemistry, physiology and ecology has been the subject of extensive research. Ethene is also found in seawater, but despite evidence that marine microalgae and seaweeds can produce ethene directly, its production is generally attributed to photochemical breakdown of dissolved organic matter. Here we confirmed ethene production in cultured samples of the macroalga Ulva (Enteromorpha) intestinalis. Ethene levels increased substantially when samples acclimatized to low light conditions were transferred to high light, and ethene addition reduced chlorophyll levels by 30%. A range of potential inhibitors and inducers of ethene biosynthesis were tested. Evidence was found for ethene synthesis via the 1-aminocylopropane-1-acrylic acid (ACC) pathway and ACC oxidase activity was confirmed for cell-free extracts. Addition of acrylate, a potential ethene precursor in algae that contain the compatible solute dimethylsulphoniopropionate, doubled the ethene produced but no acrylate decarboxylase activity was found. Nonetheless the data support active production of ethene and we suggest ethene may play a multifaceted role in algae as it does in higher plants.

Abbreviations
ACC

1-aminocylopropane-1-carboxylic acid

ADC

acrylate decarboxylase

AdoMet

S-(5′-adenosyl)- l-methionine chloride

AVG

l-alpha-[2-(2-aminoethoxy) vinyl] glycine hydrochloride

DMS

dimethyl sulphide

DMSP

dimethylsulphoniopropionate

FW

fresh weight

IAA

indole-3-acetic acid

KMBA

2-keto-4-(methylthio)butyric acid

NMHC

non-methane hydrocarbons

ROS

reactive oxygen species.

INTRODUCTION

Biogenic non-methane hydrocarbons (NMHC) have attracted research attention due to their significant contribution to the chemistry of the atmosphere. Ethene [ethylene (H2C = CH2)] is a reactive NMHC that provides a sink for hydroxyl radicals and plays a key role in the production and destruction of ozone in the troposphere (Donahue & Prinn 1990). This trace gas is well known as a hormone in higher plants (Fluhr & Mattoo 1996; Bleecker & Kende 2000; Alonso & Stepanova 2004), where the different stages of development are controlled by a range of growth regulators, for example, the five well-described growth substances abscissic acid, auxin, cytokinin, gibberellin and ethene all of which have a multiplicity of effects. Notable effects of ethene on plants include ripening of fruits, inhibition of stem and root elongation, promotion of seed germination and flowering, senescence of leaves and flowers, and sex determination (e.g. Bleecker & Kende 2000). Physiological reactions to light-stress also involve the production of ethene and it is possible that reactive oxygen species (ROS) act as secondary messengers initiating a signal cascade which can stimulate ethene synthesis (Mackerness 2000). Figure 1a shows the dominant metabolic pathway for ethene biosynthesis in higher plants that involves conversion of methionine to 1-aminocylopropane-1-carboxylic acid (ACC) via the enzyme ACC synthase and the ACC oxidase catalysis of ACC to ethene (Fluhr & Mattoo 1996; Stella, Wouters & Baldellon 1996).

Figure 1.

Possible pathways of ethene production in (a) higher plants and (b) marine algae. Numbers indicate key enzymes involved in ethene synthesis: (1) ACC synthase (Stella et al. 1996) (2) ACC oxidase (Stella et al. 1996) (3) methionine transaminase (Gage et al. 1997) (4) DMSP lyase (Steinke & Kirst 1996) (5) acrylate decarboxylase (Ghooprasert & Spencer 1975). Reactions (4) and (5) suggest a direct link between the production of DMS and ethene.

Several studies have found concentrations of dissolved ethene exceeding atmospheric equilibrium in marine surface waters, and hence the ocean is considered to be a natural source for atmospheric ethene (Seifert et al. 1999). However, the global ocean data set for ethene is relatively small and the geographical and seasonal coverage is poor. Plass-Dülmer et al. (1995) included most of the earlier published ethene data and calculated an average seawater concentration of 134 p m and an annual emission in the range of 0.89–2.17 Tg year−1. Photochemical breakdown of dissolved organic matter is often suggested to be the main pathway for marine ethene production. However, field and laboratory studies suggest marine microalgae and seaweeds as a possible source of ethene (Broadgate, Liss & Penkett 1997; Seifert et al. 1999; Broadgate et al. 2004). Emissions from a temperate coastal site were intermediate in magnitude between oceanic and terrestrial fluxes (Broadgate et al. 2004), indicating that macroalgal populations may significantly contribute to the NMHC concentration in coastal air. ACC-dependent production of ethene was documented for the marine chlorophyte Acetabularia mediterranea (Vanden Driessche, Petiau-de Vries & Guisset 1997) and the freshwater green alga Haematococcus pluvialis (Maillard, Thepenier & Gudin 1993). However, cofactor requirements in the latter species were different to those of higher plants, suggesting that additional alternative pathways are likely to exist in algae. Bacteria and fungi can use 2-keto-4-(methylthio)butyric acid (KMBA) or 2-oxoglutarate as immediate precursors for ethene (Fukuda, Ogawa & Tanase 1993) and some early branching embryophytes (bryophytes, pteridophytes and gymnosperms) release ethene via an unknown metabolic pathway and that is independent of ACC or 2-oxoglutarate (Osborne et al. 1996).

