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Phragmites australis and Quercus robur leaf extracts affect antioxidative system and photosynthesis of Ceratophyllum demersum
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Ecotoxicology and Environmental Safety 67 (2007) 240–246 www.elsevier.com/locate/ecoenv
Phragmites australis and Quercus robur leaf extracts affect antioxidative system and photosynthesis of Ceratophyllum demersum Sheku Kamara, Stephan Pflugmacher Leibniz Institute of Freshwater Ecology and Inland Fisheries, Biochemical Regulation, Mu¨ggelseedamm 301, 12587 Berlin, Germany Received 31 January 2006; received in revised form 4 July 2006; accepted 16 July 2006 Available online 22 September 2006
Abstract During senescence, leaves are deposited on aquatic bodies and decay under water releasing chemical substances that might exert physiological stress to aquatic organisms. Leaf litter alone contributes 30% of the total dissolved organic carbon (DOC) in streams. We investigated the impact of leaves extract from Phragmites australis and Quercus robur on the antioxidative system and photosynthetic rate of the aquatic macrophyte Ceratophyllum demersum exposed for 24 h. Rate of photosynthetic oxygen release and antioxidant enzyme activity (glutathione S-transferases, glutathione reductases and peroxidases) as well as lipid peroxidation in C. demersum were measured. Significant (Po0.01) elevations of antioxidative enzyme activity in C. demersum which tends to plateau at high DOC concentrations were observed. There was no detectable effect on lipid peroxidation. A significant dose-dependent reduction in photosynthetic oxygen production was measured. r 2006 Elsevier Inc. All rights reserved. Keywords: Ceratophyllum demersum; Dissolved organic carbon; Antioxidative enzymes; Physiological stress; Lipid peroxidation; Glutathione Stransferase; Glutathione reductase; Peroxidase; Phragmites australis; Quercus robur
1. Introduction Aquatic ecosystems are proliferated by a wide range of natural organic matter (NOM) originating from decomposing plant litter as well as animal and microbial debris. Organic matter can be derived both from terrestrial sources external to the aquatic system (allochthonous) and from sources within the aquatic system (autochthonous) (Meyer et al., 1998). In stream and river catchments NOM is derived primarily from allochthonous sources such as organic soil horizons (Brooks et al., 1999). On the other hand, autochthonous NOM is a significant fraction of community respiration in lake systems (Cole et al., 2000). Previous studies have focused mainly on the combined effects of organic matter (OM) and humic substances (HS) on organisms (Pflugmacher et al., 1999, 2001, 2003). However, the aquatic environment contains a wide range of autochthonous and allochthonous plant litter at various stages of decomposition and leaf litter alone contribute up Corresponding author. Fax: +49 3064 181682.
E-mail address: pfl[email protected] (S. Pflugmacher). 0147-6513/$ - see front matter r 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.ecoenv.2006.07.004
to 30% of the total dissolved organic carbon (DOC) in streams (Meyer et al., 1998). Research on the formation of reactive oxygen species (ROS) and the consequences in the cell when faced with certain chemical substances is of great importance in the elucidation of essential questions in stress physiology. Induction of antioxidative enzyme activity has been suggested as a convenient model for the investigation of stress in plants (Pflugmacher, 2004; Nimptsch et al., 2005). Leachates resulting from decomposing plant litter may or may not have adverse effects on the physiology and biochemistry of aquatic organisms depending on the nature and chemical composition of the plant exudates at the different stages of decomposition. Phragmites australis is an emergent macrophyte, cosmopolitan in distribution and makes up the most productive natural plant population in the biosphere (Wetzel, 2001). It is the dominant and most conspicuous plant species in many aquatic systems in temperate regions and often forms mono specific reed stands and huge reed-beds along many shallow lakes (Hollis and Jones, 1991). It grows perennially, with a constant turnover of population members that are senescing as new cohorts are emerging and
ARTICLE IN PRESS S. Kamara, S. Pflugmacher / Ecotoxicology and Environmental Safety 67 (2007) 240–246
growing. Cell membrane integrity collapses during senescence with major losses of organic matter into the aquatic system (Wetzel, 2001). Some oak trees (e.g., Quercus robur), which form a significant proportion in deciduous and mixed forests also lose their leaves seasonally. In aquatic bodies, plant litter usually constitutes a major fraction of the organic matter pool, which eventually undergoes microbial decomposition and mineralization (Polunin, 1984; Wetzel, 1995, 2001). This organic matter, through the release of its nutrients, may provoke stress effects on aquatic biota. Plants have developed internal physiological defense mechanisms to deal with such stressors. When susceptible plants are exposed to chemical substances, germination, growth and development may be affected. The most frequently reported gross morphological effects on plants are inhibited or retarded seed germination, effects on coleoptiles elongation and on radicles, shoot and root development (Kruse et al., 2000; Hunter and Menges, 2002; Bosch et al., 2004). In a recent study, Salminen et al. (2004) found hydrolysable tannins to be the dominant group of phenolic compounds in the leaves of the oak species Q. robur. A rare dimeric ellagitannin, cocciferin D2, was also detected for the first time in the leaves of Q. robur. A few earlier studies have also shown that the leaves of Q. robur contain flavonol glycosides, castalagin, pedunculagin, vescalagin, and casuarictin (Scalbert and Haslam, 1987; Scalbert et al., 1988; Grundho¨fer et al., 2001). One of the earliest studies of oak leaf chemistry was by Feeny (1970) studying the moth Operopthera brumata which feeds on oak leaves mainly in spring because of the decline in the nutritional value of the leaves during summer. Riipi et al. (2002) also reported a seasonal decline in birch foliage suitability for herbivores corresponding to declining concentrations of proteins and free amino acids. There was also a decrease in concentration of cell wall-bound proanthocyanidins, gallotannins and flavonoid glycosides but an increase in the soluble proanthocyanidins in the birch leaves especially in spring. There is only few literature on the effect of organic matter or humic substances on freshwater organisms (Pflugmacher et al., 1999, 2001; Pflugmacher, 2002). The synthetic organic compound, 3-chlorobiphenyl, was recently shown to induce some level of physiological stress at very low concentrations on Ceratophyllum demersum (Menone and Pflugmacher, 2005). Uptake of dissolved organic matter by C. demersum and of cyanobacterial toxins by P. australis has been shown to trigger physiological and biochemical responses (Pflugmacher et al., 1999, 2000). Like synthetic herbicides, there is no common mode of action or physiological target site for all allelochemicals. However, known sites of action for some allelochemicals include cell division, photosynthesis, nutrient uptake, pollen germination and specific enzyme function (Pflugmacher et al., 1999; Kruse et al., 2000). The present study investigated the impact, in terms of oxidative stress induction and photosynthetic oxygen production, of extract from dead P. australis and Q. robur litter on the aquatic macrophyte C.
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demersum. The submerged floating macrophyte, C. demersum was chosen for this study because of its sensitiveness in responding to slight environmental perturbations and its cosmopolitan distribution. Additionally, it plays a pivotal role in freshwater ecosystems because as a primary producer, it forms part of the baseline for materials and energy transfer in the aquatic food web. Some herbivorous migratory waterfowls eat the seeds and foliage. Furthermore, it often forms an important microhabitat for young fish, aquatic insects and small aquatic animals (Jeppeson et al., 1999).
2. Materials and method 2.1. Preparation and characterization of natural organic matter (NOM) extract Standing dead P. australis plants were collected from the littoral zone of Lake Mu¨ggelsee in April 2005. The above-ground plant parts were cut at about 1 m above the water level and composed of the leaf blade and culms. Fallen dead oak leaves (Q. robur) from top litter layer were collected within the catchments area of Lake Mu¨ggelsee in the same period. These were air-dried for 2 days in order to maintain an approximately uniform humidity level and ground using a homogeniser Mill (1094 Tecator). Eighty and 100 g, respectively of P. australis and Q. robur ground material was soaked in 1.5 L of medium containing deionized water, CaCl2 (0.2 g L1), NaHCO3 (0.103 g L1) and sea-salt (0.1 g L1) in separate plastic containers and stirred for 24 h at room temperature. The resulting mixture was centrifuged (L-60 Ultracentrifuge, Beckman LL-TB-003A) at 20,000g for 10 min at 4 1C to remove suspended materials. The supernatant was filtered using 0.8 mm cellulose-nitrate membrane filter (Sartorius AG, Germany) and total dissolved organic carbon was determined according to DIN EN 1484 (1998).
2.2. Plant material and exposure to NOM extracts C. demersum was purchased from Aqua Global (Dr. Jander & Co. OHG, Seefeld, Germany) and cultivated non-axenically in Provasoli’s medium containing de-ionized water, CaCl2 (0.2 g L1), NaHCO3 (0.103 g L1) and sea salt (0.1 g L1) in an aquarium prior to exposure experiments for acclimatization (Pflugmacher et al., 2003). Supplementary light was provided by daylight lamps with an irradiance of 12 mE m2 s, a photoperiod of 14:10 h light:dark cycle and temperature was maintained at 2371 1C. 3.5 g FW of C. demersum was exposed to P. australis and Q. robur extract in serial concentrations of 0.1, 1.0, 10.0, and 100 mg L1 DOC and placed under constant light and temperature conditions for 24 h. In control experiments, only medium water (i.e., no leaf extract) was added. The control as well as the exposures with oak and reed extracts was repeated five times independently.
