VILNIUS UNIVERSITY INSTITUTE of BOTANY of NATURE RESEARCH CENTRE Reda Grigutytė NITELLOPSIS OBTUSA (DESV.) J. GROVES CELL RESPO

VILNIUS UNIVERSITY INSTITUTE OF BOTANY OF NATURE RESEARCH CENTRE

Reda Grigutytė

NITELLOPSIS OBTUSA (DESV.) J. GROVES CELL RESPONSE TO ALLOCHTONOUS STRESSORS OF FAGUS SYLVATICA L. AND QUERCUS ROBUR L. LEAF LITTER EXTRACTS

Doctoral Dissertation Biomedical sciences, botany (04 B)

Vilnius, 2010 Dissertation was performed in 2005–2009 at the Institute of Botany of Nature Research Centre (Lithuania) and Leibniz–Institute of Freshwater Ecology and Inland Fisheries (Germany).

Scientific supervisor – Dr. Levonas Manusadžianas (Institute of Botany of Nature Research Centre, biomedical sciences, biophysics – 02 B)

Scientific advisor – PD. Dr. Stephan Pflugmacher (Leibniz–Institute of Freshwater Ecology and Inland Fisheries, biomedical sciences, biology – 01 B)

VILNIAUS UNIVERSITETAS GAMTOS TYRIMŲ CENTRO BOTANIKOS INSTITUTAS

Reda Grigutytė

NITELLOPSIS OBTUSA (DESV.) J. GROVES LĄSTELIŲ ATSAKAS Į ALOCHTONINIUS STRESORIUS – FAGUS SYLVATICA L. IR QUERCUS ROBUR L. LAPŲ PAKLOTĖS EKSTRAKTUS

Daktaro disertacija Biomedicinos mokslai, botanika (04 B)

Vilnius, 2010

Disertacija buvo rengiama 2005–2009 metais Gamtos tyrimų centro Botanikos institute ir Leibnico gėlųjų vandenų ir gėlavandenių žuvų ekologijos institute.

Mokslinis vadovas – Dr. Levonas Manusadžianas (Gamtos tyrimų centro Botanikos institutas, biomedicinos mokslai, biofizika – 02 B).

Mokslinis konsultantas – PD. Dr. Stephan Pflugmacher (Leibnico gėlųjų vandenų ir gėlavandenių žuvų ekologijos institutas, biomedicinos mokslai, biologija – 01 B).

TABLE OF CONTENTS List of abbreviations...... 8 Introduction ...... 10 1. Literature overview ...... 15 1.1. Some egzogenic sources of natural organic matter ...... 15 1.1.1. F. sylvatica leaf litter ...... 16 1.1.2. Q. robur leaf litter ...... 17 1.2. Freshwater charophyte, N. obtusa ...... 18 1.3. Reactive oxygen species ...... 19 1.4. Oxidative stress and defense mechanism in plants ...... 22 1.4.1. Glutathione S-transferase (GST: EC 2.5.1.18) ...... 24 1.4.2. Catalase (CAT: EC 1.11.1.6) ...... 26 1.4.3. Glutathione reductase (GR: EC 1.8.1.7) ...... 27 1.4.4. Glutathione peroxidase (GPx: EC 1.11.1.9) ...... 27 1.4.5. Superoxide dismutase (SOD: EC 1.15.1.1) ...... 27 1.5. Hydrogenperoxide, total antioxidant status and lipid peroxidation .... 28 1.6. Electrophysiological reaction of N. obtusa cells ...... 30 1.7. Lethality, as a measure of toxicity ...... 32 2. Methods...... 33 2.1. Plant material ...... 33 2.2. Preparation of Q. robur and F. sylvatica leaf extracts ...... 33 2.3. Leaf litter degradation extracts (LLDEs) ...... 34 2.4. Chemical characterization of leaf extracts and Lake water ...... 34 2.5. Dose- and time-dependent exposure of N. obtusa to Q. robur and F. sylvatica leaf extracts...... 35 2.6. Exposure of charophyte cells to LLDEs ...... 36 2.7. Protein extraction ...... 36 2.8. Measurement of enzymatic activities ...... 37 2.8.1. Catalase ...... 37 2.8.2. Glutathione S-transferase ...... 37 2.8.3. Glutathione reductase ...... 38 5

2.8.4. Glutathione peroxidase ...... 39 2.8.5. Calculation of specific enzymatic activity by photometric measurement ...... 39 2.8.6. Superoxiddismutase ...... 40 2.8.7. Measurement of cell internal hydrogen peroxide ...... 41 2.8.8. Measurement of the total antioxidant status ...... 42 2.8.9. Measurement of lipid peroxidation ...... 42 2.9. Electrophysiological experiments ...... 43 2.10. Cell Lethality testing ...... 45 2.11. Statistical analysis...... 45 3. Results ...... 46

3.1. Enzymatic activity, H2O2 content, total antioxidant content and lipid peroxidation in N. obtusa ...... 46 3.1.1. Dose – response effects induced by F. sylvatica and Q. robur leaf extracts ...... 46 3.1.1.1. Microsomal glutathione S-transferase assayed with the conjugating substrate 1-chloro-2,4-dintrobenzene (CDNB) .... 46 3.1.1.2. Cytosolic glutathione S-transferase assayed with the conjugating substrate CDNB...... 47 3.1.1.3. Catalase ...... 48 3.1.1.4. Glutathione reductase ...... 49 3.1.1.5. Glutathione peroxidase ...... 50 3.1.1.6. Superoxide dismutase ...... 51 3.1.1.7. Hydrogen peroxide content ...... 52 3.1.1.8. Total antioxidant content ...... 53 3.1.1.9. Lipid peroxidation ...... 53 3.1.2. Time-dependent effects induced by F. sylvatica and Q. robur leaf extracts ...... 54 3.1.2.1. Microsomal glutathione S-transferase, measured with CDNB 54 3.1.2.2. Cytosolic glutathione S-transferase, measured with CDNB .... 55 3.1.2.3. Catalase ...... 57 6

3.1.2.4. Glutathione reductase ...... 58 3.1.3. Effects induced by 30-day-decomposing of leaf litter extracts from F. sylvatica and Q. robur ...... 59 3.1.3.1. Dissolved organic carbon content ...... 59 3.1.3.2. Cytosolic glutathione S-transferase, assayed with the conjugating substrate CDNB...... 60 3.1.3.3. Catalase ...... 61 3.1.3.4. Glutathione reductase ...... 62 3.1.3.5. Hydrogen peroxide content ...... 63 3.2. The electrophysiological response of N. obtusa cells affected by F. sylvatica and Q. robur leaf extracts ...... 64 3.3. The lethality response of N. obtusa cells exposed to F. sylvatica and Q. robur leaf extracts ...... 65 4. Discussion ...... 67 Conclusions ...... 75 Acknowledgements ...... 77 References ...... 78 Research articles and congress presentations ...... 92

List of abbreviations

AFW Artificial freshwater ANOVA Analysis of variance APW Artificial pond water APX Ascorbate peroxidase BLE Fagus sylvatica leaf extract CAT Catalase CDNB 1-chloro-2,4-dinitrobenzene cGST Cytosolic GST DNA Deoxyribonucleic acid DOC Dissolved organic carbon DTE Dithioerythritol DTPA Diethylenetriaminepentaacetic acid DW Dry weight EDTA Ethylenediaminetetracetic acid g Gram GAPDH Glyceraldehyde-3-phosphate dehydrogenase GPx Glutathione peroxidase GR Glutathione reductase GSH Reduced glutathione GSSG Glutathione disulphide GST Glutathione S-transferase

H2O2 Hydrogen peroxide HNE Hydroxynonenal HS Humic substances L Litre LLDEs Leaf litter degradation extracts LPO Lipid peroxidation MDA Malondialdehyde mg Milligram

8 mGST Membrane bound GST mL Millilitre NADPH Reduced nicotinamide adenine dinucleotide phosphate NAP-5 Sephadex G-25 Column nkat Nano katal NOM Natural organic matter OLE Quercus robur leaf extract RNA Ribonucleic acid ROS Reactive oxygen species SOD Superoxide dismutase TOSC Total antioxidant status μL Microlitre

Introduction

Lake ecosystems experience various influences from terrestrial surroundings. One of these includes the input of natural organic material (NOM) originated, for example, from leaves falling into the water body

(GRIMM et al., 2003). The leaves then degrade in different steps, beginning with macro-organisms and ending with micro-organisms (BESS, 2005). Chemistry of aquatic environment partially depends on the decomposition of foliar litter and coarse woody debris as a source of dissolved organic carbon

(DOC), e.g. for the foundation of aquatic food webs (SCHLESINGER, 1997). The structural and chemical attributes of leaves can vary dramatically among species (CHAUVET, 1987; OSTROFSKY, 1997), so their breakdown may be related to the number and type of chemically unique species present in the matrix. Particulate detritus interacts with the biotic, i.e., bacteria, fungi, invertebrates (CUMMINS et al., 1989) and abiotic, i.e., temperature, water flow, soil/water chemistry (ALLARD, MOREAU, 1986; WEBSTER, BENFIELD, 1986) components of ecosystems and thus regulates the rate of breakdown. In terrestrial systems, leaf species containing low C:N content tend to stimulate the breakdown rate of leaf litter, whereas leaves with compounds such us phenolics and tannins) tend to slow breakdown rate in mixed-litter assemblages

(SWAN, PALMER, 2004). Decomposition of leaf litter has great influence on biochemical nutrient cycling, also, in all surface waters, interacts with aquatic macrophytes and is able to produce reactive oxygen species (PFLUGMACHER et al., 1999). During leaf degradation, many water-soluble chemical compounds are released into the water, which may influence aquatic organisms. A part of these compounds are quinoid structures, capable of generating reactive oxygen species (ROS) and influence, for instance, photosynthesis of aquatic plants by capturing electrons from the plastoquinone chain between photosystem II and -• photosystem I (BULYCHEV, 2001). Enhanced production of ROS (e.g. O2, O2 , • H2O2 and HO ) destroys their tiny balance in the cell. To prevent cellular damage caused by these ROS, cells have developed a protective system 10 involving antioxidative enzymes, including superoxide dismutase (SOD), catalase (CAT), glutathione S-transferase (GST), glutathione reductase (GR), etc. The enzymes take part in different phases of ROS elimination. The function of SOD is to dismutate two superoxide anion radicals into oxygen and hydrogen peroxide, while CAT reduces hydrogen peroxide to water. GR is a part of an oxido-reduction cycle involving glutathione and ascorbate (APEL,

HIRT, 2004; NOCTOR et al., 2002). GR catalyses the reduction of glutathione disulfide (GSSG) in a NADPH-dependent reaction to glutathione (GSH) and, therefore, plays a pivotal role in protection against oxidative damage and in adjustment processes of metabolic pathways (like redox homeostasis) (FOYER,

NOCTOR, 2005). Beside the biotransformation of exogenous xenobiotics, the glutathione S-transferase plays an important role in detoxifying endogenous toxic lipid peroxidation products (AWASTHI et al., 2004). Recently, the first studies on leaf litter induced oxidative stress in aquatic plants have been introduced. NIMPTSCH and PFLUGMACHER (2008) showed that leaf litter degradation extracts (LLDEs) from common oak (Quercus robur L.), European beech (Fagus sylvatica L.), and mixed Q. robur and F. sylvatica leaves had deleterious effects on the physiology of Vesicularia dubyana (C. Muell.) C. Muell., expressed as decreasing photosynthetic activity and enhancing oxidative stress response. KAMARA and

PFLUGMACHER (2007) investigated the impact of extracts from Phragmites australis (cav.) Trin. ex Steud and Q. robur leaves on the antioxidative system and photosynthetic rate of Ceratophyllum demersum L. These findings suggest that leaf litter degradation extracts may be an important environmental factor influencing specific plant species and thus community structure within freshwater ecosystems. Some water plants (V. dubyana or C. demersum) seem to be intolerant to leaf litter. We investigated responses of the cell of starry stonewort (Nitellopsis obtusa (Desv.) J. Groves). N. obtusa represents multicellular macrophytic green algae, which belongs to Charophyceae class, Nitellopsidaceae family. The internodal cells of charophytes are intensively used for studies in electrophysiology (see 11

MIMURA, 1995); ion transport (TAZAWA et al., 1987); cytoplasmic streaming

(PLIETH, HANSEN, 1992); intercellular transport (SHEPHERD, GOODWIN, 1992); electrical characteristics of plant cell membranes (SHIMMEN et al., 1994). The rapid depolarization of resting potential (RP) proved to be a relevant endpoint to assess chemicals and wastewater toxicity, and tight correlation between electrophysiological response (90-min 50%-depolarization of RP) and cell lethality (96-h LC50) induced by above chemical stressors has been shown (MANUSADŽIANAS et al., 1995; 2007). N. obtusa communities have a cosmopolitan distribution and are generally found in clear and oligo- or mesotrophic waters (PYBUS et al., 2003). They are good indicators of clear and nutrient-poor water (RUTH, PALAVAGE, 1994). The decomposition rate of leaf litter influenced by various ecological factors has been aptly investigated in terrestrial ecosystems (GESSNER, 2000;

TIEGS ET AL., 2008), however, the potential effects of the decomposition products for aquatic biota have not received parallel attention. General idea of the study: By linking lake and lake shore ecosystems and in accordance with the fact that leaf litter decomposition products are an additional source of allochthonous dissolved organic carbon, we sought to evaluate the impact of F. sylvatica and Q. robur leaf litter extracts on algae N. obtusa cell. Objectives of the study: 1. To prove the fact that N. obtusa exposed to leaf litter extracts experiences oxidative stress, by evaluating activities of the enzymes such as GST, CAT, GR, GPx and SOD, total antioxidant status of the

cells, lipid peroxidation and changes of H2O2 concentration. 2. To explore the effects of F. sylvatica and Q. robur leaf litter extracts on N. obtusa cell membrane resting potential and its lethality. 3. To investigate the influence of F. sylvatica and Q. robur leaf litter degradation extracts obtained during 30-day decomposition on oxidative

stress enzymes such as GST, CAT and GR, and H2O2 content in algae N. obtusa. 12

Defensive statements: 1. Leaf extracts from F. sylvatica and Q. robur induced dose- and time- dependent oxidative stress-like response of enzymes in N. obtusa. Detoxification reactions were expressed by changes in enzyme activities due to cleavage of reactive oxygen species. Analogous changes in enzymes activity occurred in experiments imitating natural conditions, when algae cells were affected by extracts from self-decomposing leaf litter. 2. Both leaf extracts, Q. robur (245 – 1838 mg L-1 DOC) and F. sylvatica (330–1100 mg L-1 DOC) induced fast depolarization of cell membrane resting potential after 15-min. exposure, which was permanent till the end of 24 h exposure. At lower concentrations typical to natural freshwater bodies, they invoked adaptive electrophysiological reactions. 3. Reactions of charophytic algae N. obtusa to Q. robur and F. sylvatica leaf extracts were detected at different biological levels. Relatively more sensitive reactions were detected at enzymatic level, however, to identify irreversible cell resting potential changes and lethality, higher than those of natural aquatic systems DOC concentrations or prolonged exposure are needed. Novelty and significance of the study: 1. An antioxidative status of charophyte N. obtusa cell exposed to F. sylvatica and Q. robur leaf extracts and to obtained during 30 days extracts of decomposing Q. robur and F. sylvatica leaves was investigated for the first time. 2. The responses of charophyte N. obtusa to F. sylvatica and Q. robur leaf litter extracts at different biological levels – cell, cell membrane (plasmalemma) and oxidative stress enzymes was investigated for the first time. Presentation of the results: The obtained research data were presented in 4 scientific papers published in peer-reviewed scientific journals. The investigation results were presented at 5 international conferences.