Figure 1b illustrates another possible alternative route for the production of ethene from acrylate [H2C = CH-COO] in marine algae. Acrylate has been suggested as a precursor of ethene (Abeles 1973; Watanabe & Kondo 1976) and can be cleaved from the secondary metabolite β-dimethylsulphoniopropionate [DMSP (CH3)2S+CH2CH2COO)] in an enzymatic reaction that also produces the volatile trace gas dimethyl sulphide (DMS [(CH3)2S]). Activity of this enzyme, DMSP lyase (enzyme classification number 4.4.1.3), has been demonstrated in several algal taxa including benthic macroalgae and pelagic phytoplankton (Steinke & Kirst 1996; Steinke, Wolfe & Kirst 1998). DMSP is an important metabolite in many marine algae and can contribute 48–100% of sulphur fluxes and 5–15% of carbon fluxes in marine microbial ecosystems (Simóet al. 2002). The biosynthesis of acrylate in marine organisms has attracted little research attention but the potential for its production by DMSP-containing algal taxa is high. DMSP is produced from methionine (Gage et al. 1997), the same amino acid that ultimately produces ethene in higher plants. Among the benthic algae, red and green macroalgae (rhodophytes and chlorophytes) are important producers of DMSP and DMS (e.g. Blunden et al. 1992), and the same algal taxa are also known to release ethene. Broadgate et al. (2004) tested 10 species of seaweed from rocky shores in western Ireland; all produced ethene, but, the highest ethene production rates were found in the chlorophyte Ulva intestinalis (62.93 pmol g−1 dry weight h−1), a species that uses DMSP as its principal osmolyte (approximately 25 mmol kg−1; Edwards et al. 1987). Watanabe & Kondo (1976) conducted experiments with the green alga Codium latum, the red algae Porphyra tenera and P. aborescens, and several brown algae (phaeophytes). Only the red and green algae showed ethene synthesis and its production was increased by exogenous addition of the auxin indole-3-acetic acid (IAA), and increased further with acrylate. Porphyra perforata has also been shown to produce ethene (Zhang, Yamane & Chapman 1993) and the acellular macroscopic chlorophyte A. mediterranea uses ethene for developmental differentiation (Vanden Driessche et al. 1997). The production of ethene from acrylate involves the equimolar release of CO2 and requires the presence of acrylate decarboxylase, an enzyme previously purified from the wax bean Phaseolus vulgaris (Ghooprasert & Spencer 1975) and yeast (Shimokawa & Kasai 1970). Provided that this enzyme is present in marine algae, the production of the climate relevant trace gases DMS and ethene could be linked in marine algae, as outlined in Fig. 1b.

In the present study we investigated ethene production in the green macroalgae Ulva intestinalis (formerly known as Enteromorpha intestinalis; see Hayden et al. 2003). Specifically, we wanted to:

  • 1confirm production of ethene in unialgal laboratory cultures of U. intestinalis;
  • 2investigate whether production of ethene involves the ACC pathway;
  • 3examine the influence of light intensity on ethene release; and
  • 4test how ethene may affect chlorophyll a concentrations.

Experiments were carried out to quantify ethene release during growth or when thalli were exposed to high light conditions. Potential inhibitors and inducers were used to study possible routes of ethene synthesis in U. intestinalis. We hypothesized that acrylate may be a direct precursor of ethene in this species and conducted measurements of DMS in tandem with our ethene analyses to look for evidence for the coproduction of these two volatiles.