2.3. Measurement of photosynthesis Photosynthetic oxygen production of the plant was measured using the device Plant Vital 5000 (Inno-Concept, Straussberg). Measurements were made at 100% light intensity (approximately 2000 1 ), a dark/light/dark cycle of 10/12/10 min and at 20 1C. Rates of photosynthetic oxygen production were calculated as mmoles O2sh1g fw1. After measuring photosynthesis, plants were immediately frozen in liquid N2 and kept at 80 1C for enzyme preparation.
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2.4. Enzyme preparation
2.7. Statistical analyses
Enzyme preparation was done according to the method described by Pflugmacher and Steinberg (1997) with slight modification on the steps leading to the microsomal fraction. The frozen plants were ground to a fine powder, suspended in sodium phosphate buffer (0.1 M, pH 6.5) containing 20% glycerol, 1.4 mM dithioerythritol (DTE), and 1 mM ethylenediamine tetraceticacid (EDTA). Cell debris was removed by centrifuging the slurry at 10,000g for 10 min. The residue was suspended in the same buffer and centrifuged again at 5000g for 5 min. The supernatant from the first and second centrifugation was then centrifuged at 40,000g for 60 min. The pellets, containing membrane fractions and defined as the microsomes, was suspended in sodium phosphate buffer (20 mM, pH 7.0) containing 20% glycerol and homogenized using glass homogenizer. The supernatant was precipitated with solid ammonium sulphate in two saturation steps, 0–35% and 35–80%, followed, respectively by two centrifugation steps at 30,000g for 30 min each. The precipitate from the second precipitation which contains soluble proteins were suspended in sodium phosphate buffer (20 mM, pH 7.0) and desalted by gel filtration using NAP-5 columns (Amersham Pharmacia, Germany). Both microsomal and soluble protein extracts were frozen in liquid N2 and stored at –80 1C for enzyme activity assay.
One-way analysis of variance (ANOVA) was performed to test for differences among treatments. Planned post hoc comparisons (Po0.05) were performed using Turkeys HSD test (STASTICA).
2.5. Enzyme activity determination Activity of glutathione S-tranferases (GST) in soluble (cytosol) and microsomal fractions was measured with 1-chloro-2,4-dinitrobenzene (CDNB) as substrate according to Habig et al. (1974). Additionally, GST activity in the cytosol was measured with 4-hydroxynonenal (4HNE) as substrate according to Alin et al. (1984). Peroxidase activity in the soluble fraction was measured using guajacol as substrate (Drotar et al., 1985). Glutathione reductase (GR, E.C. 1.6.4.2) activity in the soluble fraction was measured spectrophotometrically according to Carlberg and Mannervik (1985) based on the oxidation of NADPH to NADP+ which is accompanied by a decrease in absorbance at 340 nm. Enzyme activity was calculated in nano-katals per milligram protein (katal ¼ conversion rate of one mole substrate per second). Protein content of the samples was determined according to Bradford (1976). In special cases where very low protein amounts were achieved in the samples, ‘‘Advanced Protein Assay (ADV01)’’ from Cytoskeleton was used. The assay principle is the same as for Bradford, with the difference that the reagent is five times more concentrated. Serum bovine albumin (initial fraction 98%, Sigma) was used as protein standard for calibration in both assays.
3. Results Direct exposure of the aquatic macrophyte, C. demersum, to extracts of P. australis and Q. robur leaves showed a clear adverse effect on photosynthetic oxygen production (Fig. 1). The effects were significant at all tested concentrations in both leaf extracts. At the highest concentrations (10 and 100 mg L1 DOC), very strong effects were observed with a highly significant (Po0.01) reduction in the rate of photosynthetic oxygen release. Photosynthetic rate of C. demersum was more reduced in Q. robur extract than in P. australis extract especially at the 1 and 10 mg L1 DOC concentrations compared to controls. Measurement of both microsomal (Fig. 2A) and soluble (cytosolic) (Fig. 2B) GST in exposed C. demersum using CDNB as substrate showed a significant (Po0.01) elevation in activity compared to control. Soluble and microsomal GST activity increased with increase in DOC concentration and tends to reach a plateau at higher (10 and 100 mg L1 DOC) concentrations in Q. robur extracts. On the other hand, soluble GST activity in C. demersum exposed to P. australis extract begins to decrease at higher DOC concentrations (10 and 100 mg L1). A concentration-dependent significant (Po0.01) increase in soluble GST activity, using 4-hydroxynonenal (4-HNE) as substrate, was detected in C. demersum exposed to Q. robur extract (Fig. 3). In C. demersum exposed to P. australis extract the increase in activity was also significant but not concentration dependent. Glutathione reductase activity in C. demersum exposed to P. australis and Q. robur extracts increased significantly (Po0.01) compared to untreated control, with a tendency 50
Lipid peroxidation (LPO) was determined according to the User protocol of the lipid peroxidation assay kit (CALBIOCHEM, Cat. No. 437634) with minor modifications based on the method of Botsoglou et al. (1994). The aldehyde, malondialdehyde (MDA), is an end product derived from the breakdown of lipid peroxides, which are often unstable. Estimation of MDA concentration serves as an index of lipid peroxidation. The reaction between N-methyl-2-phenylindole and MDA produces a stable chromophore with maximum absorbance at 586 nm. Plant material was weighed, homogenized in sodium phosphate buffer (20 mM, pH 7.0) and centrifuged for 10 min at 14,000g and at 4 1C. In order to prevent sample oxidation, 10 mL of 0.5 mM butylated hydroxytoluene (BHT) was added. The supernatant was then used for the assay. Six hundred microlitres of N-methyl-2-phenylindole in acetonitrile was added to 200 mL of the sample and gently mixed. One hundred and fifty microlitres of 12 N HCl was then added, mixed again and incubated for 1 h at 45 1C. Samples were cooled on ice and the absorbance measured at 586 nm. A calibration curve was made using MDA.
µmol O2 (gFW)-1 . sh-1
2.6. Measurement of lipid peroxidation 40 ∗ 30
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Dissolved organic carbon (mg . L-1) Fig. 1. Dose-response photosynthetic oxygen production in Ceratophyllum demersum after 24 h exposure to 5 different concentrations of two plant extracts; n ¼ 5, error bars represent7standard deviation, Phragmites australis extract, Quercus robur extract, *Po0.05, **Po0.01.
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Fig. 3. Dose-response activity of soluble (cytosolic) glutathione Stransferase (measured with 4-hydroxynonenal) in Ceratophyllum demersum after 24 h exposure to 5 different concentrations of two plant extracts; n ¼ 5, error bars represent7standard deviation, Phragmites australis extract, Quercus robur extract; *Po0.01, **Po0.001.
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Fig. 4. Lipid peroxidation (measured as malondialdehyde, MDA, concentration) in Ceratophyllum demersum after 24 h exposure to 5 different concentrations of two plant extracts; n ¼ 3, error bars represent7standard deviation, Phragmites australis extract, Quercus robur extract.
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Dissolved organic carbon (mg . L-1)
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Dissolved organic carbon (mg . L-1) Fig. 2. Dose-response enzyme activity in Ceratophyllum demersum after 24 h exposure to 5 different concentrations of two plant extracts. (A) microsomal glutathione S-transferase measured with CDNB; (B) cytosolic glutathione S-transferase measured with CDNB; (C) glutathione reductase; (D) guajacol peroxidase; n ¼ 5, error bars represent7standard deviation, Phragmites australis extract, Quercus robur extract, *Po0.01, **Po0.001.