Volume and the structure of the dissertation: The doctoral dissertation is written in English and includes List of abbreviations, Introduction, Methods, Results and Discussion, Conclusions, Acknowledgements, References (139 citations), Research articles and congress presentations. The dissertation is illustrated with 43 figures, 2 tables. Volume of dissertation is 93 pages.

1. Literature overview

1.1. Some egzogenic sources of natural organic matter

The main sources of natural organic matter in aquatic systems depend on the location and land-use patterns in terrestrial environment. Headwaters of lotic systems may be dominated by erosional and allochthonous inputs, whereas natural lake systems are mostly dominated by autochthonous inputs. Particulate organic matter in forested headwater streams and rivers is derived mainly from terrestrial vegetation and may contribute up to 800 g m-2 year-1

(WARD et al., 1994). Leaf litter and woody debris constitute a significant component of the organic matter inputs and may reach the aquatic system directly by dropping from overhanging vegetation or may be transported from forest floor. FRANCE (1996) reported that riparian deforestation due to clear- cutting resulted in considerable reductions in dissolved organic carbon, which in turn could potentially result in decreased metabolic processes of lake plankton and vegetation communities too. The decomposition of leaf litter originates natural organic matter within freshwater ecosystems. NOM is present in all surface waters and moreover it is interacting with aquatic macrophytes and is able to produce reactive oxygen species (PFLUGMACHER et al., 1999). Around 90% of the dissolved carbon in water ecosystem consists of humic substance (LINKOV et al., 2007). Humic substance (HS) can be divided into three components: fluvic acids, humic acids and humin. HS adversely affect the quality of water since they impart colour and serve as precursors for the formation of chlorinated compounds (MAMBA et al., 2009). They also have complexing properties that include association with toxic elements and pollutants (DE WUILLOUD et al., 2003). NOM is usually found in drinking water at concentration levels of between 2 and -1 15 mg L (HEPPLEWHITE et al., 2004). In most freshwater ecosystems, dissolved organic carbon (DOC) concentrations often range between 1 and -1 100 mg L (WETZEL, 2001), but in some case can be and higher concentrations -1 (up to 300 mg L ) (BLODAU et al., 2004). Basic structures are created from 15 cellulose, tannin, cutin and lignin, along with other various proteins, lipids, and sugars. NOM is very important in the movement of nutrients in the environment and plays a role in water retention on the surface of the planet. The only information scientists have is that NOM is heterogeneous and very complex. Generally, NOM, in terms of weight, is: 45-55% Carbon, 35-45%

Oxygen, 3-5% Hydrogen, 1-4% Nitrogen (CABANISS et. al., 2007). The molecular weights of these compounds can vary drastically, depending on if they repolymerize or not. It is also important to know that 10-35% of the carbon present forms aromatic rings (CABANISS et. al., 2007). These rings are very stable due to resonance stabilization, so they are difficult to break down. The aromatic rings are also susceptible to electrophilic and nucleophilic attack from other electron-donating or electron-accepting material, which explains the possible polymerization to create larger molecules of NOM. In the present study, F. sylvatica and Q. robur species were used as allochthonous sources of NOM.

1.1.1. F. sylvatica leaf litter

Figure 1.1 F. sylvatica leaf (photo by T. Arboretum).

Leaf litter is one of the main inputs of dissolved organic carbon and nutrients in freshwater ecosystems. The majorities of leaf litter enter aquatic systems during the fall as autumn-shed leaves; although large quantities of leaf litter can also enter aquatic systems during the spring melt (POPE et al., 1999). Leaves are making an important allochthonous contribution to the organic

16 matter input in streams, rivers and lakes. The change of nutrient circulation and litter decomposition can be chemical, physical and biotic, individually or in combination. Freshwater ecosystems experience various influences from terrestrial surroundings (HANLON, 1982). One of the terrestrial species that enter into the aquatic environment via leaf litter fall is F. sylvatica. These leaves (Figure 1.1) are known to contain phenolic compounds, have high nitrogen concentrations, low concentrations of carbon-based plant protection compounds such as tannins and lignin when grown on fertile soils

(SARIYILDIZ, ANDERSON, 2005). When leaves are falling into water and degrade, biotic and abiotic processes are involved in the formation and removal of dissolved organic matter during the decomposition process (SAID-

PULLICINO et al., 2007). Many compounds are released into the water, which may have direct or indirect influence on the organisms within the aquatic ecosystem.

1.1.2. Q. robur leaf litter

Figure 1.2 Q. robur leaf (photo by H. Kettunen).

Leaf litter falling into the aquatic environment is only one of many components of the connectivity between terrestrial and aquatic ecosystems. Due to its seasonality, this phenomenon is repeated every year. Deciduous Q. robur trees leaf (Figure 1.2) makes an important allochthonous contribution to the organic matter input in aquatic ecosystems. It is found that hydrolysable tannins are the dominant group of phenolic compounds in the leaves of the

Q. robur species Q. robur (SWAN, PALMER, 2004; SALMINEN et al., 2004). A rare dimeric ellagitannin, cocciferin D2, was also detected for the first time in the leaves of Q. robur. A few earlier studies have shown that the leaves of Q. robur contain flavonol glycosides, castalagin, vescalagin and casuarictin

(SCALBERT et al., 1988; GRUNDHOFER et al., 2001).

1.2. Freshwater charophyte, N. obtusa

N. obtusa is a plant which is known to be intolerant to humic substrates, is easy to analyze at different organism levels, and is in detail described in the literature on ecotoxicology and, therefore, an excellent object for our study.

Figure 1.3 N. obtusa (photo by Klaus van de Weyer).

Taxonomic hierarchy Kingdom Protista Phylum Chlorophyta Class Charophyceae Order Charales Family Characeae Genus Nitellopsis Species N. obtusa Characeae (charophytes or stoneworts) are anatomically highly developed green algae, easily recognized by their typical morphology (Figure 1.3). Stoneworts used to be classified as members of the plant kingdom, but it

18 is now agreed that they belong, with other green algae, in the kingdom Protista. Put simply, the protistas are simple multi-celled or single celled organisms, descended from some of the earliest life-forms that appeared on Earth. Algae are generally found in oligo- or mesotrophic waters (PYBUS et al., 2003; KAIRE et al., 2004). Characeae are recognizable by long, uneven-length branches that appear angular at each node, and may have one cream-colored bulb at the base of each cluster of branches. The trunk of the plant consists of several ‘giant’ cells, up to several centimeters long, which effectively make up the stem. The stonewort anchors itself, not with roots like a plant, but with rhizoids, colorless, hair-like filaments. Like roots, these can absorb nutrients, but the organism can absorb them through its entire surface. Characeae communities have a cosmopolitan distribution and are commonly known as indicators of clear and nutrient-poor water, but high in calcium, consequently being good indicators of water quality. The bioelectrical response of charophyte cell is rapid and highly sensitive to chemical compounds and can be used as a relevant end point to assess aquatic sample toxicity (MANUSADŽIANAS et al.,

1999; MANUSADŽIANAS et al., 2002; JURKONIENĖ et al., 2004). N. obtusa is usually a summer annual, but in some years, if the winter is mild, it may not fully die back. This species very rarely produces spores. It is thought that spore production is controlled by light levels, and tends to take place from July to September.

1.3. Reactive oxygen species

Reactive oxygen species have been of interest for many years in all areas of biology. ROS are ions or very small molecules that include oxygen ions, free radicals, and peroxides, both inorganic and organic (GRENE, 2002). They are highly reactive electrons. Various enzyme systems produce ROS, including the mitochondrial electron transport chain, cytochrome P450, lipoxygenase, cyclooxygenase, the NADPH oxidase complex, xanthine oxidase, and peroxisomes (INOUE et al., 2003).

NAD(P)H o xidases

xanthine oxidases catalase •− O2 O H O H O 2 superoxide 2 2 GSH 2 hypoxanthine xanthine dismutase persoxidase xanthine uric acid (SOD)

semiubiquinone * GSH GSSG

mitochondria GSH reductase NADP+ NADPH

Figure 1.4 Pathways of reactive oxygen species (ROS) production and defenses.

ROS form as a natural byproduct of the normal metabolism of oxygen and have important roles in cell signaling. However, during times of environmental stress ROS levels can increase dramatically, which can give damage to cell structures (Figure 1.4). This cumulates into a situation known as oxidative stress (FOYER, NOCTOR, 2005).

O − + 2 H+ + e Hydrogen peroxide 2 H2O2 Hydroxil radical H2O2 + e OH − + • OH

+ • O2 + e + H HO2 Hydroperoxyl radical • HO2 H+ + O − Superoxide radical 2 Figure 1.5 Reactions generating reactive oxygen species (ROS).

Superoxide and hydrogen peroxide are much less reactive than OH•. The main risk for a cell that produces the two former reactive oxygen intermediates may be posed by the intermediates interaction, leading to the generation of highly reactive hydroxyl radicals (Figure 1.5). Because there are no known scavengers of hydroxyl radicals, the only way to avoid oxidative damage through this radical is to control the reactions that lead to its generation. Thus, cells had to evolve sophisticated strategies to keep the concentrations of superoxide, hydrogen peroxide, and transition metals such as 20

Fe and Cu under tight control. ROS are not only dangerous; it can be also functioning as signaling molecules (AWASTHI et al., 2004). It may interact selectively with a target molecule that perceives the increase of ROS concentration and translates this information into signals that direct the plant’s responses to stress.

ROS are small and can diffuse short distances (APEL, HIRT, 2004). Small molecule antioxidants such as ascorbic acid (vitamin C), tocopherol (vitamin E), uric acid and glutathione play important roles as cellular antioxidants (Figure 1.6). Similarly, polyphenol antioxidants assist in preventing ROS damage by scavenging free radicals (DROGE, 2002).

a) b)

O NH O H 2 N NH HO NH O NH O OOHH OH N H N O c) H d) SH Figure 1.6 ROS antioxidants: a) ascorbic acid; b) tocopherol; c) uric acid; d) glutathione.

In addition to energy, reactive oxygen species are produced which have the potential to cause cellular damage (APEL, HIRT, 2004; FOYER, NOCTOR,

2005; SCANDALIOS, 2005): damage of DNA, RNA, oxidations of polydesaturated fatty acids in lipids, oxidations of amino acids in proteins, oxidatively inactivate specific enzymes by oxidation of co-factors.

One major contributor to oxidative damage is hydrogen peroxide (H2O2), which is converted from superoxide that leaks from the mitochondria. To prevent cellular damage caused by these ROS, cells have developed a protective system involving antioxidative enzymes, including CAT, GST and GR between others

(BLOKHINA, et al., 2003). CAT reduces hydrogen peroxide to water. GR is part of an oxido-reduction cycle involving glutathione and ascorbate (APEL, HIRT,

2004; NOCTOR et al., 2002), catalyzing the reduction of glutathione disulfide (GSSG) in a NADPH-dependent reaction to glutathione (GSH), playing a pivotal role in protection against oxidative damage and adjustment processes of metabolic pathways (FOYER, NOCTOR, 2005), however this conversion is not 100% efficient, and residual peroxides persist in the cell.

1.4. Oxidative stress and defense mechanism in plants

Many environmental conditions induce an oxidative stress in plant cells by the generation of abnormal concentrations of reactive-oxygen species in cell compartments. It can cause damage on membrane systems, proteins and nucleic acids. ROS are tightly regulated and participating in the control of processes such as programmed cell death, stress response, and pathogen defence. Under optimal conditions the production of ROS in cells is estimated -1 – at a constant rate of 240 μM s O2 , under stress the production of ROS can raise -1 – up to 720 μM s O2 . These include drought and desiccation, salt, chilling, heat shock, heavy metals, UV radiation, air pollutants such as ozone, mechanical stress, nutrient deprivation, pathogen attack, and high light. During stress ROS may be viewed as cellular by-products of stress metabolism, as well as secondary messengers involved in the stress-response signal transduction pathway. ROS are produced by plant cells via the enhanced enzymatic activity of plasma membrane-bound NADPH oxidases, cell wall-bound peroxidases and amine oxidases in the apoplast. H2O2 produced during this response is thought to diffuse into cells through aquaporins and together with salicylic acid and nitric oxide activate many of the plant defences, including the induction of

22 programmed cell death. To protect themselves against ROS potentially damaging molecules, plants have evolved several enzymic mechanisms such like SOD, CAT, GPx, ascorbate peroxidase (APX), lipid peroxidase, GR, thioredoxin, and peroxiredoxin. They are considered the main natural antioxidant enzymes (ASADA, 1999). The balance between SOD and APX (and/or CAT) activity in cells is considered to be crucial for determining the – steady state level of O2 and H2O2. This balance, together with sequestering of metal ions such as Fe and Cu by ferritin and copper-binding proteins, is thought to be important to prevent the formation of the highly toxic HO• via the metal-dependent Haber–Weiss or the Fenton reactions (FRIDOVICH, 1997;

KARPINSKA et al., 2000). Peroxiredoxin catalyses the breakdown of alkyl hydroperoxides into water and their corresponding alcohols (ROUHIER, – JACQUOT, 2002). SOD, which catalyzes the disproportionation of O2 to O2 and

H2O2, has been called the cell’s first line of defence against ROS (HASSAN, – SCANDALIOS, 1990). This is because O2 is a precursor to several other highly – reactive species, so that control over the steady state O2 concentration by SOD constitutes an important protective mechanism (FRIDOVICH, 1997). In addition, over-expression of ROS-scavenging enzymes increases the tolerance of plants to abiotic stresses. ROS production can also be decreased in cells by the alternative channeling of electrons in the electron transport chains of the chloroplasts and mitochondria by a group of enzymes called alternative oxidases. Plants also have developed a wide range of non-enzymatic protective, mechanisms that serve to remove ROS before they can damage sensitive parts in cell. Clearly, it is of great importance to understand the quenching ability of biological compounds, such as carotenoids, in the context in which they are normally found to adequately evaluate the role they may play in protection against chlorophyl-induced photosensitization. Other low molecular weight compounds include a tocopherol (vitamin E), flavonoids of various types, ascorbate, and GSH. Flavonoids are widely distributed in higher plants and, by directly scavenging -OH, ONOOH and HOCl, are associated with strong inhibition of lipid peroxidation. The scavenging efficiency of flavonoids is 23 directly proportional to the number of hydroxyl groups bound to the phenols because this allows the flavonoids to bind metal ions. Ascorbate is of particular interest as an electron donor for hydroxyl radicals (for which there are no enzymatically catalyzed detoxification mechanisms) and as a substrate for APX. The reductant GSH, a tripeptide composed of glutamate, cysteine, and glycine, is nonspecific and is also both a general reductant and a substrate for enzymatically catalyzed reactions. It serves as a cofactor for some enzymatic reactions and as an aid in the rearrangement of protein disulfide bonds. The role of GSH as a reductant is extremely important, particularly in the highly oxidizing environment of photosynthetic cells. The sulfhydryl of GSH can be used to reduce peroxides formed during partial reduction of oxygen. The resulting oxidized form of GSH consists of two molecules disulfide-bonded together (abbreviated GSSG). The enzyme GSH reductase uses NADPH as a cofactor to reduce GSSG back to two molecules of GSH. With regard to the high molecular weight compounds, aerobic organisms express a battery of enzymes that contribute to the control of cellular ROS levels.