MATERIALS AND METHODS

Macroalgal cultures

Ulva intestinalis was kindly provided by C. Wiencke (strain 1089, isolated in Greenland in 1990; Alfred-Wegener-Institute for Polar and Marine Research, Bremerhaven, Germany). Standard cultivation of algal thalli was at 12 °C in 0.5 L borosilicate bottles filled with 450 mL culture medium. This medium consisted of aged, 0.2 µm filtered and autoclaved North Sea seawater enriched with sterile f/2 nutrients and vitamins after autoclaving (Guillard 1975). Light was supplied by fluorescent tubes at an intensity of 25 µmol photons m−2 s−1 (quantum radiometer photometer with a flat cosine irradiance sensor; Ramsden Scientific Instrument Co., United Kingdom) and a light : dark cycle of 14 : 10 h. These growth conditions provided sufficient algal growth for our experiments and minimized sporulation. All glass-ware was acid rinsed and autoclaved. Depending on algal growth, thalli were transferred into fresh culture medium every 7–14 d using sterile techniques. Microscopic checks confirmed that only few bacteria were present and thalli of similar size, shape and age were used for the experiments.

Growth experiments

Five gas-tight medical flat bottles (110 mL volume) were filled with medium leaving minimal headspace. Approximately 240 mg of algal thallus was added to three of the bottles and then all five bottles were incubated for a total of 6–7 d. During this period, samples were taken every 2–3 d for ethene measurements and algal thalli were carefully dried on tissue paper to remove water before determining the fresh weight. To avoid nutrient limitation, algae were kept in semi-continuous culture and transferred into new bottles filled with fresh medium for the next incubation period of 2–3 d. Rates of ethene production in three replicate bottles with algae were calculated by adding individual measurements and subtracting abiotic ethene production in control bottles without algae (two replicates). Growth conditions were similar to those used during normal cultivation of U. intestinalis but we added 2.38 m m NaHCO3 as additional carbon source and 3 m m HEPES for further pH-buffering (average pH after one 2–3 d incubation period was 8.7).

Light experiments

Macroalgae from higher latitudes (U. intestinalis strain 1089 was isolated at Disko Island, Greenland) have very low light requirements for compensation (e.g. 0.3–2.5 µmol m−2 s−1; Markager & Sand-Jensen 1992) and can survive substantial periods in complete darkness (Karsten, Wiencke & Kirst 1992). To maximize the light-stress U. intestinalis was acclimatized to low light conditions (0.5 µmol m−2 s−1) using black mesh 14 d before the start of the experiment. Continuous light was used to eliminate the possible effects of day–night cycles on ethene production. Incubation bottles and media were the same as for the growth experiments. Half of the bottles were kept at low light, the others were transferred into continuous high light at 120 µmol photons m−2 s−1 from the start of the experiment. Bottles without algae were incubated at low and high light intensities and used to quantify abiotic production of ethene and DMS. At each time-point a series of low and high light bottles were sacrificed so that each data-point represents a measurement from one individual bottle.

A size-fractionation was used to check for the possible contribution of abiotic processes and bacterial activity to total ethene production in this experiment. Ulva intestinalis incubated at high light for 48 h was carefully removed from the medium. Immediately afterwards the medium was filtered with low vacuum (< 13.3 kPa) using glass fibre filters (1.2 µm pore size; GF/C, Whatman International, Maidstone, UK) in a first filtration step and cellulose nitrate filters (0.2 µm pore size; Sartorius Ltd., Surrey, UK) as the second filtration step. Most bacteria will pass through the 1.2 µm filter but will be retained on the 0.2 µm filter; therefore the 1.2 µm filtrates included dissolved organic components from the U. intestinalis growth medium plus bacteria, whereas 0.2 µm filtered medium should contain only dissolved components. For technical reasons, the incubation times for the filtrates at high light conditions were different: 0.2 µm filtrates had an incubation time of 24 h and the 1.2 µm filtrates were incubated for 60 h. Initial samples were measured directly after filtration and the data were corrected for ethene concentrations in control bottles with fresh medium.