to decrease at the highest (100 mg L1) DOC concentration (Fig. 2C). Activity of guajacol peroxidase in C. demersum increased significantly (Po0.01) at all doses compared with control and in both P. australis and Q. robur extract (Fig. 2D). Elevations in POD activity were higher in C. demersum exposed to Q. robur than that exposed to P. australis extract. Highest elevation compared to control was by a factor of 12 in P. australis exposure and by a factor of 47 in
Q. robur exposure. C. demersum exposed to P. australis showed a gradual decrease in POD activity as the concentration increased from 0.1 to 100 mg L1 DOC. No significant differences (P40.05) in concentration of malondialdehyde (MDA), an index of lipid peroxidation, were detected between control and C. demersum exposed to both Q. robur and P. australis extract at all DOC concentrations tested (Fig. 4). 4. Discussion Photosynthetic oxygen production is an important physiological parameter often used to test the sensitivity of aquatic plants to changes in their surrounding (Sersen et al., 1998; Menone and Pflugmacher, 2005). In the present study, a significant concentration-dependent reduction in rate of photosynthesis of C. demersum after a 24 h
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pre-exposure to both Q. robur and P. australis extracts was observed. Similar effects have been demonstrated in earlier studies using humic substances isolated from soils (Pflugmacher et al., 1999). The mechanism of action for this observation is not clear. However, aromatic structures like phenols and aromatic hydrocarbons are known to have toxic effects on organisms. For example the oxidation of quinone leading to the formation of radicals is known to interfere with electron transfer chain in photosystem II (PSII) in plant chloroplasts (Oettmeier et al., 1988). Using the pure quinone, 1-aminoanthraquinone, photosynthetic oxygen release in Vesicularia dubyana was reduced to almost zero (Pflugmacher et al., 2003). It was also shown in a recent study that the aromatic organic compound, 3-chlorobiphenyl, significantly inhibited the photosynthesis of C. demersum at a concentration of 5 mg L1 (Menone and Pflugmacher, 2005). The significant (Po0.01) reduction in rate of photosynthetic oxygen release due to the oak (Q. robur) extract, which is well known for its high tannin and phenolic contents (Salminen et al., 2004), suggests a possible interference with electron transport chain from PSII to PSI. The adverse effect caused by Q. robur extract was indeed greater than that caused by P. australis extract. Additionally, it is known that when plants are exposed to stress conditions, there is an increase in ROS (Pflugmacher, 2004) and the antioxidant capacity of the plant may be overwhelmed. Organelles such as the peroxisomes and chloroplast (site of photosynthesis), where ROS are being produced at a relatively high rate, are especially at risk (Grene, 2002). Overproduction of ROS in chloroplasts of plants under drought-stress has been reported (Price et al., 1989), and a similar mechanism may occur when plants are exposed to certain concentrations of leaf extracts. The induced activity of antioxidative enzymes observed in the present study suggests an increase in ROS, and the susceptibility of the chloroplasts to high ROS could be an additional factor contributing to the corresponding reduction in the rate of photosynthesis. ROS such as O2d, OH, H2O2 are often internally formed as products of normal plant metabolism (Foyer et al., 1994). Under normal habitat conditions, the rate at which ROS are formed by plants is in dynamic equilibrium with the rate at which they are further utilized or broken down. Leaf extracts might be able to promote oxidative stress by inducing the production of more ROS. To combat the ROS, plants have developed an antioxidative defense system. Parts of this system are the peroxidases (POD), GST and glutathione reductase (GR). In the present study, there was a significant (Po0.01) increase in POD activity in the aquatic macrophyte C. demersum after exposure to both Q. robur and P. australis extracts for 24 h. POD catalyses the reduction of H2O2 to water for detoxification. Exposure of the macrophyte to the leaf extracts may have resulted in an increase in the production of H2O2 beyond the normal detoxifying capacity of the plant. In order to deal with this high level of H2O2 and to prevent or minimize damage to the plant cells,
the plant POD enzymes increased their activity many folds thereby catalyzing the conversion of the highly reactive H2O2 into harmless water molecules. Previous studies with other plant species such as Hydrilla verticillata also demonstrated a significant increase in POD activity after exposure to anthracene in concentrations above 0.01 mg L1 (Byle et al. 1994). Using humic substances from different soil and water sources, Pflugmacher et al. (1999) also showed an increase in POD activity on C. demersum but after 7 days exposure in a concentration of 0.5 mg L1. The very high increase (up to 47 folds) in POD activity observed in the present study was after 24 h exposure of C. demersum to crude leaf extracts. DOC concentrations as low as 0.1 mg L1 provoked significant (Po0.01) increases in POD activity. Aromatic structures and plant growth hormones like gibberellic acid are known to cause an elevation of POD activity (Kwak et al., 1996, Oberg et al., 1990). Response of POD activity upon exposure to leaf extracts from P. australis and Q. robur suggests that these chemicals may contain similar structures. Oak, for example, is well known for its high tannin and phenolic contents (Salminen et al., 2004). The antioxidant activity of phenols is mainly due to their redox properties, which allow them to act as reducing agents, hydrogen donors and singlet oxygen quenchers (RiceEvans et al., 1995). Soluble and microsomal glutathione S-transferase (sGST and mGST) activity was also found to be significantly elevated upon exposure to extracts from Q. robur and P. australis. These enzymes are invariably involved in cellular biotransformation when plants are faced with xenobiotic or toxic substances. They do so by conjugating toxicants or xenobiotics to the sulphurhydryl (–SH) group of glutathione (GSH) which enhances their excretion from cells through cell membranes. Increase in GST activity is thus considered a chemical stress signal. Similarly, Menone and Pflugmacher (2005) demonstrated that soluble GST activity in C. demersum was significantly elevated when treated with lower concentrations (up to 0.5 mg L1) of the organic PCB compound 3-chlorobiphenyl. Imposition of oxidative stress often leads to the conversion of the existing pool of reduced glutathione (GSH) to the oxidized form, glutathione disulphide (GSSG), thereby stimulating glutathione biosynthesis (May and Leaver, 1993). After GSH has been oxidized to GSSG, the recycling of GSSG to GSH is accomplished mainly by the enzyme GR. An elevated glutathione reductase activity detected in the present study was a further indication of an oxidative stress condition in the exposed C. demersum. GSH is used by glutathione peroxidase to detoxify hydrogen peroxide and in the process is converted to GSSG, which is then recycled to GSH by GR. The regenerated GSH is then available for the detoxification of more hydrogen peroxide. An increase in GSH biosynthetic capacity has been shown to enhance resistance to oxidative stress (Zhu et al., 1999). ROS are extremely reactive and unstable chemical species, which react with proteins, nucleic acids, carbohydrates,
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and lipids in cells. The latter often result in lipid peroxidation. Malondialdehyde (MDA) is one of the end products of lipid peroxidation. The findings in the present study did not reveal any significant (P40.05) increase in MDA levels in exposed (treated) C. demersum in all concentrations tested. This is presumably due to activation of defense and detoxification mechanisms, as evident by high GST, GR and POD activities observed in the present study (Figs. 2 and 3). These antioxidant enzymes can decompose ROS and prevent the damage to cellular constituents and initiation of lipid peroxidation (Yang et al., 2003). Furthermore, GST conjugates MDA and 4-HNE using GSH as substrate in the biotransformation process. 5. Conclusion Our results indicate that leaf extracts have the capacity to impose physiological stress and provoke acute antioxidative responses in aquatic organisms, in this case C. demersum. The effects observed within one photoperiodic cycle (24 h) are quite strong. This means that aquatic macrophytes could serve as early warning systems in water quality assessment procedures. Furthermore, we demonstrated that even relatively low DOC concentrations of leaf extracts from P. australis and Q. robur can induce oxidative stress as well as slow down the rate of photosynthesis in the coontail, C. demersum. The effects tend to level off or in some cases start to decrease at high DOC concentrations indicating a threshold on the antioxidative ability of the macrophyte. One of the damaging effects often observed in organisms exposed to stress is lipid peroxidation. In this study, it seems evident that the antioxidative system in C. demersum was sufficiently activated to prevent or minimise this damage since no significant effect on lipid peroxidation was detected in treated plants. Our findings thus suggest that leaf extracts may be an important environmental factor affecting aquatic biota and contributing to the overall ecological dynamics in shallow lakes and rivers. Acknowledgments Our thanks to J. Nimptsch for laboratory assistance and useful discussions on the statistical analyses. The German academic exchange service (DAAD) is gratefully acknowledged for providing financial support for S. Kamara. References Alin, P., Danielson, U.H., Mannervik, B., 1984. 4-hydroxyalk-2-enals are substrates for glutathione transferase. FEBS Lett. 179 (2), 267–270. Bosch, E.V. D., Ward, B.G., Clarkson, B.D., 2004. Woolly Nightsahde (Solanum mauritianum) and its allelopathic effects on New Zealand native Hebe stricta seed germination. In: Proceedings of the 57th New Zealand Plant Protection Society Incorporation Conference, pp. 98–101. Botsoglou, N.A., Fletouris, D.J., Papageorgiou, G.E., Vassilopoulos, V.N., Mantis, A.J., Trakatellis, A.G., 1994. Rapid, sensitive, and