1.4.1. Glutathione S-transferase (GST: EC 2.5.1.18)

Plants developed mechanisms to defend themselves against insects, herbivores, pathogens and chemical compounds that are harmful to them. These enzymes carry out a wide range of functions in cells, such as the removal of reactive oxygen species and regeneration of S-thiolated proteins (both of which are consequences of oxidative stress), catalysis of conjugations with endogenous ligands, and catalysis of reactions in metabolic pathways not associated with detoxication. These strategies have been classified in two distinct groups: constitutive and induced defences. Induced mechanism is activation of specific enzymes that inactivate chemical compounds or detoxify heavy metals in plants. Glutathione (L-glutamyl-L-cisteinyl-glycine), a submajor constituent of all cells and almost always the major nono protein thiol compound present in cells, was first reported in 1888 by De Rey-Pailhade

24 as a philothion (RH2) and shown to be responsible for formation of hydrogen sulphide when yeast cells were crushed with elemental sulphur (BOYLAND,

CHASSEAUD, 1969). He also reported that philothion was present in beef muscle, beef liver, sheep brain, lamb small intestine, fish muscle, egg white, fresh sheep blood, and in freshly picked asparagus tips. Hopkins renamed philothion in 1921 as glutathione (MEISTER, 1989). Glutathione (GSH) shows two peptide bonds, two carboxylic acid groups, one amino group, and one thiol group. Naturally occurring toxic compounds include plant and fungal toxins and reactive oxygen species, such as the superoxide radical and H2O2. Glutathione that is distributed in the intracellular space of plants, animals, and microorganisms has two general functions: to remove toxic metabolites from the cell and to maintain cellular sulphhydril groups in their reduced form

(LIEBMAN, GREENBERG, 1988). Glutathione metabolism involves many reactions where glutathione is synthesized, degraded, conjugated or oxidized. In one of these pathways, the glutathione S-transferase (GST, E.C.2.5.1.18), a family of multifunctional isozymes in vertebrates, plants, insects and aerobic microorganisms (ARMSTRONG, 1993), catalyzes both GSH-dependent conjugation and reduction (KETTERER et al., 1993). Glutathione S-transferase detoxifies endobiotic and xenobiotic compounds by covalently linking glutathione to a hydrophobic substrate, forming less reactive and more polar glutathione S-conjugate (NEUEFEIND et al., 1997). Glutathione S-transferases are dimeric enzymes that show five independent classes, alpha, mu, pi, sigma

(BUETLER, EATON, 1992; DROGG et al., 1995; KETTERER et al. 1993), and theta

(BUETLER, EATON, 1992; DROGG et al., 1995). Classes alpha, mu and pi initially proposed by Mannervik in 1988 (BUETLER, EATON, 1992), are present only in animals and yeasts but absent in bacteria and plants (PEMBLE, TAYLOR, 1992).

The very heterogeneous theta class, reported by MEYER et al. 1991 (BUETLER,

EATON, 1992) are present in yeasts, plants, bacteria, rats, humans, chickens, salmon, and non vertebrates such as flies and apparently absent in lower animals such as molluscs, nematodes, and platyhelminthes (TAYLOR et al., 1993).

Detoxification of chemical compounds can be separated into two sequential processes: chemical (modification) transformation, and compartmentation. These two processes are divided into three phases: phase I (activation) reactions, phase II (conjugation), and phase III (internal compartmentation and storage processes) (MARRS, 1996). Phase I usually involves hydrolysis catalyzed by esterases and amidases or oxidation catalyzed by the cytochrome P-450 system. Phase II involves the deactivation synthesis of xenobiotic or a phase I-activated metabolite by covalent linkage to an endogenous hydrophilic molecule, such as glucose, malonate or glutathione resulting in a nontoxic or less toxic compound. Phase II is catalyzed by glucosyl-, malonyl, or glutathione transferases. In phase III, the inactive water-soluble conjugates formed in phase II, are exported from the cytosol by membrane-located proteins, which initiate the compartmentalization and storage in the vacuole (soluble conjugates) or in the cell wall (insoluble conjugates) (MARRS, 1996). Glutathione S- transferase in plants discovered in many plants such as maize (DIXON et al., 1997;

MARRS, WALBOT, 1997), wheat (RIECHERS et al., 1996; RIECHERS et al., 1997), dwarf pine (SCHRODER, RENNENBERG, 1992), soybean (ANDREWS et al., 1997), barley (ROMANO et al., 1993; WOLF et al., 1996).

1.4.2. Catalase (CAT: EC 1.11.1.6)

Catalase was first noticed as a substance in 1811 by Louis Jacques

Thenard who discovered H2O2 and suggested that its breakdown is caused by an unknown substance. In 1900, Oscar Loew was the first to give it the name catalase, and found its presence in many plants and animals. CAT is a heme containing redox enzyme. It is found in high concentrations in a compartment in peroxisomes cells. Catalase is a tetramer of four polypeptide chains, each over 500 amino acids long (BOON et al., 2007). It contains four porphyrin heme (iron) groups that allow the enzyme to react with the hydrogen peroxide to catalyze the formation of water and oxygen:

2 H2O2 → 2 H2O + O2

Catalase has one of the highest turnover numbers of all enzymes; one molecule of catalase can convert millions of molecules of hydrogen peroxide to water and oxygen per second. CAT can also oxidize different toxins, such as formaldehyde, formic acid, phenols and alcohols. It uses hydrogen peroxide according to the following reaction:

H2O2 + H2R → 2H2O + R The mechanism of this reaction is not known.

1.4.3. Glutathione reductase (GR: EC 1.8.1.7)

Glutathione reductase is an enzyme which reduces glutathione disulfide (GSSG) to the sulfhydryl form GSH, which is an important cellular antioxidant

(MEISTER, 1988; MANNERVIK, 1987). For every mole of oxidized glutathione (GSSG) one mole of NADPH is required to reduce GSSG to GSH. The enzyme forms a FAD bound homodimer. The glutathione reductase is conserved between all kingdoms. In plant genomes two GR genes are encoded.

1.4.4. Glutathione peroxidase (GPx: EC 1.11.1.9)

Glutathione peroxidase reduces hydrogen peroxide by transferring the energy of the reactive peroxides to a very small sulphur containing protein called glutathione. The selenium contained in these enzymes acts as the reactive center, carrying reactive electrons from the peroxide to the glutathione. Peroxiredoxins also degrade H2O2, within the mitochondria, cytosol and nucleus:

2GSH + H2O2 → GS-SG + 2H2O

1.4.5. Superoxide dismutase (SOD: EC 1.15.1.1)

Three major SOD isoforms have been described in eukaryotic photosynthetic organisms (ASADA, 1999): (1) CuZnSOD located in the thylakoid membranes and cytosol of higher plants, certain dinoflagellates, and charophycean green algae; (2) MnSOD isoform within mitochondria; and (3) FeSOD isoform in the chloroplast stroma. FeSOD is considered to be the 27

- major O2 scavenger in chloroplasts, whereas MnSOD is the most active scavenger in mitochondria (FRIDOVICH, 1997). Interestingly, SOD is induced by its substrate, and thus activation of specific SOD isoforms can serve as an - indicator of the cell compartment experiencing pollutant-induced O2 levels. Although SOD genes have been isolated from many different species, the FeSOD isoform has been reported from only a few. Recently, the isolation and molecular cloning of the FeSOD isoform from the marine dinoflagellate L. polyedrum, and the changes in expression of this enzyme during algal growth have been described (SIGAUD-KUTNER et al., 2002). The genes that control the formation of SOD are located on chromosomes 21, 6, and 4. When superoxide dismutase comes in contact with superoxide, it catalyzes the formation of hydrogen peroxide. Hydrogen peroxide is dangerous for the cell because it can easily be converted into hydroxyl radicals, one of the most destructive free radicals, by interacting with Fe2+ (this process is known as a Fenton reaction): - 2O2 (aq) → H2O2 (aq) 2+ 3+ • - Fe + H2O2 → Fe +OH + OH (Fenton reaction) The superoxide dismutase gene families appear to be specialized in function with respect to subcellular location and other as yet unknown factors

(ALSCHER et al., 2002). In the case of peroxisomes, the imposition of oxidative stress gives rise to organelle proliferation, thus adding another layer of complexity to stress responses (LOPEZ-HUERTAS et al., 2000). Defence mechanisms involving molecular chaperones and methionine sulfoxide reductase are becoming recognized as important players in resistance to oxidative stress throughout the cell.

1.5. Hydrogenperoxide, total antioxidant status and lipid peroxidation

H2O2 is a powerful oxidizing agent and is potentially damaging to cells.

By preventing excessive H2O2 build up сatalase allows important cellular processes, which produce H2O2 as a byproduct, to take place safely.

Peroxisomes partially oxidize fatty acids producing H2O2 as a byproduct. This peroxisomal oxidation shortens the fatty acids to length C8 or longer and facilitates an energy efficient degradation in the mitochondrion. In plants, H2O2 is generated in photorespiration – a process that apparently wastefully undoes some of the work of photosynthesis. Photorespiration occurs through the synthesis of glycolate from 3PG in the chloroplast and conversion of glycolate back to 3PG via the peroxisome. This uses energy in NADH and ATP hydrolyzing reactions, consuming O2 and releasing CO2 in the process. One hypothesis is that photorespiration protects plants from photooxidative damage when light levels are high and there is insufficient CO2. Antioxidant systems are capable of removing free radicals, thereby providing protection from free radical attack from such destructive molecules • • • as H2O2 and the RO , ROO and O2 radicals. Three main groups of antioxidants make up the antioxidant defence system. These include the primary, secondary, and tertiary defence. The total antioxidant status test measures the total antioxidant effect of these three defence systems in circulation. The three systems are as follows: primary antioxidants work by preventing the formation of new free radical species (these include superoxide dismutase, glutathione peroxidase and metal-binding proteins (e. g., ferritin or ceruloplasmin)); secondary antioxidants trap radicals thereby preventing chain reactions (include Vitamin E, vitamin C, beta-carotene, uric acid, bilirubin, and albumin); tertiary antioxidants repair biomolecules damaged by free radicals. These include DNA repair enzymes. One of the consequences of overproduction of reactive oxygen species beyond the plant’s defence capability is damage to cell membranes – described as lipid peroxidation. Polyunsaturated fatty acids of membrane phospholipids are highly susceptible to peroxidation by ROS. An autocatalytic chain of free radical reactions can produce various aldehydes, alkenals and hydroxyalkenals such as melondialdehyde (MDA) (SCHNEIDER et al., 2001). The resulting alkyl radicals may rearrange to a more stable conjugated diene, which enters the autocatalytic lipid peroxidation cascade (YANG et al., 2003). Membrane damage is mitigated 29 by terminating the reaction chain through the removal of lipid hydroperoxides. Glutathione S-transferase and glutathione peroxidase are the main enzymes involved in protecting the cells against ROS-induced lipid peroxidation.

1.6. Electrophysiological reaction of N. obtusa cells

N. obtusa cells have proved to be the most suitable material for analyzing the electrical characteristics of membranes of plant cells

(TAKABATAKE, SHIMMEN, 1997). Changes in the electrical characteristics of cell membrane (membrane potential/resting potential, membrane resistance and excitability) may show the degree and rate of toxic agent penetration into the cells. The electrical potential across a membrane is determined to a large extent by the difference in ion concentrations across the membrane. Therefore, it is important to know the concentration of the major ions on each side of a membrane. The concentrations (in mol/m3) of Cl-, Na+, K+, and Ca 2+ in the external medium and protoplasm of N. obtusa cells are following: external Cl- =0.4; Na+ =0.1; K+ =0.1; Ca2+ =0.1; protoplasm Cl- =22; Na+ =5; K+ =110; 2+ Ca <0,001 (LUNEVSKY et al., 1983). The difference in concentration of each ion across a differentially permeable membrane creates the driving force for diffusion across the membrane. As each permeate cation diffuses across the membrane, it leaves behind a negative charge (e. g., protein with ionized carboxyl groups) that will tend to attract the cation back to the original side. Thus, there is a chemical force that causes the cations to move from high concentrations to low concentrations and at the same time sets up an electrical force that tends to move the cations back. Because the membrane is permeable to some ions (e. g., K+) and impermeable to others (e.g., protons), at equilibrium there is no net movement of charge and the chemical force equals the electrical force. The unequal distribution of ions thus gives rise to a transmembrane potential difference that is electrically equivalent to a battery. The unequal distribution of charge is only possible because the nonconducting lipid bilayer that separates two conducting aqueous solutions

30 acts as a capacitor, that is a component that has the capacity to separate charges and resists changes in voltage when current flows through it. The specific membrane capacitance is a measure of how many charges (ions) must be transferred from one side of membrane to the other side either to set up or to dissipate a given membrane potential. The specific membrane capacitance of 2 all membranes is approximately 0.01 (F/m ) (FINDLAY, 1970). As a consequence of a membrane capacitance, and the diffusion of ions across a differentially permeable membrane, an equilibrium potential (or Nernst potential) is established. Changes in the membrane potential and resistance are related to the concentration of types (organic or inorganic) of toxic substances. Depending on the concentration, the change is reversible or irreversible; e. g., ammonium chloride changes the membrane potential of Charophyta cells already in a concentration of 0.27 mg. dm-3, and beginning from 53.5 mg dm-3 the change becomes irreversible (TYRAWSKA et al., 1994). The excitation of the Chara cell is due to a series of opening events of three different diffusion channels. 1) Opening of the Ca2+ channel which increases cytoplasmic Ca2+ concentration. 2) Opening of the Ca2+ -activated Cl- channel that depolarizes the membrane largely toward the equilibrium potential of Cl- ions (+60 to +90 mV). The patch clamp study on the Ca2+ - - activated Cl channel (OKIHARA et al., 1991) supports strongly that calmodulin should be involved in the opening of this channel. 3) Opening of the voltage- dependent K+ channel. The peak of action potential is usually determined by the ratio of the K+ conductance to the Cl- conductance. In addition to closing of the three diffusion channels, conductance of the pump contributes to the large repolarization beyond the diffusion potential (KISHIMOTO et al., 1985). If toxic substances (TS) block the diffusion channel alone in the Charophyta cells plasmalemma, they should hyperpolarize the plasmalemma decreasing membrane conductance (=conductance of the diffusion channels + conductance of the pump). If toxic substances block the H+ -pump alone, they should depolarize the pasmalemma decreasing the membrane conductance. If TS 31 inhibit the activity of calmodulin existing in the cells (TOMINAGA et al., 1985), they will not only depress the excitability by inhibiting the opening of the excitable Ca2+ -activated Cl- channel, but also depolarize the plasmalemma by + affecting the electrogenic H -pump (TSUTSUI, 1990). If algae cells were washed after chemical treatment, it should be possible to remove (completely or partially) effects of TS on the Charophyta action potential with washout. There are preliminary questions on the TS effect on the Charophyta plasmalemma. Electrophysiological investigation has proved useful to obtain new information on the mode of action of chemical compounds in plant cells. We make theories on plant physiology and their response to environment.