Experiments with inhibitor and inducer compounds

The following compounds have been used in experiments with higher plants and were tested as possible inhibitors or inducers for ethene production in three separate tests with U. intestinalis (see Table 1 for expected response): l-alpha-[2-(2-aminoethoxy) vinyl] glycine hydrochloride (AVG), indole-3-acetic acid (IAA), S-(5′-adenosyl)- l-methionine chloride (AdoMet), 1-aminocyclopropane-1-carboxylic acid (ACC), acrylate, dimethylsulphoniopropionate (DMSP) and 2-keto-4-(methylthio)butyric acid sodium salt (KMBA). Stock solutions were prepared in distilled water, with the exception of IAA which was prepared in 90% acetone. Acetone (0.1% final concentration) was also added to the medium for the IAA addition controls. Tests indicated that 0.1% acetone addition does not affect abiotic or biotic ethene production (data not shown). Compounds were added to the algae in vivo before 30–48 h incubation of bottles. However, in our AVG addition experiment incubation time had to be extended to 4 d due to technical problems. We selected final concentrations of compounds that others found effective for higher plants (see Table 1). To minimize abiotic production of ethene, incubations were carried out in the dark using the same bottles and medium as in our growth experiments. Three series of control treatments were included and consisted of medium, medium plus tested compound and medium plus algae. Rates of ethene production in these controls were subtracted from the relevant treatments.

Table 1.  Effect of inhibitors, inducers and possible precursors on ethene production in Ulva intestinalis
 CompoundExpected responseConcentration
(µm)
nEthene production in
comparison to untreated control
  1. See methods for full details of the compounds. Shown are concentrations used and relative ethene production in comparison to untreated controls (n = 2 for each experiment) with ranges in parentheses.

Experiment 1AVGInhibition of ACC synthase activity  503 0.6 (0.49–0.64)
Experiment 2IAAPromotion of ACC synthase activity  502 2.1 (1.40–2.75)
IAAPromotion of ACC synthase activity10002 3.1 (2.88–3.27)
ACCPromotion of ACC oxidase activity 2004  33 (20–47)
DMSPPrecursor of acrylate via DMSP lyase activity 2002 1.0 (0.57–1.26)
KMBAPrecursor of bacterial pathway for ethene production 2004 1.0 (0.56–1.50)
Experiment 3AcrylatePromotion of acrylate decarboxylase activity  503 2.0 (1.31–2.71)

Ethene incubations

To investigate the effect of ethene on levels of chl a, algal thalli were transferred into glass vials (60 mL volume) with 1.5 mL medium. A separate small vial (2 mL volume) containing either 100 µL distilled water (controls) or a 0.2- m stock solution of ethephone (2-chloro-ethyl phosphonic acid) in 0.1% HCl (treatments), was carefully placed into the larger vials with medium and U. intestinalis. At the start of the experiments, 1 drop of 10 N NaOH was added to the small vial immediately before closing the large vial with a Teflon-coated silicone septum and a crimp-seal. At high pH, ethephone rapidly cleaves into equimolar concentrations of ethene, phosphate and chloride (Tseng, Chang & Chou 2000). Vials were incubated for 3 d at 12 °C and 25 µmol photons m−2 s−1. At the end of the experiment the ethene concentrations in the vials were measured using headspace analysis. The average concentration in the gas phase was 7 ppt (n = 10) which corresponds to an aqueous concentration of approximately 40 µm. At the end of the incubations, algae were removed from the vials, weighed and snap-frozen in liquid N2 for subsequent chlorophyll analysis.

Chlorophyll analysis

The frozen thalli were ground to fine powder in 1.5 mL reaction tubes using a micropestle and liquid N2. The powder was transferred into glass vials, 5 mL ice-cold 90% acetone was added and the samples incubated overnight at −15 °C in the dark. Chlorophylls a and b were determined with a Lambda 35 UV/VIS Spectrometer [Perkin Elmer LAS (formerly Instruments), Bucks., UK] after Jeffrey & Humphrey (1975).

Enzyme assays

Ethene measurements in combination with a range of ACC concentrations (0–2 m m final concentrations) were used to investigate in vitro and in vivo ACC oxidase activity according to the methods described in Jung et al. (2000) and Okumura et al. (1999), respectively. In vitro ADC activity was tested after Ghooprasert & Spencer (1975) using 0–10 m m acrylate as substrate. A temperature of 20 °C was used for the in vitro assays and in vivo activity was tested at 20 and 30 °C to check for temperature dependency of the reaction. For the in vitro assays, fresh thalli were carefully dried with tissue paper, snap-frozen with liquid N2 and immediately ground to fine powder using a mortar and pestle. Sand was added to aid homogenization. Extraction buffer consisting of 100 m m MES pH 6.5, 10 m m MgCl2 · 6 H2O, 0.5% sucrose, 5% glycerol, 30 m m sodium ascorbate and 0.25% Triton X-100 was added and the extract incubated on ice for 15 min. Cell debris was removed by centrifugation at 3000 × g and 4 °C for 30 min. In all cases the controls consisted of medium (in vivo) or buffer (in vitro) only, medium or buffer plus ACC, medium or buffer plus U. intestinalis (n = 3 in all control treatments).