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specific thiobarbituric acid method for measuring lipid peroxidation in animal tissue, food, and feedstuff samples. J. Agric. Food Chem. 42, 1931–1937. Bradford, M.M., 1976. A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein dye-binding. Anal. Biochem. 72, 248–254. Brooks, P.D., McKnight, D.M., Bencala, K.E., 1999. The relationship between soil heterotrophic activity, soil dissolved organic carbon (DOC) leachate, and catchment-scale DOC export in headwater catchments. Water Resour. Res. 35, 1895–1902. Byle, T.D., Sutton, H.D., Klaine, S.J., 1994. Evaluation of peroxidase as a biochemical indicator of toxic chemical exposure in the aquatic plant Hydrilla verticillata. Royle. Environ. Toxicol. Chem., 509–515. Carlberg, I., Mannervik, B., 1985. Glutathione reductase. Methods Enzymol. 113, 484–490. Cole, J.J., Pace, M.L., Carpenter, S.R., Kitchell, J.F., 2000. Persistence of net heterotrophy in lakes during nutrient addition and food web manipulations. Limnol. Oceanogr. 45 (8), 1718–1730. DIN EN 1484, 1998. Anleitung zur Bestimmung des gesamten organischen Kohlenstoffs (TOC) und des gelo¨sten organischen Kohlenstoffs. DEV, H3, 40, Lieferung. Drotar, A., Phelps, P., Fall, R., 1985. Evidence for glutathione peroxidase activities in cultured plant cells. Plant Sci. 42, 35–40. Feeny, P.P., 1970. Seasonal changes in oak leaf tannins and nutrients as a cause of spring feeding by winter moth caterpillars. Ecology 51, 565–581. Foyer, C.H., Lescure, J.C., Lefebvre, C., Morot-Guadry, J.F., Vincent, M., Vaucheret, H., 1994. Adaptations of photosynthetic electron transport, carbon assimilation and carbon partitioning in transgenic Nicotiana plumbaginifolia plants to changes in nitrate reductase activity. Plant Physiol. 104, 171–178. Grene, R., 2002. Oxidative stree and acclimation mechanisms in plants. The Arabidopsis Book. Am. Soc. Plant Biol., 1–20. Grundho¨fer, P., Niemetz, R., Schilling, G., Gross, G.G., 2001. Biosynthesis and subcellular distribution of hydrolyzable tannins. Phytochemistry 57, 915–927. Habig, W., Pabst, M.J., Jakoby, W.B., 1974. Glutathione S-transferase: the first step in mercapturic acid formation. J. Biol. Chem. 249, 1730–1739. Hollis, G.E., Jones, T.A., 1991. Europe and the Mediterranean Basin. In: Finlayson, M., Moser, M. (Eds.), Wetlands, pp. 27–56. Hunter, M.E., Menges, E.S., 2002. Allelopathic effects and root distribution of Ceratiola ericoides (Empetraceae) on seven Rosemary scrub species. Am. J. Bot. 89 (7), 1113–1118. Jeppeson, E., Sondergaard, M., Kronvang, B., Jensen, J.P., Svendsen, L.M., Lauridsen, T.L., 1999. Lake and catchment management in Denmark. Hydrobiologia 395/396, 419–432. Kruse, M., Strandberg, M., Strandberg, B., 2000. Ecological Effects of Allelopathic Plants—a Review. National Environmental Research Institute, Silkeborg, Denmark, 66pp., NERI Technical Report No. 315. Kwak, S.S., Kim, S.K., Park, I.H., Liu, J.R., 1996. Enhancement of peroxidase activity by stress-related chemicals in sweet potato. Phytochemistry 43, 565–568. May, M.J., Leaver, C.J., 1993. Oxidative stimulation of glutathione synthesis in Arabidopsis thaliana suspension cultures. Plant Physiol. 103, 621–627. Menone, M.L., Pflugmacher, S., 2005. Effects of 3-chlorobiphenyl on photosynthetic oxygen production, glutathione content and detoxication enzymes in the aquatic macrophyte Ceratophyllum demersum. Chemosphere 60, 79–84. Meyer, J.L., Wallace, J.B., Eggert, S.L., 1998. Leaf litter as a source of dissolved organic carbon in streams. Ecosystems 1, 240–249. Nimptsch, J., Wunderlin, D.A., Dollan, A., Pflugmacher, S., 2005. Antioxidant and biotransformation enzymes in Myriophyllum quitense as biomarkers of heavy metal exposure and eutrophication in Suquia River basin (Cordoba, Argentina). Chemos 61, 147–157.
ARTICLE IN PRESS 246
S. Kamara, S. Pflugmacher / Ecotoxicology and Environmental Safety 67 (2007) 240–246
Oberg, L.G., Glas, B., Swanson, S.E., Rappe, C., Paul, K.G., 1990. Peroxidas-catalyzed oxidation of chlorophenols to polychlorinated dibenzo-p-dioxins and dibenzofurans. Arch. Environ. Contam. Toxicol. 19, 930–938. Oettmeier, W., Masson, K., Donner, A., 1988. Anthraquinone inhibitors of photosystem II electron transport. FEBS Lett. 231, 259–262. Pflugmacher, S., Steinberg, C.E.W., 1997. Activity of phase I and phase II detoxication enzymes in aquatic macrophytes. J. Appl. Bot. 71, 144–146. Pflugmacher, S., Spangenberg, M., Steinberg, C.E.W., 1999. Dissolved organic matter (DOM) and effects on the aquatic macrophyte Ceratophyllum demersum in relation to photosynthesis, pigment pattern and activity of detoxication enzymes. J. Appl. Bot. 73, 184–190. Pflugmacher, S., Wiegand, C., Beattle, K.A., Krause, E., Steinberg, C.E.W., Codd, G.A., 2000. Uptake, effects and metabolism of cyanobacterial toxins in the emergent reed plant Phragmites australis (Cav.). Trin. Ex. Steud. Environ. Toxic. Chem. 20 (4), 846–852. Pflugmacher, S., 2002. Possible allelopathic effects of cyanotoxins, with reference to Microcystin LR, in aquatic ecosystems. Wiley Period, Inc. Environ. Toxicol. 17, 407–413. Pflugmacher, S., Tidwell, L.F., Steinberg, C.E.W., 2001. Dissolved humic substances directly affect freshwater organisms. Acta Hydrochim. Hydrobiol. 29, 34–40. Pflugmacher, S., Pietsch, C., Reiger, W., Paul, A., Preuer, T., Zwirnmann, E., Steinberg, C.E.W., 2003. Humic substances and their direct effects on the physiology of aquatic plants. In: Ghabbour, E.A., Davies, G. (Eds.), Humic Substances: Nature’s Most Versatile Materials. Taylor & Francis, New York. Pflugmacher, S., 2004. Promotion of oxidative stress in the aquatic macrophyte Ceratophyllum demersum during biotransformation of the cyanobacterial toxin microcystin-LR. Aquat. Toxicol. 70, 169–178. Polunin, N.V.C., 1984. The decomposition of emergent macrophytes in freshwater. Adv. Ecol. Res. 14, 115–166.
Price, A.H., Atherton, N.M., Hendry, G.A.F., 1989. Plants under drought-stress generate activated oxygen. Free Radical Res. Commun. 8, 61–66. Rice-Evans, C.A., Miller, N.J., Bolwell, P.G., Bramley, P.M., Pridham, J.B., 1995. The relative antioxidant activities of plant-derived polyphenolic flavonoids. Free Radical Res. 22, 375–383. Riipi, M., Ossipov, V., Lempa, K., Haukioja, E., Koricheva, J., Ossipova, S., Pihlaja, K., 2002. Seasonal changes in birch leaf chemistry: are there trade-offs between leaf growth and accumulation of phenolics. Oecologia 130, 380–390. Salminen, J., Roslin, T., Karonen, M., Sinkkonen, J., Pihlaja, K., Pulkkinen, P., 2004. Seasonal variation in the content of hydrolizable tannins, flavonoid glycosides and proanthocyanidins in oak leaves. J. Chem. Ecol. 30 (9), 1693–1711. Scalbert, A., Haslam, E., 1987. Plant polyphenols and chemical defense. Part 2: Polyphenols and chemical defense of the leaves of Quercus robur. Phytochemistry 26, 3191–3195. Scalbert, A., Monties, B., Favre, J.M., 1988. Polyphenols of Quercus robur: adult tree and in vitro grown calli and shoots. Phytochemistry 27, 3483–3488. Sersen, F., Kralova, K., Bumbalova, A., 1998. Action of mercury on the photosynthetic apparatus of spinach chloroplasts. Photosynthetica 35, 551–559. Wetzel, R.G., 1995. Death, detritus and energy flow in aquatic ecosystems. Freshwater Biol. 33, 83–89. Wetzel, R.G., 2001. Limnology: lake and river ecosystems, third ed. Academic Press, San Diego, pp. 527–664. Yang, Y., Sharma, R., Sharma, A., Awasthi, S., Awasthi, Y.C., 2003. Lipid peroxidation and cell cycle signaling: 4-hydroxynonenal, a key molecule in stress mediated signaling. Acta Biochim. Polon. 50 (2), 319–336. Zhu, Y.L., Philon-Smits, E.A.H., Tarun, A.S., Weber, S.U., Jouanin, L., Terry, N., 1999. Cadmium tolerance and accumulation in Indian mustard is enhanced by overexpressing gamma-glutamylcysteine synthetase. Plant Physiol. 121, 1169–1178.