1.7. Lethality, as a measure of toxicity

A lethal concentration is an indication of the lethality of a given substance or type of chemical. Because resistance varies from one individual to another, the 'lethal concentration' represents a concentration at which a given percentage of subjects will die.

The most commonly-used lethality indicator is the LD50 (or LD50), a concentration at which 50% of subjects will die. LD figures depend not only on the species, but also on the mode of administration. For instance, a toxic substance inhaled or injected into the bloodstream may require a much smaller dosage than if the same substance is swallowed. The assessment of aquatic toxicity for the classification of chemicals and environmental risk assessments is typically based on toxicity test data for fish, crustacea (daphnids), and algae/aquatic plants. Toxicity data from freshwater and marine species are considered equivalent, although this is not true for all substances. The data showing the highest toxicity (the lowest acute toxicity values) is to be used in classifying a substance.

2. Methods

2.1. Plant material

The freshwater charophyte, N. obtusa was harvested in Lake Švenčius (Lithuania) in autumn 2004-2007 by long metal hook from a depth of about 5 m. The plants were transported to the laboratory in plastic bags filled with lake water. Prior to experiment, internodal cells (second or third internode below the tip of growing plants) were separated from neighboring cells. After separation from the bulk, single cells were kept at room temperature (20±2°C) in glass aquariums filled with equal parts of non chlorinated tap water, lake water and artificial pond water (APW) containing 0.1 mM KH2PO4, 1.0 mM

NaHCO3, 0.4 mM CaCl2, 0.1 mM, Mg(NO3)2, and 0.1 mM MgSO4 (pH 7-7.4) (VOROBIOV, MANUSADŽIANAS, 1983). Most of the cells had axial internodes about 1 mm in diameter and up to 15 cm long.

2.2. Preparation of Q. robur and F. sylvatica leaf extracts

Q. robur and F. sylvatica leaves from top litter layer were collected within the catchments area of Lake Müggelsee (2004–2007 years). The leaves were air-dried for two days in order to maintain an approximately uniform humidity level and ground using a homogeniser. 400 g of ground material was -1 soaked in 1.5 L of medium containing de-ionized water, CaCl2 (0.2 g L ), -1 -1 NaHCO3 (0.103 g L ) and sea-salt (0.1 g L ) in separate glass containers (Figure 2.1) and stirred for 24 h at room temperature. The resulting mixture was centrifuged at 20 000 × g for 10 minutes at 4°C to remove suspended materials. The supernatant was filtered using 0.8 µm cellulose-nitrate membrane filter and total dissolved organic carbon was determined according to DIN EN 1484, 1998.

Control Beech Oak

Figure 2.1 Control, F. sylvatica and Q. robur leaf extracts (photo by S. Pflugmacher).

2.3. Leaf litter degradation extracts (LLDEs)

Dry, fallen leaves belonging to the species F. sylvatica and Q. robur were collected during autumn season 2007 around Lake Mügelsee, Germany. The leaf litter was weighed to achieve a total mass of 40.0 g of leaves per treatment. For this experiment three treatments were chosen (F. sylvatica 100%, Q. robur 100% and control treatments without leaf litter). The leaf litter was placed into glass tanks with 10 litre capacity filled with artificial freshwater (AFW), where AFW was prepared according to NIMPTSCH and

PFLUGMACHER, 2005. Evaporated medium of each tank was restocked weekly with fresh AFW to reach the previously level and then homogenized mechanically with a glass stirrer. Exposures to LLDEs were set up in quintuplicate. The total period of the degradation experiment was 30 days. Total dissolved organic carbon from all treatments was determined according to DIN EN 1484, 1998.

2.4. Chemical characterization of leaf extracts and Lake water

Total dissolved organic carbon of both leaf extracts was determined by high temperature combustion (TOC-5000A, Shimadzu, Australia) after removal of inorganic carbonates by acidification (DIN EN 1484, 1998). From the total organic carbon concentrations, different dilutions were prepared from both leaf extracts and used for exposure experiments.

Dissolved organic carbon fractions were characterized by Liquid Chromatography and Organic Carbon Detection according to LST ISO 8245,

2003 and to HUBER and FRIMMEL (1996) as described in detail by SACHSE et al. (2001). Shortly, the chromatographable DOC portion of the filtered samples passes through a size-exclusion column packed with resin (Toyopearl HW 50S, volume of 250 x 20 mm). Phosphate buffer (0.029 mM, pH 6.5) was used as eluent at a flow rate of 1 mL min-1. Detectors were used for both DOC and absorbance (254 nm) measurements. DOC was detected by infrared absorbance of CO2 after UV oxidation of DOC at 185 nm in a cylindrical UV thin-film reactor (Graentzel-reactor).

2.5. Dose- and time-dependent exposure of N. obtusa to Q. robur and F. sylvatica leaf extracts

To investigate dose-effect relationship, the cells of N. obtusa were exposed to F. sylvatica and Q. robur leaf extracts DOC concentrations for 24 h. The treatments contained 0.11, 1.1, 5.5, 11, 55 and 0.245, 2.45, 12.25, 24.5, 122.5 mg L- 1 DOC, respectively. Alternatively, to investigate time-dependent activity changes, the cells were exposed to F. sylvatica and Q. robur leaf extract of 1.1 and 2.45 mg L-1 DOC, respectively, for 15 min, 0.5, 1, 1.5, 2, 4 and 24 h. In control experiments, only APW (no leaf extract) was added. All exposures were done in quintuplicate where each replicate contained 25 algae cells (Figure 2.2). The temperature was maintained at 20±1°C and the photoperiod at 14:10 (dark/light). After exposure, the N. obtusa cells were frozen in liquid nitrogen.

Figure 2.2 N. obtusa cells exposure (photo by L. Manusadžianas). 35

2.6. Exposure of charophyte cells to LLDEs

The charophyte cells were exposed to F. sylvatica and Q. robur LLDEs of various DOC concentrations (50 mL) for a period of 2 h. In control treatment, only APW (no extract) was added. All exposures were done in quintuplicate where each replicate contained 25 individual algae cells. The temperature was maintained at 20±1°C and the photoperiod at 14:10 (dark/light). After exposure, the N. obtusa cells were frozen in liquid nitrogen and stored at -80ºC until enzyme preparation.

2.7. Protein extraction

25 N. obtusa cells, homogenized in buffer

Centrifuge at 10 000 rpm Pellets, Supernatant discarded

Centrifuge at 40 000 rpm Supernatant, Pellets + 35% NH4SO4 Resuspend& Cent rifuge at homogenise in buffer Pellets, 20 000 rpm Supernatant, discarded + 80% NH4SO4 Microsomal fraction Cent rifuge at 30 000 rpm Supernatant Pellets discarded Resuspend in buffer & desalt

Cytosolic fraction Figure 2.3 Schema describing the procedure steps for enzyme preparations according to Pflugmacher and Steinberg (1997).

Protein extraction was done according to PFLUGMACHER and

STEINBERG (1997) with minor modification (Figure 2.3). For each treatment the algae (25 cells) were homogenized in a glass potter by adding 10 mL ice- cold 0.1 M sodium phosphate buffer pH 6.5 containing 1.4 mM dithioerythritol (DTE) and 1 mM ethylenediaminetetracetic acid (EDTA). After removing cell debris by centrifuging at 10 000 x g for 10 min, the supernatant was centrifuged again at 40 000 x g for 60 min to collect the microsomal fraction. The microsoms were resuspended in sodium-phosphate buffer (20 mM, 36 pH 7.0) containing 20% glycerol, 1.4 mM of DTE, and homogenized in a glass potter. The first protein precipitation was achieved by a 35% ammonium sulphate saturation followed by a centrifugation at 20 000 x g for 20 min where the pellet was discarded (Figure 2.3). The second protein precipitation was achieved by an 80% ammonium sulphate saturation followed by a centrifugation at 30 000 x g for 30 min. The pellet from the last precipitation step was suspended in 20 mM sodium phosphate buffer pH 7.0 and the resulting extract was desalted using NAP-5 columns (Amersham Pharmacia, Uppsala, Sweden) (Figure 2.3). Immediately after extraction the protein preparations extracts were shock frozen using liquid nitrogen, and then stored at -80°C until enzyme activity assays.

2.8. Measurement of enzymatic activities

2.8.1. Catalase

Catalase (EC 1.11.1.6) was measured photometrically according to

BAUDHUIN, 1964, where a decrease of H2O2 is measured obtaining a decrease of absorption at 240 nm (extinction coefficient ε = 0.0361 L*mmol-1*cm-1). The assay consisted of: 1250 μL NaP-buffer (50 mM; pH=7.0),

100 μL H2O2 (150 mM),

25 μL H2O / sample. Reaction:

CAT 2H O 2H O+ O 2 2 2 2

2.8.2. Glutathione S-transferase

GST (EC 2.5.1.18) activities were measured in the soluble (cytosolic and microsomic) fraction applying described methods using 1-chloro-2,4- dinitrobenzene (CDNB) (HABIG et al., 1974) as substrates. Co-substrate is

GSH. Substrates were diluted in ethanol. Wave length 340 nm, ε = 9.6 (L*mmol-1*cm-1). The assay consisted of: 1100 μL NaP-buffer (0.1 M; pH=6.5), 40 μL GSH, 40 μL CDNB,

40 μL H2O / sample. Reaction:

NH2 O Cl HO NH O NH NO2 OOHH OH GST S S O + HCl + GSH + pH 6.5 N - O

NO2

+ - N O O 1-chloro-2,4-dinitrobenzene (CDNB) GS-DNB conjugate 2.8.3. Glutathione reductase

Glutathione reductase (EC 1.8.1.7) activity was assayed photometrically by measuring NADPH (β-nicotinamide adenine dinucleotide phosphate) consumption during the enzymatic reaction. In the presence of GSSG (glutathione disulfide) and NADPH, GR reduces GSSG and oxidizes NADPH to yield NADP+ resulting in a decrease of absorbance at 340 nm (extinction -1 coefficient ε = 6.4 1*mM*cm ) (CARLBERG, MANNERVIK, 1985). The assay consisted of: 850 μL NaP-buffer (0.1 M; pH=7.5), 50 μL GSSG (20 mM), 50 μL NADPH (2 mM),

50 μL H2O / sample. Reaction:

NH2 O HO NH O NH NH2 O OOHH OH GR HO NH O S + S ++NADPH + H 2 NH NADP+ OOHH OH pH 7.5 OOHH OH NH HO NH O SH

NH2 O GSSG GSH

2.8.4. Glutathione peroxidase

The glutathione peroxidase (EC 1.11.1.9) activity was assayed photometrically as described by DROTHAR et al. (1985), through the reduction of H2O2 by GPx in expend of reduced glutathione (GSH) which results in the formation reaction of the glutathione reductase, reducing GSSG to GSH and oxidizing NADPH to yield NADP+ resulting in a decrease of absorbance at 340 nm. One unit of GPx is defined as the activity that converts 2 μmol of reduced glutathione (corresponding to 1 μmol NADPH) per minute at 25°C (extinction coefficient ε = 6.4 1*mmol-1*cm-1). The assay consisted of: 800 μL NaP-buffer (0.1 M; pH=7.5), 10 μL Glutathione reductase (2 U), 40 μL GSH (50 mM), 40 μL NADPH (2 mM),

10 μL H202 (9 mM),

100 μL H2O / sample. Reaction:

NH2 O HO NH O NH2 O NH HO NH O GPx OOHH OH 2 NH + 2 NADPH + 2 H2O2 S pH 7.5 S + 2 NADP + 4 H2O OOHH OH OOHH OH SH NH HO NH O

NH2 O GSH GSSG 2.8.5. Calculation of specific enzymatic activity by photometric measurement

The measurement of the catalytic activity of an enzyme (Z) occurs through the assessment of the velocity of the catalyzed reaction (i.e. substrate turnover per time unit) expressed in katal (μmol s-1) and calculated using the following equation.

(∆E * V) z = (ε*d*∆t*v)

Where, ∆E – extinction change per minute, V – test volume (μL), ε – molar extinction coefficient (L*mM*cm-1), d – cuvette width (cm), v – volume of enzyme extract in the test (μL), ∆t – measuring interval (s). In order to facilitate the comparison between different samples the specific enzyme activity was divided by the total protein content of the enzyme extract resulting the definitive equation used in this research for activity assessment.

(∆E * V)___ z = (ε*d*∆t*v*c)

Where, c – protein content of enzyme extract (μg*μL-1). Therefore all specific enzymatic activities were expressed in nkat mg-1protein.

2.8.6. Superoxiddismutase

SOD activity was determined by photometrical method utilizing the SOD assay kit from Calbiochem®. The assay is based on the SOD-mediated increase in the rate of autoxidation of 5,6,6a,11b-tetra hydro-3,9,10- trihydroxybenzo[c]fluorine (BXT-01050) (0.66 mM) in aqueous alkaline solution (32 mM HCl; 0.5 DTPA; 2.5% ethanol) (R1). This autoxidation yields a chromophore with a maximal absorbance wavelength of 525 nm. The optimized assay of SOD activity is performed at pH 8.8, 37°C, in 50 mM air- saturated 2-amino-2-methyl-1,3-propanediol buffer (SOD-buffer) containing 3 mM boric acid and 0.11 mM diethylenetriaminepentaacetic acid (DTPA) which is mixed with the mercaptan scavenger 1,4,6-trimethyl-2- vynilpyridinium trifuoromethanesulfonate (R2) (33.3 mM, in 1 mM HCl), according to NEBOT et al., 1993. The assay consisted of: 900 μL SOD-buffer,

40 μL H2O / sample, 30 μL R2 solution, 30 μL R1 solution.