Ethene measurements

Samples for headspace measurements of ethene were taken with a gas-tight syringe (250 µL) and on-column injection was carried out via the septum port of a gas chromatograph equipped with a flame ionization detector (GC-FID; GC-2010; Shimadzu UK, Milton Keynes, UK). Water samples were taken using Teflon tubing and gas-tight glass syringes (100 mL volume). Cryo-focusing of ethene was carried out in a purge and trap system similar to the one described in Broadgate et al. (1997). In brief, 80 mL volume samples were purged with oxygen-free nitrogen at 60 mL min−1 for 30 min and trace gases were cryogenically preconcentrated in a 1/8 inch (approximately 3.2 mm od) stainless steel trap filled with 80-mesh glass beads that was held at −185 °C. The gases were released from the trap by rapid heating to > 90 °C and swept into the gas chromatograph using CP-grade helium (BOC, Guildford, UK) as the carrier. For technical reasons two columns and different methods were used during this study. First, for the growth experiments, a CarboBOND fused silica column (50 m × 0.53 mm; DF 5 µm; Varian UK, Walton-on-Thames, UK) was used with injector and detector temperatures set to 150 and 250 °C, respectively. The helium make-up flow and the flame gases air and hydrogen were supplied at 30, 400 and 47 mL  min−1. Oven temperature was 120 °C for 5 min and then increased to 230 °C at a rate of 50 °C min−1 The final temperature was held for 5 min. Second, all other measurements were conducted using an Al2O3-plot fused silica column (50 m × 0.53 mm; Varian UK) and an oven temperature program starting at 33 °C for 5 min followed by a 50 °C min−1 increase to 200 °C that was held for 12 min. Injector and detector were kept at 250 °C and gas-flows were the same as above. In both cases calibrations were conducted with commercial standard gas-mixtures (98 ppm ethene in helium) and a series of either 3–100 µL (direct on-column injections) or 3–1000 µL (purge and trap measurements) of gas was used.

DMS measurements

Pre-concentrating of DMS and subsequent analysis was carried out using a purge-and-trap system coupled to a gas chromatograph with flame-photometric detection (GC-FPD; GC-2010, Shimadzu UK). Water samples of 3 mL volume were purged with N2 at 60 mL min−1 before cryofocusing DMS in a cold-trap made from 1/8 inch (3.2 mm od) Teflon tubing at −150 °C. Details of this method can be found in Turner et al. (1990). Rapid heating with hot water (> 90 °C) was used to desorb DMS while flushing it into the carrier gas flow (He at 2.3 mL min−1) and onto a capillary column (CP SIL 5CB, 50 m × 0.53 mm, Varian UK). The oven temperature was 80 °C, and the injector and detector were kept at 250 °C with flame gases air and H2 supplied at 70 and 60 mL min−1, respectively. Stock solutions (2.5 and 25 n m) of commercial DMSP [Centre for Analysis, Spectroscopy and Synthesis (CASS), University of Groningen Chemical Laboratories, The Netherlands] were hydrolysed to DMS with 10 m NaOH (final concentration 250 m m) and used to calibrate the instrument.

RESULTS AND DISCUSSION

Ethene is well recognized as a hormone in higher plants but very little is known about its production by marine algae or possible role as an infochemical in aquatic signalling. In this study we investigated the production of ethene by the marine chlorophyte U. intestinalis during normal growth and under high light-stress, conducted assays to study possible pathways of ethene biosynthesis and studied the effect of ethene on chlorophyll a concentrations.

Ethene production during growth

Individual thalli of U. intestinalis showed specific growth rates (µ) of 0.03, 0.07 and 0.08 d−1 and doubling times (td) of 26, 10 and 8 d, respectively (Fig. 2a). In comparison with the medium-only controls, ethene concentrations were substantially higher when algae were present (Fig. 2b). The ethene production rate in the control is probably the result of photochemical breakdown of dissolved organic carbon and was linear at 4.4 pmol ethene L−1 h−1. Subtraction of this control rate from the treatments with algae suggests production rates of 6.4–10.6 pmol ethene L−1 h−1 or 2.3–2.8 pmol ethene g−1 fresh weight (FW) h−1 in U. intestinalis.

Figure 2.