Ecotoxicology and Environmental Safety 67 (2007) 240–246 www.elsevier.com/locate/ecoenv
Phragmites australis and Quercus robur leaf extracts affect antioxidative system and photosynthesis of Ceratophyllum demersum Sheku Kamara, Stephan Pflugmacher Leibniz Institute of Freshwater Ecology and Inland Fisheries, Biochemical Regulation, Mu¨ggelseedamm 301, 12587 Berlin, Germany Received 31 January 2006; received in revised form 4 July 2006; accepted 16 July 2006 Available online 22 September 2006
Abstract During senescence, leaves are deposited on aquatic bodies and decay under water releasing chemical substances that might exert physiological stress to aquatic organisms. Leaf litter alone contributes 30% of the total dissolved organic carbon (DOC) in streams. We investigated the impact of leaves extract from Phragmites australis and Quercus robur on the antioxidative system and photosynthetic rate of the aquatic macrophyte Ceratophyllum demersum exposed for 24 h. Rate of photosynthetic oxygen release and antioxidant enzyme activity (glutathione S-transferases, glutathione reductases and peroxidases) as well as lipid peroxidation in C. demersum were measured. Significant (Po0.01) elevations of antioxidative enzyme activity in C. demersum which tends to plateau at high DOC concentrations were observed. There was no detectable effect on lipid peroxidation. A significant dose-dependent reduction in photosynthetic oxygen production was measured. r 2006 Elsevier Inc. All rights reserved. Keywords: Ceratophyllum demersum; Dissolved organic carbon; Antioxidative enzymes; Physiological stress; Lipid peroxidation; Glutathione Stransferase; Glutathione reductase; Peroxidase; Phragmites australis; Quercus robur
1. Introduction Aquatic ecosystems are proliferated by a wide range of natural organic matter (NOM) originating from decomposing plant litter as well as animal and microbial debris. Organic matter can be derived both from terrestrial sources external to the aquatic system (allochthonous) and from sources within the aquatic system (autochthonous) (Meyer et al., 1998). In stream and river catchments NOM is derived primarily from allochthonous sources such as organic soil horizons (Brooks et al., 1999). On the other hand, autochthonous NOM is a significant fraction of community respiration in lake systems (Cole et al., 2000). Previous studies have focused mainly on the combined effects of organic matter (OM) and humic substances (HS) on organisms (Pflugmacher et al., 1999, 2001, 2003). However, the aquatic environment contains a wide range of autochthonous and allochthonous plant litter at various stages of decomposition and leaf litter alone contribute up Corresponding author. Fax: +49 3064 181682.
E-mail address: pfl[email protected] (S. Pflugmacher). 0147-6513/$ - see front matter r 2006 Elsevier Inc. All rights reserved. doi:10.1016/j.ecoenv.2006.07.004
to 30% of the total dissolved organic carbon (DOC) in streams (Meyer et al., 1998). Research on the formation of reactive oxygen species (ROS) and the consequences in the cell when faced with certain chemical substances is of great importance in the elucidation of essential questions in stress physiology. Induction of antioxidative enzyme activity has been suggested as a convenient model for the investigation of stress in plants (Pflugmacher, 2004; Nimptsch et al., 2005). Leachates resulting from decomposing plant litter may or may not have adverse effects on the physiology and biochemistry of aquatic organisms depending on the nature and chemical composition of the plant exudates at the different stages of decomposition. Phragmites australis is an emergent macrophyte, cosmopolitan in distribution and makes up the most productive natural plant population in the biosphere (Wetzel, 2001). It is the dominant and most conspicuous plant species in many aquatic systems in temperate regions and often forms mono specific reed stands and huge reed-beds along many shallow lakes (Hollis and Jones, 1991). It grows perennially, with a constant turnover of population members that are senescing as new cohorts are emerging and
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growing. Cell membrane integrity collapses during senescence with major losses of organic matter into the aquatic system (Wetzel, 2001). Some oak trees (e.g., Quercus robur), which form a significant proportion in deciduous and mixed forests also lose their leaves seasonally. In aquatic bodies, plant litter usually constitutes a major fraction of the organic matter pool, which eventually undergoes microbial decomposition and mineralization (Polunin, 1984; Wetzel, 1995, 2001). This organic matter, through the release of its nutrients, may provoke stress effects on aquatic biota. Plants have developed internal physiological defense mechanisms to deal with such stressors. When susceptible plants are exposed to chemical substances, germination, growth and development may be affected. The most frequently reported gross morphological effects on plants are inhibited or retarded seed germination, effects on coleoptiles elongation and on radicles, shoot and root development (Kruse et al., 2000; Hunter and Menges, 2002; Bosch et al., 2004). In a recent study, Salminen et al. (2004) found hydrolysable tannins to be the dominant group of phenolic compounds in the leaves of the oak species Q. robur. A rare dimeric ellagitannin, cocciferin D2, was also detected for the first time in the leaves of Q. robur. A few earlier studies have also shown that the leaves of Q. robur contain flavonol glycosides, castalagin, pedunculagin, vescalagin, and casuarictin (Scalbert and Haslam, 1987; Scalbert et al., 1988; Grundho¨fer et al., 2001). One of the earliest studies of oak leaf chemistry was by Feeny (1970) studying the moth Operopthera brumata which feeds on oak leaves mainly in spring because of the decline in the nutritional value of the leaves during summer. Riipi et al. (2002) also reported a seasonal decline in birch foliage suitability for herbivores corresponding to declining concentrations of proteins and free amino acids. There was also a decrease in concentration of cell wall-bound proanthocyanidins, gallotannins and flavonoid glycosides but an increase in the soluble proanthocyanidins in the birch leaves especially in spring. There is only few literature on the effect of organic matter or humic substances on freshwater organisms (Pflugmacher et al., 1999, 2001; Pflugmacher, 2002). The synthetic organic compound, 3-chlorobiphenyl, was recently shown to induce some level of physiological stress at very low concentrations on Ceratophyllum demersum (Menone and Pflugmacher, 2005). Uptake of dissolved organic matter by C. demersum and of cyanobacterial toxins by P. australis has been shown to trigger physiological and biochemical responses (Pflugmacher et al., 1999, 2000). Like synthetic herbicides, there is no common mode of action or physiological target site for all allelochemicals. However, known sites of action for some allelochemicals include cell division, photosynthesis, nutrient uptake, pollen germination and specific enzyme function (Pflugmacher et al., 1999; Kruse et al., 2000). The present study investigated the impact, in terms of oxidative stress induction and photosynthetic oxygen production, of extract from dead P. australis and Q. robur litter on the aquatic macrophyte C.
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demersum. The submerged floating macrophyte, C. demersum was chosen for this study because of its sensitiveness in responding to slight environmental perturbations and its cosmopolitan distribution. Additionally, it plays a pivotal role in freshwater ecosystems because as a primary producer, it forms part of the baseline for materials and energy transfer in the aquatic food web. Some herbivorous migratory waterfowls eat the seeds and foliage. Furthermore, it often forms an important microhabitat for young fish, aquatic insects and small aquatic animals (Jeppeson et al., 1999).
2. Materials and method 2.1. Preparation and characterization of natural organic matter (NOM) extract Standing dead P. australis plants were collected from the littoral zone of Lake Mu¨ggelsee in April 2005. The above-ground plant parts were cut at about 1 m above the water level and composed of the leaf blade and culms. Fallen dead oak leaves (Q. robur) from top litter layer were collected within the catchments area of Lake Mu¨ggelsee in the same period. These were air-dried for 2 days in order to maintain an approximately uniform humidity level and ground using a homogeniser Mill (1094 Tecator). Eighty and 100 g, respectively of P. australis and Q. robur ground material was soaked in 1.5 L of medium containing deionized water, CaCl2 (0.2 g L1), NaHCO3 (0.103 g L1) and sea-salt (0.1 g L1) in separate plastic containers and stirred for 24 h at room temperature. The resulting mixture was centrifuged (L-60 Ultracentrifuge, Beckman LL-TB-003A) at 20,000g for 10 min at 4 1C to remove suspended materials. The supernatant was filtered using 0.8 mm cellulose-nitrate membrane filter (Sartorius AG, Germany) and total dissolved organic carbon was determined according to DIN EN 1484 (1998).
2.2. Plant material and exposure to NOM extracts C. demersum was purchased from Aqua Global (Dr. Jander & Co. OHG, Seefeld, Germany) and cultivated non-axenically in Provasoli’s medium containing de-ionized water, CaCl2 (0.2 g L1), NaHCO3 (0.103 g L1) and sea salt (0.1 g L1) in an aquarium prior to exposure experiments for acclimatization (Pflugmacher et al., 2003). Supplementary light was provided by daylight lamps with an irradiance of 12 mE m2 s, a photoperiod of 14:10 h light:dark cycle and temperature was maintained at 2371 1C. 3.5 g FW of C. demersum was exposed to P. australis and Q. robur extract in serial concentrations of 0.1, 1.0, 10.0, and 100 mg L1 DOC and placed under constant light and temperature conditions for 24 h. In control experiments, only medium water (i.e., no leaf extract) was added. The control as well as the exposures with oak and reed extracts was repeated five times independently.