The specific activity of SOD was calculated using the following equation:

0.93 (Vs/Vc-1)__ SOD = 1.073-0.073(Vs/Vc) Where, Vs – reaction rate of sample containing SOD, Vc – reaction rate of controls (without SOD). Reaction:

SOD .- + 2 O2 + 2 H 2 H O + O 2 2 2

2.8.7. Measurement of cell internal hydrogen peroxide

The levels of cell internal H2O2 were measured by spectrophotometric method according to JANA and CHOUDHURI (1981), where 5 cells of the algae were homogenized in 3 ml 50 mM sodium phosphate buffer (pH 7.0) and centrifuged at 20 000 g (4°C) for 5 min. Supernatant was mixed with 0.1% titanium sulphate in 20% (v/v) H2SO4 (250 μL) and the absorbance was measured at 410 nm (extinction coefficient ε = 0.281*mmol-1*cm-1). The assay consisted of: 0.1% Titanchloride in 20%,

750 μL H2SO4,

250 μL H20 / sample. -1 The content of H2O2 was expressed in μmol*g FW, where FW corresponds to the cells of plant material used for extraction, was calculated using the following equation.

(∆E*V)____ H O [μmol * gFW] = 2 2 (ε*d*∆t*v*gFW)

Where, ∆E – extinction change per minute, V – test volume (μL), ε – molar extinction coefficient (1*mmol-1*cm-1), d – cuvette width, ∆t – measuring interval (time unit), v – volume of sample (μL).

2.8.8. Measurement of the total antioxidant status

Total antioxidant status (TOSC) was determined by a photometrical method utilizing the TOSC assay kit from Calbiochem®. This method is based on the formation of blue-green cation radical of 2,2’-azino-di-[3- ethylbenzthiazoline sulphonate] cation (ABTS+) in the presence of metmyoglobine and H2O2. The low molecular weight antioxidants inhibit colour production proportionally to their concentration (MILLER et al., 1993). The absorbance was measured at 600 nm lengths. The assay consisted of: 1000 μL Chromogen,

20 μL H2O / standard / sample. The total antioxidant status was calculated using the following equation:

Antioxidant Concentration (mM) = [Standard](∆A Blank-∆A Sample) (∆A Blank-∆A Sample)

2.8.9. Measurement of lipid peroxidation

Lipid peroxidation (LPO) products malondialdehyde (MDA) and 4- hydroxy-2-nanenal (4-HNE) were determined by a photometrical method utilizing the lipid peroxidation assay kit from Calbiochem®. This method is based on the method of Esterbauer and Cheeseman (ESTERBAUER,

CHEESEMAN, 1990). The colorimetric reaction is a condensation of the respective aldehyde with 1-methyl-2-phenylindole yielding chromophores with absorption maxima at 586 nm. Both aldehydes react with 1-methyl-2- phenylindole in the presence of methanesulfonic acid and ferric iron, while subsituation of methanesulfonic acid by hydrochloric acid allows measurement of MDA alone due to an irreversible cyclization reaction of hydroxyalkenals

(GERARD-MONNIER et al., 1998). To calculate the amount of MDA + 4-HNE a standard curve of MDA was established. The amount of total lipid peroxidation was calculated using the following equation:

[MDA+HAE] = (A-Ao)D/ε, where, A – the absorbance at 586 nm of the sample, Ao – the absorbance of the blank, D – dilution factor, ε – molar extinction coefficient -1 -1 (1*mmol *cm ) obtained using MDA standard. Reaction mechanism:

H O + 2 Ph + OH N N N Ph Ph Ma londia ldhehyde N-methyl-2-phenylindole Chromophore

2.9. Electrophysiological experiments

The changing of resting potential of the cells (n≈120) was measured during a 24-h and 3-h rewash in control solution. The treatments contained 7.5–1838 mg L-1 and 33–1100 mg L-1 DOC concentrations from Q. robur and F. sylvatica leaf extracts respectively. In control experiments, APW (i.e., no leaf extract) was used. Bioelectrical activity of cells was measured simultaneously according to + K -anaesthesia method (SHIMMEN et al., 1976), modified for multichannel recording with extra cellular chlorinated silver wire electrodes. The discrete RP values from distinct cells were taken every second. After amplification, the output signals were channelled into a PC by means of interface system. The data on kinetics of RP of all cells were potted on a graphic display for visual control and stored for further analysis. The cells were placed in a plexiglas chamber having distinct yet identical sections, as seen in Figure 2.4. One end of the cell was paced in the larger pool. The node and small segment (about 1 mm) of each cell was positioned in the common central pool, filled with 100 mM KCl in control solution. Electrical insulation between the pools was achieved with 5% boric vaseline junctions, as well as by an air gap between them (Figure 2.4). Throughout the study, electrodes made of chlorinated silver wire with both

43 high stability and low electrode potential were used. The electrical contact between the pools was enabled through the Π-shaped glass tube bridge plugged with 3% (w/v) agar solidified in a 3 M KCl solution. The placing of reference electrode in the pool reference instead of in the common central pool was necessary to avoid the appearance of chlorine ions as a potential component that may interfere in later measuring of RP value. Such interference could occur due to alteration of the Cl- concentration difference between the central pool and changing test solutions. Electrical signals from electrodes were led through the preamplifier module and multiplexer to the input of the differential amplifier (Figure 2.4). The discrete RP meanings were taken every second. The output signal was channeled into an IBM PC computer by the means (Figure 2.4 and Figure 2.5).

Figure 2.4 The set-up of electrophysiological measurements.

Figure 2.5 Electrophysilogical testing system.

2.10. Cell Lethality testing

Lethality response of algal cells was investigated during 42-days. The treatments contained 5.5–1100 mg L-1 and 24.5–1225 mg L-1 DOC concentrations from F. sylvatica and Q. robur leaf extracts respectively. In control experiments, only APW was used. Single internodal cells (each 4– 15 cm in length) were placed in Petri dishes (10 cells per dish, 4 replicates), preadapted for 1–2 days in artificial pond water containing 0.1 mM KH2PO4,

1.0 mM NaHCO3, 0.4 mM CaCl2, 0.1 mM Mg(NO3)2 and 0.1 mM MgSO4 (pH 7.0–7.4, unbuffered), and then were kept at room temperature (18–24oC) in the dark. The preadaptation in APW before the test allowed discarding occasionally dead cells, which had been injured during the transfer to the Petri dishes. Survival of the cells was checked daily by gently picking up each cell with a spatula. A cell was judged to be dead when picked up if there was disappearance of turgor pressure, a state in which a cell bends on the spatula and loses its cylindrical shape (Figure 2.6a and b). a) b)

Figure 2.6 N. obtusa cells checked with the spatula, a- with turgor pressure, b- without turgor pressure.

2.11. Statistical analysis

All values are expressed as mean ± S.D. except DOC values. Significant differences between the control and respective treatments were analyzed by one-way ANOVA followed by Tukey’s post-hoc test, at p < 0.05 using Statistica v.5.5 (StatSoft, Inc. 2000).

3. Results

3.1. Enzymatic activity, H2O2 content, total antioxidant content and lipid peroxidation in N. obtusa

3.1.1. Dose – response effects induced by F. sylvatica and Q. robur leaf extracts

3.1.1.1. Microsomal glutathione S-transferase assayed with the conjugating substrate 1-chloro-2,4-dintrobenzene (CDNB)

Microsomal glutathione S-transferase activity, measured with CDNB as a substrate, was elevated in N. obtusa cells exposed for 24 h compared to F. sylvatica and Q. robur with control (Figure 3.1 and Figure 3.2). In F. sylvatica extract (Figure 3.1), while the increase in activity was significant at only 5.5 mg L-1 DOC concentration, in Q. robur extract, the increase in enzymatic activity was significant at all tested DOC concentrations (0.245 – 122.5 mg L-1) (Figure 3.2).

8.00 mGST- Beech 7.00 *

6.00 protein] -1 5.00

4.00

3.00

2.00

Enzyme activity [nkat mg activity Enzyme 1.00

0.00 Control 0.11 1.1 5.5 11 55 Dissolved organic carbon [mg L-1] Figure 3.1 The activity of microsomal glutathione S-transferase, measured with CDNB, in N. obtusa after 24 h exposure to different DOC concentrations from F. sylvatica leaf extract. * Significantly different from the control, p < 0.05.

8.00 mGST- Oak 7.00 *** * * protein] 6.00 -1

5.00

4.00

3.00

2.00 Enzyme activity [nkat mg activity Enzyme 1.00

0.00 Control 0.245 2.45 12.25 24.5 122.5 -1 Dissolved organic carbon [mg L] Figure 3.2 The activity of microsomal glutathione S-transferase, measured with CDNB, in N. obtusa after 24 h exposure to different DOC concentrations from Q. robur leaf extract. * Significantly different from the control, p < 0.05.

3.1.1.2. Cytosolic glutathione S-transferase assayed with the conjugating substrate CDNB

In comparison to mGST, the elevation of cytosolic GST activity induced by F. sylvatica extracts was evident in broader range of DOC concentrations from 1.1 up to 5.5 mg L-1 (Figure 3.3). At highest tested DOC concentrations (55 mg L-1), the activity of cGST decreased to control level.

8.00 cGST- Beech 7.00 * ** 6.00 protein]

-1 5.00

4.00

3.00

2.00

1.00 Enzyme activity [nkat mg [nkat activity Enzyme

0.00 Control 0.11 1.1 5.5 11 55 Dissolved organic carbon [mg L-1] Figure 3.3 The activity of cytosolic glutathione S-transferase, measured with CDNB, in N. obtusa after 24 h exposure to different DOC concentrations from F. sylvatica leaf extract. * Significantly different from the control, p < 0.05. 47

As in the case of mGST, the significant increase in activity of sGST in N. obtusa cells exposed to Q. robur extract was detected in all tested DOC concentrations (0.245 – 122.5 mg L-1) (Figure 3.4).

8.00 cGST- Oak * * 7.00 * * *

protein] 6.00 -1 -1

5.00

4.00

3.00

2.00

Enzyme activity [nkat mg [nkat activity Enzyme 1.00

0.00

Control 0.245 2.45 12.25 24.5 122.5 Dissolved organic carbon [mg L-1] Figure 3.4 The activity of cytosolic glutathione S-transferase, measured with CDNB, in N. obtusa after 24 h exposure to different DOC concentrations from Q. robur leaf extract. * Significantly different from the control, p < 0.05.

3.1.1.3. Catalase

The catalase activity within F. sylvatica DOC dose dependent charophyte algae exposures increased significantly in the treatment of 1.1 mg L-1 DOC, statistically significantly decreased CAT activities were observed as the DOC concentrations increased from 5.5 to 55 mg L-1 DOC (Figure 3.5).

350.0 CAT- Beech * 300.0

protein] 250.0

-1 **

200.0

* 150.0

100.0

50.0 nyeatvt na mg [nkat activity Enzyme

0.0 Control 0.11 1.1 5.5 11 55 Dissolved organic carbon [mg L-1] Figure 3.5 The activity of catalase, in N. obtusa after 24 h exposure to different DOC concentrations from F. sylvatica leaf extract. * Significantly different from the control, p < 0.05. 48

The activity of catalase in the algae preparations increased significantly at the lowest DOC concentrations of Q. robur extract, 0.245 and 2.45 mg L-1 (Figure 3.6). No significant differences were observed as the concentration increased from 12.25 to 122.5 mg L-1 DOC.

600.00 CAT- Oak

500.00 * * protein]

-1 400.00

300.00

200.00

100.00 Enzyme activity [nkat mg [nkat activity Enzyme

0.00 Control 0.245 2.45 12.25 24.5 122.5 Dissolved organic carbon [mg L-1] Figure 3.6 The activity of catalase, in N. obtusa after 24 h exposure to different DOC concentrations from Q. robur leaf extract. * Significantly different from the control, p < 0.05.

3.1.1.4. Glutathione reductase

The glutathione reductase activity in algae cells increased significantly by factors 1.4 to 2.6 at all DOC concentrations exposed to F. sylvatica leaf extract (Figure 3.7). The highest elevation of GR by a factor 2.6 was observed at a DOC concentration of 5.5 mg L-1 DOC and the lowest one by a factor 1.4 at 0.11 mg L-1 DOC.

3.00 GR- Beech *

2.50

protein] 2.00 * -1 * * * 1.50

1.00

0.05 Enzyme activity [nkat mg [nkat activity Enzyme

0.00 Control 0.11 1.1 5.5 11 55 Dissolved organic carbon [mg L-1] Figure 3.7 The activity of glutathione reductase, in N. obtusa after 24 h exposure to different DOC concentrations from F. sylvatica leaf extract. * Significantly different from the control, p < 0.05. 49

The activity of GR in the algae exposed to Q. robur leaf extract increased significantly at all DOC concentrations (0.245 and 122.5 mg L-1) (Figure 3.8).

3.00 GR- Oak *

2.50 * * * protein] * -1 2.00

1.50

1.00

0.50 Enzyme activity [nkat mg [nkat activity Enzyme

0.00 Control 0.245 2.45 12.25 24.5 122.5 Dissolved organic carbon [mg L-1] Figure 3.8 The activity of glutathione reductase, in N. obtusa after 24 h exposure to different DOC concentrations from Q. robur leaf extract. * Significantly different from the control, p < 0.05.

3.1.1.5. Glutathione peroxidase

Activity of glutathione peroxidase in N. obtusa exposed to F. sylvatica leaf extract decreased significantly, compared to untreated cells in control, from 5.5 up to 55 mg L-1 (Figure 3.9).

GPx- Beech 2.00

1.50 protein] -1

1.00 * *

0.50 * nyeatvt na mg [nkat activity Enzyme

0.00 Control 0.11 1.1 5.5 11 55 Dissolved organic carbon [mg L-1] Figure 3.9 The activity of glutathione peroxidase, in N. obtusa after 24 h exposure to different DOC concentrations from F. sylvatica leaf extract. * Significantly different from the control, p < 0.05.

In N. obtusa exposed to Q. robur extract, the increase in GPx activity was significant at all DOC concentrations tested but not in a concentration- dependent manner (Figure 3.10). At higher DOC concentrations the activity of GPx decreased slightly.

GPx- Oak 2.00 * * *

* *

protein] 1.50 -1

1.00

0.50 Enzyme activity [nkat mg activity Enzyme 0.00 Control 0.245 2.45 12.25 24.5 122.5 Dissolved organic carbon [mg L-1] Figure 3.10 The activity of glutathione peroxidase, in N. obtusa after 24 h exposure to different dissolved organic carbon concentrations from Q. robur leaf extract. * Significantly different from the control, p < 0.05.

3.1.1.6. Superoxide dismutase

Significant decrease (p < 0.05) in SOD activity was detected between control and N. obtusa exposed for 24 h to F. sylvatica at all DOC concentrations tested, at higher DOC concentrations the activity of SOD increased (Figure 3.11).