Ethene production in Ulva intestinalis during semicontinuous culture. (a) Fresh weight and growth parameters of three algal thalli. (b) Sum of ethene concentrations in bottles with seawater medium (control; range of data indicated by error bars) and algae. (c) Sum of ethene concentrations normalized to algal fresh weight.

Light-induced production of ethene and DMS

Irradiance levels affected both ethene and DMS production in U. intestinalis. Thalli acclimatized to low light conditions (0.5 µmol photons m−2 s−1) showed an increase in ethene and DMS production when exposed to high light conditions (120 µmol photons m−2 s−1; Fig. 3). Ethene release into the medium was linear over the first 6 h of the experiment and the production rate observed in high light conditions was more than six times higher (96.6 pmol ethene g−1 FW h−1) than the low light control (14.5 pmol ethene g−1 FW h−1; Fig. 3a). Ethene concentrations in the high light treatment increased to 893 pmol g−1 FW after a further 24 h but then decreased to 509 pmol g−1 FW. In comparison, the production rate for DMS under high light was linear for the whole 24 h incubation period (1797 pmol DMS g−1 FW h−1) and there was no clear trend in DMS production at low irradiance (Fig. 3b). As a result, the ratio of DMS-to-ethene production was similar and indicated a co-production of ethene and DMS in both light treatments over the first 24 h (range: 14–36 in low light, 21–46 in high light; Fig. 3c) but this ratio diverged after 24 h. At low irradiances the DMS-to-ethene ratio remained similar to the earlier values at 19, whereas in high light and with reduced ethene production the ratio increased to 167.

Figure 3.

Light-induced production of (a) ethene and (b) DMS, and (c) DMS : ethene ratio in Ulva intestinalis. Filled symbols show production in low light-adapted controls (0.5 µmol m−2 s−1) in contrast to thalli that were exposed to high light (120 µmol m−2 s−1) at 0 h (open symbols).

In higher plants, light-induced production of ROS has a role as secondary messenger acting up-stream of pathways involving ethene (Mackerness 2000). It is therefore possible that light-stress and ethene release are interconnected and part of the physiological adaptation of algae to high light intensities. To be an effective infochemical, ethene is not required at elevated concentrations for an extended time, rather, signalling compounds such as ethene mediate and accelerate a sequence of events that result in corresponding physiological changes such as chlorophyll degradation (Pandey et al. 2000). In contrast to this, DMS concentration continued to increase after 24 h in high light. Recently, DMS was linked to photo-oxidative stress reactions in marine algae: Sunda et al. (2002) showed that DMSP and its breakdown products (DMS, acrylate, dimethylsulphoxide, and methane sulphinic acid) readily scavenge harmful hydroxyl radicals in vitro, and thus may serve as an antioxidant system, regulated in part by enzymatic cleavage of DMSP, in vivo. Our results on light-dependent DMS production support this hypothesis.

The ethene production rates reported here ranged from 2.3 to 96.6 pmol ethene g−1 FW h−1 and are similar to previous findings. Broadgate et al. (2004) reported production rates of 62.93 pmol ethene g−1 dry weight for freshly collected U. intestinalis. This rate is equivalent to 20.98 pmol ethene g−1 FW when assuming a dry to fresh weight ratio of 1 : 3 (Karsten et al. 1994). Additionally, ethene production rates in this seaweed are comparable to findings for higher plants such as sunflowers (Helianthus annuus; approximately 20–60 pmol g−1 FW h−1; Kathiresan et al.  1997)  or  cleavers  (Galium  aparine;  59 pmol g−1 FW  h−1; Hansen & Grossmann 2000). This suggests that marine seaweeds may significantly add to atmospheric ethene concentrations in coastal air.

Effect of potential inhibitors and inducers

As mentioned earlier, ACC synthase and ACC oxidase are key enzymes of ethene synthesis in higher plants but acrylate may provide a link between DMS and ethene production in DMSP-producing algae. Here we found that exogenous addition of potential inhibitors and inducers affected ethene production rates in U. intestinalis. IAA, ACC and acrylate stimulated and AVG inhibited ethene release. DMSP, the precursor of acrylate in some marine algae, and KMBA, the intermediate in bacterial ethene production, appeared not to affect ethene production in our experiments (Table 1). The ACC oxidase enzyme converts ACC into ethene and addition of 200 µm ACC significantly stimulated ethene release with a 33-fold increase. The inhibitor AVG and inducer IAA affect the ACC synthase enzyme in the ACC-pathway (Fig. 1). Addition of 50 and 1000 µm IAA additions increased ethene production two- to three-fold whereas 50 µm AVG resulted in a 45% reduction in ethene production. We note that AVG inhibition of ACC synthase is incomplete in some higher plants (e.g. Hansen & Grossmann 2000), however, our experiments with potential inhibitors and inducers suggest that alternative pathways for ethene production in marine algae remain a possibility.