2.3. Measurement of photosynthesis Photosynthetic oxygen production of the plant was measured using the device Plant Vital 5000 (Inno-Concept, Straussberg). Measurements were made at 100% light intensity (approximately 2000 1 ), a dark/light/dark cycle of 10/12/10 min and at 20 1C. Rates of photosynthetic oxygen production were calculated as mmoles O2sh1g fw1. After measuring photosynthesis, plants were immediately frozen in liquid N2 and kept at 80 1C for enzyme preparation.
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2.4. Enzyme preparation
2.7. Statistical analyses
Enzyme preparation was done according to the method described by Pflugmacher and Steinberg (1997) with slight modification on the steps leading to the microsomal fraction. The frozen plants were ground to a fine powder, suspended in sodium phosphate buffer (0.1 M, pH 6.5) containing 20% glycerol, 1.4 mM dithioerythritol (DTE), and 1 mM ethylenediamine tetraceticacid (EDTA). Cell debris was removed by centrifuging the slurry at 10,000g for 10 min. The residue was suspended in the same buffer and centrifuged again at 5000g for 5 min. The supernatant from the first and second centrifugation was then centrifuged at 40,000g for 60 min. The pellets, containing membrane fractions and defined as the microsomes, was suspended in sodium phosphate buffer (20 mM, pH 7.0) containing 20% glycerol and homogenized using glass homogenizer. The supernatant was precipitated with solid ammonium sulphate in two saturation steps, 0–35% and 35–80%, followed, respectively by two centrifugation steps at 30,000g for 30 min each. The precipitate from the second precipitation which contains soluble proteins were suspended in sodium phosphate buffer (20 mM, pH 7.0) and desalted by gel filtration using NAP-5 columns (Amersham Pharmacia, Germany). Both microsomal and soluble protein extracts were frozen in liquid N2 and stored at –80 1C for enzyme activity assay.
One-way analysis of variance (ANOVA) was performed to test for differences among treatments. Planned post hoc comparisons (Po0.05) were performed using Turkeys HSD test (STASTICA).
2.5. Enzyme activity determination Activity of glutathione S-tranferases (GST) in soluble (cytosol) and microsomal fractions was measured with 1-chloro-2,4-dinitrobenzene (CDNB) as substrate according to Habig et al. (1974). Additionally, GST activity in the cytosol was measured with 4-hydroxynonenal (4HNE) as substrate according to Alin et al. (1984). Peroxidase activity in the soluble fraction was measured using guajacol as substrate (Drotar et al., 1985). Glutathione reductase (GR, E.C. 1.6.4.2) activity in the soluble fraction was measured spectrophotometrically according to Carlberg and Mannervik (1985) based on the oxidation of NADPH to NADP+ which is accompanied by a decrease in absorbance at 340 nm. Enzyme activity was calculated in nano-katals per milligram protein (katal ¼ conversion rate of one mole substrate per second). Protein content of the samples was determined according to Bradford (1976). In special cases where very low protein amounts were achieved in the samples, ‘‘Advanced Protein Assay (ADV01)’’ from Cytoskeleton was used. The assay principle is the same as for Bradford, with the difference that the reagent is five times more concentrated. Serum bovine albumin (initial fraction 98%, Sigma) was used as protein standard for calibration in both assays.
3. Results Direct exposure of the aquatic macrophyte, C. demersum, to extracts of P. australis and Q. robur leaves showed a clear adverse effect on photosynthetic oxygen production (Fig. 1). The effects were significant at all tested concentrations in both leaf extracts. At the highest concentrations (10 and 100 mg L1 DOC), very strong effects were observed with a highly significant (Po0.01) reduction in the rate of photosynthetic oxygen release. Photosynthetic rate of C. demersum was more reduced in Q. robur extract than in P. australis extract especially at the 1 and 10 mg L1 DOC concentrations compared to controls. Measurement of both microsomal (Fig. 2A) and soluble (cytosolic) (Fig. 2B) GST in exposed C. demersum using CDNB as substrate showed a significant (Po0.01) elevation in activity compared to control. Soluble and microsomal GST activity increased with increase in DOC concentration and tends to reach a plateau at higher (10 and 100 mg L1 DOC) concentrations in Q. robur extracts. On the other hand, soluble GST activity in C. demersum exposed to P. australis extract begins to decrease at higher DOC concentrations (10 and 100 mg L1). A concentration-dependent significant (Po0.01) increase in soluble GST activity, using 4-hydroxynonenal (4-HNE) as substrate, was detected in C. demersum exposed to Q. robur extract (Fig. 3). In C. demersum exposed to P. australis extract the increase in activity was also significant but not concentration dependent. Glutathione reductase activity in C. demersum exposed to P. australis and Q. robur extracts increased significantly (Po0.01) compared to untreated control, with a tendency 50
Lipid peroxidation (LPO) was determined according to the User protocol of the lipid peroxidation assay kit (CALBIOCHEM, Cat. No. 437634) with minor modifications based on the method of Botsoglou et al. (1994). The aldehyde, malondialdehyde (MDA), is an end product derived from the breakdown of lipid peroxides, which are often unstable. Estimation of MDA concentration serves as an index of lipid peroxidation. The reaction between N-methyl-2-phenylindole and MDA produces a stable chromophore with maximum absorbance at 586 nm. Plant material was weighed, homogenized in sodium phosphate buffer (20 mM, pH 7.0) and centrifuged for 10 min at 14,000g and at 4 1C. In order to prevent sample oxidation, 10 mL of 0.5 mM butylated hydroxytoluene (BHT) was added. The supernatant was then used for the assay. Six hundred microlitres of N-methyl-2-phenylindole in acetonitrile was added to 200 mL of the sample and gently mixed. One hundred and fifty microlitres of 12 N HCl was then added, mixed again and incubated for 1 h at 45 1C. Samples were cooled on ice and the absorbance measured at 586 nm. A calibration curve was made using MDA.
µmol O2 (gFW)-1 . sh-1
2.6. Measurement of lipid peroxidation 40 ∗ 30
∗ ∗ ∗∗
∗∗
20
∗∗
∗∗
10
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0 control
0.1
1
10
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Dissolved organic carbon (mg . L-1) Fig. 1. Dose-response photosynthetic oxygen production in Ceratophyllum demersum after 24 h exposure to 5 different concentrations of two plant extracts; n ¼ 5, error bars represent7standard deviation, Phragmites australis extract, Quercus robur extract, *Po0.05, **Po0.01.
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4000
35
** 3000 **
**
2000 *
*
*
*
(D)
Enzyme activity * 10-4 (nkat mg-1 protein)
**
1000
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∗
25 20 ∗
15
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∗
∗
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Dissolved organic carbon (mg .
100 L-1)
**
1.6 1.2
0.4
*
**
**
**
0.8
**
Fig. 3. Dose-response activity of soluble (cytosolic) glutathione Stransferase (measured with 4-hydroxynonenal) in Ceratophyllum demersum after 24 h exposure to 5 different concentrations of two plant extracts; n ¼ 5, error bars represent7standard deviation, Phragmites australis extract, Quercus robur extract; *Po0.01, **Po0.001.
*
*
(C) 0.0 6.0
80 ** 60
4.0
**
** ** 2.0
MDA conc. (μ μM)
Enzyme activity (nkat. mg-1 protein)
∗∗
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** *
*
* (B)
40
20
0.0 4.0 *
3.0 *
**
0.0 1
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Fig. 4. Lipid peroxidation (measured as malondialdehyde, MDA, concentration) in Ceratophyllum demersum after 24 h exposure to 5 different concentrations of two plant extracts; n ¼ 3, error bars represent7standard deviation, Phragmites australis extract, Quercus robur extract.
*
*
0.1
control
Dissolved organic carbon (mg . L-1)
(A) control
0
*
*
2.0 1.0
*
100
Dissolved organic carbon (mg . L-1) Fig. 2. Dose-response enzyme activity in Ceratophyllum demersum after 24 h exposure to 5 different concentrations of two plant extracts. (A) microsomal glutathione S-transferase measured with CDNB; (B) cytosolic glutathione S-transferase measured with CDNB; (C) glutathione reductase; (D) guajacol peroxidase; n ¼ 5, error bars represent7standard deviation, Phragmites australis extract, Quercus robur extract, *Po0.01, **Po0.001.