21.00 SOD- Beech 18.00

15.00 protein]

-1 *

12.00 * 9.00 * ** 6.00

3.00 Enzyme activity [nkat mg activity Enzyme 0.00

Control 0.1 1.1 5.5 11 55 Dissolved organic carbon [mg L-1] Figure 3.11 The activity of superoxide dismutase, in N. obtusa after 24 h exposure to different DOC concentrations from F. sylvatica leaf extract. * Significantly different from the control, p < 0.05. 51

The activity of superoxide dismutase in N. obtusa cells when exposed to Q. robur leaves extract for 24 h remained unchanged in all treatments with the DOC concentrations from 0.245 to 24.5 mg L-1 (Figure 3.12). Statistically significant inhibition of SOD compared to control by a factor 1.4 was observed only at the highest, 122.5 mg L-1 DOC concentration (Figure 3.12).

21.00 SOD- Oak 18.00

15.00 protein] -1 12.00 *

9.00

6.00

3.00 Enzyme activity [nkat mg [nkat activity Enzyme 0.00 Control 0.245 2.45 12.25 24.5 122.5 Dissolved organic carbon [mg L-1] Figure 3.12 The activity of superoxide dismutase, in N. obtusa after 24 h exposure to different DOC concentrations from Q. robur leaf extract. * Significantly different from the control, p < 0.05.

33 3.1.1.7. Hydrogen peroxide content

N. obtusa cells exhibited decrease in H2O2 content when exposed to F. sylvatica and Q. robur leaf extracts for 24 h (Figure 3.13). In both leaf extracts, H2O2 levels tend to decrease at tested DOC concentrations.

1.00 Control H2O2 Beech -3 Oak

0.50 )/cm* cell] 10 cell] )/cm* 2 O 2 [µmol (H [µmol

0.00 Control 1.1 5.5 2.45 12.25 Dissolved organic carbon [mg L-1] Figure 3.13 Intracellular hydrogen peroxide content in N. obtusa cells after 24 h exposure to different DOC concentrations from F. sylvatica and Q. robur leaf extracts. 52

3.1.1.8. Total antioxidant content

The total antioxidant concentration in algae exposed to the different DOC concentrations showed substantially lower values in treatments with leaf extracts as than those detected in control (Figure 3.14). Decrease in TOSC, when exposed to F. sylvatica and Q robur leaf extracts for 24 h, was observed at all tested DOC concentrations.

0.60 TOSC Control Beech 0.50 Oak

0.40

0.30

0.20

0.10 Antioxidant concentration (mM)

0.00 Control 1.1 5.5 2.45 12.25 Dissolved organic carbon [mg L-1] Figure 3.14 Total antioxidant content in N. obtusa algae after 24 h exposure to different DOC concentrations from F. sylvatica and Q. robur leaf extracts.

3.1.1.9. Lipid peroxidation

The lipid peroxidation in charophyte algae exposed to F. sylvatica leaf extract increased (not significantly). The LPO increase, exposed to F. sylvatica and Q. robur leaf extracts for 24 h, was observed at both respective DOC concentrations (Figure 3.15).

14.00 LPO Control 12.00 Beech Oak 10.00

8.00

6.00 MDA [ µM ] µM [ MDA 4.00

2.00

0.00 Control 1.1 5.5 2.45 12.25

Dissolved organic carbon [mg L-1]

Figure 3.15 Lipid peroxidation measured as the concentration of MDA in N. obtusa algae after 24 h exposure to different DOC concentrations from F. sylvatica and Q. robur leaf extracts.

3.1.2. Time-dependent effects induced by F. sylvatica and Q. robur leaf extracts

3.1.2.1. Microsomal glutathione S-transferase, measured with CDNB

Time-dependent exposure (from 0 – 24 h) of N. obtusa cells to F. sylvatica leaf extracts did not result in significantly different from control mGST activity compared to the control in the first exposures at 15 and 30 min, whereas a statistically significant decrease of mGST activity by a factor from 1.4 to 1.7, below control level was observed in the period of 1 to 4 h (Figure 3.16). After 24 h - exposure the activity of mGST in the algae preparations increased to the control level again (Figure 3.16).

7.00 mGST- Beech 6.00

protein] 5.00 -1 * * 4.00 * * 3.00

2.00

1.00 Enzyme activity [nkat mg activity Enzyme

0.00 Control 15 min 30 min 1 h 1.5 h 2 h 4 h 24 h Exposure time Figure 3.16 Kinetics of microsomal glutathione S-transferase (mGST) activity, measured with CDNB, in N. obtusa after exposure to 1,1 mg L-1 DOC from F. sylvatica. * Significantly different from the control, p < 0.05 (n=5).

Q. robur leaves extract induced statistically significant decrease of microsomal GST in the algae at all time exposure (Figure 3.17).

8.00 mGST- Oak 7.00

protein] 6.00 -1 -1

5.00

4.00 *

* * 3.00 * * * * 2.00

Enzyme activity [nkat mg activity Enzyme 1.00

0.00 Control 15 min 30 min 1 h 1.5 h 2 h 4 h 24 h Exposure time Figure 3.17 Kinetics of microsomal glutathione S-transferase (mGST) activity, measured with CDNB, in N. obtusa after exposure to 2.45 mg L-1 DOC from Q. robur. * Significantly different from the control, p < 0.05 (n=5).

3.1.2.2. Cytosolic glutathione S-transferase, measured with CDNB

Cytosolic glutathione S-transferase (cGST) activity in N. obtusa exposed to F. sylvatica leaf extracts with 1.1 mg L-1 DOC concentration, 55 presented a significant increase by a factor approximately 2.5 in exposures between 15 min and 4 h. Even after 24 h of exposure to F. sylvatica leaf extracts, cGST activity remained increased by a factor 2 when compared to control (Figure 3.18).

10.00 cGST- Beech * 9.00 * * * 8.00 * *

protein] 7.00

-1 * 6.00

5.00

4.00

3.00

2.00

Enzyme activity [nkat mg activity Enzyme 1.00

0.00 Control 15 min 30 min 1 h 1.5 h 2 h 4 h 24 h Exposure time Figure 3.18 Kinetics of cytosolic glutathione S-transferase (cGST) activity, measured with CDNB, in N. obtusa after exposure to 1.1 mg L-1 DOC from F. sylvatica. * Significantly different from the control, p < 0.05 (n=5).

The response of cytosolic glutathione S-transferase showed an increase with variability exposed to 2.45 mg L-1 DOC in all time-range investigated (Figure 3.19).

10.00 cGST- Oak 9.00

8.00

protein] 7.00 -1 * * 6.00 * * 5.00 * * * 4.00

3.00

2.00

Enzyme activity [nkat mg activity Enzyme 1.00

0.00 Control 15 min 30 min 1 h 1.5 h 2 h 4 h 24 h Exposure time Figure 3.19 Kinetics of cytosolic glutathione S-transferase (cGST) activity, measured with CDNB, in N. obtusa after exposure to 2.45 mg L-1 DOC from Q. robur. * Significantly different from the control, p < 0.05 (n=5). 56

3.1.2.3. Catalase

F. sylvatica leaf extracts induced statistically significant elevation of CAT activity in N. obtusa during the whole exposure period (Figure 3.20). Overall, the CAT activity increased by a factor from 1.4 to 2.0 compared to control, where the highest elevation by a factor 2.0 was observed after 4 h of exposure to F. sylvatica leaf extract. After 24 h of exposure to the extract the CAT activity tended to decrease (Figure 3.20).

600.0 CAT- Beech *

500.0 * * protein] ***

-1 400.0 *

300.0

200.0

100.0 Enzyme activity [nkat mg [nkat activity Enzyme

0.0 Control 15 min 30 min 1 h 1.5 h 2 h 4 h 24 h Exposure time Figure 3.20 Kinetics of catalase (CAT) activity, in N. obtusa after exposure to 1.1 mg L-1 DOC from F. sylvatica. * Significantly different from the control, p < 0.05 (n=5).

A time response an experiment was conducted using 2.45 mg L-1DOC from Q. robur leaf extract and results for CAT activity are shown in Figure 3.21. Thought the CAT activity detected during the experiment was not significant compared to the control.

600.00 CAT- Oak 500.00 protein]

-1 400.00

300.00

200.00

100.00 Enzyme activity [nkat mg activity Enzyme

0.00 Control 15 min 30 min 1 h 1.5 h 2 h 4 h 24 h Exposure time Figure 3.21 Kinetics of catalase (CAT) activity, in N. obtusa after exposure to 2.45 mg L-1 DOC from Q. robur. * Significantly different from the control, p < 0.05 (n=5).

3.1.2.4. Glutathione reductase

F. sylvatica leaf extract (1.1 mg L-1 DOC) induced significant increase in GR activity after 1 h of exposure and this tendency was evident until the end of 24 h-period (Figure 3.22). The highest increase of GR by a factor 2, was achieved after 2 h of exposure. After 4 h and 24 h of exposure to F. sylvatica leaf extracts, GR activity tends to decrease, but still remained significantly different from control (Figure 3.22).

2.50 GR- Beech

2.00 * * protein] * -1 * 1.50 *

1.00

0.50 Enzyme activity [nkat mg activity Enzyme

0.00 Control 15 min 30 min 1 h 1.5 h 2 h 4 h 24 h Exposure time Figure 3.22 Kinetics of glutathione reductase (GR) activity, in N. obtusa after exposure to 1.1 mg L-1 DOC from F. sylvatica. * Significantly different from the control, p < 0.05 (n=5). 58

Time-dependent exposure of N. obtusa to Q. robur leaf extract at 2.45 mg L-1 DOC concentration did not reveal a relationship between glutathione reductase activity and exposure duration. Significant increase in GR activity by a factor 2 was detected after 4 h, while the largest difference from control was achieved after 24 h exposure to Q. robur extract (Figure 3.23).

3.00 GR- Oak

2.50 *

protein] 2.00 -1 * 1.50

1.00

0.50 Enzyme activity [nkat mg [nkat activity Enzyme 0.00 Control 15 min 30 min 1 h 1.5 h 2 h 4 h 24 h Exposure time Figure 3.23 Kinetics of glutathione reductase (GR) activity, in N. obtusa after exposure to 2.45 mg L-1 DOC from Q. robur. * Significantly different from the control, p < 0.05 (n=5).

3.1.3. Effects induced by 30-day-decomposing of leaf litter extracts from F. sylvatica and Q. robur

3.1.3.1. Dissolved organic carbon content

Prior to exposure experiments, both leaf litter decomposing extracts (LLDE) as well as control were characterized by the amounts of dissolved organic carbon (Table 1; Figure 3.24).

Control Beech Oak Day DOC (mg L-1) 00.5- - 1 0.7 94.8 86.0 2 0.4 128.7 137.3 7 1.0 166.5 177.7 14 3.5 151.2 156.1 21 0.7 136.3 156.2 30 0.9 127.2 144.1 Table 1 Dissolved organic carbon content in control, F. sylvatica and Q. robur leaf extracts 59

10 180 Control 160 Beech DOCL (mg 8 Oak 140

120 (grey stripes) & 6

100 -1 ) from oak & beech & oak from ) ) from control ) from -1

(white) 80 4 60

DOC (mg L 40 2

0 0 0 1 2 7 14 21 30 Time (days) Figure 3.24 Dissolved organic carbon content in control ( -artificial freshwater), F. sylvatica and Q. robur leaf degradation extracts.

3.1.3.2. Cytosolic glutathione S-transferase, assayed with the conjugating substrate CDNB

The activity of cGST measured in N. obtusa cells exposed to F. sylvatica and Q. robur leaf litter degradation extracts (LLDEs) for 2 h, using CDNB as substrate, showed a significant elevation in activity compared to control (Figure 3.25). Significant activation of cGST by a factor from 1.4 to 2.7 compared to control was observed in cells exposed to F. sylvatica and by a factor from 1.2 to 2.7 exposed to Q. robur LLDEs. One-day-old F. sylvatica and Q. robur LLDEs already induced the highest significant elevation of cGST activity by a factor 2.7 (at DOC concentration 94.81 and 86.01 mg L-1respectively (Table 1; Figure 3.24, Figure 3.25).

1.00 cGST *** 0.90 * * * Control 0.80 Beech * * *

protein] 0.70 Oak -1 0.60 * 0.50

0.40

0.30

0.20

0.10 Enzyme [nkat activity mg

0.00 0 127142130 Time of leaf degradation (days) Figure 3.25 Kinetics of cytosolic glutathione S-transferase (cGST) activity response measured, using CDNB as substrate, in N. obtusa cells exposed to F. sylvatica and Q. robur leaf litter degradation extracts (LLDEs). * Significantly different from the control, n=5, p < 0.05.

3.1.3.3. Catalase

The 30 days exposure of N. obtusa cells to F. sylvatica and Q. robur LLDEs showed neither significant increase nor decrease in catalase activity. An increase of CAT activity was observed in cells exposed to 2 day old F. sylvatica LLDE, however, at increasing leaf degradation time, starting from day 7 a slight decrease in CAT activity in N. obtusa cells was observed, though without significant differences to the respective controls (Figure 3.26) Cells exposed to second day Q. robur LLDE showed the increased CAT activity by a factors 1.6, but CAT activities decreased by factors 1.75 and 2.1 compared to the control after 21 and 30 days of leaf degradation, respectively (Figure 3.26).

600.0 CAT Control 500.0 Beech

protein] Oak

-1 400.0

300.0

200.0

100.0 Enzyme activity [nkat mg [nkat Enzyme activity

0.0 0 1 2 7 14 21 30 Time of leaf degradation (days) Figure 3.26 Kinetics of catalase (CAT) activity measured, in N. obtusa cells exposed to F. sylvatica and Q. robur leaf litter degradation extracts (LLDEs). * Significantly different from the control, n=5, p < 0.05.

3.1.3.4. Glutathione reductase

The GR activity in N. obtusa cells increased significantly compared to control when exposed up to 7 day old F. sylvatica and Q. robur LLDEs, whereas no significant difference in GR activity was detected in the cells exposed to 21- and 30-day old F. sylvatica and Q. robur LLDEs (Figure 3.27). The glutathione reductase activity in the algae preparations increased significantly by factor 1.4 to 2.1 and by factors 1.3 to 1.8 in cells exposed to LLDEs of F. sylvatica and Q. robur degradation extracts in 30 day period (Figure 3.27). The highest elevation by a factor 2.1 was observed in N. obtusa exposed to 7 day old F. sylvatica LLDE containing 166.5 mg L-1 of DOC (Figure 3.27; Table 1) and the lowest one by a factor 1.4, in cells exposed to 2 day old LLDE containing 94.8 mg L-1 of DOC. After 2 h of exposure to Q. robur LLDE the highest GR activity was detected by a factor 1.8 in exposures to 7 day old LLDE and the lowest by the factor 1.5 in cells exposed to 2 day old LLDE (Figure 3.27; Table 1).

3.50 GR *

3.00 Control Beech * * * * protein] 2.50 Oak -1 * *

2.00

1.50

1.00

0.50 Enzyme activity [nkat mg [nkat activity Enzyme

0.00 0 1 2 7 14 21 30 Time of leaf degradation (days) Figure 3.27 Kinetics of glutathione reductase (GR) activity measured, in N. obtusa cells exposed to F. sylvatica and Q. robur leaf litter degradation extracts (LLDEs). * Significantly different from the control, n=5, p < 0.05.