Abiotic production from photochemical processes (summarized by Riemer et al. 2000) and bacterial activity (Fukuda et al. 1993) can add to ethene concentrations in the field and may have contributed to ethene production in our experiments. However, in comparison with the release of ethene by U. intestinalis in low light and high light (14.5 and 96.9 pmol ethene g−1 FW h−1, respectively; Fig. 3) incubation of 0.2 and 1.2 µm filtrates of algal culture indicated little production of ethene when algae were absent (0.15–0.83 pmol ethene g−1 FW h−1; Table 2). Our inducer experiments with KMBA (the precursor of ethene production in bacteria; Table 1) provide further evidence that bacterial production of ethene was small in our tests. However, the physiology and morphology of Ulva sp. can radically alter under the presence of bacteria (Dobretsov & Qian 2002; Hayden et al. 2003) and separating the effect of bacteria on ethene production would be extremely difficult. In marine ecosystems ethene is probably a result of several biotic and abiotic processes and interactions combined, and their relative importance for total ethene production may alter under different conditions.

Table 2.  Ethene production in medium preconditioned with Ulva intestinalis in high light for 48 h and size-fractionated to contain dissolved compounds (0.2 µm filtration) or dissolved compounds plus bacteria (1.2 µm filtration)
TreatmentPreconditioningnEthene production
(pmol g−1 FW h−1)
  1. The ethene production rate derived from the initial slope of our light experiment shown in Fig. 3 is provided for comparison.

0.2 µm filtrate – low lightHigh light2 0.18
0.2 µm filtrate – high lightHigh light2 0.15
1.2 µm filtrate – high lightHigh light2 0.83
Ulva intestinalisLow light314.5
Ulva intestinalisHigh light396.6

Enzyme assays

Experiments conducted with cell-free extracts of U. intestinalis indicated that acrylate decarboxylase, the enzyme required for the production of ethene from acrylate, is absent or inactive under the conditions used (assay after Ghooprasert & Spencer 1975; Table 3). In contrast, in vitro and in vivo assays suggested the presence of ACC oxidase. Enzyme activity was proportional to the amount of protein in the in vitro assay and showed an average activity of 6.8 ± 0.7 pmol ethene g−1 FW h−1 at 20 °C. This level of activity is in agreement with our in vivo assays for ACC oxidase where we found production rates of 8.6 and 28.5 pmol ethene g−1 FW h−1 at 20 and 30 °C, respectively. The enzyme tests were conducted with thalli cultivated under standard conditions. It is possible that enzyme expression and activity may have been different had the algae been tested using high-light conditions where we found highest ethene production rates (see Figs 2 & 3). Direct evidence for the existence of the acrylate-dependent alternative pathway for ethene biosynthesis in algae is still lacking and our experiments suggest that the majority of ethene is produced via an algal ACC pathway.

Table 3.  Activities of acrylate decarboxylase (ADC) and 1-aminocylopropane-1-carboxylic acid (ACC) oxidase in cell-free extracts (in vitro) and in vivo incubations with Ulva intestinalis
EnzymeFW U. intestinalis
(mg)
Protein in assay
(mg)
nEnzyme activity ± SD
(pmol ethene g−1 FW h−1)
  1. Shown are fresh weight (FW) equivalents and protein concentrations used in the assays, number of replicates and enzyme activities. Results indicate the existence of ACC oxidase, but absence of ADC (ND, not determined; ncontrol = 3)

ADC in vitro130–1901.32–3.314not detected
ACC oxidase in vitro 300.3347.0 ± 0.5
 600.6676.8 ± 0.8
(average = 6.8 ± 0.7)
ACC oxidase in vivo170 ± 20ND38.6 at 20 °C
28.5 at 30 °C