to decrease at the highest (100 mg L1) DOC concentration (Fig. 2C). Activity of guajacol peroxidase in C. demersum increased significantly (Po0.01) at all doses compared with control and in both P. australis and Q. robur extract (Fig. 2D). Elevations in POD activity were higher in C. demersum exposed to Q. robur than that exposed to P. australis extract. Highest elevation compared to control was by a factor of 12 in P. australis exposure and by a factor of 47 in
Q. robur exposure. C. demersum exposed to P. australis showed a gradual decrease in POD activity as the concentration increased from 0.1 to 100 mg L1 DOC. No significant differences (P40.05) in concentration of malondialdehyde (MDA), an index of lipid peroxidation, were detected between control and C. demersum exposed to both Q. robur and P. australis extract at all DOC concentrations tested (Fig. 4). 4. Discussion Photosynthetic oxygen production is an important physiological parameter often used to test the sensitivity of aquatic plants to changes in their surrounding (Sersen et al., 1998; Menone and Pflugmacher, 2005). In the present study, a significant concentration-dependent reduction in rate of photosynthesis of C. demersum after a 24 h
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pre-exposure to both Q. robur and P. australis extracts was observed. Similar effects have been demonstrated in earlier studies using humic substances isolated from soils (Pflugmacher et al., 1999). The mechanism of action for this observation is not clear. However, aromatic structures like phenols and aromatic hydrocarbons are known to have toxic effects on organisms. For example the oxidation of quinone leading to the formation of radicals is known to interfere with electron transfer chain in photosystem II (PSII) in plant chloroplasts (Oettmeier et al., 1988). Using the pure quinone, 1-aminoanthraquinone, photosynthetic oxygen release in Vesicularia dubyana was reduced to almost zero (Pflugmacher et al., 2003). It was also shown in a recent study that the aromatic organic compound, 3-chlorobiphenyl, significantly inhibited the photosynthesis of C. demersum at a concentration of 5 mg L1 (Menone and Pflugmacher, 2005). The significant (Po0.01) reduction in rate of photosynthetic oxygen release due to the oak (Q. robur) extract, which is well known for its high tannin and phenolic contents (Salminen et al., 2004), suggests a possible interference with electron transport chain from PSII to PSI. The adverse effect caused by Q. robur extract was indeed greater than that caused by P. australis extract. Additionally, it is known that when plants are exposed to stress conditions, there is an increase in ROS (Pflugmacher, 2004) and the antioxidant capacity of the plant may be overwhelmed. Organelles such as the peroxisomes and chloroplast (site of photosynthesis), where ROS are being produced at a relatively high rate, are especially at risk (Grene, 2002). Overproduction of ROS in chloroplasts of plants under drought-stress has been reported (Price et al., 1989), and a similar mechanism may occur when plants are exposed to certain concentrations of leaf extracts. The induced activity of antioxidative enzymes observed in the present study suggests an increase in ROS, and the susceptibility of the chloroplasts to high ROS could be an additional factor contributing to the corresponding reduction in the rate of photosynthesis. ROS such as O2d, OH, H2O2 are often internally formed as products of normal plant metabolism (Foyer et al., 1994). Under normal habitat conditions, the rate at which ROS are formed by plants is in dynamic equilibrium with the rate at which they are further utilized or broken down. Leaf extracts might be able to promote oxidative stress by inducing the production of more ROS. To combat the ROS, plants have developed an antioxidative defense system. Parts of this system are the peroxidases (POD), GST and glutathione reductase (GR). In the present study, there was a significant (Po0.01) increase in POD activity in the aquatic macrophyte C. demersum after exposure to both Q. robur and P. australis extracts for 24 h. POD catalyses the reduction of H2O2 to water for detoxification. Exposure of the macrophyte to the leaf extracts may have resulted in an increase in the production of H2O2 beyond the normal detoxifying capacity of the plant. In order to deal with this high level of H2O2 and to prevent or minimize damage to the plant cells,
the plant POD enzymes increased their activity many folds thereby catalyzing the conversion of the highly reactive H2O2 into harmless water molecules. Previous studies with other plant species such as Hydrilla verticillata also demonstrated a significant increase in POD activity after exposure to anthracene in concentrations above 0.01 mg L1 (Byle et al. 1994). Using humic substances from different soil and water sources, Pflugmacher et al. (1999) also showed an increase in POD activity on C. demersum but after 7 days exposure in a concentration of 0.5 mg L1. The very high increase (up to 47 folds) in POD activity observed in the present study was after 24 h exposure of C. demersum to crude leaf extracts. DOC concentrations as low as 0.1 mg L1 provoked significant (Po0.01) increases in POD activity. Aromatic structures and plant growth hormones like gibberellic acid are known to cause an elevation of POD activity (Kwak et al., 1996, Oberg et al., 1990). Response of POD activity upon exposure to leaf extracts from P. australis and Q. robur suggests that these chemicals may contain similar structures. Oak, for example, is well known for its high tannin and phenolic contents (Salminen et al., 2004). The antioxidant activity of phenols is mainly due to their redox properties, which allow them to act as reducing agents, hydrogen donors and singlet oxygen quenchers (RiceEvans et al., 1995). Soluble and microsomal glutathione S-transferase (sGST and mGST) activity was also found to be significantly elevated upon exposure to extracts from Q. robur and P. australis. These enzymes are invariably involved in cellular biotransformation when plants are faced with xenobiotic or toxic substances. They do so by conjugating toxicants or xenobiotics to the sulphurhydryl (–SH) group of glutathione (GSH) which enhances their excretion from cells through cell membranes. Increase in GST activity is thus considered a chemical stress signal. Similarly, Menone and Pflugmacher (2005) demonstrated that soluble GST activity in C. demersum was significantly elevated when treated with lower concentrations (up to 0.5 mg L1) of the organic PCB compound 3-chlorobiphenyl. Imposition of oxidative stress often leads to the conversion of the existing pool of reduced glutathione (GSH) to the oxidized form, glutathione disulphide (GSSG), thereby stimulating glutathione biosynthesis (May and Leaver, 1993). After GSH has been oxidized to GSSG, the recycling of GSSG to GSH is accomplished mainly by the enzyme GR. An elevated glutathione reductase activity detected in the present study was a further indication of an oxidative stress condition in the exposed C. demersum. GSH is used by glutathione peroxidase to detoxify hydrogen peroxide and in the process is converted to GSSG, which is then recycled to GSH by GR. The regenerated GSH is then available for the detoxification of more hydrogen peroxide. An increase in GSH biosynthetic capacity has been shown to enhance resistance to oxidative stress (Zhu et al., 1999). ROS are extremely reactive and unstable chemical species, which react with proteins, nucleic acids, carbohydrates,
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and lipids in cells. The latter often result in lipid peroxidation. Malondialdehyde (MDA) is one of the end products of lipid peroxidation. The findings in the present study did not reveal any significant (P40.05) increase in MDA levels in exposed (treated) C. demersum in all concentrations tested. This is presumably due to activation of defense and detoxification mechanisms, as evident by high GST, GR and POD activities observed in the present study (Figs. 2 and 3). These antioxidant enzymes can decompose ROS and prevent the damage to cellular constituents and initiation of lipid peroxidation (Yang et al., 2003). Furthermore, GST conjugates MDA and 4-HNE using GSH as substrate in the biotransformation process. 5. Conclusion Our results indicate that leaf extracts have the capacity to impose physiological stress and provoke acute antioxidative responses in aquatic organisms, in this case C. demersum. The effects observed within one photoperiodic cycle (24 h) are quite strong. This means that aquatic macrophytes could serve as early warning systems in water quality assessment procedures. Furthermore, we demonstrated that even relatively low DOC concentrations of leaf extracts from P. australis and Q. robur can induce oxidative stress as well as slow down the rate of photosynthesis in the coontail, C. demersum. The effects tend to level off or in some cases start to decrease at high DOC concentrations indicating a threshold on the antioxidative ability of the macrophyte. One of the damaging effects often observed in organisms exposed to stress is lipid peroxidation. In this study, it seems evident that the antioxidative system in C. demersum was sufficiently activated to prevent or minimise this damage since no significant effect on lipid peroxidation was detected in treated plants. Our findings thus suggest that leaf extracts may be an important environmental factor affecting aquatic biota and contributing to the overall ecological dynamics in shallow lakes and rivers. Acknowledgments Our thanks to J. Nimptsch for laboratory assistance and useful discussions on the statistical analyses. The German academic exchange service (DAAD) is gratefully acknowledged for providing financial support for S. Kamara. References Alin, P., Danielson, U.H., Mannervik, B., 1984. 4-hydroxyalk-2-enals are substrates for glutathione transferase. FEBS Lett. 179 (2), 267–270. Bosch, E.V. D., Ward, B.G., Clarkson, B.D., 2004. Woolly Nightsahde (Solanum mauritianum) and its allelopathic effects on New Zealand native Hebe stricta seed germination. In: Proceedings of the 57th New Zealand Plant Protection Society Incorporation Conference, pp. 98–101. Botsoglou, N.A., Fletouris, D.J., Papageorgiou, G.E., Vassilopoulos, V.N., Mantis, A.J., Trakatellis, A.G., 1994. Rapid, sensitive, and