3.1.3.5. Hydrogen peroxide content

The intracellular hydrogen peroxide content significantly decreased over controls by a factor ca. 2.2 in N. obtusa exposed to 1 day old F. sylvatica LLDE

(Figure 3.28). Significant decreases of H2O2 content were observed by a factor 9.1 in treatments exposed to 30 day old Q. robur LLDE. Other N. obtusa samples exposed to differently aged Q. robur LLDE also showed decrease in H2O2 concentration, however, it was not significant compared to control (Figure 3.28).

0.10 H O Control 0.09 2 2 Beech 0.08 Oak 0.07 per cell)

-1 0.06 * 0.05

0.04 (µmolcm

2 *

O 0.03 2 H 0.02

0.01

0.00 0 12 7142130 Time (days) Figure 3.28 Alterations of intracellular hydrogen peroxide in N. obtusa cells exposed to F. sylvatica and Q. robur leaf litter degradation extracts (LLDEs).* Significantly different from the control, n=5, p < 0.05. 63

3.2. The electrophysiological response of N. obtusa cells affected by F. sylvatica and Q. robur leaf extracts

F. sylvatica leaf extract (BLE) (33-1100 mg L-1 DOC) induced a rapid cell depolarization during the first 15 min, however, cells that were treated with 33-245 mg L-1 DOC of BLE tended to restore their initial RP values within 3-24 h period of prolonged treatment exposure (Figure 3.29a). Similar rapid depolarization (25-60% of control level) of the cell resting potential during the first 15 min was induced, when algae cells were treated with Q. robur leaf extract (OLE), which remained relatively unchanged until the end of 24 h exposure (Figure 3.29b). Small depolarization (up to approximately 10%) could be indicated at 73.5 and 245 mg L-1 DOC, while stimulation of electrogenic activity of the cells was expressed by 10-20% hyperpolarization at 7.35 and 24.5 mg L-1 DOC, the lowest tested OLE concentrations (Figure 3.29b).

100 a) b) 1100 mg/L 75 550 1837.5 mg/L

770 1225 50 245 900

735 25 110 330 245 77 0 73.5 Cell % m embrane depoliarization, 33 7.35

24.5 -25 0612182406121824 0 6 12 18 24 Time, h Time, h Figure 3.29 Average of cell resting potential induced by F. sylvatica (a) and Q. robur (b) leaf extracts, n≈120 for each treatment.

Figure 3.30 a and b present kinetics of RP response of N. obtusa cells to F. sylvatica and Q. robur leaf extracts with different DOC concentrations during 24 h experiment and 90 min rewash in APW solution. Results reported in Figure 3.30a showed two different sets of rewash curves. In the first set, we

64 observed effecting 90 min rewash from BLE up to 330 mg L-1, in other set, BLE concentrations from 550 mg L-1 DOC, we determinate the rewash till 25- 60% of control levels. It is interesting to note that the rewash and reached RP level which was only 12% less than that of controls, except the highest DOC (1837.5 mg L-1) (Figure 3.30b). It was observed that the rewash from Q. robur leaf extract was better than from F. sylvatica.

100 a) Rewash b) Rewash 75

1100 mg/L 50 550

1837.5 mg/L 25 770

RP depoliarization, % RP depoliarization, 735 1225 900 0 330 245 245 73.5 110 33 7.35 77 24.5 -25

0524(0)120524 (0)12 Time, h Time, h Figure 3.30 The rewash response (starting at 24 (0) h) of algae cells exposed to F. sylvatica (a) and Q. robur (b) leaf extracts, n≈120 for each treatment.

3.3. The lethality response of N. obtusa cells exposed to F. sylvatica and Q. robur leaf extracts

All charophyte cells were able to survive in all tested concentrations of F. sylvatica (5.5–1100 mg L-1 DOC) and Q. robur (24.5–1225 mg L-1 DOC) leaf extracts for one and two days, respectively (Figure 3.31 a–b). In general, F. sylvatica leaf extract was more toxic than that of the Q. robur. After 4-day exposure, mortality of the cells treated with BLE exceeded 20% at 61– 1100 mg L-1 DOC concentrations (Figure 3.31a), while the same effect level for OLE was reached only at the highest 1225 mg L-1 DOC concentration (Figure 3.31b). Within 8 days, contrary to OLE, the mortality curves for the cells treated with BLE have already reached relatively stable level (61-1100 mg L-1 DOC), which did not further increased significantly until the

65 end of experiment at the 42nd day, though slight mortality increase was detected at the lowest BLE concentrations (5.5–55 mg L-1) within 30–42 days (Figure 3.31a). The difference in toxicity between F. sylvatica and Q. robur leaf extracts was evident starting from the second day and throughout the -1 exposure duration (30-d LC20 = 55 and 490 mg L , respectively). During 42 day-long mortality experiment the controls stayed unchanged up to 40 days (Figure 3.31 a-b).

100 1100 mg/L a) 367.5 b) 1225 mg/L 245 735 490

980 735 122.5 50 Mortality, % Mortality, 61

490 55 27.5 245 5.5 24.5 Control Control 0

1101004 8 20 30 40 14 8 1020 30 40 100

Time, days Time, days Figure 3.31 The lethality response of algae cells exposed to F. sylvatica (a) and Q. robur (b) leaf extracts, n=40 for each treatment.

In order to detour data on amounts of DOC in the natural habitats of Charophytes, water samples were collected from various lakes of Lithuania. DOC in Lakes variated from 7.2 to 35.1 mg L-1 (Table 2). N. obtusa are generally found in clear and oligo- or mesotrophic waters (PYBUS et al., 2003). N. obtusa is extremely intolerant to presence of DOC in high concentrations, as known that in Lake Žuvintas (DOC - 20.4 mg L-1), during 3 last decades the algae N. obtusa almost disappeared (SINKEVIČIENĖ, 2007) (Table 2).

No. Lake name DOC, mgL-1 1. Smalvas 7.2 2. Skritelis 9.9 3. Tauragnas 9.9 4. Tapeliai 10.3 5. Kampinis 11.3 6. Seirijis 11.4 7. Juodis 11.6 8. Metelys 11.6 9. Burgis 11.9 10. Dusia 12.2 11. Platelių 12.7 12. Alksnas 13.2 13. Rubikių 13.3 14. Vilkas 14.3 15. Burgailis 14.5 16. Germantas 15.5 17. Žuvintas 20.4* 18. Juodikis 27.0** 19. Bedugnis 35.1**

Table 2 Dissolved organic carbon concentrations (mg L-1) in Lithuanian lakes (*disappearing and ** not growing N. obtusa).

4. Discussion

Oxidative stress is produced in cells by ROS resulting from cellular metabolism and from interaction with cells of exogenous sources (ALSCHER et al., 2002, BLOKHINA et al., 2003). Throughout the degradation of leaves, a lot of chemical compounds are set free into water, which may promote oxidative stress via generation of ROS and/or by disturbance of the intracellular oxidative balance in aquatic organisms (BLOKHINA et al., 2003). Oxidative stress can be detected by measuring activities of oxidative stress enzymes, such us GST, CAT, GR, GPx, SOD etc. We investigated dose-response relationships for the activities of oxidative stress enzymes in macrophytic algae cells of N. obtusa modulated by F. sylvatica and Q. robur, leaf extracts after 24 h exposure at 0.11–55.0 mg L-1 and 0.245–122.5 mg L-1 DOC, respectively. Under 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 (FOYER et al., 1994). It is known that when plants are exposed to

67 stress conditions there is an increase in ROS (PFLUGMACHER, 2004), and then the antioxidative ability of the plant may be exhausted. Organelles such as the peroxisomes and chloroplasts, where ROS are also produced under ‘normal’ conditions, are especially at risk (ALSCHER, 2002). Overproduction of ROS in chloroplasts of plants under drought-stress has been reported (PRICE et al., 1989). It might be suggested that a similar mechanism occurs when plant cells are exposed to certain concentrations of leaf extracts. To combat excess of ROS, plants have developed an antioxidative defense system, which includes enzymes such as superoxide dismutases, glutathione S-transferases, glutathione reductase, catalase, glutathione peroxidase etc. Within a cell, the superoxide dismutases, the family of metalo- enzymes, constitute the first line of defence against ROS (ALSCHER et al.,

2002). According to summarizing redox cycling scheme (DI GIULIO et al., 1989) (Figure 1.4), SOD catalyses the disproportionation of superoxide 2 O•- to molecular oxygen and H2O2. SOD removes superoxide and hence decreases the risk of hydroxyl radical formation from superoxide via the metal-catalysed

Haber–Weiss-type reaction (ARORA at al., 2002). In our study, SOD activity in N. obtusa cells exposed to F. sylvatica extract was modified by significant suppression at all tested DOC concentrations whereas Q. robur leaf extract was effective just at the highest, 122.5 mg L-1 DOC concentration. Indirectly, the absence of suppressive effect on SOD at lower concentrations of Q. robur extract coincides with previously obtained data on green algae

Scenedesmus armatus (Chod.) Chod. (AKSMANN, TUKAJ, 2004). It was shown that neither anthracene nor phenanthrene cause significant changes in the activity of superoxide dismutase in algal cells after 24 h exposure to 0.7- 5.6 µM concentrations, although these hydrocarbons strongly inhibit their growth. These ring organic structures are similar to structures such as ellagic acid found in Q. robur leaf extracts (BIANCO, SAVOLAINEN, 1997). Another enzyme invariably involved in cellular biotransformation when plants are faced with toxic substances is GST. It participates in the conjugation of toxicants or xenobiotics to the sulphurhydryl (–SH) group of glutathione 68

(GSH), which enhances their water solubility and therefore their excretion from cells through cell membranes, incorporation into cell wall or compartmentation into vacuoles or vessicles. Increase in GST activity is thus considered as a stress signal. In our study, we detected a significant increase in cGST and mGST activities in N. obtusa at 1.1-11.0 mg L–1 DOC of F. sylvatica leaf extracts and 5.5 mg L–1 DOC of Q. robur leaf extract (Grigutyte et al., 2009). Previous investigations showed that humic substances (HS) and natural organic matter do have an influence on the physiology, namely on photosynthesis, biotransformation and antioxidant enzymes in plants (PFLUGMACHER et al., 2001, PFLUGMACHER et al., 2003). MENONE and

PFLUGMACHER (2005) demonstrated that cytosolic GST activity in C. demersum was significantly elevated when treated with concentrations (up to 0.5 mg L-1) of the organic PCB compound 3-chlorobiphenyl. Likewise, herbicide diclofop-methyl caused a rapid and sustained increase in GST activity, in Euphorbia esula L. (ANDERSON, DAVIS, 2004). Similarly, GST activity was significantly increased for untreated E. esula plants at 5.0 mM paraquat and for 2-aminoethanol pre-treated plants at 0.1–5.0 mM paraquat

(MASCHER et. al., 2005). The organic PCB structures are similar to structures that can be found in leaves (BIANCO, SAVOLAINEN, 1997). 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, LEAVER, 1993). After GSH has been oxidized to GSSG, the recycling of GSSG to GSH is accomplished mainly by the enzyme glutathione reductase. An elevated GR activity measured in the present study was a further indication of an oxidative stress condition in exposed to F. sylvatica and Q. robur leaf extracts N. obtusa cells. 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

69 to enhance resistance to oxidative stress in Brassica juncea (L.) Czern. (ZHU et al., 1999). A similar increase of GR activity, enzyme responsible for maintaining the glutathione homeostasis in the cell, found in our study in N. obtusa has been previously reported for C. demersum after exposure to 3-chlorobiphenyl

(MENONE, PFLUGMACHER, 2005). An increase of GR has been shown in

E. esula plants treated with herbicide diclofop-methyl (ANDERSON, DAVIS, 2004). Similarly, herbicide flumioxazin increased activity of glutathione reductase too (SHAIKH et. al., 2006). Increases in GR activity were also found in maize leaves incubated with paraquat (MALAN et al. 1990). It is known that various structurally dissimilar xenobiotics are capable to induce an oxidative stress in plants. Some of these xenobiotic structures are similar what can be found in Q. robur leaf extracts, so increase of GR activity in N. obtusa exposed to the extracts clearly shows the enhanced protection against oxidative stress in these charophyte algae.

The activity of CAT, an enzyme that detoxifies H2O2 in the cells (Figure 1.4), increased significantly when charophyte algae calls were treated by 1.1 mg L–1 DOC, decreased by factor 1.2 at F. sylvatica leaf litter extract DOC concentrations (5.5-55 mg L–1). The significantly increased CAT activity in N. obtusa exposed to Q. robur leaf extract was found at the lowest tested concentrations, 0.245 and 2.45 mg L-1 DOC. Data, confirming the suppression of CAT in other plants: decrease in Betula pubescens Ehrh. leaves by factor 1.6

(RUUHOLA, YANG, 2006); decrease by 48 % in B. pubescens leaves treated with paraquat (ITURBE-ORMAETXE et. al., 1998). Paraquat, a well known promoter of oxidative stress, is an aromatic structure similar to that which might also be found in leaf extracts. Diverse effects on catalase activity were observed in Arachis hypogaea L.: a significant decrease when exposed to salicylic acid, flumioxazin and imazapic (herbicides), and the increase caused by phorate (SHAIKH et. al., 2006). ROS are extremely reactive and unstable chemical species, which react with proteins, nucleic acids, carbohydrates, and lipids in cells. The latter often 70 results in lipid peroxidation. Malondialdehyde is one of the end products of lipid peroxidation (NOWAK, JANCZAK, 2006). The findings in the present study did not reveal any significant increase in MDA levels in exposed N. obtusa in all concentrations tested. This is presumably due to activation of defence and detoxification mechanisms, as evidenced by high GST, GR and CAT activities. These antioxidant enzymes can enhance the removal of ROS and thus prevent damage of cellular constituents and initiation of lipid peroxidation (YANG et al., 2003). The other part of study was to monitor enzyme kinetics in order to assess the primary response of enzymatic activity of N. obtusa to leaf litter extracts. Hence, N. obtusa cells were exposed for the following durations: -1 15 min, 30 min, 1h, 1.5 h, 2, 4 and 24 h to 1.1 and 2.45 mg L DOC of F. sylvatica and Q. robur extracts, respectively, because the strongest effects were, in most cases, observed at these concentrations. The biotransformation enzyme, glutathione S-transferase, was significantly increased after 15 min of exposure to F. sylvatica extract. The onset of GST response in N. obtusa upon exposure to Q. robur extract was statistically different from control at all tested time durations. mGST activity of the algae exposed to F. sylvatica extract significantly decreased during 1 h to 4 h and subsequently returned to control values in 24 h (GRIGUTYTĖ et al., 2008).