Effects of ethene on algal physiology

Ethene plays a central role as a hormone for growth and development of higher plants. For example, ripening usually starts in one region of a fruit and spreads to the other regions that lag behind. This process is initiated, accelerated and integrated by ethene that diffuses from cell to cell throughout the fruit (Bleecker & Kende 2000). This may also apply to differentiation in algae. Circadian rhythms were identified for ethene production, its retention in the cell wall and ACC conversion to ethene in A. mediterranea (Vanden Driessche et al. 1997). Ethephone additions (≥ 10 µm) affected cap formation in this species and could be stimulatory or inhibitory depending on the timing of application (Vanden Driessche et al. 1988). In our experiments with U. intestinalis we used ethephone to investigate the effect of exogenous ethene on the concentrations of chlorophyll a. A 3 d incubation of U. intestinalis in 40 µm ethene resulted in 30% loss of chlorophyll a in comparison to untreated controls. This may suggest that ethene can have a physiological role in marine algae and could indicate the presence of ethene receptors and possible signal transduction pathways that affect chlorophyll a concentrations in U. intestinalis. Further experiments will be needed to investigate the physiological role of ethene in seaweeds more fully.

Ethene in aquatic signalling

Ethene is involved in neighbour sensing in tobacco plants (Pierik et al. 2004) and similar mechanisms could help macroalgae to adjust their physiology and/or morphology to environmental conditions. For example, it is possible that the synchronization of growth, morphogenesis, and gamete production and release involves ethene signalling. Despite its relatively high diffusivity, the diffusion of ethene through water is 10 000-times slower than through air (Bleecker & Kende 2000). Hence, ethene gradients can be established resulting in locally increased ethene concentrations that may be high enough to affect physiological processes in other parts of the alga or in organisms close by.

Little is known about signalling in aquatic environments and the field of pelagic infochemistry is still in its infancy. Biogenic trace gases may be involved in trophic interactions (for review see Steinke, Malin & Liss 2002) but the utility of ethene as a signal depends on the ability of cells to monitor the changing concentrations of this gas and to transmit this information onto appropriate physiological or behavioural responses (e.g. Bleecker & Kende 2000). Our experiments with ethephone indicate that ethene elicits a physiological response (chlorophyll a degradation) in U. intestinalis. This response may not be exclusive to macroalgae and ethene signalling could have a role in phytoplankton as well. However, receptors for ethene have not yet been found in marine algae. The structure and role of six different putative ethene receptors (ETR1, ETR2, EIN4, ERS1, ERS2 and NR) are known from higher plants (e.g. Hua et al. 1998). The open reading frame slr1212 from the genome of the freshwater cyanobacterium Synechocystis sp. (strain PCC 6803) shows homology to the ethene-binding domain of ETR1 and encodes a functional ethene-binding protein (Rodriguez et al. 1999). It has been suggested that Synechocystis, Gloeotrichia and Nostoc also release ethene (May 2001) but the biological function of ethene in these species is unexplained and the production of ethene in cyanobacteria requires further investigations. Annotation of the Thalassiosira pseudonana (Bacillariophyceae) genome has revealed that diatoms may also utilize ethene for regulating stress responses and putative diatom genes encoding ACC synthase, the rate-limiting step in ethene biosynthesis in plants, have been found (Dr Chris Bowler and Dr Assaf Vardi, personal communication). Future efforts could use molecular approaches in combination with physiological experiments to elucidate the role of ethene in aquatic signalling.

CONCLUSIONS

Our results with U. intestinalis add to the evidence from the freshwater chlorophyte Haematococcus pluvialis (Maillard et al. (1993) and the marine chlorophyte A. mediterranea (Vanden Driessche et al. 1988) that ethene is produced via the ACC-pathway in several chlorophyte taxa. This is contrary to the suggestion that this pathway arose late in plant evolution (Osborne et al. 1996). We propose that ethene production from ACC may be common among marine and freshwater algae and suggest that this plant hormone is an important infochemical in aquatic environments. It is likely that algae use similar complex signalling pathways that are based on hormones or hormone-like substances comparable to the infochemicals in higher plants. Cytokinins have recently been identified in several species of macroalgae (Stirk et al. 2003) and green microalgae (Ördög et al. 2004) and these hormone-like compounds provide further evidence on the role of infochemicals in algal physiology.

ACKNOWLEDGMENTS

Christian Wiencke kindly provided a culture of Ulva intestinalis and Jamie Kettle helped with the calculations of ethene solubility in seawater. We thank Anthony Trewavas and two anonymous referees for their comments on an earlier version. Financial support was provided by the UK Natural Environment Research Council (NERC contracts NER/I/S/2000/00897 and NE/B500282/1 to M.S., NER/M/S/2002/00122 to M.S and G.M., GT5/98/8/MS and NE/B501039/1 to G.M).

Ancillary

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