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specific thiobarbituric acid method for measuring lipid peroxidation in animal tissue, food, and feedstuff samples. J. Agric. Food Chem. 42, 1931–1937. Bradford, M.M., 1976. A rapid and sensitive method for the quantification of microgram quantities of protein utilizing the principle of protein dye-binding. Anal. Biochem. 72, 248–254. Brooks, P.D., McKnight, D.M., Bencala, K.E., 1999. The relationship between soil heterotrophic activity, soil dissolved organic carbon (DOC) leachate, and catchment-scale DOC export in headwater catchments. Water Resour. Res. 35, 1895–1902. Byle, T.D., Sutton, H.D., Klaine, S.J., 1994. Evaluation of peroxidase as a biochemical indicator of toxic chemical exposure in the aquatic plant Hydrilla verticillata. Royle. Environ. Toxicol. Chem., 509–515. Carlberg, I., Mannervik, B., 1985. Glutathione reductase. Methods Enzymol. 113, 484–490. Cole, J.J., Pace, M.L., Carpenter, S.R., Kitchell, J.F., 2000. Persistence of net heterotrophy in lakes during nutrient addition and food web manipulations. Limnol. Oceanogr. 45 (8), 1718–1730. DIN EN 1484, 1998. Anleitung zur Bestimmung des gesamten organischen Kohlenstoffs (TOC) und des gelo¨sten organischen Kohlenstoffs. DEV, H3, 40, Lieferung. Drotar, A., Phelps, P., Fall, R., 1985. Evidence for glutathione peroxidase activities in cultured plant cells. Plant Sci. 42, 35–40. Feeny, P.P., 1970. Seasonal changes in oak leaf tannins and nutrients as a cause of spring feeding by winter moth caterpillars. Ecology 51, 565–581. Foyer, C.H., Lescure, J.C., Lefebvre, C., Morot-Guadry, J.F., Vincent, M., Vaucheret, H., 1994. Adaptations of photosynthetic electron transport, carbon assimilation and carbon partitioning in transgenic Nicotiana plumbaginifolia plants to changes in nitrate reductase activity. Plant Physiol. 104, 171–178. Grene, R., 2002. Oxidative stree and acclimation mechanisms in plants. The Arabidopsis Book. Am. Soc. Plant Biol., 1–20. Grundho¨fer, P., Niemetz, R., Schilling, G., Gross, G.G., 2001. Biosynthesis and subcellular distribution of hydrolyzable tannins. Phytochemistry 57, 915–927. Habig, W., Pabst, M.J., Jakoby, W.B., 1974. Glutathione S-transferase: the first step in mercapturic acid formation. J. Biol. Chem. 249, 1730–1739. Hollis, G.E., Jones, T.A., 1991. Europe and the Mediterranean Basin. In: Finlayson, M., Moser, M. (Eds.), Wetlands, pp. 27–56. Hunter, M.E., Menges, E.S., 2002. Allelopathic effects and root distribution of Ceratiola ericoides (Empetraceae) on seven Rosemary scrub species. Am. J. Bot. 89 (7), 1113–1118. Jeppeson, E., Sondergaard, M., Kronvang, B., Jensen, J.P., Svendsen, L.M., Lauridsen, T.L., 1999. Lake and catchment management in Denmark. Hydrobiologia 395/396, 419–432. Kruse, M., Strandberg, M., Strandberg, B., 2000. Ecological Effects of Allelopathic Plants—a Review. National Environmental Research Institute, Silkeborg, Denmark, 66pp., NERI Technical Report No. 315. Kwak, S.S., Kim, S.K., Park, I.H., Liu, J.R., 1996. Enhancement of peroxidase activity by stress-related chemicals in sweet potato. Phytochemistry 43, 565–568. May, M.J., Leaver, C.J., 1993. Oxidative stimulation of glutathione synthesis in Arabidopsis thaliana suspension cultures. Plant Physiol. 103, 621–627. Menone, M.L., Pflugmacher, S., 2005. Effects of 3-chlorobiphenyl on photosynthetic oxygen production, glutathione content and detoxication enzymes in the aquatic macrophyte Ceratophyllum demersum. Chemosphere 60, 79–84. Meyer, J.L., Wallace, J.B., Eggert, S.L., 1998. Leaf litter as a source of dissolved organic carbon in streams. Ecosystems 1, 240–249. Nimptsch, J., Wunderlin, D.A., Dollan, A., Pflugmacher, S., 2005. Antioxidant and biotransformation enzymes in Myriophyllum quitense as biomarkers of heavy metal exposure and eutrophication in Suquia River basin (Cordoba, Argentina). Chemos 61, 147–157.
ARTICLE IN PRESS 246
S. Kamara, S. Pflugmacher / Ecotoxicology and Environmental Safety 67 (2007) 240–246
Oberg, L.G., Glas, B., Swanson, S.E., Rappe, C., Paul, K.G., 1990. Peroxidas-catalyzed oxidation of chlorophenols to polychlorinated dibenzo-p-dioxins and dibenzofurans. Arch. Environ. Contam. Toxicol. 19, 930–938. Oettmeier, W., Masson, K., Donner, A., 1988. Anthraquinone inhibitors of photosystem II electron transport. FEBS Lett. 231, 259–262. Pflugmacher, S., Steinberg, C.E.W., 1997. Activity of phase I and phase II detoxication enzymes in aquatic macrophytes. J. Appl. Bot. 71, 144–146. Pflugmacher, S., Spangenberg, M., Steinberg, C.E.W., 1999. Dissolved organic matter (DOM) and effects on the aquatic macrophyte Ceratophyllum demersum in relation to photosynthesis, pigment pattern and activity of detoxication enzymes. J. Appl. Bot. 73, 184–190. Pflugmacher, S., Wiegand, C., Beattle, K.A., Krause, E., Steinberg, C.E.W., Codd, G.A., 2000. Uptake, effects and metabolism of cyanobacterial toxins in the emergent reed plant Phragmites australis (Cav.). Trin. Ex. Steud. Environ. Toxic. Chem. 20 (4), 846–852. Pflugmacher, S., 2002. Possible allelopathic effects of cyanotoxins, with reference to Microcystin LR, in aquatic ecosystems. Wiley Period, Inc. Environ. Toxicol. 17, 407–413. Pflugmacher, S., Tidwell, L.F., Steinberg, C.E.W., 2001. Dissolved humic substances directly affect freshwater organisms. Acta Hydrochim. Hydrobiol. 29, 34–40. Pflugmacher, S., Pietsch, C., Reiger, W., Paul, A., Preuer, T., Zwirnmann, E., Steinberg, C.E.W., 2003. Humic substances and their direct effects on the physiology of aquatic plants. In: Ghabbour, E.A., Davies, G. (Eds.), Humic Substances: Nature’s Most Versatile Materials. Taylor & Francis, New York. Pflugmacher, S., 2004. Promotion of oxidative stress in the aquatic macrophyte Ceratophyllum demersum during biotransformation of the cyanobacterial toxin microcystin-LR. Aquat. Toxicol. 70, 169–178. Polunin, N.V.C., 1984. The decomposition of emergent macrophytes in freshwater. Adv. Ecol. Res. 14, 115–166.
Price, A.H., Atherton, N.M., Hendry, G.A.F., 1989. Plants under drought-stress generate activated oxygen. Free Radical Res. Commun. 8, 61–66. Rice-Evans, C.A., Miller, N.J., Bolwell, P.G., Bramley, P.M., Pridham, J.B., 1995. The relative antioxidant activities of plant-derived polyphenolic flavonoids. Free Radical Res. 22, 375–383. Riipi, M., Ossipov, V., Lempa, K., Haukioja, E., Koricheva, J., Ossipova, S., Pihlaja, K., 2002. Seasonal changes in birch leaf chemistry: are there trade-offs between leaf growth and accumulation of phenolics. Oecologia 130, 380–390. Salminen, J., Roslin, T., Karonen, M., Sinkkonen, J., Pihlaja, K., Pulkkinen, P., 2004. Seasonal variation in the content of hydrolizable tannins, flavonoid glycosides and proanthocyanidins in oak leaves. J. Chem. Ecol. 30 (9), 1693–1711. Scalbert, A., Haslam, E., 1987. Plant polyphenols and chemical defense. Part 2: Polyphenols and chemical defense of the leaves of Quercus robur. Phytochemistry 26, 3191–3195. Scalbert, A., Monties, B., Favre, J.M., 1988. Polyphenols of Quercus robur: adult tree and in vitro grown calli and shoots. Phytochemistry 27, 3483–3488. Sersen, F., Kralova, K., Bumbalova, A., 1998. Action of mercury on the photosynthetic apparatus of spinach chloroplasts. Photosynthetica 35, 551–559. Wetzel, R.G., 1995. Death, detritus and energy flow in aquatic ecosystems. Freshwater Biol. 33, 83–89. Wetzel, R.G., 2001. Limnology: lake and river ecosystems, third ed. Academic Press, San Diego, pp. 527–664. Yang, Y., Sharma, R., Sharma, A., Awasthi, S., Awasthi, Y.C., 2003. Lipid peroxidation and cell cycle signaling: 4-hydroxynonenal, a key molecule in stress mediated signaling. Acta Biochim. Polon. 50 (2), 319–336. Zhu, Y.L., Philon-Smits, E.A.H., Tarun, A.S., Weber, S.U., Jouanin, L., Terry, N., 1999. Cadmium tolerance and accumulation in Indian mustard is enhanced by overexpressing gamma-glutamylcysteine synthetase. Plant Physiol. 121, 1169–1178.