As observed in previous study (KAMARA, PFLUGMACHER, 2007), mGST activity in C. demersum affected to P. australis and Q. robur extracts for longer exposures (48 h and 168 h) returned to control values, so plants may accommodate to oxidative stress generated by leaf extracts. GR represents the exceptional case in the enzymatic response of charophyte algae, since F. sylvatica extract provoked a notably sooner (after 1 h) elevation in its activity than did Q. robur extract (after 4 h). Comparison of enzymatic response (GST, CAT and GR) as a whole suggests that N. obtusa was more sensitive to F. sylvatica than to Q. robur leaf litter extract and therefore could invoke an earlier de novo biosynthesis of GSH.

One of the study approaches consisted of imitating as much as possible the environmental conditions, thereby 30-day-decomposing leaf litter extracts from F. sylvatica and Q. robur were used. We observed more than double increase of soluble GST activity in the macroalgae N. obtusa exposed for 2 h to F. sylvatica and Q. robur leaf litter extracts that were obtained within a whole 30-day period of leaves decay. Similar statistically significant sGST elevation was found in C. demersum and Lemna minor L. exposed for 24 h to artificial -1 extracts from Q robur and P. australis (0.1–100 mg L DOC) (KAMARA,

PFLUGMACHER, 2007; KAMARA, 2008). In parallel to the stimulation of sGST measured in the present study, glutathione reductase was also found to be significantly increased upon exposure of N. obtusa cells to F. sylvatica and Q. robur LLDEs of up to 14-day-old. Similarly, paraquat increased GR activity in Pisum sativum L. (MADAMANCHI et al., 1994) and in Anabaena cylindrica

A. Juss. (EKMEKCI, TERZIOGLU, 2005). The stimulation of GST and GR activities in N. obtusa cells detected in our survey gives the impression that the LLDEs have an effect on the biotransformation system of charophyte algae similar to that on mentioned above aquatic and terrestrial plants. In our experiments, we were not able to demonstrate any significant alteration of CAT activity against the simultaneous control in algal cells exposed to the extracts of both leaf litters that have been decaying from 1 to 30 days.

Similarly, the content of H2O2 measured in the cells treated by LLDEs showed a scarce potential to correlate clearly the catalase activity and the leaf litter degrading time or to differ from the respective controls. Nevertheless, the tendency of decreased H2O2 concentrations during the first 7 days for F. sylvatica and during the latest 15 days for Q. robur LLDEs of 30 day degradation period could be confirmed statistically, thus it might be suggested that the observed minor changes of H2O2 content were due to LLDEs-induced decrease in SOD activity. This suggestion is further strengthened by the finding in previous study (GRIGUTYTĖ et al., 2009; GRIGUTYTĖ et al., in press), when SOD activity in N. obtusa cells exposed to Q. robur leaf extract has been inhibited significantly at the DOC concentrations above 100 mg L-1 (in the 72 present study, DOC ranged from 95–166 and 86–177 mg L-1, respectively for F. sylvatica and Q. robur LLDEs). Additionally, the explanations that the absence of the increase in H2O2 production under the affect of leaf litter degradation extracts could be due to some other ROS, might not be excluded. In general, our results indicate that leaf extracts from F. sylvatica and Q. robur, have evoked oxidative stress in green algae N. obtusa, reflected in activity alterations of such enzymes as SOD, CAT, GST and GR. Surprisingly, relatively low DOC concentrations that normally can be found in lakes were able to induce oxidative stress. DOC concentrations in Lithuania’s freshwater lakes with prospering macrophytes vary from 7 to 21 mg L-1. When there is a higher concentration as, for example, in Lake Žuvintas (20.4 mg L-1), algae became almost extinct during last 30 years, although they were growing abundantly earlier. We demonstrated that in some cases enzyme activities started to decrease at higher DOC concentrations indicating an intense impact on the antioxidative ability of algae. Since the enzymatic response observed within first 24 h significantly differs from activity of enzymes in normal conditions, it can be suggested that the antioxidative system in N. obtusa was sufficiently activated to prevent or minimize oxidative stress damage. This, fairly active algae antioxidative system is also confirmed by the fact, that no significant effect on lipid peroxidation was detected in leave extract-treated algae. To supress stress, the antioxidative system, which functions in N. obtusa responded relatively sooner to F. sylvatica leaf extracts than to Q. robur ones. The effects have a tendency to decrease to control levels at elevated DOC concentrations (mainly after exposure for 24 h), thus representing a limit on the antioxidative ability of the charophyte N. obtusa. A strong increase of sGST and GR activity observed in N. obtusa might be caused by the oxidation of phenolic compounds that are being released as soluble organic materials during leaf degradation in the water already within the first 48 h (POPE et al., 1999). Overall, this study indicates that the antioxidative system in N. obtusa cells was activated when these algae get in contact with investigated leaf litter and it’s decomposing extracts. 73

During the research of effects of leaf extracts at higher than enzymatic regulation level, i.e. at the level of electrochemical regulation of membrane transport, we have found that charophyte cell of N. obtusa treated with Q. robur and F. sylvatica leaf extracts reacted by a rapid (15-30 min) concentration-dependent cell membrane depolarization, which was relatively stable during 24 h in Q. robur and at least 3-4 h in F. sylvatica leaf extract. Under experimental conditions, the average value of measured resting potential of the cells equaled -220±5 mV (mean±sd, n=240) in control solution, which represents a normal physiological state of plasma membrane with electrically evident component of electrogenic ion pump (P-state). In charophyte cells this electrogenic component, in addition to the diffusion potential of K ions across the plasma membrane, is known to be related with the operating of the enzymatic H+-pump, particularly K+, Mg2+-specific H+-ATPase

(MIMURA, 1995). It was shown by ecotoxicological studies, that depolarization of N. obtusa cells affected by chemical stressors is tightly correlated to inhibition of K+, Mg2+-ATPase and later observed cells mortality

(MANUSADŽIANAS et al., 2002, 2007). Thus, alterations of membrane bioelectrical parameters serve as a relevant indicator of adverse impacts on the cell homeostasis. Our results show, that by exposing N. obtusa cells to F. sylvatica and Q. robur leaf extracts we get different electrophysiological responses, ones are reversible and the others are not. Reversibility was observed during the exposure to low concentrations of both extracts. This tendency was more obvious at rewashing cells after 90 min. exposure. However, at higher than natural (770-1100 and 1837 mg L-1 DOC) concentrations, level of residual depolarization suggests that electrogenic ion pumps were inhibited. Reactions of algae N. obtusa to Q. robur and F. sylvatica extracts were found at different biological levels. Reactions at enzymatic level when cells were exposed to DOC concentrations common to natural water bodies, were observed after 15 min. Cell membrane reactions were also observed within 15 min, but effects were evident only at higher concentrations (from 550 74 mg L-1 DOC). Cell lethality was evoked by similar concentrations, like in electrophysiological experiments: larger than 20% RP depolarization was observed at higher than 77 mg L-1 DOC concentration of F. sylvatica extracts, and this correlated with higher than 20% lethality at 61 mg L-1. The ecological importance of chemical stressor proves to be evident when environmental conditions become impossible to exist for particular aquatic species. In our case, it was known that macrophytes being indicators of oligotrophic freshwater bodies, do not grow in lakes with higher organic carbon concentration. In the study, we succeeded to show that changes in oxidative stress enzyme activities and cells membrane potential evoked by F. sylvatica and Q. robur extracts relate with cells mortality, thus presuming a possible causality between more sensitive earlier reactions and later occurring lethality. Herewith it was concluded that leaf litter may be important environmental factor, which has impact on diversity of vegetation in freshwater ecosystems.

Conclusions

1. Q. robur leaf water-extracts induced statistically significant elevation of cytosolic (sGST) and microsomal glutathione S-transferase (mGST), glutathione reductase (GR) and glutathione peroxidase (GPx) activities in N. obtusa cells at all tested DOC concentrations (from 0.245 to 122.5 mg L-1 DOC, 24 h exposure). The activity of catalase (CAT) increased at 0.245 and 2.45 mg L-1 DOC, and SOD was inhibited only at the highest tested DOC concentrations. Time-dependent exposure during 24 h to the extract at 2.45 mg L-1 DOC revealed a decrease in mGST, increase in sGST and the lack of changes for CAT activities. A significant increase in GR activity was detected after 4 h and lasted until the end of 24 h exposure. 2. F. sylvatica leaf water-extracts induced significant increase in sGST activity in algal cells exposed for 24 h to 1.1, 5.5 and 11.0 mg L-1 DOC,

while mGST was activated only at 5.5 mg L-1 DOC. The activity of CAT increased at 1.1 mg L-1 DOC, however its inhibition was observed as the concentration increased from 5.5 to 55 mg L-1 DOC. The activities of SOD and GPx were decreased and GR increased at all DOC concentrations tested (0.11-55 mg L-1 DOC). Time-dependent exposure during 24 h to the extract at 1.1 mg L-1 DOC showed a significant increase in sGST and CAT, as well as GR, however, 1 h later. In case of mGST, inhibition of the enzyme was evident within 1 to 4 h. 3. Both leaf extracts – Q. robur (245 – 1838 mg L-1DOC) and F. sylvatica (330–1100 mg L-1 DOC) – induced a fast depolarization of cell membrane of N. obtusa already after 15 min exposure, which was evident up to the end of 24 h exposure. The extracts characterised by the DOC concentrations attributive to natural freshwater bodies induced similar fast depolarization of RP, however, electrophysiological reactions demonstrated adaptive response character. A better recovery from Q. robur than of F. sylvatica leaf extracts within a 1.5 h in control solution was observed and it was related with the mortality of N. obtusa cells – they were able to survive for 1 and 2 days in F. sylvatica and Q. robur leaf extracts, respectively. In general, F. sylvatica leaf extract was more toxic than Q. robur: at 245 mg L-1 DOC, larger than 50 % mortality was reached after 4-day exposure in F. sylvatica leaf extract, while the same concentration of Q. robur leaf extract was nontoxic. 4. Responses of algae N. obtusa to Q. robur and F. sylvatica leaf extracts were detected at different biological levels. Changes in activities of oxidative stress enzymes and cell membrane potential invoked by leaf extracts predicted cell mortality, which allows us to presume causality between more sensitive initial reactions and lethality. Leaf litter degradation extracts obtained from Q. robur and F. sylvatica leaves during 30 days of their decomposing enhanced oxidative stress response in N. obtusa cell, too. These findings suggest that that leaf litter might 76

be an important environmental factor impacting aquatic vegetation species diversity in freshwater ecosystems.

Acknowledgements

I am grateful to my scientific supervisor Dr. Levonas Manusadžianas for all care, support and for invaluable suggestions, discussions and help with the preparation of the doctoral thesis. I am grateful to my scientific advisor PD. Dr. Stephan Pflugmacher for advices and guiding in the biochemical studies. I thank Dr. Jorge Nimptsch for useful lessons in laboratory, for valuable remarks in writing my publications and for advice on statistical data analysis. I thank prof. Dr. Claudia Wiegand and Dr. Jan Peter Lay for the support and encouragement in doctoral studies and continuous support. My thanks to all the colleagues in the research group of biochemical regulation and laboratory of hidrobotany, especially to dr. Sheku Kamara, dr. Anja Peuthert, dr. Rolandas Karitonas, dr. Valeska Contardo, Viola Viehmann, Julija Stefanovič, Irma Vitonytė, Jūratė Karosienė for advices, discussions, help and good environment in the laboratory. I would like to express my appreciation to Dr. Sigutė Jurkonienė and Dr. Kazys Sadauskas for the academic advice and support. Finally and the most important thanks go to my all family, boyfriend Tomas, his family. My friends, specially Inga and Donatas Antanevičiai, and Agnė Stonkutė, Laura Steponienė for their patience, help, understanding, concern and support. The work presented in this thesis was carried out in the Institute of Botany and Leibniz-Institute of Freshwater Ecology and Inland Fisheries. Special thanks to chemical laboratory at the IGB for carrying out the DOC analysis. The work was financial supported by the Deutsche Bundesstiftung Umwelt (DBU, grant No. 19000/059) and Lithuania Science Foundation (LMSF).

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Research articles and congress presentations

Research articles: Steinberg C. E. W., Manusadžianas L., Grigutytė R., Karitonas R., Jurkonienė S., Pflugmacher S., 2004: Membrane depolarization and elevation of ROS-defensive mechanisms due to the impact of dissolved natural organic matter (NOM) in the charophyte Nitellopsis obtusa. Humic Substances and Soil and Water Environment. ISBN 85-86463-12-4. Martin-Neto, L, Milori, D. M. B., Lopes da Silva, W. T. (eds). Proc. XII Intern. Meeting of IHSS, São Pedro, São Paulo, Brazil, July 25–30, Embrapa Instrumentação Agropecuária: 135–137. Manusadžianas L., Steinberg C. E. W., Grigutytė R., Jurkonienė S., Karitonas R., Pflugmacher S., 2005: Effects of humic substances on charophyte Nitellopsis obtusa at the stress enzyme, cell membrane and cellular endpoints – 12th International Symposium on Toxicity Assessment, – Proceedings. Ed. A. Kungolos: 73. ISBN 960-99067-6-3 Grigutytė R., Nimptsch J., Manusadžianas L., Pflugmacher S., 2008: Effects of decomposing leaf litter from Fagus sylvatica and Quercus robur of different degradation levels on oxidative stress response in the charophyte Nitellopsis obtusa. Botanica Lithuanica, 14 (4): 233–240. Grigutytė R., Nimptsch J., Manusadžianas L., Pflugmacher S., 2009: Response of oxidative stress enzymes in charophyte Nitellopsis obtusa exposed to allochthonous leaf extracts from beech Fagus sylvatica. – Biologija, 55 (3– 4): 142–149.

Congress presentations: Grigutytė R., Manusadžianas L., Pflugmacher S.: Promotion of oxidative stress response in Nitellopsis obtusa during exposure to leaf extract from oak and beech. 10th Society of Environmental Toxicology and Chemistry (10 SETAC), Basel, Switzerland, 28–30 September 2005, German Language Branch. Manusadžianas L., Grigutytė R., Karitonas R., Sadauskas K., Vitkus R.: Combining of exposure and recovery parameters in charophyte cell response as an approach to reveal remote toxic effects. III tarptautinė metaloekologų konferencija: Metalai aplinkoje, – Vilnius, 2006. Grigutytė R., Manusadžianas L., Pflugmacher S.: Promotion of oxidative stress response in Nitellopsis obtusa during exposure to leaf extract from oak and beech. SETAC Europe 16th Annual Meeting, The Hague, The Netherlands, 7–11 May 2006. Grigutytė R., Pflugmacher S., Manusadžianas L.: Electrophysiological and survival responses of the charophyte cell of Nitellopsis obtusa during prolonged exposure to leaf extract from oak Quercus robur. Society of Environmental Toxicology and Chemistry, Leipzig, Germany, 12–14 September 2007, German-Language-Branch. Manusadžianas L., Grigutytė R., Pflugmacher S.: The impact of oak and beech leaf extracts on freshwater charophyte cell Nitellopsis obtusa. International Symposium on Green Chemistry for Environment and Health, – Munchen, Germany, 13–16 October 2008.